We thank Dr. J. Vergara (University of California, Los Angeles) for sharing the EGFP-CaV1.1 (rabbit) wild-type plasmid. We thank the Yale Department of Physiology Electronics Laboratory and especially Henrik Abildgaard for the design and construction of the photodiode with track and hold circuit. This work was supported by the National Institutes of Health grants R01-AR075726 and R01-NS103777

Functional Site-Directed Fluorometry in Native Skeletal Muscle Cells

The authors report no conflict of interest.

Here, a step-by-step protocol to conduct FSDF in muscle fibers for the study of individual voltage sensor motions from the CaV1.1 channel is described. Even though the number of steps and the diversity of approaches that are combined in this technique may appear complex, most of these techniques are often routinely used in biophysicist/cell biologist laboratories. Thus, the apparent complexity resides mainly in the combination of all the various approaches in a single integrated technique. Often when carrying out a multi-step method, the impact of small modifications conducted at the beginning are only detectable later in the following steps. With rigor in the development of each step, and precise inclusion and exclusion criteria, it is possible to obtain multiple recordings from the same preparation or even the same fiber.
The approaches described here could be adapted to other cell types, including cardiac cells and neurons. In addition, different detection systems, such as high-speed cameras, could be used. Currently, the use of voltage clamp (patch clamp or two-electrode voltage clamp) is limited due to the additional level of complexity intrinsic to these techniques44. Improvements on the method presented here (such as brighter and more photostable probes) will facilitate the concurrent use of FSDF with voltage-clamp methods.
Field stimulation in native dissociated skeletal muscle fibers has the advantage of having a higher throughput than voltage-clamp and can be applied for the study of other signals, such as action potential propagation45 or action potential-evoked calcium transients41.
The main advantage of this native cell FSDF when compared to the heterologous expression system is the ability to detect protein conformational changes in response to induced depolarization in its physiological environment and by its endogenous stimulus. It is especially relevant for the study of CaV1.1, which acts as the voltage sensor for another calcium channel, the ryanodine receptor type 1 (RyR1), inserted in the distinct sarcoplasmic reticulum membrane domain5,33,35. In practice, it could be possible to study the simultaneous measurement of fluorescent signals from a voltage sensor with calcium release from the sarcoplasmic reticulum using a suitable calcium fluorescent indicator. Moreover, since it has been shown that many proteins from the triadic environment could impact CaV1.1 gating46,47,48,49,50, the study of the voltage sensor motion in a muscle fiber is of high physiological relevance. Two major disadvantages of the approach are: 1) small amplitude of the measured signals, and 2) photobleaching, which precludes the use of repetitive and/or prolonged stimuli. Improvements to this technique are needed and will allow to combine FSDF with voltage clamp experiments in adult muscle fibers.
The application of FSDF to other channels (or receptors) in muscle fibers is feasible, provided that: (1) they are expressed at a sufficient level, (2) they are accessible (from extracellular or intracellular space), and (3) the cysteine modification via mutagenesis does not impair the channel function. The complementary use of FSDF in both heterologous and native muscle fibers is necessary and will be critical for addressing unanswered questions regarding voltage dependent CaV1.1 transitions that are critical for muscle function.

The ability to track ion channel conformational rearrangements in response to a known electrical stimulus in a living cell is a source of valuable information for molecular physiology1. Voltage-gated ion channels are membrane proteins that sense changes in transmembrane voltage, and their function is also affected by voltage changes2. The development of voltage clamp techniques in the last century allowed physiologists to study, in real-time, ionic currents carried by voltage-gated ion channels in response to membrane depolarization3. The use of voltage clamp technology has been crucial in understanding the electrical properties of excitable cells such as neurons and muscle. In the 1970s, voltage clamp refinement allowed for the detection of gating currents (or charge movement) in voltage-gated calcium (CaV) and sodium (NaV) channels4,5. Gating currents are non-linear capacitive currents that arise from the movement of voltage sensors in response to changes in the electric field across the cell membrane6. Gating currents are considered an electrical manifestation of molecular rearrangements that precede or accompany ion channel opening7. While these current measurements provide valuable information regarding the channel&#39;s function, both ionic currents and gating currents are indirect readouts of inter- and intra-molecular conformational rearrangements of voltage-gated channels7.
Functional site-directed fluorometry (FSDF; also referred to as voltage clamp fluorometry, VCF) was developed in the early 1990s8 and, for the first time, provided the ability to directly view local conformational changes and the function of a channel protein in real time. Using a combination of channel mutagenesis, electrophysiology, and heterologous expression systems, it is possible to fluorescently tag and track the moving parts of specific channels or receptors in response to the activating stimulus9,10. This approach has been extensively used to study the voltage-sensing mechanisms in voltage-gated ion channels8,10,11,12,13,14,15,16,17,18,19. For authoritative reviews, see10,20,21,22,23.
The CaV and NaV channels, critical for the initiation and propagation of electrical signals, are composed of a main &#945;1 subunit, which possesses a central pore and four non-identical voltage sensing domains2. In addition to their distinct primary structure, CaV and NaV channels are expressed as multisubunit complexes with auxiliary subunits24. Voltage-dependent potassium channels (KV) consist of four subunits that look like a single domain of NaV or CaV25. The pore-forming and voltage-sensing &#945;1 subunit of CaV and NaV channels is formed by a single polypeptide coding for four individual domains of six unique transmembrane segments (S1-S6; Figure 1A)24,26. The region comprised by S1 to S4 transmembrane segments form the voltage sensing domain (VSD) and S5 and S6 transmembrane segments form the pore domain26. In each VSD, the S4 &#945;-helix contains positively charged arginine or lysine (Figure 1A,B) that move in response to membrane depolarization7. Several decades of research and the results from highly diverse experimental approaches support the premise that S4 segments move outward, generating gating currents, in response to membrane depolarization6.
FSDF measures the fluorescence changes of a thiol-reactive dye conjugated to a specific cysteine residue (i.e., the S4 &#945;-helix) on an ion channel or other protein, engineered via site-directed mutagenesis, as the channel functions in response to membrane depolarization or other stimuli10. In fact, FSDF was originally developed to investigate whether the S4 segment in KV channels, proposed to be the main voltage sensor of the channel, moves when the gating charges move in response to changes in membrane potential8,10. In case of voltage-gated ion channels, FSDF can resolve independent conformational rearrangements of the four VSDs (tracking one VSD at any given time), concurrently with channel function measurements. Indeed, using this approach, it has been shown that individual VSDs appear to be differentially involved in specific aspects of channel activation and inactivation12,27,28,29,30. Identifying the contribution of each VSD to the channels&#39; function is of high relevance and can be used to further elucidate channel operation and potentially identify new targets for drug development.
The use of FSDF in heterologous expression systems has been extremely helpful in furthering our understanding of channel function from a reductionist perspective10,23. Like many reductionist approaches, it presents advantages but also has limitations. For instance, one major limitation is the partial reconstitution of the channel nano environment in the heterologous system. Often, ion channels interact with numerous accessory subunits and numerous other proteins that modify their function31. In principle, different channels and their accessory subunits can be expressed in heterologous systems with the use of multiple protein coding constructs or polycistronic plasmids, but their native environment cannot be fully reconstituted30,32.
Our group recently published a variant of FSDF in native dissociated skeletal muscle fibers for the study of early steps of excitation-contraction coupling (ECC)33,34, the process by which muscle fiber electrical depolarization is linked to the activation of muscle contraction35,36. For the first time, this approach allowed the motion tracking of individual S4 voltage-sensors from the voltage-gated L-type Ca2+ channel (CaV1.1, also known as DHPR) in the native environment of an adult differentiated muscle fiber37. This was accomplished by considering multiple characteristics of this cell type, including the electrical activity of the cell allowing fast stimulation-induced self-propagated depolarization, the ability to express cDNA plasmid through in vivo electroporation, the natural high expression and compartmental organization of the channels within the cell, and its compatibility with high-speed imaging and electrophysiological recording devices. Previously, we used a high-speed line scanning confocal microscope as a detecting device37. Now, a variation of the technique is presented using a photodiode for signal acquisition. This photodiode-based detection system could facilitate the implementation of this technique in other laboratories.
Here, a step-by-step protocol to utilize FSDF in native cells for the study of individual voltage-sensor movement from CaV1.1 is described. While the CaV1.1 channel has been used as an example throughout this manuscript, this technique could be applied to extracellularly accessible domains of other ion channels, receptors, or surface proteins.

<table><tbody><tr><td>Hyaluronidase</td><td>SIGMA ALDRICH</td><td>H3884-50mg</td><td></td></tr><tr><td>0.5 mL Eppendorf tube</td><td>Millipore Sigma</td><td>EP022363719-500EA</td><td></td></tr><tr><td>1 mL syringe</td><td>Millipore Sigma</td><td>Z683531-100EA</td><td>tuberculine slip tip</td></tr><tr><td>1/2&amp;rdquo; long 29-gauge sterile insulin needle and syringe</td><td>Becton Dikinson</td><td>324702</td><td></td></tr><tr><td>35 mm non coated plastic plate</td><td>Falcon, Corning</td><td>353001</td><td></td></tr><tr><td>60 mm non coated plastic plate</td><td>Falcon, Corning</td><td>351007</td><td></td></tr><tr><td>Alcoholic whip</td><td>PDI</td><td>B60307</td><td></td></tr><tr><td>Alexa-533 cube LP</td><td>Chroma</td><td>49907</td><td>Ex: 530/30x; BS: 532; Em: 550lp</td></tr><tr><td>Arc lamp</td><td>Sutter Instrumets</td><td>LB-LS 672</td><td></td></tr><tr><td>Artificial tears cream</td><td>Akorn</td><td>NDC 59399-162-35</td><td></td></tr><tr><td>Borosilicate glass Pasteur pipet 5 3/4&quot;</td><td>VWR</td><td>14672-200</td><td></td></tr><tr><td>BTS (N-benzyl-p-toluene sulphonamide)</td><td>SIGMA ALDRICH</td><td>203895</td><td></td></tr><tr><td>collagenase type I</td><td>SIGMA ALDRICH</td><td>C0130-1g</td><td></td></tr><tr><td>Cotton tip</td><td>VWR</td><td>VWR-76048-960-BG</td><td></td></tr><tr><td>Double electrode array&amp;nbsp; (for electroporation)</td><td>BTX harvard apparatus</td><td>45-0120</td><td>10mm 2 needle array tips</td></tr><tr><td>EGFP cube</td><td>Chroma</td><td>39002AT</td><td>Ex: 480/30x; BS 505; Em: 535/40m</td></tr><tr><td>Electroporation apparatus device</td><td>BTX harvard apparatus</td><td>ECM 830</td><td></td></tr><tr><td>EPC10</td><td>HEKA Elektronik GmbH&amp;nbsp; (Harvard Bioscience)</td><td>895000</td><td></td></tr><tr><td>FBS</td><td>Biotechne,&amp;nbsp; R&amp;amp;D Systems</td><td>RND-S11150H</td><td>Fetal Bovine Serum - Premium, Heat Inactivated</td></tr><tr><td>glass coverslip 35 mm dish</td><td>MatTek Life Science</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>Isoflurane</td><td>Fluriso (Isoflurane) Liquid for Inhalation</td><td>502017-250ml</td><td></td></tr><tr><td>Isothermal heating pad</td><td>Braintree scientific inc</td><td>39DP</td><td></td></tr><tr><td>Laminin</td><td>Thermo Fisher</td><td>INV-23017015</td><td>Laminin Mouse Protein, Natural</td></tr><tr><td>Latex bulb</td><td>VWR</td><td>82024-554</td><td></td></tr><tr><td>LED 530 nm</td><td>Sutter Instrumets</td><td>5A-530</td><td></td></tr><tr><td>Low binding protein 0.2 &amp;mu;m sterile filter</td><td>Pall</td><td>FG4579</td><td>acrodisk&amp;nbsp; syringe filter 0.2um supor membrane low protein binding non pyrogenic</td></tr><tr><td>MEM</td><td>Invitrogen</td><td>INV-11380037</td><td></td></tr><tr><td>MTS-5-TAMRA</td><td>Biotium</td><td>89410-784</td><td>MTS-5-TAMRA</td></tr><tr><td>OriginPro Analysis Software</td><td>OriginLab Corporation</td><td>OriginPro 2022 (64-bit) SR1</td><td></td></tr><tr><td>Photodiode</td><td>Custom Made</td><td>NA</td><td></td></tr><tr><td>PlanApo 60x oil&amp;nbsp; 1.4 N.A/&amp;infin;/0.17</td><td>Olympus</td><td>BFPC2</td><td></td></tr><tr><td>Platinum wire 0.5 mm, 99.9 % metals basis</td><td>SIGMA</td><td>267228-1G</td><td>To manufacture field stimulation electrode</td></tr><tr><td>Pulse Generator</td><td>WPI</td><td>Pulsemaster A300</td><td></td></tr><tr><td>Shutter drive controller</td><td>Uniblitz</td><td>100-2B</td><td></td></tr><tr><td>Shuttter</td><td>Uniblitz</td><td>VS2582T0-100</td><td></td></tr><tr><td>S-MEM</td><td>Invitrogen</td><td>INV-11380037</td><td></td></tr><tr><td>Sterile bench pad</td><td>VWR</td><td>DSI-B1623</td><td></td></tr><tr><td>Sterile saline</td><td>SIGMA ALDRICH</td><td>S8776</td><td></td></tr><tr><td>Sylgard 184 Silicone Elastomer kit</td><td>Dow Corning</td><td>1419447-1198</td><td></td></tr><tr><td>Vaporizer for Anesthesia</td><td>Parkland Scientific</td><td>V3000PK</td><td></td></tr><tr><td>Voltage generator</td><td>Custom Made</td><td>NA</td><td></td></tr></tbody></table>

functional site-directed fluorometry in native cells to study skeletal muscle excitability

Functional site-directed fluorometry has been the technique of choice to investigate the structure-function relationship of numerous membrane proteins, including voltage-gated ion channels. This approach has been used primarily in heterologous expression systems to simultaneously measure membrane currents, the electrical manifestation of the channels&#39; activity, and fluorescence measurements, reporting local domain rearrangements. Functional site-directed fluorometry combines electrophysiology, molecular biology, chemistry, and fluorescence into a single wide-ranging technique that permits the study of real-time structural rearrangements and function through fluorescence and electrophysiology, respectively. Typically, this approach requires an engineered voltage-gated membrane channel that contains a cysteine that can be tested by a thiol-reactive fluorescent dye. Until recently, the thiol-reactive chemistry used for the site-directed fluorescent labeling of proteins was carried out exclusively in Xenopus oocytes and cell lines, restricting the scope of the approach to primary non-excitable cells. This report describes the applicability of functional site-directed fluorometry in adult skeletal muscle cells to study the early steps of excitation-contraction coupling, the process by which muscle fiber electrical depolarization is linked to the activation of muscle contraction. The present protocol describes the methodologies to design and transfect cysteine-engineered voltage-gated Ca2+ channels (CaV1.1) into muscle fibers of the flexor digitorum brevis of adult mice using in vivo electroporation and the subsequent steps required for functional site-directed fluorometry measurements. This approach can be adapted to study other ion channels and proteins. The use of functional site-directed fluorometry of mammalian muscle is particularly relevant to studying basic mechanisms of excitability.

When propagating action potentials are triggered in response to repetitive field stimulation, it is possible to track specific voltage sensor motion in response to a specific frequency of depolarization. As shown in Figure 6A, the motion of VSD-II-tagged helices can be tracked in response to each of two successive depolarizations applied at 10 Hz (i.e., spaced by 100 ms). Signal bleaching can be corrected by subtracting baseline to the trace (Figure 6B). Further time magnification on the first and second responses (Figure 6C) enables precise observation of these fast-developing VSD-II helix movements. The time marker from the field-stimulation trigger allows relative temporal alignment of the first and second responses at a precision of 10 &#181;s. Both signals can be normalized by their respective minimum value (i.e.,&#160;minimum peak) to allow a kinetic comparison between the two responses (Figure 6D). From these recordings of relative fluorescence, it can be observed that the time to peak is comparable between successive depolarizations for this voltage sensor.
This approach could be used to track individual voltage sensors&#39; motion with various frequencies of depolarization and multiple action potentials to study their activation and relationship with charge movement, calcium current, or calcium transients, which can be studied in parallel in a different set of fibers, as previously reported37.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65311/65311fig06.jpg" /> 
Figure 6: Representative fluorometric recording from EGFP-CaV1.1 VSD-II triggered by two successive depolarizations at 10 Hz. (A) &#916;F/F0 fluorometric signal in a fiber expressing EGFP-Cav1.1 VSD-II stimulated twice at 10 Hz and a baseline correction curve (purple). The top black line indicates induced depolarization. (B) &#916;F/F0 baseline corrected signal of the two independent responses (C). Values on the plot correspond to the minimum reach at the peak. (D) Signal alignment relative to depolarization and normalization by their respective minimum peak shows VSD-II domain motion with comparable kinetics for both stimuli. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65311/65311fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagents</td>
			<td>Final Concentration (mM)</td>
			<td>Molecular weight</td>
			<td>g/L</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>150</td>
			<td>58.44</td>
			<td>8.766</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>4</td>
			<td>74.55</td>
			<td>0.298</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>2</td>
			<td>110.9</td>
			<td>0.222</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>1</td>
			<td>95.22</td>
			<td>0.095</td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>5</td>
			<td>180.2</td>
			<td>0.901</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>10</td>
			<td>238.3</td>
			<td>2.383</td>
		</tr>
		<tr>
			<td colspan="4">pH adjusted to 7.4 with 1 M NaOH</td>
		</tr>
	</tbody>
</table>
Table 1: Modified Ringer&#39;s solution composition.

Watch this Scientific Journal Video about Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability at JoVE.com

Author Spotlight: Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability  

Functional site-directed fluorometry is a method to study protein domain motions in real time. Modification of this technique for its application in native cells now allows the detection and tracking of single voltage-sensor motions from voltage-gated Ca2+ channels in murine isolated skeletal muscle fibers.

Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability

Functional site-directed fluorometry has been the technique of choice to investigate the structure-function relationship of numerous membrane proteins, ...

functional-site-directed-fluorometry-native-cells-to-study-skeletal

Nanyang Technological University

Research

JoVE Journal

Biology

907 Views.  University of Maryland School of Medicine. CaV1.1, een lid van de spanningsafhankelijke ionkanalen, heeft vier verschillende spanningssensoren. Er zijn aanwijzingen dat sommige spanningssensoren meer bijdragen aan ryanodinereceptoractivering of calciumstroom. We streven ernaar om de precieze rol van elke spanningssensor in excitatie-contractiekoppeling en calciumkanaalactivering te kunnen identificeren.Excitatie-contractiekoppeling is sinds het begin van de jaren 50 bestudeerd, maar de moleculaire details van hoe dit proces pla...

Video: Auteur in de schijnwerpers: functionele plaatsgerichte fluorometrie in inheemse cellen om de prikkelbaarheid van skeletspieren te bestuderen  

Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for muscle regeneration. Upon injury, which occurs frequently in skeletal muscles, SCs are activated. They proliferate as myoblasts and differentiate to repair muscle lesions. Among many events that take place during muscle differentiation, cytosolic Ca2+ signals are of great importance. These Ca2+ signals arise from Ca2+ release from internal Ca2+ stores, as well as from Ca2+ entry from the extracellular space, particularly the store-operated Ca2+ entry (SOCE). This paper describes a methodology used to obtain a pure population of human myoblasts from muscle samples collected after orthopedic surgery. The tissue is mechanically and enzymatically digested, and the cells are amplified and then sorted by flow cytometry according to the presence of specific membrane markers. Once obtained, human myoblasts are expanded and committed to differentiate by removing growth factors from the culture medium. The expression levels of specific transcription factors and in vitro immunofluorescence are used to assess the myogenic differentiation process in control conditions and after silencing proteins involved in Ca2+ signaling. Finally, we detail the use of Fura-2 as a ratiometric Ca2+ probe that provides reliable and reproducible measurements of SOCE.

Here, we describe a methodology to obtain a pure population of human myoblasts from adult muscle tissue. These cells are used to study in vitro skeletal muscle differentiation and, in particular, to study proteins involved in Ca2+ signaling.

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for ...

Developmental Biology

Homemade Site Directed Mutagenesis of Whole Plasmids

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. With this method the cloned target gene can be altered by substitution, deletion or insertion of a few bases directly into a plasmid. It works by simply amplifying the whole plasmid, in a non PCR-based thermocycling reaction. During the reaction mutagenic primers, carrying the desired mutation, are integrated into the newly synthesized plasmid. In this video tutorial we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid. The protocol works with standard reagents and is independent from commercial kits, which often are very expensive. Applying this protocol can reduce the total cost of a reaction to an eighth of what it costs using some of the commercial kits. In this video we also comment on critical steps during the process and give detailed instructions on how to design the mutagenic primers.

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. Here we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid using standard reagents.

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. With this method the cloned ...

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

Adult and Embryonic Skeletal Muscle Microexplant Culture and Isolation of Skeletal Muscle Stem Cells

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a robust and reliable method for isolating relatively large numbers of proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. The authors have used micro-dissected explant cultures to analyse the growth characteristics of skeletal muscle cells in wild-type and dystrophic muscles. Each of the components of tissue growth, namely cell survival, proliferation, senescence and differentiation can be analysed separately using the methods described here. The net effect of all components of growth can be established by means of measuring explant outgrowth rates. The micro-explant method can be used to establish primary cultures from a wide range of different muscle types and ages and, as described here, has been adapted by the authors to enable the isolation of embryonic skeletal muscle precursors.
Uniquely, micro-explant cultures have been used to derive clonal (single cell origin) skeletal muscle stem cell (SMSc) lines which can be expanded and used for in vivo transplantation. In vivo transplanted SMSc behave as functional, tissue-specific, satellite cells which contribute to skeletal muscle fibre regeneration but which are also retained (in the satellite cell niche) as a small pool of undifferentiated stem cells which can be re-isolated into culture using the micro-explant method.

The micro-dissected explants technique is a robust and reliable method for isolating proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. Uniquely, these cells have been clonally derived to produce skeletal muscle stem cell lines used for in vivo transplantation.

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a ...

Mitochondrial Isolation from Skeletal Muscle

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and regulate a number of signaling cascades2,3. Past and current researchers have isolated mitochondria from rat and mice tissues such as liver, brain and heart4,5. In recent years, many researchers have focused on studying mitochondrial function from skeletal muscles.
Here, we describe a method that we have used successfully for the isolation of mitochondria from skeletal muscles 6. Our procedure requires that all buffers and reagents are made fresh and need about 250-500 mg of skeletal muscle. We studied mitochondria isolated from rat and mouse gastrocnemius and diaphragm, and rat extraocular muscles. Mitochondrial protein concentration is measured with the Bradford assay. It is important that mitochondrial samples be kept ice-cold during preparation and that functional studies be performed within a relatively short time (~1 hr). Mitochondrial respiration is measured using polarography with a Clark-type electrode (Oxygraph system) at 37&deg;C7. Calibration of the oxygen electrode is a key step in this protocol and it must be performed daily. Isolated mitochondria (150 &mu;g) are added to 0.5 ml of experimental buffer (EB). State 2 respiration starts with addition of glutamate (5mM) and malate (2.5 mM). Then, adenosine diphosphate (ADP) (150 &mu;M) is added to start state 3. Oligomycin (1 &mu;M), an ATPase synthase blocker, is used to estimate state 4. Lastly, carbonyl cyanide p-[trifluoromethoxy]-phenyl-hydrazone (FCCP, 0.2 &mu;M) is added to measurestate 5, or uncoupled respiration 6. The respiratory control ratio (RCR), the ratio of state 3 to state 4, is calculated after each experiment. An RCR &ge;4 is considered as evidence of a viable mitochondria preparation.
In summary, we present a method for the isolation of viable mitochondria from skeletal muscles that can be used in biochemical (e.g., enzyme activity, immunodetection, proteomics) and functional studies (mitochondrial respiration).

This protocol describes a procedure to study the respiration of mitochondria isolated from skeletal muscles. This method was adapted from Scorrano et al. (2007). The mitochondrial isolation procedure requires about 2 hours. The mitochondrial respiration can be completed in about 1 hour.

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

Neuroscience

DNA Electroporation, Isolation and Imaging of Myofibers

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently labeled proteins in the flexor digitorum brevis (FDB) muscle. Conditions have been optimized to provide a large number of high quality myofibers expressing the electroporated plasmid while minimizing muscle damage. The method employs fluorescent tags on various proteins. Combining this expression method with high resolution confocal microscopy permits live cell imaging, including imaging after laser-induced damage. Fluorescent dyes combined with imaging of fluorescently-tagged proteins provides information regarding the basic structure of muscle and its response to stimuli.

This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure.

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently ...

Enzymatic Isolation and Characterization of Variable Length Mouse Muscle Fibers

Intact Short, Intermediate, and Long Skeletal Muscle Fibers Obtained by Enzymatic Dissociation of Six Hindlimb Muscles of Mice: Beyond Flexor Digitorum Brevis

Skeletal muscle fibers obtained by enzymatic dissociation of mouse muscles are a useful model for physiological experiments. However, most papers deal with the short fibers of the flexor digitorum brevis (FDB), which restrains the scope of results dealing with fiber types, limits the amount of biological material available, and impedes a clear connection between cellular physiological phenomena and previous biochemical and dynamical knowledge obtained in other muscles.
This paper describes how to obtain intact fibers from six muscles with different fiber type profiles and lengths. Using C57BL/6 adult mice, we show the muscle dissection and fiber isolation protocol and demonstrate the suitability of the fibers for Ca2+ transient studies and their morphometric characterization. The fiber type composition of the muscles is also presented. When dissociated, all muscles rendered intact, living fibers that contract briskly for more than 24 h. FDB gave short (&#60;1 mm), peroneus digiti quarti (PDQA) and peroneus longus (PL) gave intermediate (1-3 mm), while extensor digitorum longus (EDL), extensor hallucis longus (EHL), and soleus muscles released long (3-6 mm) fibers.
When recorded with the fast dye Mag-Fluo-4, Ca2+ transients of PDQA, PL, and EHL fibers showed the fast, narrow kinetics reminiscent of the morphology type II (MT-II), known to correspond to type IIX and IIB fibers. This is consistent with the fact that these muscles have over 90% of type II fibers compared with FDB (~80%) and soleus (~65%). Moving beyond FDB, we demonstrate for the first time the dissociation of several muscles, which render fibers spanning a range of lengths between 1 and 6 mm. These fibers are viable and give fast Ca2+ transients, indicating that the MT-II can be generalized to IIX and IIB fast fibers, regardless of their muscle source. These results increase the availability of models for mature skeletal muscle studies.

Author Spotlight: Isolation of Long Muscle Fibers from Mouse Hindlimb Muscles for Studying Excitation-Contraction Coupling Across Fiber Types

We describe a protocol to obtain enzymatically dissociated fibers of different lengths and types from six muscles of adult mice: three of them already described (flexor digitorum brevis, extensor digitorum longus, soleus) and three of them successfully dissociated for the first time (extensor hallucis longus, peroneus longus, peroneus digiti quarti).

Skeletal muscle fibers obtained by enzymatic dissociation of mouse muscles are a useful model for physiological experiments. However, most papers deal ...

Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability

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