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

Summary

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.

Abstract

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' 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 (Ca_V1.1) 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.

Introduction

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 (Ca_V) and sodium (Na_V) channels^4,5. 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'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 stimulus^9,10. This approach has been extensively used to study the voltage-sensing mechanisms in voltage-gated ion channels^{8,10,11,12,13,14,15,16,17,18,19}. For authoritative reviews, see^{10,20,21,22,23}.

The Ca_V and Na_V channels, critical for the initiation and propagation of electrical signals, are composed of a main α1 subunit, which possesses a central pore and four non-identical voltage sensing domains². In addition to their distinct primary structure, Ca_V and Na_V channels are expressed as multisubunit complexes with auxiliary subunits²⁴. Voltage-dependent potassium channels (K_V) consist of four subunits that look like a single domain of Na_V or Ca_V²⁵. The pore-forming and voltage-sensing α1 subunit of Ca_V and Na_V 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 α-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 α-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 K_V channels, proposed to be the main voltage sensor of the channel, moves when the gating charges move in response to changes in membrane potential^8,10. 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 inactivation^{12,27,28,29,30}. Identifying the contribution of each VSD to the channels' 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 perspective^10,23. 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 reconstituted^30,32.

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 contraction^35,36. For the first time, this approach allowed the motion tracking of individual S4 voltage-sensors from the voltage-gated L-type Ca²⁺ channel (Ca_V1.1, 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 Ca_V1.1 is described. While the Ca_V1.1 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.

Protocol

This protocol was approved by the University of Maryland Institutional Animal Care and Use Committee. The following protocol has been divided into multiple subsections, consisting of (1) molecular construct design and cysteine reacting dye selection, (2) in vivo electroporation, (3) muscle dissection and fiber isolation, (4) acquisition setup description, (5) assessment of enhanced green fluorescent protein (EGFP) positive fiber electrical activity and cysteine staining, and (6) signal acquisition and processing. Additionally, at the beginning of each section, some relevant considerations are detailed when applying FSDF in a skeletal muscle fiber. All protocol sections should be carried out with proper personal protective equipment, including a laboratory coat and gloves.

1. Molecular construct design and cysteine reacting dye selection

1. Construct design is a critical part of the experiment's success. First, generate a wild-type, fluorescently tagged Ca_V1.1 cDNA construct and evaluate its expression in the appropriate cell type. For muscle fibers, a strong transfection efficiency can be achieved by using a plasmid carrying a cytomegalovirus (CMV) promoter. In this protocol, an already characterized rabbit EGFP-Ca_V1.1 plasmid was used³⁸.
   NOTE: When engineering the cDNA construct to introduce a cysteine residue into a voltage-gated ion channel, the cysteine position is critical and should be carefully considered. The cysteine should be accessible from the extracellular space to allow a thiol-conjugated dye reaction (Figure 1C,D) and should be proximal to the S4 region to precisely track its motion in response to depolarization. However, to allow dye fluorescence quenching in response to protein motion, the introduced cysteine conjugated fluorophore should be at the interface of two different environments (e.g., membrane and extracellular fluid; Figure 1E). In addition, it is critical to ensure that the inserted cysteine does not interfere with protein function.
2. To obtain an idea about proper cysteine localization, gather information on channel structure or from other fluorometry experiments of other related channel proteins. For the design of cysteine-engineered Ca_V1.1 constructs, evaluate the resolved cryo-electron microscopy (cryo-EM) structure of the channel²⁶ and compare the cysteine insertion of previous work from related channels such as Ca_V1.2¹², or other channels such as Shaker¹¹, and NaChBac³⁹.
3. Once the proper cysteine position is chosen, use a commercial site-directed mutagenesis kit to introduce cysteine substitutions. In the present protocol, the following cysteines modifications were independently engineered on each cytosolic end of the S4 voltage-sensing segment of Ca_V1.1: VSD-I (L159C), VSD-II (L522C), VSD-III (V893C), VSD-IV (S1231C), and UniProtKB: P07293 (Figure 1B).
4. For skeletal muscle fibers, use 5-carboxytetramethylrhodamine methanethiosulfonate (MTS-5-TAMRA), which exhibits proper and rapid diffusion into the transverse tubule membrane system, which is an invagination of the surface membrane and the predominant location of Ca_V1.1 channels. MTS-5-TAMRA displays a decrease in fluorescence when transitioning from the lipid membrane to an aqueous environment (Figure 1E).
   NOTE: Dylight or Alexa-maleimide derivatives stain the surface membrane but not the transverse tubule system.

Figure 1: Schematic of thiol-cysteine reaction at the interface of a transmembrane α-helix. (A) L-type Ca_V1.1 membrane topology. The plus signs represent basic residues within the S4 α-helix and the orange stars indicate the location where cysteine was introduced via site-directed mutagenesis. (B) Sequence alignment of S4_I to S4_IV of rabbit Ca_V1.1 (UniProtKB: P07293). Positively charged arginine and lysine residues critical for voltage sensing are highlighted in red, while engineered cysteine substitutions are indicated in orange. This panel has been adapted from reference³⁷. (C) Cysteine-thiol fluorescent molecule reaction. (D) Diagram illustrating cysteine mutagenesis insertion within a transmembrane voltage-sensitive α-helix. Cysteine should be buried in the membrane at rest and be accessible extracellularly after depolarization (ΔV; I). Cysteine tracking is typically unlikely to occur if target cysteine is already accessible from the extracellular space before depolarization (II) or if the cysteine is not accessible from the extracellular space after depolarization (III). (E) After the reaction with the thiol fluorescent molecule, α-helix motion in response to depolarization decreases MTS-5-TAMRA fluorescence emission. The fluorometric signal is generated by the movement of the S4 helix and the subsequent dye movement relative to the plane of the membrane and the aqueous environment. Please click here to view a larger version of this figure.

2. In vivo electroporation

NOTE: Electroporation experiments were conducted as previously described³⁸ with modifications. In the following section, the protocol is designed for the electroporation of one foot pad of the mouse. Volumes need to be adjusted if both paws are prepared.

1. Aliquot 25-100 µL of plasmid solution at 2-5 µg/µL in a 1.5 mL tube placed on ice.
2. Prepare a 0.5 mL solution of 2 mg/mL hyaluronidase in sterile saline and filter the solution through a low binding protein 0.2 µm sterile filter mounted to a 1 mL syringe. Store in a 1.5 mL tube at room temperature.
3. Using a calibrated anesthesia apparatus, anesthetize a mouse using 3%-4.5% isoflurane in O₂ (1 L/min) by placing the mouse into the anesthesia chamber. Confirm adequate anesthesia of the animal by pinching the tip of the tail using a pair of tweezers. No reaction should be observed when optimal anesthesia is reached.
4. Remove the mouse from the anesthesia chamber and place an anesthesia nose mask over the mouse. Place the animal on its back on an isothermal heating pad covered with a sterile bench pad. Continue the anesthesia using the rodent mask with 3% isoflurane in O₂ (1 L/min).
5. To prevent eye dryness during the procedure, apply a fine layer of artificial tear cream on the eyes of the animal with a sterile cotton tip. Disinfect the paw of the animal using a sterile wipe saturated in ethyl alcohol.
6. Using a 0.5 in long 29 G sterile insulin needle, aspirate 20 µL of hyaluronidase solution. Penetrate the skin at the level of the heel and slide the needle subcutaneously toward the base of the toes (Figure 2A). Slowly inject the solution while progressively moving the needle backward. A bolus or bump should be observed under the paw (Figure 2B).
   NOTE: Depending on the age of the animal and the size of the paw, it is likely that the full amount of solution may not be injected. Often, a small leak through the point of injection may occur.
7. Repeat step 2.6 with the other paw, if desired, using another sterile needle after proper paw disinfection, as in step 2.5.
8. Disconnect the anesthesia by removing the animal from the nose mask and return the mouse to the cage with access to food and water ad libitum. Full recovery from anesthesia should be observed in ~5 min. Place the tube containing the plasmid solution on the bench to allow it to reach room temperature.
9. After 1 h, anesthetize the animal a second time, place it on the heating pad, and disinfect the paw as described in steps 2.3-2.5.
10. Inject 10-20 µL of the cDNA construct, using the same technique described in step 2.6. The total amount of construct injected is 50-100 µg per paw. Repeat the procedure with the contralateral paw if desired using another sterile syringe.
11. Keep the animal under anesthesia on the isothermal heating pad for 5 min to allow the cDNA solution to disperse evenly through the tissue.
12. Turn on the electroporation apparatus device and connect it to the double electrode array, as recommended by manufacturer.
13. Disinfect the double electrode array using a wipe saturated in ethyl alcohol. Stabilize the paw with one hand and first insert one electrode under the skin at the back of the heel. Then, insert the second electrode at the base of the toes, ensuring that the orientations of both electrodes are perpendicular to the axis of the foot (Figure 2C). Orient the probe in a position that does not constrain the foot or leg in an extreme angular orientation.
    NOTE: Electrode insertion could be facilitated with the use of tweezers and by regularly sharpening the tips of the electrodes. Depending on the age of the animal, the size of the paw can vary and electrode spacing should be adapted accordingly.
14. Electroporate the muscles by applying 20 pulses, 20 ms in duration/each, at 1 Hz. For electrode needles spaced at 1 cm, set the voltage to ~100 V. This should be adapted if electrode spacing is modified to reach ~100 V/cm. Slight flexion of the digits should be observed during pulse delivery if the electrodes are positioned properly.
15. Repeat steps 2.13 and 2.14 with the contralateral paw if desired.
16. Disconnect the anesthesia and place the animal in a cage, isolated from its non-electroporated counter mate with food and water access ad libitum for 2 h. Full recovery from anesthesia should be observed in ~10 min. Place the animal back inside the cage.
    NOTE: Expression of cDNA construct(s) is highly dependent on the protein encoded. Protein turnover, quantity and quality of cDNA, plasmid promoter, and other variables can impact construct expression. In this experiment, optimal expression of the α1S subunit of Ca_V1.1 with a CMV promoter requires 4 to 6 weeks but can be detected from 2 weeks for up to 12-15 months.

Figure 2: Diagram of cDNA injection and electroporation electroporation electrode positioning in a mouse foot pad for electroporation. (A) Needle position for hyaluronidase and cDNA injection under a mouse foot pad. The arrow indicates the point of insertion through the skin. (B) Slight discoloration of the skin and slight increase of the paw size should be transiently observed after injection. (C) Electrode array positioning for electroporation. Please click here to view a larger version of this figure.

3. Muscle dissection and fiber isolation

NOTE: Skeletal muscle fiber dissociation was carried out as previously described^37,40,41 with modifications. In the following section, the protocol is suited for two foot pads of the mouse.

1. Ahead of the fiber dissociation, prepare the sylgard-covered plate by adding one part of curing agent to 10 parts of elastomer (% w/w) in a 60 mm plastic Petri dish to reach a thickness of ~5 mm. Allow the elastomer covered plate to cure overnight before use. This can be reused multiple times if properly stored and disinfected with 70% ethyl alcohol before and after use.
2. Prepare 4 mg of collagenase type I in 2 mL of spinner minimum essential eagle's medium (S-MEM) supplemented with 10% fetal bovine serum (FBS; final concentration of 2 mg/mL). Transfer the solution to a 35 mm non-coated plastic plate and place the dish in an incubator at 37 °C, 5% CO₂.
   NOTE: S-MEM is a modified MEM formulation without glutamine and Ca²⁺. The absence of Ca²⁺ at this step reduces fiber contracture during enzymatic digestion and trituration.
3. Add 5 mL of S-MEM supplemented with 10% FBS in a 60 mm non-coated plastic plate and store in the incubator at 37 °C, 5% CO₂.
4. Euthanize the animals by asphyxiation via CO₂ followed by cervical dislocation. To reduce contamination during primary cell isolation, submerge the animal carcass in 70% ethyl alcohol for ~10 s. Remove and dry the carcass with absorbing paper and cut the feet with a pair of scissors between the ankle and the knee (Figure 3A).
5. Pin down one foot of the animal, with the paw facing up, with dissection pins on the elastomer covered plate under a dissection microscope at 10x magnification.
6. With dissection scissors and fine tweezers, remove the skin from the paw to expose the flexor digitorum brevis (FDB) muscle (Figure 3B). To prevent tissue dryness, add a drop of S-MEM 10% FBS to the muscle using a 1,000 µL pipette.
7. At the level of the heel, incise the tendon and carefully dissect the FDB muscle from heel to toe (Figure 3B). Avoid tension on the muscle as much as possible while performing the dissection. Too much force applied on the tissues will result in muscle fiber damage. Place the muscle immediately in the collagenase solution.
8. Repeat steps 3.5-3.7 with the contralateral foot. Place the dissected muscle in the collagenase solution in a 37 °C, 5% CO₂ incubator for 2 h 45 min to 3 h 15 min. Adapt the incubation time depending on the collagenase enzymatic activity and the age of the animal.
9. While the muscle tissue is incubating, add 300 µL of cold MEM without serum or antibiotics at the center of a 35 mm glass bottom dish. Add 2 µL of cold laminin at 1.20 mg/mL directly to the MEM. Repeat the process for the number of dishes desired. Place the dishes into the cell culture incubator at 37 °C, 5% CO₂ for at least 1 h to allow laminin polymerization.
10. When the enzymatic digestion is complete, transfer the muscle into the 60 mm cell culture dish containing S-MEM 10% FBS with the use of a large bore (5 mm) fire-polished glass Pasteur pipette and a latex bulb.
11. Using a smaller bore (2 mm) fire-polished glass Pasteur pipette and a latex bulb, gently triturate the muscle under a dissection microscope (Figure 3C). Dissociated muscle fibers should start detaching from the tissue and be released in the solution.
    NOTE: As a rule of thumb, less trituration (15-30 pipette passages) is always preferred, as too much prolongated trituration can stress or even damage the fibers.
12. Using a pair of fine tweezers, remove any non-muscle tissue, such as nerves, tendons, or blood vessels (Figure 3D).
13. Add 2 mL of warm MEM 2% FBS to each 35 mm laminin-coated glass bottom dish. Using a 200 µL pipette and a sterile plastic pipette tip, transfer the dissociated muscle fibers to the 35 mm laminin-coated glass bottom dish (Figure 3E).
    NOTE: Achieving a low fiber density and allowing the fibers to be well separated from each other is important to prevent fiber overlapping.
14. Place the 35 mm glass bottom dish in the incubator at 37 °C, 5% CO₂. Fibers can be used in a time frame of 2-20 h.

Figure 3; Muscle FDB fiber dissection and dissociation. (A) After foot dissection above the ankle articulation (dashed line), the skin under the foot paw is removed, following the dashed line to expose the FDB muscle (B). The muscle is dissected and placed in collagenase solution. (C) After incubation, the muscle is triturated to dissociate and obtain individual muscle fibers. (D) Fine tweezers are used to remove non-muscle tissue and debris before transferring muscle fibers to the laminin-coated glass bottom culture dish. Please click here to view a larger version of this figure.

4. Acquisition setup description

NOTE: The acquisition setup is comparable to the one described before⁴² with modifications (Figure 4A).

1. Turn on all the components: computer, AD/DA converter and path-clamp amplifier, microscope, motorized stage, manipulator(s), power supply for the photodiode, light source, light shutter, pulse generator, and protocol editor.
2. Activate the transistor-transistor logic (TTL) triggering OUT signal from the AD/DA to control the pulse generator, the light shutter, and the protocol editor.
3. Use the TTL output signal from the pulse generator and connect to an AD channel of the amplifier to verify precise temporal triggering. Adequate control of the triggering consistency should be evaluated carefully ahead of the experiment to ensure adequate apparatus synchronization.
   NOTE: In the case of Ca_V1.1, expect to see maximal signal in less than 4-10 ms after stimulation (time to develop maximal charge movement⁵). The signal is fast and precise temporal resolution is critical to compare with other measures, such as calcium transient or charge movement measured via voltage clamp.
4. To focus the excitation light in a specific area or spot of the fiber (Figure 4B), use a diaphragm positioned in the excitation light path. This enables signal acquisition only in an area where the EGFP-Ca_V1.1 signal is maximal (Figure 4B).

Figure 4: Description of the recording system. (A) Diagram illustrating the connection between the different components of the recording system. The setup consists of an inverted microscope with a motorized stage, a light emitting diode (LED) light source, a light shutter, a custom-made photodiode-based light monitoring circuit with a track and hold function⁴³, an AD/DA converter (from a patch clamp amplifier), an analog pulse generator, an external field stimulation unit coupled to field stimulation electrodes, motorized manipulators, and commercial software for the acquisition, synchronization, and generation of protocols. The electrode for field stimulation is made of two platinum wires welded to copper cables linked to the pulse generator via a BMC connector. Specific excitation and emission filters are used to detect both EGFP and MTS-5-TAMRA signals. To excite EGFP, a xenon lamp with a 488 nm (± 20 nm) excitation (Ex) filter and an LP510 nm Em filter is used. For MTS-5-TAMRA, a 530 nm LED light source and an LP550 nm Em filter is used. (B) View of a fiber expressing an EGFP-Ca_V1.1-cys construct with the two-field stimulation electrodes (black circles) orientated properly (left) and improperly (right) in the main axis of the fibers (dashed line). The black unfilled circle represents the area of acquisition with a diameter controlled by the diaphragm opening, placed in front of the light source. Please click here to view a larger version of this figure.

5. Assessment of EGFP-positive fiber electrical activity and cysteine staining

NOTE: Skeletal muscle fiber field stimulation is carried out as described before⁴¹ with modifications. This approach is used to (1) identify healthy, functional, and electrically responsive fibers, (2) stain the fibers with the cysteine-reactive fluorescent dye, and (3) record the fluorescent signal in response to the propagated action potential. Every step of this section and the following one should be conducted in a low-light environment to reduce fluorescent dye bleaching.

1. Place the 35 mm glass bottom dish containing dissociated muscle fibers on the stage of the microscope. Carefully remove the culture media with a 1,000 µL pipette and replace with 2 mL of room temperature Ringer's solution (see Table 1 for composition). Multiple rounds of media replacement may be required to fully remove the original cell culture media containing free cysteines.
2. Using a mechanical or motorized manipulator, place the two platinum wires perpendicular to the bottom of the dish. Ensure that the electrode terminals are aligned relative to the fiber's longitudinal axis and a few millimeters away from fiber's ends, and that the separation between the electrodes is 5 mm (Figure 4B). Further adjust the electrode positioning by either rotating the dish or mounting each electrode on an independent micromanipulator (Figure 4B).
3. Turn on the transmitted light and find the fibers in the field of view using a 20x objective. Move the EGFP filter cube into the light pathway.
   NOTE: Using a microscope equipped with epifluorescence and a low-magnification (2x) objective, it is possible to assess the construct transfection efficiency in the whole muscle before fiber dissociation by assessing EGFP expression (Figure 5A).
4. Using a remote-controlled light shutter, activate the 488 nm excitation light to identify the EGFP-positive fibers. Store the fiber x-y location on the dish using a motorized microscope stage. The EGFP signal is often heterogeneous within the fiber (Figure 4B). Center the saved position to the brightest EGFP signal.
5. After identifying EGFP-positive fibers, return to the first saved localization. Using a manual trigger switch, deliver two sequential stimulation pulses with a 1 ms duration and 20 V amplitude. Set the polarity of the pulses to alternating.
6. After the stimulation, observe two concentric homogenous fiber contractions in response to the two pulses of opposite polarity. A local contraction or the absence of contraction in responses to pulses of alternate polarity indicate local non-propagated passive responses or unexcitabilty⁴¹. Exclude these fibers for the rest of the experiment.
7. Add 2 µL of 10 mM MTS-5-TAMRA solution directly into the dish and gently mix with a 1,000 µL pipette (10 µM final concentration). Take care to not move the dish, or the stored fiber positions will be lost. Incubate for 4-5 min to allow diffusion of the fluorescent thiol molecule into the transverse tubule system lumen.
8. Apply bipolar repetitive stimulations to evoke successive action potential trains at a rate of 50 Hz for 300 ms every 1 s for 5 min.
   NOTE: The pulse trains give accessibility to the cysteines inserted into the S4 of EGFP-Ca_V1.1 to react with MTS-5-TAMRA. The fiber's ability to mechanically contract in response to stimulation is important for transverse tubule lumen content to cycle with the extracellular environment.
9. Remove the staining solution from the dish with a 1,000 µL pipette and replace with 2 mL of room temperature Ringer's solution. Two or three rounds may be required to fully remove unconjugated MTS-5-TAMRA. Let the stained fiber recover from the staining protocol for at least 10 min.
10. As in step 5.6, re-assess the fiber health and electrical activity by observing symmetrical fiber contraction in response to alternating polarity. Exclude fibers that do not respond to both stimulations from the rest of the experiment.
11. Move the MTS-5-TAMRA filter cube into the light pathway. Using a remote-controlled light shutter, activate the 533 nm excitation light to confirm homogenous MTS-5-TAMRA staining on the fibers.
    NOTE: Upon staining with MTS-5-TAMRA, both engineered and endogenous cysteines react with the maleimide derivative (Figure 5B). Thus, it is difficult to evaluate the proper reaction with the cysteine of interest. Ca_V1.1 is mainly expressed in the transverse tubule, forming a distinguishable double band pattern. Using a confocal or epifluorescence microscope, an x-y image could be used to confirm proper staining and transverse tubule entry and diffusion of MTS-5-TAMRA (Figure 5C).

6. Signal acquisition and processing

NOTE: Before performing fluorometric measurements, signal acquisition must be carefully designed to obtain the optimal signal/noise ratio. Slower sampling rates allow more light detection while reducing the number of points that would be acquired during protein conformational rearrangement. In the case of EGFP-Ca_V1.1-cys, charge movement induced by an action potential waveform occurs in ~1-10 ms³⁷. To obtain multiple points to track the evolution of the motion over time, the acquisition was set to 50 µs per point.

1. Place the fiber in the middle of the field of view with a proper magnification system. For these experiments, a 60x oil 1.4 numerical aperture (NA) inverted objective was used. Optimize the illumination and the fiber position with the motorized stage and the diaphragm to illuminate a circular area of the diameter of the fiber, where the EGFP signal is maximum (Figure 4B).
2. Once the fiber is positioned for acquisition, orient the two independently mounted field stimulation platinum wires at each end of the fiber. Align the wires on the main axis of the fibers in a straight line and space them 5 mm apart with the fiber in the center (Figure 4B).
3. Set the acquisition excitation and emission filters to the proper settings for MTS-5-TAMRA. Start the experiment by executing the protocol written in the acquisition software. This step triggers all the downstream devices (i.e., protocol editor, light shutter, pulse generator).
   NOTE: This protocol allows for a brief period (i.e., 10 ms) of baseline acquisition before the delivery of the field stimulus to allow subsequent measurements of resting fluorescence.
4. Initiate a single or a train of action potentials with a 0.5 or 1 ms, 20 V square pulse. Minimize the total acquisition time as much as possible to avoid signal bleaching.
   NOTE: Even if recording at the center of the fiber, a movement-related fluorescence signal could occur and may be confounded with the fluorescent signal due to protein-fluorophore conformational change (Figure 5D). The contraction-induced signal should be delayed compared to the expected time of charge movement post-stimulation³⁷.
5. To further distinguish the signal arising from S4 motions from that due to fiber contraction, add 1 µL of 100 mM N-benzyl-p-toluene sulphonamide (BTS; 50 µM final concentration) to the recording solution to minimize contractile responses and repeat step 6.4. Signals detected for a second time after fiber pharmacological immobilization correspond to the molecular motion of the tagged S4 helix (Figure 5D).
   NOTE: No signal should be detected in the control EGFP-Ca_V1.1 without engineered cysteine after movement suppression with BTS.
6. Using the same settings, acquire a similar signal in a location within the dish where no muscle fiber nor debris are present to obtain a value of background fluorescence.
7. Import the files that contain the time course of the raw fluorescence [Fr(t)] from the fiber and background fluorescence [Fb(t)] into the data analysis software. Average the column that contains Fb(t) to obtain a homogeneous Fb value. Subtract Fb from the Fr(t) to obtain the absolute fluorescence [F(t)] values. Smooth the resulting signal with a smoothing function if needed.
   NOTE: With this detection system and the frequency of acquisition, we decided to use an adjacent-averaging function with a window of 50 points for smoothing.
8. Average the baseline F(t) values in a time interval of 10 ms before the stimulation to obtain a resting fluorescence value (F0). Subtract F0 from the smoothed F(t) to obtain the absolute change in fluorescence [ΔF(t)]. Then, to express fluorescence change over time relative to resting fluorescence (ΔF/F0), divide ΔF(t) by F0.
9. To evaluate the extent of signal bleaching, define two points from the ΔF/F0 over time signal, before and after the stimulation and away from the fluorometric signal. Fit a linear function to these two points to obtain a baseline trace. Subtract the baseline to ΔF/F0 over time to correct for signal bleaching.
10. Subtract from the time column the delay between acquisition initiation and the feedback signal from the stimulation, to have t = 0 corresponding to the start of the electrical stimulus.
    NOTE: To allow multiple signal comparisons, it is often required to normalize the amplitude of the signal. Different approaches can be used depending on the purpose of the experiment. In the following results section, we were interested in signal kinetics, so we used a simple method consisting of normalizing each signal by the minimal reached value (i.e., the negative going peak).

Figure 5: Imaging muscle fiber expressing EGFP-Ca_V1.1-cys without and with MTS-5-TAMRA staining and representative raw fluorometric record. (A) Examples of transmitted (left) and fluorescent (right) images of the dissected, not dissociated muscle expressing an EGFP-Ca_V1.1 VSD-III construct. Scale bar: 100 µm. (B) Representative image of a muscle fiber expressing an EGFP-Ca_V1.1 VSD-III construct before (left) and after (right) MTS-5-TAMRA staining. Endogenous cysteines of non-transfected fibers are also stained by the dye. Scale bar: 30 µm. (C) Confocal image of an EGFP-Ca_V1.1 VSD-III construct (left) and MTS-5-TAMRA staining (right) show a classic double band pattern characteristic of Ca_V1.1 localization on the transverse tubule system of the muscle fiber (bottom). Scale bar: 25 µm. (D) Representative fluorometric recording in response to two stimuli and measured with a photodiode before (blue trace) and after (red trace) fiber immobilization with N-benzyl-p-toluene sulphonamide (BTS). The top black line indicates the protocol for fiber depolarization via external field stimulation. Please click here to view a larger version of this figure.

Results

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...

Discussion

Here, a step-by-step protocol to conduct FSDF in muscle fibers for the study of individual voltage sensor motions from the Ca_V1.1 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 ...

Materials

List of materials used in this article

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