This work was supported by NIH-R01DK106964 and P20DK123971 grants to Georgi V. Petkov. The authors thank Dr. Viktor Yarotskyy and Ms. Sarah Maxwell for critical evaluation of the manuscript. We are also grateful to urology staff surgeons at MUSC and UTHSC: Drs. Thomas Keane, Harry Clarke, Stephen Savage, Ross Rames, Sandip Prasad, Jonathan Picard, Christopher Ledbetter, and Anthony Patterson as well as the MUSC and UTHSC Urology residents: Drs. Taylor Vaughan, Samuel Walker Nickles, Matthew Young, Erin Burns, Justin Ellett, Ryan Levey, Austin Younger, Mark Currin, Nima Baradaran, Olugbemisola McCoy, Tracy Tipton, Bryce Wyatt, Alyssa Greiman, Sarah Starosta, Aaron Bloch, Christine Callaway, Lucille Cox, Christian Dewan, Erin Heitman, Bradley Houston, Stephen Legg, Robert S. Libby, Cole Locklear, Kristen Marley, Monica O&#39;Hanlon, Patrick Probst, Cynthia Sharadin, Elizabeth Tourville, Daniel Zapata for their help with human tissue collection.

All methods described here have been approved by Institutional Review Board Committees of the University of Tennessee Health Science Center (Memphis, TN, IRB# 17-05714-XP), and the Medical University of South Carolina (Charleston, SC, IRB# 00045232). The approved procedures allow whole thickness urinary bladder specimens (&#62;1 cm by &#62;1 cm) - containing all layers including mucosa, detrusor smooth muscle, and serosa as well are attached blood vessels and adipose tissue) - to be collected from patients-donors undergoing surgical partial extraction of bladder. Patient-donors are adult (age range studied until now: 25 to 87 years old), either male or female, with or without symptoms of overactive bladder (as classified by American Urological Association I-PSS score41). Surgical procedures involve a variety of medical conditions including radical cystectomy for urothelial carcinoma, and adenocarcinoma. In such cases, the collected urinary bladder specimen is remote from the site of tumor.
1. Dissection of DSM tissues and preparation of mucosa-free DSM pieces
<ol>
	<li>Examine the whole thickness urinary bladder specimen that arrived in the lab from the operating room in a tightly sealed container filled with the cold dissection/digestion solution (Figure 1 and Table 2 for composition of DS). 
	NOTE: The specimen is usually kept in cold DS from a few hours to overnight prior to arrival in the laboratory. For longer storage, DS (Table 2) is supplemented with 1 mM CaCl2.</li>
	<li>Remove and rinse the human whole thickness DSM specimen (containing all layers including mucosa, DSM, and serosa) with ice-cold DS to wash out attached debris and blood.</li>
	<li>Pin the urinary bladder specimen, mucosa facing upward and serosa down, onto a silicone enantiomer-coated (Table of Materials) 150 mm diameter round dish filled with ice-cold DS (Figure 1B).</li>
	<li>Remove the adjacent adipose tissue, blood vessels, epithelium (urothelium) and muscularis mucosa from the specimen by sharp dissection using microscissors and forceps.</li>
	<li>Cut out several mucosa-free DSM pieces (~2-3 mm long and 4-6 mm wide) (Figure 1C).</li>
</ol>
2. Enzymatic dissociation of DSM pieces yielding freshly isolated single DSM cells
<ol>
	<li>Place 3 to 6 DSM pieces into a tube containing 1 to 2 mL of pre-warmed (~37 &#176;C) DS containing papain and dithiothreitol (DS-P, Table 2) and incubate DSM pieces in DS-P for 30-45 min at ~37 &#176;C gently shaking the tube occasionally (once every 10-15 min). 
	NOTE: To optimally control the temperature for enzymatic treatment, tubes with tissue pieces and enzyme solutions are placed in either a glass tissue chamber filled with water connected to a circulating heated water bath (Figure 1D) or a high-precision temperature-controlled shaking water bath (Figure 1E).</li>
	<li>Remove DS-P from the tube, briefly wash DSM pieces with ice cold DS, discard cold DS from the tube leaving DSM pieces sitting at the bottom of the tube.</li>
	<li>Add 1 to 2 mL of DS-containing collagenase type II (DS-C, Table 2) to the tube with DSM pieces, gently mix; and incubate for 25-40 min at ~37 &#176;C gently shaking the tube occasionally (every 10-15 min).</li>
	<li>Discard DS-C and wash enzyme-treated DSM pieces 5-10 times with ice-cold DS.</li>
	<li>Following the last wash, leave DS solution inside the tube; gently triturate with a fire-polished Pasteur pipette several times to release single DSM cells.</li>
	<li>Place a few drops of DS solution containing dispersed DSM cells onto a glass-bottom chamber or a coverslip and visually inspect for the quality under a microscope (using a 20x or 40x objective) after at least 5 min following the application to allow the cells to adhere to the bottom.</li>
	<li>Immediately use freshly isolated DSM cells for electrophysiological experiments or store the cells in a tube containing DS at ~4 &#176;C either on ice or in a refrigerator until use (typically for up to 8 h of preparation). 
	NOTE: Within the same preparation the quality of cells varies from highly viable to over-digested, dead DSM cells (Figure 2). When the sequential papain-collagenase method yields a very high number of unviable cells, the preparation is discarded, and a new digestion of DSM pieces is carried out but with reduced incubation intervals. If the procedure results in too few DSM cells then for the subsequent digestion of DSM pieces, the incubation intervals are increased. Positive immunoreactivity to &#945;-smooth muscle actin confirms the identity of DSM cells (Figure 3).</li>
</ol>
3. Recording voltage-step induced cation currents from DSM cells using amphotericin-B perforated whole-cell voltage patch-clamp technique
<ol>
	<li>Pipette 0.25-1 mL of cell suspension onto a glass bottom chamber sitting on a stage of an inverted microscope and allow the cells to adhere to the glass bottom.</li>
	<li>After incubation for at least 45 min, remove DS from the bath and replace with the E solution (Table 2) by superfusion where the solution flow aided by gravity via inlet tubing slowing replaces DS with the new solution while outlet tubing connected to a vacuum waste vessel removes the chamber solution and prevents overflow. Note that E solution contains tetraethylammonium (TEA+) and cesium (Cs+) ions to inhibit K+ currents.</li>
	<li>Prepare a working stock solution of amphotericin-B in dimethyl sulfoxide (DMSO) (1 mg per 10 &#181;L of DMSO). To fully dissolve amphotericin powder, sonicate (at least 15 min) and vortex the solution well. 
	NOTE: This step usually takes less than 10 min. Dissolving 3-4 mg of amphotericin-B in 30-40 &#181;L of DSMO in a 1.5 mL microcentrifuge tube works well. Higher quantities of amphotericin-B require more DSMO solvent typically resulting in a longer interval for mixing and incomplete solubilization of the amphotericin-B solid particles present in the tube.</li>
	<li>Dissolve stock solution of amphotericin-B in the pipette solution (solution P, Table 2) to obtain a final concentration of 200-500 &#181;g/mL. This step requires extensive sonication and vortexing at a high-speed setting (8-10/10) for ~30 to 60 min per step to ensure optimal mixing and prevention of amphotericin-B precipitate formation in the pipette solution. 
	NOTE: Amphotericin-B will precipitate over-time and is light-sensitive. The working pipette solution containing Amphotericin-B is checked for solubility, hand-mixed prior to pipette filling, and kept in the dark.</li>
	<li>Pull multiple patch electrodes, fire-polish electrode tips, and (if needed) coat the tips with dental wax.</li>
	<li>Fill the tip of a patch electrode with the pipette solution (solution P, Table 2) without amphotericin-B by briefly dipping the electrode in the solution.</li>
	<li>Backfill the electrode with the same pipette solution containing amphotericin-B.</li>
	<li>Mount the electrode onto a holder connected to a patch-clamp amplifier headstage.</li>
	<li>Using a micromanipulator, place the electrode just below the surface of the extracellular solution so that the tip of the electrode is just submerged.</li>
	<li>In the voltage-clamp mode, set the holding potential to 0 mV and adjust the current to 0 pA with the pipette offset dial on the commercial amplifier (Table of Materials).</li>
	<li>Determine the electrode resistance using the Membrane Test window/function of the commercial acquisition software (Table of Materials). To activate click Tools&#62;Membrane Test&#62;Play or a shortcut icon in the software. The determined electrode resistance should be in the range of 2 to 5 M&#937;. 
	NOTE: Membrane Test function provided in the commercial acquisition software or Seal Test option on the amplifier can be used to monitor electrode resistance by applying voltage steps repetitively.</li>
	<li>Continue monitoring electrode resistance while advancing the electrode toward a chosen DSM cell with a micromanipulator (Figure 4A). 
	NOTE: To be considered a viable DSM cell, the cell must show spindle-shaped elongated morphology, a well-defined halo around the cell, crisp edges, and semi-contractile (serpentine) appearance.</li>
	<li>When touching the cell surface with the electrode - indicated by a rapid increase in the electrode resistance measured with the Membrane Test function - form a giga-seal by applying gentle rapid negative pressure to the electrode holder via tubing. This results in negative pressure created at the tip of the electrode that pulls cell membrane into electrode aiding in the formation of a giga-seal or a very tight contact between the electrode and plasma membrane (Figure 4B).</li>
	<li>Once the giga-seal forms, compensate pipette capacitance by adjusting fast and slow dials on the commercial amplifier and monitor giga-seal stability (leak current) using the Membrane Test function.</li>
	<li>Allow time, typically 30-60 min, for amphotericin-B to diffuse down the pipette and be inserted into the plasma membrane forming pores primarily selective to monovalent cations. During this step, continue monitoring the giga-seal with the Membrane Test function. As cell perforation increases so does the amplitude of the capacitance transients (compare Figure 4B versus Figure 4C displaying no and effective cell perforation, respectively) measured with the Membrane Test function.</li>
	<li>When the patch perforation is optimal (judged by stable series resistance typically below 50 M&#8486;), cancel out the capacitance transients by adjusting the dials for cell capacitance and series resistance on the amplifier. Series resistance compensation can also be performed at this time (Figure 4D).</li>
	<li>Once stable voltage-step induced cation currents evoked by the specified protocol are observed, apply a compound or physiological condition to test by superfusion and record the responses for the control-, test-condition, and washout (if possible) with the commercial acquisition software.
	<ol>
		<li>Record currents with a routine voltage-step protocol that involves holding DSM cells at -64 or -74 mV and stepping the voltage in 10 mV increments for 400 or 500 ms from -94 to +96 or +106 mV and returning to the holding potential. 
		NOTE: The membrane potential values are adjusted for a liquid junction potential of 14 mV (using P and E solutions, Table 2). The liquid junction potential is obtained in the commercial acquisition software (Table of Materials) by clicking (Tools&#62;Junction Potentials) and entering the concentrations of solution ion components. A ramp protocol can also be used to obtain current recordings.</li>
		<li>Run the voltage-protocol in continuous ~1 min interval during an experiment recording currents for pre-addition control, test condition, and washout.</li>
	</ol>
	</li>
</ol>
4. Data analysis and visualization
<ol>
	<li>Open recorded files in the commercial data analysis software (Table of Materials) for the control, test condition, and washout by clicking File&#62;Open Data and selecting files of interest for opening. 
	NOTE: For analysis typically three files (each containing a single set of traces to a single protocol run) for each condition are opened and analyzed. The responses are subsequently averaged to obtain a mean response for each condition. The software used for data acquisition contains an option to automatically collect user-specified multiple test runs and average them for single output file that can be used as an alternative.</li>
	<li>Obtain the average response over the last 200 ms for the current trace measured at each voltage; the duration interval chosen reflects a steady-state level of voltage-step current activation. To do so follow the steps below.
	<ol>
		<li>Select a file of interest for analysis in the commercial analysis software (Table of Materials). 
		NOTE: The software positions the most recently imported file into its active display window. The open file displays a series of overlapping traces obtained with a voltage-step protocol. By default, within the active window four cursors (displaying x and y values for the highlighted trace) are shown.</li>
		<li>Choose the range for analysis by positioning cursor 2 at the end of the 400 or 500 ms voltage step and cursor 1 at the interval of 200 ms before that so that the analysis range is 200 ms.</li>
		<li>Obtain responses for each voltage by clicking Analyze&#62;Quick Graph&#62;IV (or the IV shortcut icon) (in the prompt window before generating data confirm that for y-axis (Current) signal Region options &#34;Cursors 1..2&#34; and &#34;Mean&#34; are selected). Click OK to generate the I-V graph and to place the data into a Results column sheet that can be viewed by accessing Windows&#62;Results.</li>
		<li>Analyze additional files by repeating steps 4.2.1 - 4.2.3. Click Analyze&#62;Quick Graph&#62;IV or the IV icon shortcut, select Append rather than Replace to add additional data to the Results sheet when processing the traces.</li>
		<li>Copy data to a spreadsheet by selecting the columns of interest and pressing CTRL+C to copy and CTRL+V to paste. Store the Results worksheet in the commercial analysis software (Table of Materials) in (*.rlt) format by clicking File&#62;Save As.</li>
	</ol>
	</li>
	<li>For each cell, normalize the responses for all three conditions to the value of the maximum voltage step for the pre-addition control (according to the formula: Response/ResponseControl-Max) and graph the responses as current (or current density) values versus voltage relationships (Figure 6).
	<ol>
		<li>In a worksheet processor, determine the average responses as either currents (pA) or current densities (pA/pF) for control, test condition (in this example, 9-phenanthrol), and washout at each voltage step.</li>
		<li>Divide values for each voltage of each condition (control, 9-phenanthrol, and washout) by the maximum control response obtained at the highest voltage (+96 mV in Figure 6) for the pre-addition control following the formula: Normalized response for condition(x) = Response(x)/ ResponseControl-Max for (x).</li>
	</ol>
	</li>
	<li>For summary analysis, arrange the data in a format that can be easily copied to a graphical program (e.g. GraphPad Prism) for visualization.</li>
</ol>

The procedures described here explain the steps involved in the preparation of viable, freshly isolated DSM cells from whole thickness human urinary bladder specimens using enzymatic digestion and in the recording of whole-cell cation currents sensitive to the TRPM4 channel inhibitor 9-phenanthrol employing the amphotericin-B perforated patch-clamp approach. The enzymatic procedure relies on a two-step sequential exposure referred to herein as the sequential papain-collagenase digestion method. DSM tissues are first treated with papain and dithiothreitol (an enzyme stabilizing agent) under a nominal Ca2+-free condition, followed in the second step by collagenase type II in the presence of low Ca2+. The rationale for carrying out papain digestion under low Ca2+ conditions in smooth muscle cells dates back to the late 1980&#39;s. Freshly isolated carotid artery smooth muscle cells prepared with papain displayed an elongated shape, showed viability (resistance to Trypan Blue uptake) and responded to contractile stimuli (higher Ca2+ and histamine)65. Years later, this method was applied in the preparation of DSM cells (see Table 1). The choice of collagenase type II rather than other types relates to its relatively high proteolytic activity ideally suited for smooth muscle tissues including DSM. Indeed, collagenase treatment alone could yield single DSM cells albeit requiring extensive enzyme exposure (&#8805;60 min)53,54. Since collagenase activity depends on Ca2+ and the enzyme is inactive under the Ca2+-free conditions, optimum enzymatic digestion of DSM pieces requires the presence of Ca2+ 66. In our case, DS-C contains 100-200 &#181;M [Ca2+] (Table 2). After enzymatic treatment, digested DSM pieces are washed gently a number of times with cold DS without enzymes or Ca2+ to remove any enzyme bound to tissues. The ice-cold DS helps to preserve DSM cell integrity and to limit enzymatic activity of any remaining papain or collagenase. In the last step, trituration of enzyme-treated DSM pieces with a fire polished Pasteur pipette releases single DSM cell. DSM cells are either immediately placed onto a recording chamber for patch-clamp studies or other types of experimentation or stored on ice in DS for use later on the same day (typically within 8 hrs of preparation, but the cells remain viable for up to 24 h).
We identified several important considerations for successfully obtaining single DSM cells. The first pertains to the human DSM specimen source quality. To optimally preserve tissue integrity, DSM samples obtained from open bladder surgeries are placed in ice-cold DS as soon as possible and maintained in a cold environment. Specifically, upon surgical extraction from the patient, the bladder specimen is immediately placed on a fully prepared side table in the operating room. Gross examination of the whole specimen (usually obtained during radical or simple cystectomy) and its opening follow. After visual inspection, a piece of whole thickness urinary ladder specimen is removed from a remote area of the specimen grossly uninvolved with tumor and immediately placed into a cup (either 50 or 100 mL) containing cold (~4 &#176;C) Dissection Solution (DS) (Table 2) and then tightly closed with a lid. Due to the planned nature of the harvesting of the tissue, the operating room personnel and auxiliary staff doing the harvesting are alerted at the start of the surgical case in order to have the materials available in the operating room at the time of tissue extraction. These precautions along with the routine, repetitive nature of the processing steps keep the warm ischemia time for the tissue - from extraction to placement in the chilled container with DS solution - to less than 5 min. The container is then placed in a refrigerator or on ice in a cooler to maintain the cold environment and transported (ice-cold) to the laboratory. Once the specimen arrives in the laboratory, dissection and enzymatic dissociation steps commence. It is very difficult to predict whether a given DSM specimen will yield high quality DSM cells following enzymatic dissociation, so we proceed with the enzymatic dissociation steps. In many cases, in parallel with electrophysiological experiments, our group conducts isometric tension recordings on DSM strips prepared from the same DSM specimens. We have found that we can usually obtain high quality DSM cells from preparations that also successfully provide viable strips for isometric contraction studies (our unpublished observation).
The second factor relates to different enzyme lot variabilities. We observed that for both papain and collagenase type II, each time a new lot of enzyme arrives from a supplier, the enzyme activity in DS for tissue digestion can vary. We, hence, routinely optimize the enzyme concentration and incubation intervals for each new lot. To minimize the lot variability contribution, we order larger quantities of the same lot and make a large batch of stock solutions in 2 mL aliquots of enzymes and store them at ~-20 &#176;C until use. Over time, however, frozen stocks (stored up to 2 weeks) can lose their enzymatic activity. The third variable relates to the temperature of enzyme digestion treatments. Enzymatic activities of both papain and collagenase display temperature-dependency. Papain and collagenase type II exhibit activity in the temperature ranges encompassing normal body physiology67,68. Therefore, we aim to maintain the enzyme treatments stable at ~37 &#176;C avoiding higher temperatures to preserve DSM cell integrity. The fourth consideration concerns the variability of the quality of DSM cells present within each preparation ranging from highly viable (exhibiting excellent classical smooth muscle characteristics) to non-healthy, over-digested cells. A prolonged enzyme incubation interval is one of the main reasons for obtaining a high number of damaged cells. Excessive enzyme treatments also impair protein structures of ion channels, receptors, and transporters, adversely affecting their functionality. Interpretation of results obtained from enzymatically obtained, freshly isolated cells should bear this consideration in mind. Optimization of enzyme digestion conditions aims to increase the percentage of highly viable cells. Experimental approaches relying on a higher number of viable cells such as microarray analyses require more robust optimizations than those successfully carried out on fewer cells such as single-cell patch-clamp electrophysiology or Ca2+ imaging. Consideration of the aforementioned factors has guided our research efforts over the last decade in obtaining high quality single DSM cells.
The perforated patch-clamp technique has been a mainstay electrophysiological approach for over a quarter of a century. Several publications provide details of the technical considerations69,70,71,72,73. Cell perforation can be obtained with amphotericin-B, nystatin, gramicidin, or &#946;-escin (see reference32 for overview of each). The main advantage of perforated patch-clamp recordings over other electrophysiological approaches is that the native intracellular environment - including intracellular Ca2+ and signaling molecules (e.g., cAMP, PKA, phosphates, and phosphodiesterases) - is preserved. This technique is, hence, ideally suited for investigating whole-cell ion channel currents and their regulatory mechanisms under near physiological conditions. A key caveat is that intracellular cell composition cannot be precisely controlled unlike other electrophysiological methods such as conventional whole-cell and single-channel excised-patch (inside- and outside-out) recordings. In our experience, three factors routinely contribute to successful experimental outcomes of amphotericin-B-perforated patch-clamp experiments. The first is the quality of the DSM cell chosen to attempt a recording. When DSM cells are highly viable displaying a semi-contractile (serpentine-like), high-contrast shiny appearance with a well-defined halo around the cell surface and attach tightly to the glass bottom of the recording chamber then giga-seal formation and cell perforation occur relatively easy. The second and third factors for success, respectively, relate to quality of source and solubilization of amphotericin-B (in dimethyl sulfoxide/DMSO and intracellular pipette solution). We observed discrepancies among different suppliers in terms of source and lot variabilities. Each day we prepare a fresh solution of amphotericin-B stock solution from powder followed by its dilution in intracellular pipette solution. These steps require extensive sonication and vortexing. With freshly made amphotericin-B-containing pipette solution, successful cell perforation (&#60;50 M&#8486;) following giga-seal formation can be usually obtained within 30 min. The concentration of amphotericin-B in the pipette solution also requires optimization dependent on the amphotericin-B lot and source. Under our experimental conditions, the final amphotericin-B concentrations in the pipette range from 200 to 500 &#181;g/mL. Since amphotericin-B exhibits light sensitivity, its solutions need to be kept in the dark. Using the amphotericin-B perforated patch-clamp technique, our group recorded voltage-step induced K+, Ca2+, and non-selective cation currents from human, guinea pig, mouse, and/or rat DSM cells17,21,22,23,29,30,31,35,60. Here, we describe conditions for recording non-selective cation currents in human DSM cells. 9-Phenanthrol, a blocker of TRPM4 channels, attenuated voltage-step induced currents supporting the role of these channels in the control of DSM excitability. As a note, it usually requires at least 45 min after obtaining a giga-seal and initiation of perforation to record optimum stable voltage-step induced non-selective cation currents. Voltage ramps can also be used as an alternative to voltage-step protocols30,64. Here, a voltage-step protocol from a hyperpolarized holding membrane potential was preferred rather than a ramp protocol since the former approach minimizes the effect of voltage-dependent inactivation and allows for averaging of evoked current over a duration of a voltage&#8211;step where the ramp provides a single data point per voltage. The latter point especially applies to human DSM cells as the currents show variable activity during voltage steps (Figure 5 and Figure 6). The amphotericin-B perforated patch-clamp technique has been essential in identifying the properties of DSM cells and other cell types and will continue to aide in providing novel discoveries in the future. Furthermore, freshly isolated single DSM cells can be successfully used for measuring whole-cell K+, Cl-, and Ca2+ currents with the conventional mode of the patch-clamp technique, membrane potential recording with the current clamp, and single channel recordings as exemplified by our prior reports23,29,35,64.
In addition to single cell patch-clamp methods, freshly isolated DSM cells can be studied with other technical approaches including Ca2+ imaging, RT-PCR/q-RT-PCR, immunocytochemistry, in situ proximity ligation assay, and genomic approaches (e.g., microarray, RNA-seq, CHIP-seq)15,18,30,33,34. As single cell transcriptome determination methods continue to evolve and become highly sensitive, we envision in a future ability to routinely and specifically link electrical or pharmacological properties of individual DSM cells with their transcriptome/proteome profiles. This will be achieved by first recording from a DSM cell and then extracting mRNA or protein followed by transcriptomic/proteomic analyses. Although such methods have been tested already in non-DSM cells, they are at present technically challenging, lack sensitivity to be considered routine, and limited to successful detection of a few selected gene products74. Function-molecular profile expression linking studies when done on DSM cells obtained from urinary bladders derived from control and diseased patient-donors will provide insights into physiological processes essential for driving normal DSM functions, pathogenesis, and in identifying effective novel therapeutic approaches.

Detrusor smooth muscle (DSM) cells constitute the most abundant cell type in the urinary bladder and ultimately control urine storage and voiding through relaxation and contraction, respectively. DSM cells form smooth muscle bundles that intertwine with adjacent connective tissue, nerve processes, interstitial cells, and other cell types1. The current understanding of the role of DSM cells in urinary bladder function has been achieved through a multi-level integrated approach. Each experimental method - whether based on isolated single cells in vitro, tissue strips containing smooth muscle bundles in vitro/ex vivo, or in vivo determinations (such as cytometry and voiding function assessments) - provides important and specific insights into physiological and pharmacological properties of DSM (please see reviews1,2,3,4,5,6 for details). However, interpretation of results obtained from isolated single cells allows conclusions to be specifically attributed to the single cell type itself. This realization has been the driving force for establishing a reliable and reproducible method for obtaining freshly isolated DSM cells from the whole thickness urinary bladder specimens. Unlike many other cell types, smooth muscle cells cannot be reliably cultured due to the loss of their native phenotype including specific changes in their electrophysiological and contractile properties7,8. This fact further reinforces the importance of studies carried out on physiologically active freshly isolated DSM cells.
In the late 1980&#39;s and early 1990&#39;s, Isenberg&#39;s group (Germany) published a series of electrophysiological studies on freshly isolated DSM cells obtained from guinea pig urinary bladders9,10,11,12,13 (Table 1). The method highlighted two important observations that aided in obtaining vital cells and served as an initial guideline for others to follow. They were 1) pre-treating isolated DSM pieces with Ca2+-free solution/medium prior to enzymatic treatment and 2) tissue digestion with a solution containing collagenase. These two critical steps have been incorporated into all the subsequent variants of DSM cell dissociation procedures (Table 1). Currently, our group employs a two-step sequential papain-collagenase dissociation approach. DSM pieces are first treated with an enzyme solution containing papain and then with collagenase type II solubilized in the same solution (DS, dissection/digestion solution). This approach yields single DSM cells from various species including guinea pig, pig, rat, mouse, and importantly human (Table 1).
Single DSM cells provide a source for multiple molecular biology and physiological experiments. So far, protein and mRNA expressions studied with immunocytochemistry, or RT-PCR/qRT-PCR determinations revealed high levels of detection for various ion channels including the large conductance voltage- and Ca2+-activated (BK), small conductance Ca2+-activated K+ type 3 (SK3), voltage-gated K+ (Kv), L-type voltage-gated Ca2+ (Cav), and transient receptor potential melastatin type 4 (TRPM4) channels, as well as a Na/Ca2+ exchanger14,15,16,17,18,19,20,21,22. They are all thought to control DSM excitability, intracellular Ca2+ levels and contractility. Patch-clamp electrophysiological approaches, performed directly on guinea pig, mouse, rat, or human DSM cells, provided direct demonstration of biophysical and pharmacological properties of L-type Cav, Kv (Kv2.x. Kv7), SK, BK, and TRPM4 channels17,19,20,21,22,23,24,25,26,27,28,29,30,31. The approaches included a conventional whole-cell voltage-clamp, a perforated voltage-clamp, and single-channel recordings (cell attached, inside-out and outside-out configurations). Additionally, membrane potential recording of DSM using a current-clamp provided evidence that target-engaging pharmacological agents alter the cell excitability. For example, the TRPM4 inhibitor 9-phenanthrol induced hyperpolarization in DSM cells obtained from humans, guinea pig, and rat urinary bladders19,20,22,31. Among the various electrophysiological methods, the amphotericin-B (and nystatin, gramicidin, and &#946;-escin) perforated patch-clamp recordings provide a key advantage by preserving intrinsic intracellular signaling molecules and pathways. Only low molecular weight cations and to a lesser extent, Cl- - but not proteins or signaling molecules including Ca2+ - are permeable through the plasma membrane pores formed by amphotericin-B or nystatin32. The successful outcome of perforated patch-clamp experiments depends on several general variables unique to this technique. Here, we describe the details of the procedure utilizing amphotericin-B that our group has used successfully over the years15,22,33,34,35,36,37,38,39.
Arguably, non-selective cation channels remain one of the least understood channel types in DSM cells. The first report of a non-selective cation-like channel dates back to 1993. The paper by Wellner and Isenberg11 described a 33 pS stretch-activated single channel displaying the following rank order of ion permeability: K+&#62;Na+&#62;Cs+&#62;&#62;&#62;Ba2+&#62;Ca2+, and inhibition of channel activity by Gd3+, a general inhibitor of non-selective cation channels. Almost a decade later, Thorneloe and Nelson40 described Na+ permeable cation currents in mouse DSM cells, inhibited by Gd3+, using whole-cell recordings. Since molecular identities of non-selective cation channels and their biophysical characterizations remain to be determined, future investigations in this research area are warranted. The protocol described herein for the recording of non-selective cation channel currents - using extracellular and pipette intracellular solutions containing Cs+, TEA+, and nifedipine (Table 2) that physiologically and pharmacologically mitigate Kv and Cav currents - has been and will continue to be useful in electrophysiological investigations of non-selective cation channels. We have utilized this specific protocol to determine the extent of inhibition of whole-cell cation currents by the TRPM4 channel blocker 9-phenanthrol in guinea pig, rat and human DSM cells19,20,22.
Taken together, the method described here for obtaining freshly isolated single DSM cells from human urinary bladder provides viable cells highly suitable for electrophysiological investigations using various configurations of the patch-clamp technique, Ca2+-imaging, immunocytochemistry, in situ proximal litigation assay, and single cell RT-PCR/qRT-PCR as well as advanced molecular biology techniques including microarray analysis, RNA-seq, and CHIP-seq. The use of the amphotericin-B perforated patch-clamp method preserves the native cell environment unlike other configurations. When carried out using the specific conditions outlined here, designed to negate contributions of K+ and Ca2+ currents in DSM cells, voltage-step induced currents display the properties of non-selective cation currents suitable for biophysical and pharmacological characterizations.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>5 ml polystyrene round-bottom tube</td>
			<td>Falcon</td>
			<td>352054</td>
			<td>Tubes for DS containing enzymes used in digestion steps</td>
		</tr>
		<tr>
			<td>9-Phenanthrol</td>
			<td>Sigma-Aldrich</td>
			<td>211281</td>
			<td>TRPM4 channel inhibitor</td>
		</tr>
		<tr>
			<td>Amphotericin-B</td>
			<td>Fisher</td>
			<td>BP928-250</td>
			<td>Used for patch/cell perforation</td>
		</tr>
		<tr>
			<td>Amphotericin-B</td>
			<td>European Pharmacopoeia Reference Standards</td>
			<td>5</td>
			<td>Used for patch/cell perforation</td>
		</tr>
		<tr>
			<td>Amphotericin-B</td>
			<td>Sigma-Aldrich</td>
			<td>A9528-100MG</td>
			<td>Used for patch/cell perforation</td>
		</tr>
		<tr>
			<td>Analog vortex mixer</td>
			<td>VWR</td>
			<td>58816-121</td>
			<td></td>
		</tr>
		<tr>
			<td>Aspartic acid</td>
			<td>Sigma-Aldrich</td>
			<td>A9006</td>
			<td>Intracellular pipette solution</td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A7906</td>
			<td>DS</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma-Aldrich</td>
			<td>C1016</td>
			<td>Extracellular solution and DS</td>
		</tr>
		<tr>
			<td>Capillary Glass</td>
			<td>Sutter</td>
			<td>BF150-110-7.5</td>
			<td>Capillary for preparation of pulled patch electrodes</td>
		</tr>
		<tr>
			<td>Cesium hydroxide hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>C8518</td>
			<td>Intracellular pipette solution</td>
		</tr>
		<tr>
			<td>Clampex ver. 10 software includes data acqusition (Clampex) and analysis (Clampfit) programs</td>
			<td>Axon Instruments/ Molecular Devices</td>
			<td>pCLAMP-10</td>
			<td>Commerical software and part of patch-clamp rig setup</td>
		</tr>
		<tr>
			<td>Collagenase type 2</td>
			<td>Worthington Biochemical Corporation</td>
			<td>LS004177</td>
			<td>DS-C</td>
		</tr>
		<tr>
			<td>CsCl</td>
			<td>Sigma-Aldrich</td>
			<td>203025</td>
			<td>Extracellular and intracellular solutions</td>
		</tr>
		<tr>
			<td>Dental wax</td>
			<td>Miltex Dental Wax Technologies, Inc.</td>
			<td>18058351</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital Thermometer with Probe</td>
			<td>Fisher Scientific</td>
			<td>15-077-32</td>
			<td>Placed in tissue bath to monitor temperature</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Sigma-Aldrich</td>
			<td>D2650</td>
			<td>Solvent</td>
		</tr>
		<tr>
			<td>DL-Dithiothreitol (DDT)</td>
			<td>Sigma-Aldrich</td>
			<td>D9779</td>
			<td>Reducing agents used together with Papain</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Sigma-Aldrich</td>
			<td>E3889</td>
			<td>Ca2+ chelator, used in intracellular pipette solution</td>
		</tr>
		<tr>
			<td>Flaming/Brown micropipette puller</td>
			<td>Sutter</td>
			<td>P-97</td>
			<td>Required to pull electrodes with very fine tips</td>
		</tr>
		<tr>
			<td>Floating foam tube rack/holder</td>
			<td>VWR Scientific</td>
			<td>82017-634</td>
			<td>Used for holding tubes with enzymes for temperature control</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G8270</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamic acid (Na salt)</td>
			<td>Sigma-Aldrich</td>
			<td>G1626</td>
			<td>DS</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H3375</td>
			<td>pH Buffer</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fisher Scientific</td>
			<td>BP366-1</td>
			<td>Extracellular solution</td>
		</tr>
		<tr>
			<td>Low Noise Data Acquisition System</td>
			<td>Axon Instruments/ Molecular Devices</td>
			<td>Digidata 1440A</td>
			<td>Part of patch-clamp rig setup</td>
		</tr>
		<tr>
			<td>Magnetic stirrer</td>
			<td>VWR</td>
			<td>01-442-684</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2 (hexahydrate)</td>
			<td>Sigma-Aldrich</td>
			<td>M2670</td>
			<td>Extracellular and intracellular solutions</td>
		</tr>
		<tr>
			<td>MicroForge</td>
			<td>Narishige</td>
			<td>MF-830</td>
			<td>Used for fire-polishing electrodes</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td>Extracellular and intracellular solutions</td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Sigma-Aldrich</td>
			<td>S8045</td>
			<td></td>
		</tr>
		<tr>
			<td>Nifedipine</td>
			<td>Sigma-Aldrich</td>
			<td>N7634</td>
			<td>L-type voltage-gated Ca2+ channel blocker</td>
		</tr>
		<tr>
			<td>Nikon inverted microscope, TS100 with T1-SM stage with 5x, 10x, 20x, and 40x objectives</td>
			<td>Nikon</td>
			<td>Discontinued</td>
			<td>Part of Patch-clamp rig setup</td>
		</tr>
		<tr>
			<td>Non-metalic syringe needle, MicroFil</td>
			<td>WPI</td>
			<td>MF-34G-5</td>
			<td>Filling of intracellular pipette solution</td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Worthington Biochemical Corporation</td>
			<td>LS003126</td>
			<td>DS-P</td>
		</tr>
		<tr>
			<td>Pasteur pipette</td>
			<td>FisherBrand</td>
			<td>13-678-20A</td>
			<td>Tips are broken off and fire-polished and used for titration of enzymatically treated tissues to release single DSM cells from pieces</td>
		</tr>
		<tr>
			<td>Patch-clamp amplifier</td>
			<td>Axon Instruments/ Molecular Devices</td>
			<td>Axon Axopatch 200B</td>
			<td>Part of patch-clamp rig setup</td>
		</tr>
		<tr>
			<td>PC computer</td>
			<td>DELL</td>
			<td>Custom configuration</td>
			<td>Part of patch-clamp rig setup</td>
		</tr>
		<tr>
			<td>pH Meter</td>
			<td>Aspera Instruments</td>
			<td>PH700</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene tubing</td>
			<td>Intramedic</td>
			<td>427-436</td>
			<td>Tubing for superfusion of extracellular bath connected to glass-bottom recording chamber</td>
		</tr>
		<tr>
			<td>Tetraethylammonium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>T2265</td>
			<td>Ion channel blocker of Kv and BK channels added to the extracellular bath solution</td>
		</tr>
		<tr>
			<td>Thermo Scientific Precision shaking water bath (model 2870)</td>
			<td>Thermo Scientific</td>
			<td>Discontinued</td>
			<td>Water bath for temperature control of enzymatic digestion employed as an alternative to tissue chamber-circulating bath setup</td>
		</tr>
		<tr>
			<td>Tissue bath, 100 mL</td>
			<td>Radnoti</td>
			<td>1583-101</td>
			<td>Connected to a circulating bath and filled with water, tubes with DS and DSM pieces are placed in the setup to control the temperature of digestion steps</td>
		</tr>
		<tr>
			<td>Vinyl tubing</td>
			<td>ColePalmer</td>
			<td>06405-3</td>
			<td>Multiple uses including for connecting tissue bath to circulating water bath</td>
		</tr>
		<tr>
			<td>Water circulator bath, Haake D1 L</td>
			<td>Haake</td>
			<td>Discontinued</td>
			<td>Connected to tissue bath</td>
		</tr>
		<tr>
			<td>Weighting scale</td>
			<td>Mettler Toledo</td>
			<td>XS64</td>
			<td></td>
		</tr>
		<tr>
			<td>ZeissAxiovert 40C inverted microscope with 10x and 40x objectives</td>
			<td>Carl-Zeiss</td>
			<td>Discontinued</td>
			<td>Part of patch-clamp rig setup</td>
		</tr>
	</tbody>
</table>

preparation and utilization of freshly isolated human detrusor smooth muscle cells for characterization of 9-phenanthrol-sensitive cation currents

Detrusor smooth muscle (DSM) cells present within the urinary bladder wall ultimately facilitate urine storage and voiding. Preparation of the viable, fresh, and isolated DSM cells presents an important technical challenge whose achievement provides optimal cells for subsequent functional and molecular studies. The method developed and elaborated herein, successfully used by our group for over a decade, describes dissection of human urinary bladder specimens obtained from open bladder surgeries followed by an enzymatic two-step treatment of DSM pieces and mechanical trituration to obtain freshly isolated DSM cells. The initial step involves dissection to separate the DSM layer (also known as muscularis propria) from mucosa (urothelium, lamina propria, and muscularis mucosa) and the adjacent connective, vascular, and adipose tissues present. The DSM is then cut into pieces (2-3 mm x 4-6 mm) in nominal Ca2+-containing dissection/digestion solution (DS). DSM pieces are next transferred to and sequentially treated separately with DS containing papain and collagenase at ~37 &#176;C for 30-45 min per step. Following washes with DS containing enzyme-free bovine serum and trituration with a fire-polished pipette, the pieces release single DSM cells. Freshly isolated DSM cells are ideally suited for patch-clamp electrophysiological and pharmacological characterizations of ion channels. Specifically, we show that the TRPM4 channel blocker 9-phenanthrol reduces voltage-step evoked cation currents recorded with the amphotericin-B perforated patch-clamp approach. DSM cells can also be studied by other techniques such as single cell RT-PCR, microarray analysis, immunocytochemistry, in situ proximity ligation assay, and Ca2+ imaging. The main advantage of utilizing single DSM cells is that the observations made relate directly to single cell characteristics revealed. Studies of freshly isolated human DSM cells have provided important insights characterizing the properties of various ion channels including cation-permeable in the urinary bladder and will continue as a gold standard in elucidating DSM cellular properties and regulatory mechanisms.

Enzymatic dissociation of DSM pieces provides healthy freshly isolated DSM cells routinely used in functional and molecular studies such as: patch-clamp electrophysiology and immunocytochemistry. Figure 1 summarizes the dissection steps and visualizes the setups employed for temperature control of enzymatic treatment steps. Figure 2 illustrates bright-field images of DSM cells obtained from three human urinary bladder specimens each from a different patient-donor. Healthy single DSM cells are characterized by spindle-shaped morphology, crisp well-defined edges, a well-defined halo around the cell, and semi-contractile (serpentine-like) appearance when viewed under the microscope (see DSM cells demarked by white arrows in Figure 2). They also respond to contraction stimulating agents such as the muscarinic agonist carbachol or high K+ (60 mM) applications. DSM cells show positive immunoreactivity for &#945;-smooth muscle actin confirming their identity (Figure 3).
DSM cells are ideally suited for patch-clamp electrophysiological investigations of ion channel properties. Here, we describe the amphotericin-B perforated patch-clamp recording method using pipette and extracellular solutions (Table 2) to optimally record voltage-step induced cation channels. Under the specific conditions employed, blockade of Kv and L-type Ca2+ currents with Cs+/TEA+ and nifedipine, respectively, ensured the elimination of contribution of these ionic components to the whole-cell voltage-evoked currents. Figure 4 and Figure 5 show, respectively, the experimental steps of the amphotericin-B perforated patch-clamp method and representative whole-cell currents measured either with a voltage-step induced or a ramp protocol in three different human DSM cells, each from a different patient donor. Note that the recordings display a certain degree of variability in terms of current amplitudes and outward rectification. Additional experiments revealed that 9-phenanthrol, a TRPM4 channel inhibitor, effectively and reversibly inhibited human DSM cation currents at negative and positive voltages (Figure 6). The 9-phenanthrol-sensitive current component illustrates a stronger inhibition at positive voltages and outward rectification (Figure 6C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59884/59884fig01.jpg" /> 
Figure 1: Summary of dissection steps resulting in the preparation of detrusor smooth muscle (DSM) pieces and setup used for enzymatic dissociation. Shown are images of: (A) a whole thickness human urinary bladder specimen provided from an open bladder surgery as extraneous surgical material in ice-cold DS, (B) the same preparation after pinning with a partially dissected DSM layer, (C) DSM pieces of variable dimensions cut out from the DSM layer ready for enzymatic digestion (smaller pieces) or other experimental investigations (larger pieces), (D, E) alternative setups used for enzymatic digestion of DSM pieces consisting of either (1) a temperature-controlled circulating water bath connected via tubing to a large glass tissue chamber filled with water, a rubber holder for tubes, plastic tubes containing DSM pieces and enzyme solutions prepared in dissection/digestion solution (DS, either DS-P or DS-C, Table 2) and a temperature probe linked to a display allowing for continuous monitoring (D), or (2) a large water-filled temperature-controlled bath containing a holder and tubes with DSM pieces and enzyme solutions (E). <a href="https://www.jove.com/files/ftp_upload/59884/59884fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59884/59884fig02.jpg" /> 
Figure 2: Representative bright-field images of human freshly isolated DSM cells obtained using the sequential papain-collagenase digestion method. (A-F) Displayed are images of viable, physiologically active DSM cells considered suitable candidates for attempting perforated patch-clamp recordings. (G, H) Images of non-viable or over-digested cells; such cells were avoided for patch-clamp experiments. White and black arrows in panels (A-H) point to DSM cells considered viable and non-viable, respectively, for attempting patch-clamp recordings. Note that the black arrows in panels (A, C, and G) point to cell fragments (circular pieces) or small cells lacking DSM morphology and in (H) the cells appear pale and dilated. Images are from three different urinary bladder specimens (A and B : patient-donor source one, C and D : patient-donor source two, and E-H : patient-donor source three). <a href="https://www.jove.com/files/ftp_upload/59884/59884fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59884/59884fig03.jpg" /> 
Figure 3: Expressions of transient receptor potential melastatin type 4 (TRPM4) channel and &#945;-smooth muscle specific actin immunoreactivities in single human DSM cells by immunocytochemistry analysis. (A) Shown are confocal images showing immunocytochemical detection of TRPM4 channel protein expression in a human DSM cell. Red staining (bottom left) indicates TRPM4 channel proteins; blue (DAPI) staining detects cell nuclei (top left); green staining indicates &#945;-smooth muscle actin (&#945;-SMA, top right); the merged image (bottom right) illustrates the overlap of all three images. (B) Confocal images illustrating attenuation of immunocytochemical detection of TRPM4 channel protein expression in the presence of a TRPM4-specific competing peptide (CP) in isolated human DSM cells. Blue (DAPI) staining indicates cell nuclei (top left); green staining is for &#945;-smooth muscle actin (&#945;-SMA, top right); the merged image (bottom right) illustrates the overlap of all three images. The results were verified in four separate experiments using DSM whole tissue or multiple DSM cells isolated from four patients. Images are from Hristov et al. (2016)22 and used with permission. <a href="https://www.jove.com/files/ftp_upload/59884/59884fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59884/59884fig04.jpg" /> 
Figure 4: Schematic illustration of steps involved in giga-seal formation and amphotericin-B perforation of human DSM cells. Illustrated are spatial positions of an amphotericin-B containing pipette and a DSM cell along with associated responses for membrane tests obtained in the commercial acquisition software (Table of Materials) by altering voltage steps (either -10 or -20 mV in this example) determining resistance. Configurations are: (A) prior to the cell approach with an electrode, (B) after giga-seal formation obtained by positioning the amphotericin-B containing pipette (amphotericin-B represented by red dots) onto the cell surface and applying negative pressure, (C) the on-cell configuration shown ~45 min after giga-seal formation, at this time-point amphotericin-B has diffused down the pipette and its molecules have inserted into the plasma membrane at the tip of the electrode forming cation permeable pores, and (D) the same configuration as in (C) but with capacitance transients canceled out using dials for whole-cell capacitance and series resistance on the amplifier. <a href="https://www.jove.com/files/ftp_upload/59884/59884fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59884/59884fig05.jpg" /> 
Figure 5: Whole-cell cation currents recorded with amphotericin-B perforated patch-clamp method in human DSM cells. (A) A diagram of the voltage-step protocol illustrates a holding potential of -74 mV and voltage-steps of 400 ms duration from -94 to +96 mV carried out in 10 mV increments and then returned to -74 mV. (B, C) Representative current traces along with current density-voltage plots from two different human DSM cells, each from a different urinary bladder specimen/patient-donor obtained with the voltage-step protocol described in (A). (D) An example of a current trace obtained with a ramp protocol (graphically represented in the top inset as voltage change from -94 to +116 mV over a 1 s duration at 0.21 mV/ms, holding potential was -94 mV). On the right in panels (B-D), the graphs display the current density-voltage relationship for each DSM cell recorded. <a href="https://www.jove.com/files/ftp_upload/59884/59884fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59884/59884fig06.jpg" /> 
Figure 6: TRPM4 channel blocker 9-phenanthrol-mediated inhibition of voltage-step induced cation currents in human DSM cells. (A) Shown are representative currents measured with the voltage-step protocol described in Fig. 5A for control, 9-phenanthrol, and washout. (B) Summary of normalized responses versus voltage for control, 9-phenanthrol, and washout in seven DSM cells (from seven different patient-donors). (C) Difference current for 9-phenanthrol-sensitive component obtained by subtracting the values in the presence of 9-phenanthrol (30 &#181;M) from those of the control shown in (B). Data in (B) and (C) are displayed as means with error bars for SEM, * depicts significance (p&#60;0.05, paired Student&#39;s test) for the comparison of control vs 9-phenanthrol at each voltage. Panels (A) and (B) have been reproduced from Hristov et al. (2016)22 and used with permission. <a href="https://www.jove.com/files/ftp_upload/59884/59884fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Species</td>
			<td colspan="2">Procedure Details</td>
			<td>References</td>
		</tr>
		<tr>
			<td>Guinea pig</td>
			<td colspan="2">DSM pieces rinsed with Ca2+-free medium (in mM: 100 NaCl, 10 KCl, 1.2 KH2PO4, 5 MgCl2, 20 glucose, 50 taurine, measured pCa=6 (or 1 mM) and then cut into pieces and treated typically 90-120 min (4 periods of 30 min) with enzyme medium (in mM: 130 KOH, 20 taurine, 5 pyruvate, 5 creatine, 10 mM HEPES, adjusted with methansulfonic acid to pH 7.4, 1 mg/ml collagenase, 0.2 mg/ml pronase E, 1 mg/ml fatty acid free albumin, pCa=4.2 (63 mM) or 3.7 (200 mM). Single DSM cells were stored in Kraft-Bruhe (KB)-medium (in mM: 85 KCl, 30 K2PO4, 5 MgSO4, 5 Na2ATP, 5 K-pyruvate, 5 creatine, 20 taurine, 5 beta-OH-butyrate, 1 mg/ml fatty acid free albumin, adjusted with KOH to pH 7.2). 
			Alternative method: DSM pieces rinsed for 10 min in a Ca2+-free medium (in mM): 140 NaCl, 5 KCl, 1.2 MgCl2, 10 glucose, 20 taurine, 5 HEPES, adjusted with NaOH to pH 7.4). Then DSM pieces, incubated in the same Ca2+-free medium complemented with 5 mg% collagenase, 2 mg% pronase and 100 &#956;M CaCl2 for 2x20 min stirring.</td>
			<td>Klockner &#38; Isenberg (1985)13,34 Klockner &#38; Isenberg (1986)35 Schneider et al. (1991)10 Bonev &#38; Isenberg (1992)9 Weidelt &#38; Isenberg (2000)36 Moore et al. (2004)14</td>
		</tr>
		<tr>
			<td>Guinea pig, Landrace pig, and human</td>
			<td colspan="2">DSM pieces pre-incubated for 5 min in Ca2+-free Krebs solution then cut into pieces and enzymatically digested in Ca2+-free Krebs solution containing 0.5-2 mg/ml collagenase type I and 0.1-0.5 mg/ml pronase at 36&#176;C for 20-30 min constantly stirred. In some cases, digested pieces were further agitated by a blunt tipped pipette or by spinning until yielding cells. Isolated cells were stored in modified Krebs solution (described in Klockner &#38; Isenberg13) and were normally used within 3 h. The composition of the Krebs solution was (mM): 140 Na+, 6 K+, 2 Ca2+, 1.2 Mg2+, 152.4 Cl-, 10 glucose, 10 HEPES, pH 7.35-7.4 with Tris. For Ca2+-free solution, Ca2+ and Mg2+ were omitted from the Krebs solution.</td>
			<td>Inoue &#38; Brading (1990)37 Inoue &#38; Brading (1991)38 Nakayama &#38; Brading (1995)39,40</td>
		</tr>
		<tr>
			<td>Human</td>
			<td colspan="2">DSM pieces placed in Ca2+-free HEPES Tyrode&#39;s solution (in mM: 105.4 NaCl, 20.0 or 22.3 NaHCO3, 3.6 KCl, 0.9 MgCl2, 0.4 NaH2PO4, 19.5 or 4.9 HEPES, 5.4 or 5.5 glucose, 4.5 or 5.5 Na-pyruvate) and cut into DSM pieces. DSM pieces soaked in an enzyme solution (Ca2+ free HEPES solution with 0.7 mg/ml collagenase type I, 0.7 mg/ml papain, 1 mg/ml albumin) overnight at 4&#176;C. The strips were then heated at 36.5&#176;C for 15 to 30 min, washed and gently triturated in fresh solution. Isolated cells that were stored in Ca2+ containing HEPES Tyrode&#39;s solution or used immediately for experiments.</td>
			<td>Montgomery &#38; Fry (1992)24 Gallegos and Fry (1994)41 Fry et al. (1994)42 Sui et al. (2001)43 Wu et al. (2002)44</td>
		</tr>
		<tr>
			<td>Guinea pig</td>
			<td colspan="2">DSM cut into pieces in PSS (in mM: 137 NaCl, 5.4 KCl, 2 MgCl2, 2 CaCl2, 0.42 KH2P04, 4.17 NaHCO3, 10 glucose, 10 HEPES, pH 7.4 with NaOH). DSM pieces placed for 10 min in in the following digestion solution (in mM: 80 Na-glutamate, 55 NaCl, 6 KCl, 10 HEPES, 11 glucose, 2 MgCl2, and 0.2 CaCl2) and then transferred to a vial containing the same solution but with 1 mg/ml collagenase 2, 1 mg/ml trypsin inhibitor (sometimes omitted), 1 mg/ml fat-free bovine albumin, for ~ 70 min at 35&#176;C or ~60 min at 37&#176;C. Single DSM cells were obtained by trituration through a Pasteur pipette in the same solution without calcium and enzymes. After trituration, Ca2+ (1 mM) was added and the cells were stored at 4&#176;C. The cells were always used on the same day.</td>
			<td>Bonev &#38; Nelson (1993)53,54 Heppner et al. (1997)26 Petkov et al. (2001)47 Shieh et al. (2001, 2007)48,49</td>
		</tr>
		<tr>
			<td>Guinea pig, mouse, rat, and human</td>
			<td colspan="2">Protocol utilizes a two-step enzymatic dissociation treatment following sharp dissection in Ca2+-free digestion solution (in mM: 80 Na-glutamate, 55 NaCl, 6 KCl, 10 HEPES, 11 glucose, and 2 MgCl2). First, DSM pieces were treated for 25-45 min at 37&#176;C with 1-2 mg/ml papain, 1 mg/ml dithioerythreitol and 1 mg/ml bovine serum albumin in dissociation solution (in mM: 80 monosodium glutamate, 55 NaCl, 6 KCl, 2 MgCl2, 10 HEPES, and 10 glucose, adjusted to pH 7.3 with NaOH) and then DSM pieces transferred to digestion solution containing 1-5 mg/ml collagenase XI (Sigma) or collagenase type 2, 1 mg/ml bovine serum albumin, 0 or 1 mg/ml trypsin inhibitor and 100 &#956;M Ca2+, for 6-30 min. Following incubation, the digested tissue was washed several times in digestion solution without enzymes and Ca2+ and then gently triturated to yield single smooth muscle cells.</td>
			<td>Petkov et al. (2001)50 Thorneloe &#38; Nelson (2003)51 Thorneloe &#38; Nelson (2004)33 Petkov &#38; Nelson (2005)27 Hristov et al. (2008)52 Layne et al. (2010)53 Hristov et al. (2011)15 Xin et al. (2012)54 Parajuli et al. (2012)25 Malysz et al. (2013)29 Parajuli et al. (2013)31 Lee et al. (2013)55 Malysz et al. (2014)23 Smith et al. (2013)19,20 Hristov et al. (2016)22 Lee et al. (2017)56 Yarotskyy et al. (2018)57</td>
		</tr>
	</tbody>
</table>
Table 1: Summary of enzymatic approaches used for isolating single DSM cells from urinary bladders of various species.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Solution Type</td>
			<td>Composition (in mM)</td>
		</tr>
		<tr>
			<td>DS (Dissection/Digestion Solution)</td>
			<td>80 Na-glutamate, 55 NaCl, 6 KCl, 10 HEPES, 2 MgCl2, and 11 glucose, pH adjusted to 7.4 (with 10 M NaOH)</td>
		</tr>
		<tr>
			<td>DS-P (Papain-containing DS)</td>
			<td>DS containing 1-2 mg/ml papain, 1 mg/ml dithiothreitol and 1 mg/ml bovine serum albumin</td>
		</tr>
		<tr>
			<td>DS-C (Collagenase-containing DS)</td>
			<td>DS solution containing 1-2 mg/ml collagenase type II, 1 mg/ml bovine serum albumin, 0 or 1 mg/ml trypsin inhibitor and 100-200 &#956;M Ca2+</td>
		</tr>
		<tr>
			<td>P (Pipette)</td>
			<td>110 CsOH, 110 aspartic acid, 10 NaCl, 1 MgCl2, 10 HEPES, 0.05 EGTA, and 30 CsCl,pH adjusted to 7.2 with CsOH, and supplemented with amphotericin-B (300-500 &#956;g/ml)</td>
		</tr>
		<tr>
			<td>E (Extracellular)</td>
			<td>10 tetraethylammonium chloride (TEA), 6 CsCl, 124 NaCl, 1 MgCl2, 2 CaCl2, 10 HEPES, and 10 glucose, pH adjusted to 7.3-7.4 with NaOH or CsOH, and 0.002-3 (2-3 mM) nifedipine</td>
		</tr>
	</tbody>
</table>
Table 2: Compositions of dissection/digestion solution (DS), and pipette and extracellular solutions used in perforated patch-clamp experiments.

Watch this Scientific Journal Video about Preparation and Utilization of Freshly Isolated Human Detrusor Smooth Muscle Cells for Characterization of 9-Phenanthrol-Sensitive Cation Currents at JoVE.com

Preparation and Utilization of Freshly Isolated Human Detrusor Smooth Muscle Cells for Characterization of 9-Phenanthrol-Sensitive Cation Currents

We describe a method for preparation of the single freshly isolated detrusor smooth muscle cells from human urinary bladder specimens employing a two-step enzymatic procedure. The obtained viable DSM cells can be studied by various single cell techniques including the described amphotericin-B patch-clamp electrophysiology to reveal physiological and pharmacological properties.

Detrusor smooth muscle (DSM) cells present within the urinary bladder wall ultimately facilitate urine storage and voiding. Preparation of the viable, ...

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7.1K Views.  University of Tennessee Health Science Center. We describe a method for preparation of the single freshly isolated detrusor smooth muscle cells from human urinary bladder specimens employing a two-step enzymatic procedure. The obtained viable DSM cells can be studied by various single cell techniques including the described amphotericin-B patch-clamp electrophysiology to reveal physiological and pharmacological properties.

Video: Preparation and Utilization of Freshly Isolated Human Detrusor Smooth Muscle Cells for Characterization of 9-Phenanthrol-Sensitive Cation Currents

Focal Ca2+ Transient Detection in Smooth Muscle

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ from internal stores, and allows multiple cells to be monitored simultaneously to assess, for example, coupling in syncytial tissue. Subcellular Ca2+ transients are common in smooth muscle, yet are difficult to measure accurately because of the problems caused by their stochastic occurrence, over an often wide field of view, in an organ that it prone to contract. To overcome this problem, we've developed a series of imaging protocols and analysis routines to acquire and then analyse, in an automated fashion, the frequency, location and amplitude of such events. While this approach may be applied in other contexts, our own work involves the detection of local purinergic Ca2+ transients for locating transmitter release with submicron resolution. 
ATP is released as a cotransmitter from autonomic nerves, where it binds to P2X1 receptors on the smooth muscle of the detrusor and vas deferens. Ca2+ enters the smooth muscle, resulting in purinergic neuroeffector Ca2+ transients (NCTs). The focal Ca2+ transients allow the optical monitoring of neurotransmitter release in a manner that has many advantages over electrophysiology. Apart from the greatly improved spatial resolution, optical recording has the additional advantage of allowing the recording of transmitter release from many distinguishable sites simultaneously. Furthermore, the optical plane of focus is easier to maintain or correct during long recording series than is the repositioning of an intracellular sharp microelectrode. 
In summary, a method for imaging of Ca2+ fluorescence is outlined which details the preparation of tissue, and the acquisition and analysis of data. We outline the use of several scripts for the analysis of such Ca2+ transients.

Focal Ca2+ Transient Detection in Smooth Muscle

Details methods for high-resolution Ca2+ imaging of smooth muscle within isolated organs, including: preparation of the tissue, image acquisition and data analysis.

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ ...

Isolation of Human Umbilical Arterial Smooth Muscle Cells (HUASMC)

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded and their collection does not imply any added risk to the newborn or mother s health. Moreover no ethical concerns are raised. The UC is composed by one vein and two arteries from which both endothelial cells (ECs) 1 and smooth muscle cells (SMCs) 2, two of the main cellular components of blood vessels, can be isolated. In this project the SMCs were obtained after enzymatic treatment of the UC arteries accordingly the experimental procedure previously described by Jaffe et al 3. After cell isolation they were kept in t-flash with DMEM-F12 supplemented with 5% of fetal bovine serum and were cultured for several passages. Cells maintained their morphological and other phenotypic characteristics in the different generations. The aim of this study was to isolate smooth muscle cells in order to use them as models for future assays with constrictor drugs, isolate and structurally characterize L-type calcium channels, to study cellular and molecular aspects of the vascular function 4 and to use them in tissue engineering.

Isolation of Human Umbilical Arterial Smooth Muscle Cells HUASMC

The umbilical cords are used to isolate smooth muscle cells by different ways. In this work we used the enzymatic treatment to isolated smooth muscle cells.

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded ...

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+ handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+ signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e., mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

We describe a method to directly measure muscle force, muscle power, contractile kinetics and fatigability of isolated skeletal muscles in an in vitro system using field stimulation. Valuable information on Ca2+ handling properties and contractile machinery of the muscle can be obtained using different stimulating protocols.

 
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, ...

Isolation of Pulmonary Artery Smooth Muscle Cells from Neonatal Mice

Pulmonary hypertension is a significant cause of morbidity and mortality in infants. Historically, there has been significant study of the signaling pathways involved in vascular smooth muscle contraction in PASMC from fetal sheep. While sheep make an excellent model of term pulmonary hypertension, they are very expensive and lack the advantage of genetic manipulation found in mice. Conversely, the inability to isolate PASMC from mice was a significant limitation of that system. Here we described the isolation of primary cultures of mouse PASMC from P7, P14, and P21 mice using a variation of the previously described technique of Marshall et al.26 that was previously used to isolate rat PASMC. These murine PASMC represent a novel tool for the study of signaling pathways in the neonatal period. Briefly, a slurry of 0.5% (w/v) agarose + 0.5% iron particles in M199 media is infused into the pulmonary vascular bed via the right ventricle (RV). The iron particles are 0.2 &#956;M in diameter and cannot pass through the pulmonary capillary bed. Thus, the iron lodges in the small pulmonary arteries (PA). The lungs are inflated with agarose, removed and dissociated. The iron-containing vessels are pulled down with a magnet. After collagenase (80 U/ml) treatment and further dissociation, the vessels are put into a tissue culture dish in M199 media containing 20% fetal bovine serum (FBS), and antibiotics (M199 complete media) to allow cell migration onto the culture dish. This initial plate of cells is a 50-50 mixture of fibroblasts and PASMC. Thus, the pull down procedure is repeated multiple times to achieve a more pure PASMC population and remove any residual iron. Smooth muscle cell identity is confirmed by immunostaining for smooth muscle myosin and desmin.

We have developed a novel and reproducible technique to isolate primary cultures of pulmonary artery smooth muscle cells (PASMC) from mice as young as P7, thereby allowing better study of the signaling pathways involved in neonatal smooth muscle cell contraction and relaxation.

Pulmonary hypertension is a significant cause of morbidity and mortality in infants. Historically, there has been significant study of the signaling ...

Measurement of Smooth Muscle Function in the Isolated Tissue Bath-applications to Pharmacology Research

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues. While this technique has been implemented for over 100 years, the versatility, simplicity and reproducibility of this assay helps it to remain an indispensable tool for pharmacologists and physiologists alike. Tissue bath systems are available in a wide array of shapes and sizes, allowing a scientist to evaluate samples as small as murine mesenteric arteries and as large as porcine ileum &#8211; if not larger. Central to the isolated tissue bath assay is the ability to measure concentration-dependent changes to isometric contraction, and how the efficacy and potency of contractile agonists can be manipulated by increasing concentrations of antagonists or inhibitors. Even though the general principles remain relatively similar, recent technological advances allow even more versatility to the tissue bath assay by incorporating computer-based data recording and analysis software. This video will demonstrate the function of the isolated tissue bath to measure the isometric contraction of an isolated smooth muscle (in this case rat thoracic aorta rings), and share the types of knowledge that can be created with this technique. Included are detailed descriptions of aortic tissue dissection and preparation, placement of aortic rings in the tissue bath and proper tissue equilibration prior to experimentation, tests of tissue viability, experimental design and implementation, and data quantitation. Aorta will be connected to isometric force transducers, the data from which will be captured using a commercially available analog-to-digital converter and bridge amplifier specifically designed for use in these experiments. The accompanying software to this system will be used to visualize the experiment and analyze captured data.

This protocol describes the measurement of isometric contraction in an isolated smooth muscle preparation, using an isolated tissue bath system and computer-based data acquisition.

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues.  ...

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on changes in intracellular calcium ([Ca2+]i) and nitric oxide (NO). In order to understand how [Ca2+]i, NO and downstream molecules are handled by a blood vessel in response to vasoconstrictors and vasodilators, we developed a novel technique that applies calcium-labeling (or NO-labeling) dyes with two photon microscopy to measure calcium handling (or NO production) in isolated blood vessels. Described here is a detailed step-by-step procedure that demonstrates how to isolate an aorta from a rat, label calcium or NO within the endothelial or smooth muscle cells, and image calcium transients (or NO production) using a two photon microscope following physiological or pharmacological stimuli. The benefits of using the method are multi-fold: 1) it is possible to simultaneously measure calcium transients in both endothelial cells and smooth muscle cells in response to different stimuli; 2) it allows one to image endothelial cells and smooth muscle cells in their native setting; 3) this method is very sensitive to intracellular calcium or NO changes and generates high resolution images for precise measurements; and 4) described approach can be applied to the measurement of other molecules, such as reactive oxygen species. In summary, application of two photon laser emission microscopy to monitor calcium transients and NO production in the endothelial and smooth muscle cells of an isolated blood vessel has provided high quality quantitative data and promoted our understanding of the mechanisms regulating vascular function.

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Vascular cell functiondepends on activity of intracellular messengers. Described here is an ex vivo two photon imaging method that allows the measurement of intracellular calcium and nitric oxide levels in response to physiological and pharmacological stimuli in individual endothelial and smooth muscle cells of an isolated aorta.

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on ...

Single-channel Analysis and Calcium Imaging in the Podocytes of the Freshly Isolated Glomeruli

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells in the pathogenesis of various renal diseases has been established since podocyte injury leads to proteinuria and foot process effacement. It was previously reported that various endogenous agents may cause a dramatic overload in intracellular Ca2+ concentration in podocytes, presumably leading to albuminuria, and this likely occurs via calcium-conducting ion channels. Therefore, it appeared important to study calcium handling in the podocytes both under normal conditions and in various pathological states. However, available experimental approaches have remained somewhat limited to cultured and transfected cells. Although they represent a good basic model for such studies, they are essentially extracted from the native environment of the glomerulus. Here we describe the methodology of studying podocytes as a part of the freshly isolated whole glomerulus. This preparation retains the functional potential of the podocytes, which are still attached to the capillaries; therefore, podocytes remain in the environment that conserves the major parts of the glomeruli filtration apparatus. The present manuscript elaborates on two experimental approaches that allow 1) real-time detection of calcium concentration changes with the help of ratiometric confocal fluorescence microscopy, and 2) the recording of the single ion channels activity in the podocytes of the freshly isolated glomeruli. These methodologies utilize the advantages of the native environment of the glomerulus that enable researchers to resolve acute changes in the intracellular calcium handling in response to applications of various agents, measure basal concentration of calcium within the cells (for instance, to evaluate disease progression), and assess and manipulate calcium conductance at the level of single ion channels.

Changes in the intracellular calcium levels in the podocytes are one of the most important means to control the filtration function of glomeruli. Here we explain a high-throughput approach that allows detection of real-time calcium handling and single ion channels activity in the podocytes of the freshly isolated glomeruli.

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells ...

Measurement of Calcium Fluctuations Within the Sarcoplasmic Reticulum of Cultured Smooth Muscle Cells Using FRET-based Confocal Imaging

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall health. A significant portion of intracellular Ca2+ content is found within the SR stores in VSMCs. As the only intracellular organelle with a close association to the surrounding extracellular space through plasma membrane-SR junctions, the SR can be considered to constitute the first line of response to any irregularity in Ca2+ transients, or stress experienced by the cell. Among its many functions, one of the most important is its role in the transmission of Ca2+-regulated signals throughout the cell to induce further cell-wide reactions downstream. The more common use of cytoplasmic Ca2+ indicators in this regard is overall insufficient for research into the highly dynamic changes to the intraluminal SR Ca2+ store that have yet to be fully characterized. Here, we provide a detailed protocol for the direct and clear measurement of luminal SR Ca2+. This tool is useful for investigation into the nuanced changes in SR Ca2+ that have significant subsequent effects on the normal function and health of the cell. Fluctuations in SR Ca2+ content specifically can provide us with a significant amount of information pertaining to cellular responses to disease or stress conditions experienced by the cell. In this method, a modified version of a SR-targeted Ca2+ indicator, D1SR, is used to detect Ca2+ fluctuations in response to the introduction of agents to cultured rat aortic smooth muscle cells (SMCs). Following incubation with the D1SR indicator, confocal fluorescence microscopy and fluorescence resonance energy transfer (FRET)-based imaging are used to directly observe changes to intraluminal SR Ca2+ levels under control conditions and with the addition of agonist agents that function to induce intracellular Ca2+ movement.

Currently, most available calcium indicators are used to quantify cytoplasmic calcium transients as indirect measures of calcium released from the sarcoplasmic reticulum in cultured smooth muscle cells. This protocol describes the use of a specific FRET-based indicator that allows direct measurement of calcium signals within the sarcoplasmic reticulum lumen.

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall ...

Isolation of Murine Coronary Vascular Smooth Muscle Cells

While the isolation and culture of vascular smooth muscle cells (VSMCs) from large vessels is well established, we sought to isolate and culture VSMCs from the coronary circulation. Hearts with intact aortic arches were removed and perfused via retrograde Langendorff with digestion solution containing 300 Units/ml of collagenase type II, 0.1 mg/ml soybean trypsin inhibitor and 1 M CaCl2. The perfusates were collected at 15 min intervals for 90 min, pelleted by centrifugation, resuspended in plating media, and plated on tissue culture dishes. VSMCs were characterized by presence of SM22&#945;, &#945;-SMA, and vimentin. One of the main advantages of using this technique is the ability to isolate VSMCs from the coronary circulation of mice. Although the small number of cells obtained can limit some of the applications for which the cells can be utilized, isolated coronary VSMCs can be used in a variety of well-established cell culture techniques and assays. Studies investigating VSMCs from genetically modified mice can provide further information about structure-function and signaling processes associated with vascular pathologies.

The purpose of this protocol is to demonstrate the isolation and culture techniques of murine primary vascular smooth muscle cells (VSMCs) from the coronary circulation. Once VSMCs have been isolated, they can be used for many standard culture techniques.

While the isolation and culture of vascular smooth muscle cells (VSMCs) from large vessels is well established, we sought to isolate and culture VSMCs ...

MitoCeption: Transferring Isolated Human MSC Mitochondria to Glioblastoma Stem Cells

Mitochondria play a central role for cell metabolism, energy production and control of apoptosis. Inadequate mitochondrial function has been found responsible for very diverse diseases, ranging from neurological pathologies to cancer. Interestingly, mitochondria have recently been shown to display the capacity to be transferred between cell types, notably from human mesenchymal stem cells (MSC) to cancer cells in coculture conditions, with metabolic and functional consequences for the mitochondria recipient cells, further enhancing the current interest for the biological properties of these organelles.
Evaluating the effects of the transferred MSC mitochondria in the target cells is of primary importance to understand the biological outcome of such cell-cell interactions. The MitoCeption protocol described here allows the transfer of the mitochondria isolated beforehand from the donor cells to the target cells, using MSC mitochondria and glioblastoma stem cells (GSC) as a model system. This protocol has previously been used to transfer mitochondria, isolated from MSCs, to adherent MDA-MB-231 cancer cells. This mitochondria transfer protocol is adapted here for GSCs that present the specific particularity of growing as neurospheres in vitro. The transfer of the isolated mitochondria can be followed by fluorescence-activated cell sorting (FACS) and confocal imaging using mitochondria vital dyes. The use of mitochondria donor and target cells with distinct haplotypes (SNPs) also allows detection of the transferred mitochondria based on the concentration of their circular mitochondrial DNA (mtDNA) in the target cells. Once the protocol has been validated with these criteria, the cells harboring the transferred mitochondria can be further analyzed to determine the effects of the exogenous mitochondria on biological properties such as cell metabolism, plasticity, proliferation and response to therapy.

Here, a protocol (MitoCeption) is presented to transfer mitochondria, isolated from human mesenchymal stem cells (MSC), to glioblastoma stem cells (GSC), with the goal of studying their biological effects on GSC metabolism and functions. A similar protocol can be adapted to transfer mitochondria between other cell types.