The authors thank Mrs. Lyndsay Busby for administrative assistance and Mr. Robert Highet from Mayo Clinic Division of Engineering for design and 3D printing of the culture dish insert. This work was supported by NIH K08 to AB (DK106456), Pilot and Feasibility Grant to AB from Mayo Clinic Center for Cell Signaling in Gastroenterology (NIH P30DK084567) and 2015 American Gastroenterological Association Research Scholar Award (AGA RSA) to AB and NIH R01 to GF (DK52766).

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of Mayo Clinic. The primary cell culture portion of this protocol is based on previously published methods that can be referenced for further details10. All experimental procedures must be approved by the (IACUC), and all experiments must be performed in accordance with relevant guidelines and regulations.
1. Culture Preparation
<ol>
	<li>Media.
	<ol>
		<li>Prepare EC cell complete culture media with high glucose [4500mg/L] Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 5% Fetal Bovine Serum (FBS), 1% L-Glutamine and 1% Penicillin-Streptomycin (Table 1). 
		NOTE: Media can be stored at 4&#160;&#176;C for up to one month.</li>
		<li>Filter the EC cell complete culture media and place 25 mL of media into a 50 mL tube at 37 &#176;C.</li>
		<li>Prepare digestion media with high glucose [4,500 mg/L] DMEM supplemented with 0.1% Bovine Serum Albumin (BSA) (Table 1).</li>
		<li>Filter the digestion media and place four 10 mL aliquots of media into separate 50 mL tubes and incubate in a 37 &#176;C water bath. 
		NOTE: Media can be stored at 4 &#176;C for up to one month.</li>
	</ol>
	</li>
	<li>Enzyme.
	<ol>
		<li>Weigh out 4 mg (for jejunum) or 24 mg (for colon) of Collagenase XI [&#8805;800 units/mg] in a 1.5 mL microcentrifuge tube. 
		NOTE: Collagenase is stable at -20 &#176;C for up to one year.</li>
		<li>Resuspend the collagenase aliquot in 1 mL of ice cold PBS with Ca2+ and Mg2+. Vortex the solution to thoroughly combine, and place on ice.</li>
	</ol>
	</li>
	<li>Coating Culture Plates.
	<ol>
		<li>Thaw 150 &#181;L of extracellular matrix overnight at 4 &#730;C on ice.</li>
		<li>Make a 5% w/v extracellular matrix solution by mixing 100 &#181;L of thawed extracellular matrix with 2 mL of ice-cold serum-free high-glucose DMEM. 
		NOTE: Extracellular matrix solidifies at room temperature and should be handled with pre-chilled pipettes, tubes, and media until ready to coat dishes.</li>
		<li>Immediately coat the glass inner circle of glass-bottom 35 mm dishes with 250 &#181;L of the 5% extracellular matrix solution. This recipe will cover up to 8 dishes.</li>
		<li>Place the glass-bottom dishes in a 37 &#176;C incubator during the remaining cell culture isolation steps. The extracellular matrix takes a minimum of 1 h to set but can be incubated for up to 2.5 h while performing other isolation steps. 
		NOTE: Incubations longer than 3 h can create a build-up of extracellular matrix which could impede seal formation during whole cell electrophysiology.</li>
	</ol>
	</li>
</ol>
2. Tissue Isolation
<ol>
	<li>According to ethical guidelines, sacrifice a 3to 6 week-old male or female Tph1-CFP transgenic mouse (The Jackson Laboratory Stock: 028366)9 or another reporter strain. 
	NOTE: We use a fatal dose of rising levels of carbon dioxide and a cervical dislocation as a secondary means to ensure death. The AVAM Committee on Euthanasia also considers cervical dislocation an allowable primary euthanasia method by individuals trained in its use if the animals are first sedated.</li>
	<li>Pin the euthanized mouse out on a polystyrene foam dissection stage using 21 G needles. Decontaminate the mouse abdomen with 70% ethanol.</li>
	<li>Pull mouse skin taut using micro-dissection forceps (#5), and then use sharp surgical scissors to make a transverse incision at the bottom of the mouse abdomen to expose the intraperitoneal cavity. Make another incision vertically up the abdomen until the rib cage is exposed.</li>
	<li>Use the micro-dissection forceps to move the cecum to the left of the mouse to attain a clear view of the colon.</li>
	<li>Use the micro-dissection scissors to cut the most distal portion of the colon (but still proximal to the rectum) and make a second cut distal to the stomach. Place the extracted intestines into a 100 x 15 mm2 Petri dish filled with 20 mL of ice-cold phosphate buffered saline (PBS).</li>
	<li>Use a small ruler to measure and cut out 10 cm of either mid colon or jejunum. Extract colon by measuring out the 10 cm segment of intestine located proximally to the first cut taken above the rectum. Identify the jejunum by folding the entire small intestine in half and making a first incision at the halfway point and a second incision 10 cm proximal to the first cut. 
	NOTE: Continue to perform all subsequent isolation steps on ice.</li>
	<li>Fill a 6 mL syringe with ice cold PBS and place the syringe needle into the lumen of the extracted intestinal segment. Use the micro-dissection forceps to clamp and seal the intestines around the syringe needle, and gently flush 10 mL of PBS through the lumen to rinse away fecal matter.</li>
	<li>Invert the intestinal tissue by gently weaving the micro-dissection forceps through the lumen, pinching an intestinal wall, and retracting the tissue proximal to the tip of the forceps. Repeat this process until the segment is inside-out with the lumen facing outward.</li>
	<li>Use micro-dissection scissors to cut the tissue segment into 1 cm pieces.</li>
	<li>Use micro-dissection forceps to transfer the tissue segments in a 50 mL beaker filled with 5 mL sterile PBS. Use the micro-dissection scissors to mince the tissue pieces to a liquified consistency.</li>
	<li>Place the minced tissue and 15 mL of ice cold PBS in a clean 50 mL tube by transfer pipette. Triturate 2 times with the transfer pipette, wait for the tissue to settle, remove 15 mL of the PBS supernatant, and replace with fresh PBS. Repeat this process three times until the PBS is clear.</li>
	<li>Place the 50 mL tube with washed tissue pieces on ice and bring the ice bucket into a tissue culture hood.</li>
</ol>
3. Enzymatic and Mechanical Digestion
<ol>
	<li>First digestion
	<ol>
		<li>From the water bath, retrieve one of the 50 mL tubes containing 10 mL of pre-warmed digestion media. To the 50 mL tube, add 250 &#181;L of the PBS collagenase solution and minced intestinal tissue pieces. 
		NOTE: Take care to avoid including any PBS from previous washing steps.</li>
		<li>Place the 50 mL tube in a water bath for 5 minutes (colon or jejunum) at 37 &#176;C. Shake the tube 2x per min. 
		NOTE: The optimal timing and intensity of mechanical digestion may vary and will need to be determined by the user. To determine the optimal digestion conditions, a 10 &#181;L aliquot of cells can be viewed after each digestion. During Digestion 1, the supernatant should consist of clumps of cells.</li>
		<li>Slowly triturate the tissue one time with a 10 mL serological pipette. Briefly let the tissue pieces settle, then use a transfer pipette to remove the entire 10 mL of supernatant.</li>
	</ol>
	</li>
	<li>Second digestion: Repeat steps 3.1.1 to 3.1.3. Increase the incubation time to 10 min if digesting colon and keep the incubation time at 5 min for jejunum. 
	&#8203;NOTE: Upon observation, the supernatant should still consist of large clumps of cells.</li>
	<li>Third digestion
	<ol>
		<li>Repeat step 3.1.1</li>
		<li>Place the 50-mL tube in a 37 &#730;C water bath for 10 min (colon or small jejunum). Shake the tube 2-3x per min at a higher intensity than Digestions 1 and 2.</li>
		<li>Slowly pipette the tissue up and down with a 10-mL serological pipette. Allow the tissue pieces to settle for a short time. 
		NOTE: The supernatant should consist of a combination of single cells and small clumps.</li>
		<li>Use a transfer pipette to place 10 mL of the supernatant into a new 15 mL tube, add 2 mL of warmed EC cell complete culture media, and invert once to mix.</li>
		<li>Spin at 100 x g for 5 min at room temperature, then use a transfer pipette to remove the supernatant. Use a transfer pipette to resuspend the remaining pellet in 2.5 mL of warmed EC cell complete culture media.</li>
	</ol>
	</li>
	<li>Fourth digestion: Repeat steps 3.3.1 to 3.3.5. Increase the incubation time to 15 min for colon and remain at 10 min for jejunum. 
	NOTE: The supernatant should now consist of single cells.</li>
</ol>
4. Cell Culture
<ol>
	<li>Use a transfer pipette to combine the cell suspensions collected from Digestions 3 and 4 into a new 15 mL tube (5 mL total volume).</li>
	<li>Remove a 10 &#181;L aliquot of cells to count with a hemocytometer and spin the remaining cell suspension at 100 x g for 5 min at room temperature. 
	NOTE: The final cell suspension will consist of small clumps and single cells.</li>
	<li>Remove the supernatant with a transfer pipette and resuspend the pellet in EC cell complete culture media at a density of 1,000,000 cells per mL. 
	NOTE: The final volume of cell suspension will depend on the final cell count. Typical cell counts range from 2,000,000 to 4,000,000.</li>
	<li>Remove coated glass-bottom culture dishes from the incubator. Use a P1000 pipette to replace extracellular matrix from each culture dish with 250 &#181;L of the final cell suspension. 
	NOTE: Extracellular matrix coating tends to stick to the edges of the glass-bottom dish. Special care must be taken to remove the coating from along the edge to prevent gel buildup.</li>
	<li>Make a stock solution of ROCK inhibitor Y-27632 at 1 mM. Add 2.5 &#181;L of stock solution to each glass bottom dish to achieve a working concentration of 10 &#181;M ROCK inhibitor in each dish. 
	NOTE: This step is critical for the survival of jejunum cultures but is optional for colon.</li>
	<li>Place each culture dish in a 37 &#176;C and 5% CO2 incubator for 24 to 72 h (Figure 1).</li>
</ol>
5. Preparation of EC Cells for Whole Cell Electrophysiology
<ol>
	<li>Line the inner diameter of the microscope stage with two 5 x 0.5-cm strips of wax film to create an O-ring. Mount the 35-mm cell culture dish within the O-ring (Figure 2A-B).</li>
	<li>Rinse both floating debris, such as unattached extracellular matrix or dead cells, and culture media serum completely away from EC cells to prevent either from impeding seal formation between the EC cell and electrode. Use of the three following options to achieve thorough cell washing sufficient for electrophysiology:
	<ol>
		<li>Option 1: Solution exchange is more efficient in a long chamber more so than a circular chamber. Since the EC cells were cultured in round 35 mm dishes, create a plastic elliptical insert for the dish.
		<ol>
			<li>Create the insert by 3D printing or by traditional milling methods. The insert we used was milled from acrylic plastic with the following dimensions (in mm): outer diameter, 34.5; inner ellipse, 20x9; outer height, 10; inner height, 1.5; bridge span, 3x3; bridge clearance, 1.5; inlet and outlet, 4 x 4 each.</li>
			<li>Lower the insert into the culture dish (Figure 2A-B) and secure it to the top of the microscope stage with two 0.15&#160;to 0.2 g pieces of modeling clay pressed into 1.2 x 0.3 x 0.1 cm rectangles (Figure 2A-C).</li>
			<li>With a plastic transfer pipette, slowly add extracellular solution to one side of the dish (inlet) while aspirating from the other side (outlet).</li>
		</ol>
		</li>
		<li>Option 2: Without an engineered plastic insert, add extracellular solution by transfer pipette from one side of the dish (inlet) while aspirating from the opposite side (outlet). Slowly rotate the location of the transfer pipette (e.g., N to E to S) while mirroring the placement of the aspiration needle on the opposite side (e.g., S to W to N).</li>
		<li>Option 3: Coat extracellular matrix onto rectangular coverslips in step 1.3.3., and culture the cell suspension on these coverslips in step 4.4. Transfer this coverslip to a rectangular chamber filled with extracellular solution, omitting step 5.1 and skipping step 5.3. Rinse the chamber with extracellular solution from a plastic transfer pipette, as described in Option 1 (5.2.1.3).</li>
	</ol>
	</li>
	<li>Re-attach the stage with the cell culture above the inverted microscope (Figure 2D). Incubate the EC cell culture in serum-free extracellular solution at room temperature.</li>
	<li>After 4 hours, rinse the culture again with extracellular solution as described above. Proceed with whole cell electrophysiology.</li>
</ol>
6. Whole Cell Electrophysiology of EC Cells from Primary Culture
<ol>
	<li>Achieving a Whole Cell Giga-seal
	<ol>
		<li>Pull one dozen electrodes to a resistance of 1&#8211;5 M&#8486;. 
		NOTES: Pipettes of 1&#8211;2 M&#8486;&#160;resistance that yield 10-100 G&#8486;&#160;seals perform best. Fire polishing the electrode tips is not necessary to achieve G&#8486; seals.</li>
		<li>Coat evenly the nose cone of all pipettes with polymer. Hold the pipette horizontal to the ground and perpendicular to the front of a heat gun. Slowly rotate the pipette until the polymer hardens.</li>
		<li>Pour an aliquot of intracellular solution into a 15-mL conical tube. Transfer 1.2 mL of this solution into a 1.5 mL microcentrifuge tube via 1-mL syringe. Withdraw another 0.7 mL of intracellular solution into the syringe. Dispel air from the end of the syringe. Attach a flexible 34 G, 67 mm needle to the syringe, and flush out any remaining air from this needle.</li>
		<li>Identify a Tph1-CFP+ cell that is not in contact with any other cell or debris.</li>
		<li>Fill the electrode tip with intracellular solution. Backfill the pipette by syringe. Carefully flick the pipette with a fingernail to dispel the air bubble interface.</li>
		<li>With the plunger drawn up with 0.3 mL of air, attach a 6 mL syringe to the outlet of the holder tubing. Mount the electrode into the holder. 
		NOTE: Connect the syringe to the tubing before the electrode enters the bath solution to maintain neutral or positive pressure within the intracellular solution prior to sealing.</li>
		<li>Load a two-step voltage-clamp protocol that records steady-state activating currents by a voltage ladder on the first step and records the available currents by a single voltage on the second step. Open the seal test (5 mV at 50 kHz).</li>
		<li>Lower the electrode into the bath solution. With the micromanipulators, maneuver the electrode tip in-plane with and directly horizontal from the cell. Expel the 0.3 mL of air from the syringe.</li>
		<li>Gently move the electrode horizontally along the x-axis to touch the cell. Watch for a dimple to appear on the EC cell and/or an increase in membrane resistance on the seal test. If necessary, readjust the electrode down along the z-axis and/or horizontally along the x-axis, not moving past the midline of the cell.</li>
		<li>Apply 0.1&#8211;0.2 mL suction on the 6-mL syringe. Hold the plunger steady until 100 M&#8486; is achieved, then release grip from the plunger. Wait for the cell to giga-seal. Gently disconnect the syringe with an unscrewing motion. 
		NOTE: If an EC cell does not seal initially, it might be possible to re-seal it with a subsequent attempt. To do this, gently pull the electrode away from an EC cell with &#60;20 M&#8486; seal resistance. Steering the spent electrode with the micromanipulators, circumnavigate the targeted EC cell to manually clear away matrix or debris. Then, attempt to seal the EC cell with a fresh electrode.</li>
	</ol>
	</li>
	<li>Record whole cell voltage-gated Na+ current.
	<ol>
		<li>Attach a 3 mL syringe drawn up with 0.5&#8211;1.0 mL air. Apply a quick tap of suction (0.1&#8211;0.5 mL) on a 3 mL syringe. Repeat until whole cell access is achieved, then gently disconnect the syringe. 
		NOTE: As the performance of a syringe can vary initially and change gradually with time and use, practice beforehand with the 3 mL syringe (sealing the end with an index finger) to ensure that the plunger will recoil within 0.1&#8211;0.2 mL of its starting position. If not, then exchange the syringe for a new one.</li>
		<li>Turn on whole cell capacitance compensation. Adjust capacitance and series resistance. Turn on series resistance compensation. With lag set at 60 ms, adjust predicted then corrected series resistance compensation. Close the seal test.</li>
		<li>Record an average of 5 runs of whole-cell voltage-gated Na+ current (Figure 3A).</li>
		<li>Quickly take note of two parameters of voltage-gated Na+ current: the voltage at window current (Figure 3B, red symbols, the intersection of steady-state activation and inactivation curves) and the peak Na+ current density. 
		NOTE: EC cells with peak Na+ currents more than -50 pA in voltage clamp mode commonly will go on to fire elicited potentials in current clamp mode. Additionally, currents greater than -200 pA in voltage clamp (Figure 3A) then usually fire action potentials spontaneously in current clamp from membrane potentials (red line, Figure 3D) near the voltage of the window current (red symbols, Figure 3B).</li>
	</ol>
	</li>
	<li>Record elicited action potentials.
	<ol>
		<li>Turn off the series resistance compensation. Turn off the external command. Switch the mode from V-clamp to I-clamp. Adjust the gain. Load an episodic current clamp protocol. Adjust the amount of holding current to 0 pA. Turn on the external command. 
		CAUTION: Do not switch from V-clamp to I-clamp (or vice versa) without first turning off the external command.</li>
		<li>Note the resting membrane potential. Adjust the holding current input so that the membrane potential reads below the window current (e.g., -80 to -100 mV). Adjust the episodic current clamp protocol so that the step interval of current input is less than the value of the holding current (Figure 3C inset, +2 pA steps for -3 pA holding current).</li>
		<li>Record elicited action potentials. Note the least amount of current injected to fire off an action potential (Figure 3C, sweep 3 of 5, a +4 pA step from -3 pA holding = +1 pA).</li>
	</ol>
	</li>
	<li>Record spontaneous action potentials.
	<ol>
		<li>Turn off the external command. Load a gap-free (non-episodic) current clamp protocol. 
		NOTE: Do not turn the external command back on until the amount of holding current in the gap-free current clamp protocol has been verified to be suitable for the cell.</li>
		<li>Change the holding current to be the amount of current injected in the last sweep in the elicited protocol that did not fire an action potential (Figure 3D, the current input during the second sweep, a +2 pA step from -3 pA holding = -1 pA).</li>
		<li>Turn on the external command. Double check that the predicted holding current brings the membrane potential below the threshold to fire an action potential. Adjust the holding current if necessary. Record spontaneous activity.</li>
	</ol>
	</li>
</ol>

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Fully understanding EC cell function requires a high-quality primary culture method to generate cells for whole cell electrophysiology. Primary cultures of GI epithelium had traditionally been difficult due to low survival. Alleviating this confounding factor, the present method yields cultures of singular EC cells from either small or large bowel that are capable of surviving for several days.
We found that the following were critical for improving the quality of cultures to be suitable for e...

The gastrointestinal (GI) epithelium is a diverse community consisting of several cell types. Enteroendocrine cells comprise roughly 1% of all epithelial cells, and enterochromaffin (EC) cells are the largest enteroendocrine cell population1. Recent studies show that enteroendocrine2 and EC3,4 cells are electrically excitable. We are interested in understanding primary EC cell electrophysiology. Thus, the purpose of this study was to establish primary EC cell cultures optimized for whole cell electrophysiology.
Existing EC cell lines that produce and secrete 5-HT (e.g.,&#160;QGP-15, BON6, KRJ-17) and have been used to examine electrophysiology5,8 are typically generated from immortalized neoplastic tissues. While the information acquired from these cell lines is valuable5,8, studies of primary cell electrophysiology are necessary to properly understand EC cell physiology. The electrophysiology of primary EC cells requires the isolation and culture of single epithelial cells, which has been limited by low viability of epithelial cultures.
The culture method presented in this study relies on transgenic mice with fluorescently labeled EC cells, like Tph1-CFP9, used in this study. The method optimizes mixed primary epithelial cultures developed previously2,10 for single cell electrophysiology3. Previous methods used combinations of Trypsin/EDTA plus Collagenase A or EDTA and DTT for enzymatic digestion and used density gradients to specifically isolate and culture guinea-pig11 and rat EC cells12. More recently, intestinal organoids were generated and mechanically disrupted for electrophysiological recordings4. Cultures using these methods are well suited for serotonin release experiments, RT-qPCR analysis and, while they could be used for electrophysiology, are reliant on the success of time consuming organoid generation, density gradients to identify EC cells, and the cellular disruptive effects of Trypsin and EDTA. By contrast, the protocol described here improves enzymatic and mechanical treatments, optimizes culturing conditions, and streamlines procedures to produce single and healthy EC cells suitable for the high cellular standards needed for electrophysiology.
This method will be useful for scientists who want to work on primary EC cells in mixed cultures rather than immortalized cultures and want to investigate the electrophysiology of single cells. However, it may prove less appropriate for studying cell populations that need sorting or long-term cultures that require survival past 72 h.

<table><tbody><tr><td>100 mm x 15 mm culture dish</td><td>Corning (Falcon)</td><td>351029</td><td></td></tr><tr><td>Transfer Pipette</td><td>Fisherbrand</td><td>13-711-7M</td><td></td></tr><tr><td>10 mL serological pipette</td><td>Corning (Falcon)&amp;nbsp;</td><td>357551</td><td></td></tr><tr><td>50 mL Conical</td><td>Corning (Falcon)&amp;nbsp;</td><td>352098</td><td></td></tr><tr><td>15 mL Conical</td><td>Corning (Falcon)&amp;nbsp;</td><td>352097</td><td></td></tr><tr><td>1,000 mL Barrier Pipet Tips</td><td>Thermo Scientific (Art)</td><td>2079E</td><td>Keep sterile</td></tr><tr><td>MatTek Glass Bottom Dishes</td><td>MatTek Corporation</td><td>P35G-1.5-14-C</td><td>Keep sterile</td></tr><tr><td>Micro-dissection Forceps - Very Fine, Extra-Long Points</td><td>Fine Science Tools</td><td>SB12630M</td><td></td></tr><tr><td>Surgical Scissors-Sharp</td><td>Nasco</td><td>14002-14</td><td></td></tr><tr><td>Micro-Dissection Scissors</td><td>Nasco</td><td>SB46568M</td><td></td></tr><tr><td>Monoject Hypo Needle, 21 G x 1&quot; A, 100/BX</td><td>Covidien</td><td>8881250172</td><td></td></tr><tr><td>&amp;nbsp;Tg(Tph1-CFP)1Able/J Mice</td><td>Jackson</td><td>Stock # 028366</td><td>Use male or female mice between the ages of 4-7 weeks</td></tr><tr><td>Microfil nonmetallic syringe needles (34 gauge, 67 mm)</td><td>World Precision Instruments</td><td>MF34G-5</td><td></td></tr><tr><td>Parafilm &quot;M&quot; Laboratory Film</td><td>Pechiney Plastic Packaging</td><td></td><td></td></tr><tr><td>R6101</td><td>Dow Corning</td><td></td><td></td></tr><tr><td>6 mL syringe with regular luer tip</td><td>Monoject</td><td></td><td></td></tr><tr><td>3 mL syringe with regular luer tip</td><td>Monoject</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Electrophysiology Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Amplifier</td><td>Molecular Devices</td><td>Axopatch 200B</td><td></td></tr><tr><td>Data acquisition system (daq)</td><td>Molecular Devices</td><td>Digidata 1440A</td><td></td></tr><tr><td>Signal conditioner</td><td>Molecular Devices</td><td>CyberAmp 320</td><td></td></tr><tr><td>Software</td><td>Molecular Devices</td><td>pClamp 10.6</td><td></td></tr></tbody></table>

whole cell electrophysiology of primary cultured murine enterochromaffin cells

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory cells, EC cells sense luminal stimuli and convert them into serotonin (5-hyroxytryptamine, 5-HT) release events. However, the electrophysiology of these cells is poorly understood because they are difficult to culture and to identify. The method presented in this paper outlines primary EC cell cultures optimized for single cell electrophysiology. This protocol utilizes a transgenic cyan fluorescent protein (CFP) reporter to identify mouse EC cells in mixed primary cultures, advancing the approach to obtaining high-quality recordings of whole cell electrophysiology in voltage- and current-clamp modes.

Cultures: 
Primary murine epithelial cells made from a transgenic Tph1-CFP model attach to dishes after 4 hours and are ready for physiological experiments between 24 to 72 h. When culture conditions had not been optimized, the epithelial cell culture consisted of large clumps, floating cellular debris, damaged membranes, and weak CFP signal in the EC cells (Figure 1A). Cultures that have been optimized for electrophysio...

Watch this Scientific Journal Video about Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells at JoVE.com

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Enterochromaffin (EC) cells comprise a small subset of gastrointestinal epithelial cells. EC cells are electrically excitable and release serotonin, yet difficulties in culturing and identifying EC cells have limited physiological studies. The method presented here establishes a primary culture model amenable to examination of single EC cells by electrophysiology.

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory ...

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Research

JoVE Journal

Biology

10.2K Views.  Mayo Clinic. Deze methode kan helpen bij het beantwoorden van belangrijke vragen in de enterische neurowetenschappen en gastro-intestinale fysiologie, zoals, wat zijn de mechanismen van enterochromaffin cel excitability en reactie op luminale stimuli, en wat zijn de mechanismen van gastro-intestinale hormoon release? Het belangrijkste voordeel van deze techniek is dat het ons in staat stelt om enterochromaffincellen in detail te bestuderen met behulp van eencellige technieken, zoals elektrofysiologie. Het...

Video: Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Whole-cell Patch-clamp Recordings of Isolated Primary Epithelial Cells from the Epididymis

The epididymis is an essential organ for sperm maturation and reproductive health. The epididymal epithelium consists of intricately connected cell types that are distinct not only in molecular and morphological features but also in physiological properties. These differences reflect their diverse functions, which together establish the necessary microenvironment for the post-testicular sperm development in the epididymal lumen. The understanding of the biophysical properties of the epididymal epithelial cells is critical for revealing their functions in sperm and reproductive health, under both physiological and pathophysiological conditions. While their functional properties have yet to be fully elucidated, the epididymal epithelial cells can be studied using the patch-clamp technique, a tool for measuring the cellular events and the membrane properties of single cells. Here, we describe the methods of cell isolation and whole-cell patch-clamp recording to measure the electrical properties of primary dissociated epithelial cells from the rat cauda epididymides.

We present a protocol that combines cell isolation and whole-cell patch-clamp recording to measure the electrical properties of the primary dissociated epithelial cells from the rat cauda epididymides. This protocol allows for investigation of the functional properties of primary epididymal epithelial cells to further elucidate the physiological role of the epididymis.

The epididymis is an essential organ for sperm maturation and reproductive health. The epididymal epithelium consists of intricately connected cell types ...

Developmental Biology

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls gastrointestinal motility. A procedure for the isolation and culture of a mixed population of neurons and glia from the myenteric plexus is described. The primary cultures can be maintained for over 7 days, with connections developing among the neurons and glia. The longitudinal muscle strip with the attached myenteric plexus is stripped from the underlying circular muscle of the mouse ileum or colon and subjected to enzymatic digestion. In sterile conditions, the isolated neuronal and glia population are preserved within the pellet following centrifugation and plated on coverslips. Within 24-48 hr, neurite outgrowth occurs and neurons can be identified by pan-neuronal markers. After two days in culture, isolated neurons fire action potentials as observed by patch clamp studies. Furthermore, enteric glia can also be identified by GFAP staining. A network of neurons and glia in close apposition forms within 5 - 7 days. Enteric neurons can be individually and directly studied using methods such as immunohistochemistry, electrophysiology, calcium imaging, and single-cell PCR. Furthermore, this procedure can be performed in genetically modified animals. This methodology is simple to perform and inexpensive. Overall, this protocol exposes the components of the enteric nervous system in an easily manipulated manner so that we may better discover the functionality of the ENS in normal and disease states.

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

We demonstrate a cell culture protocol for the direct study of neuronal and glial components of the enteric nervous system. A neuron/glia mixed culture on coverslips is prepared from the myenteric plexus of adult mouse providing the ability to examine individual neuron and glia function by electrophysiology, immunohistochemical, etc.

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls ...

Neuroscience

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

In vitro Functional Characterization of Mouse Colorectal Afferent Endings

This video demonstrates in detail an in vitro single-fiber electrophysiological recording protocol using a mouse colorectum-nerve preparation. The approach allows unbiased identification and functional characterization of individual colorectal afferents. Extracellular recordings of propagated action potentials (APs) that originate from one or a few afferent (i.e., single-fiber) receptive fields (RFs) in the colorectum are made from teased nerve fiber fascicles. The colorectum is removed with either the pelvic (PN) or lumbar splanchnic (LSN) nerve attached and opened longitudinally. The tissue is placed in a recording chamber, pinned flat and perfused with oxygenated Krebs solution. Focal electrical stimulation is used to locate the colorectal afferent endings, which are further tested by three distinct mechanical stimuli (blunt probing, mucosal stroking and circumferential stretch) to functionally categorize the afferents into five mechanosensitive classes. Endings responding to none of these mechanical stimuli are categorized as mechanically-insensitive afferents (MIAs). Both mechanosensitive and MIAs can be assessed for sensitization (i.e., enhanced response, reduced threshold, and/or acquisition of mechanosensitivity) by localized exposure of RFs to chemicals (e.g., inflammatory soup (IS), capsaicin, adenosine triphosphate (ATP)). We describe the equipment and colorectum&#8211;nerve recording preparation, harvest of colorectum with attached PN or LSN, identification of RFs in the colorectum, single-fiber recording from nerve fascicles, and localized application of chemicals to the RF. In addition, challenges of the preparation and application of standardized mechanical stimulation are also discussed.

In vitro Functional Characterization of Mouse Colorectal Afferent Endings

This video demonstrates a protocol for conducting single-fiber electrophysiological recordings on an in vitro mouse colorectum-nerve preparation.

This video demonstrates in detail an in vitro single-fiber electrophysiological recording protocol using a mouse colorectum-nerve preparation. The ...

Medicine

Prolonged Incubation of Acute Neuronal Tissue for Electrophysiology and Calcium-imaging

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the experimental time, and increases the number of animals that are utilized per study. This limitation specifically impacts protocols such as calcium imaging that require prolonged pre-incubation with bath-applied dyes. Exponential bacterial growth within 3 - 4 h after slicing is tightly correlated with a decrease in tissue health. This study describes a method for&#160;limiting the proliferation of bacteria in acute preparations to maintain viable neuronal tissue for prolonged periods of time (&#62;24 h) without the need for antibiotics, sterile procedures, or tissue culture media containing growth factors. By cycling the extracellular fluid through UV irradiation and keeping the tissue in a custom holding chamber at 15 - 16 &#176;C, the tissue shows no difference in electrophysiological properties, or calcium signaling through intracellular calcium dyes at &#62;24 h postdissection. These methods will not only extend experimental time for those using acute neuronal tissue, but will reduce the number of animals required to complete experimental goals, and will set a gold standard for acute neuronal tissue incubation.

Once removed from the body, neuronal tissue is greatly affected by environmental conditions, leading to eventual degradation of the tissue after 6 - 8 h. Using a unique incubation method, which closely monitors and regulates the extracellular environment of the tissue, tissue viability can be significantly extended for &#62;24 h.

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the ...

Isolation and Kv Channel Recordings in Murine Atrial and Ventricular Cardiomyocytes

KCNE genes encode for a small family of Kv channel ancillary subunits that form heteromeric complexes with Kv channel alpha subunits to modify their functional properties. Mutations in KCNE genes have been found in patients with cardiac arrhythmias such as the long QT syndrome and/or atrial fibrillation. However, the precise molecular pathophysiology that leads to these diseases remains elusive. In previous studies the electrophysiological properties of the disease causing mutations in these genes have mostly been studied in heterologous expression systems and we cannot be sure if the reported effects can directly be translated into native cardiomyocytes. In our laboratory we therefore use a different approach. We directly study the effects of KCNE gene deletion in isolated cardiomyocytes from knockout mice by cellular electrophysiology - a unique technique that we describe in this issue of the Journal of Visualized Experiments. The hearts from genetically engineered KCNE mice are rapidly excised and mounted onto a Langendorff apparatus by aortic cannulation. Free Ca2+ in the myocardium is bound by EGTA, and dissociation of cardiac myocytes is then achieved by retrograde perfusion of the coronary arteries with a specialized low Ca2+ buffer containing collagenase. Atria, free right ventricular wall and the left ventricle can then be separated by microsurgical techniques. Calcium is then slowly added back to isolated cardiomyocytes in a multiple step comprising washing procedure. Atrial and ventricular cardiomyocytes of healthy appearance with no spontaneous contractions are then immediately subjected to electrophysiological analyses by patch clamp technique or other biochemical analyses within the first 6 hours following isolation.

Kv channel dysfunction is associated with cardiac arrhythmias. In order to study the molecular mechanisms that lead to such arrhythmias we utilize a systematic protocol for isolation of atrial and ventricular cardiomyocytes from Kv channel ancillary subunit knockout mice. Isolated cardiomyocytes can then immediately be used for cellular electrophysiological studies, biochemical or immunofluorescence (IF) assays.

KCNE genes encode for a small family of Kv channel ancillary subunits that form heteromeric complexes with Kv channel alpha subunits to modify their ...

Confocal Microscopy to Measure Three Modes of Fusion Pore Dynamics in Adrenal Chromaffin Cells

Dynamic fusion pore opening and closure mediate exocytosis and endocytosis and determine their kinetics. Here, it is demonstrated in detail how confocal microscopy was used in combination with patch-clamp recording to detect three fusion modes in primary culture bovine adrenal chromaffin cells. The three fusion modes include 1) close-fusion (also called kiss-and-run), involving fusion pore opening and closure, 2) stay-fusion, involving fusion pore opening and maintaining the opened pore, and 3) shrink-fusion, involving shrinkage of the fusion-generated &#937;-shape profile until it merges completely at the plasma membrane.
To detect these fusion modes, the plasma membrane was labeled by overexpressing mNeonGreen attached with the PH domain of phospholipase C &#948; (PH-mNG), which binds to phosphatidylinositol-4,5-bisphosphate (PtdIns(4,5)P2) at the cytosol-facing leaflet of the plasma membrane; vesicles were loaded with the fluorescent false neurotransmitter FFN511 to detect vesicular content release; and Atto 655 was included in the bath solution to detect fusion pore closure. These three fluorescent probes were imaged simultaneously at ~20-90 ms per frame in live chromaffin cells to detect fusion pore opening, content release, fusion pore closure, and fusing vesicle size changes. The analysis method is described to distinguish three fusion modes from these fluorescence measurements. The method described here can, in principle, apply to many secretory cells beyond chromaffin cells.

This protocol describes a confocal imaging technique to detect three fusion modes in bovine adrenal chromaffin cells. These fusion modes include 1) close-fusion (also called kiss-and-run), involving fusion pore opening and closure, 2) stay-fusion, involving fusion pore opening and maintaining the opened pore, and 3) shrink-fusion, involving fused vesicle shrinkage.

Dynamic fusion pore opening and closure mediate exocytosis and endocytosis and determine their kinetics. Here, it is demonstrated in detail how confocal ...

Mixed Primary Cultures of Murine Small Intestine Intended for the Study of Gut Hormone Secretion and Live Cell Imaging of Enteroendocrine Cells

The gut is the largest endocrine organ of the body, with hormone-secreting enteroendocrine cells located along the length of the gastrointestinal epithelium. Despite their physiological importance, enteroendocrine cells represent only a small fraction of the epithelial cell population and in the past, their characterization has presented a considerable challenge resulting in a reliance on cell line models. Here, we provide a detailed protocol for the isolation and culture of mixed murine small intestinal cells. These primary cultures have been used to identify the signaling pathways underlying the stimulation and inhibition of gut peptide secretion in response to a number of nutrients and neuropeptides as well as pharmacological agents. Furthermore, in combination with the use of transgenic fluorescent reporter mice, we have demonstrated that these primary cultures become a powerful tool for the examination of fluorescently-tagged enteroendocrine cells at the intracellular level, using methods such as patch clamping and single-cell calcium and cAMP-FRET imaging.

This protocol describes the isolation and culture of mixed murine small intestinal cells. These primary intestinal cultures enable the investigation of signaling pathways underlying gut peptide secretion using a number of techniques.

The gut is the largest endocrine organ of the body, with hormone-secreting enteroendocrine cells located along the length of the gastrointestinal ...

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.

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, ...

Two-dimensional Porcine Intestinal Organoids Reflecting the Physiological Properties of Native Gut

The gastrointestinal tract (GIT) serves both in the digestion of food and the uptake of nutrients but also as a protective barrier against pathogens. Traditionally, research in this area has relied on animal experiments, but there's a growing demand for alternative methods that adhere to the 3R principles-replace, reduce, and refine. Porcine organoids have emerged as a promising tool, offering a more accurate in vitro replication of the in vivo conditions than traditional cell models. One major challenge with intestinal organoids is their inward-facing apical surface and outward-facing basolateral surface. This limitation can be overcome by creating two-dimensional (2D) organoid layers on transwell inserts (from here on referred to as insert(s)), providing access to both surfaces. In this study, we successfully developed two-dimensional cultures of porcine jejunum and colon organoids. The cultivation process involves two key phases: First, the formation of a cellular monolayer, followed by the differentiation of the cells using tailored media. Cellular growth is tracked by measuring transepithelial electrical resistance, which stabilizes by day 8 for colon organoids and day 16 for jejunum organoids. After a 2-day differentiation phase, the epithelium is ready for analysis. To quantify and track active electrogenic transport processes, such as chloride secretion, we employ the Ussing chamber technique. This method allows for real-time measurement and detailed characterization of epithelial transport processes. This innovative in vitro model, combined with established techniques like the Ussing chamber, provides a robust platform for physiologically characterizing the porcine GIT within the 3R framework. It also opens opportunities for investigating pathophysiological mechanisms and developing potential therapeutic strategies.

This study outlines a protocol for generating 2D monolayers of porcine organoids derived from the small and large intestines. The growth of these monolayers is marked by increasing TEER values, indicating robust epithelial integrity. Additionally, these monolayers exhibit physiological secretory responses in Ussing chamber experiments following the application of forskolin.

The gastrointestinal tract (GIT) serves both in the digestion of food and the uptake of nutrients but also as a protective barrier against pathogens. ...

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

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