Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Summary

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.

Abstract

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.

Introduction

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 EC^3,4 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., QGP-1⁵, BON⁶, KRJ-1⁷) and have been used to examine electrophysiology^5,8 are typically generated from immortalized neoplastic tissues. While the information acquired from these cell lines is valuable^5,8, 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 previously^2,10 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.

Protocol

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

1. Media.
   1. Prepare EC cell complete culture media with high glucose [4500mg/L] Dulbecco'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 °C for up to one month.
   2. Filter the EC cell complete culture media and place 25 mL of media into a 50 mL tube at 37 °C.
   3. Prepare digestion media with high glucose [4,500 mg/L] DMEM supplemented with 0.1% Bovine Serum Albumin (BSA) (Table 1).
   4. Filter the digestion media and place four 10 mL aliquots of media into separate 50 mL tubes and incubate in a 37 °C water bath.
      NOTE: Media can be stored at 4 °C for up to one month.
2. Enzyme.
   1. Weigh out 4 mg (for jejunum) or 24 mg (for colon) of Collagenase XI [≥800 units/mg] in a 1.5 mL microcentrifuge tube.
      NOTE: Collagenase is stable at -20 °C for up to one year.
   2. 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.
3. Coating Culture Plates.
   1. Thaw 150 µL of extracellular matrix overnight at 4 ˚C on ice.
   2. Make a 5% w/v extracellular matrix solution by mixing 100 µL 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.
   3. Immediately coat the glass inner circle of glass-bottom 35 mm dishes with 250 µL of the 5% extracellular matrix solution. This recipe will cover up to 8 dishes.
   4. Place the glass-bottom dishes in a 37 °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.

2. Tissue Isolation

1. 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.
2. Pin the euthanized mouse out on a polystyrene foam dissection stage using 21 G needles. Decontaminate the mouse abdomen with 70% ethanol.
3. 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.
4. Use the micro-dissection forceps to move the cecum to the left of the mouse to attain a clear view of the colon.
5. 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).
6. 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.
7. 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.
8. 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.
9. Use micro-dissection scissors to cut the tissue segment into 1 cm pieces.
10. 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.
11. 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.
12. Place the 50 mL tube with washed tissue pieces on ice and bring the ice bucket into a tissue culture hood.

3. Enzymatic and Mechanical Digestion

1. First digestion
   1. 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 µL of the PBS collagenase solution and minced intestinal tissue pieces.
      NOTE: Take care to avoid including any PBS from previous washing steps.
   2. Place the 50 mL tube in a water bath for 5 minutes (colon or jejunum) at 37 °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 µL aliquot of cells can be viewed after each digestion. During Digestion 1, the supernatant should consist of clumps of cells.
   3. 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.
2. 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.
   ​NOTE: Upon observation, the supernatant should still consist of large clumps of cells.
3. Third digestion
   1. Repeat step 3.1.1
   2. Place the 50-mL tube in a 37 ˚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.
   3. 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.
   4. 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.
   5. 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.
4. 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.

4. Cell Culture

1. 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).
2. Remove a 10 µL 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.
3. 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.
4. Remove coated glass-bottom culture dishes from the incubator. Use a P1000 pipette to replace extracellular matrix from each culture dish with 250 µL 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.
5. Make a stock solution of ROCK inhibitor Y-27632 at 1 mM. Add 2.5 µL of stock solution to each glass bottom dish to achieve a working concentration of 10 µM ROCK inhibitor in each dish.
   NOTE: This step is critical for the survival of jejunum cultures but is optional for colon.
6. Place each culture dish in a 37 °C and 5% CO₂ incubator for 24 to 72 h (Figure 1).

5. Preparation of EC Cells for Whole Cell Electrophysiology

1. 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).
2. 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:
   1. 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.
      1. 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.
      2. Lower the insert into the culture dish (Figure 2A-B) and secure it to the top of the microscope stage with two 0.15 to 0.2 g pieces of modeling clay pressed into 1.2 x 0.3 x 0.1 cm rectangles (Figure 2A-C).
      3. With a plastic transfer pipette, slowly add extracellular solution to one side of the dish (inlet) while aspirating from the other side (outlet).
   2. 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).
   3. 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).
3. 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.
4. After 4 hours, rinse the culture again with extracellular solution as described above. Proceed with whole cell electrophysiology.

6. Whole Cell Electrophysiology of EC Cells from Primary Culture

1. Achieving a Whole Cell Giga-seal
   1. Pull one dozen electrodes to a resistance of 1–5 MΩ.
      NOTES: Pipettes of 1–2 MΩ resistance that yield 10-100 GΩ seals perform best. Fire polishing the electrode tips is not necessary to achieve GΩ seals.
   2. 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.
   3. 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.
   4. Identify a Tph1-CFP+ cell that is not in contact with any other cell or debris.
   5. 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.
   6. 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.
   7. 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).
   8. 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.
   9. 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.
   10. Apply 0.1–0.2 mL suction on the 6-mL syringe. Hold the plunger steady until 100 MΩ 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 <20 MΩ 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.
2. Record whole cell voltage-gated Na⁺ current.
   1. Attach a 3 mL syringe drawn up with 0.5–1.0 mL air. Apply a quick tap of suction (0.1–0.5 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 0.1–0.2 mL of its starting position. If not, then exchange the syringe for a new one.
   2. 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.
   3. Record an average of 5 runs of whole-cell voltage-gated Na⁺ current (Figure 3A).
   4. 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).
3. Record elicited action potentials.
   1. 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.
   2. 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).
   3. 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).
4. Record spontaneous action potentials.
   1. 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.
   2. 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).
   3. 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.

Results

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...

Discussion

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...

Materials

Name	Company	Catalog Number	Comments
High Glucose DMEM	Sigma	D6546	Store at 4˚ C for no longer than 1 month
Fetal Bovine Serum (FBS), Heat Inactivated	Sigma	F4135	Freeze in 25 mL aliquots  and store long term at -20˚ C
L-Glutamine	Life Technologies	25030-081	Freeze in 5 mL aliquots and store long term at -20˚ C
Penicillin/ Streptomycin	Sigma	P0781	Freeze in 5 mL aliquots and store long term at -20˚ C
Bovine Serum Albumin (BSA)	Sigma	A2153	Store at 4˚ C for long term storage
Collagenase XI	Sigma	C9407	Store at -20˚ C in aliquots of 100mg, avoid freeze thaw and avoid lot to lot variation
Phosphate Buffered Saline (PBS), w/ Ca and Mg	Sigma	D8662	Store long term at   4˚ C
Y-27632 (Rock Inhibitor)	StemCell	72304	Resuspend at 5mM and store in small aliquots at -80˚ C
100x15mm culture dish	Corning (Falcon)	351029
Transfer Pipette	Fisherbrand	13-711-7M
10 mL serological pipette	Corning (Falcon)	357551
50 mL Conical	Corning (Falcon)	352098
15 mL Conical	Corning (Falcon)	352097
1000 mL Barrier Pipet Tips	Thermo Scientific (Art)	2079E	Keep sterile
Matrigel, Growth Factor Reduced	Corning	354230	Store at 150uL aliquots at -20˚ C. Thaw on ice and keep cold until ready for plating
MatTek Glass Bottom Dishes	MatTek Corporation	P35G-1.5-14-C	Keep sterile
Micro-dissection Forceps - Very Fine, Extra-Long Points	Fine Science Tools	SB12630M
Surgical Scissors-Sharp	Nasco	14002-14
Micro-Dissection Scissors	Nasco	SB46568M
Monoject Hypo Needle, 21G x 1" A, 100/BX	Covidien	8881250172
Tg(Tph1-CFP)1Able/J Mice	Jackson	Stock # 028366	Use male or female mice between the ages of 4-7 weeks
Microfil nonmetallic syringe needles (34 gauge, 67 mm)	World Precision Instruments	MF34G-5
Parafilm "M" Laboratory Film	Pechiney Plastic Packaging
R6101	Dow Corning
6-mL syringe with regular luer tip	Monoject
3-mL syringe with regular luer tip	Monoject
Electrophysiology Equipment
Amplifier	Molecular Devices	Axopatch 200B
Data acquisition system (daq)	Molecular Devices	Digidata 1440A
Signal conditioner	Molecular Devices	CyberAmp 320
Software	Molecular Devices	pClamp 10.6