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 electrophysiology: (1) inverting the mucosa rather than peeling it, (2) adding a small amount of BSA to the digestion media and FBS to the culture media, (3) lowering the Collagenase XI concentration, (4) adjusting the duration and intensity of mechanical agitation to increase the number and health of single cells, (5) plating the extracellular matrix at a low concentration, and (6) culturing with ROCK inhibitor.
Similarly, the following adaptations were critical for obtaining whole cells while patch clamping: (1) thoroughly rinsing the cultures with serum-free extracellular solution, (2) letting the EC cells stand in the extracellular solution for at least 4 h, (3) re-purposing any spent electrodes-like those from failed seals-to clear away debris en route to the next EC cell, (4) adding 10 &#181;M Gd3+ to the extracellular solution to block leak currents (if present) and to facilitate adherence of mammalian cell suspensions to glass. We caution, however, that Gd3+ could block voltage-gated Ca2+ and/or mechanosensitive channels expressed in EC cells.
Despite having carefully optimized the culturing conditions, we found a few limitations: EC cells need 24 h to fully attach yet expire after ~72 h in culture; therefore, they have a 48-hour window when they would be suitable for electrophysiological experiments. Because clumped cells often introduce capacitance transients that cannot be compensated from recordings, generation of single EC cells are critical. However, by separating EC cells from the epithelium, the cells then live in a non-native environment (i.e., high glucose media with fetal bovine serum and ROCK inhibitor atop the extracellular matrix as an artificial basement membrane) and could respond to stimuli not entirely representative of EC cell function in vivo. Despite these limitations, the primary culturing of EC cells still provides insight into their physiology.
EC cells play a critical role in maintaining a well-functioning GI tract and body, and as such, there have been many methods developed to gain an understanding about EC cell electrophysiology. Enteroendocrine and EC cell electrophysiology was investigated in immortalized cell models5,8,13, primary EC cell cultures from guinea pig11, and mouse3,4, Our method utilizes cell quality-preserving methods to create a culture that has been minimally processed and reflects the native EC cells. We use a transgenic mouse model, Tph1-CFP, that shows a cyan fluorescence in the cytoplasm when Tph1 is present9. This method can also be used for other reporter or lineage-traced enteroendocrine cell types. Our method also accomplishes the isolation of single EC cells in minimal time and enzymatic treatment, as the cells are only exposed to a gentle Collagenase XI at a minimal yet effective concentration of 0.1 mg/mL (jejunum) and 0.6 mg/mL (colon) for less than one hour. The result of these modifications is a culture that has been optimized for electrophysiological studies of the EC cells.
We have successfully used this method to study electrophysiology and serotonin release in murine EC cells3. The ability to successfully isolate, identify, and culture single EC cells facilitates important future directions in studies on EC cell physiology. This method can be used to further determine critical pathways and channels involved in the EC cell exocytosis of serotonin and other EC cell-specific transmitters. The method could also be modified to isolate and characterize the physiology of the human EC cell along with the use of acridine orange for EC cell identification13.
As with most scientific protocol development, communication and unbiased observations were critical in the troubleshooting of this protocol. When we tested new culture methods for the survival of our cultures, it was helpful to count the number of fluorescent cells as a measure of viability, estimate cell densities in each dish, and collect representative pictures and videos of the cultures across many time points. After the cells were able to survive past 24 h in culture, we worked on modifying our cultures for electrophysiology experiments. Along with collecting images and detailed observations, it was important to maintain clear communication between lab personnel regarding adjustments to mechanical digestion and supplement and/or substrate concentrations, so that healthy EC cells were ready for electrophysiological characterization.

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.

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			<td height="20" style="height:20px;width:276px;">High Glucose DMEM</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">D6546</td>
			<td style="width:564px;">Store at 4&#730; C for no longer than 1 month</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Fetal Bovine Serum (FBS), Heat Inactivated</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">F4135</td>
			<td>Freeze in 25 mL aliquots&#160; and store long term at -20&#730; C</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">L-Glutamine</td>
			<td style="width:179px;">Life Technologies</td>
			<td style="width:175px;">25030-081</td>
			<td>Freeze in 5 mL aliquots and store long term at -20&#730; C</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Penicillin/ Streptomycin</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">P0781</td>
			<td>Freeze in 5 mL aliquots and store long term at -20&#730; C</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Bovine Serum Albumin (BSA)</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">A2153</td>
			<td>Store at 4&#730; C for long term storage</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Collagenase XI</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">C9407</td>
			<td>Store at -20&#730; C in aliquots of 100mg, avoid freeze thaw and avoid lot to lot variation</td>
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			<td height="20" style="height:20px;width:276px;">Phosphate Buffered Saline (PBS), w/ Ca and Mg</td>
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			<td>Resuspend at 5mM and store in small aliquots at -80&#730; C</td>
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			<td height="20" style="height:20px;width:276px;">100x15mm culture dish</td>
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			<td>351029</td>
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			<td style="width:179px;">Fisherbrand</td>
			<td style="width:175px;">13-711-7M</td>
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			<td style="width:175px;">357551</td>
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			<td height="20" style="height:20px;width:276px;">50 mL Conical</td>
			<td style="width:179px;">Corning (Falcon)&#160;</td>
			<td style="width:175px;">352098</td>
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			<td height="20" style="height:20px;width:276px;">15 mL Conical</td>
			<td style="width:179px;">Corning (Falcon)&#160;</td>
			<td style="width:175px;">352097</td>
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			<td height="20" style="height:20px;width:276px;">1000 mL Barrier Pipet Tips</td>
			<td style="width:179px;">Thermo Scientific (Art)</td>
			<td style="width:175px;">2079E</td>
			<td>Keep sterile</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Matrigel, Growth Factor Reduced</td>
			<td style="width:179px;">Corning</td>
			<td style="width:175px;">354230</td>
			<td>Store at 150uL aliquots at -20&#730; C. Thaw on ice and keep cold until ready for plating</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">MatTek Glass Bottom Dishes</td>
			<td style="width:179px;">MatTek Corporation</td>
			<td style="width:175px;">P35G-1.5-14-C</td>
			<td>Keep sterile</td>
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		<tr height="35">
			<td height="35" style="height:35px;width:276px;">Micro-dissection Forceps - Very Fine, Extra-Long Points</td>
			<td style="width:179px;">Fine Science Tools</td>
			<td>SB12630M</td>
			<td></td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Surgical Scissors-Sharp</td>
			<td style="width:179px;">Nasco</td>
			<td>14002-14</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Micro-Dissection Scissors</td>
			<td style="width:179px;">Nasco</td>
			<td style="width:175px;">SB46568M</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Monoject Hypo Needle, 21G x 1&#34; A, 100/BX</td>
			<td style="width:179px;">Covidien</td>
			<td style="width:175px;">8881250172</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:276px;">&#160;Tg(Tph1-CFP)1Able/J Mice</td>
			<td style="width:179px;">Jackson</td>
			<td>Stock # 028366</td>
			<td>Use male or female mice between the ages of 4-7 weeks</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:276px;">Microfil nonmetallic syringe needles (34 gauge, 67 mm)</td>
			<td style="width:179px;">World Precision Instruments</td>
			<td>MF34G-5</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Parafilm &#34;M&#34; Laboratory Film</td>
			<td style="width:179px;">Pechiney Plastic Packaging</td>
			<td></td>
			<td></td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">R6101</td>
			<td style="width:179px;">Dow Corning</td>
			<td></td>
			<td></td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">6-mL syringe with regular luer tip</td>
			<td style="width:179px;">Monoject</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:276px;">3-mL syringe with regular luer tip</td>
			<td style="width:179px;">Monoject</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:276px;">Electrophysiology Equipment</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Amplifier</td>
			<td style="width:179px;">Molecular Devices</td>
			<td>Axopatch 200B</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Data acquisition system (daq)</td>
			<td style="width:179px;">Molecular Devices</td>
			<td>Digidata 1440A</td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Signal conditioner</td>
			<td style="width:179px;">Molecular Devices</td>
			<td>CyberAmp 320</td>
			<td></td>
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		<tr height="20">
			<td height="20" style="height:20px;width:276px;">Software</td>
			<td style="width:179px;">Molecular Devices</td>
			<td>pClamp 10.6</td>
			<td></td>
		</tr>
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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 electrophysiology consisted of single cells and small clumps. Healthy EC cells were readily identified by bright CFP fluorescence and had still adhered to the dish after vigorous washing (Figure 1B).
Electrophysiology: 
EC whole cells were obtained on just 15 &#177;4% of attempts from colon cultures without ROCK inhibitor (29 whole cells out of 182 attempts from 22 cultures) but 30 &#177;7% of attempts from colon cultures with ROCK inhibitor (27 whole cells out of 102 attempts from 50 cultures) (Figure 4A; p &#60;0.05 by a non-parametric two-tailed t-test, colon cultures without vs. with ROCK inhibitor). Similarly, jejunum EC cells without ROCK inhibitor did not survive long enough for electrophysiology, while EC whole cells were obtained on 41 &#177;3% of attempts from jejunum culture with ROCK inhibitor (186 whole cells out of 447 attempts from 50 cultures) (Figure 4A; p &#62;0.05 by a non-parametric two-tailed t-test, jejunum vs. colon with ROCK inhibitor).
By whole cell electrophysiology, Na+ currents were recorded in voltage-clamp mode and action potentials (AP) in current-clamp mode. Na+ currents were recorded from 81.3 &#177;4.0% of TPH1-CFP+ cells from jejunum or 64.1 &#177;9.2% from colon3. Of 59 EC cells from jejunum cultures confirmed by voltage-clamp to have Na+ peak currents larger than -25 pA, 19 EC cells in current-clamp mode fired spontaneous AP (&#34;spontaneous&#34;), 29 EC cells fired AP only when elicited by a step protocol in current-clamp mode (&#34;elicited-only&#34;), and 11 EC cells did not fire AP spontaneously or by step protocol (&#34;non-firing&#34;) (Figure 4B). Spontaneous cells had an average peak Na+ current density (IPEAK) of -108.0 &#177;19.3 pA/pF, elicited-only cells had an IPEAK of -60.2 &#177;9.2 pA/pF, while non-firing cells had an IPEAK of -43.3 &#177;14 pA/pF (Figure 4B; p &#60; 0.05, spontaneous vs. elicited-only or non-firing groups; p &#62;0.05 elicited-only vs. non-firing groups). The difference in current densities were not a reflection of whole cell capacitance, as the spontaneous (3.3 &#177;0.2 pF), elicited-only (2.9 &#177;0.2 pF), and non-firing (3.1 &#177;0.3 pF) cells had similar whole cell capacitances (P&#160;&#62;0.05). EC cell Na+ currents and elicited AP were sensitive to [Na+]o concentration and NaV1.3 inhibition3.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58112/58112fig1.jpg" /> 
Figure 1: Representative range of culture results. DIC with epifluorescence images of a primary epithelial jejunum culture from a Tph1-CFP mouse plated on a glass bottom dish and cultured for 24 h. These two images demonstrate a range of culture outcomes from this protocol. CFP fluorescent cells (white) are enterochromaffin (EC) cells. (A)&#160;representative image from a culture that is poor quality for electrophysiology. The culture consists of large clumps, damaged cell membranes, low fluorescence signal, and floating groups of dead cells. (B)&#160;representative image from a culture that has been optimized for electrophysiology. Culture consists mainly of single, healthy cells and small clumps that have adhered to the plate. EC cells in this culture maintain a strong fluorescence signal. Scale bar = 100 &#181;m; panels are scaled 1:1. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58112/58112fig1large.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/58112/58112fig2.jpg" /> 
Figure 2: Preparation of cell cultures for electrophysiology. A strategy for efficient and thorough rinsing of EC cell cultures with serum-free extracellular solution is critical for reducing extracellular matrix and serum which would impede seal formation. (A)&#160;materials. Clockwise, from top left: Stage, two 5 mm wide strips of wax film, two 200 mg pieces of modeling clay, plastic transfer pipette, #5 forceps, elliptical recording chamber, 35 mm culture dish with glass bottom. (B)&#160;adhere wax film to the inner diameter of the stage, lower the elliptical chamber into the culture dish, and flatten the clay into rectangular strips. (C)&#160;gently press the culture dish into the wax film-lined stage and immobilize the recording chamber by stretching the modeling clay over the edge of the dish, from the chamber to the stage. Rinse EC cells with several volumes of extracellular solution. (D)&#160;return the stage to the mounting brackets above the inverted microscope. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58112/58112fig2large.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/58112/58112fig3.jpg" /> 
Figure 3: Representative electrophysiology results.&#160;(A)&#160;whole cell Na+ currents, recorded by a two-step voltage-clamp protocol (inset). 50 ms depolarizations from -120 mV activate channels during step 1 (&#9679;), while a further depolarization to 0 mV during step 2 (&#9675;) activates channels that had not yet inactivated (i.e., are still available to open). Red trace, current elicited by voltage steps from -120 to -40 to 0 mV. (B)&#160;current-voltage relationship of peak Na+ currents from panel A, step 1 (&#9679;, steady-state activation) or step 2 (&#9675;, steady-state inactivation). The intersection of the steady-state activation and inactivation curves reveals a small window current at -40 mV (red symbols). (C)&#160;action potentials elicited by depolarizing current stimuli. An action potential can fire past 0 mV (gray line) when the membrane potential reaches the voltage of Na+ window current (red line, -40 mV). Inset, current-clamp protocol, in which 0 to +8 pA was injected for 50-ms depolarizations from a baseline of -3 pA. (D)&#160;action potentials fire from atop spontaneous events that have depolarized to -40 mV. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58112/58112fig3large.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/58112/58112fig4.jpg" /> 
Figure 4: Throughput of whole cell electrophysiology.&#160;(A)&#160;mean percentage of whole cells per attempt for n days of work in the absence (-) or presence (+) of ROCK inhibitor (10 &#181;M). EC cells from jejunum did not survive 24 h in culture without ROCK inhibitor. (B)&#160;Na+ current densities measured in voltage-clamp from n EC cells, in which action potentials did not fire (Non-firing), fired only when elicited by an episodic current-clamp protocol (Elicited-only), or fired spontaneously in gap-free current-clamp mode (Spontaneous) (error bars are SEM). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58112/58112fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

whole-cell-electrophysiology-primary-cultured-murine-enterochromaffin

Nanyang Technological University

Research

JoVE Journal

Biology

10.2K Views.  Mayo Clinic. 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.

Video: Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Derivation of Hematopoietic Stem Cells from Murine Embryonic Stem Cells

A stem cell is defined as a cell with the capacity to both self-renew and generate multiple differentiated progeny.  Embryonic stem cells (ESC) are derived from the blastocyst of the early embryo and are pluripotent in differentiative ability.  Their vast differentiative potential has made them the focus of much research centered on deducing how to coax them to generate clinically useful cell types.  The successful derivation of hematopoietic stem cells (HSC) from mouse ESC has recently been accomplished and can be visualized in this video protocol.  HSC, arguably the most clinically exploited cell population, are used to treat a myriad of hematopoietic malignancies and disorders.  However, many patients that might benefit from HSC therapy lack access to suitable donors.  ESC could provide an alternative source of HSC for these patients.  The following protocol establishes a baseline from which ESC-HSC can be studied and inform efforts to isolate HSC from human ESC.  In this protocol, ESC are differentiated as embryoid bodies (EBs) for 6 days in commercially available serum pre-screened for optimal hematopoietic differentiation.  EBs are then dissociated and infected with retroviral HoxB4.  Infected EB-derived cells are plated on OP9 stroma, a bone marrow stromal cell line derived from the calvaria of  M-CSF-/- mice, and co-cultured in the presence of hematopoiesis promoting cytokines for ten days.  During this co-culture, the infected cells expand greatly, resulting in the generation a heterogeneous pool of 100s of millions of cells.  These cells can then be used to rescue and reconstitute lethally irradiated mice.

This protocol details the derivation of transplantable hematopoietic stem cells from mouse embryonic stem cells (ESC) and their subsequent injection into lethally irradiated recipient mice. Briefly, ESC are differentiated as embryoid bodies, which are then infected with retroviral HoxB4 and co-cultured with OP9 stromal cells and hematopoietic cytokines.

A stem cell is defined as a cell with the capacity to both self-renew and generate multiple differentiated progeny.  Embryonic stem cells (ESC) are ...

Counting and Determining the Viability of Cultured Cells

Determining the number of cells in culture is important in standardization of culture conditions and in performing accurate quantitation experiments. A hemacytometer is a thick glass slide with a central area designed as a counting chamber. Cell suspension is applied to a defined area and counted so cell density can be calculated. 

Determining the number of cells in culture is important in standardization of culture conditions and in performing accurate quantitation experiments. In this video, we demonstrate how cells are counted using a hemacytometer.

Determining the number of cells in culture is important in standardization of culture conditions and in performing accurate quantitation experiments. A ...

The Preparation of Primary Hematopoietic Cell Cultures From Murine Bone Marrow for Electroporation

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser MXcell  electroporation system and Gene Pulser  electroporation buffer were specifically developed to transfect nucleic acids into mammalian cells and difficult-to-transfect cells, such as primary and stem cells.This video demonstrates how to establish primary hematopoietic cell cultures from murine bone marrow, and then prepare them for electroporation in the MXcell system. We begin by isolating femur and tibia. Bone marrow from both femur and tibia are then harvested and cultures are established. Cultured bone marrow cells are then transfected and analyzed.

This procedure describes how to establish primary hematopoietic cell cultures from murine bone marrow and is followed by transfection using the Gene Pulser MXCell electroporation system.

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser ...

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Spinal Cord Electrophysiology

The neonatal mouse spinal cord is a model for studying the development of neural circuitries and locomotor movement. We demonstrate the spinal cord dissection and preparation of recording bath artificial cerebrospinal fluid used for locomotor studies. Once dissected, the spinal cord ventral nerve roots can be attached to a recording electrode to record the electrophysiologic signals of the central pattern generating circuitry within the lumbar cord.

A demonstration of the isolation of neonatal mouse spinal cord for electrophysiologic studies.

The neonatal mouse spinal cord is a model for studying the development of neural circuitries and locomotor movement. We demonstrate the spinal cord ...

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 ...

Adenovirus-mediated Genetic Removal of Signaling Molecules in Cultured Primary Mouse Embryonic Fibroblasts

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction analysis. One particular method to achieve this conditional deletion is via the use of the Cre-loxP system. This method involves flanking the gene of interest with loxP sites, which are specific recognition sequences for the Cre recombinase protein. Exposure of the so-called floxed (flanked by loxP site) DNA to this enzyme results in a Cre-mediated recombination event at the loxP sites, and subsequent excision of the intervening gene3. Several different methods exist to administer Cre recombinase to the site of interest. In this video, we demonstrate the use of an adenovirus containing the Cre recombinase gene to infect primary mouse embryonic fibroblasts (MEFs) obtained from embryos containing a floxed Rac1 allele1. Our rationale for selecting Rac1 MEFs for our experiments is that clear morphological changes can be seen upon deletion of Rac1, due to alterations in the actin cytoskeleton2,5. 72 hours following viral transduction and Cre expression, cells were stained using the actin dye phalloidin and imaged using confocal laser scanning microscopy. It was observed that MEFs which had been exposed to the adeno-Cre virus appeared contracted and elongated in morphology compared to uninfected cells, consistent with previous reports2,5. The adenovirus method of Cre recombinase delivery is advantageous as the adeno-Cre virus is easily available, and gene deletion via Cre in nearly 100% of the cells can be achieved with optimized adenoviral infection.

In this video we use an adenovirus carrying the Cre recombinase gene to infect primary mouse embryonic fibroblasts carrying a floxed Rac1 allele.

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction ...

Chromosome Preparation From Cultured Cells

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as fluorescence in situ hybridization (FISH), this assay has the powerful ability to analyze individual cells for aberrations that involve gains or losses of portions of the genome&#160;and rearrangements involving one or more chromosomes. In humans, chromosome abnormalities occur in approximately 1 per 160 live births1,2, 60-80% of all miscarriages3,4, 10% of stillbirths2,5, 13% of individuals with congenital heart disease6, 3-6% of infertility cases2, and in many patients with developmental delay&#160;and birth defects7. Cytogenetic analysis of malignancy is routinely used by researchers and clinicians, as observations of clonal chromosomal abnormalities have been shown to have both diagnostic and prognostic significance8,9.&#160; Chromosome isolation is invaluable for gene therapy and stem cell research of organisms including nonhuman primates and rodents10-13.
Chromosomes can be isolated from cells of live tissues, including blood lymphocytes, skin fibroblasts, amniocytes, placenta, bone marrow, and tumor specimens. Chromosomes are analyzed at the metaphase stage of mitosis, when they are most condensed and therefore more clearly visible. The first step of the chromosome isolation technique involves the disruption of the spindle fibers by incubation with Colcemid, to prevent the cells from proceeding to the subsequent anaphase stage. The cells are then treated with a hypotonic solution and preserved in their swollen state with Carnoy&#39;s fixative. The cells are then dropped on to slides and can then be utilized for a variety of procedures. G-banding involves trypsin treatment followed by staining with Giemsa to create characteristic light and dark bands. The same procedure to isolate chromosomes can be used for the preparation of cells for procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY)14,15.

Chromosomes can be isolated from live cells such as lymphocytes or skin fibroblasts, and from organisms including humans or mice. These chromosome preparations can be further utilized for routine G-banding and molecular cytogenetic procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY).

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as ...

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...