NIH R01 DK55619, NIH R21 CA139063

1. Animals
<ol>
	<li>All procedures were approved by the Institutional Animal Care and Use Committee at the University of Medicine and Dentistry of New Jersey.</li>
	<li>Group house male C57BL/6 mice on a 12 hr light/12 hr dark schedule and allow ad libitum access to water and food. Sacrifice at 4-5 months old.&#160;Euthanasia of the mice was performed using the surgical plane of anesthesia and a secondary form of euthanasia (i.e.&#160;penetrating incision into the chest cavity via the diaphragm). This is consistent with the AVMA Guidelines on Euthanasia.</li>
</ol>
2. Preparation of Perfusion Solution, Coverslips, Glass Pipettes, and Media
<ol>
	<li>Make 1L perfusion solution consisting of 2.5 mM KCl, 7 mM MgCl2, 1.25 mM NaH2PO4, 28 mM NaHCO3, 0.5 mM CaCl2, 7 mM glucose, 1 mM ascorbate, and 3 mM pyruvate in distilled dH2O. Adjust the osmolarity to ~300 mOsm using approximately 80 g/L sucrose. Oxygenate by bubbling with 95% O2 /5% CO2 and adjust the pH to 7.4. Each experiment will require approximately 200 ml&#160;of perfusion solution. Aliquots can be stored at -20 &#176;C for up to 2 months.</li>
	<li>Make 250 ml&#160;Neurobasal stock containing Neurobasal media without glucose, 2.5 mM glucose, and 100 U/ml&#160;penicillin streptomycin. Adjust the osmolarity of the Neurobasal stock to 280 mOsm using sucrose. Make 500 ml&#160;Hibernate stock containing Hibernate-A media without glucose, 2.5 mM glucose, and 100 U/ml&#160;penicillin streptomycin.</li>
	<li>Stir both stocks well and filter-sterilize using Stericup vacuum filter units. Be careful not to introduce glucose or other contaminants through the glassware or stir bars. Wrap the bottles in foil to protect the media from light and store at 4 &#176;C for up to 2 months.</li>
	<li>Flame the tips of 4 autoclaved glass pipettes (9 in) so that the tips are no longer sharp and the opening diameters are large (~0.9 mm), medium-large (~0.7 mm), medium (~0.5 mm), and small (~0.3 mm). The largest should be barely polished and the smallest should be about a third of that diameter.</li>
	<li>On the day before harvesting neurons, clean four 25 mm glass coverslips using 70% ethanol and passing over a Bunsen burner flame. Place each coverslip in a sterile 35 mm culture dish and add 2 ml&#160;sterile 1% poly-D-lysine in dH2O. Put the dishes in an incubator (37 &#176;C) overnight. The next day, wash with sterile dH2O 3x and allow coverslips to fully dry.</li>
	<li>Make 75 &#181;l aliquots of 1 M lactic acid and 200 &#181;l aliquots of GlutaMAX; store at -20 &#176;C. Fully thaw aliquots of lactic acid and GlutaMAX before use to ensure homogeneity.</li>
	<li>On the day of harvesting neurons, make 30 ml&#160;of fresh culture media consisting of Hibernate-A stock containing 1mM lactic acid, 0.5mM GlutaMAX, and 2% B27 without insulin. Vortex and adjust pH to 7.4 using 1 N NaOH.</li>
	<li>Put 10 ml&#160;of culture media in a 60 mm glass Petri dish on ice, put 2 ml&#160;in a 15 ml&#160;conical tube on ice and keep the remaining media at room temperature.</li>
	<li>On the day of harvesting neurons, make 25 ml&#160;of fresh growth media consisting of Neurobasal stock containing 1 mM lactic acid, 0.5 mM GlutaMAX, and 2% B27 without insulin. Vortex and adjust pH to 7.4 using 1 N NaOH. Allow the fresh growth media to equilibrate in an incubator (37 &#176;C, 5% CO2) for at least 30 min.</li>
</ol>
3. Cardiac Perfusion
<ol>
	<li>Fully thaw 200 ml&#160;perfusion solution and oxygenate with 95% O2/ 5% CO2 on ice for at least 15 min.</li>
	<li>Wash a vibratome blade with acetone, 70% ethanol, and dH2O. Position the blade in the vibratome.</li>
	<li>Rinse the vibratome cooling chamber and perfusion tubing with dH2O. Fill the cooling chamber (4 &#176;C) with perfusion solution and continue to oxygenate.</li>
	<li>Anesthetize a mouse with pentobarbital sodium 50 mg/ml, diluted 1:1 with sterile water, injected intraperitoneally. Verify the mouse is fully anesthetized by pinching the tail and observing no response.</li>
	<li>Pour cold perfusion solution into the perfusion reservoir and tubing just prior to dissection. Save 20-30 ml&#160;for brain dissection and continue to oxygenate on ice. Gravity flow from an elevated 60 ml&#160;syringe through tubing and a 20 G needle provides sufficient flow. Allow solution to run until air bubbles are washed out. Continue to oxygenate.</li>
	<li>Cut back skin above the ribcage of the mouse. Lift the ribcage and carefully make a horizontal cut through the diaphragm. Make two lateral cuts through the ribcage up toward the arms to expose the heart. Use forceps to hold back the ribcage.</li>
	<li>Make a small 2 mm cut in the right atrium to provide an exit point for blood to be washed out of the vascular system.</li>
	<li>Puncture the left ventricle with the perfusion needle, just enough to insert the needle opening. Hold the needle in place with one hand and start the flow of perfusion solution with the other hand. After 10-15 ml&#160;of perfusion solution, the blood should be washed out and the effluent will clear.</li>
</ol>
4. Brain Slicing and Dissection
<ol>
	<li>After cardiac perfusion, transfer oxygenated cold perfusion solution to a Petri dish on ice just prior to use.</li>
	<li>Quickly decapitate, remove the brain from the skull and place the brain into perfusion solution. To remove the brain: pull skin forward, make a midline cut in the skull toward the eye sockets, make 2 lateral cuts toward each eye, use forceps to pull back each side of the skull, make a coronal cut between the eye sockets, and use a spatula to gently coax the brain out into perfusion solution. Use scissors to cut the optic tracts if necessary. Do not to touch the hypothalamic area and do not pull the optic tracts because this puts too much pressure on the median eminence and underlying hypothalamic tissue.</li>
	<li>Use a razor blade and needle to laterally trim the brain tissue approximately 3 mm posterior and anterior to the hypothalamus while the tissue is submerged (bregma -3.52 mm). It is especially important for the anterior side cut to be level so that the brain will sit straight on the vibratome chuck.</li>
	<li>Remove the remaining brain tissue from solution and use filter paper to wick away solution from the mounted brain section.</li>
	<li>Place a dab of super glue on a vibratome chuck and spread the glue to the size of the brain. Put the brain tissue on top of the glue with the anterior side facing down and the hypothalamus facing the blade. Wick away excess glue and place the chuck into the vibratome cooling chamber. Use caution not to leave much glue as it will bubble up around the brain when placed into solution.</li>
	<li>With the vibratome set to a slow speed (level 2) and high amplitude (level 9), make 300-500 &#181;m slices until reaching the hypothalamic area. Now make thin 100 &#181;m slices cutting from the posterior to the anterior. The visual cues are as follows. Posterior to the hypothalamus the third ventricle will be very small (bregma -2.70 to -2.30 mm). At bregma -2.30 mm, the third ventricle is separated into dorsal and ventral sections on the coronal slice.</li>
	<li>Once sections of the third ventricle fuse, make two 500 &#181;m slices which will contain the correct region of the VMH (bregma -2.18 to -1.22 mm). Anterior to the VMH, the third ventricle no longer extends to the ventral floor of the brain (bregma -1.06 to -0.82)8.</li>
	<li>Transfer these slices to the Petri dish on ice containing culture media. Dissect the VMH (VMN + ARC) as shown in Figure 1. Use a pipette to gently transfer the VMH pieces in 2 ml&#160;of culture media on ice.</li>
</ol>
5. Dissociation and Culture
<ol>
	<li>Remove the VMH from ice and place at room temperature. Add 20 U/ml&#160;papain to 4 ml&#160;of culture media. Invert to mix and place in a 34 &#176;C water bath. Check and mix by inversion every minute until the digestion media is no longer cloudy. Filter (0.22 &#956;m) into a sterile flask and transfer the tissue to the digestion media.</li>
	<li>Digest the tissue by shaking at 100 rpm at 34 &#176;C for 30 min.</li>
	<li>Prepare 1 ml&#160;of media containing 8% bovine serum albumin (BSA) during the digestion. Dissolve 80 mg BSA in 1 ml&#160;of culture media and filter (0.22 &#956;m) into a conical tube.</li>
	<li>Wash the tissue by transferring to 5 ml&#160;of culture media in a conical tube and slowly inverting once.</li>
	<li>Add 30 &#181;l DNase enzyme to 6 ml&#160;of culture media and mix to make trituration media. Aspirate the wash media and add 3 ml&#160;of trituration media.</li>
	<li>Use the glass pipettes to triturate in the order of largest to smallest. Gently triturate 10x and then wait 4 min&#160;for larger pieces to settle. Use the second pipette to transfer the top 2 ml&#160;containing a dissociated cell suspension to a new conical tube. Add 2 ml&#160;of trituration media, triturate 10x and wait 3 min. Use the third pipette to transfer the top 2 ml&#160;to the cell suspension. Add 1 ml&#160;of trituration media, triturate 5x and wait 2 min. Use the fourth pipette to transfer the top 2ml&#160;to the cell suspension.</li>
	<li>Layer the dissociated cell suspension on top of the 8% BSA, being careful not to mix. Centrifuge at 1,000 rpm for 5 min.</li>
	<li>Place autoclaved 6 mm x 8 mm cloning cylinders in the center of each coverslip. Coverslips must be completely dry. Aspirate and resuspend the pellet in 440 &#181;l warm growth media. Fill cloning cylinders with the neuronal suspension and place in the incubator for 20 min.</li>
	<li>Remove the cloning cylinders and very gently wash away debris. Fill each dish with 2 ml&#160;of growth media. Allow cells to recover for at least 1 hr before any assays are performed.</li>
</ol>
6. Preparation for MPD Imaging
<ol>
	<li>Make recording solution consisting of 121 mM NaCl, 4.7 mM KCl, 2 mM CaCl2, 0.1 mM MgCl2, 1.2 mM MgSO4, 0.97 mM K2HPO4, 0.23 mM KH2PO4, 5 mM NaHCO3, and 25 mM HEPES. Adjust the pH to 7.4.</li>
	<li>Add glucose to make the desired low glucose test solution (0.1-2.0 mM glucose). Mix thoroughly and reserve 400 ml&#160;low glucose recording solution. Add glucose to the remaining 600 ml&#160;to make 2.5 mM glucose recording solution and mix thoroughly.</li>
	<li>Protect dye from light as much as possible. Add 4 ml of room temperature Buffer to a Blue MPD vial. Mix thoroughly and add 5 &#181;l dye per ml&#160;recording solution. Keep solution homogenous by mixing with pipette in between solutions. The dye contains fluorescent molecules, which enter cells with depolarization, and quencher molecules, which cannot cross the cell membrane. Aliquots may be stored at -20 &#176;C and used within 4 days. Do not freeze-thaw more than once.</li>
	<li>Incubate cells in 2 ml&#160;of 2.5 mM glucose recording solution plus dye at 34 &#176;C for 30 min. A dry heating block can be used. Protect from light.</li>
	<li>Prepare syringe pumps and perfusion system with 2.5 mM and 0.1 mM recording solutions plus dye. Tubing from 60 ml&#160;syringes should be connected to a manifold.</li>
	<li>After stopping any pump, wait for ~10 sec&#160;before switching solutions at the manifold to prevent pressure buildup in the syringe. Pressure changes alter fluid delivery to the closed chamber containing the neurons, causing changes in microscope focus during the experiment and distortion of images.</li>
	<li>The manifold output tubing should be connected to an in-line heater and then polyethylene tubing , which connects to a closed chamber. Bubbles should be cleared and the perfusion rate should be set to 0.5 ml/min. Protect from light.</li>
	<li>Transfer 25 mm coverslip with adherent neurons to the closed chamber, being careful not to allow coverslip to dry and not to introduce air bubbles. The slip is aligned in the grooves of the bottom piece, a couple drops of recording solution is placed along the sides, and the top piece is gently fitted to hold the coverslip in place.</li>
	<li>Use a dropper to fill the chamber with recording solution and then place a 18 mm coverslip on top. Gently fit white ring into the chamber to hold the smaller coverslip in place. Press down very slowly and lightly to prevent air bubbles from being introduced into the closed chamber.</li>
	<li>Start the syringe pump for the 2.5 mM glucose recording solution, press down on the white ring, and connect to the closed chamber. Slowly release pressure from white ring and connect to tubing leading to a waste container.</li>
</ol>
7. MPD Imaging
<ol>
	<li>Center the neurons using low bright field light. Try to minimize the amount of time the cells are being exposed to light.</li>
	<li>Allow the perfusion system to run for 10 min&#160;to allow for stabilization of temperature, focus, and dye equilibrium.</li>
	<li>While priming, open the MetaMorph program and take a bright field image (10X) of the neurons. Use the Auto-Focus function to take 3 stacks of fluorescent images (150 msec, Narrow Cy3 filter, 10X) and determine the focus within 5 &#956;m. At the end of the 10 min&#160;priming period, check the focus once more.</li>
	<li>Begin to record fluorescent images every 30 sec&#160;for 40 min. Establish a baseline at 2.5 mM glucose for 10 min, then decrease glucose for 15 min, and finally return to 2.5 mM glucose for 15 min. Solutions are changed by stopping a syringe pump, waiting 10 sec, turning a port on the manifold, and starting another syringe pump. Changes should occur ahead of desired time points based upon the lag between the manifold and cells.</li>
	<li>After recording all fluorescent images, take a bright field image of the neurons.</li>
</ol>
8. MPD Imaging Analysis
<ol>
	<li>Use MetaMorph to create oval regions around each cell in the bright field image taken at the end of the experiment. Regions should be slightly larger than each neuron and can be made 1 pixel outside of the cell&#39;s dark edge. Do not choose cells that are unhealthy, overlapping, or close to debris. Unhealthy cells often appear dark. Lastly, create 5 large oval regions in areas with no cells or debris for background measurements.</li>
	<li>Transfer the regions to all fluorescent images and inspect to verify that no movement/shift has occurred. Use the Measure function to record the fluorescence intensity of each region for all images in an Excel file. The use of Journals in Metamorph is advised.</li>
	<li>In Excel, create an average of the 5 background regions for each fluorescent image. This represents the average background value at each time point. For each image/time point, subtract the background from all region measurements. The background-subtracted fluorescence intensity will be referred to as Intensity hereafter.</li>
	<li>For each cell, calculate the average Intensity during minutes 2-8 of recording. This represents the Baseline of the cell in 2.5 mM glucose. A 2 minute buffer on both ends of each treatment is used to minimize noise.</li>
	<li>For each cell, calculate the percent change from baseline: (Intensity-Baseline)/Baseline</li>
	<li>Create a line graph with the percent change from baseline vs time.</li>
	<li>Calculate the average percent change from baseline during minutes 18-23. Calculate the average percent change from baseline during minutes 35-40. While image processing is not used, noise is reduced through the averaging of intensity within each cell region, background subtraction, and the averaging of region intensity over 5 min&#160;or longer.</li>
	<li>Control experiments in which 2.5 mM glucose recording solution is exchanged with 2.5 mM glucose recording solution must be performed to determine the noise threshold. Calculate the average percent change from baseline during minutes 18-23 for at least 6 dishes and 3 mice. Determine the mean and standard deviation of these values.</li>
	<li>Set the threshold for a positive depolarization response to the mean plus 2 standard deviations. The mean plus 2 standard deviations corresponds to a difference with p&#60;0.05. Our control experiments revealed a 10% change from baseline as the appropriate threshold value.</li>
	<li>For each cell, assess the reversible depolarization response based upon 2 criteria. First, the average percent change from baseline during minutes 18-23 must be greater than 10%, the threshold determined in the previous step. Second, the average percent change from baseline during minutes 35-40 must be less than half of the average percent change from baseline during minutes 18-23. The second criterion was chosen since it was found to be more rigorous than stimulation with glutamate or KCl. Unhealthy neurons often respond to stimulation with glutamate or KCl even though their responses to decreased glucose are not reversible.</li>
	<li>For each dish, calculate the % of depolarized neurons. This value is used to quantify the % of GI neurons among different treatment groups.</li>
</ol>

<ol>
	<li>Routh, V. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glucose-sensing+neurons:+are+they+physiologically+relevant?.">Glucose-sensing neurons: are they physiologically relevant?.</a> Physiol. Behav. 76, 403-413 (2002).</li><li>Canabal, D. D., Potian, J. G., Duran, R. G., McArdle, J. J., Routh, V. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hyperglycemia+impairs+glucose+and+insulin+regulation+of+nitric+oxide+production+in+glucose-inhibited+neurons+in+the+ventromedial+hypothalamus.">Hyperglycemia impairs glucose and insulin regulation of nitric oxide production in glucose-inhibited neurons in the ventromedial hypothalamus.</a> Am. J. Physiol. 293, 592-600 (2007).</li><li>Canabal, D. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glucose,+insulin,+and+leptin+signaling+pathways+modulate+nitric+oxide+synthesis+in+glucose-inhibited+neurons+in+the+ventromedial+hypothalamus.">Glucose, insulin, and leptin signaling pathways modulate nitric oxide synthesis in glucose-inhibited neurons in the ventromedial hypothalamus.</a> American journal of physiology. Reg. Integr. Comp. Physiol. 292, 1418-1428 (2007).</li><li>Murphy, B. A., Fakira, K. A., Song, Z., Beuve, A., Routh, V. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AMP-activated+protein+kinase+and+nitric+oxide+regulate+the+glucose+sensitivity+of+ventromedial+hypothalamic+glucose-inhibited+neurons.">AMP-activated protein kinase and nitric oxide regulate the glucose sensitivity of ventromedial hypothalamic glucose-inhibited neurons.</a> Am. J. Physiol. Cell Physiol. 297, C750-C758 (2009).</li><li>Murphy, B. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fasting+enhances+the+response+of+arcuate+neuropeptide+Y-glucose-inhibited+neurons+to+decreased+extracellular+glucose.">Fasting enhances the response of arcuate neuropeptide Y-glucose-inhibited neurons to decreased extracellular glucose.</a> Am. J. Physiol. Cell Physiol. 296, C746-C756 (2009).</li><li>Kang, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glucokinase+is+a+critical+regulator+of+ventromedial+hypothalamic+neuronal+glucosensing.">Glucokinase is a critical regulator of ventromedial hypothalamic neuronal glucosensing.</a> Diabetes. 55, 412-420 (2006).</li><li>Kang, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prior+hypoglycemia+enhances+glucose+responsiveness+in+some+ventromedial+hypothalamic+glucosensing+neurons.">Prior hypoglycemia enhances glucose responsiveness in some ventromedial hypothalamic glucosensing neurons.</a> Reg. Integr. Comp. Physiol. 294, R784-R792 (2008).</li><li>Paxinos, G., Franklin, K. B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Mouse Brain in Stereotaxic Coordinates. , (2004).</li><li>Paxinos, G., Watson, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Rat Brain in Stereotaxic Coordinates. , (1998).</li><li>Song, Z., Levin, B. E., McArdle, J. J., Bakhos, N., Routh, V. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Convergence+of+pre-+and+postsynaptic+influences+on+glucosensing+neurons+in+the+ventromedial+hypothalamic+nucleus.">Convergence of pre- and postsynaptic influences on glucosensing neurons in the ventromedial hypothalamic nucleus.</a> Diabetes. 50, 2673-2681 (2001).</li><li>Song, Z., Routh, V. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+effects+of+glucose+and+lactate+on+glucosensing+neurons+in+the+ventromedial+hypothalamic+nucleus.">Differential effects of glucose and lactate on glucosensing neurons in the ventromedial hypothalamic nucleus.</a> Diabetes. 54, 15-22 (2005).</li></ol>

We have nothing to disclose.

The key to being able to study activity of neurons from adult mice is the ability to dissociate healthy neurons. Dissociation of hypothalamic neurons from adult mice is more difficult at several key steps in the protocol compared to neurons from juvenile mice. We have overcome this problem in a number of ways. Making thick 500 &#181;m brain slices minimizes mechanical damage to neurons compared to the usual 250-350 &#181;m slices used for brain tissue from younger mice. However, thicker slices require greater attention t...

The brain regulates energy homeostasis through the neuroendocrine and autonomic nervous systems. The ventromedial hypothalamus (VMH), comprised of the ventromedial nucleus (VMN) and the arcuate nucleus (ARC), is important for the central regulation of energy homeostasis. Specialized glucose sensing neurons, within the VMH, link neuronal activity and peripheral glucose homeostasis1. There are two types of glucose sensing neurons; glucose excited (GE) neurons increase while glucose inhibited (GI) neurons decrease their activity as extracellular glucose increases. VMH glucose sensing neurons are generally studied using electrophysiology or calcium/membrane potential sensitive dye imaging.
The electrophysiological patch clamp technique is considered to be the gold standard in the study of ex vivo neuronal activity. In this technique, a glass micropipette electrode is attached to the cell membrane via a high resistance (G&#937;) seal. Patch clamp electrodes allow real time recording of action potential frequency (current clamp) or ion conductance (voltage clamp) changes within a single neuron. While the patch clamp technique provides detailed information regarding changes in specific ion channel conductances, a major drawback is that only one neuron may observed at a time. It takes approximately 30-45 min&#160;of recording to verify that one is recording from a glucose sensing neuron before even beginning a specific experimental treatment. Moreover, GI and GE neurons comprise &#60;20% of the total VMH neuronal population. Compounding this issue is the lack, in many cases, of an identifying cellular marker for these neurons. Thus, it is clear that despite providing valuable electrical information that other techniques cannot, patch clamp analysis is laborious, time consuming and low yield.
The use of fluorescence imaging of dissociated VMH neurons allows for the study of hundreds of neurons simultaneously. Calcium sensitive dyes can be used to measure intracellular calcium changes, which indirectly correlate to changes in neuronal activity. Membrane potential sensitive dyes are used to monitor membrane potential changes. Measuring cellular membrane potential is a more direct index of neuronal activity compared to changes in intracellular calcium levels. Furthermore, membrane potential dye (MPD) imaging potentially detects smaller changes in membrane potential where action potential firing is not altered and intracellular calcium levels might not change. Both of these fluorescence imaging techniques have been used to study VMH glucose sensing neurons from juvenile mice2-7. While results are less detailed than those obtained with patch clamp electrophysiology, the strength of imaging experiments is that they simultaneously evaluate a large population of cells which inevitably include a significant number of glucose sensing neurons. MPD imaging is particularly useful for studying GI neurons which are more uniformly localized throughout the entire VMH; thus providing an adequate population to study in the dissociated VMH (~15% GI). In contrast, while GE neurons are densely localized to the ventrolateral-VMN and cell poor region between the ARC and VMN, they do not represent a significant number of neurons within the VMH (&#60;1% GE). Moreover, by studying isolated neurons, astrocytic and presynaptic effects are eliminated. This can be an advantage in studying first order neuron effects, as well as a disadvantage since physiological connections and processes are lost.
A limiting factor in both patch clamp electrophysiology and MPD/calcium dye imaging is the need to use younger animals (e.g.&#160;mice or rats &#60;8 weeks of age). This is predominantly due to increased connective tissue in combination with decreased neuronal viability that occurs with age. In brain-slice electrophysiology studies, increased connective tissue makes it more difficult to visualize the neurons. Increased connective tissue also makes it harder to dissociate a large number of healthy neurons for imaging studies. Furthermore, neurons from younger animals survive longer during either patch clamp recording or imaging. However, the use of young mice can be a major limitation. Neuronal activity and/or responses to neurotransmitters or circulating nutrients change with age. For example, since energy balance is closely tied to reproductive status, the hypothalamic neurons regulating energy balance may respond differently in pre- vs postpubescent animals. Additionally, many diseases require long term treatment or do not manifest until adulthood. Prime examples of such diseases are dietary obesity or Type 2 Diabetes Mellitus. Since glucose sensing neurons are believed to play a role in these diseases we developed a methodology for successfully culturing healthy adult VMH neurons for use in MPD imaging experiments.

<table border="0" cellpadding="0" cellspacing="0" width="947">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Neurobasal-A Medium (Custom)</td>
			<td style="width:115px;">Invitrogen</td>
			<td style="width:180px;">0050128DJ</td>
			<td style="width:427px;">custom made glucose free</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Hibernate-A Medium (Custom)</td>
			<td style="width:115px;">BrainBits</td>
			<td style="width:180px;"></td>
			<td>custom made glucose free</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:225px;">Penicillin streptomycin (20,000 U/ml)</td>
			<td style="width:115px;">Invitrogen</td>
			<td style="width:180px;">15140</td>
			<td>other vendors acceptable</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:225px;">Stericup vacuum filter units (0.22 &#956;m)</td>
			<td style="width:115px;">Millipore</td>
			<td style="width:180px;"></td>
			<td>other vendors acceptable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">25 mm Glass coverslips</td>
			<td style="width:115px;">Warner</td>
			<td style="width:180px;">#1 25mm round</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">18 mm Glass coverslips</td>
			<td style="width:115px;">Warner</td>
			<td style="width:180px;">#1 18mm round</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">GlutaMAX</td>
			<td style="width:115px;">Invitrogen</td>
			<td style="width:180px;">35050</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">B27 minus insulin (50x)</td>
			<td style="width:115px;">Invitrogen</td>
			<td style="width:180px;">0050129SA</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Razor blade</td>
			<td style="width:115px;">VWR</td>
			<td style="width:180px;">55411</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:225px;">Vibratome &#38; cooling chamber</td>
			<td style="width:115px;">Vibratome</td>
			<td style="width:180px;">Series 1000 Sectioning system</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Vibratome blades</td>
			<td style="width:115px;">Polysciences</td>
			<td style="width:180px;">22370</td>
			<td>injector or double edge blades from other vendors acceptable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Papain, suspension</td>
			<td style="width:115px;">Worthington</td>
			<td style="width:180px;">LS003124</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">BSA, suitable for cell culture</td>
			<td style="width:115px;">Sigma</td>
			<td style="width:180px;"></td>
			<td>other vendor acceptable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">DNAse, for cell culture</td>
			<td style="width:115px;">Invitrogen</td>
			<td style="width:180px;"></td>
			<td>other vendor acceptable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">cloning cylinders, 6 mm x 8 mm</td>
			<td style="width:115px;">Bellco Glass</td>
			<td style="width:180px;">2090-00608</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:225px;">Membrane Potential Dye (blue)</td>
			<td style="width:115px;">Molecular Devices</td>
			<td style="width:180px;">R8042</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">In-line heater</td>
			<td style="width:115px;">Warner</td>
			<td style="width:180px;">SF-28</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Syringe pumps</td>
			<td style="width:115px;">WPI</td>
			<td style="width:180px;">sp100i</td>
			<td>other vendor acceptable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Closed chamber</td>
			<td style="width:115px;">Warner</td>
			<td style="width:180px;">RC-43C</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Polyethylene tubing</td>
			<td style="width:115px;">Warner</td>
			<td style="width:180px;">PE-90</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:225px;">Metamorph</td>
			<td style="width:115px;">Molecular Devices</td>
			<td style="width:180px;"></td>
			<td>alternate image analysis software acceptable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Microscope</td>
			<td style="width:115px;">Olympus</td>
			<td style="width:180px;">BX61 WI</td>
			<td>
			used with 10X&#160;objective
			</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Camera</td>
			<td style="width:115px;">Photometrics</td>
			<td style="width:180px;">Cool Snap HQ</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:225px;">Narrow Cy3 Filter Set</td>
			<td style="width:115px;">Chroma</td>
			<td style="width:180px;">41007a</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:225px;">Illumination System</td>
			<td style="width:115px;">Sutter Instruments</td>
			<td style="width:180px;">Lambda DG-4</td>
			<td></td>
		</tr>
	</tbody>
</table>

membrane potential dye imaging of ventromedial hypothalamus neurons from adult mice to study glucose sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The precise dissection of the VMH away from other hypothalamic areas is important to obtain consistent results. The inclusion of other areas could dilute the VMH neuronal population, changing the % of depolarized neurons calculated. Furthermore, glucose sensing neurons have been identified in other hypothalamic regions, such as the lateral hypothalamus, which may differ functionally and mechanistically from VMH glucose sensing neurons. Figure 1 illustrates the correct anatomical locations for proper diss...

Watch this Scientific Journal Video about Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing at JoVE.com

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

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Neuroscience

13.6K Views.  Rutgers New Jersey Medical School. The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Video: Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Isolation and Culture of Hippocampal Neurons from Prenatal Mice

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual neurons, researchers are able to analyze properties related to cellular trafficking, cellular structure and individual protein localization using a variety of biochemical techniques. Results from such experiments are critical for testing theories addressing the neural basis of memory and learning. However, unambiguous results from these forms of experiments are predicated on the ability to grow neuronal cultures with minimum contamination by other brain cell types. In this protocol, we use specific media designed for neuron growth and careful dissection of embryonic hippocampal tissue to optimize growth of healthy neurons while minimizing contaminating cell types (i.e. astrocytes). Embryonic mouse hippocampal tissue can be more difficult to isolate than similar rodent tissue due to the size of the sample for dissection. We show detailed dissection techniques of hippocampus from embryonic day 19 (E19) mouse pups. Once hippocampal tissue is isolated, gentle dissociation of neuronal cells is achieved with a dilute concentration of trypsin and mechanical disruption designed to separate cells from connective tissue while providing minimum damage to individual cells. A detailed description of how to prepare pipettes to be used in the disruption is included. Optimal plating densities are provided for immuno-fluorescence protocols to maximize successful cell culture. The protocol provides a fast (approximately 2 hr) and efficient technique for the culture of neuronal cells from mouse hippocampal tissue.

We provide a protocol for the culture of highly purified hippocampal neurons from prenatal mouse brains without the use of a feeder glial cell layer.

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual ...

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion channel, and transporter cell surface presentation on a minutes time scale.&#160;There is a broad diversity of mechanisms that control endocytic trafficking of individual proteins. Studies investigating the molecular underpinnings of trafficking have primarily relied upon surface biotinylation to quantitatively measure changes in membrane protein surface expression in response to exogenous stimuli and gene manipulation.&#160;However, this approach has been mainly limited to cultured cells, which may not faithfully reflect the physiologically relevant mechanisms at play in adult neurons.&#160;Moreover, cultured cell approaches may underestimate region-specific differences in trafficking mechanisms.&#160;Here, we describe an approach that extends cell surface biotinylation to the acute brain slice preparation.&#160;We demonstrate that this method provides a high-fidelity approach to measure rapid changes in membrane protein surface levels in adult neurons.&#160;This approach is likely to have broad utility in the field of neuronal endocytic trafficking.

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Neuronal membrane trafficking dynamically controls plasma membrane protein availability and significantly impacts neurotransmission.&#160;To date, it has been challenging to measure neuronal endocytic trafficking in adult neurons.&#160;Here, we describe a highly effective, quantitative method to measure rapid changes in surface protein expression ex vivo in acute brain slices.

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion ...

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with enhanced [Ca2+]i signals. It has been proposed that astrocytes in turn regulate neurons and blood vessels through calcium-dependent mechanisms, such as the release of signaling molecules. However, [Ca2+]i imaging in entire astrocytes has only recently become feasible with genetically encoded calcium indicators (GECIs) such as the GCaMP series. The use of GECIs in astrocytes now provides opportunities to study astrocyte [Ca2+]i signals in detail within model microcircuits such as the striatum, which is the largest nucleus of the basal ganglia. In the present report, detailed surgical methods to express GECIs in astrocytes in vivo, and confocal imaging approaches to record [Ca2+]i signals in striatal astrocytes in situ, are described. We highlight precautions, necessary controls and tests to determine if GECI expression is selective for astrocytes and to evaluate signs of overt astrocyte reactivity. We also describe brain slice and imaging conditions in detail that permit reliable [Ca2+]i imaging in striatal astrocytes in situ. The use of these approaches revealed the entire territories of single striatal astrocytes and spontaneous [Ca2+]i signals within their somata, branches and branchlets. The further use and expansion of these approaches in the striatum will allow for the detailed study of astrocyte [Ca2+]i signals in the striatal microcircuitry.

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ.