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Michael AC, Borland LM, editors. Electrochemical Methods for Neuroscience. Boca Raton (FL): CRC Press/Taylor & Francis; 2007.

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Electrochemical Methods for Neuroscience.

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Chapter 16Amperometric Detection of Dopamine Exocytosis from Synaptic Terminals

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Introduction to Dopamine Neurotransmission

Dopamine (DA) was discovered as the precursor of the neurotransmitter norepinephrine in 1957 (Montagu 1957). Carlsson demonstrated that DA levels in the striatum were much higher than in the rest of the brain despite its low level of norepinephrine, suggesting that DA was not merely a norepinephrine precursor but was itself a neurotransmitter (1959) (Carlsson 1959). Subsequently, DA’s role as a neurotransmitter has been confirmed and populations of DA neurons have been identified in various regions of the central nervous system (olfactory bulb, substantia nigra, ventral tegmental area, and the retina) and the periphery (enteric nervous system of the gut). These neurons are involved in modulating intestinal motility, the sense of smell, light sensitivity, and control of fine motor movements. They are also integral to higher brain functions such as salience, euphoria, pleasure and appetitive and consummatory aspects of reward. Their demise or dysfunction plays vital roles in neuropathologies such as Parkinson’s disease, schizophrenia and depression. Thus, understanding how DA neurons function normally in neurotransmission and in pathological processes is essential for elucidating their roles in behavior and disease.

Electrochemical Detection of DA

The ability to detect DA and its metabolites electrochemically (Kissinger, Hart, and Adams 1973) greatly advanced the study of dopaminergic transmission. Microdialysis provided the first studies of DA levels in the brain in response to pharmacological treatments and during behavioral tests. Limitations of microdialysis include the relatively slow time resolution (on the order of several minutes) and the size of the probes, which lead to local tissue damage (Borland et al. 2005). The development of the carbon fiber electrode (CFE) enabled the detection of oxidizable transmitters with much faster time resolution and at the cellular and quantal level (transmitter released by a single vesicle) (Gonon et al. 1978). Initial studies performed by Wightman and colleagues on adrenal chromaffin cells which contain large dense core vesicles (LDCVs) (Leszczyszyn et al. 1990; Wightman et al. 1991) were followed by other studies utilizing carbon fiber amperometry to detect oxidizable transmitter release from both LDCVs and small synaptic vesicles (SSVs) from various cell types (Alvarez de Toledo, Fernandez-Chacon, and Fernandez 1993; Chen, Luo, and Ewing 1994; Bruns and Jahn 1995; Chen and Ewing 1996; Zhou, Misler, and Chow 1996; Pothos, Davila, and Sulzer 1998). The techniques outlined here are the first to provide for measurement of neurotransmitter release by SSV exocytosis from a central nervous system synaptic terminal.

Quantal DA Release

A major challenge in the study of DA neurotransmission has been the development of a system (cell culture, slice or other) in which it is possible to record quantal DA release. Most studies of quantal transmitter release have been performed on neuroendocrine cells such as adrenal chromaffin cells. Stimulation of these cells yields many readily detectable release events, although the release occurs from LDCVs in the cell body (Wightman et al. 1991), not from SSVs on axons as in neurons. Brain slices are widely used to study DA overflow, but are not widely used for amperometric recordings of quantal DA release because it is virtually impossible to record directly from varicosities due to steric hindrance by glia or other cells and structures. There is a single report of quantal recordings in the slice by Jaffe, Marty, and Schulte (1998), in which the surfaces of cell bodies were “cleaned” by a pressure stream, providing evidence for quantal release from substantia nigra cell bodies. This approach has proven technically challenging, and it is impossible to differentiate between release from cell bodies versus release from axonal terminals that contact these cell bodies, such as serotonergic inputs from the raphe. The laboratory of John Dani also has had some success in recording relatively small spontaneous DA release events in the slice (2005), and Mark Wightman’s group has produced analogous recordings in vivo (2004). These events, however, are composed of multiple overlapping quanta that cannot be resolved.

Another challenge exists in developing a preparation to study the exocytosis of SSVs from synaptic terminals due to their much smaller quantal size and rapid kinetics. We have shown that with available hardware, the amperometric signal resulting from the oxidation of transmitter release from a SSV is lost in the baseline noise of the current due to diffusion at distances greater than 300 nm (Sulzer and Pothos 2000). Therefore, the CFE must essentially touch the release site in order to minimize the filtering effect of diffusion on the amperometric signal. In addition, the background noise must be kept to a minimum and the sampling rate must be fast enough to obtain a reliable report of the event. Finally, the release site may also be blocked by the presence of a postsynaptic cell or by glial cells that cover the terminal. Despite this, there are a sufficient number of exposed release sites to make studies of quantal DA release possible, although patience is required to locate exposed sites. While it remains unproven, we suspect that SSVs may release from multiple sites at or near a varicosity, and this in part increases the chances of successful quantal recordings (Gonon et al. 2000; Sulzer and Pothos 2000).

Culture systems have several advantages for the monitoring of quantal DA release from SSVs. First, the DA neurons are intact and can survive for long periods during which they can be treated with drugs, transfected or otherwise manipulated. Second, ascorbate concentrations in the media are very low, and projections from other neurons with oxidizable transmitters are rare. Third, the absence of brain waves and the reduced electrical activity greatly reduce the background noise. The only system that has to date consistently provided recordings of quantal DA release from SSVs is the postnatally-derived ventral midbrain culture system. Amperometric studies of DA release from this preparation have already yielded important insights into the regulation of pre-synaptic quantal size and modes of transmitter release in DA neurons (Pothos, Davila, and Sulzer 1998; Pothos et al. 2000; Staal, Mosharov, and Sulzer 2004).

Source of DA Neurons: Selection of Animals and Brain Regions

Several factors need to be taken into consideration when determining the source of DA neurons. The choice of SN, VTA or both (ventral midbrain, VM) depends mostly on the neuropathology or neurological system to be modeled (Parkinson’s disease, SN; schizophrenia and depression, VTA; etc.) as well as the type of experiments (multi-dish toxicity studies, VM). For toxicity studies, the source of DA neurons is often less important than producing an adequate yield of DA neurons. If a high percentage of DA neurons within a dish are needed, only the VTA is dissected. This may be necessary for assays where measured parameters need to be maximized to reach detectable levels (DA levels, TH activity etc). If a high yield (many cultures) is desired, the entire VM is used. This would be appropriate for studies with many treatment groups and an easily measured parameter (such as number of cells/dish). Since we are trying to increase our success rate of finding active release sites, we usually opt for VTA cultures which have a very high density of DA neurons and DAergic axons and varicosities versus SN or VM cultures. Approximately 1 rat cortex yields enough glia to plate ~125 dishes, and one rat VTA yields enough DA neurons to plate 3–4 dishes. There is no distinct boundary between the lateral VTA and the medial SN, however; Figure 16.1 and Figure 16.2 show the approximate location of the incisions we make when dissecting these nuclei. In those studies where a comparison between VTA and SN is required, we dispose of the border regions between the nuclei (Burke, Antonelli, and Sulzer 1998).

FIGURE 16.1. (See color insert following page 272.

FIGURE 16.1

(See color insert following page 272.) Sagittal section of P1 mouse brain stained for TH using horseradish peroxidase and diaminobenzidine. Black lines indicate the incisions made during the dissection. (MF, midbrain flexure; VTA, ventral tegmental area.) (more...)

FIGURE 16.2. (See color insert following page 272.

FIGURE 16.2

(See color insert following page 272.) Coronal section of P1 mouse brain stained for TH using horseradish peroxidase and diaminobenzidine. Black lines indicate the incisions made during the dissection. (A, aqueduct; C, cortex; H, hypothalamus; MF, midbrain (more...)

The choice of animal species and strain is also critical for successful experiments. For example, amperometric recordings from cultured DA neurons from rats yield readily detectable amperometric spikes, whereas those from mice are much smaller, making their detection difficult if not impossible (unpublished observations). This may be due to lower levels of VMAT2 in striatal SSVs in mice (Staal et al. 2000). One drawback to using rats, however, is that it is currently impossible to visually differentiate between varicosities from DA neurons and other neurons. In our experiments to date (Pothos, Davila, and Sulzer 1998; Staal, Mosharov, and Sulzer 2004), axonal varicosities were located without knowing whether the varicosity was DAergic or not. Electrodes were placed onto the varicosity and secretagogue was applied and only the presence of amperometric events confirmed the varicosity as DAergic. Thus, recordings may be attempted at dozens of sites before finding an active DAergic release site. One potential solution is to perform recordings on cultured DA neurons derived from strains of mice expressing fluorescent proteins under appropriate promoters, such as the TH or DAT promoter or expressing proteins tagged with fluorescent proteins (GFP-hDAT, Staal et al, unpublished) (Matsushita et al. 2002; Zhang et al. 2004; Zhuang et al. 2005).

Methods

Various methods for preparing dissociated cultures of DA neurons are available. Most preparations produce cultures of embryonically-derived neurons of which approximately 1% are dopaminergic. The postnatally derived cultures developed by Rayport and Sulzer (1992) have several advantages, the most striking being that the fraction of DA neurons is usually at least 20% for VM and as high as 70% for VTA. Another advantage of postnatally derived neurons is that they display higher levels of the DA uptake transporter (Staal, unpublished observations).

Under optimal conditions, these cultures can survive up to 6 months, although most survive about 2 months. We suspect that the death is mostly due to increased osmolarity of the medium, and could probably be controlled by partially exchanging the older media with fresh conditioned media, testing the osmolarity of the medium over time with an osmometer and compensating with the addition of water or using a dish cover that allows air transfer but not evaporation. The latter approach has recently been introduced by Steve Potter (2001), but has not been attempted to date for the postnatal DA neurons.

We have cultured DA neurons on various types of surfaces and dishes, including multi-well plates, although we generally prefer to use glass coverslips that are glued to petri dishes using SYLGARD (SYLGARD® 184 Elastimer kit, Dow Corning Corporation, Midland, MI), so that the neurons produce small, dense cultures. This system maintains the cultures in a healthy state for weeks to months without requiring feeding with new media (for up to several weeks). The glass allows optimal microscopy with DIC/Nomarski imaging and fluorescent imaging. The central glass well allows most of the neurons to be approached with electrodes (except those within ~50 μm of the edge of the well).

Preparation of Glass Coverslips

  1. The utmost care should be taken to always maintain a sterile environment and sterile technique.
  2. Drop 100 coverslips [15×15 mm coverslips (deckglöser), Assistant, Germany purchased through Carolina Biological Supply Co., Burlington, NC] one at a time into a 200 ml beaker containing 100 ml of 95% ethanol (leave for at least 1 min).
  3. Prepare a solution of poly-d-lysine (PDL) in water (H2O). Always use ultra-pure water such as tissue culture water or water purified by reverse osmosis to a resistance of ≥18 Ω. The final concentration should be 40 μg/ml, and 250 ml are adequate for 100 coverslips. Place the PDL solution in a glass tray (~4 cm H×100 cm2) so that the PDL solution is at least 2 cm deep.
  4. Pour off excess ethanol and place wet cover slips on to a paper or glass container.
  5. Using forceps, pass individual coverslips through the flame of a Bunsen burner once.
  6. Cool the coverslip by gently waving it in the air in the air for approximately 10 s.
  7. Place the coverslip into the PDL solution (cracking or sizzling sounds mean it did not cool enough). Repeat for all coverslips.
  8. Let the slips sit for 1 h, then pour off the PDL and allow the slips to dry.
  9. Punch 10 mm holes into 50×9 mm Petri culture dishes (or whichever dishes fit the desired dish holder for the physiology set up) using a bench punch (Roper and Whitney No. XX, Rockford, IL).
  10. Wearing gloves, prepare about 5 ml of SYLGARD in a large weighing boat by combining 10 parts resin with 1 part catalyst (by weight) and stirring extensively with a glass stirrer. Pour contents into new weighing boat to avoid use of resin without catalyst.
  11. Dip an empty 15 ml conical culture tube or glass scintillation vial upside-down into the SYLGARD, using it like a rubber stamp to apply a circle of SYLGARD to the underside of the dish, around the hole. Use caution when applying the SYLGARD as too much will result in excess SYLGARD covering the culturing surface of the coverslip whereas inadequate SYLGARD will result in a leaky well.
  12. Place (drop) the coverslips on the circle of SYLGARD taking care to prevent bubbles.
  13. Allow SYLGARD to spread (about 15 min).
  14. Cure the SYLGARD, place dishes, upside-down, at 37°C overnight, or at room temperature for 2–3 days (cured SYLGARD is no longer sticky to the touch). Sterlize dishes under UV light for 2 h.
  15. Note that these glass coverslips must be coated with laminin before culturing (see glia dissection protocol). Once the dishes are cured they can be stored in zip-lock bags at 4°C until needed.

Preparation of Pipette “Tech-Tips” for Triturating Cells

  1. Obtain a box of 1000 μl pipette tips (no-filters) and flame all tips in order to seal off the hole.
  2. Using either a 21G 1½ (Large), 22G 1½ (Medium) or 26G 1½ (Small) needle, poke a hole as close to the tip as possible passing straight through, in one side and out the other. This will result in two holes on opposite sides of the tip. The purpose of piercing through both sides is to prevent cells from being caught in the pipette during trituration.

SYLGARD Circles

SYLGARD circles are used as a dissection platform enabling one to pin down the brain using forceps without damaging the tips of the forceps or blunting the scalpel blade. SYLGARD that is left over from the preparation of dishes can be used to make SYLGARD circles.

  1. Weigh out SYLGARD, mix the components well and pour into wells of a 12 well plate to a depth of approximately 3–4 mm.
  2. Cure in oven at 60°C for 30 min or at room temperature for 2–3 days.
  3. Pull circles out of wells using razor and individually package each circle in aluminum foil and autoclave.

Glia (Using Rat Pups)

  1. Add 100 μl laminin (10 μg/ml) per round well for a minimum of 1 h at room temperature before plating cells and leave in flow bench (or leave overnight at 4°C). Aspirate off laminin and wash each well with 200 μl MEM. Aspirate off all remaining MEM and place into 37°C incubator (CO2 5%) to warm up before plating cells.
  2. Obtain one or two P1–P3 mouse or rat pups (animals which are 1–3 days old).
  3. Pre-heat the water bath to 37°C (Mistral Multi Stirrer, Lab-Line, Melrose Park IL; Neslab EX-7, Thermo Electron Corp, Austin, TX).
  4. Prepare the papain dissociation solution (Table 16.1B). Each vial should hold tissue from a maximum of 4–6 animals.
  5. Sterilize the top of each Nunc specimen vial (25 ml) with ethanol. Punch two holes in the top with an 18 gauge needle, using a separate needle to bore holes. In one hole place a new needle with a 0.22 μm filter attached. Leave the other hole empty for ventilation.
  6. Add an HCl-cleaned, autoclaved micro-stir bar to the vial. Fill each tube with 5–10 ml of freshly made papain solution (sterile filtered with 0.22 μm filter). Place in temperature-controlled 37°C water bath with magnetic stirrers.
  7. Arrange dissection tools, microscope, dissection light, transfer pipettes, sterile Petri dish with SYLGARD square, beaker, and 70% ethanol in a 15 ml tube in the sterile hood.
  8. Prepare two buckets of ice to chill the PBS and for the pups.
  9. Cut aluminum foil squares (about 10×10 cm).
  10. Perfuse dissociation tube (papain tube) with a steady flow of humidified carbogen (95% oxygen, 5% CO2) delivered through the filter/needle (flow rate adjusted to about 1 bubble per second).
  11. When the papain has reached a pH of 7.2–7.4 in about ½ –1 h (indicated by a red color; purple indicates that the papain solution is too alkaline and orange that it is too acidic), anesthetize pups and begin dissection.
  12. Anesthetize pups with an intraperitoneal injection of ketamine (0.01 and 0.05 ml of 100 mg/ml for mouse and rat pups respectively).
  13. When pups begin to show sedation, put them on ice until unresponsive and hypothermic.
  14. Rinse aluminum foil square, scissors and pup with 70% ethanol.
  15. Decapitate pup, allowing head to fall onto aluminum foil square and move to laminar flow hood. Using (toothed) forceps hold head by the eyes (using left hand if you are right handed). Using the other hand and a curved or angled forceps, pinch the scalp just behind the eyes and pull back, tearing it off. Next, use a micro-scissor to cut around the circumference of the skull, and gently peel the skull off with a forceps, being careful in case any tissue still connects it to the rest of the skull. Gently remove brain (using a small scoop shaped scalpel) and place into Petri dish with one SYLGARD circle stuck onto the bottom and filled ice cold sterile PBS (use enough PBS so that the brain lying on the SYLGARD circle is submerged).
  16. Dissect out the cortex taking care to remove the hippocampus. Cut into 1 mm3 chunks.
  17. Place chunks in papain (Table 16.1A), without kynurenate (1 cortex/vial). Chunks should be digested until the tissue looks a bit fluffy like cotton candy (10–30 min, so it can be easily triturated). In the meantime, warm up 45 ml sterile M10C-G for rinsing and trituration (Table 16.2).
  18. From this point forward, everything should be done in a sterile laminar hood with sterile technique.
  19. Remove the chunks from enzyme vial using a transfer pipette and place in 15 ml conical test tube.
  20. Rinse the chunks using transfer pipettes 3 times with 2 ml M10C-G, allowing them to settle each time, and discard supernatant.
  21. Using 1 ml warm M10C-G triturate up to a maximum of 10 times (3–5 times should be sufficient) using a regular 1 ml pipette tip.
  22. Remove supernatant to a separate tube, and replace the volume with 1 ml fresh M10C-G.
  23. Repeat trituration, first using a large, then a medium, then a small bore “tech-tip” until the chunks are dissociated.
  24. Pellet the glial cells at 1000×g for 5 min.
  25. Resuspend the pellet in 8 ml M10C-G, triturate 5 times to mix, and then count the cells.
  26. Dilute the cell suspension to a plating density of 1,000,000–1,500,000 cells/ml and plate about 80,000 cells per well.
  27. Plate by dripping the cell suspension from about 10 cm above the dish to help kill neurons.
  28. Tap the sides of the trays in order to evenly distribute the cell suspension.
  29. Allow cells to settle for 1½ –2 h in incubator. Chill the M10C-G in a 4°C refrigerator or on ice.
  30. For monolayers: add 2 ml of cold MEM into each well, moving the pipette around in a circular motion to wash each well evenly.
  31. Aspirate off MEM and repeat the wash. Aspirate off the MEM and feed with 2 ml of cold M10C-G (Table 16.3Table 16.4).
  32. Add FDU (Table 16.5) to the dishes when glia are ~70% confluent (about 3–5 days). If there are still neurons present, remove the medium and cold-feed with 2.5 ml M10C-G before adding FDU. If neurons still persist, place cells at 4°C for 1 h.
  33. Astrocytes dishes should be at least one week old and at most 4 weeks old at the time the neurons are plated. Feed with cold M10C-G at one month if not already used.
TABLE 16.1B

TABLE 16.1B

Papain Solution for Neurons (Prepare Just prior to Culturing)

TABLE 16.1A

TABLE 16.1A

Papain Solution for Glia from 1 to 2 Cortices (Prepare Just prior to Culturing)

TABLE 16.2

TABLE 16.2

Cysteine Water

TABLE 16.3

TABLE 16.3

H&B Concentrate (5×)

TABLE 16.4

TABLE 16.4

M10C-G

TABLE 16.5

TABLE 16.5

5-Fluorodeoxyuridine (FDU) Solution

DA Neurons

  1. The day before culturing neurons, wash glial cultures twice with 2 ml of cold MEM. Replace MEM with 2 ml of SF1C (Rosenberg 1988). (Table 16.7, Table 16.8) Alternatively, neurobasal medium A/B27 can also be used (Table 16.9). While A/B27 is easier to make, we have had slightly better success with SF1C in toxicity assays (unpublished observations).
  2. Place sterile rings inside the dish but to the side of the well, not around the well, using forceps with sterile tips (Thomas Scientific, catalog no. 6705R12). The rings will later be placed around the wells to ensure that when the neurons are plated they settle into the well and do not float or wash away into the rest of the dish. Rings should be cleaned by washing with 0.1 M HCl for at least 1 h, and then rinsed at least 10 times with distilled water before autoclaving. Any remaining acid will kill neurons the next time the rings are used!
  3. Leave dishes with SF1C in the incubator overnight.
  4. Pre-heat the water bath to 37°C.
  5. Make the papain dissociation solution (Table 16.1B). Each vial should hold tissue from a maximum of 4–6 animals.
  6. Sterilize the top of each Nunc specimen vial with ethanol. Make two holes in the top with an 18 gauge needle, using a separate needle to bore holes. In one hole place a new needle with a 0.22 μm filter attached. Leave the other hole empty for ventilation.
  7. Add an HCl-cleaned, autoclaved micro-stir bar to the vial. Fill each tube with 5–10 ml for neuronal cultures (2 ml for neuronal cultures from individual pups) of freshly made papain solution (sterile filtered with 0.22 μm filter). Place in temperature-controlled 37°C water bath with magnetic stirrers.
  8. Arrange dissection tools, microscope, dissection light, a bunch of transfer pipettes, sterile Petri dish with SYLGARD square, beaker, and 70% ethanol in a 15 ml tube in the sterile hood.
  9. Get two buckets of ice to chill the PBS and for pups.
  10. Cut aluminum foil squares (about 10 cm2).
  11. Obtain as many P1–P3 rat or mouse pups as needed (animals which are one to three days old).
  12. Perfuse dissociation tube (papain tube) with a steady flow of humidified carbogen delivered through the filter/needle (flow rate adjusted to about 1 bubble per second).
  13. When the papain has reached a pH of 7.2–7.4 in about ½ –1 h (see Glia step 11), anesthetize pups and begin dissection.
  14. Anesthetize pups with an intraperitoneal injection of ketamine (0.01 ml and 0.05 ml of 100 mg/ml for mouse and rat pups).
  15. Gently remove brain into ice cold PBS-filled Petri dish as described for glia. With anterior of brain facing left on the SYLGARD circle (dissection will be described for right handed people) use a fine forceps in the left hand to pin the two hemispheres of the brain to the SYLGARD. Remove any remaining meninges.
  16. Place the brain, ventral side down, anterior facing left onto the SYLGARD. Using forceps, pin the brain to the SYLGARD to immobilize. Make the initial cut through the entire brain caudal to the midbrain flexure (right line, Figure 16.1), and the second cut rostral to the flexure, including the caudal aspect of the hypothalamus. Lay the resulting slice flat on the SYLGARD with the ventral side facing right, and the dorsal side to the left and the caudal aspect up. Remove any remaining cortex. Place the forceps through the aqueduct and push into the SYLGARD to immobilize the slice. Cut the ventral edge of the slice along the top of the hypothalamus as indicated by the line in Figure 16.2. Next cut approximately halfway between the new ventral edge of the slice and the aqueduct. Cut into 1 mm3 segments and place all three segments into the vial with papain solution for ventral midbrain cultures. For VTA cultures only use the two outer segments and for SN only use the inner segment.
  17. Set the magnetic stirrer on low. Chunks should gently swirl around vial.
  18. Digest for approximately 10–20 min or until chunks have fallen apart and look fluffy, like cotton candy. Periodically examine the tubes to see if segments are sticking to the stir bars; if so, tap vials to knock the tissue loose.
  19. Place tubes containing SF1C for rinsing and triturating in the incubator until ready to triturate, so the pH stays between 7.2 and 7.4. The media must be at the correct pH (indicated by the red color) or the neurons will die.
  20. Use forceps with sterile pipette tips at the end (so they can be changed between dishes) to slide the rings from the side of the dish over the well so the ring acts as a wall extending the height of the well. The rings should be removed 2 h after plating.
  21. From this point forward, everything should be done in a sterile laminar hood with sterile technique.
  22. Remove chunks from enzyme vial using transfer pipette and place in 15 ml conical test tube. Take great care to prevent bubble formation because bubbles are harmful to cells. If bubbles occur, aspirate them off.
  23. Rinse chunks 3 times using about 2 ml pre-warmed SF1C. After adding SF1C for each wash, flick the tube so that the tissue swirls around and is washed. Allow the tissue to settle each time and discard as much of the supernatant as possible without exposing the tissue to air.
  24. Triturate in 1 ml of room temperature SF1C (for tissue from 4 to 6 pups or 0.5 ml for individual pups) with large-bore “tech-tips” up to 10 times, preferably fewer (when done properly the medium becomes slightly cloudy with dissociated cells). Let the tube stand until the cells settle, remove the supernatant to a new tube and replace with equal amount of fresh SF1C.
  25. Repeat the trituration process up to three times with a medium-, then a small-bore “tech-tip.” The chunks should now be completely dissociated into individual cells.
  26. Centrifuge cells at 1000×g for 5 min. Re-suspend the pellet in 0.2–0.5 ml (for individual and pooled mouse cultures) or 1 ml (for cultures from 4 rat pups) of SF1C media, mix and count (see below). Dilute the cell suspension until 75–120 μl of media contain 80,000 cells. Add the appropriate volume to the well (with the slide ring around it) of the SF1C-containing dishes so that each well contains 80,000 cells (use a circular motion to spread the cells around). Again, the rings should ensure that the neurons settle into the well and do not float away or get washed away when moving the dishes.
  27. After 1½ –2 h, cells should have attached to the well. Remove rings from each well using forceps with sterile pipette tips at the end, again, changing tips between dishes.
  28. 1½ –2 h after plating, add 100 μl GDNF (200 ng/ml, frozen aliquots) to all dishes (containing 2 ml SF1C media), for a final concentration of 10 ng/ml. We have shown that GDNF enhances quantal size (Pothos, Davila, and Sulzer 1997).
  29. Allow cells to settle in a 37°C incubator overnight (5% CO2).
  30. The next day add 20 μl of FDU (diluted from stock) to inhibit the growth of non-neural cells (use 1000× stock).
  31. SF1C media can be changed when necessary (change of color) by aspirating half the media and replacing an equal volume of fresh media (pre-warmed in incubator!).
  32. Disturb cultures as little as possible.
TABLE 16.7

TABLE 16.7

Stock Solutions for DiPorzio Concentrate

TABLE 16.8

TABLE 16.8

DiPorzio Concentrate (10 ml, 100×)

TABLE 16.9

TABLE 16.9

Neurobasal A/B27 Medium

Media and Reagents

Table 16.1 through Table 16.9

Carbon Fiber Amprometry

Electrode Fabrication

  1. A carbon fiber bundle (individual fibers are 5 μm in diameter, Type T650-42, grade 12K, finish 309, Thornel® carbon fibers, Cytec Industries, West Paterson, NJ) is cut into approximately 10 cm sections (the length of the glass capillary tube) and placed into the middle of a folded piece of paper so that the bundle protrudes about 1 cm. This is placed onto another sheet of white paper (so that the black carbon fibers are more easily seen). The protruding fibers may be gently rolled between thumb and forefinger or otherwise manipulated with the other hand until the carbon fiber cable is frayed. While one hand holds down the folded piece of paper containing the carbon fiber bundle, the index finger of the other hand pins down an individual carbon fiber. The folded paper holding the cable of fibers is pulled back, leaving behind a single carbon fiber.
  2. A glass capillary filament tube (12 mm×0.68 mm, 10 cm long, A–M Systems, Inc Everett, WA) which has been kept in 70% ethanol is attached to a vacuum socket which dries the capillary tube. Pinning the carbon fiber down with an index finger at one end, the other end of the carbon fiber is sucked into the capillary tube which is moved towards the base of the fiber/index finger until the entire fiber is in the tube and tube is touching the index finger pinning down the fiber. Disconnect the tube from the vacuum before releasing the carbon fiber! Then, trim the protruding ends of the carbon fiber with a razor or scalpel blade.
  3. The carbon fiber-containing capillary tube is then pulled into two equal parts using a Sutter Flaming/Brown Micropipette Puller (P-97; Novato, CA). Usually the carbon fiber remains intact, so it is necessary to cut the fiber using a scissor before removing electrodes from the puller. Otherwise it will pull out of one side. Settings on the micropipette puller vary, depending on the installation of the filament as well as the filament geometry. The pipette should be pulled so that the tip is gently tapered from the original capillary diameter to the point where the carbon fiber protrudes (approximately 5 mm). A short length (approximately 0.1–0.3 mm) of the carbon fiber should be tightly encased by the glass pulled tightly around the carbon fiber. In order to reduce noise, the pipette tips are quickly dipped into molten stickly wax (Grobet, Carlstadt, NJ), held for about 5 seconds to allow the wax to seal any remaining gaps or cracks between the pipette and carbon fiber and slowly removed to allow the excess wax to run off the pipette and carbon fiber, thus preventing beading up of the wax along the fiber. Pipettes are then beveled (marked side up) at approximately 45° (the same angle that will be used for recording when the electrode is in the holder). Electrodes may be purchased commercially, although we have found to date that commerically produced electrodes are too nosiy to record from neurons.

Testing and Selection of Electrodes

Electrodes are backfilled with K+ acetate (4 N). Silver wires for the pipette holder and ground electrodes (0.25 mm diameter, Warner Instrument Corp., Hamden, CT) are coated with chloride using 1 N HCl and a 9 V battery. Electrodes are tested by running a voltammogram (−200– 800 mV at 200 mV/s) with the tip of the CFE immersed in 100 μM DA in physiological saline (in mM:128 NaCl, 2 KCl, 10 HEPES, 1 Na2HPO4, 2 MgCl2, 1.2 CaCl2, 10 d-glucose; ~300 mOsm and pH 7.4) using a Ensman potentiostat (Ensman instruments, Bloomington IN). Electrodes with a sigmoid shaped voltammogram are then mounted on the headstage of an Axopatch 200 B amplifier (Axon Instruments, Foster City, CA). A voltage of +700 mV (filtered with a built in 4-pole 10 kHz Bessel filter) is applied with the tip of the CFE and the reference electrode immersed in physiological saline. The root mean square of the background noise should be less than 1.1 pA for neuronal recordings.

Recording

Neurons cultured for at least 3 weeks are used for amperometric recordings as it takes this long for VMAT2 and DAT to reach maximum expression levels (Staal unpublished observations). We have rarely had success obtaining quantal DA release before 2 weeks. Neuronal cultures are typically incubated for at least 1 h with 100 μM l-DOPA prior to changing to physiological saline (listed above), which enhances quantal size (Pothos, Davila, and Sulzer 1998). 100 μM l-DOPA may be added to the saline while recording. This will raise the baseline current but will not increase the noise. Using micromanipulators (Newport micromanipulator MX300R, Irvine, CA), electrodes are positioned above the desired recording site and slowly lowered until minimal depression of the structure (cell body or varicosity) is seen. Carbon fibers are held at +700 mV and the resulting current is filtered using a low pass 4-pole, 10 kHz Bessel filter built into an Axopatch 200B amplifier. The current is digitized and sampled at 100 kHz (PCI-6052E, National Instruments, Austin, TX). Start recording as soon as possible followed by application of secretagogue (usually after 10 s). Data is recorded using IGOR PRO (WaveMetrics, Lake Oswego, OR) and saved as an IGOR binary file for further analysis.

We generally use potent secretagogues, such as high potassium (in mM: 52 NaCl, 80 KCl, 10 HEPES, 1 Na2HPO4, 2 MgCl2, 1.2 CaCl2) or α-latrotoxin (a generous gift from A. Petrenko, New York University, New York, NY) and potassium (80 mM K+ solution and 20 nM α-LTX). Secretagogues are applied by local perfusion through a glass micropipette (Picospritzer, General Valve, Fairfield, NJ) for 3–10 s at 10–15 psi and ~5–15 μm from the recording site depending on response. Data should be acquired for several minutes as events often appear at long intervals and sometimes after a significant delay following secretagogue application.

Since amperometric spikes are so small and fast they are often difficult to distinguish from action potentials or a spike due to a noisy electrode. While electrodes with any current spikes should be discarded, an occasional noise-related spike may occur even with good electrodes. However, non-amperometric spikes often consist of a single point (when recording at 100 kHz), are typically symmetrical (like an isosceles triangle) or have a component right before or after the spike that drops below the baseline (a negative current component). Another method to verify that the amperometric events are not mere artifacts depends on finding a site that continuously releases DA. First, the electrode can be raised off the recording site. All spikes should cease, confirming that they are not due to current leaks in a bad electrode. Second, the holding potential can be slowly reduced from +700 to 0 mV and after 30 s or a min, brought back to +700 mV. All amperometric events should disappear as the potential goes to zero and reappear as it approaches the oxidation potential of DA. Only spikes due to action potentials will persist while the voltage is 0 mV.

On rare occasions, varicosities with a small fraction of very large amperometric events have been observed. The quantal size of these spikes resembled large vesicles as utilized by serotonergic neurons (Bruns and Jahn 1995). Due to their infrequency in DA neurons, they could also be eliminated statistically as they were more than 5 standard deviations larger than the average quantal size (Staal, Mosharov, and Sulzer 2004). These events could also correspond to exocytosis of “small” dense core vesicles (~100 nm diameter) that we have noted in electron micrographs, which typically represent less than 1% of the vesicles in synaptic terminals (Pothos et al. 1998).

Analysis

The detection and analysis of amperometric peaks has been reviewed by Mosharov and Sulzer (in presss), and our analysis program, written for Igor Pro (4.07 or later), can be downloaded from our laboratory web site:

http://cumc.columbia.edu/dept/neurology/sulzer/download.html

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Copyright © 2007, Taylor & Francis Group, LLC.
Bookshelf ID: NBK2577PMID: 21204391

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