Nafion Exclusion Layer

Nafion^® is an anionic Teflon^® derivative and is one of the most widely used methods for improving the selectivity of voltammetric recordings in CNS tissue. Historically, Nafion is used on the surface of carbon fiber microelectrodes for the detection of catecholamines [14–19]. The negatively charged sulfonic acid groups substituted into the polymer matrix of Nafion repel anions from the Carbon or Pt recording surfaces. If these anions are not blocked, they cause high background signals that interfere with reliable recordings. At the same time, the negatively charged sulfonic acid groups on Nafion concentrate cationic species such as DA, NE and 5-HT. With the development of the multisite microelectrodes, Nafion is a logical choice for an exclusion layer and is used reliably in our studies of L-glutamate [1–5,12]. However, because Nafion does attract catecholamines to the Pt recording surface, which are oxidizable on Pt, other exclusion layers are used in CNS regions with high monoamine levels.

Nafion (5%, Sigma-Aldrich, Catalog #27,470-4) is aliquoted into a 300 μl centrifuge tube. The microelectrode tip is lowered halfway into the aliquot of Nafion and rotated in a circle five times lasting approximately 1 s per rotation. Next the tip is pulled straight out of the Nafion solution. Finally, microelectrodes are placed into an oven at 165°C–175°C for 4 min to allow the Nafion layer to cure on the tip. In general, lower temperature curing produces thicker films compared to higher temperature curing [20]. Also, longer drying times at higher temperatures produce microelectrodes with lower responses to both AA and to analytes such as L-glutamate. Microelectrodes coated with excessive Nafion show little or no detectable response to electrochemically active molecules. Microelectrodes coated with not enough Nafion record high levels of background AA once inserted into the brain. After the curing procedure, microelectrodes are allowed to cool to room temperature for 30 min before enzyme coating (described in the section on Enzyme Coating Layers). An important concept to remember is that once a prepared Nafion coated microelectrode is placed into an aqueous solution (either a beaker or the brain), then the tip must remain wet or at the very least, removed from solution for only a limited time. When Nafion dries after soaking, it can crack allowing areas where interferents pass through and are oxidized on the Pt recording surfaces. If this occurs, the electrode is no longer selective for the analyte being measured.

1,3-Phenylenediamine

The organic molecule 1,3-phenylenediamine (mPD) is another chemical used to create an exclusion layer for Pt recording electrodes [21,22]. A potential is applied to a solution of mPD, thus causing mPD to electropolymerize onto the Pt recording surfaces. Historically, mPD or its derivate 1,2-phenylenediamine (o -phenylenediamine), is used on carbon fiber microelectrodes for selectivity and to prevent electrode fouling—a process where oxidizable molecules such as 5-HT can adhere to the carbon fiber surface thus preventing detection. Electropolymerized mPD selectivity is achieved by forming a size exclusion layer that prevents larger molecules such as AA, DA, and DOPAC from reaching the recording surface. Smaller molecules, such as nitric oxide and peroxide, are still able to pass through the matrix [23]. Since peroxide is a reporter molecule for oxidase enzymes, it makes mPD an ideal exclusion layer for our enzyme-based multisite microelectrodes [21,24].

The electropolymerizing procedure is completely different from Nafion coating but has four distinct advantages: (1) The matrix formed by mPD blocks DA, NE, and 5-HT from reaching the Pt recording sites. This helps make the microelectrode more selective for measuring the analyte of interest; (2) The mPD is electropolymerized onto the microelectrode after the microelectrode is coated with an enzyme layer. The mPD electropolymerizes through the enzyme layer to form the matrix. Since mPD is electropolymerized after enzyme coating, additional coatings of mPD can be applied to the microelectrode surface without having to clean the microelectrode. If the matrix degrades during an experiment, the microelectrode can be replated, recalibrated, and re-used to finish the experiment; (3) Because a potential must be applied, Pt recording sites can also be selectively coated with mPD. Removing the applied potential between a Pt recording site and the headstage prevents mPD from being electropolymerized onto that site; and (4) Once mPD has been successfully electropolymerized onto the Pt recording sites, the solution does not have to remain wet. In fact, our laboratory has found that exposure to aqueous solutions slowly degrades the matrix over time. The procedure for electropolymerizing mPD to the Pt recording sites is outlined below.

Our laboratory has found that 1,3-phenylenediamine dihydrochloride, 99% (Sigma-Aldrich, Catalog #235903-25g) works best for electroplating and matrix formation on our Pt recording sites. First, a solution of 5 mM mPD is prepared in a degassed solution of 0.05 M phosphate buffered saline (PBS). Degassing is accomplished by bubbling nitrogen gas through the 0.05 M PBS solution for 20 min to remove oxygen before dissolving mPD. Once the 5 mM mPD is dissolved in the degassed 0.05 M PBS, the mPD solution is stored in a brown glass bottle to prevent oxidation from light. If the solution turns yellow, it has oxidized and will no longer electropolymerize. Even with these storage methods, the 5 mM mPD solution can oxidize after 3 h so it is recommended using the solution immediately after preparation. When the mPD solution is prepared, the microelectrode is connected to the FAST-16 recording system and approximately 40 ml of the 5 mM mPD is poured into a 50 ml beaker. The microelectrode tip is lowered halfway into the solution along with a glass, Ag/AgCl reference electrode (Bioanalytical Systems, Inc. RE-5B, Catalog #MF-2079). A constant potential of + 0.5 V versus a Ag/AgCl reference electrode is applied for 15 min to allow the mPD to electropolymerize onto the Pt recording sites. At 15 min, the microelectrode tip is removed from the 5 mM mPD solution, rinsed with ddH₂ O, and stored at room temperature for twenty-four hours prior to calibration.

Oxidase Enzymes

Enzymes provide a means to convert a molecule that is not inherently electroactive and thus not measurable, into a reporter molecule such as peroxide that is oxidized at the Pt recording surfaces of the microelectrodes. The current measured from the oxidation of peroxide generated during the enzymatic breakdown is directly proportional to the analyte concentration. Table 19.1 provides a list of available oxidase enzymes and their potential uses. Some compounds require multiple enzymes to convert them to a reporter molecule, such as acetylcholine and γ-aminobutyric acid (GABA). Oxidase enzymes that our laboratory has used to measure neurochemicals are highlighted in Table 19.1 and include L-glutamate oxidase [1–5], choline oxidase [25,26], L-lactate oxidase [27], and L-glucose oxidase. Furthermore, acetylcholine esterase is used to measure acetylcholine (in conjunction with choline oxidase) and catalase is used to prevent peroxide detection.

TABLE 19.1

Available Oxidase Enzymes with Their Substrates, Products, and Neurochemicals

As previously mentioned, the O₂ dependence of oxidase enzyme-coated microelectrodes is also a concern. It is widely known that O₂ is required by the enzymes to measure the analyte, but a portion is returned to the tissue by the oxidation of peroxide to O₂ and H⁺ at the microelectrode surfaces. Often oxidase enzyme-coated microelectrodes are relatively free from O₂ dependence over a useful range in vivo [25], but certain applications, such as those during recordings of stroke episodes, may require correction by use of an O₂ sensing microelectrode.

Immobilization of enzymes to the Pt recording surface helps to stabilize the enzymes and makes them active for longer periods of time. Our laboratory has immobilized several of the oxidase enzymes onto the Pt recording surfaces. While the general procedure remains the same for cross-linking the enzymes to the Pt recording sites, relative ratios of the enzyme to protein mixtures may vary. Our laboratory focuses extensively on L-glutamate oxidase for the detection of L-glutamate dynamics in the mammalian CNS whose immobilization procedure is outlined in detail below.

L-Glutamate Oxidase Coating Procedure

Properly cleaned microelectrodes are ready for enzyme immobilization. A stock solution of L-glutamate oxidase (Seikagaku America, Catalog #100645-1) is prepared by adding 50 μl of ddH₂ O to the lyophilized, purified enzyme in a vial to make a final concentration of 1 U/μl. All proteins and enzymes are brought to room temperature and 0.10 g of bovine serum albumin (BSA) Fraction V, 99% (Sigma-Aldrich, Catalog #A-3059) is dissolved in a 1.5 ml microcentrifuge tube containing 985 μl ddH₂ O by manual agitation. Once dissolved, 5 μl of glutaraldehyde solution, Grade I, 25% (Sigma-Aldrich Catalog #G-6257) is added to the BSA mixture and manually mixed by inversion five times. The solution is then set aside for 5 min until it turns a faint yellow color. Glutaraldehyde crosslinks proteins to the microelectrode surface when cured. The BSA serves as a matrix to protect the oxidase enzyme activity during immobilization. Next, 9 μl of the BSA/glutar-aldehyde mixture is removed and added to a 300 μl microcentrifuge tube and to this 1 μl of the L-glutamate stock solution (1 U/μl) is added and mixed by pipette agitation. This 10 μl solution has a final concentration of 1% BSA, 0.125% glutaraldehyde, and approximately 1% L-glutamate oxidase. An extensive series of parametric studies were carried out to determine this optimized stoichiometry for enzyme immobilization.

All enzymes are currently applied to the Pt recording sites by hand. Solutions are drawn up into a 25 μl Gastight^® Hamilton microsyringe (Hamilton Co., Catalog #80065) and slowly dispensed to form a small droplet of solution at the tip of the microsyringe. Using a dissecting microscope (and a steady set of hands), the droplet is applied briefly to the Pt recording sites. The solution quickly dries, leaving behind a thin, translucent layer of enzyme that is visible underneath a dissecting microscope. Two additional coats of enzyme are applied in the same manner with 1-min dry times in between each coat. This procedure is complete once a visible film remains following coating. The advantage of using multisite microelectrodes is that a pair of recording sites is coated with the enzyme mixture and the adjacent sites are coated with the inactive protein matrix (BSA and glutaraldehyde) in the same manner as applying the enzyme mixture. The laboratory refers to this technique as “self-referencing” [1], and this approach has many advantages, which are discussed later in this chapter.

Once the enzyme and/or inactive protein matrix is applied, microelectrodes are stored at room temperature. Enzyme-coated microelectrodes must cure at room temperature for 48–72 h prior to calibration and experimentation. Complete curing increases enzyme layer adhesion to the micro-electrode surface. This process provides better sensitivity to L-glutamate as well as increases the shelf-life of the microelectrodes. Our laboratory recommends using the microelectrodes within three weeks after coating. The enzyme/protein layers on “uncured microelectrodes” can dissolve when put into solution. Optimal curing times may vary depending on temperature, humidity, and the type of enzyme. Appendix A provides a step-by-step procedure for cleaning and preparing L-glutamate microelectrodes.

Since photolithographic procedures can place adjacent Pt recording sites within tens of microns from one another, coating individual Pt sites with enzyme layers by hand is often difficult if not an impossible task. For this reason, our laboratory is developing a microcoater that can precisely control the application of microliter amounts of enzyme solutions. This instrument can precisely coat individual recording sites while significantly decreasing the variability caused by hand coating procedures. Additional details of this new technology will be published elsewhere once work is completed.

Calibration Preparation

Nafion coated microelectrode tips are soaked in a solution of 0.05 M PBS at 37°C for at least one hour prior to calibration. These microelectrodes can be placed in the PBS solution overnight for calibration the next morning. The soaking time allows for better diffusion of analytes through the Nafion layer as well as activation of the enzyme layer. The mPD-coated microelectrodes, on the other hand, do not require soaking. The time period that the microelectrode tip sits in the calibration solution is sufficient for proper enzyme activation without additional soaking.

All stock solutions for L-glutamate calibrations are prepared in ddH₂ O and listed below:

1. 20 mM AA (Ascorbic acid powder, Fisher Scientific, Catalog #A62)
2. 20 mM L-glutamic acid sodium salt hydrate, minimum 99% (Sigma-Aldrich, Catalog #G1626)
3. 2 mM DA (3-Hydroxytyramine hydrochloride, Sigma-Aldrich, Catalog #H-8502)
4. 8.8 mM hydrogen peroxide (Rite Aide, 3% Hydrogen Peroxide Topical Solution)

Solutions are stored in a refrigerator at 4°C. AA and hydrogen peroxide are made fresh daily since they oxidize in solution. L-glutamate is made fresh on a weekly basis. DA is kept for a month providing the stock solution contains 0.01 M perchloric acid.

Calibration Procedures

Using the FAST-16 recording system, a combination headstage gain of 2 nA/V and secondary gain of 10X generates a final 200 pA/V gain for all calibrations. Forty milliliters of 0.05 M PBS is measured out with a graduated cylinder and added to a 50 ml beaker. This beaker is placed in a recirculating water bath (Quanteon, L.L.C.) set at 37°C resting upon a battery operated, portable magnetic stir plate (Barnant Co., Model #700-0153). The calibration temperature is maintained at 37°C to activate L-glutamate oxidase for physiologically relevant temperatures. Our laboratory uses a battery operated stir plate to reduce potential AC (60 Hz) current that can affect the recordings. A 10×3 mm² stir bar is added to the 0.05 M PBS and the solution is slowly stirred to prevent forming a vortex in the solution. A glass Ag/AgCl reference electrode is placed in the buffer solution. Finally, the microelectrode tip is lowered halfway into the buffer solution. Figure 19.6a shows a photograph of the calibration system.

FIGURE 19.6

(a) Photograph of a microelectrode immersed in the calibration chamber. The calibration solution is maintained at 37°C with a heated waterbath. (b) Calibration of a Nafion coated L-glutamate (Glu) self-referencing microelectrode. During calibration, (more...)

After initial system settings are completed on the FAST-16 software, the FAST-16 recording system applies a potential of + 0.7 V versus the Ag/AgCl reference to the microelectrode recording surfaces. Once calibration has begun, the current is allowed to reach a stable baseline (approximately 10–15 min, but may take longer). When a stable baseline is achieved, the user marks the baseline. (When marking, the user selects a time point for an event, such as baseline, interferent, or analyte addition, which is recorded by the FAST-16 software. A series of data points before the mark are averaged and used in the calibration analysis.) Next, the user adds 500 μl of 20 mM A (interferent) for a final beaker concentration of 250 μM. When a new stable baseline is reached the user marks the interferent. Next, three 40 μl additions of 20 mM L-glutamate are added for final buffer concentrations of 20, 40, and 60 μM L-glutamate. Analyte marks are recorded after each addition to create the calibration curve. After 3 additions of analyte, 40 μl of 2 mM DA (2 μM, final beaker concentration) is added to the solution as a test substance and marked. Finally, 40 μl of 8.8 mM (8.8 μM, final beaker concentration) hydrogen peroxide is added to the solution to confirm microelectrode sensitivity, and the addition is marked. The test substances are used to check selectivity and can be used to normalize the enzyme coated and inactive protein matrix sites for self-referencing recordings. The additions of test substances do not factor into calculating the standard curve for the analyte of interest. All chemicals used in vivo should be tested in vitro to ensure that they are not electrochemically active on the Pt recording surface. It is important to know whether a chemical is electrochemically active in vitro prior to its local application near the Pt recording sites during experimentation. This ensures that in vitro or in vivo analyte concentration changes are from the analyte of interest rather than due to local application of drugs. Upon completion of the calibration, the program is stopped, the data are saved, and the microelectrode is removed from the buffer solution and stored appropriately. Nafion coated microelectrode tips are placed in 0.05 M PBS while mPD electroplated microelectrodes are kept dry. Graphs of typical self-referencing L-glutamate calibrations with either Nafion or mPD applied as exclusion layers are shown in Figure 19.6b and Figure 19.6c, respectively.

Microelectrode Calibration Criteria

During calibration, the FAST-16 recording software automatically calculates selectivity ratios for L-glutamate over AA as well as the slope (microelectrode sensitivity for L-glutamate), limit of detection (LOD), and linearity (R²). What do all these numbers mean, and how does one know if they have a satisfactory L-glutamate microelectrode? A poor R² is seen rarely with good micro-electrodes. Since the microelectrode array fabrication procedure is highly reproducible, our research has found that L-glutamate responses are extremely linear and should result in linear regression curve fits with R² ≥0.99.

Slope or sensitivity of the microelectrode refers to how well it can measure the change in L-glutamate. The number is used to equate a change in current to a change in L-glutamate concentration. The slope also is used to calculate LODs, which are the most important criteria for determining if a microelectrode is satisfactory for use. Here LODs are used to select microelectrodes because slopes can be misleading. LOD refers to the limit of detection for the microelectrodes defined as the analyte concentration that yields an electrode response that is equivalent to three times the background noise of the recording system. This is the lowest detectable change in analyte concentration that cannot be attributed to noise. The LODs for the study’s microelectrodes range from 0.2 to 1.0 μM, depending on the recording enzyme composition and stability of the sites. The user must select microelectrodes with LODs that are lower than the response they expect to observe. Normally it is recommended using microelectrodes with LODs ≤ 1 μM.

Selectivity refers to a ratio of the microelectrodes sensitivity for L-glutamate over interferents such as AA. It is calculated by dividing the L-glutamate slope by the AA slope. A microelectrode with a selectivity of 100:1 means that a 100 μM concentration increase of AA results in an apparent 1 μM concentration increase in L-glutamate. With a selectivity of 100:1, the user knows that the microelectrode is 99% effective at blocking AA. Selectivity ratios of 100:1 or greater are ideal, but selectivity ratios of 20:1 are used and are still extremely effective because this ratio is not a linear correlation. In other words, a microelectrode with a selectivity of 50:1 is blocking approximately 98% of the AA while a microelectrode with a selectivity of 20:1 is still blocking approximately 95% of the AA signal.

Cell Culture Techniques

The culturing of cells does not change and standard culture dishes are used with the enzyme-based multisite microelectrodes. In fact the microelectrodes are capable of recording from a variety of cell culture well sizes.

Culture plates are placed on top of a heated water pad (Gaymar Industries, Inc Model #TP12E) that holds the temperature constant at 37°C and insulated with towels to help prevent heat loss from the air. An electrode manipulator on a stereotaxic frame (Kopf Instruments, Model #960) is positioned next to the culture dish, and the microelectrode connector with attached microelectrode is secured to the electrode manipulator. The glass micropipette attached to the microelectrode is filled with solution as previously described. The tubing from the Picospritzer III is laced through the electrode manipulator and secured onto the glass micropipette. Lacing the tubing through the electrode manipulator prevents breakage of the glass micropipette by an accidental tug on the tubing. The solution flow through the glass micropipette is tested with several pressure ejections before lowering into a well. If an air bubble is found, the solution is removed, and the glass micropipette is refilled and tested again. Finally, the microelectrode is lowered into a culture well with the aid of a stereomicroscope to ensure that the Pt recording sites are immersed in the culture media, without breaking the tip on the bottom of the plate. The coated tip of the Ag/AgCl reference electrode is placed into the same well and held in place with tape. Figure 19.14 shows an in vitro cell culture recording set-up used in the laboratory.

FIGURE 19.14

Photograph of an in vitro cell culture recording set-up. (a) The microelectrode and connector are attached to an electrode manipulator on a stereotaxic frame (not shown) and positioned over a culture dish. The reference wire is attached to the culture (more...)

The laboratory has used cell culture techniques with enzyme-based multisite microelectrodes to examine L-glutamate uptake in astrocyte cell cultures. By pressure ejecting a solution of 5 mM L-glutamate into the culture media, one can determine the rate of uptake of the exogenously supplied L-glutamate into astrocytes. Furthermore, by making serial dilutions of the astrocyte cultures, one can examine the efficiency of L-glutamate uptake. These studies hold great promise for a better understanding of L-glutamate uptake.

Brain Slice Techniques

Our laboratory uses the R1 microelectrode for brain slice experiments. The slice is only thick enough for 1 Pt recording site to be inserted, and the R1 configuration is best for this recording method. Because only one Pt recording site is inserted, a slightly different coating method is employed. The bottom and top sites are coated with L-glutamate oxidase while the two middle sites are left uncoated and used qualitatively for self-referencing recordings. If a micropipette is attached, the pipette is centered over the bottom recording site since this site is inserted into the brain slice.

Brain slices are harvested using standard procedures [23,24]. The artificial cerebral spinal fluid (aCSF) used consists of the following (mM): NaCl (124); KCl (5); MgCl₂ (1.5); CaCl₂ (2.5); NaH₂ PO₄ (1.4); D-glucose (10); NaHCO₃ (26). aCSF is bubbled continuously with 95% O₂ /5% CO₂. After the brain is removed, it is placed in ice cold oxygenated aCSF, blocked, and then, using a Vibratome^® Series 1000 Sectioning System (Technical Products International, Inc., Product #054018), the brain is sectioned to form coronal slices that are 350–400 μm in thickness. Brain slices containing brain regions of interest are immersed in a holding chamber filled with bubbled room temperature aCSF and allowed to acclimate for at least 1h prior to recording.

An immersion-style perfusion chamber maintains the brain slices during the recordings at a temperature of 33°C–35°C (Figure 19.15). Pre-warmed solution is perfused at a rate of 1.5–2.0 ml/min through the chamber with the aid of a peristaltic pump. Prior to placing the slice in the chamber, the coronal slices are cut along the midline and only one hemisphere is immersed in the chamber. After securing the microelectrode-connector-headstage assembly to a micromanipulator, the microelectrode is lowered at a 60° angle into the slice. Once in the slice, a stereomicroscope is used to visualize the lowering of the microelectrode such that site 1 (the site closest to the microelectrode tip) is completely in the slice (Figure 19.16a and b). Slices are allowed to acclimate and the recordings are allowed to reach a steady, level baseline prior to studies.

FIGURE 19.15

(a) Recording system for measuring L-glutamate in brain slices. (b) An immersion-style chamber helps maintain brain slices during L-glutamate measures. The slice is perfused from below and only a reference electrode and the microelectrode tip are placed (more...)

FIGURE 19.16

Microelectrode array in a brain slice. (a) Photograph of a microelectrode above a brain slice. The microelectrode is lowered at a 60° angle to provide adequate contact with the tissue. (b) An L-glutamate oxidase coated site is placed into the (more...)

Delivery of drugs is provided through superfusion of the slices or local application from glass micropipettes. L-glutamate release is often evoked by stimulation through one of two means: (1) bath application of 40 mM KCl (40 mM KCl, 109 mM NaCl, 2.5 mM CaCl₂, pH 7.4) or (2) direct, local application of 70 mM KCl (70 mM KCl, 79 mM NaCl, 2.5 mM CaCl₂, pH 7.4) via pressure ejection from glass micropipettes. Bath application of 40 mM KCl (aCSF where the KCl concentration is raised to 40 mM with an equivalent decrease in NaCl concentration) for 2 min elicits an increase in L-glutamate concentration (Figure 19.17). A 30 min recovery period is carried out before a second KCl stimulation is repeated. A comparison between the ratio of the amplitudes of the second response to that of the first response in control slices versus slices treated with drugs (e.g., transporter or presynaptic receptor modulators) serves as a useful technique for assessing the effects of these treatments.

FIGURE 19.17

Representative L-glutamate signals seen following perfusion of the brain slice with 40 mM KCl (black bars). The KCl solution depolarizes cells and evokes the release of extracellular L-glutamate.

A second method employed to stimulate release is pressure ejection of excess KCl directly into the brain slices. A glass micropipette filled with Isotonic 70 mM KCl (pH 7.4) is attached to the microelectrode as described above and pressure ejections of 70 mM KCl are used to evoke L-glutamate responses in brain areas (Figure 19.18). Both of these methods provide reliable, reproducible means of measuring L-glutamate in brain slices. The ability to measure the effects of known and highly controlled concentrations of drugs on L-glutamate signaling in the living brain slice provides a helpful compliment to L-glutamate measurements in vivo.

FIGURE 19.18

Pressure ejection induced L-glutamate release in the rat frontal cortex. Repeated ejections of 70 mM KCl (25–50 nl) produced rapid L-glutamate responses in the frontal cortex. Inset, L-glutamate release after a single pressure ejection of potassium (more...)

Anesthetized Rats and Mice

Once the microelectrode is calibrated, rodents are anesthetized with urethane (Sigma, U-2500) (1.25 g/kg, i.p.) or another suitable anesthetic such as chloral hydrate. Our laboratory began employing urethane as an anesthetic during DA studies using carbon fiber microelectrodes. Urethane, unlike other anesthetics, does not greatly affect the clearance of DA from the extracellular space [35]. Furthermore, little is known regarding the mechanism of action of urethane. Urethane is also a useful anesthetic for long-term surgeries. Other anesthetics that are used for these types of experiments, such as ketamine, are documented to affect glutamatergic function [36–38]. As the recording technology has evolved, and the understanding of the glutamatergic system has increased, the laboratory has noticed that urethane does decrease basal levels of L-glutamate (unpublished data).

Once fully anesthetized, the rodent is placed in a stereotaxic frame (Kopf Instruments). A Deltaphase^® Isothermal Pad (Braintree Scientific, Inc. Model #39DP) is placed between the metal frame and the animal to maintain its body temperature at 37°C. In the case of mice, these heating pads are bulky so a heated water pad (Gaymar Industries, Inc., TP3E) connected to a water bath (Gaymar Industries, Inc, T/Pump, TP500) is used to maintain body temperature. An incision is made on the midline of the scalp and the skin is reflected. Once the skull is exposed, a Dremel^® (local hardware retailer) with bit size 107 (rats) or 105 (mice) is used to remove a portion of the skull over the recording sites. With forceps, the overlying dura is pulled laterally to expose the surface of the brain. The dura is removed gently as it is strong enough to break the microelectrode tip if it is not reflected prior to micro-electrode insertion. Finally, a small hole is drilled in a remote location from the recording site for a Ag/AgCl reference electrode placement. The coated tip of the Ag/AgCl reference electrode is inserted into the brain and held in place using Durelon^® Carboxylate dental cement (CMA/microdialysis Catalog #030 0003801).

It is recommended that a micromanipulator (Narishige Scientific Instrument Lab, RO-10) that attaches to the microelectrode manipulator on the stereotaxic frame is used to lower or raise the microelectrode by small increments. The microelectrode attached to the connector is fitted onto the micromanipulator by means of a Lexan^® microelectrode holder designed by our lab. Once fitted to the stereotaxic frame, the glass micropipette attached to the microelectrode is filled with solution. Next, the tubing from the Picospritzer III is laced through the electrode manipulator and secured onto the glass micropipette. The flow of the solution through the glass micropipette is tested with several pressure ejections before implantation. Once properly configured, the microelectrode is lowered into the brain region of interest. The calibration parameters are recalled on the FAST-16 recording software and an experimental recording is initiated. The microelectrode equilibrates until a steady baseline is reached (usually 15–20 min) at which time the experimental study begins. Figure 19.19a and b show photographs of the microelectrode inserted into the striatum of a Fischer 344 (F344) rat and C57BL/6 mouse, respectively.

FIGURE 19.19

Photographs of the anesthetized rodent recording apparatus. (a) A close-up view of a F344 rat placed in a stereotaxic frame with a microelectrode array inserted into the striatum. (b) A far view of a C57BL/6 mouse placed in a mouse stereotaxic frame designed (more...)

This laboratory has examined the effects of aging on the glutamatergic system using the F344 rat model of aging [4]. Using the microelectrodes the differences in L-glutamate release and re-uptake in the striatum of anesthetized six, eighteen and twenty-four-month-old F344 rats have been determined. Nickell et al. [4] found significant changes in L-glutamate release and re-uptake in young versus aged rats. Figure 19.20 shows representative tracings for 70 mM KCl-evoked L-glutamate release in six and twenty-four-month old F344 rats. Six-month-old F344 rats released more L-glutamate with less stimulus compared to the aged animals (note scale bars). Furthermore, the uptake of L-glutamate was significantly decreased in the twenty-four-month-old F344 animals compared to the six and eighteen-month-old F344 animals as shown in Figure 19.21.

FIGURE 19.20

Tracings of 70 mM potassium-evoked L-glutamate release in the young, 6 month (a), and aged, 24-month (b), F344 rats. Both age groups produced robust, reproducible L-glutamate responses. However, larger responses in the six-month rats were obtained with (more...)

FIGURE 19.21

Representative traces of 70 mM potassium-evoked L-glutamate signals in 6, 18, and 24-month F344 rats. Note that the 24-month F344 rats had a significantly longer rate of L-glutamate clearance compared to the 6-month and 18-month F344 rats.

Anesthetized Rhesus Monkeys

Much of the basic science that is conducted in rodents is assumed to be applicable to human conditions; however, important biological species differences may exist that limit the relevance of experimental findings [39]. Species that are more closely related to humans, such as rhesus monkeys, represent a logical step along the pathway toward applying the technology in the clinical setting. Additionally, differences exist between working with rodents under acute recording conditions in the laboratory setting and chronically investigating monkeys in the veterinary operating room. The tasks of general animal handling, anesthesia, surgery, and recovery fall on teams of experts.

The primary concern centered on creating a mobile FAST-16 recording system that is easily moved between settings and can be closely positioned to the anesthetized monkey in the operating room without compromising the sterile field. It was sought to make the system self-contained, to move all components as a single unit, and to require minimal setup and additional equipment. As shown in Figure 19.22, the FAST-16 recording system was placed onto a standard push cart that is easily moved into and around the operating room.

FIGURE 19.22

Photograph of the mobile FAST-16 recording system used for anesthetized monkey recordings.

Similar to anesthetized rodent studies, microelectrodes are coated and calibrated as previously described. However, due to the expensive costs of these monkeys and the limited time allotted for these procedures, four microelectrodes are prepared for these experiments. Based on calibration data, the four microelectrode arrays are ranked by LOD, slope, and selectivity. The top two ranking microelectrode arrays are assigned for use with KCl induced L-glutamate release and local applications of exogenous L-glutamate, respectively. The other two microelectrodes are backups in case of accidental breakage or other unforeseen problems.

Rhesus monkey surgical procedures are handled by qualified technicians and outlined below. Following initial sedation with ketamine hydrochloride ( ~ 20 mg/kg; i.m.) plus atropine sulfate ( ~ 0.04 mg/kg; i.m.), each rhesus monkey is intubated via the orotracheal method, and intravenous lines are secured. Then, the animal is anesthetized with isoflurane (1%–3%) and placed in a magnetic resonance image-compatible stereotaxic apparatus (Kopf Instrument, Model #1430M) in a ventral–lateral position. The animal is maintained on a heated blanket and has cardiac and respiratory parameters monitored during the procedure. Coordinates for microelectrode array implantation are determined by magnetic resonance imaging prior to the surgery. After being shaved, the scalp area is cleaned with antiseptic. Starting at the incision site and moving circum-ferentially to the periphery, the shaved area is gently scrubbed with sterile 4 ″×4 ″ sponges soaked in chlorhexidine diacetate surgical scrub followed by 70% isopropyl alcohol. After the alcohol dries, Betadine^® prep is applied, and the animal is covered with sterile drapes. Then, a 5–7 cm incision is made through the scalp, and the skin and muscles are reflected to expose the skull. Small (2 m diameter) holes are drilled in the skull directly over the targeted areas using a dental drill and a rounded drill bit, and the overlying meninges are removed to expose the surface of the brain. A lateral pocket is made in the fascia at the posterior extent of the incision using blunt dissection. A 70% isopropyl alcohol-cleaned glass-bodied Ag/AgCl reference electrode (Bioanalytical Systems, Inc., RE-5b) is inserted into the pocket. Sterile saline irrigation is used to maintain ionic contact between the reference electrode and the exposed brain.

Microelectrode/micropipette assemblies are slowly lowered into each brain region using a stereotaxic holder as previously described. Figure 19.23 shows photographs of the anesthetized rhesus monkey set-up in the operating room. After completion of the recording session, bone wax is placed in the drill holes, and the scalp incision is sutured over the exposed areas as per normal procedures. The animal then is given an analgesic (buprenorphine, 0.01 mg/kg, i.m.) and prophylactic antibiotics (combination Penicillin G Benzathine and Penicillin G Procaine, 20,000 U/kg, s.c.). Temperature, heart rate, respiratory rate, and femoral pulse are monitored until the animal recovers from anesthesia. Full recovery is defined as self-sustained balance and posture.

FIGURE 19.23

Photographs of L-glutamate recordings in anesthetized rhesus monkeys. (a) A photograph of the operating room. Rhesus monkeys are anesthetized and prepared for electrochemical recordings under sterile field conditions. The mobile FAST-16 is positioned (more...)

Our laboratory has conducted several recordings in the anesthetized monkey. We have found that stimulus-evoked release of L-glutamate in the motor cortex and prefrontal cortex is similar to that seen in anesthetized rodents. Figure 19.24 shows potassium-evoked L-glutamate release in the cortex of young and aged anesthetized rhesus monkeys.

FIGURE 19.24

Rapid potassium induced release of L-glutamate in the motor cortex of young (top trace) and aged (bottom trace) rhesus monkeys. Potassium ejections at one minute intervals (black arrows) in the monkey motor cortex evoked fast L-glutamate release and uptake. (more...)

Awake, Freely Moving Rats and Mice

Our laboratory is transitioning from anesthetized animal recordings to recordings in awake freely moving animals. Recordings of L-glutamate and other neurotransmitters in awake freely moving animals have several distinct advantages. First, this approach allows for monitoring L-glutamate neurotransmission without the effects of anesthesia. Anesthetics are known to alter basal and phasic L-glutamate release dynamics [37,40,41]. Second, freely moving animal recordings can be coupled to behavioral studies to investigate changes in L-glutamate that are dynamically related to behaviors. Third, the microelectrodes can reliably record L-glutamate for over two weeks in freely moving animals. This means that multiple measures can be made over multiple days instead of only one day as in anesthetized and microdialysis studies.

Implantation and Recording Procedure

First, the microelectrode used for in vitro or anesthetized in vivo animal recordings is modified to make it smaller and lighter weight for use in chronic implantations. Refer to Rutherford et al. for a detailed description of freely moving microelectrode preparation and implantation (manuscript in preparation) [42]. The PCB holder of the enzyme-based microelectrodes is scored just beneath the first set of pinholes on the PCB (Figure 19.3c), without splitting the microelectrode into two pieces. The microelectrode is handled easily as a single unit, but the scoring allows for the top part of the PCB to be easily broken off at a later time. Since scoring produces dust that can cling to the Pt recording sites, the microelectrodes are then cleaned as previously described. Once cleaned, the scored microelectrode is coated with Nafion (it has been found this exclusion layer works longer for chronic implantations compared to mPD) and enzyme solution.

Next, microelectrode connecting wires are prepared by first stripping both ends of 30 AWG varnished copper wire (Radio Shack). One end of the copper connecting wire is soldered to a pinhole on the row closest to the microelectrode tip. The other end is attached to a gold-plated socket amphenol (Ginder Scientific Part #220-S02). Four copper wires are soldered to the PCB, each corresponding to one of the Pt recording sites. Once all wires are soldered, the microelectrode is coated with 5-min epoxy, which protects the microelectrode connection from potential water or other liquid damage during the remaining portion of the procedure. After the epoxy dries for approximately 30 min, the remaining portion of the PCB is split in two using pliers to bend the PCB at the score mark. Now, the modified microelectrode is ready for enzyme coating and allowed to cure for 48 h before final modifications.

Once the enzyme layer has properly cured to the microelectrode surface, the socket amphenol attached ends of the reference and microelectrode wires are inserted into a miniature connector (Ginder Scientific, 9-pin ABS Plug, Part #GS09PLG-220). Pliers are used to push the socket amphenols into the connector so that the socket amphenols are completely encased. A miniature Ag/AgCl reference electrode is prepared and inserted into the connector. The copper wires are wrapped around the connector and the microelectrode tip is positioned parallel to the connector. Correct positioning of the microelectrode tip is essential for accurate stereotaxic placement into the brain. Five-minute epoxy is used to secure the microelectrode array and wires, making sure that all wires and solder points are covered with epoxy. The openings on the end of the connector closest to the microelectrode tip are also covered with epoxy to ensure that moisture does not penetrate the connector and disrupt the recorded signal.

Similar to anesthetized studies, drugs are locally applied close to the Pt recording sites for pharmacological studies in freely moving animals. Instead of a glass micropipette, a 26-gauge stainless steel guide cannula (Plastics One Inc., C315G Cannula Guide 26GA 38172, PO: 052604AA, SO: 41123-1) is centered among the recording sites with sticky wax, Figure 19.25. A stainless steel dummy cannula (Plastics One Inc., C315DC Cannula dummy, PO: 121409, SO: 45203-3), slightly longer than the guide, is inserted into the guide cannula and screwed into place. The dummy cannula tip is positioned among the four recording sites of the microelectrode at a distance of approximately 100 μm above the microelectrode tip. The stainless steel dummy cannula remains in the guide at all times, except during ejection when it is replaced with a 33 gauge internal acute cannula (Plastics One Inc., C315IA Cannula Internal Acute, PO: 121409, SO: 45203-1) through which solutions are locally applied. This cannula is the same length as the dummy cannula so it is positioned in the same location. The dummy cannula helps prevent particulate matter from clogging the guide cannula.

FIGURE 19.25

Photograph of the modified microelectrode with attached guide cannula used for freely moving recordings in rats and mice. The attached guide cannula is centered among the 4 recording sites and held approximately 100 μm above the ceramic surface (more...)

The recording headstage is slightly different from those used for in vitro and in vivo anesthetized animal recordings. This headstage screws directly onto the rat microelectrode assembly connecting the chronically implanted microelectrode to the FAST-16 recording system. The head-stage consists of a miniature connector with five connector pins (one connecting each of the four channels and one for the Ag/AgCl reference electrode). The connector pins lead to the four channel mini-amplifier, which is positioned as close as possible to the animal in order to minimize noise artifacts as shown in Figure 19.26. Shielded connecting wire, leads to an electrical swivel (Airflyte Electronics Co., P/N 1001460-012) at the top of the recording chamber. The headstage is connected at the top center of the recording apparatus to the electrical swivel (commutator) that contains twelve electrical contacts as seen in Figure 19.27a. The commutator allows the animal to move freely to all areas of the behavior chamber.

FIGURE 19.26

Photographs of the freely moving rat and mouse connector. The circuit board at the center of the photograph is the smaller, lighter headstage. This is then wired to the tether and shielded (upper left). Finally, the fully assembled tether and modified (more...)

FIGURE 19.27

Photographs of an awake rat connected to the commutator and FAST-16 recording system. (a) Close-up of the commutator with the attached tether for freely moving recordings. (b) A freely moving Long Evans rat is connected to the FAST-16 recording system. (more...)

For chronic implantation surgeries, animals are anesthetized with sodium pentobarbital solution (50 mg/ml), administered in one or two intraperitoneal (i.p.) doses, and placed in a stereotaxic apparatus. Animal body temperature is maintained with an isothermal heating pad at 37°C. The animals’ eyes are coated with artificial tears (www.medivet.com, Catalog #11970) to help maintain fluids and prevent infection. All surgeries are performed in a Vertical Laminar Flow Workstation (Microzone Corp., VLF-2–4). Prior to incision, excess fur on the head is shaved and the skin directly on top of the animal’s head (between the ears and from just behind the eyes to the neck) is wiped with an iodine solution to keep the incision area clean and prevent infection. The skin on top of the head is reflected, making as small an incision as necessary. The rat undergoes a craniotomy to remove a 2 mm×2 mm portion of the skull for rat microelectrode implantation. Additionally, four small holes ( < 0.5 mm) are drilled in the skull with Dremel engraving cutting bit (#105) for placement of the reference electrode and three stainless steel skull screws (Small Parts Inc., Part #MPX-080-02-M) that help hold the assembly in place. Similar to the anesthetized studies, a Ag/AgCl reference electrode is implanted remotely from the recording site. The assembly is secured with approximately four layers of Ortho-Jet Powder acrylic resin (Lang Dental Manufacturing, Co., Inc., Reference No. 1330) mixed with Jet Acrylic Liquid (Lang Dental Manufacturing Co., Inc., Reference #1406), which the skull screws help anchor in place. The dental acrylic covers as much of the microelectrode PCB holder as possible, without adhering to the threads of the cannula or the microelectrode connector. The dental acrylic has a smooth texture so as not to promote the rodent to scratch the implanted microelectrode. Figure 19.28a is a photograph of a Long Evans rat with an implanted microelectrode held in place with dental acrylic.

FIGURE 19.28

Photographs of a Longs Evan rat (a) and a c57BL/6 mouse (b) with implanted microelectrode arrays held in place with dental acrylic. Note that the animal in (a) is connected to the miniature recording headstage seen in Figure 19.26.

Following surgery, rats are placed on a heating pad to help maintain body temperature until the animal recovers from the anesthesia. Three subcutaneous injections of 1 ml each of Ringer’s Solution, Mammal (Fisher Scientific, Catalog #S77939) are administered subcutaneously immediately following surgery. Animals are given three days to recover prior to the start of a recording session.

Also our laboratory is adapting this technology for use in the freely moving mouse. The smaller size and thinner skull of the mouse makes implantation more difficult than in the rat. Although the basic microelectrode preparation remains similar, the implantation is slightly different. Mice are anesthetized with a diluted dose of the sodium pentobarbital (5 mg/ml in 0.9% physiological saline, pH~ 7.4). Once again, a 2 mm×2 mm craniotomy is performed for microelectrode implantation. Next, three holes are drilled in the skull, two for the skull screws placed diagonal from one another and the third for a Ag/AgCl reference placed diagonal from the craniotomy. Because the skull of the mouse is considerably thinner than the rat, the skull screws are cut in half with wire cutters prior to implantation. Again, the assembly is held in place with the Hygenic cold cure denture resin and Jet acrylic liquid. Because the brain of the mouse does not have the same volume of the rat, the microelectrode tip is not lowered to the same depths in rats [43,44]. The assembly rests higher on the head of the mouse than the rat; therefore, additional layers of dental resin are needed to secure the assembly. Figure 19.28b shows a photograph of a C57BL/6 mouse chronically implanted with an L-glutamate microelectrode. Despite the current size of the microelectrode, the mouse is not hindered by the additional weight.

On experiment days, the animal is allowed to explore the behavior chamber for 15–20 min before being connected to the FAST-16 recording system. Usually, the first day of recordings involves locally applying 5 mM L-glutamate (in 0.9% physiological saline, pH ~ 7.4) to determine the microelectrode response before behavioral studies are conducted. Locally applied solutions are filtered and then delivered into the internal guide cannula with the aid of 28-gauge tubing (Small Parts Inc.) fitted over a Hamilton 10 μl microsyringe to provide accurate volume ejections of locally-applied exogenous 5 mM L-glutamate. The microsyringe and tubing are taped to the tether so it can freely rotate with the rat. The dummy cannula is removed, and the internal cannula is inserted in its place. The animals should not be restrained to insert the cannula as restraining the rat alters behavioral studies by stressing the rodent.

The freely moving rodent recording system allows the effects of behavior on L-glutamate neurotransmission to be examined. In particular, different types of stressors to elicit L-glutamate signals have been used successfully. Rats exposed to a 5 min tail pinch (a clothes pin around the base of the tail) exhibit novel L-glutamate responses from the striatum and the prefrontal cortex. Figure 19.29 shows a representative tracing of an L-glutamate signal from a 5 min tail pinch in the striatum of a F344 rat. A typical tail pinch produces a bi-modal response in the striatum of a F344 rat—a fast L-glutamate spike with an elevated plateau. The spike occurs immediately at the start of the tail pinch while the plateau lasts for the duration of the tail pinch stressor, followed by a slow return to baseline. The clothespin does not hurt the rodent, but the rodent normally gnaws and/or flips around in the recording chamber in an attempt to remove the clothes pin.

FIGURE 19.29

Representative tracing of an L-glutamate response elicited by a five minute tail pinch stress in the freely moving F344 rat. The duration of the tail pinch is denoted by the solid black line under the trace.

Additionally, predatory stressors [45,46] have been used to evoke L-glutamate release in both rats and mice. Fox urine applied to a cotton ball and introduced into the recording chamber for 5 min produces increases in L-glutamate release in the mouse striatum. Figure 19.30 shows a typical L-glutamate signal from a 5 min fox urine exposure in the right striatum of a c57BL/6 mouse. The L-glutamate concentration steadily increases throughout the duration of the odor stressor. Introduction of the fox urine soaked cotton ball also causes the rodent to move to a far corner of the recording chamber at the same time as an increase in L-glutamate is measured. When the odor stressor is removed from the behavior chamber, the L-glutamate levels decrease until the response returns to baseline (approximately 15 min later).

FIGURE 19.30

Representative tracings from a typical L-glutamate signal recorded in the striatum of a freely moving c57BL/6 mouse from a fox urine stressor. The duration of the fox urine exposure is indicated by the black line under the trace.

Chronic Implantation Histopathology

Finally, the chronically implanted L-glutamate microelectrodes have been seen to produce minimal to no damage to the surrounding brain tissue in rats. Rats were chronically implanted with micro-electrodes for either two, four or eight weeks. At the end of these time points, the rats were perfused; and the brains removed, sectioned, and stained for both astrocytes (GFAP) and microglia (Iba1). Staining showed a slight increase in astrocytes immediately surrounding the implant site and minimal activated microglia (rounded microglia) surrounding the implant site for all three time periods compared to the contralateral control hemisphere. Figure 19.31 shows photomicrographs of the GFAP and Iba1 stained brain sections. Our lab is continuing to study tissue reactivity in longer term implants and to better understand the effects of extended (i.e., six month) microelectrode implants. However, there is encouragement to be found by the lack of scarring from these devices, thus, what factors (ceramic material, shape, etc) contribute to the limited tissue changes will continued to be studied.

FIGURE 19.31

( See color insert following page 272.) Immunohistochemical staining of a microelectrode recording tract of a Long Evans rat 8 weeks post implantation. The brain was sectioned and stained for GFAP (left, green) and Iba1 (right, red). The microelectrode (more...)

Telemetry

The tether attaching the freely moving rodents to the recording hardware hinders the ability to perform certain type of behaviors, such as learning tasks in a radial arm maze. Due to this limitation, our laboratory is developing a wireless transmitter for multisite recordings from freely moving rodents as well as primates. This wireless transmitter includes a smaller, lighter-weight version of the current headstage. We envision being able to chronically implant a multisite microelectrode as previously discussed. The wireless transmitter with head-stage connects to the microelectrode and is strapped to the animals back with a harness. The distance the animal moves no longer is limited by the length of the tether, but rather by the range of the wireless transmitter (currently 20 ft.). Additionally, the headstage now is closer to the multisite microelectrode, thus improving the signal-to-noise ratio recording by the microelectrodes.

Despite the fact that much of this technology is currently available, it is not yet practical for neurobiological studies. Battery life is a major limiting issue. The multisite microelectrodes are capable of recording L-glutamate in the CNS for over two weeks, and current battery life falls well short of this. In order to replace the battery the animal must be sedated every few days in order to prevent undue harm or stress. The continued work on this project has made inroads into resolving these issues and progressing towards the ultimate goal of eliminating the need for restraining cable connections.