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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 19Second-by-Second Measures of L-Glutamate and Other Neurotransmitters Using Enzyme-Based Microelectrode Arrays

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L-Glutamate is the major excitatory neurotransmitter in the mammalian central nervous system (CNS) and is implicated in a number of brain disorders including Parkinson’s disease (PD), cognitive disturbances, epilepsy, schizophrenia, attention deficit hyperactivity disorder (ADHD) and drug abuse. While microdialysis methods have been used extensively over the last decade to investigate minute-by-minute measures of L-glutamate, the rapid time dynamics of L-glutamate signaling in the CNS has warranted a technique to measure L-glutamate release on a second-by- second basis. A major goal of the research is to develop a recording technology for recording second-by-second measurements of L-glutamate and other neurotransmitters—specifically a mass-fabricated microelectrode technology that could be (1) mass produced such that other laboratories could utilize the same recording technology and (2) configured for “self-referencing” recordings, which allows for second-by-second cross-checking of the selectivity of the micro-electrode measures and improved signal-to-noise of the recording methods. The present chapter documents current capabilities of measuring L-glutamate and several other neurotransmitters on a second-by-second basis using mass-fabricated microelectrode arrays formed on ceramic. While not a comprehensive assessment of the technology, this chapter contains a large amount of information regarding the fabrication, use, and potential pitfalls of this technology. The reader should refer to numerous articles [1–6] for additional details regarding measuring neurotransmitters in the CNS.

Principles of in Vivo Electrochemistry

In vivo electrochemistry utilizes microelectrodes that can be implanted into the mammalian CNS, thus providing a means to record chemical signaling of neurons. The microelectrode’s recording surface, or working electrode (normally an inert metal such as platinum (Pt) or carbon), can oxidize or reduce compounds of interest. A potential is applied versus a reference electrode, normally a Ag/AgCl reference electrode that is in ionic contact with the working microelectrode. Low noise headstages, which include potentiostats, apply this potential and are computer-controlled with multiple inputs to simultaneously record from several microelectrodes or several recording surfaces on a single microelectrode array. If the potential at the microelectrode surface is sufficient, then molecules directly at the recording surface of the working electrode are either oxidized or reduced depending upon their intrinsic electrochemical properties. Oxidized molecules give up one or more electrons to the recording surface while reduced molecules receive electrons from the recording surface. The currents generated from those Faradaic reactions are linear with respect to concentration of the electroactive molecule(s) in the tissue surrounding the microelectrode. This basic principle allows for in vitro calibration methods for in vivo studies. (For a more complete overview, please see Gerhardt and Burmeister’s chapter in the Encyclopedia of Analytical Chemistry.)

One of the simplest electrochemical techniques is amperometry, which involves the measurement of current at a constant fixed potential. The current can be monitored continuously, thus, events can be measured as quickly as ≤1 ms [6]. Because the voltage is applied continuously to the working electrode, the non-Faradaic background current recorded from the microelectrode is low allowing for sensitive measurements of electrochemically active molecules [6,7].

Several neurochemicals reside in the extracellular space of the CNS that are electrochemically active on Pt recording surfaces at low or high oxidation potentials including ascorbic acid (AA), dopamine (DA), norepinephrine (NE), serotonin (5-HT), 5-hydroxyindoleacetic acid (5-HIAA), homovanillic acid (HVA), nitric oxide, uric acid, and 3,4-dihydroxyphenylacetic acid (DOPAC). Additionally, enzymes are applied to the recording surface to produce an electrochemically active reporter molecule (usually hydrogen peroxide) to allow measurements of molecules that are not inherently electrochemically active. When a potential of +0.7 V versus a Ag/AgCl reference is applied to the Pt recording sites, the newly formed peroxide oxidizes, and the resulting change in current from the transfer of electrons to the Pt recording surface is detected. While O2 is required as a cofactor for oxidase enzymes, the oxidation of the reporter molecule, hydrogen peroxide, generates a portion of the O2 for continued reactions. This, however, does not allow measures using oxidase enzymes in zero O2 environments.

An important and often poorly considered aspect of microelectrodes is that analytes from only a small area (microns) of tissue surrounding the recording surface are detected. This aspect is useful for studying small or layered structures in the brain or the spinal cord. Additionally, micro-electrodes with multiple recordings surfaces can be geometrically arranged to measure analyte concentrations from two or more distinct brain regions. Since microelectrodes coupled with amperometric techniques can measure analytes of interest on rapid time scales (1–1000 ms), uptake and release kinetics of neurotransmitters are easily studied.

Enzyme-Based Multisite Microelectrode Arrays


Enzyme-based multisite microelectrode arrays are mass fabricated using photolithographic methods [8–11]. An advantage of photolithography allows routine production of reproducible recording surfaces as small as 5–10 μm. In addition to this advantage, multiple microelectrodes are patterned onto a single fabrication substrate (usually 2.5 cm×2.5 cm) allowing for increased fabrication number at a decreased cost. Finally, photolithographic methods are used to manufacture numerous microelectrode designs with multiple recordings sites in well-defined, highly reproducible geometrical configurations.

The multisite microelectrodes used in the research are currently constructed in conjunction with Thin Films Technology, Inc. (Buellton, California). The fabrication process is explained thoroughly in previously published papers [1,2,11–13]. Initially, microelectrode photographic masks are designed on a computer aided design (CAD) program where arrays of 4–16 recording sites are arranged on templates. A 2.5×2.5 cm × 125 m thick ceramic wafer (alumina, Al2 O3, Coors Ceramic, Coors Superstrate 996) serves as a common substrate for the microelectrode arrays. Ceramic reduces the cross-talk from adjacent connecting lines. Additionally, ceramic is strong and rigid, which aids in precise stereotaxic placement into tissues without flexing or breaking. The ceramic substrate may be polished or lapped down to achieve microelectrodes as thin as 37.5 μm [12].

Following cleaning, photoresist is spun onto the ceramic wafer. Collimated light passing through the photomask exposes the photoresist, thus transferring the micro-electrode images onto the wafer. Spaces for the recording sites, connecting lines and bonding pads are not exposed to light. Solvents are used to remove unexposed photoresist from the wafer. Many 1-cm long microelectrodes (34–64) can be patterned onto the wafer simultaneously to facilitate production and to increase the number of microelectrodes that are made from the ceramic substrate (Figure 19.1). Patterning also can be performed on the reverse side of the ceramic wafer to increase recording site density.

FIGURE 19.1. Photograph of a ceramic wafer containing 34 microelectrodes with Pt bonding pads, connecting lines and recording sites (not visible) are patterned onto the 2.


Photograph of a ceramic wafer containing 34 microelectrodes with Pt bonding pads, connecting lines and recording sites (not visible) are patterned onto the 2.5 cm× 2.5 cm, 125 μm thick substrate.

Next, an adhesion layer of Titanium (Ti) is used to allow Noble metals (Pt or Ir) to adhere to the ceramic substrate. Titanium (500 A ¢ª thick) is sputtered onto the developed photoresist covered ceramic wafer. Following the adhesion layer, the active recording metal layer, Pt for most of the work, is sputtered onto the substrate ( ~ 1.5 μm thick). Solvents are used to remove the developed photoresist and unwanted metals leaving the recording pads, connecting lines, and connecting pads.

Once the Pt recording sites, connecting lines, and bonding pads are in place, the connecting lines are coated with an insulator using another photolithographic step. The connecting lines act as wires to connect the bonding pads to the recording sites and must be insulated from aqueous environments. To accomplish this, the microelectrodes are coated again with photoresist. A second photomask is used to define the areas where polyimide is to be placed. Polyimide acts as an insulator to define the recording site active area as well as the bonding pads. It also reduces cross talk between the connecting lines. After the photoresist is developed, polyimide is spun onto the wafer (2–4 μm thick). Once the insulating layer is applied, only the recording sites and bonding pads are exposed. The photoresist and excess polyimide are removed and the remaining polyimide is cured at 200°C. The complete fabrication process is depicted in Figure 19.2.

FIGURE 19.2. Schematic of the fabrication sequence for the ceramic wafers with Pt recording sites.


Schematic of the fabrication sequence for the ceramic wafers with Pt recording sites.

After the formation of the microelectrode on the ceramic wafer, a diamond saw or laser is used to form or “cut out” the individual microelectrodes. A major advantage of the diamond saw is that it produces highly polished edges for reduced tissue damage during implantation. The ceramic wafer with Pt recording sites is attached to a printed circuit board (PCB) holder for handling and connection to recording equipment. To connect the ceramic wafer to the PCB holder, each bonding pad is wire bonded to an individual Pt recording site on the ceramic wafer. The tips are epoxied onto the paddle for stability and to insulate the wire bonds. Cutting and assembly is performed currently in conjunction with Hybrid Circuits, Inc. (Sunnyvale, CA). The PCB holder and fully constructed multisite microelectrode are shown in Figure 19.3a and b, respectively. Figure 19.3c is a version of the microelectrode for awake animal recordings (see the section in this chapter on Awake, Freely-Moving Rats and Mice for further details).

FIGURE 19.3. Photograph of the microelectrode PCB with and without attached ceramic microelectrode tip.


Photograph of the microelectrode PCB with and without attached ceramic microelectrode tip. (a) Close-up view of the PCB. Numbers beneath the pinholes indicate the specific Pt recording site connection. (b) Fully assembled microelectrode array. Black epoxy (more...)

Multisite Microelectrode Array Designs

Due to the flexibility of the fabrication process, the microelectrodes can be manufactured with different recording site geometric configurations. For this research, the most commonly used configurations have four Pt recording sites. However, microelectrodes with eight recording sites are in use and those for sixteen recording sites are being developed as discussed in later sections. Figure 19.4 shows two popular four site configurations designated as the R1 (a row of four in-line sites, Figure 19.4a) and S2 (two side-by-side site pairs, Figure 19.4b). Individual microelectrodes are selected based on the type of recordings and brain regions of interest. The R1s provide a larger recording distance that is useful for large brain regions or layered structures, while the S2s provide dual detection in smaller brain structures. Furthermore, the advantage of multiple recording sites allows for self-referencing techniques (see the section on Amperometric Recordings Utilizing Self-Referencing) and/or detection of multiple analytes. Additional designs are presented in Gerhardt and Burmeister’s work in the Encyclopedia of Sensors.

FIGURE 19.4. Photomicrographs of two ceramic-based multisite microelectrode designs.


Photomicrographs of two ceramic-based multisite microelectrode designs. Each has four Pt recording sites arranged in different geometrical configurations. (a) Photomicrograph of an R1 microelectrode with a row of four Pt recording sites. Each site measures (more...)

Headstage /Recording System

A headstage/potentiostat creates a potential difference between the microelectrode array and a reference electrode to carryout electrochemical recordings. The recording system used also amplifies the electrochemical signal and is part of the Fast Analytical Sensing Technology (FAST-16) system (Quanteon L.L.C, Nicholasville, KY). Figure 19.5 shows a photograph of the current FAST-16 recording system. In conjunction with other hardware elements, including the control box and a 16 bit A/D card in the computer, the FAST-16 system amplifies the electrical current generated by oxidation or reduction reactions and digitizes this current. The Windows™ based FAST-16 system software records, creates and displays second-by-second (1 Hz), or faster (1–40 Hz), data files (nominal 1–10 Hz). The software is written for eight simultaneous channel recordings, which can be increased as the number of Pt recording sites increases in future generations of microelectrodes. Recorded files are exported to other Windows™ based applications, such as Excel™, for easier data processing; or processed directly through the data analysis software package on the FAST-16 system software. The low noise design and software oversampling allow the FAST-16 recording system to be used for bench top experimentation without the need of a Faraday cage. However, greater performance can be achieved with the use of a Faraday cage or appropriate shielding for most applications

FIGURE 19.5. Photograph of the control box (bottom), headstage (right) and A/D board (left) encompassing the Fast Analytical Sensing Technology (FAST-16).


Photograph of the control box (bottom), headstage (right) and A/D board (left) encompassing the Fast Analytical Sensing Technology (FAST-16). The A/D board has a sampling rate of 1.25 million samples per second (MS/s). This value refers to the speed at (more...)

Microelectrode Preparation

Many CNS neurotransmitters are not inherently electrochemically active and therefore must be converted to a reporter molecule by cross-linking enzymes to our microelectrode surface. Below are the general procedures involved in preparing a microelectrode for use in vitro or in vivo.

Cleaning Procedures

Microelectrodes received from Hybrid Circuits, Inc., undergo a cleaning procedure to remove any particulate matter or residue from the Pt recording sites that may have been deposited during the manufacturing process. New microelectrodes are placed into a stirred solution of Fisherbrand Citrisolv™ (Fisher Scientific, Catalog #22-143–975) for 5 min followed by a 5-min rinse in ddH2 O. After the chemical cleaning procedure, a thin film can develop on the Pt recording sites, so a Kimwipe® (Kimberly-Clark Professional) is swiped delicately across the tip of the micro-electrodes to remove this film and any excess ddH2 O. Then microelectrodes are allowed to dry for 15 min in an oven at 105°C–115°C prior to coating.

Typically, enzyme-based microelectrodes are reused for more than one experiment, provided they meet calibration criteria. Over prolonged use, applied enzyme layers (described below) may degrade analyte detection. A slightly different cleaning procedure is employed for microelectrodes that have been previously coated with an enzyme layer. This enzyme layer must first be removed prior to re-use. The microelectrode tip is placed into a stirred solution of ddH2 O at 80°C to soften the protein matrix on the Pt recording sites. Following 30 min of soaking, the microelectrodes are cleaned as described in the preceding paragraph and tested for response to hydrogen peroxide to ensure good sensitivity of the microelectrode recording surfaces.

Exclusion Layer Coatings

Exclusion layer coatings are applied after successful cleaning of the microelectrode and are used to alter the recording properties of the microelectrode arrays. Exclusion layers are applied to block or minimize undesirable electrochemically active compounds found in high concentrations in the CNS such as AA (250–500 μM) or DOPAC. By blocking these undesired electrochemically active molecules from the Pt recording sites, the microelectrode has better selectivity for the analyte of interest. The planar geometry of the multisite microelectrodes often affects how materials adhere to the recording surfaces. It should be noted that exclusion and enzyme layer coatings can inhibit compounds from diffusing to the microelectrode surface by creating a diffusion barrier, thus limiting the response time of the microelectrode. For this reason, coating parametrics are performed to determine optimal coating procedures for both exclusion layer and enzyme layer coatings.

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.


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.

Enzyme Layer Coatings

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

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.


Our laboratory has observed that the micro-fabricated microelectrode arrays have highly reproducible Pt recording surfaces. However, manufacturing procedures only investigate the geometric surface area of the microelectrodes. Since each Pt recording site on a microelectrode can respond differently to peroxide and L-glutamate, they must be calibrated in vitro prior to experimentation to determine standard curves. Thus, calibrations are used to equate a change in current from the oxidation of peroxide to a proportional change in analyte concentration from the L-glutamate generating the peroxide. A known concentration of analyte is added to 40 ml of 0.05 M PBS solution at 37°C to generate a current in picoamperes (pA) that is measured by the FAST-16 system. The FAST-16 system software records the current for each addition of analyte, creates a calibration curve for each Pt recording site, and stores the slope of this calibration. Also, known interferents such as AA are added during the calibration to test the selectivity of the recording sites to the analyte of interest versus interferents. When the experiment is being performed, the calibration data are recalled and used to determine the concentration of analyte being measured from the change in current during experiments in vivo.

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.


(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.

Glass Micropipettes for Local Drug Delivery

After calibrations are completed, a glass micropipette often is attached to the PCB holder so the tip lies among the Pt recording sites. Through this glass micropipette, drug solutions are pressure ejected for local pharmacological manipulations of the CNS. Our laboratory uses a vertical pipette puller (David Kopf Instruments, Model 700C, Catalog #730) to prepare single barrel glass (1 mm o.d., 0.58 mm i.d., A-M Systems, Inc., Catalog #601500) micropipettes. Using a dissecting microscope, the tip of the micropipette is bumped against a glass stir rod to create an internal diameter of 12–15 μm. Next, modeling clay is attached to the PCB holder. The glass micropipette is attached to the modeling clay and used to hold the glass micropipette steady while it is positioned between the recording sites. Once the glass micropipette is properly positioned, it is held more firmly in place by applying melted Sticky Wax (Kerr Corporation, Catalog #00625) onto the PCB between the ceramic wafer and the modeling clay. The drying of the Sticky Wax may reposition the glass micropipette relative to its placement amongst and above the Pt recording sites. If necessary, the tip of a spatula can be heated to help remodel the Sticky Wax and thus reposition the glass micropipette. Figure 19.7a shows a glass micropipette attached to the PCB with modeling clay and Sticky Wax. Figure 19.7b and Figure 19.7c show the tip of a glass micropipette positioned within the recording sites for an R1 and S2 microelectrode, respectively. The tip of the glass micropipette must not touch the ceramic surface. Instead it should be positioned between 50 and 100 μm above the Pt recording sites (as shown in Figure 19.7d). This distance allows an optimum amount of CNS tissue between the glass micropipette and microelectrode when it is lowered into the brain. Additionally, if the microelectrode is too close to the Pt recording sites, pressure ejection of fluid causes a dilution effect in vivo (an actual decrease in analyte concentration due to the introduction of exogenous fluid in the area of the Pt recording site). If positioned too far, the rapid uptake of L-glutamate into glia limits its diffusion to the Pt recording sites on the microelectrode surface. To properly position the glass micropipette on the microelectrode, the microelectrode must be viewed at two angles underneath a dissecting microscope. Our lab has designed a special holder for this application consisting of two glass microscope slides glued perpendicular to one another with an eight pin DIP socket glued onto one of the flat surfaces (Figure 19.8). Additionally, double and triple barrel glass micropipettes (World Precision Instruments Catalog #2B150F-6, and #3B120F-6, respectively) can be used to locally apply two or three different drug solutions into the same brain region. These are harder to utilize because the adjoining micropipettes prevent tubing from easily sliding over the openings of the glass micropipette. Instead, smaller tubing (Small Parts, Inc., PTFE Tube Regular Wall, 28-gauge, Catalog #SWTT-28) is wedged into the open end of each micropipette, which attaches to the tubing from a pressure ejection system such as a Picospritzer® III (Parker Hannifan, Corp., General Valve Operation).

FIGURE 19.7. Photographs of a glass micropipette attached to a multisite microelectrode.


Photographs of a glass micropipette attached to a multisite microelectrode. (a) Photograph of the glass micropipette held in place with modeling clay and Sticky Wax. (b) Photomicrograph of a glass micropipette tip positioned between sites 2 and 3 on an (more...)

FIGURE 19.8. Photograph of the microelectrode holder used for attaching glass micropipettes to the micro-electrode arrays.


Photograph of the microelectrode holder used for attaching glass micropipettes to the micro-electrode arrays. The microscope slides fit any standard dissection microscope and allows for easier placement and viewing of the micropipette at both angles. (more...)

Attaching the glass micropipette is probably one of the most difficult tasks of an experiment and takes practice to properly achieve. Practicing is also ideal for success of experiments because positioning the glass micropipette on a microelectrode must occur within a short time period for optimum microelectrode performance.

The glass micropipette is filled with solution using a 30-gauge, 4-in. long hypodermic needle (Popper and Sons, Inc., Standard Female Luer Hub, Catalog #7400). Glass micropipettes always are filled prior to implantation. First, drug solutions are drawn into a 1 ml tuberculin tip syringe (Beckton-Dickinson, and Co., Catalog #309602), which is then attached to the intake end of a μStar 0.22 μm sterile filter (Costar Corp, Catalog #8110). The other end of the filter attaches to a 4-in., 30-gauge, hypodermic needle. The solution is dispensed through the filter and ultimately through the hypodermic needle. The needle is inserted into the glass micropipette and filled from the pulled tip up to the open end making sure not to create air bubbles within the glass micropipette. Air bubbles prevent effective delivery of drugs by pressure ejection and make it difficult to determine how much fluid is dispensed. Solutions are normally pressure ejected prior to micro-electrode implantation into the brain to determine that there is a smooth fluid flow out of the glass micropipette. Solutions are removed from a glass micropipette by inserting the 30-gauge filling needle with an attached tuberculin syringe and drawing up the solution.

Ag/AgCl Reference Electrode

While larger glass Ag/AgCl reference electrodes are suitable for mPD electropolymerization and microelectrode calibration, this reference electrode is too large for many biological applications. For this reason, a smaller Ag/AgCl reference electrode must be made at the start of every experiment. Miniature Ag/AgCl reference electrodes are prepared by first stripping 0.25 in. of the Teflon coating from each end of a silver wire (0.008 in. bare, 0.011 in. coated; A-M Systems, Inc., Catalog #786500). One of the stripped ends is soldered to a wire crimp pin (Mill-Max Mfg. Corp., Part #3603) for connection to the FAST-16 headstage. The other end is placed into a solution of 1 M HCl saturated with NaCl. A Pt wire acts as the counter electrode and is also placed into the solution. The laboratory routinely uses a 9V DC adapter fitted with alligator clips on both the positive and negative leads. However, DC adapters ranging from 1.5 to 9 V can be used. The negative lead is attached to the Pt counter electrode while the positive lead is connected to the Ag wire. With both wires in solution, the applied potential attracts Cl on the wire to form AgCl thus making the Ag/AgCl reference. The plating potential is applied for approximately 10–15 min. When properly connected, bubbles are seen around the Pt counter electrode. These miniature Ag/AgCl references are soaked in 3 M NaCl prior to use in vivo.

Signal Analysis

These are the basic steps necessary for successfully measuring L-glutamate in the CNS using our multisite microelectrode arrays. The steps for setting up different experiments using this technology will be further discussed later in this chapter. Before there is an explanation of the experimental protocol, there must be an examination of the features of a typical, rapid L-glutamate signal and how it is analyzed.

The Picospritzer III or related hardware connects to the FAST-16 recording system, so when solutions are pressure ejected through the glass micropipette, or experimental events occur, the software records these events. The FAST-16 recording system automatically saves amperometric or chronoamperometric data, time, and pressure ejections marks (external events) for all recording sites for a specified time period. Figure 19.9 shows a typical L-glutamate signal with some of the analysis terms. A stereomicroscope fitted with a reticule monitors the change in solution in the glass micropipette after pressure ejection. Pressure ejection of 70 mM KCl (70 mM KCl, 79 mM NaCl, 2.5 mM CaCl2, pH 7.4) leads to a depolarization event and release of L-glutamate into the extra-cellular space, causing a robust and reproducible rise in the L-glutamate signal compared to baseline. The maximum change in concentration compared to baseline is referred to as maximum amplitude or amplitude of the L-glutamate signal. This value is normally recorded in μM units. Next, the maximum amplitude of the signal is divided by the amount of stimulus (i.e. volume ejected from the glass micropipette) necessary to elicit the depolarization event. The stimulus ejected is visualized via a stereomicroscope [28] mounted on a universal boom stand (Meiji EMZ Zoom, Labtek, Inc.) fitted with a reticule. Volume displacement is calibrated according to the pipette inner diameter. (Note: the single barrel glass micropipettes used have a solution volume of approximately 250 nl/mm of glass, but this varies with the inner diameter/total volume of the glass micropipette. For this reason, one should use the type of glass micropipettes from A-M Systems, Inc., Catalog #601500 as discussed in the section on glass micropipettes for local drug delivery.) The volumes of stimulus ejected are recorded in nanoliters, so this calculation is referred to as amplitude per nanoliter ( μM/nl) and gives an interpretation of the excitability of the neurons. Finally, the time period for the signal to reach the maximum amplitude from baseline is recorded (in s) and referred to as rise time (Tr ).

FIGURE 19.9. A schematic diagram of an L-glutamate signal is shown to illustrate how different signal parameters are measured and calculated.


A schematic diagram of an L-glutamate signal is shown to illustrate how different signal parameters are measured and calculated. See text for details. Typically L-glutamate or other neurotransmitter signals are plotted as concentration versus time. The (more...)

Signal decay refers to the point of maximum amplitude of the signal to its return to baseline. The clearance of L-glutamate is a result of the rapid uptake of L-glutamate into glial and neuronal transporters [29]. L-Glutamate uptake follows an apparent first order decay rate. The decay of the L-glutamate signal is fitted to the slope of the linear regression of the natural log transformation of the data over time and this is referred to as k−1 and has units of s−1. When k−1 and the maximum amplitude of the signal are multiplied, the uptake rate, or micromolar of L-glutamate removed from the extracellular space per second ( μM/s), is obtained. Finally, the decay of the signal can be examined at different percentages of decay. These often include T50, T80, and T100, and follow the general format of TD (s), where T refers to the time from maximum amplitude for the signal to decay by D percent. Typically T50 and T80 decay times are used.

Amperometric Recordings Utilizing Self-Referencing Microelectrodes

Importance of Self-Referencing Microelectrode Arrays

An advantage of photolithographic fabrication of microelectrode designs is the precise control of the layout of the microelectrodes allowing for multiple recording sites geometrically configured in a small area. Multiple recording sites on a single microelectrode can be used to remove chemical interferents that may contribute to a portion of the analyte signal [1]. Our laboratory refers to this principle as self-referencing, and it is commonly used in the data analysis. As mentioned in the coating section, the L-glutamate oxidase is applied to a pair of Pt recording sites while the chemically inactive protein matrix (BSA and glutaraldehyde) is applied to the adjacent pair of recording sites referred to as “sentinel sites”. A schematic of the coating procedure is shown in Figure 19.10. The L-glutamate oxidase coated Pt recording sites detect extracellular L-glutamate concentrations (the analyte) in addition to molecules that the inactive protein matrix coated recording sites can detect (interferents). These additional molecules detected can include interferents such as AA and DOPAC as well as other neurotransmitters such as DA, NE, 5-HT. Figure 19.6b and Figure 19.6c show a typical L-glutamate self-referencing calibration where the L-glutamate oxidase coated sites respond to L-glutamate while the inactive protein matrix coated sites do not.

FIGURE 19.10. Schematic and photomicrograph of an S2, Nafion coated, self-referencing, L-glutamate selective multisite microelectrode.

FIGURE 19.10

Schematic and photomicrograph of an S2, Nafion coated, self-referencing, L-glutamate selective multisite microelectrode. (a) Schematic showing the bottom pair of Pt recording sites coated with L-glutamate oxidase (Glu-ox), BSA, and glutaraldehyde (Glut). (more...)

The purpose of the inactive protein matrix on the self-referencing, or sentinel sites, is to make the response times of the sentinel sites similar to the L-glutamate oxidase recording sites. As mentioned earlier, protein coats can alter the diffusional properties of molecules to the Pt recording surfaces. Without this inactive protein matrix coat, the sentinel sites would respond faster to interferents than the L-glutamate oxidase sites. Protein layers on both pairs of Pt recordings sites are necessary to minimize differences between the temporal recording properties of the different microelectrode surfaces. Ideally, the sentinel vs. analyte recording sites should be positioned side-by- side. However, the current, manual enzyme coating procedures preclude this arrangement.

Experimentally, self-referencing recordings are beneficial for providing qualitative as well as quantitative assessment of the analyte of interest. During an experiment, the FAST-16 recording software allows the user to simultaneously view changes in L-glutamate concentration for all four recording sites. If a peak is seen on the L-glutamate oxidase coated sites, but not the inactive protein matrix coated sites, this provides a good qualitative assessment that L-glutamate is being measured. Figure 19.11 shows a screen capture of an L-glutamate self-referencing microelectrode during an in vivo experiment.

FIGURE 19.11. Screen capture of the FAST-16 system with a self-referencing L-glutamate microelectrode measuring potassium-evoked L-glutamate release in the motor cortex of a rhesus monkey.

FIGURE 19.11

Screen capture of the FAST-16 system with a self-referencing L-glutamate microelectrode measuring potassium-evoked L-glutamate release in the motor cortex of a rhesus monkey. Each line corresponds to a different Pt recording site. Sites 1 and 2 are coated (more...)

Quantitatively, the self-referencing method is even more powerful. Since some of the response on the L-glutamate oxidase coated sites is attributable to interferents the sentinel sites are used to record the interferents contributing to the L-glutamate response. If the recording sites’ measure of Faradaic responses to DA and H2 O2 are close ( < 10%), the raw current data collected for a sentinel recording site is subtracted from the raw current obtained from the L-glutamate oxidase coated sites. This new raw current, once divided by the slope of the corresponding L-glutamate oxidase recording site, provides the adjusted concentration of L-glutamate that is free of interferents. If the recording sites’ calibration non-Faradaic currents and Faradaic response to DA and H2 O2 are not close (> 10%), then a normalization procedure is used to correct for recording site coating inconsistencies by factoring in the DA and/or the peroxide responses during calibration [1].

Besides being able to remove interferents that contribute to an analyte signal, self-referencing recordings can remove periodic or random noise [2]. This is an obvious advantage because smaller changes in current (lower detection limits) can be achieved. Figure 19.12 illustrates removal of noise using self-referencing recordings. The noisy upper trace is a response from an L-glutamate oxidase coated recording site, while the lower trace is the resulting sentinel site response. When the sentinel site raw current is subtracted from the L-glutamate raw current and divided by the slope of the L-glutamate oxidase coated site, the signal-to-noise ratio of the peaks is greatly improved. This dual-channel noise subtraction approach is widely used in other methodologies such as electro-physiology and spectroscopy and can now be used with the precise microelectrode arrays.

FIGURE 19.12. Pressure ejection of 70 mM KCl (black arrowheads) causes release of L-glutamate in the striatum of a C57BL/6 mouse.

FIGURE 19.12

Pressure ejection of 70 mM KCl (black arrowheads) causes release of L-glutamate in the striatum of a C57BL/6 mouse. Non-Faradaic current decreases the signal-to-noise ratio making it difficult to distinguish between current fluctuations and low L-glutamate (more...)

Phasic Release of Neurotransmitters

Depolarization of neurons using increased KCl as a stimulus is a common method for in vivo analysis of neurotransmitter release [3,4,24–26,30–33]. While a stimulus does depolarize neurons, nerve fibers, and nerve terminals it does not selectively release neurotransmitters. On the contrary, the solution causes release of a multitude of neurotransmitters. These additional neurotransmitters released into the extracellular space have the potential to contribute to a portion of the analyte response. Luckily, few neurotransmitters are electrochemically active (DA, NE, and 5-HT) without having to be converted to a reporter molecule. DA and 5-HT usually exist in much higher levels than NE in the CNS and therefore are of greater concern for contributing to a portion of the KCl-evoked L-glutamate response. Using the self-referencing technique, contributions from DA, NE, and 5-HT to the L-glutamate response are easily removed.

Interestingly, the present studies support that while release of L-glutamate and DA are relatively fast and can occur simultaneously with applications of KCl, DA has a much slower uptake process compared to L-glutamate [2,3]. This slower uptake gives DA a very distinguishable response signal compared to L-glutamate in vivo. The decay of the DA signal may last minutes while L-glutamate decay takes only seconds. This DA response coincides with the L-glutamate response on the L-glutamate oxidase coated sites, and gives a very distinguishable response that is referred to as a spike-dome response. The spike corresponds to the quick release and re-uptake of L-glutamate. However, as this L-glutamate response begins to decay, DA release and diffusion continues and its much slower uptake creates a dome response. Essentially, the DA response contributes to a portion of the L-glutamate response making analysis difficult.

As mentioned before, Nafion coated microelectrodes can actually attract DA, NE, and 5-HT to the Pt recording sites and mPD coatings may not always block the DA, NE, and 5-HT responses. Therefore, self-referencing recordings are crucial for removing interferent responses from the analyte response. Since interferent responses occur on the sentinel and L-glutamate oxidase coated sites, the raw current from the sentinel site can be subtracted from the L-glutamate oxidase coated sites. This generates the true L-glutamate response without interferents contributing to the signal. Figure 19.13 shows a typical spike-dome response in a mouse striatum from an L-glutamate oxidase coated site, the DA response on the sentinel site, and the subtracted L-glutamate response.

FIGURE 19.13. Stimulus evoked L-glutamate release sometimes results in measuring both L-glutamate (the fast spike) and DA (the slow dome) on the L-glutamate oxidase coated Pt recording sites.

FIGURE 19.13

Stimulus evoked L-glutamate release sometimes results in measuring both L-glutamate (the fast spike) and DA (the slow dome) on the L-glutamate oxidase coated Pt recording sites. The sentinel Pt recording sites only measure the DA contribution to the response. (more...)

Tonic or Basal Release of Neurotransmitters

The photolithographic fabrication methods used allow the multiple recording sites to be patterned in close proximity to one another. This provides a novel opportunity to measure basal levels of neurotransmitters. Basal measures refer to the low levels of neurotransmitters constantly found in the extracellular space that are important for CNS homeostasis. High in vivo concentrations of interferents, such as AA may contribute to a portion of the L-glutamate basal responses. Once again, self-referencing is utilized to subtract off the portion of the interferent signal that contributes to the L-glutamate signal thus providing basal L-glutamate measurements.

Furthermore, basal L-glutamate regulation is studied using pharmacological methods. Local applications of Tetrodotoxin (TTX), a sodium ion channel blocker, or DL-threo-beta-benzyloxyas-partate (TBOA), an excitatory amino acid uptake inhibitor, are utilized to study their effects on basal L-glutamate concentrations. The laboratory has demonstrated successfully that local application of TTX decreases basal L-glutamate while TBOA increases basal L-glutamate on the L-glutamate oxidase coated microelectrodes [5]. Sentinel sites do not show changes in their basal recordings after administration of either drug. In this scenario, a self-referencing microelectrode qualitatively validates the effects of pharmacological manipulations. This assessment is made directly after local application of the drug and is easily seen using the FAST-16 recording system. Once again, self-referencing can quantitatively determine the change in basal L-glutamate since the “normalized” raw currents from the sentinel sites are subtracted from the raw currents of the L-glutamate oxidase coated sites to obtain changes in basal L-glutamate. This is a unique recording feature of the microelectrode arrays that cannot be carried out using single microelectrode recordings.

The principle of self-referencing holds great promise for the future of in vivo electrochemical recordings. Self-referencing involves a control recording site introduced in close proximity to the analyte recording sites. This technique allows the user to qualitatively assess the validity of an analyte signal during an experiment. Additionally, the ability to subtract off interferents and noise during off-line data analysis allows the user to obtain accurate and detect smaller concentrations of analytes

Application to Biological Systems

In Vitro Systems

In vitro cell culture techniques provide scientists with simpler, more controllable conditions for studying isolated systems. The multisite microelectrodes are small enough for basic cell culture studies as well as brain slice measures. Below the experimental procedures and techniques used for both are discussed.

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.

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.

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.

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).

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.

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...)

In Vivo Anesthetized Animal Recordings

The ability to perform in vivo recordings in anesthetized animals with enzyme-based multisite microelectrodes is a hallmark of this technology. The laboratory has recorded successfully L-glutamate (as well as other neurotransmitters including L-lactate, acetylcholine and choline) from the brains of rats, mice, and monkeys as well as L-glutamate in the rat spinal cord. Outlined below is the procedure for preparing an animal for in vivo brain recordings. Spinal cord recordings follow a similar procedure except a laminectomy is performed rather than a craniotomy.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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...)

Future Directions

Eight Site Microelectrode Arrays

Most of the applications discussed center around the use of enzyme-based microelectrodes with four Pt recording surfaces. Photolithography allows the design of microelectrodes with even more Pt recording sites. Now there has been the fabrication of microelectrodes that contain eight Pt recording sites, and those with sixteen Pt recording sites are being developed. Furthermore, a more flexible multisite microelectrode for recording from deep brain structures in monkeys and humans is being tested. These new electrode designs can further the understanding of L-glutamate and other neurotransmitters in the CNS.

The eight site microelectrodes coupled with the new FAST-16 recording system can simultaneously record amperometric data from eight sites on a second-by-second basis. Figure 19.32 shows photomicrographs of three current eight site microelectrode designs. There are two advantages of the increased number of Pt recording sites: (1) multiple analytes can be measured from a single microelectrode in the same brain region; and (2) multisite microelectrodes can be designed specifically to simultaneously record from several brain regions.

FIGURE 19.32. Photomicrographs of three, eight site microelectrode arrays.

FIGURE 19.32

Photomicrographs of three, eight site microelectrode arrays. Pt recording site dimensions are labeled at the bottom left of each photomicrograph while the dimensions between groups of recording sites are labeled between the hash marks.

The long term purpose of designing these multisite microelectrodes is to simultaneously record multiple neurotransmitters in the same brain region. The eight site microelectrodes allow the user to coat a single microelectrode as if it were two microelectrodes. For example, on the W2 and W3 designs the bottom set of Pt recording sites are configured for L-glutamate with self-referencing measures. The top set of Pt recording sites is coated with a different enzyme/protein matrix also configured for self-referencing recordings. This allows for simultaneous recording of both L-glutamate and other neurotransmitters and observing any possible interactions during a depolarization or behavioral event. With future designs that incorporate sixteen Pt recording sites on the same microelectrode, four or more analytes can be simultaneously measured. Furthermore, with the development of a microcoater to facilitate more precise enzyme coating, side-by-side pairs of Pt recording sites can be configured for self-referencing recordings. This allows measurements of four different analytes on an eight site microelectrode array and eight different analytes on a 16 site microelectrode.

The second advantage of eight Pt recording sites per array is that the Pt recording sites can be geometrically positioned to encompass two separate anatomical brain structures. For example, 600 μm spacing between sets of Pt recordings sites on the W2 enzyme-based microelectrode is the average distance between the rat CA1 and CA3 cell regions in the rat hippocampus. The bottom set of Pt recordings sites rests in the CA3 regions while the top set of Pt recording sites can reside in the CA1 region. By configuring both sets of Pt recordings sites for L-glutamate/self-referencing recordings, neurons in the CA3 region can be depolarized and the subsequent L-glutamate signaling to the CA1 region can be recorded.

Using Microelectrode Microdrives for Chronic Implantations

A microdrive (Figure 19.33) developed at Wake Forest University (courtesy of Dr. Sam Dead-wyler) allows one to replace and lower microelectrodes for freely moving animal studies. Freely moving rodent craniotomies are performed as described earlier in the chapter. However, here a base plate is attached to the skull. The microelectrode fits into the microdrive; this assembly attaches to the base plate. Once connected to the base plate, the microelectrode is lowered to the desired depth. This microdrive allows one to measure multiple brain regions within the same animal by raising or lowering the microelectrode as desired. Additionally, if a microelectrode loses sensitivity for L-glutamate, the microelectrode can be replaced allowing for longer recording capability in a single animal. This has applicability for studying L-glutamate dynamics throughout the lifespan of aging or disease animals.

FIGURE 19.33. Photograph of a microelectrode inserted into a microdrive.

FIGURE 19.33

Photograph of a microelectrode inserted into a microdrive. The microdrive is used to reposition a microelectrode array once chronically implanted in a freely moving rodent. The microdrive unit is attached to the base plate, which is attached to the rodent (more...)


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.

Deep Brain Structure Enzyme-Based Microelectrodes

A version of the enzyme-based multisite microelectrodes has been designed for recording from deep brain structures such as the hippocampus and temporal lobe in monkeys as well as humans. This new microelectrode called the Spencer-Gerhardt electrode is modeled after the Spencer electrode (AD-TECH®) currently used for monitoring electrophysiological signals in humans. The advantage of this design increases the flexibility of the microelectrode so it can be positioned and attached to the skull with little concern for tissue damage as the brain moves. This version of the microelectrode is being utilized in the nonhuman primate testing before being used in human clinical trials. Future designs will incorporate both components of the two electrodes so that a single electrode can simultaneously monitor electrophysiological recordings of multiple single neurons and L-glutamate dynamics. This application has important benefits for helping neurosurgeons excise brain tissues in patients with epilepsy.


Special thanks to Ingrid Strömberg for her help with the sectioning and staining of brains from chronically implanted rats. Special thanks to AD-TECH Medical (David Putz) and Dr. Dennis Spencer (Yale University) for their help with designing new microelectrode arrays for nonhuman primate and human studies. In addition, special thanks to Drs. Sam Deadwyler and Robert Hampson

(Wake Forest University) for the design of the microdrive system for recordings in awake animals. Finally, we thank Andrew Allen for her dedication to these on-going projects as well as Heather Davis for her help with the photographs.

None of this work would be possible without generous support from the following agencies:

DARPA: DAMD17-99-1-9480

NIDA: 5 R01 DA017186

NIA: 2 P01 AG013494

NINDS: 1P50 NS39787

NSF: DBI-0352848

Appendix A Preparing Glutamate Microelectrodes

  1. Cleaning (first time use):
    • 5 min in Citrisolv (constant stirring)
    • 5–10 min in filtered (0.22 μm) DI water (constant stirring)
    • Dry at low temp (105–115°C) oven ( ~ 15–20 min)
      • (When reusing electrodes, stir 30 min in 80°C filtered DI water and continue as above)
  2. Nafion coating: (This step is skipped if electropolymerizing with mPD)
    • Use fresh Nafion weekly
      • (Store in fridge~ 1ml aliquots in glass container)
    • Use Nafion at room temp; Insert tip (more than half-way), swirl 5 times, pull out
    • Keep level (horizontal) and bake 4 min @ 165–175°C
      • (This time will vary according to humidity levels in your area i.e. more humid = more time)
      • ***Do not heat microelectrodes above 175°C.
  3. Enzyme coating:
  4. BSA (Sigma A-3059), Glutaraldehyde (Sigma G-5882), Glutamate oxidase (Glu-ox)
  5. (Seikagaku America, 100645)
  6. (Dissolve Glu-Ox @ 1U/μl in filtered DI water)
  7. 3.1 Bring all chemicals to room temp before use
  8. 3.2 Weigh 0.01 g BSA in 1.5 ml microcentrifuge tube
    • – Add 985 μl of filtered DI water, DO NOT vortex vigorously
    • – Add 5 μl of glutaraldehyde
    • (Final concentrations: BSA; 1%, glut; 0.125%)
  9. 3.3 Transfer 9 μl of BSA/Glut into a 500 μl microcentrifuge tube
    • – Add 1 μl of Glu-Ox
    • (Final concentration: 2%)
  10. 3.4 Apply enzyme solution using a 10 μl Hamilton syringe with a dome tip needle.
    • – Draw up~ 1 μl of solution
    • – Create a bead (drop) of solution at the tip of the syringe
    • – Under a dissecting stereoscope, apply bead on surface of the electrode, wait 2–3 s, pull tip away
    • – Let dry for a minute, typically 3 applications are enough
    • 3.5 Set aside in a clean dry place
      • – Room temp for at least 48–72 h—Time required for full cross-linking of the protein to the surface of the electrode.
  11. mPD electropolymerization 1,3 phenylinediamine, dihydrochloride 99% (Sigma-Aldrich, Cat #235903)
    • 4.1 Degas 100 ml of 0.05 M PBS with nitrogen for 20 min
    • 4.2 Dissolve 0.0905 g mPD in 100 ml 0.05 M PBS (5 mM)
    • 4.3 Add~ 40 ml of 5 mM mPD to a 50 ml beaker
      • – Lower microelectrode tip into 5 mM mPD solution
      • – Place glass Ag/AgCl reference electrode into beaker
      • – Apply a potential of + 0.5 V vs. Ag/AgCl reference for 15 min
      • – Remove microelectrode from 5 mM mPD and rinse tip with filtered, DI H2 O.
      • – Cure at room temperature for 24 h prior to calibration.


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Copyright © 2007, Taylor & Francis Group, LLC.
Bookshelf ID: NBK2567PMID: 21204381


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