Assaying the functional effects of demyelination and remyelination: Revisiting field potential recordings

Journal of Neuroscience Methods 182 (2009) 25–33

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Journal of Neuroscience Methods journal homepage: www.elsevier.com/locate/jneumeth

Assaying the functional effects of demyelination and remyelination: Revisiting field potential recordings Daniel K. Crawford a , Mario Mangiardi a , Seema K. Tiwari-Woodruff a,b,∗ a Multiple Sclerosis Program at UCLA, Department of Neurology, UCLA School of Medicine, Neuroscience Research Building 1, 475C, 635 Charles E Young Drive, Los Angeles, CA 90095, USA b Brain Research Institution, University of California, Los Angeles, Los Angeles, CA 90095, USA

a r t i c l e

i n f o

Article history: Received 6 April 2009 Received in revised form 12 May 2009 Accepted 20 May 2009 Keywords: Functional remyelination Brain slice electrophysiology Extracellular field potential recording Compound action potential Corpus callosum Demyelination Pathophysiology

a b s t r a c t The occurrence and histopathological characteristics of demyelination and neurodegeneration have been well described in different demyelinating mouse models. However, histopathological analysis is limiting in that it is unable to describe the functional consequences of demyelination and recovery after remyelination. Establishing the functional correlates of axon demyelination and remyelination is an important goal and can be used to measure axon function and develop neuroprotective therapies. This report describes a previously established, simple, easily applied method of electrophysiological measurement that can characterize white matter axonal dysfunction following demyelination and potential recovery after remyelination. It is designed to study in vitro stimulated compound action potentials in the corpus callosum of superfused brain slices at various time points and can be similarly used on white matter tracts in the optic nerve, spinal cord and cerebellum. Since behavioral testing can be performed prior to the brain slice electrophysiology, and the recorded slices can be post-fixed and subjected to histological analysis, correlates between behavior, axon function, and pathology can be determined. A temporal pattern of white matter functional deterioration and recovery can also be established to study mechanisms of demyelination-induced white matter injury and repair. Published by Elsevier B.V.

1. Introduction Early in the twentieth century, Erlanger et al. (1924) used a cathode-ray oscilloscope to show that a brief electrical stimulus to a peripheral nerve evoked a series of conducted electrical waves. By the mid-twentieth century, electronic equipment capable of amplifying small electrical signals had markedly improved, facilitating electrophysiological recording from myelinated and unmyelinated axon fibers. To demonstrate and explain electrical activity in nerves and muscles, numerous undergraduate physiology classes conduct a laboratory experiment in which field potential recordings of compound action potentials (CAPs) are performed (e.g., in the sciatic nerve of a frog). These laboratories represent a methodology to illustrate various characteristics of axonal conduction, such as recruitment, velocity, refractoriness, and pharmacological or physical conduction block. A stimulus pulse is given at one end of the freshly isolated nerve bundle and the combined action potential responses are then recorded at some distance away from the stimu-

∗ Corresponding author at: Multiple Sclerosis Program at UCLA, Department of Neurology, UCLA School of Medicine, Neuroscience Research Building 1, 475C, 635 Charles E Young Drive, Los Angeles, CA 90095-1769, USA. Tel.: +1 310 267 4778; fax: +1 310 206 7282. E-mail address: [email protected] (S.K. Tiwari-Woodruff). 0165-0270/$ – see front matter. Published by Elsevier B.V. doi:10.1016/j.jneumeth.2009.05.013

lus. Analysis of these CAPs yields information on the overall function of these axons. Application of this methodology to fiber tracts in the central nervous system provides us with an important tool that can be applied to the study of complex neurological disorders, including those characterized by damage or degeneration of white matter tracts (Waxman, 1980, 1988). Here, we describe a methodology for measuring CAPs in the mouse corpus callosum (CC), the large bundle of nerve fibers connecting the left and right cerebral hemispheres. These techniques have been applied in the study of traumatic brain injury in rats (Reeves et al., 2005, 2007), and in establishing the role of epidermal growth factor in myelination in mice (Aguirre et al., 2007). We recently utilized these CC CAP recording techniques to study axon demyelination induced by cuprizone diet (Crawford et al., submitted). In this model, ingestion of the cuprizone diet results in significant demyelination of the CC and arbor vitae (Blakemore, 1973, 1981; Matsushima and Morell, 2001; Skripuletz et al., 2008), along with some demyelination also occurring in the hippocampus and cortical layers (Koutsoudaki et al., 2009; Skripuletz et al., 2008). Subsequent investigation into the resulting axonal dysfunction and pathology could then provide information that may be useful for understanding demyelination-induced axon damage as it occurs in MS and would be helpful in other neurological diseases where axon dysfunction is suspected. Therefore, taken together with

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behavioral analysis, imaging, electron microscopy and immunohistochemistry methodologies utilized to study white matter tracts, the functional measurement of CC CAPs provides an essential application of an established experimental paradigm. Electrophysiological CAP analysis of axons can enhance our understanding of the timing of axon damage due to demyelination, as well as recovery of axon function during remyelination. Thus, CAP recordings represent the most sensitive assay available for analyzing the functional axon conduction deficit due to oligodendrocyte death and demyelination and the axon conduction recovery following remyelination by oligodendrocytes, stem cells, or other remyelination enhancing therapeutic agents.

drocyte glycoprotein (MOG; 1:1000, Abcam Inc., Cambridge, MA) antibodies were used to assess the fate of myelin before and after cuprizone diet and visualized with a TRITC-conjugated secondary antibody (Vector Laboratories, Burlingame, CA).

2. Materials and methods

2.5. Tissue preparation

2.1. Animals

Slice incubation unit: For proper incubation of brain slices, a slice incubation unit is either pre-made or purchased prior to the dissection. A slice incubation unit can be purchased from Digitimer Limited (Hertfordshire, England). We use a home-made slice incubation chamber with an insert to bubble carbogen as first described by Zhou et al. (1995) and adapted from the lab of Dr. Tom Otis at UCLA (Smith and Otis, 2005). We make the insert as follows: Obtain two 60 mm × 15 mm plastic Petri dishes (VWR, San Dimas, CA). Cut away the bottom sections from the lower half of each of the Petri dishes and keep the surrounding side pieces. Also, cut off and discard the lower two-thirds of a plastic 15 mL tube (Fisher Scientific, Pittsburgh, PA). On one of the Petri dish side pieces, put superglue around the entire edge circumference. Then stretch cotton gauze across this piece and place the other side piece on top. Press firmly together and let these pieces dry for at least one day. Cut away excess gauze from around the edges. Use a long, thin strip of Parafilm to wrap around the Petri dish/gauze setup and attach the cut upper portion of the 15 mL tube. The height of the cut 15 mL tube should extend a few millimeters beyond the height of the stacked Petri dishes. The insert along with the cut tube should fit snugly side by side into a 400 mL beaker and makes up the slice incubation chamber. Prior to an experiment, 200 mL cold ACSF is poured into the slice incubation chamber. The slice incubation chamber is immersed part way into a 37 ◦ C water bath. With a bubbler tip inserted into the cut 15 mL tube of the incubation chamber, carbogen gas is slowly bubbled and maintained throughout the course of the experiment.

Breeding pairs of PLP EGFP mice on the C57BL/6J background were a kind gift from Dr. Wendy Macklin (Cleveland Clinic Foundation, Ohio). The generation, characterization and genotyping of PLP EGFP transgenic mice have been reported previously (Fuss et al., 2001; Mallon et al., 2002). Mice were bred in house at the University of California, Los Angeles animal facility. All procedures were conducted in accordance with the National Institutes of Health and were approved by the Institutional Guide for the Care and Use of Laboratory Animals, UCLA. Mice were 8–10 weeks of age prior to the start of treatment. 2.2. Treatment of mice Two groups of five male mice were fed 0.2% cuprizone [oxalic bis(cyclohexylidenehydrazide)] (Sigma–Aldrich, St. Louis, MO) mixed into milled chow (Harlan Teklad 2918, Madison, WI). The diet was packed in 0.5 kg vacuum-sealed bags and stored at −80 ◦ C for long-term storage. One pouch of cuprizone pellets was brought to room temperature at least 24 h before use. Three to four pellets/mouse were placed in the cages every other day for 3 weeks to study cuprizone diet-induced demyelination (3wkCup). Following cuprizone treatment, one group of five mice was returned to a diet of normal pellet chow for 3 weeks, which allowed us to analyze remyelination (3wkCup + 3wkN). A group of five mice were untreated and were maintained on a diet of normal chow (normal). 2.3. Materials and solutions Artificial cerebrospinal fluid (ACSF) was prepared with the following ingredients (in mM): NaCl 124, KCl 5, NaH2 PO4 1.25, NaHCO3 26, MgSO4 1.3, CaCl2 2, glucose 10; adjusted to pH 7.4 (stored at 4 ◦ C, 2 weeks maximum). An hour before starting the experiment, 200 mL of pre-made ACSF is warmed to 37 ◦ C and an additional 125 mL ACSF is chilled at 4 ◦ C until ice crystals begin to form (approximately 10 min before starting). ACSF can be supplemented with various pharmacological agents, such as the sodium channel blocker tetrodotoxin (TTX, 1 ␮M, Alomone Laboratories, Jerusalem, Israel) or the potassium channel blocker 4-aminopyridine (4-AP, 10 ␮M, Sigma, St. Louis, MO), to verify and characterize the compound action potentials measured. A gelatin mixture for brain slice embedding was prepared by adding 7.5 g gelatin (BD Biosciences, San Jose, CA) and 15 g sucrose (Sigma, St. Louis, MO) to 100 mL of double distilled water. The mixture was heated to 50 ◦ C until the gelatin and sugar were dissolved; then the gelatin solution was aliquoted into 50 mL conical tubes (Fisher Scientific, Pittsburgh, PA) and stored at 4 ◦ C. Myelin basic protein (MBP; 1:1000, Chemicon, Temecula, CA), proteolipid protein (PLP; 1:750, Chemicon) or myelin oligoden-

2.4. Carbogen source Prior to starting the experiment, tubing, valves (WPI Inc., Sarasota, FL), and bubbler tips (Fisher Scientific, Pittsburgh, PA) were connected to a carbogen gas source (95% O2 and 5% CO2 gas cylinder) so one or more lines could be independently run near the bench area designated for dissection and to the electrophysiology setup.

2.5.1. Brain dissection A covered bench area next to the vibratome should be setup for dissection with the following arrangement of tools: a scalpel (number 3 with a number 10 blade), curved-blade scissors, pinch scissors, forceps, spatula (with one end bent 45◦ ), a single-edged razor blade, a cell strainer (40 ␮m nylon, BD Biosciences, San Jose, CA), cyanoacrylate glue (KrazyGlue, Columbus, OH), and a small paint brush (Loew-Cornell 1812 #2, Cincinnati, OH). Cut a 10 mL plastic pipette so that it can contain approximately 3 mL of liquid and attach a 3 mL dropper bulb (VWR, San Dimas, CA). Briefly pass the open end of the pipette through a flame to slightly melt the edges in order to remove any jagged borders. This pipette/bulb will be used to transfer the brain slices. A 100 mm × 15 mm Petri dish filled with water frozen in advance will serve as a cold surface to perform the dissection and initial gross cutting steps described below. Set up a vibrating-knife microtome (we use a Leica, model VT1000S, Wetzlar, Germany) by placing the cutting chamber into the vibratome and packing ice into the space provided around the chamber. Remove the beaker with 125 mL ice-cold ACSF and the Petri dish (cold cutting surface described above) from the freezer. Place a small piece of filter paper (Whatman Inc., Florham Park, New Jersey) onto the Petri dish and add a few milliliters of ACSF into the

D.K. Crawford et al. / Journal of Neuroscience Methods 182 (2009) 25–33

filter paper. The cold temperature of the ACSF solution and from the ice around the cutting chamber aids in decreasing protease activity in the brain and is essential for tissue health and longevity (Moyer and Brown, 1998; Watson and Pittman, 1987). A surgical dressing jar (Profex Inc., Memphis, TN) can be used as an anesthesia chamber. We anesthetize one mouse at a time with isoflurane (Baxter Healthcare Corporation, Deerfield, IL) for 15–20 s. The anesthetized mouse is promptly removed from the chamber and is decapitated with a small animal guillotine (Kent Scientific Corporation, Torrington, CT). The decapitated head is placed onto the filter paper on top of the frozen Petri dish. A scalpel is used to cut down through the midline of the scalp to separate the halves. The curved scissors are used to trim away the muscle around the skull. The pinch scissors are used to carefully cut once (less than 1 mm) into the intraparietal bone from the foramen magnum. The forceps are used to remove the intraparietal bone. The pinch scissors are used again to cut once (less than 1 mm) between the parietal bones. The forceps are then used to pry apart the left and right halves of the skull. The forceps should be used at this point to peel back the meninges surrounding the brain. The flat end of the spatula is inserted under the brain and the brain is gently lifted out of the skull and placed into the cell strainer. The brain-containing cell strainer is completely immersed into the beaker of ice-cold ACSF. The dissection starting from decapitation and extending up to this point should take less than 2 min. 2.5.2. Slice cutting Before the brain is placed in the vibratome, it needs to be blocked to access the region of interest quickly. The brain is removed from the cell strainer and placed on the frozen Petri dish which has fresh filter paper wet down with a few milliliters of ice-cold ACSF. Using a razor blade, a coronal cut is made approximately 1–2 mm from the rostral end of the brain, as well as at the fissure between the cerebrum and cerebellum. A drop of cyanoacrylate glue is placed onto the specimen disc of the vibratome cutting chamber. The brain is oriented such that the caudal end is placed down onto the drop of superglue and that the dorsal surface of the brain faces the cutting blade. The specimen disc with the brain is secured on to the vibratome and the remaining ice-cold ACSF is poured in so that the brain is completely immersed. For the Leica vibroslicer, a speed setting of 3.5 out of 10 (corresponding to approximately 0.18 mm/s) and a frequency setting of 10 (maximum setting, equivalent to 100 Hz) are used. Speed settings should be low to moderate so that the blade can safely cut through the tissue, while not moving forward too fast and pushing the tissue away. Frequency settings should be high so that the blade vibrates back and forth quickly to yield a cleaner cut. Coronal slices of 400 ␮m thickness are then cut with the vibrating-knife microtome. Slices, corresponding approximately to Plates 29–48 in the Paxinos and Franklin atlas (Fig. 1A), are transferred using the modified pipette and bulb to the incubation chamber containing oxygenated ACSF. By this time, the ACSF in the incubation chamber should be at the proper temperature and the water bath heat can be turned off. The slices are then allowed to equilibrate under these conditions for at least 1 h prior to recording.

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2.6. Electrophysiology setup There are many different ways that an electrophysiology rig can be set up. Our lab has a Faraday cage mounted onto an air table (30 deep × 48 wide, model 63-543, Technical Manufacturing Corporation, Peabody, MA). We use a submersion-type recording chamber that is mounted to the stage of an upright Olympus microscope (model BX51WI, Center Valley, PA). A manual micromanipulator (WPI Inc., Sarasota, FL) is used for the stimulating electrode and an electronic micromanipulator (model MP-225, Sutter Instrument Co., Novato, CA) is used to position the recording electrode. An amplifier (Axon Axopatch 200A Molecular Devices, Sunnyvale, CA) and a stimulus isolator (Iso-flex unit set to generate current, A.M.P.I., Jerusalem, Israel) are connected to a data acquisition system (Axon Digidata A-D converter, model 1322A, Molecular Devices, Sunnyvale, CA), which is then connected to a computer running pClamp software (Molecular Devices, Sunnyvale, CA). Recording electrodes are pulled from borosilicate glass capillaries (1.5/0.84 OD/ID, 4 long, WPI Inc., Sarasota, FL) using an electrode tip puller with a two-step protocol (model P-30, Sutter Instrument Co., Novato, CA) and back-filled with 3 M NaCl. Micropipettes with a resistance of 1–3 M are used. Ensure that no air bubbles are present in the tip of the micropipette by tapping on the side with your finger and/or immersing the tip into a warm solution. The micropipette is then placed onto the silver/silver chloride wire of the recording electrode holder. Bipolar stimulating electrodes (teflon-insulated tungsten, 1 M impedance, WPI Inc., Sarasota, FL) originally come with tips that have a sharp point. We carefully cut off approximately 0.5 mm of this tip, without separating the halves. This allows for more efficient current delivery during stimulation. This stimulating electrode is connected to the output of the stimulus isolator and inserted into approximately 3 cm of glass capillary to permit easier mounting onto the manipulator and to provide some stability since less than 1 cm of the electrode protrudes from the end of the capillary. For recording, a slice is transferred to the submersion-type chamber and perfused at a rate of approximately 2 mL/min with room temperature ACSF bubbled with carbogen gas through a gravity feed perfusion system. The stimulating electrode is lowered into the CC at approximately 1 mm lateral to midline (Fig. 1A). The recording electrode is to be placed into the contralateral CC at a distance of 2 mm away from the stimulating electrode. Each electrode is inserted up to a depth of approximately 200 ␮m below the surface of the slice. Alternatively, the stimulating electrode may be placed onto the surface of the slice to minimize damage to the CC fibers. Adjustments are made in the depths of both stimulating and recording electrodes to optimize the signal amplitude. 2.7. Measurements We use a protocol with an acquisition mode of episodic stimulation that sweeps every 5 s, with eight sweeps being run during a single trial. Samples are taken every 5 ␮s (200 kHz) over a 12-ms period of recording in a sweep. In this protocol, allocate two samples for a square wave step to 10 V from the digital out of the A-D

Fig. 1. Callosal CAP recording. (A) Coronal brain slices (400 ␮m thick), corresponding approximately to Plates 29–48 in the atlas of Paxinos and Franklin. A tungsten bipolar electrode is used for stimulation in the corpus callosum (CC) of one hemisphere and a glass electrode filled with 3 M NaCl is placed in the contralateral hemisphere for recording. (B) The last four responses, out of eight stimulation pulses, are averaged for amplitude measurement. (C) Boxed area from (B) on a larger scale. CAP amplitude is measured as the vertical distance from the local negative peak to a tangent joining preceding and following positivities (N1-myelinated axons and N2-unmyelinated axons).

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converter into the stimulus isolator, which will convert the voltage into current to be delivered into the slice for stimulation. 2.7.1. Generating the CAP Standardized input–output functions are generated for each slice in current clamp mode by varying the intensity of stimulus pulses (using the amplitude dial on the stimulus isolator) in steps from approximately threshold level to an asymptotic maximum (e.g., 0.3–4.0 mA) for the short-latency negative CAP component. Evoked callosal CAPs are then filtered (Lowpass Bessel filter set to 10 kHz), amplified (output gain ˛ = 50) and stored on disk for offline analysis. To enhance the signal-to-noise ratio, all quantitative electrophysiological analyses are conducted on waveforms that are the average of four successive data acquisition sweeps (Fig. 1B). These last four responses have the lowest baseline drift and highest consistency among the eight responses. Waveform analysis is performed using Microcal Origin (Northhampton, MA). 2.7.2. CAP amplitude determination CAP stimulation results in a large stimulus artifact (Fig. 1B) and a response waveform primarily consisting of two components: ‘N1’ representing faster depolarization from mostly large myelinated axons and ‘N2’ representing slower depolarization from non-myelinated axons. The stimulus artifact results from the generation of an electrical field which passes current directly through the buffer instead of through the slice tissue. The artifact generally lasts less than 0.2 ms and can be disregarded for the purpose of CAP analysis. The N1 and N2 components generally appear between 1–2 and 3–6 ms, respectively, following stimulation. The amplitude of the N1 and N2 components are determined at each current step by measuring the vertical distance from the local negative peak to a tangent joining preceding and following positivities (Fig. 1C). The CAP amplitude can then be graphed versus the stimulus level. Nonlinear regression analysis (sigmoidal dose response with variable slope) is performed for each CAP component, followed by statistical comparison (e.g., one-way analysis of variance using GraphPad Prism, La Jolla, CA) of the stimulus–response curves. Normalization to the highest response can also be performed, along with subsequent non-linear regression analysis, for an indication of the health and function of experimental animal axons relative to control. 2.7.3. Conduction velocity CC conduction velocity can be estimated by changing the distance between the stimulating and recording electrodes from 0.5 to 2.5 mm, while holding the stimulus intensity constant. The stimulating electrode is set in the CC at a distance of 1 mm away from midline, while the recording electrode is moved from 2.5 mm away from the stimulating electrode in the contralateral hemisphere to the closest distance of 0.5 mm in 0.5 mm steps. Recordings are performed using the protocol described above for standard CAP measurements. For analysis, the peak latency of the N1 and N2 components are measured at each point and graphed versus this distance. Linear regression analysis can then be performed for each CAP component to yield a slope that is the inverse of the velocity, followed by statistical comparison of the velocities. 2.7.4. Axon refractoriness Axon refractoriness is defined as the reduced excitability of an axon following an action potential. Axon damage can modify refractoriness through different mechanisms, including changes in the expression and distribution of ion channels involved in axonal conduction and a reduced ability to re-establish energy-dependent ionic homeostasis following an action potential. Therefore, the measurement of axon refractoriness represents a diagnostic tool to measure axon health. To quantify refractoriness, the suppression of a second CAP response in paired stimulus trials is determined as

previously described (Reeves et al., 2005). Initially, a single stimulating pulse is given at a defined strength to establish a control response (C1). Following this response, a protocol is setup to generate two pulses of equal intensity and duration which are separated by a variable time window, starting with an interpulse interval (IPI) of 8 ms and decreasing in 0.5 ms steps down to 1.5 ms. The total recording time for a sweep is adjusted to 20 ms to accommodate responses to both stimulus pulses. For analysis, the control response is subtracted from the paired stimulus responses (PSR) at each IPI, as shown in Fig. 4. This results in the response, which can be attributed to the second pulse (C2). The estimated N1 and N2 responses for the second pulse (C2) are then measured. Refractoriness is calculated for both N1 and N2 by dividing these C2 CAP component amplitudes by their respective C1 CAP component amplitudes and multiplying by 100%. The results are then graphed versus the IPI and analyzed using non-linear regression analysis, with specific use of the Boltzmann sigmoid function. The IPI that results in a 50% reduction in the CAP component is then used as a standard measure when making statistical comparisons between groups. 2.7.5. Troubleshooting Slices that have been properly incubated can have recordings performed for a period of about 4–5 h past dissection and cutting. Healthy individual slices can usually be recorded for an hour. Unhealthy slices will show a rapid decrease in CAP amplitude over time, eventually resulting in an absence of CAPs. The absence of CAPs from the start of a recording could indicate that one or both electrodes were not placed properly in the CC. If adjusting the electrodes does not yield recordable CAPs, then there could be various other reasons including damaged recording electrodes or inadequate perfusion. 2.8. Post-fixing, slice embedding and subsequent immunohistochemistry Once recordings are complete, the slices can be dropped into a 15-mL tube containing 4% paraformaldehyde (PFA) and stored for 12 h at 4 ◦ C. Following this fixation period, the PFA is removed, replaced with 30% sucrose and stored again at 4 ◦ C. Once the slices sink in the bottom of the tube, they are gelatin embedded to assure that the brain slices are flat before cutting into thinner sections. A 150 mm × 15 mm plastic Petri dish is wrapped in aluminum foil and placed with the open side down onto a bed of dry ice. In addition, a glass microscope slide (25 mm × 75 mm, VWR, San Dimas, CA) and a plastic embedding mold (22 mm × 22 mm × 20 mm deep, Peel-A-Way® , Polysciences Inc., Warrington, PA) are placed on top of cold Petri dish for 5 min. While waiting for the objects to cool, a 50-mL conical tube of solid gelatin (described in Section 2.3) is heated in a microwave for 20–30 s and placed in a 35–40 ◦ C water bath to maintain proper viscosity of the liquid gelatin. Higher gelatin temperatures may cause damage to the tissue and will cause previously embedded tissue layers to melt with a subsequent loss of section orientation in the gelatin block. Using a disposable 3 mL plastic transfer pipette (BD Labware, Franklin Lakes, NJ), place a 5-mm layer of gelatin in the bottom of the embedding mold to give adequate space between the bottom slice and the chuck when cryosectioning. Allow the gelatin layer to harden, but do not permit the gelatin to freeze. If the gelatin begins to discolor, remove the gelatin mold from the dry ice and thaw. Remove the tissue section from the sucrose solution using a brush (Loew-Cornell 1812 # 3, Cincinnati, OH) and gently dab away the excess solution using a Kimwipe (Kimberly-Clark, Roswell, GA). Flatten the section on the glass slide and “sandwich” the tissue between the aluminum foil covered dish and glass slide (on top) for 1 min. Take care not to press down on the tissue, but gently rest

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the slide with tissue side down on top of foil. Use the warmth of a finger on the underside of the glass slide to slightly unfreeze the tissue from the slide and then remove the slice with the brush. Quickly center the tissue section on top of the chilled gelatin layer in a specific orientation relative to the embedding mold (e.g., the dorsal end of the slice placed close to the notched side of the embedding mold). Apply a thin layer of gelatin (three to four drops using the disposable pipette) over the tissue section and let it cool on dry ice for roughly 1 min. Once the gelatin has solidified, make another layer of gelatin (∼3 mm) on top of the embedded tissue section and repeat the previous step to embed the next slice. After all of the tissue sections are embedded (usually four sections to a block), the mold is set at 4 ◦ C for 1 h. The sides of the embedding mold are now cut away using a single-edged razor blade and the gelatin block is carefully removed from the mold. The gelatin block may separate at certain points corresponding to the prepared layers. Addition of a few drops of warm gelatin and additional storage at 4 ◦ C may ‘glue’ the pieces back together. The block can then be placed into a 50-mL conical tube for post-fixation in PFA for 24 h at 4 ◦ C. Before placing the block into the tube, 1–2 mm of gelatin may be trimmed from the sides of the gelatin block using a single-edged razor blade. This allows the block to fit inside the tube better and float freely in the PFA. Following this fixation period, the PFA is removed, replaced with 30% sucrose and stored again at 4 ◦ C to cryoprotect the slices. For sectioning, the orientation of the block must be tracked since the slices will not be visible within the block past this point. This can be accomplished by cutting a single corner off of the gelatin block. The trimmed gelatin block is now placed onto a sheet of aluminum foil on dry ice for 30 min to allow the entire block to freeze. The block and aluminum foil are then placed in the cryostat for 1.5–2 h to adjust to the temperature inside. The gelatin block can now be mounted onto the chuck using O.C.T. compound (TissueTek, Sakura Finetek, Torrance, CA) and sectioned further. Slight adjustments in chuck orientation will be needed for each section within a block for optimal slice appearance. These re-cut slices are then processed for immunohistochemical analysis as described previously (Tiwari-Woodruff et al., 2007). In addition a few animals can be directly perfused with glutaraldehyde or PFA for ultrastructure electron microscopy analysis and immunohistochemical analysis (Tiwari-Woodruff et al., 2006). These can be compared to sections that were recorded and then post-fixed. 3. Results 3.1. Electrophysiological analysis Compound action potentials (CAPs) were recorded in brain slices corresponding to Plates 29–48 in the atlas of Paxinos and Franklin

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(Fig. 1A). The typical evoked CAP consists of three or more negative peaks. The stimulus artifact is the first negative peak (Fig. 1B). The remaining peaks represent evoked conduction that is dependent on axon myelination and diameter. The evoked CAP amplitude is quantified as a peak-to-peak measurement between the stable second negative peak and the positive one preceding it. ‘N1’ represents fast depolarization from mostly large myelinated axons and ‘N2’ represents slower depolarization from non-myelinated axons (Fig. 1C). Additional peaks may also be observed, but are not present in all slices (Swanson et al., 1998). During initial recording sessions, we found that the shortlatency component, N1 in the biphasic callosal CAP was obscured by the stimulus artifact when the ACSF was near physiological temperature (35–37 ◦ C) as previously confirmed by Reeves et al. (2005). When the recordings were performed with the ACSF at room temperature (21–23 ◦ C), conduction was slowed enough to allow separation of the N1 component from the stimulus artifact. All subsequent recordings in our lab have been performed with the ACSF at room temperature. Cuprizone diet for 3 weeks (3wkCup) completely abolished N1 conduction, suggestive of significant CC demyelination (p < 0.001, Fig. 2A and B). This lack of N1 component could also reflect asynchrony in CAP conduction, thereby effectively eliminating N1 CAP conduction. Cuprizone diet also reduced the amplitude of the N2 component and produced a delay in peak latency suggestive of slower conduction relative to normal (p < 0.001, Fig. 2A and B). These findings suggest that the CC was significantly demyelinated during the 3 weeks of cuprizone diet. The N2 amplitude reduction and latency delay also suggest that the demyelination process may have directly or indirectly caused damage to unmyelinated CC axons. When cuprizone diet animals were switched to normal diet (3wkCup + 3wkN), N1 returned to approximately 75% of normal levels (*p < 0.5, **p < 0.001, Fig. 2A and B). N2 amplitude recovered to about 70% of normal and peak latency returned to normal (* p < 0.05, Fig. 2A and C). These results suggest that although remyelination was initiated following cuprizone-induced demyelination, recovery to normal levels was not completely possible. Conduction velocity measurement of myelinated CC axons in normal mice was 1.67 ± 0.21 m s−1 , whereas non-myelinated axon conduction was 0.54 ± 0.02 m s−1 (Fig. 3). Cuprizone diet for 3 weeks eliminated myelinated N1 conduction and the conduction velocity of the N2 component was decreased to 0.37 ± 0.12 m s−1 . Switching to a normal diet in 3wkCup + 3wkN produced recovery of both N1 and N2 components, with conduction velocities of 1.72 ± 0.36 m s−1 and 0.53 ± 0.04 m s−1 , respectively. To assess the status of axon health, axon refractoriness was measured. As per protocol, the individual CAP component amplitudes elicited by the second pulse (C2) in paired stimulation (PSR) were divided by the respective CAP component amplitude resulting from single pulse stimulation (C1) and this value was plotted (Fig. 4B).

Fig. 2. Cuprizone diet induced demyelination and subsequent normal diet remyelination. (A) Typical CC CAPs from male PLP EGFP C57Bl/6 mice evoked at the highest stimulus level (4 mA) from normal, 3-week Cuprizone fed (3wkCup), and 3-week Cuprizone fed switched to normal diet for 3 weeks (3wkCup + 3wkN). (B, C) Quantification of the average stimulus–response from each animal reveals that 3wkCup decreased both N1 and N2 amplitude compared to normal mice and that switching to normal diet promotes remyelination and recovery. (n = 4–7, p < 0.001 by one-way analysis of variance).

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Fig. 3. Callosal conduction velocity. Representative CC tracings from (A) Normal (black), (B) 3wkCup (gray), and (C) 3wkCup + 3wkN (blue) mice. The recordings at different distances (0.5–2.5 mm) and the measurement of peak latency allow for an estimation of the conduction velocity for both N1 and N2 components (n = 4).

The IPI that results in a 50% reduction in the CAP component was then measured for both N1 and N2. This value was 3.4 ± 0.2 ms for N1 and 3.5 ± 0.2 ms for N2 in normal mice. The absence of N1 and significant reduction in N2 prevented meaningful determination of axon refractoriness in 3wkCup

mice. Therefore, we focused on axon health in the remyelination paradigm. Here, the N1 component was readily measurable in 3wkCup + 3wkN mice, but exhibited a slower refractoriness than normal at 4.1 ± 0.2 ms. The 3wkCup + 3wkN N2 refractoriness value of 3.2 ± 0.2 ms was not significantly different than normal.

Fig. 4. Callosal axon refractoriness. (A) Equation for calculating refractoriness. The response to a control stimulus (C1) is subtracted from the paired stimulus response (PSR) at each interpulse interval (IPI). This results in the response, which can be attributed to the second pulse (C2). The estimated N1 and N2 responses for the second pulse (C2) are then measured. Refractoriness is calculated for both N1 and N2 by dividing these C2 CAP component amplitudes by their respective C1 CAP component amplitudes and multiplying by 100%. (B) Plots of the IPI versus the C2/C1 ratio for Normal and 3wkCup + 3wkN mice. The C2/C1 ratio calculation shown in (A) is performed at each IPI. C2/C1 ratios were fitted to Boltzmann sigmoid curves. Rightward shifts in these curves correspond to increases in the refractory recovery cycle in callosal axons and are indicative of axonal damage (n = 3–6).

D.K. Crawford et al. / Journal of Neuroscience Methods 182 (2009) 25–33

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Fig. 5. Immunohistochemistry of brain slices after electrophysiology analysis. Recorded brain slices from 8-week-old normal, 3wkCup, and 3wkCup + 3wkN PLP EGFP C57Bl/6 male mice were post-fixed, embedded, re-cut and processed for immunohistochemical analysis for PLP EGFP+ green oligodendrocytes, or myelin proteins MBP (red), PLP (red), or MOG (red). Slices from 3wkCup show a substantial decrease in PLP EGFP+ cells as well as for all the myelin proteins. A substantial recovery of PLP EGFP+ cell numbers as well as myelin proteins was observed in slices from 3wkCup + 3wkN animals. Representative EM photographs of CC are shown at 4800× magnification. Cuprizone diet induced demyelination in the 3wkCup group had many demyelinated axons (absence of myelin around majority of axons). In contrast, many more myelinated axons were observed in the 3wkCup + 3wkN group similar to normal animals. Scale bar = 1 ␮m.

Collectively, the CAP amplitude, velocity and refractoriness results suggest that cuprizone diet-induced demyelination causes axon conduction changes that are partially reversible with normal diet-induced remyelination. 3.2. Immunohistochemical and electron microscopy analysis Recorded brain slices were either post-fixed in PFA, embedded, re-cut and processed for immunohistochemical analysis or post-fixed in glutaraldehyde, re-cut and processed for EM analysis. Because these mice have enhanced green fluorescent protein expressing under a proteolipid protein promoter (PLP), green oligodendrocytes scattered all over CC can be visualized under fluorescent microscope. When the sections are immunostained with antibodies against myelin proteins MBP, PLP or MOG antibodies, CC white matter myelinated axons can be visualized (Fig. 5). Cuprizone diet for 3 weeks caused a substantial decrease in PLP EGFP+ cell numbers and in the immunostaining of different myelin proteins. Switching back to normal diet for 3 weeks (3wkCup + 3wkN mice) induced a substantial recovery in PLP EGFP+ oligodendrocyte numbers and recovery in myelin staining to near normal levels. To confirm the complete recovery achieved in myelin staining via immunohistochemical analysis, which contrasts with the partial recovery observed with electrophysiology CAP recording in the 3wkCup + 3wkN group, we performed electron microscopy (EM). Images at 4800× magnification revealed normal callosal axons with thick, compact myelin layers (Fig. 5). A marked decrease (∼50%) in the number of myelinated axons was observed for 3wkCup, which recovered to near normal levels in 3wkCup + 3wkN. A more detailed

analysis of the ultrastructure of CC axons by EM after cuprizone diet has been previously done by others and more recently by our group (Stidworthy et al., 2003); reviewed in Crawford et al. (submitted) and Matsushima and Morell (2001). It is important to note that subtle changes in axon function (shown by CAP recordings) cannot be assessed by immunohistochemical studies. Detailed ultrastructure analysis via EM and functional electrophysiology analysis may be more useful to ascertain true changes in myelin and axon integrity. We show here a disconnect between the functional outcome and immunohistochemical protein expression during remyelination. 4. Discussion The myelin sheath provides protection and insulation of axons, as well as increases the rate of nerve impulse transmission. Demyelination impairs axon conduction and can lead to deficits in motor, sensory and/or cognitive functions depending on the location of the affected axons. Demyelinating diseases can occur due to genetic abnormalities (leukodystrophies), infectious reactions (Epstein-Barr virus) or by autoimmune responses (multiple sclerosis). In recent years, various genetically dysmyelinating (e.g., quaking, jimpy and shiverer) and inducible demyelinating (e.g., cuprizone-diet, lysolecithin, Theiler’s murine encephalomyelitis, and experimental autoimmune encephalomyelitis) mouse models have enhanced our knowledge of the importance of myelination of axons. Demyelination and remyelination have been assessed by decreases in various myelin proteins by RT-PCR (changes in RNA levels), by Western analysis (changes in protein levels), and/or by

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visualization under the microscope using histology, immunohistochemistry and ultrastructure analysis by electron microscopy. However, these assays are limiting, as they are unable to measure the consequences of demyelination and remyelination in the context of axon function. In fact, myelination during development (myelination), disease (demyelination) or during repair (remyelination) should be analyzed in the context of its functional effect on axon conduction. To this end, CAP recording of axons as described in this paper fills the void left by other experimental methodologies and can provide functional information that would be otherwise unattainable. In vivo analysis of interhemispheric antidromic latencies in the rhesus monkey cortex and a decrease in callosal axon conduction of lysolecithin injected rabbits, were performed by the Waxman group nearly 30 years ago (Foster et al., 1980; Swadlow et al., 1978). Recently, these techniques have been used to measure changes in CC axon function during traumatic brain injury in rodents (Baker et al., 2002; Reeves et al., 2005). The present report revisits this electrophysiological recording technique while investigating the effect of cuprizone diet-induced demyelination and subsequent normal diet-induced remyelination on callosal axon conduction. It is designed to study in vitro stimulated CAP recordings in the CC of superfused brain slices following cuprizone diet and can demonstrate a temporal pattern of white matter functional deterioration or recovery during demyelination or remyelination. Similar to previous results (Baker et al., 2002; Preston et al., 1983), two CAP components from distinct populations of faster conducting myelinated fibers (N1) and slower unmyelinated fibers (N2) were observed in normal control group. The CAP recordings at room temperature allow the separation of stimulus artifact from the fast conducting myelinating N1 component that is lost if the recordings are performed at 35–37 ◦ C (Baker et al., 2002; Reeves et al., 2005). Cuprizone diet induced a significant decrease in the myelinated CAP component. This decrease in CAP amplitude of cuprizone diet group may reflect alterations to callosal axons such as pathological depolarization or reduced numbers of responsive axons due to demyelination-induced damage. Some of the damage persists, as the CAP amplitudes of mice switched to normal diet do not show complete recovery in the myelinated N1 component. The slower CAP component, N2 also showed a significant decrease in amplitude during cuprizone diet which recovered in groups that were switched back to normal diet. A more detailed effect of cuprizone diet on axon conduction was analyzed by measuring axon velocity and refractoriness measurements. Conduction velocity is the rate at which a nerve impulse is propagated. The myelin status, axon diameter, distribution of ion channels at nodal regions and temperature all affect conduction velocity. In our experiments, cuprizone diet-induced demyelination in the CC eliminated the N1 component. These demyelinated fibers likely conduct similar to the non-myelinated axons. In addition, cuprizone diet also produced a delay in peak latency for nonmyelinated conduction (N2). This delay in peak latency is suggestive of axon damage and could result from adaptive changes in ion channel distribution. Similarly, changes have been seen in antidromic response latency in cortical neurons after cuprizone diet have also been seen (Bando et al., 2008). This delay could also occur through the loss of a sub-population of non-myelinated fibers of smaller diameter that conduct slightly faster than non-myelinated fibers of larger diameter. Switching cuprizone-fed mice back to a normal diet produced a complete recovery of axon conduction velocity, a result which is more consistent with changes in ion channel distribution altering peak latency than the loss of a CC fiber sub-population, resulting from the cuprizone diet-induced demyelination. A more detailed analysis of axon damage can be assessed by measuring axon refractoriness in cuprizone-fed mice. Refractoriness, long recognized to depend largely on recovery of Na+ channels

from inactivation after an action potential (Hodgkin and Huxley, 1952) is sensitive to changes in membrane potential which could occur due to demyelination-induced Na+ channel reorganization that has been observed in EAE and MS (Craner et al., 2004, 2005). Cuprizone diet-induced changes in refractoriness are consistent with this altered sodium channel function and persisting axonal depolarization after injury. This depolarizing shift in membrane potential increases the extent of Na+ channel inactivation in normal diet-induced remyelinated callosal axons which exhibit a persistently reduced capability for firing repeated action potentials, denoting permanent deleterious changes in axon that have undergone demyelination in the presence of cuprizone. Previous changes in refractoriness of unmyelinated axons due to traumatic brain injury were observed by Reeves et al. (2005). Our results indicate for the first time demyelination induced axon function changes by refractoriness analysis. The electrophysiological measured deficits were investigated by immunohistochemical analysis of the recorded slices. Thus, immunostained recorded slices allow us to correlate functional and immunohistochemical data directly. Along with the precision and care taken throughout dissection and slice sectioning, incubation and recording of slices in carbogen gas-bubbled ACSF seems to preserve tissue integrity throughout the course of the electrophysiology experiments. After fixation, cryoprotection, and re-cutting, immunostained recorded slices did not show significant differences from perfusion fixed slices. Myelin immunohistochemistry with anti-MOG antibody was analyzed in combination with PLP EGFP OL/myelin fluorescence. Similar to previous reports, a significant decrease in OL numbers and myelin proteins was observed in the cuprizone diet group. A recovery of OL numbers and myelin density to normal control levels was observed in the group of animals switched to normal diet similar to previously reported (Lindner et al., 2008; Matsushima and Morell, 2001; Torkildsen et al., 2008). Taken together with the present functional CAP results, we demonstrate that callosal axons may have significant remyelination, but the new myelin is unable to restore callosal axon conduction to normal levels. The CAPs can be recorded from nearly all white matter tracts in the brain or spinal cord and the results are consistent and reproducible. Best of all, the same recorded slices can be subjected to immunohistochemical analysis. This CAP recording methodology is easily applied and can be used to assay changes in axon function during developmental myelination, disease and during repair strategies (e.g., therapeutic drug treatment or stem cell therapy) in various transgenic and disease rodent models.

Acknowledgements The authors wish to acknowledge their deep appreciation to Xiaoyu Xia and Dr. Héctor López-Valdés for their valuable suggestions.

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