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Frostig RD, editor. In Vivo Optical Imaging of Brain Function. 2nd edition. Boca Raton (FL): CRC Press/Taylor & Francis; 2009.

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In Vivo Optical Imaging of Brain Function. 2nd edition.

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Chapter 11Optical Imaging of Neuronal Activity in the Cerebellar Cortex Using Neutral Red

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Over the last decade our laboratory has developed and used optical imaging techniques to monitor neuronal activity in the cerebellar cortex in vivo. These techniques include the use of voltage sensitive dyes [1–3], neutral red [4,5], flavoprotein autofluorescence [6,7], and Ca++ imaging [7]. Optical imaging has allowed us to address a spectrum of questions about both normal and abnormal cerebellar cortical physiology.

This chapter focuses on the use of pH imaging, specifically neutral red, as a measure of neuronal activation in the cerebellar cortex. Our earliest attempts to use voltage sensitive dyes in the cerebellar cortex in vivo were disappointing due to the small size of the signal [3]. Therefore, we turned to other approaches to optically map neuronal activity in the cerebellum. The first of these approaches was neutral red imaging [4]. Neutral red imaging exploits the known close coupling between pH changes and neuronal activation [8,9]. In this chapter we discuss the theory underlying pH imaging with neutral red, describe the techniques used to image the cerebellar cortex in vivo, and provide examples of how neutral red imaging has generated insights into the functioning of cerebellar cortical circuits.


A number of excellent reviews have extensively covered regulation of pH in the nervous system, pH and neuronal activity, and changes in pH during pathophysiological conditions [9–11]. Both a rich and complex topic, a general discussion of pH and the nervous system is beyond the scope of the present chapter. It suffices to state that neuronal activity is accompanied by extra- and intracellular pH shifts in neurons and glia across a wide range of species. In most preparations, including the cerebellar cortex and cerebral cortex, neuronal activity leads to neuronal acidification and glial alkalization. Extracellularly, the pH changes consist of a transient alkaline shift followed by a prolonged acidic shift. Also, large pH shifts, particularly acidification, occur in many pathological events in the central nervous system (CNS), including stroke, traumatic brain injury, epilepsy, and spreading depression [11–14]. Therefore, pH-based optical imaging is potentially an effective approach to map neuronal function and dysfunction in vivo.

A major issue is the choice of pH fluorescent dye for use in the CNS. Ideal properties of a pH indicator include maximal sensitivity within the physiological range in which pH shifts occur (~6.8–7.6), large signal-to-noise ratio, no toxicity, and deliverability. Another highly desirable property is sequestration to the compartment of interest (e.g., extracellular versus intracellular space). Characteristics, such as selectivity for specific cell classes (e.g., neurons versus glia) and for specific cell types (e.g., Purkinje cells versus granule cells), would be a major advance in pH imaging.

A number of fluorescent pH indicators have been developed for use in cells. The most widely used pH fluorescent dyes are BCECF (2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein) and SNARF (seminaphtharhodafluor) [15–18]. Both dyes have near-neutral pKa (7 and 7.5, respectively), in the physiological pH range of neurons (6.8–7.4). BCECF and SNARF have been primarily used in ratiometric imaging studies of extracellular or intracellular pH in vitro. We are unaware of published results using these pH indicators in the CNS in vivo, likely due to the difficulty of staining cells in the intact brain. Due to poor staining and toxicity, we were never successful in using these dyes to monitor neuronal activity in the rat cerebellar cortex in vivo. The only published pH optical imaging studies of activity patterns in the CNS have used the pH dye, neutral red [4,19].

Neutral red (3-amino-m-dimethylamino-2-methylphenazine hydrochloride) has long been recognized as a vital dye that could be useful as an indicator of cellular pH [20,21]. Neutral red has a pKa in the physiological pH range (6.8 in saline, 7.0 in rat brain) and changes its absorbance inversely with changing pH [22,23]. Therefore, the intracellular acidification of neurons that results from depolarization/excitatory stimulation will produce an increase in fluorescence [4]. The optimal excitation is centered on 540 nm, and the maximal emission is 630 nm. Large variations in the concentration of ions that have major effects on the excitability of neurons and glia (Na+, K+, and Ca++) have little effect on the emission spectra of neutral red [4]. Therefore, neutral red is relatively selective for shifts in pH.

Neutral red is highly lipophilic, uncharged at neutral pH, readily crosses the blood-brain barrier, and enters cells easily. Early uses of neutral red included as an assay of viability in cultured fish cells [24]. On entry into cells, neutral red is protonated, allowing it to be quickly trapped in cells and concentrated; as a consequence, little dye is associated with the cell membrane itself [20,21]. In slices of the cerebellar cortex, neutral red stains Purkinje cell bodies and their dendritic trees intensely [20]. However, the staining is not uniform, with interneurons exhibiting little staining while the molecular layer shows diffuse staining, presumably due to the densely packed parallel fibers [20]. Confocal microscopy of fresh sections of the cerebellar cortex stained with neutral red show that the dye is sequestered intracellularly in Purkinje cells (Chen and Ebner, unpublished data). Neutral red exhibits little toxicity in vitro and in vivo. [4,20] In agreement with La Manna’s assessment [21], neutral red is primarily located in the intracellular compartment and has many of the properties needed for measurement of intracellular pH, independent of extracellular pH.

Neutral red also has a measurable membrane potential dependent fluorescence that was attributed to changes in hydrophobicity [25]. It has been argued that this hydrophobic property could potentially be used to monitor faster changes in membrane potential, presumably by altering the association of neutral red with the plasma membrane [22]. However, this non-pH sensitivity is modest and unlikely to account for the large, slow fluorescent changes reported for neutral red in vivo. [4,25]


11.3.1. Animal Preparation

All our neutral red imaging studies have used either adult rats or FVB mice. Anesthesia is induced by intramuscular injection of a cocktail solution of ketamine (60 mg/kg), xylazine (3 mg/kg), and acepromazine (1.2 mg/kg), and monitored by EKG and corneal and foot pinch reflexes. Supplemental injections of the ketamine and xylazine solution are given as needed. The animal is placed in a stereotaxic frame and body temperature is maintained at 37°C by a feedback-controlled homeothermic blanket system. The trachea is cannulated below the cricoid cartilage to allow for artificial respiration with 95% O2 and 5% CO2. A craniotomy exposes the cerebellar cortex, usually Crus I and II, and a watertight chamber of acrylic is constructed around the opening and filled with Ringer’s solution containing (in mM): 123 NaCl, 3 KCl, 26 NaHCO3, 2 CaCl2, 2 MgSO4, and 10 D-Glucose bubbled with 95% O2 and 5% CO2. If needed, gallamine triethiodide (0.05 ml, 20 mg/mL, intramuscular injection) is used to paralyze the animal.

11.3.2. Dye Application

Because of its lipid solubility, neutral red readily penetrates the blood–brain barrier and cell membranes, easily staining the brain [20,21]. We have used two dye loading methods in the cerebellar cortex [4,5]. First, a 10 mM solution of neutral red is superfused over the exposed cortical surface for ~2 hours, supplemented with either intravenous (1 ml, 35 mM, for rats) or intraperitoneal (i.p.) injection of neutral red (0.3 ml, 35 mM, for mice). After staining, the neutral red solution in the chamber is thoroughly washed out and refilled with Ringer’s solution. Second, we subsequently learned that two i.p. injections of neutral red work equally well (35 mM, 2 ml/injection; first injection, 30 m before imaging and during surgery; second injection, immediately prior to imaging). The i.p. staining method offers two distinct advantages. First, there is no precipitation of neutral red on the cerebellar surface that requires extensive washout. Second, the preparation time is shortened by 1–2 h, permitting a longer recording period.

11.3.3. Electrical Stimulations, Electrophysiological Recording, and Pharmacology

Stimulation of parallel fibers is achieved using a train of pulses (typically 50–200 μA, 100–200 μs pulse width, 5–35 Hz for 10 s) via an epoxylite-coated tungsten micro-electrode (1–2 MΩ) placed at or just below the surface of the cerebellar cortex. To activate the climbing fiber projections to the cerebellar cortex a tungsten microelectrode is stereotaxically placed through the dorsal foramen magnum into the contralateral inferior olive and stimulated with a train of pulses (50–350 μA, 100 μs pulse width, 10 Hz for 5–10 s). Electrical stimulation of the ipsilateral face is delivered by two closely spaced, platinum needle electrodes placed subdermally using frequencies from 2 to 20 Hz for 5–20 s at 5–30 V with pulse widths of 100–500 μs. The voltage for face stimulation is adjusted to just below the threshold to evoke observable muscle twitches.

In some experiments, extracellular recordings of either field potentials or single cerebellar neurons are obtained with glass microelectrodes (2 M NaCl, 2–5 MΩ) using conventional electrophysiological techniques [4,7]. One of the advantages of optical imaging is that it allows precise placement of recording electrodes into the desired locations. The electrophysiological recordings are digitized (50 kHz), averaged on-line and stored for off-line analysis.

To examine the contributions of various receptors, neural signaling pathways, and pH modulation in the cerebellar cortex, various drugs were added to the Ringer’s superfusing the chamber.

11.3.4. Optical Imaging

The animal in the stereotaxic frame is placed on an x-y stage mounted on a modified Zeiss or Nikon epifluorescence microscope. Either a 2× or 4× objective coupled with a 0.63× reducing lens is used to view the exposed cerebellar cortex. Images are acquired with either a PXL (512 × 512 pixels) or Quantix 57 (530 × 512 pixels) cooled charge coupled device (CCD) frame transfer camera with 12-bit digitization (Roper Scientific, Tucson, Arizona). The camera exposure time is 100–200 ms. A 100 W mercury–xenon lamp (Hamamatsu Photonics, Shizouka, Japan) with a direct current-controlled power supply (Opti Quip model 1600, Highland Mills, New York) was used as the excitation light source. The filter set for neutral red includes an excitation filter at 546 ± 10 nm, an emission filter at >620 nm, and a dichroic mirror of 580 nm [4,21]. Images are binned to yield resolutions of typically 10 × 10 μm2 or 14 × 14 μm2. Imaging flavoprotein autofluorescence used a band-pass excitation filter (455 ± 35 nm), an extended reflectance dichroic mirror (500 nm), and a >515 nm long-pass emission filter [6]. The Ca++ filter set was customized to exclude the majority of autofluorescence signal (excitation 490–510 nm), a long-pass dichroic mirror of 515 nm, and emission 520–530 nm [7].

11.3.5. Data Processing and Analysis

The imaging protocol includes obtaining control images followed by a sequence of images with stimulation or other manipulations. The total number of frames acquired depends on the specific experimental question. Typically, the frame duration is 100–200 ms. Due to the high signal-to-noise ratio of neutral red imaging, the signal is easily detected and studied without averaging. The fluorescence change in each image, Fi, is quantified on a pixel basis by subtracting a background fluorescence frame (FB) and then dividing by FB, that is ΔF/F = (Fi–FB)/FB. The average of 10–20 control frames was used as FB. Next, the average ΔF/F in regions of interest is determined. For example, parallel fiber stimulation evokes a beam-like response and inferior olive stimulation evokes a band-like response that are defined as regions of interest. For spreading acidification and depression, the region of interest is the folium stimulated. To establish the significance of any changes in the amplitude of the optical response within a region of interest either paired student’s t-test (within animals) or ANOVA (among animals) using a randomized complete blocked design followed by Duncan’s post hoc is used.

More specialized analysis can be used as needed. For example, to quantify the optical responses evoked by face and inferior olive stimulation, a 2D Fourier analysis is used [26]. Based on the assumption that the parasagittal bands evoked are of lower spatial frequencies than the background noise and that the vasculature is primarily horizontally aligned, this analysis extracted the power in the parasagittal bands.

Two types of visualization methods are used. The first is simply based on the ΔF/F for a select image, in which either a grey or pseudocolor scale depicts the changes in fluorescence relative to background (examples shown in Figures 11.4A,B, 11.5C, and 11.6A). Given the large amplitudes of the optical responses, this first approach to visualization has a number of advantages. These include being relatively simple to implement, minimizing assumptions about the underlying statistics of the data, as well as showing the amplitudes of both the response and the background noise. The second involves an activation map constructed by statistical “thresholding.” As described in detail previously [4,26], statistically significant changes in fluorescence relative to background fluorescence are determined (examples shown in Figure 11.5A,B). Pixels with intensity levels greater than or equal to a threshold (e.g., 2–5 standard deviations of the mean background fluorescence) are defined as statistically significant, pseudocolored and superimposed on an image of the background. Statistical thresholding has the advantage of removing the background noise as well as visualization of the optical signal in relation to the underlying anatomy. However, this methodology has the intrinsic problem of selecting a threshold that distinguishes between the noise and the signal of interest. Ideally, both methods of visualization can be used to fully convey the nature of the optical responses.

FIGURE 11.4. Properties of responses to parallel fiber stimulation.


Properties of responses to parallel fiber stimulation. (A) Responses evoked by parallel fiber stimulation based on neutral red, flavoprotein autofluorescence and Ca++ imaging. (Figure 4a, panels 3,4 from Gao, W. et al., J. Neurosci. 26: 8377–8387, (more...)

FIGURE 11.5. (A) Ipsilateral face stimulation evokes parasagittal bands in Crus I and II.


(A) Ipsilateral face stimulation evokes parasagittal bands in Crus I and II. Activation map shows statistically significant optical signals (> 4 SD above or below control) pseudocolored and superimposed onto a gray scale image of the folia. Color (more...)

FIGURE 11.6. Spreading acidification and depression.


Spreading acidification and depression. (A) Series of optical images (stimulation minus background) illustrating the propagation of the optical response initiated by surface stimulation (150 μA, 150 μs pulses at 10 Hz for 10 s). The first (more...)


Our initial studies focused on understanding the pH dependence and origin of the neutral red signal. Fluorescence changes were determined when the rat cerebellar cortex stained with neutral red was superfused with Ringer’s of varying pH (6.0–8.8). Bathing the cerebellar cortex in alkaline Ringer’s results in a decrease in the mean fluorescence across the entire image. Conversely, bathing in acidic Ringer’s results in an increase in the fluorescence. Mean fluorescence increased 5.6% with a decrease of 1 pH unit and decreased 7.1% with an increase of 1 pH unit. To exclude the possibility that changes in neuronal activity and/or excitability contributed to the optical signals, these experiments were repeated after blocking voltage-gated Na+ and Ca++ channels, as well as AMPA receptors. The fluorescence changes were comparable, showing that the optical signals are primarily due to the manipulation of the pH.

A large number of experimental findings support the hypothesis that the pH dependent changes in fluorescence are primarily intracellular in origin. First, super-fusion with Ringer’s containing 20 mM sodium propionate produces an increase in fluorescence (Figure 11.1A). This increase in fluorescence is due to the uncharged form of the weak acid propionate acidifying the cytoplasm by carrying H+ into cells [27–29]. Second, replacing normal Ringer’s solution with a nominally CO2 free Ringer’s solution results in an abrupt decrease in fluorescence (Figure 11.1B). This decrease in fluorescence is consistent with a rapid efflux of CO2 from the cytoplasm, equivalent to a net extrusion of protons [30,31]. The increase in intracellular pH results in a decrease in neutral red fluorescence. Third, bathing neurons in NH4Cl produces an initial, brief intracellular alkalosis due to entry of NH3 and is followed by a much longer intracellular acidification due to the passive influx of NH4+ and subsequent efflux of NH3, leaving H+ behind when the NH4Cl is washed out [28,29]. Short duration perfusion with NH4Cl results in the expected longer duration increase in fluorescence (Figure 11.1C). The recovery from this intracellular acid load can be substantially reduced by amiloride, an inhibitor of Na+/H+ transporter that facilitates the removal of protons to the interstitial space [32]. This transporter is found in Purkinje neurons and granule cells [31,33,34]. Accordingly, recovery from the fluorescence increase that follows a pulse of NH4Cl was completely blocked by the addition of amiloride (Figure 11.1D). Finally, the cell permeant carbonic anhydrase inhibitor, acetazolamide, enhances the optical response to parallel fiber stimulation but the cell impermeant inhibitor, benzolamide, has no effect (Figure 11.1E,F). Blocking carbonic anhydrase intracellularly reduces the capacity of cells to buffer the excess protons generated [9], and a similar increase in intracellular acidification has been observed in Purkinje cells [35]. These observations demonstrate that the neutral red signals in the cerebellar cortex reflect primarily intracellular shifts in pH, in agreement with LaManna’s earlier observations [21].

FIGURE 11.1. Neutral red fluorescence changes are due to intracellular pH shifts.


Neutral red fluorescence changes are due to intracellular pH shifts. A–F illustrate the changes in fluorescence in the cerebellar cortex as a function of addition of various drugs to the bath. (A) Sodium propionate (20 mM) produces intracellular (more...)

The next question we addressed is whether the changes in fluorescence arise from neurons or glia. Elevated extracellular potassium produces an intracellular acidosis in neurons and an extracellular alkalosis. The neuronal acidification due to depolarization involves a number of mechanisms, including production of metabolic acid, mitochondrial Ca++/H+ exchange, glutamate-mediated H+ influx, and Ca++-related acidification in Purkinje cell dendrites [8,35–39]. In astrocytes the depolarization activates an electrogenic Na+-HCO3 cotransport that produces an alkaline shift [40–42]. The superfusion of the cerebellar cortex with Ringer’s solution with 8 mM KCl evokes a rapid and dramatic increase in neutral red fluorescence (Figure 11.2A). The increase in fluorescence due to KCl is converted to a decrease in fluorescence when neuronal activity is blocked with a cocktail including glutamate receptor antagonists, blockers of transmitter release and voltage-gated Na+ channels (Figure 11.2B). The decrease in fluorescence on depolarization with KCl was attributed to alkalosis of cerebellar glia and confirmed by blocking the glial depolarization evoked decrease with Ba++ (Figure 11.2C). Therefore, neurons and glia contribute optical signals of opposite sign to depolarizing stimuli (increase in fluorescence for neurons and a decrease for glia), consistent with the characteristics of activity dependent neuronal and glial pH modulation [8,9].

FIGURE 11.2. Neurons and glial cells respond differentially to depolarization.


Neurons and glial cells respond differentially to depolarization. (A) Superfusion of the cerebellar cortex with 8 mM KCl evokes a large increase in fluorescence. (B) The addition of TTX (10 μM), CNQX (10 μM) and MnCl2 (6 mM) to the Ringer’s (more...)

Several additional experiments ruled out other possible factors as contributing to the signal observed with neutral red. Neuronal activity results in an increase in neuronal and glial cell volume and a reduction in the extracellular space [43,44]. That this increase in cell volume is responsible for the increase in fluorescence was excluded by demonstrating that fluorescent dyes that easily enter cells but lack pH sensitivity, show no measurable optical responses. Additionally, furosemide, an anion transport inhibitor that blocks extracellular space and cellular volume changes [44,45], does not affect the optical responses [46]. Also, parallel fiber stimulation in the unstained cerebellar cortex does not produce detectable epifluorescence optical signals.

Finally, we demonstrated that the neutral red signal is neither the intrinsic hemodynamic signal [47,48] nor due to flavoprotein autofluorescence [6,49]. For differentiation from hemodynamic signals we showed that the reflectance signal at the wavelengths used to monitor neutral red epifluorescence is undetectable in the unstained cerebellar cortex [4]. In response to parallel fiber stimulation, the largest hemodynamic reflectance signal is on the order of 0.5% ΔR/R, almost an order of magnitude smaller than the epifluorescence change obtained with neutral red. Finally, the neutral red signal consists of an initial increase in fluorescence, not a decrease, as expected for the intrinsic hemodynamic optical signal. To differentiate between neutral red and flavoprotein autofluorescence, we demonstrated there is no flavoprotein signal in the cerebellar cortex at the excitation and emission wavelengths used to monitor neutral red [6]. Therefore, activity-dependent signals generated by neutral red primarily reflect intracellular pH shifts.


One of the intriguing aspects of the cerebellar cortex is its highly ordered and stereotypic circuitry and functional architectures (Figure 11.3). Optical imaging provides a power tool to study these well-defined circuits and architectures. The parallel fibers define the first of these architectures. Parallel fibers are the molecular layer extension of granule cell axons and project medial and lateral along the long axis of a folium for several millimeters [50,51]. This massive system of fibers forms excitatory glutamatergic synapses on the dendrites of Purkinje cells and cerebellar interneurons (Figure 11.3A,B). Activating parallel fibers using a brief stimulus train evokes an optical response consisting of a longitudinal beam running parallel to the long axis of the folium [4,5,46,52]. The optical signal consists of an initial increase in fluorescence (acidic shift) that returns to baseline in approximately 60 s, followed by a beam of decreased fluorescence (alkaline shift) for up to 120 s. A major advantage of the neutral red optical signal is that its amplitude is 5–20 times larger (1–5% ΔF/F) than the epifluorescence signals obtained from most voltage sensitive dyes (0.1–0.5% ΔF/F) [3,53,54] or the reflectance change from the hemodynamic intrinsic signal (0.1–0.5% ΔR/R) [4,47,48]. The optical signal obtained with neutral red in the cerebellar cortex is equal to or larger than the recently described flavoprotein autofluorescence signal or in vivo Ca++ imaging (Figure 11.4A) [6,7]. The larger signal-to-noise ratio eliminates the need for averaging the responses to multiple stimulations. Clearly, a robust signal is a great advantage, particularly in the intact CNS in which there are numerous sources of noise, including cardiovascular and respiratory motion artifacts.

FIGURE 11.3. Cerebellar cortical circuitry and functional architectures.


Cerebellar cortical circuitry and functional architectures. (A) Diagram of the parallel fiber-Purkinje cell and climbing fiber-Purkinje cell circuitry. Parallel fibers, which extend medial and lateral along the long axis of a folium, are the branched (more...)

The neutral red optical signal has several additional important characteristics. First, it is monotonically and linearly related to the stimulus parameters over a wide range of amplitudes and frequencies (Figure 11.4B–D). Second, the optical signal originates primarily from the postsynaptic elements (i.e., Purkinje cells and interneurons) and involves both iontotropic and metabotropic glutamate receptor (mGluR) activation. Both receptor types are present postsynaptically at parallel fiber-Purkinje cell synapses and provide for the vast majority of the Purkinje cells response to parallel fiber inputs [55–57]. Application of the non-NMDA glutamate receptor antagonist, CNQX, decreases the optical signals by 50–60%, and the mGluR receptor antagonist MCPG reduces the signal by an additional 15–25% (Figure 11.4E) [5,46,58]. Therefore, approximately 80–85% of the increase in fluorescence is due to activation of the postsynaptic targets of the parallel fibers.

Purkinje cells are likely to contribute a large fraction of this postsynaptic signal as the depolarization opens voltage-gated Ca++ associated with large acidic shifts in the dendrites of these cells [35]. The Ca++ is exchanged for protons via the plasma membrane calcium ATPase, which is expressed at high density in Purkinje cell dendrites [59]. Neutral red preferentially stains Purkinje cells [20]. What remains unresolved is the exact contribution of cerebellar interneurons, including stellate, basket, and Golgi cells, all of which have dendrites in the molecular layer that are excited by parallel fibers [50,51]. Cerebellar interneurons were shown to be only weakly stained by neutral red [20], suggesting these cells may make only a small contribution to the optical response. Further studies are needed to address this question.

Lastly, the spatial resolution obtainable using neutral red is excellent. Stimulation at low amplitudes evokes parallel fiber-like optical responses approximately 40–80 μm in width (anterior–posterior extent), considerably smaller than the ~300 μm width of a Purkinje cell dendritic tree (Figure 11.4B). The anterior–posterior extent of the optical response to surface stimulation is more constrained than the field potential recordings [4,26]. This is not unexpected, given that the neutral red response is primarily intracellular, and the field potentials are volume conducted through the extracellular space. Also, the optical response to parallel fiber stimulation is as spatially precise with neutral red as the responses obtained using other imaging techniques (Figure 11.4A and B). Therefore, the neutral red optical response is precise enough to monitor the activation of only a region of the dendritic tree of a Purkinje cell.

The above comments refer to the initial phase of increased fluorescence evoked by parallel fiber stimulation. What is the source of the subsequent decrease in fluorescence (Figure 11.1E,F)? One possibility is the decrease is due to an alkaline shift in neurons following the initial acidic shift. This would require that neuronal pH increase below resting levels following depolarization. The decreased fluorescence is most likely due to a depolarization-induced alkalosis occurring in cerebellar astrocytes [40,60,61]. Our studies did not tackle this important problem. One potential approach to examining this question would be to selectively block the Na+-HCO3 cotransporters or mGluRs on astrocytes.

One limitation of neutral red imaging is the apparent inability to map postsynaptic inhibition. Stimulation of the parallel fibers activates molecular layer interneurons (stellate and basket cells) that in turn produce a powerful GABAergic inhibition of both Purkinje cells and other molecular layer interneurons as diagramed in Figure 11.3B [62–64]. The inhibition occurs within the beam of activated parallel fibers (on-beam inhibition) and lateral to the beam (off-beam inhibition). Both on- and off-beam inhibition is mediated via GABAA receptors [64,65]. On-beam inhibition can be detected indirectly as GABAA antagonists increase the amplitude and width of the activated beam [5,46]. However, there is no obvious signal lateral to the beam suggestive of off-beam inhibition (Figure 11.4). Recently, we demonstrated using flavoprotein and Ca++ imaging that off-beam inhibition can be detected and it is organized in parasagittal bands [7,66]. However, the failure to observe off-beam inhibition with neutral red is not unanticipated. The opening of GABAA receptors results in an intracellular acidification due to a HCO3 efflux, a finding that has been documented in a variety of preparations [67,68]. Therefore, the shifts in pH resulting from postsynaptic GABAA receptor mediated inhibition would not be distinguishable from postsynaptic glutamatergic excitation. Also, the pH effects are likely to be smaller and more difficult to detect.

The other major limitation of any form of pH imaging in the CNS is the relatively slow time course relative to electrophysiological time scales (Figure 11.1E). The pH changes, and therefore the fluorescence changes, occur over seconds and not milliseconds. As a result, neutral red imaging will be best for mapping spatial patterns of activity or processes with slower time courses, as detailed in the following sections. However, the latency of the fluorescence increase with neuronal activation is excellent, with changes detected in less than 100 ms. As a method for monitoring pH changes in the CNS, the response time is considerably faster than the 1–2 s response time of even the fastest pH microelectrodes [9].


11.6.1. Parasagittal Organization of the Cerebellar Cortex

In addition to the medial-lateral organization of the parallel fibers, another major functional architecture of the cerebellum is the parasagittal zonation of its afferent and efferent projections and molecular compartmentalization. Climbing fibers arise solely from the inferior olive, synapse monosynaptically on Purkinje cells and produce low frequency, all-or-none complex spikes [51,69]. The climbing fiber projection from the inferior olive makes excitatory, glutamatergic synapses on Purkinje cells in parasagittal zones in the cerebellar cortex (Figure 11.3A) [70]. The output projections of the Purkinje cells to the deep cerebellar nuclei are also organized into parasagittal zones [71]. Further, numerous markers reveal a parasagittal molecular compartmentalization of Purkinje cells [72,73]. How this parasagittal architecture functions is a major question in cerebellar research.

We have used neutral red imaging to map the spatial patterns of activation in the rat and mouse cerebellar cortex evoked by peripheral stimulation [19,26,74]. Electrical stimulation of the vibrissae area of the ipsilateral face in the rat evokes optical responses in Crus I and II consisting of parasagittal bands (Figure 11.5A). The bands are 100–500 μm in width, elongated in the anterior–posterior direction, commonly extend across at least two folia, and vary in number from 1–7. The optical responses evoked by peripheral stimulation are due to activation of postsynaptic elements, as they are essentially blocked by CNQX. At the mossy fiber-granule cell, parallel fiber-Purkinje cell and climbing fiber-Purkinje cells synapses AMPA receptors are major contributors to the post-synaptic responses [57,75,76].

We hypothesized that these parasagittal responses are due to activation of the inferior olive and the climbing fiber projections to Purkinje cells. A number of observations are in agreement with this hypothesis. The optical bands evoked by peripheral stimulation are in register with the parasagittal compartmentalization of the olivocerebellar projection and Purkinje cells as delineated by immunostaining with anti zebrin II [19,72,73]. The parasagittal bands show a preferred frequency of face stimulation (6–8 Hz), consistent with the inherent rhythmicity of inferior olivary neurons and complex spike responses in Crus I and Crus II [77–79]. Contralateral inferior olivary stimulation evokes parasagittal bands with nearly identical spatial and frequency tuning characteristics to those evoked by peripheral stimulation (Figure 11.5B) [5,19,26,52]. Finally, lidocaine injection into the inferior olive blocks the parasagittal bands evoked from the periphery [26]. These findings demonstrate that peripheral inputs activate climbing fibers to well-defined parasagittal zones in the rat cerebellar cortex. These observations support the hypothesis that the inferior olivary neurons are dynamically coupled, and that the frequency content of the stimulus is critical to engaging the inferior olive.

One unanswered question is why the peripheral stimulation evokes climbing fiber responses with little evidence of a contribution from mossy fiber activation of the granule cell-parallel fiber-Purkinje cell circuitry. Most likely the answer involves the powerful, all-or-none action of climbing fibers on Purkinje cells. The massive and long duration depolarization of Purkinje cells associated with the complex spike should result in large acidic shifts in Purkinje cell dendrites [35,80]. The mossy, fiber-granule, cell-Purkinje cell pathway has a smaller effect on Purkinje cell depolarization and simple spike firing [62,81], and would likely generate smaller pH shifts. Potentially, pH indicators with greater sensitivity could be used to monitor the responses to mossy fiber inputs.

11.6.2. Opticali Maging of Plasticity at the Parallel Fiber-Purkinje Cell Synapse

We have also used neutral red imaging to examine the plasticity of the parallel fiber-Purkinje cell synapse in vivo. Conjunctive stimulation of climbing fiber and parallel fiber inputs onto cerebellar Purkinje cells results in long-term depression (LTD) at parallel fiber-Purkinje cell synapses [82]. Parallel fiber-Purkinje cell LTD has been hypothesized to play a major role in cerebellar motor learning [83,84]. The intracellular signaling mechanisms have been extensively studied in vitro and include the activation of group 1 metabotropic glutamate receptors (mGluR1) and protein kinase C (PKC) in Purkinje cells leading to internalization of AMPA receptors [85,85–89]. While parallel fiber-Purkinje cell LTD has been shown to exist in decerebrate animals [82,90–92], there have been few studies in the intact animal and no characterization of the intracellular mechanisms or spatial aspects of LTD in vivo.

Using neutral red imaging, we optically mapped and characterized LTD of the parallel fiber-Purkinje cell synapse evoked by conjunctive stimulation of parallel fibers and the contralateral inferior olive [52]. The properties of parallel fiber-Purkinje cell LTD predict that the depression will be spatially specific and occur at the intersection of the evoked parallel fiber and climbing fiber responses [93,94], as has been demonstrated in vitro [95]. At the site of the intersection of the parallel fiber-evoked beam and the climbing fiber-induced parasagittal band (Figure 11.5C), the subsequent responses to parallel fiber stimulation are depressed for at least 60 minutes (Figure 11.5E). The depression is spatially specific, confined to the region at which the parallel fiber and climbing fiber responses overlap (Figure 11.5D). Guided by the optical imaging, electrophysiological monitoring of the postsynaptic response to parallel fiber stimulation confirmed that the LTD occurred only at the intersection of the activated parallel fibers and climbing fibers. The generation of the LTD requires the conjunction of both inputs. Also, in agreement with the in vitro results, blocking mGluRs prevents the induction of LTD. Finally, we took advantage of a transgenic mouse that expresses an inhibitor of PKC selectively in Purkinje cells [96]. Parallel fiber-Purkinje cell LTD is abolished in this mouse as is vestibular-ocular reflex adaptation and learning-dependent timing of conditioned eyeblink responses [96,97]. As predicted, the optical counterpart of parallel fiber-Purkinje cell LTD is not present in this transgenic mouse. Conversely, LTD is preserved in the mGluR4 knock-out mouse that is known to have intact parallel fiber-Purkinje cell LTD [98]. Therefore, neutral red imaging provided the first visualization of parallel fiber-Purkinje cell LTD in vivo, demonstrated its spatial specificity, and its dependence on mGluR1 and PKC.

11.6.3. Spreading Acidification and Depression

A distinct advantage of optical imaging is the ability to study the spatial and temporal proprieties of cellular activity throughout a region of the brain. In addition to visualizing the activity of populations of cells or neuronal circuits, optical imaging methodologies have proven well-suited for characterizing the propagation of waves of activity in the CNS. For example, classical spreading depression of Leao [99] and its variants have been extensively studied with intrinsic signal optical imaging [100–102]. The discovery and investigation of calcium waves has relied on calcium imaging [103,104]. Spreading depolarization in the embryonic brain has been characterized using voltage sensitive dyes [105]. In this tradition, optical imaging with neutral red allowed us to discover and characterize a novel form of propagated activity in the cerebellar cortex in vivo. [46,106]

Surface stimulation of the cerebellar cortex normally evokes a highly constrained “beam” of activity, due to the activation of the parallel fibers and their postsynaptic targets. If the surface stimulation is sufficiently intense, the evoked beam of increase in fluorescence propagates anteriorly and posteriorly beyond the beam of activated parallel fibers and Purkinje cells (Figure 11.6A) [46,49,106]. Essentially, this is an all-or-none event that is initiated when a threshold is reached. The increase in fluorescence is dramatic and attains levels as high as 30% above baseline. The propagation and increase in fluorescence continues for 1–2 minutes beyond the duration of the surface stimulation. The spread is initiated nearly simultaneously along the beam of activated parallel fibers and travels orthogonally to the beam. The average propagation speed is ~500 um/s with peak speeds up to 2000 μm/s. The wave of increased fluorescence spreads for considerable distances, traveling along the sulci to reach neighboring folia (Figure 11.6B).

The propagation of the increased fluorescence is accompanied by a transient but powerful depression of the cerebellar cortical circuitry (Figure 11.6C). Both the presynaptic and postsynaptic field potentials evoked by parallel fiber stimulation can be completely suppressed as the wave of increased fluorescence passes over a folium [46,106]. The presynaptic component recovers to normal levels within 30–60s; however, recovery of the postsynaptic component takes considerably longer (60–300 s). Therefore, we named this propagation event spreading acidification and depression (SAD). The mechanisms underlying the suppression of cerebellar cortical circuitry during SAD are not known. The large pH shifts during SAD may directly decrease neuronal excitability, possibly by depressing voltage-gated sodium channels [107] and/or attenuating glutamate channel conductance [108]. A large number of factors differentiate SAD from classical spreading depression [46,106]. SAD can also be imaged with voltage sensitive dyes and flavoprotein autofluorescence, showing that large changes in depolarization and oxidative metabolism are occurring and potentially contribute to the decrease in excitability [49].

Insights into the mechanisms underlying SAD are based on experiments assessing the contributions of presynaptic and postsynaptic elements of the cerebellar circuit. An increase in the excitability of the cerebellar circuits is needed to evoke SAD [46,49]. The greater the stimulation intensity, the greater the probability of evoking SAD. Both TTX and Ca++-free Ringer’s completely prevent evoking SAD by parallel fiber stimulation. In contrast, blocking AMPA and metabotropic glutamate receptors increases the threshold for evoking SAD but does not prevent the generation of SAD [46]. Increasing the stimulation intensity invariably results in SAD, even when combinations of glutamate receptor blockers are applied. We also demonstrated that the threshold for generating SAD is greatly lowered by blocking Kv1.1 potassium channels that play a major role in controlling neuronal excitability [49]. A single stimulus to the parallel fibers can evoke SAD in the presence of a Kv1.1 blocker, and spontaneous SAD is observed. Therefore, the activation of parallel fibers and the release of glutamate are essential for evoking SAD. Conversely, activation of Purkinje cells or molecular layer interneurons is not essential.

That the direction of SAD propagation is perpendicular to the parallel fibers raises the possibility that the molecular layer interneurons, with their axons extruding perpendicular to the parallel fibers, provide the underlying neuronal substrate (Figure 11.3B). However, cerebellar cortical interneurons are not critical elements of SAD as the threshold for SAD is actually decreased by GABAA blockers [46] and SAD occurs in mice lacking stellate cells [109]. Nor is the gaseous neurotransmitter, nitric oxide, critical, as SAD can be evoked in nitric oxide synthase deficient mice or in the presence of blockers of nitric oxide synthase.

Therefore, the mechanism underlying the generation and propagation of SAD remains unresolved. Our working hypothesis is that SAD is due to the unusual release of glutamate from a bundle of activated parallel fibers. In turn, the glutamate diffuses to and activates neighboring parallel fibers. Preliminary simulations show that this parallel fiber model accounts for the high speed, geometry and threshold properties of SAD [110]. We also hypothesized that SAD is primarily a pathophysiological process that can temporarily disrupt cerebellar function. One possibility is that a SAD-like process occurs in the Kv1.1 channelopathy, episodic ataxia type 1 (EA1) [111]. EA1 is due to a mutation in the gene encoding for the Kv1.1 α-subunit [112,113] and EA1 patients have transient attacks of cerebellar dysfunction [114,115]. We have argued that the properties of SAD are consistent with the molecular and clinical features of EA1 [49,111].


Improvements in pH indicators for imaging in vivo would be welcomed. As described in this chapter, the activity-dependent optical signals obtained using neutral red have an excellent signal-to-noise ratio and little toxicity. However, pH indicators that further increase the signal size will allow assessment of smaller perturbations in pH. Also, indicators with greater signal-to-noise ratios are needed for imaging in awake, behaving animals. Additionally, it would be useful to have a spectrum of pH dyes that allow imaging of different compartments, particularly dyes that sequester exclusively in the intracellular versus extracellular space. Indicators that are sensitive to different pH ranges and have different excitation/emission properties would allow monitoring multiple parameters in the same experiments (e.g., pH and Ca++, pH and flavoprotein fluorescence, etc). There are a host of questions concerning the interactions between neurons and glia, cerebral metabolism, and the regulation of intracellular and extracellular pH that would benefit from improved pH indicators.

Ideally, the next generation of pH dyes would provide for cell specific imaging. In the cerebellum, having the capability to image specific populations of neurons, such as Purkinje cells or granule cells, or simply differentiate between neurons and glial cells, would constitute a major advance in optical imaging in vivo. Cell-specific imaging offers the chance of solving long-standing problems in cerebellar cortical physiology. For example, the controversy about whether parallel fibers are activated in “beam-like” patterns in response to natural inputs [116] could potentially be answered. A series of pH sensitive, GFP-based genetically encodable fluorescent probes have been developed [117,118], and mice have been generated on a pH-sensitive variant of enhanced yellow-fluorescent protein [119]. An important caveat is the need to distinguish between the fluorescence changes due to intrinsic optical signals, such as flavoprotein autofluorescence, and the hemodynamic intrinsic signal [6,47]. In Section 11.1.4, Nature and Origin of the Neutral Red Fluorescence Signal, we discussed several approaches to differentiating these different optical signals. Also promising is the use of viral-vector techniques for gene transfer [120,121]. The viral-vector approach has the advantages of requiring less effort, time, and expense than generating transgenic animals, yet having the potential to deliver intrinsic pH sensors to specific cell types. Furthermore, the viral vector approach is not limited to a few species in which transgenic animals are available, but to a much wider range of species including nonhuman primates. Approaches based on the power of genetic manipulation will undoubtedly have a major impact on the optical imaging of activity dependent signals in the future, including pH imaging.

Concerning answering neurobiological questions, we would suggest using neutral red imaging (or other pH imaging modalities) in three areas. First, given its large signal size, limited toxicity, and ease of use, neutral red is an excellent tool for mapping neuronal patterns of activation in CNS circuits/structures. While our laboratory has been focused on the cerebellar cortex, regions of the nervous system in which spatial patterning is hypothesized to play a role in the information processing are also candidates for study. In addition, it may be possible to use activity-dependent pH signals to map neuronal activity in the awake mouse. Neutral-red- injected intraperitoneally readily crosses the blood–brain barrier and stains the cerebellum. With the transparency of the cranial bone in the mouse [122], it may be possible to map neuronal activation in the awake, behaving mouse.

The second set of scientific questions are those of pH regulation in the CNS, how activity modulates neuronal and glial cells, and how shifting in pH can alter neuronal function [9]. The need to study pH-related changes in the intact nervous system is obvious and pH imaging in vivo will be a powerful tool.

Finally, the third area in which pH imaging can play an important role is studying abnormal and pathophysiological processes in the CNS. Acidosis plays a central role in neuronal damage during ischemic stroke through several mechanisms including nonspecific denaturation of proteins and nucleic acids [123,124], triggering of cell swelling via Na+/H+ and Cl/HCO3 exchangers [125], inhibition of mitochondrial energy metabolism [12], stimulation of pathologic free radical formation [126], and potentiation of glutamate receptor-medicated excitotoxicity [127]. Recent findings that acidosis activates a distinct family of membrane ion channels, the acid-sensing ion channels (ASICs) in both peripheral and central nervous systems has changed the view of acidosis-associated signaling in the brain [128,129]. Activation of these cation channels by protons plays an important role in a variety of physiological as well as acidosis-mediated neuronal injury. The acidosis due to brain ischemia causes an increase in glycolysis in both glia and neurons, leading to an increase in lactic acid production and a decrease of pH. This acidosis activates ASICs, resulting in Ca++ influx and subsequent neuronal damage. Similarly, epilepsy causes local acidosis due to increased energy demand and increased glycolysis [13]. These and other pathological processes are strong candidates for the use of pH imaging. While there were some early attempts using umbelliferone (7-hydroxycoumarin) as a fluorescence indicator of brain pH in cerebral ischemia [130,131], clearly a renewed effort using modern optical imaging techniques is warranted.


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