b2-ADRENERGIC RECEPTORS PROTECT AXONS DURING ENERGETIC STRESS BUT DO NOT INFLUENCE BASAL GLIO-AXONAL LACTATE SHUTTLING IN MOUSE WHITE MATTER

G. LAUREYS, a* M. VALENTINO, b F. DEMOL, a C. ZAMMIT, c R. MUSCAT, b M. CAMBRON, a R. KOOIJMAN d AND J. DE KEYSER a,e

Abstract

In vitro studies have demonstrated that b2-adrenergic receptor activation stimulates glycogen degradation in astrocytes, generating lactate as a potential energy source for neurons. Using in vivo microdialysis in mouse cerebellar white matter we demonstrate continuous axonal lactate uptake and glial–axonal metabolic coupling of glutamate/ lactate exchange. However, this physiological lactate production was not influenced by activation (clenbuterol) or blocking (ICI 118551) of b2-adrenergic receptors. In two-photon imaging experiments on ex vivo mouse corpus callosum subjected to aglycemia, b2-adrenergic activation rescued axons, whereas inhibition of axonal lactate uptake by a-cyano-4-hydroxycinnamic acid (4-CIN) was associated with severe axonal loss. Our results suggest that axonal protective effects of glial b2-adrenergic receptor activation are not mediated by enhanced lactate production. 2014 IBRO. Published by Elsevier Ltd. All rights reserved.

Key words: glucose, lactate, glutamate, white matter, axons, b2-adrenergic receptors.

INTRODUCTION

Recent data suggest that chronic failure of white matter energy metabolism plays a role in several neurodegenerative diseases (Matute and Ransom, 2012) such as Alzheimer’s disease (Yao et al., 2011), amyotrophic lateral sclerosis (Lee et al., 2012) and multiple sclerosis (Cambron et al., 2012).
Brain neuro-energetics is a major area of scientific debate with opposing opinions on how electro-active neurons and their surrounding ‘‘supportive’’ glial cells thrive or fail by exchange of glucose versus lactate as primary energetic substrates. Pellerin and Magistretti introduced the astrocyte-neuron lactate shuttle hypothesis (ANLSH) (Pellerin and Magistretti, 1994) in which synaptic glutamate-release is balanced by astrocytic re-uptake and conversion to glutamine. This process is supposed to be driven by astrocytic glycolysis that generates lactate, subsequently taken up by active neurons to fuel their energetic demands. However, with regard to synapses in gray matter there are experimental data both in favor (Sampol et al., 2013) and against (Hall et al., 2012) the ANLSH hypothesis. Applicability of the ANLSH to the white matter setting however remains largely unexplored and most data are derived from ex vivo studies in rodent optic nerve suggesting a role for glycogen-derived lactate as an energy source for axons (Wender et al., 2000; Tekkok et al., 2005). Most of the glycogen of the central nervous system white matter resides in astrocytes (Wender et al., 2000) and is converted to lactate which is released into the extracellular space. Axons in mouse white matter take-up lactate through the monocarboxylate transporter (MCT)-2 (Tekkok et al., 2005), and metabolize it aerobically to energy. More recent ex vivo data demonstrates that lactate imported via MCT-1 transporters can rescue oligodendrocytes and prevent demyelination under low glucose conditions (Rinholm et al., 2011). Subsequently, Lee et al. demonstrated how oligodendrocytes provide axons with lactate, critical to their survival, via the same MCT-1 transporter (Lee et al., 2012). These data point to glial cell-derived lactate as a critical factor for axonal energy metabolism and survival in white matter. A summary of the supposed metabolic coupling mechanisms between axons and surrounding glial cells is illustrated in Fig. 1.
In vitro studies revealed that astrocytic glycogenolysis is under noradrenergic control, and that activation of astrocytic b2-adrenergic receptors induces glycogenolysis and increases Na+,K+-ATPase activity (Hertz et al., 2010). Little is known about these regulatory mechanisms in white matter in vivo, and we are aware of only one study showing that noradrenaline pretreatment reduces glycogen content in isolated mouse optic nerve (Wender et al., 2000). In the present study we explored the in vivo existence of astro-axonal coupling of lactate and glutamate metabolism and the potential influence of b2-adrenergic receptors in this process during axonal damage as a result of oxygen/glucose deprivation.

EXPERIMENTAL PROCEDURES

Chemicals and reagents

Aqueous solutions were made from purified water (Seralpur pro 90 CN, Belgolabo, Overijse, Belgium) and filtered through a 0.2-lm-membrane filter. The perfusion fluid for microdialysis consisted of artificial cerebrospinal fluid (aCSF) (147 mM NaCl, 3 mM KCl, 1 mM MgCl26H2O, 1.2 mM CaCl26H2O, 200 lM ascorbate, 352 mM NaH2PO4H2O, pH 7.4). ICI-118551 (200 nM), clenbuterol-hydrochloride (200 nM), DL-threo-bbenzyloxyaspartic acid (TBOA) (250 lM) and a-cyano4-hydroxycinnamic acid (4-CIN) (1 mM) (all from Sigma–Aldrich, St. Louis, MO, USA) were dissolved in aCSF and administered via the microdialysis probe. Selective in situ concentrations for each compound were determined using published affinity constants, and concentrations applied in the perfusion medium were calculated based on a microdialysis probe recovery of 10%. The 10% correction factor considers approximate recovery for small molecules considering the used flowrate and type of microdialysis probe (Grubb et al., 2002). For the two-photon imaging experiments the selective concentrations were applied without using the correction factor. The selectivity of 4-CIN for MCT-2 renders the interaction with energetic substrate uptake at the stipulated dose in astrocytes unlikely. Interaction with mitochondrial pyruvate uptake is improbable at the stipulated dose as previously determined (Erlichman et al., 2008; Newman et al., 2011).

Surgery

Protocols were in accordance with national guidelines and regulations on animal experiments and approved by the Ethics Committee on Animal Experiments of the Vrije Universiteit Brussel, Belgium and the Ethics Committee of the University of Malta. Male C57bl/6 mice (Charles Rivers, France) between 7 and 8 weeks of age (weighing 25–30 g), were anesthetized with a mixture of xylazine/ketamine (10/100-mg/kg, i.p.) and mounted on a stereotaxic frame. An intracranial guide (CMA/ Microdialysis, Stockholm, Sweden) was implanted in the bilateral cerebellar white matter (coordinates toward bregma were 5.7 mm posterior, 2.2 mm lateral and 2.3 mm ventral of the dura (Paxinos and Franklin, 2004)). Immediately after surgery, guide cannula obturators were replaced by microdialysis probes (CMA7; membrane length: 1 mm theoretical cutoff: 6000 Da; CMA/Microdialysis, Solna, Sweden). Postoperative analgesia was ensured by ketoprofen (4 mg/kg, i.p.). Animals were allowed to recover from surgery overnight receiving laboratory chow and water ad libitum. Probe localization was histologically verified postmortem (Fig. 4B). Animals with aberrant probe location were excluded from the study.

Intracerebral microdialysis

Microdialysis probes were continuously perfused with aCSF at a flow-rate of 1 ll/min (CMA/400 microdialysis pump, CMA/Microdialysis, Solna, Sweden). All experiments were performed on the day following surgery in non-anesthetized freely moving mice. Tubings were flushed with 70% ethanol and rinsed with purified water before perfusion with aCSF to exclude any bacterial interference with the glucose/lactate levels. 1 ll/min perfusion of the probes was started 2 h before the experiment to attain steady-state concentrations. Six dialysate samples (20 ll) were collected at 20-min intervals to determine the basal concentrations. TBOA, 4-CIN, clenbuterol or ICI-118551 was added to the perfusion medium at the last 20-min baseline sample. Thereafter six 20-min samples were collected during compound administration. In a separate set of experiments (n = 5) a control experiment was performed with a ‘‘sham’’ switch of syringes to exclude any effect of switching syringes on glucose, lactate and glutamate concentrations.

Liquid chromatographic assays

We used a gradient liquid chromatographic method for the quantitative simultaneous determination of aminoacids in dialysates as previously described (Van Hemelrijck et al., 2005). All substances were identified and quantified by comparing retention times and peak areas with those of external standards.

Enzymatic colorimetric assays

Microdialysate samples were analyzed for glucose and lactate content using enzymatic lactate (607-100) and glucose (606-100) assay kits (Biovision, Mountainview, CA, USA) according to the manufacturers’ guideline. Fluorescence was measured at 460 nm using a microplate reader (Model 680 Bio-Rad, Hercules, CA, USA) and an excitation wavelength of 355 nm.

Preparation of fresh slices for imaging of corpus callosum axons

Adult 12–20-week-old mice weighing 25–27 g with a C57BL6/J genetic background were used in experiments. To enhance the visualization of axon morphology, we used transgenic mice with neuronspecific expression of yellow fluorescent protein (YFP), under control of the thy1 promoter (Feng et al., 2000). We selected a mouse transgenic line (Line H), which expressed YFP in a subset of cortical neurons that project axons across the corpus callosum. After deep halothane anesthesia and decapitation, the cranium was opened and the brain rapidly removed and placed in chilled and oxygenated (95% O2/5% CO2) aCSF buffer supplemented with 75 mM sucrose. This hyperosmolar slicing solution prevents brain edema and is rich in magnesium and low in sodium and calcium, consisting of (mM) 234 sucrose, 11 glucose, 24 NaHCO3, 2.5 KCl, 1.25 NaH2PO4, 10 MgSO4 and 0.5 CaCl2. After removing the cerebellum and brainstem, the entire brain was mounted on the ice-cold platform of a Vibratome 1000 vibroslicer (Technical Products, St. Louis, MO) covered in modified ice-cold aCSF. The brain was oriented to cut coronal slices (400 lm thick) from the genu of the corpus callosum through the caudal extent of the hippocampus. Immediately after sectioning, slices were transferred to a Haas-type interface brain slice chamber (Harvard Apparatus, South Natick, MA) and allowed to recover at room temperature in oxygenated (95% O2/5% CO2) aCSF for 1 h at a flow rate of 3.5 ml/min aCSF was composed of (mM) 126 NaCl, 3.5 KCl, 1.3 MgCl2, 2 CaCl2, 1.2 NaH2PO4, 25 NaHCO3, 10 glucose, 0.43 L-lactate, pH 7.4. The osmolality (300 mOsm) was checked with a microosmometer (Precision Systems, Natick, MA).

Two-photon imaging of brain slices

A brain slice was then transferred to a submerged minichamber (0.5 ml) with a coverglass bottom (Warner Instrument Corporation, Hamden, CT) mounted on an upright BX50W1 Olympus Multiphoton microscope (Olympus, Tokyo, Japan) and perfused with room temperature oxygenated (95% O2/5% CO2) aCSF at a flow rate of approximately 3.5 ml/min. Final temperature control (33 ± 1 C) was maintained using an in-line heater (Warner Instrument Corporation, Hamden, CT) equipped with a feedback thermistor placed in the chamber and the temperature raised gradually over 1 h. Slices were continually perfused with oxygenated buffer during imaging by means of a gravity flow perfusion system and vacuum aspiration within the imaging chamber. Glucose deprivation was initiated by replacing glucose in normal oxygenated aCSF with 10 mM sucrose for a period of 15, 30 or 45 min and imaging was continued through 120 min of reperfusion with substitution to glucose-containing aCSF. In alternate experiments, aCSF in the aglycemic phase was supplemented with 0.1 mM 4-CIN (stock solution dissolved in 0.1 N NaOH), 20 nM clenbuterol or a combination of both drugs. Since 4-CIN necessitated dimethyl sulfoxide (DMSO) as a solvent, all solutions contained 0.01% DMSO. Control experiments for 15, 30 and 45 Control experiments for 15, 30 and 45 min hypoglycemia in the absence of DMSO excluded any effect on axon integrity due to DMSO alone. Control slices maintained in oxygenated superfusion medium at 33 ± 1 C demonstrated intact linear axonal morphology for at least 5 h after preparation (Fig. 4A). YFP-labeled axons were visualized with a 920-nm laser line (7% laser power) using a water-based 25 Olympus XLPLN25xWMP objective (NA 1.05, WD 2.0, IR-corrected).

Image processing

Image acquisition was performed using the Olympus FluoView software. Single-focal-plane images were collected at 30-min intervals or, more frequently, stacks of five optical sections at an incremental z-step of 1 lm apart. Subsequently, all z-stacks of images were projected along the z-axis to recreate two-dimensional representations of the three-dimensional structures within the imaged tissue. Post-acquisition images were only adjusted for brightness, contrast and background noise by using ImageJ. For brightness and contrast adjustments the depth of pixel intensities that spanned the entire 8-bit range (0–255) was readjusted for display optimization. The two-photon experiments were performed in triplicate for confirmation of reproducibility.

Scoring of axonal injury

Axon damage was quantified by visual scoring as previously described (Tekkok and Goldberg, 2001; McCarran and Goldberg, 2007). Images were divided into a 5 5 grid, and each grid box was scored by a blinded observer for the presence of axon damage using the following system: 0, no damage; 1, axon swelling and/or beading; 2, axon fragmentation. The total score for a single section (0–50) was divided by the number of grid boxes to give a mean damage score (0–2). Damage scores from three different experiments were averaged and recorded for each condition.

Statistical analysis

The average (with s.e.m.) of the 5 stable baseline dialysate levels for the animals included in the study are: 0.218 ± 0.016 mg/dl glucose, 0.036 ± 0.002 mg/dl lactate and 0.590 ± 0.046 lM glutamate. Glutamate, glucose and lactate levels are expressed relative to the stable baseline levels, which were equated to 100% with s.e.m. A correction for changes induced by syringe exchange was performed by subtracting the % change in sham conditions for each time point. Statistical analysis between pharmacologically induced levels and baseline level was performed using a Friedman test followed by Dunn’s multiple comparisons post hoc test. The basal area under the curve (AUC) was calculated as the sum of dialysate concentrations in the first six ‘‘basal’’ collections (20 min samples). AUC following drug was calculated as the sum of the dialysate concentrations in the six collections following the start of drug perfusion. AUC after drug administration was expressed as a percentage of basal AUC. Wilcoxonsigned rank test was used to compare basal and druginduced AUC. Animals were excluded from analysis when contamination of the sample or technical problems rendered measurement impossible. Statistical analysis was performed with the InStat Prism statistical package (GraphPad Software, La Jolla, USA). Axonal injury data are expressed as mean ± s.e.m., statistical significance versus controls was determined by a one-way ANOVA. p values are reported in the figure.

RESULTS

Exploration of metabolic shuttling in white matter

To address the presence of functionally coupled glutamate–lactate metabolism in white matter our first step was to pharmacologically block both ends of the presumed glial–axonal shuttle. Inhibition of axonal lactate uptake by the MCT-2 blocker 4-CIN resulted in an increased extracellular lactate concentration (Fig. 2A), whereas extracellular glucose levels were not affected (Fig. 2B). Inhibition of axonal lactate uptake increased extracellular glutamate concentrations (Fig. 2C). Inhibition of the astrocytic glutamate transporters by TBOA led to the accumulation of EC glutamate (Fig. 2C). TBOA also significantly diminished extracellular glucose levels (Fig. 2B) without influencing extracellular lactate concentration (Fig. 2A).
To evaluate the impact of astrocytic lactate production on axonal survival we performed a series of two-photon sequential imaging experiments. The effects of aglycemia on morphological damage of axons have not yet been described. Following aglycemia applied for up to 30 min, axonal structures were maintained throughout the complete recovery phase of 120 min of recirculation with glucose-containing aCSF (Fig. 3A). Forty-five minutes of glucose deprivation caused few immediate changes, but resulted in gradual axonal beading and fragmentation during reperfusion (Fig. 3B). To investigate if glycogen-derived lactate served as the protective factor during aglycemia, 4-CIN was supplemented to corpus callosum slices undergoing 30 min of aglycemia. Toward the end of the aglycemic period axonal beading was observed (Fig. 3C), followed by extensive damage during reperfusion as compared to the 30 min of aglycemia without 4-CIN (Fig. 3A).

Role of b2-adrenergic receptors in axonal neuroprotection and basal lactate production

The mean axonal injury scores for the various conditions are shown in Fig. 3. No statistically significant difference was observed in the axonal injury score throughout the duration of control experiments [F(14, 350) = 0.385, p = 0.979]. Control slices maintained in oxygenated superfusion medium demonstrated intact linear axonal morphology for at least 5 h after preparation (Fig. 4A). When compared with control slices, 30 min of GD did not result in any significant axonal injury while 45 min of GD did (p < 0.001). Addition of 0.1 mM 4-CIN during 30 min of GD resulted in considerable axonal damage, with these slices scoring a statistically significant (p < 0.001) higher axonal injury than control slices and slices following 30 min of GD without the drug. Addition of 20 nM clenbuterol offered protection to slices during 45 min of GD, with no statistically significant difference in axonal injury score between such slices and controls. This protection was lost when axonal lactate uptake was blocked by co-administration of 0.1 mM 4-CIN during 45 min GD, with slices now showing a statistically significant (p < 0.001) difference in axonal injury score when compared with control slices. Although axonal damage was clearly visible, fluorescence was relatively preserved (for comparison see Fig. 3C illustrating loss of fluorescence with administration of 4CIN only), suggesting that clenbuterol still exerted protective effects independent of a possible lactate-mediated effect blocked by 4CIN. In vivo intracerebellar administration of the selective b2-antagonist ICI-118551 or b2-agonist clenbuterol did not significantly change extracellular lactate or glucose concentrations (Fig. 2D, E).

DISCUSSION

The increase in extracellular lactate during pharmacological blockade of axonal lactate uptake demonstrates the existence of a physiological glia–axonal lactate shuttle in white matter under basal conditions. Inhibition of glycogenolysis elevates extracellular glutamate concentrations in vitro (Sickmann et al., 2009; Schousboe et al., 2010). Our data demonstrate an in vivo counterpart during 4-CIN administration since the increase in extracellular glutamate probably reflects inhibition of astrocytic lactate production by feedback inhibition (Sotelo-Hitschfeld et al., 2012). Functional glutamate reuptake by optic nerve axons has been demonstrated (Arranz et al., 2008) and may constitute a complementary mechanism of glutamate accumulation when axons are deprived of lactate influx. Inversely inhibition of glutamate reuptake with TBOA decreased extracellular glucose levels compatible with enhanced glycolytic glucose consumption. In summary, these data are in line with functional exchange machinery where glial cells produce lactate during the buffering process of extracellular glutamate, with lactate subsequently being transported to axons as energy source. In our sequential two-photon imaging experiments we found a time window of 30 min of aglycemia before axonal damage occurred following reperfusion with glucose. These findings confirm a morphological counterpart to the previously described 30-min protective glycogen-buffer in ex vivo experiments of rodent optic-nerve electrophysiology (Wender et al., 2000). When we blocked axonal lactate uptake during 30 min of aglycemia damage occurred, confirming that lactate supports axonal energetic needs.
Hypoglycemia activates Locus Coeruleus neurons (Morilak et al., 1987) leading to increased noradrenaline release throughout the brain (Bengzon et al., 1991). Noradrenaline can enhance astrocytic glycogenolysis by b2-adrenergic activation as previously described in vitro (Hertz et al., 2010). Therefore, we suggested b2-adrenergic stimulation would provide glycogenolytic support of axons in our model. Callosal slices are deprived of noradrenergic input from locus coeruleus projections necessitating pharmacological stimulation. b2-adrenergic stimulation with clenbuterol indeed sustained axonal survival during aglycemia in our two-photon experiments (Fig. 3D). Moreover this effect was found to be counteracted by the addition of 4-CIN (Fig. 3E). Since the protective effect of clenbuterol and the deleterious effect of 4-CIN might represent independent effects we investigated the role of b2-adrenergic stimulation and inhibition on in vivo extracellular levels of glucose and lactate. Neither stimulation nor inhibition led to significant changes in extracellular glucose or lactate concentrations suggesting alternative mechanisms convey b2-adrenergic axonal protection during aglycemia.

CONCLUSION

We showed that b2-adrenergic receptors exert axonal protective effects under aglycemic conditions. Although our data support the existence of a functional glutamate–lactate exchange system between white matter glia and axons, b2-adrenergic receptors did not affect this system. Therefore, we postulate that protection of axons during energetic stress by the b2-adrenergic receptor agonist clenbuterol may occur through previously described anti-inflammatory, antioxidative and neurotrophic effects of clenbuterol (Gleeson et al., 2010), and not by enhanced astrocytic lactate production.

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