Fluorescent CSF Tracers

All CSF tracers were reconstituted in artificial CSF at a concentration of 0.5%. For co-infusion experiments, small molecular weight Texas Red conjugated dextran (TR-d3; 3 kDa) and large molecular weight Alexa 647 conjugated Ovalbumin (OA-647; 45 kDa) were mixed in a 1:1 ratio of 1% solutions. All fluorescent CSF tracers were acquired from Invitrogen.

Intracisternal Tracer Infusions

Mice were anesthetized and fixed in a stereotaxic frame while the posterior atlanto occipital membrane overlying the cisterna magna was surgically exposed. For all CSF tracer experiments, tracers were infused into the subarachnoid CSF via cisterna magna puncture, at a rate of 2 µL/min for a period of 5 minutes (10 µL total volume) through a 30 GA syringe pump (Harvard Apparatus). In prior studies, we have found that this rate and duration of tracer infusion does not cause reflux of subarachnoid CSF back into the ventricular CSF compartments, suggesting that the physiological direction of CSF bulk flow is maintained (4, 6, 7). We have observed that intracisternal CSF tracer infusion mildly increases intracranial pressure (ICP), however these effects persist only for the duration of tracer infusion and ICP normalizes rapidly after the cessation of infusion (7). Additionally, we have observed that at infusion rates that do not change ICP, paravascular CSF tracer flux follows the same patterns observed in the present study (5, 8). Because paravascular CSF recirculation occurs long after the normalization of ICP after the cessation of tracer infusion, the CSF tracer fluxes observed in the present study appear to represent the physiological movement of CSF along paravascular spaces after the tracer loading that occurs during the 5 min infusion period. To visualize penetration of fluorescent CSF tracers into the brain parenchyma ex vivo, anesthetized animals were trans-cardially perfusion fixed with 4% paraformaldehyde (PFA) at 30 minutes after the start of infusion, a time point previously identified to correspond to robust tracer penetration of similar molecular weight compounds in young male C57BL/6 mice (4). Brains were then dissected and post-fixed in 4% PFA for 24 hours before being sliced into 100 µm coronal sections using a vibrotome and mounted using PROLONG anti-fade gold with DAPI (Invitrogen). Tracer movement along perivascular spaces and permeation of the brain parenchyma was visualized in ex vivo studies by conventional fluorescence microscopy and confocal laser scanning microscopy.

Ex-vivo imaging of fluorescent CSF-tracers

Tracer recirculation along perivascular pathways and penetration into the brain parenchyma was visualized by conventional fluorescence microscopy of 100 µm vibratome coronal brain sections as described previously (4). In brief, 8 brain sections per animal were imaged and analyzed by a blinded investigator using an Olympus fluorescence microscopy under 4x objective power to generate whole-slice montages (using the Virtual Slice module of Microlucida software, MicroBrightfeild).

Tracer penetration was quantified using NIH Image J software as described previously (4). Briefly, fluorescence channels from each slice were split to separately measure fluorescence of co-infused small and large molecular weight Texas Red conjugated dextran (Dex-3; 3 kDa) and Alexa 647-conjugated ovalbumin (OA-45; 45 kDa). Brain regions of interest, including whole slice, dorsal cerebral cortex, lateral cerebral cortex, ventral cerebral cortex, caudate putamen (striatum), lateral septal nucleus, hippocampus, corpus callosum, thalamus, and hypothalamus, were defined using NIH Image J ROI manager. Fluorescent tracer coverage within each region was measured by uniformly thresholding each fluorescence channel and measuring the thresholded area as a % of the overall region area, as described previously (4). The fluorescence area coverage from the 8 brain slices from each animal were averaged to define CSF penetration within a single biological replicate. The effect of age on small and large molecular weight CSF tracer penetration was evaluated independently for each tracer using a 1-way ANOVA with Tukey’s post-hoc test to determine differences between age groups. The relative changes in CSF tracer penetration were plotted by normalizing the small and large molecular weight tracer fluorescence intensity for the young, middle aged, and old groups to the fluorescence intensity of the young animals for each tracer. Size-dependent differences in the rate of decline in CSF tracer penetration were evaluated using a 2-way ANOVA with Sidak’s post-hoc test to determine individual differences in the rate of decline with respect to the young animals

Intraparenchymal radio-tracer injections

Since there was no significant difference between the clearance of Aβ from brain using radio-iodinated Aβ and non-radio labeled Aβ (ELISA) (9), iodinated Aβ was used in these studies. Briefly, a stainless steel guide cannula was implanted stereotaxically into the right caudate nucleus of anesthetized mice with the cannula tip coordinates 0.7 mm anterior and 3 mm lateral to the bregma, and 1.3 mm below the surface of the brain. Animals were allowed to emerge from anesthesia and the experiments were conducted 10–18 hrs after guide cannula implantation, a time point that precedes the start of substantial inflammatory process, as assessed by histological analysis of tissue (negative staining for reactive astrocytes, GFAP, and reactive microglia, anti-phosphotyrosine), but allowed time for the BBB repair for large molecules, as reported (10, 11).

For the injection of the mixture of tracers, artificial CSF (0.5 µL) containing ¹²⁵I-labeled test-tracers Aβ (10 nM monomer) together with ¹⁴C-inulin (0.05 µ Ci; reference molecule) was microinfused into brain ISF over 5 minutes. At the end of the experiments brain was removed and prepared for radioactivity analysis and TCA analyses. Our earlier studies with ¹²⁵I-labeled Aβ have demonstrated that radiolabeled Aβ_1–40 remain mainly intact in brain ISF (> 95%) within 30–300 min of in vivo clearance studies.

For radioactivity analysis, ¹²⁵I radioactivity was determined in a gamma counter (Wallac Vizard Gamma Counter, Perkin Elmer). For ¹⁴C- counting, the samples were solubilized in 0.5 ml tissue solubilizer (Perkin Elmer) overnight, followed by addition of 5 ml of scintillation cocktail (Packard Ultima Gold) and analyzed in a liquid scintillation counter (Beckman Coulter Counter). To differentiate between radioactivity originating from ¹⁴C and ¹²⁵I, the counting window for measuring radioactivity was set to remove ¹²⁵I spillover, while a sample containing ¹²⁵I-Aβ_1–40 equivalent to that which was present in the injectate was measured in parallel with each experiment to monitor spillover of ¹²⁵I radioactivity into the ¹⁴C counting window.

All calculations of clearance parameters were performed as reported previously (4, 6, 10). Briefly, the percentage of radioactivity remaining in the brain after microinjection was determined as % recovery in brain = 100×(N_b/N_i) (eq. 1), where, N_b is the radioactivity remaining in the brain at the end of the experiment and N_i is the radioactivity injected into the brain ISF, i.e., the d.p.m. for ¹⁴C-inulin and the c.p.m. for the TCA-precipitable ¹²⁵I-radioactivity. Inulin was used as a metabolically inert polar molecule which is neither transported across the BBB nor retained by the brain. The percentage clearance of ¹²⁵I-Aβ or ¹⁴C-inulin was calculated from the total, as 100- brain recovery). Differences between ages in of ¹²⁵I-Aβ or ¹⁴C-inulin clearance were evaluated by 1-way ANOVA with Tukey’s post hoc test for multiple comparisons.

In vivo 2-photon fluorescence imaging

Tracer penetration into the intact living brain was visualized through a closed cranial window using a custom built 2-photon laser scanning (Olympus) microscope, as described previously (4). Animals were anethestized, intubated, and ventilated with a small animal ventilator (CWE) while blood gasses and pH were monitored and body temperature maintained at physiological parameters (~100 breaths/min, tidal volume of 0.3–0.4 ml, pO2 = 80–150mmHg, pH 7.25–7.5, body temperature of 37°C). Unilateral craniotomy (3mm in diameter) was performed over the cerebral cortex, 1mm lateral and 0.5mm posterior to bregma. Care was taken to ensure the dura was not ruptured. Immediately following the craniotomy the dura was covered with aCSF and the craniotomy sealed with glass coverslip. The femoral artery was then cannulated for mean arterial blood pressure monitoring and the measurement of arterial blood gas values. Only mice with blood gases within the physiological range were included in the present study. To visualize the vasculature, 0.1ml BBB impermeable Texas Red-dextran 70 (MW 70kD; 1% in saline, Invitrogen) was injected intra-arterially immediately before imaging.

A Mai Tai laser (SpectraPhysics) attached to a confocal scanning system (Fluoview 300, Olympus) and an upright microscope (IX51W, Olympus) was used for in vivo imaging as described previously (4). A 20X (0.9NA) water immersion lens was used to image the cortex, from the surface to a depth of ~250µm. Excitation wavelength was 870nm for FITC and Texas Red. For FITC and Texas Red, emission was collected at 575–645nm. The cerebral vasculature was first imaged with 512×512 pixel frames from the surface to a depth of 240µm with 5µm z-steps. After intracisternal injection of CSF tracer, tracer movement into the cortex was conducted with dual-channel (FITC and Texas Red) 512×512 pixel image acquisition. The cortex was repeatedly scanned from the surface to 240µm below the surface with 20µm z-steps at 1 minute intervals for the duration of the experiment. Image analysis was conducted in with ImageJ software (NIH) with the UCSD plugin set. After imaging, penetrating arterioles were distinguished from penetrating venules on the basis of morphology: surface arteries passing superficially to surface veins and exhibiting less branching at superficial cortical depths. Imaging planes 100–120µm below the cortical surface were selected for the analysis of intracisternal tracer penetration. To define tracer movement into paravascular brain tissue, donut-shaped ROIs were defined that had an external diameter of 150 pixels and an internal diameter of 50 pixels (thus excluding the vessel). These were centered upon the penetrating arterioles. Mean pixel intensity within these ROIs was measured at each time point. Within each animal at each time point para-arteriolar ROIs were separately averaged to generate values for a single biological replicate. When tracer movement along penetrating arterioles was compared, a 2-way repeated measures ANOVA was used followed by Sidak’s test for multiple comparisons.

Measurement of Cerebrovascular Pulsatility

Cerebral vascular pulsatility was evaluated by in vivo 2-photon microscopy as described previously (7). To measure vessel diameters, 3000 ms X–T line scans were acquired orthogonal to the vessel axis in surface arteries, surface veins, penetrating arteries, and penetrating veins (Figure 3B). Penetrating vessel line scans were acquired 50–150 µm below the cortical surface. Vessel diameter was extracted from X–T plots (Figure 3C) by and plotted versus time using custom Matlab and ImageJ software. Steady-state vessel diameters were calculated as the mean value over the 3000 ms epoch. Vessel wall pulsatility (derived units µm*s) was calculated as the absolute value of area under the diameter–time plot, integrated about the running average over the 3000 ms epoch. Differences in vessel pulsatility between different vessel types was evaluated by one-way ANOVA, with Tukey's post hoc test to evaluate differences between vessel segments. The effect of treatment on vascular pulsatility and diameter was evaluated by two-way ANOVA, with Tukey’s post hoc test for multiple comparisons.

Tetramethylammonium (TMA) Microiontophoresis Measurement of Extracellular Volume

All experimental procedures were adapted from the previous studies (6, 12). The single barrel iontophoresis microelectrode (tip diameter of 2–3 µm) contained 150 mM tetramethylammonium (TMA)-chloride and 10 µM Alexa 488. A series of currents of 20 nA, 40 nA and 80 nA were be applied by a dual channel microelectrode pre-amplifier. For measurements of TMA, microelectrodes with an outer diameter of 2–3 µm were fabricated from double-barreled theta-glass using a tetraphenylborate-based ion exchanger. The TMA barrel was backfilled with 150 mM TMA chloride, while the reference barrel contained 150 mM NaCl and 10 µM Alexa 568. All recordings were obtained by inserting the two electrodes to a depth of 150 µm below the cortical surface. Recording electrodes were inserted 2.5 mm lateral and 2 mm posterior to bregma. The electrode tips were imaged after insertion using 2-photon excitation to determine the exact distance between the electrodes (typically ~150 µm). The TMA signal was calculated by subtracting the voltage measured by the reference barrel from the voltage measured by the ion-detecting barrel using a dual-channel microelectrode pre-amplifier. The Nikolsky equation was used for calibration of the TMA electrodes based on measurements obtained in electrodes containing 0.5, 1, 2, 4, and 8 mM TMA-chloride in 150 mM NaCl. The TMA measurements were acquired relative to similar recordings obtained in 0.3% agarose prepared from a solution containing 0.5 mM TMA and 150 mM NaCl. A custom-made MatLab software, ‘Walter’, developed by C. Nicholson was used to calculate α and λ values (6, 12). Differences in extracellular volume fraction and tortuosity were evaluated by 2-way ANOVA with Sidak’s multiple comparisons test.

Evaluating regional CSF tracer penetration, AQP4 and GFAP immunofluorescence

Free floating immunofluorescence staining of 100 µm fixed coronal brain sections was performed to evaluate protein expression patterns of aquaporin-4 (AQP4) and glial-fibrillary acidic protein (GFAP) Slices were blocked with 3% normal donkey serum for 1 hour at room temperature, incubated overnight with primary antibody (rabbit anti-AQP4, 1:500; mouse anti-GFAP, 1:500, both from Millipore) and again incubated for 2 hours at room temperature with secondary antibody (Cy3-conjugated donkey anti-mouse antibody; Cy2-conjugated donkey anti-rabbit antibody; both 1:500; Jackson Immunoresearch).

GFAP and AQP4 expression were evaluated regionally in slices taken from animals injected intracisternally with ovalbumin-conjugated ALEXA 647, then perfusion-fixed 30min post-injection. In this way, GFAP- and AQP4-immunofluorescence could be evaluated in the same animals and brain regions in which CSF tracer penetration was measured. 3-channel whole-slice montages were generated at 4X magnification as above from one anterior and one posterior brain slice from each animal. Fluorescence channels were split and CSF tracer penetration was evaluated as detailed above and as described previously in each brain region. Regional GFAP expression, AQP4 expression and AQP4 polarization were similarly evaluated as described previously (13, 14). To measure regional GFAP expression, regions were uniformly thresholded and the area coverage of GFAP-immunoreactivity (as a % of the whole region area) was measured. To evaluate global AQP4 expression levels, mean AQP4 immunofluorescence intensity was measured within each region. To measure perivascular AQP4 polarization, the median immunofluorescence intensity of perivascular regions was measured. Then a threshold analysis measured the % of the region exhibiting AQP4 immunofluorescence greater than or equal to perivascular AQP4 immofluorescence (AQP4 % Area). ‘Polarization’ was expressed as the % of the region that exhibited lower AQP4-immunoreactivity than the perivascular endfeet ( ‘Polarization’ = 100 – AQP4 % Area). Hence AQP4 ‘Polarization’ is a relative measurement of AQP4 localization: increasing polarization indicating higher perivascular AQP4-immunoreactivity relative to lower parenchymal AQP4-immunoreactivity, whereas reduced polarization indicating lower perivascular AQP4-immunoreactivity relative to higher parenchymal AQP4-immunoreactivity. Differences in regional GFAP expression, AQP4 expression, AQP4 polarization and CSF tracer penetration were evaluated by 2-way repeated measures ANOVA with Sidak’s test for multiple comparisons.

Cerebral arterial pulsatility is impaired in the aging brain

The recirculation of CSF into the brain parenchyma along paravascular pathways is driven in part by arterial pulsation (5–7, 13, 16, 17). We next evaluated pulsation of the cerebrovascular wall by in vivo 2-photon microscopy. The cerebral vasculature was visualized by intra-arterial injection of Texas Red-conjugated dextran (70kD) visualized through a thin-skull cranial window preparation (Figure 3A). Small changes in vessel diameter were measured by high-frequency linescanning of penetrating arteries, and ascending veins between 0–150µm below the cortical surface. As we have recently described (7), a value for ‘vascular pulsatility’ was extracted from X-t (diameter-time) plots (Figure 3B-C), derived from the dilation of the vessel diameter with each cardiac cycle, and integrated across a 3000 ms epoch. In agreement with our prior findings (7), we observed that within the young brain the vessel wall pulsatility of penetrating arteries was significantly greater than the pulsatility of ascending veins (Figure 3D; *P<0.05, Penetrating arteries vs. Ascending veins; 2-way ANOVA with Tukey’s post hoc test for multiple comparisons; n = 8–20 vessels from 4 animals per group). When vascular pulsatility was evaluated in the old (18 months) cortex, we observed that the pulsatility in deep penetrating arteries was significantly reduced compared to that in the young cortex (Figure 3D; *P<0.05, Young vs. Old; 2-way ANOVA with Tukey’s post hoc test for multiple comparisons; n = 8–20 vessels from 4 animals per group). In contrast to penetrating arteries, no age-related changes in vascular pulsatility were observed in ascending veins, (Figure 3E). These data suggest that age-related reductions in penetrating arterial pulsatility may underlie the impairment of paravascular CSF recirculation in the aging brain.

Extracellular volume regulation of the interstitial space is unchanged in the aging brain

Paravascular CSF-ISF exchange is a feature of the sleeping brain. In the awake state, the rate of paravascular CSF recirculation is reduced by ~95% compared to that observed in the brain undergoing natural sleep or anesthesia, while the clearance of interstitial solutes such as amyloid β from the awake brain is more than 2-fold slower than in the sleeping or anesthetized state (6). This appears to be attributable to a 60% expansion in the extracellular volume during sleep, which presumably lowers the total tissue resistance to convective CSF fluxes, facilitating the movement of fluid and solutes through the brain interstitium (6). Thus, in the present study, we sought to evaluate whether a changes in the extracellular space may account for the impaired CSF-ISF exchange in the aging brain. To test this hypothesis, we used in vivo cortical TMA iontophoresis (6, 18) to measure whether the dimensions (extracellular volume fraction, α) or characteristics (tortuosity, λ) of the extracellular space are altered in the aging brain (Figure 4A). In each animal, α and λ values were measured first in the awake state. Animals were then anesthetized and α and λ values were measured a second time to generate paired awake/anesthetized recordings. When extracellular volume fraction were measured, we found that in both the awake and the anesthetized state no significant differences were apparent between the young and the old brains (Figure 4B; P = 0.4957, Young vs. Old; 2-way ANOVA with Sidak’s multiple comparisons test; n = 9–20 per group). However, the significant expansion of the extracellular space between the waking and the anesthetized state was observed both within the young and the old brain (Figure 4B; ***P<0.001). When extracellular tortuosity was measured, there were no significant effects of age upon lambda values in either the waking and anesthetized state (Figure 4C; P = 0.3375, Young vs. Old; 2-way ANOVA with Sidak’s multiple comparisons test; n = 9–20 per group).Thus, the expansion of the extracellular space in the anesthetized state is maintained in the aging brain which suggests that extracellular volume regulation likely does not explain the dramatic reduction in CSF-ISF exchange observed in old vs young mice.

Perivascular AQP4 polarization is impaired in the aged brain

AQP4, an astroglial water channel, is localized in a highly polarized manner to endfoot processes surrounding the cerebral vasculature in addition to the glial limitans externa which abut the pia mater and bound the subarachnoid CSF compartment (19, 20). The perivascular astrocytic endfeet completely ensheath the cerebral vasculature, leaving on 20nm gaps between overlapping processes (21–23). Along these processes, as much as 50 percent of the vessel-facing membrane is populated by crystalline plaques (termed ‘orthogonal arrays of particles’) of AQP4 water channels (24). We have previously demonstrated that perivascular AQP4 facilitates paravascular CSF-ISF exchange (14), and have proposed that the loss of perivascular AQP4 polarization in the injured brain may contribute to the impairment of glymphatic pathway function (13, 14, 16). In the present experiments, we evaluated whether perivascular AQP4 polarization is lost in the aging brain, and whether changes in AQP4 localization differed between perivascular domains surrounding large penetrating arterioles and surrounding microvessels..

Figure 5 shows the differences in perivascular GFAP and AQP4 expression and localization in the young (Figure 5A) and old (Figure 5B) cortex. AQP4 and GFAP immunofluorescence intensity was measured along linear ROIs extending outward from the lumen of penetrating cerebral arterioles or cerebral capillaries, averaged between vessels in the young and old cortex, then plotted as a function of distance from the vessel wall. In the young cortex, AQP4 immunofluorescence surrounding penetrating arterioles and capillaries was highly polarized, with low levels of expression up to the margin of the perivascular endfeet where expression was high (Figure 5C-D). In the old cortex, AQP4 immunofluorescence increased markedly in the tissue immediately surrounding penetrating arterioles (Figure 5C; ***P<0.001 young vs. old, 2-way repeated measures ANOVA; n = 11–12 vessels from 4 animals per group) while AQP4 expression surrounding capillaries was generally elevated, was not as pronounced as in the tissue surrounding penetrating arterioles (Figure 5D). AQP4 expression in the perivascular endfeet immediately surrounding penetrating arterioles was significantly reduced in the old compared to the young brain (Figure 5E; *P<0.05; un-paired t-test, n = 11–18 vessels from 4 animals per group), while no significant difference in AQP4 expression were observed in perivascular endfeet surrounding cerebral capillaries.

Within the same ROIs, GFAP expression was also significantly higher surrounding penetrating arterioles (Figure 5F) and cortical capillaries (Figure 5G; *P<0.05, 2-way repeated measures ANOVA, n = 11–18 vessels from 4 animals per group). Quantification of GFAP expression in perivascular endfeet revealed significantly elevated expression in perivascular endfeet surrounding both penetrating arterioles and capillaries (Figure 5H; ***P<0.001, un-paired t-test; n = 11–18 vessels from 4 animals per group). These results show that the development of diffuse reactive astrogliosis is a general feature of the aging brain. In contrast, the changes in AQP4 expression in the aging brain is more specifically distributed, with marked upregulation of AQP4 and loss of perivascular AQP4 polarization in brain tissue immediately surrounding penetrating cortical arterioles.

We next evaluated whether different brain regions exhibited distinct patterns of AQP4 expression or localization, or reactive astrogliosis with advancing age. Slices from the anterior (Figure 6A-C) and posterior (Figure 6D-F) brains of young (2–3 months) and old (18 months) mice fixed 30 min after intracisternal CSF tracer (OA-45) injection were stained for AQP4 and GFAP. Slices were imaged by 3-channel whole-slice fluorescence microscopy and divided into region-specific ROIs for analysis of AQP4 expression (mean AQP4 immunofluorescence intensity, Figure 6A, D), AQP4 polarization (proportional localization of AQP4 to perivascular vs. parenchymal domains (13, 14), Figure 6B, E) and GFAP expression (% coverage of GFAP immunoreactivity, Figure 6C, F). To show differences in these values, we generated heat maps that show the change in mean AQP4 expression, AQP4 polarization or GFAP expression within each brain region between young and old brains. Global AQP4 expression within each brain region did not significantly change when compared between young and old anterior or posterior brain slices (Figure 6A, D, respectively). In contrast, perivascular AQP4 polarization in the old brains was consistently and significantly reduced compared to the young brains throughout both anterior (Figure 6A) and posterior (Figure 6E) brain regions (*P<0.05, ***P<0.001, Young vs. Old; 2-way repeated-measures ANOVA with Sidak’s multiple comparisons test; n = 4 animals per group). These effects were most consistent within the lateral and ventral cortex, while marked loss of AQP4 polarization was also observed in the aged versus the young hippocampus. Development of reactive astrogliosis, identified by increasing GFAP expression, was similarly apparent throughout the aging brain (Figure 6C, F; *P<0.05, **P<0.01, Young vs. Old; 2-way repeated-measures ANOVA with Sidak’s multiple comparisons test; n = 4 animals per group). As with AQP4 mis-localization, the most pronounced elevation of GFAP expression was in the lateral and ventral cortex and in the hippocampus of the aged compared to the young brain.

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Figure 6

Impairment of perivascular AQP4 polarization is greatest in the lateral and ventral cortex, hippocampus and striatum of the aging brain

AQP4 expression and polarization and GFAP expression were evaluated by immunofluorescence in fixed brain slices from young (2–3 month) and old (18 month) brains. Expression and polarization were evaluated within different regions of anterior (A-C) and posterior (D-F) brain slices. Regional heat maps depict mean change in AQP4 expression (AQP4 immunofluorescence), AQP4 polarization (% area), and GFAP expression (% area) between young and old brains. (A, D) Within both the anterior and posterior regions, global AQP4 immunofluorescence did not differ between the young and old brains. (B, E) Perivascular AQP4 polarization was significantly reduced in the aged brain, with most pronounced effects in the lateral and ventral cortex, striatum and hippocampus (*P<0.05, **P<0.01, ***<0.001, Young vs. Old; 2-way ANOVA; n = 4 per group). (C, F) GFAP expression was significantly increased in the old compared to the young brain, with the greatest effect evident within the lateral and ventral cortex, hippocampus and striatum (*P<0.05, **P<0.01, ***<0.001, Young vs. Old; 2-way ANOVA; n = 4 per group).

The authors thank Dr. Charles Nicholson (New York University) for assistance in the analysis of TMA micro-iontophoresis recordings. This work was supported by funding from NIH/National institute of Neurological Disorders and Stroke (NS078167 and NS078304 to MN and NS073373 to JI) and the American Heart Association (12SDG11820014 to JI).