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Medicine
Chemistry

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Mapping glucose-mediated gut-to-brain signalling pathways in humans

Objectives Previous fMRI studies have demonstrated that glucose decreases the hypothalamic BOLD response in humans. However, the mechanisms underlying the CNS response to glucose have not been defined. We recently demonstrated that the slowing of gastric emptying by glucose is dependent on activation of the gut peptide cholecystokinin (CCK1) receptor. Using physiological functional magnetic resonance imaging this study aimed to determine the whole brain response to glucose, and whether CCK plays a central role. Experimental design Changes in blood oxygenation level-dependent (BOLD) signal were monitored using fMRI in 12 healthy subjects following intragastric infusion (250 ml) of: 1 M glucose + predosing with dexloxiglumide (CCK1 receptor antagonist), 1 M glucose + placebo, or 0.9% saline (control) + placebo, in a single-blind, randomised fashion. Gallbladder volume, blood glucose, insulin, and GLP-1 and CCK concentrations were determined. Hunger, fullness and nausea scores were also recorded. Principal observations Intragastric glucose elevated plasma glucose, insulin, and GLP-1, and reduced gall bladder volume (an in vivo assay for CCK secretion). Glucose decreased BOLD signal, relative to saline, in the brainstem and hypothalamus as well as the cerebellum, right occipital cortex, putamen and thalamus. The timing of the BOLD signal decrease was negatively correlated with the rise in blood glucose and insulin levels. The glucose + dex arm highlighted a CCK1-receptor dependent increase in BOLD signal only in the motor cortex. Conclusions Glucose induces site-specific differences in BOLD response in the human brain; the brainstem and hypothalamus show a CCK1 receptor-independent reduction which is likely to be mediated by a circulatory effect of glucose and insulin, whereas the motor cortex shows an early dexloxiglumide-reversible increase in signal, suggesting a CCK1 receptor-dependent neural pathway.

Introduction
Obesity is now one of the major public health problems facing the world. In 2005, 400 million adults were obese, and this is projected to increase to 700 million by 2015 (www.who.int). This "epidemic" has prompted major scientific efforts to determine the mechanisms underlying the regulation of food intake in order to facilitate the development of effective therapies for obesity. A particular focus has been applied to nutrient-induced gut-to-brain signalling mechanisms. In human research, the development of blood-oxygenation-level-dependent (BOLD) functional magnetic resonance imaging (fMRI) has resulted in the identification of a number of homeostatic and non-homeostatic brain regions associated with the regulation of appetite (Gibson et al., 2010). However, to date, the focus of these human studies has largely been the evaluation of higher centre cognitive responses to food cues (such as images of palatable food), rather than the physiological responses to meal ingestion. The more recent application of the physiological MRI (physMRI) technique (Jones et al., 2012;Lassman et al., 2010) has allowed the exploration of BOLD signal responses to nutrients over time across the whole brain, in particular, in key areas involved in the regulation of food intake such as the brainstem and hypothalamus.
It is well established that the presence of glucose in the small intestinal lumen is associated with decreased hunger, increased fullness, and decreased energy intake at a subsequent meal (Cook et al., 1997;Lavin et al., 1996;Pilichiewicz et al., 2007), but the brain centres and transduction mechanisms underlying these effects are poorly defined in humans. Animal studies have identified the hypothalamus as a critical region for the regulation of energy balance (Brown and Melzack, 1969;Grossman, 1975), and in man, a number of fMRI studies have reported that oral glucose decreases hypothalamic BOLD signal, relative to water (Liu et al., 2000;Smeets et al., 2005aSmeets et al., , 2005bSmeets et al., , 2007Vidarsdottir et al., 2007). However, these studies did not investigate the responses of other sub-cortical brain regions to glucose, and the movement of the head during consumption of oral meals induced imaging artefacts that precluded evaluation of the early responses. In particular, detailed imaging of the human brainstem has not been reported and is critical as it is the first relay point responsible for integrating neural signals transmitted from the upper gut via vagal afferent neurons.
The mechanism by which the presence of glucose in the gut is signalled to the human central nervous system is likely to involve an interaction between direct responses to elevated circulating glucose concentrations (Levin et al., 2004), and indirect responses via glucose sensitive neural (e.g. vagal), or endocrine signalling pathways (e.g. insulin (Liu et al., 2000;Smeets et al., 2005b) and/or gut derived peptides, such as cholecystokinin (CCK) Little et al., 2010) and glucagon-like peptide-1 (GLP-1) (Pannacciulli et al., 2007)). Recent studies have indicated that signals arising from the gastrointestinal tract are likely to play an important role in mediating the hypothalamic response to glucose. For example, the effect of glucose on hypothalamic BOLD signal has been reported to be greater when administered orally compared to intravenously . The hypothalamic response has also been related to fasting insulin concentrations (Liu et al., 2000;Matsuda et al., 1999), and additionally a relationship has been found between postprandial plasma GLP-1 concentrations and increases in BOLD signal in the left dorsolateral prefrontal cortex and the hypothalamus, following ingestion of a liquid meal (Pannacciulli et al., 2007). However, in these previous studies, whole brain responses, and time courses of the response could not be evaluated.
Using physMRI to investigate whole brain responses, we have previously shown that intragastrically administered lipid (the fatty acid, lauric acid (C12)) increases BOLD signal in key human brain areas, including the brainstem and hypothalamus . These activations were abolished by the administration of the CCK 1 receptor antagonist, dexloxiglumide , and modulated by ghrelin (Jones et al., 2012).
We, therefore, decided to apply the same imaging technique to identify and define the brain regions responsive to glucose. Like lipid, glucose ingestion is known to slow gastric emptying (Hunt and Knox, 1968), and to induce gallbladder contraction (de Boer et al., 1993). In addition, dexloxiglumide attenuates the inhibitory effects of glucose on gastric emptying (Fried et al., 1991;Little et al., 2010), suggesting that CCK is also involved in this, and possibly mediating the gut-to-brain signalling response to glucose. Therefore, the CNS response to different macronutrients may be predicted to have significant overlaps.
This study therefore had two aims: first, to determine whether intragastric glucose would induce changes in BOLD signal in the same nutrient-sensitive regions, robustly observed in response to lipid, namely the brainstem and hypothalamus, and second, to determine whether any effects of glucose observed in the CNS were also CCK 1 receptor-dependent.

Methods
Subjects 12 healthy volunteers (7 males, mean (SEM) age: 38 (3.4) (range: 23-60) years) with a non-obese body mass index (BMI: 24 (1) (range: 19.7-28.9) kg/m 2 ) were recruited from the staff and local population of Salford Royal NHS Foundation Trust. Exclusion criteria comprised: presence of any gastrointestinal, endocrine or metabolic disease; consumption of medications that may affect gastrointestinal function, sensation or appetite; metallic implants or objects (e.g. hearing aid, pacemaker), and consumption of greater than 20 g of alcohol or 10 cigarettes/day. All were instructed to abstain from alcohol and vigorous physical activity for 24 h prior to the study. In both the gastric emptying and the fMRI study arms, participants were blinded both to the nature of the intragastric infusions, and to the drug, and studies were performed in a randomised order. Randomisation was performed using a random number generator. Participants provided informed written consent. The study was approved by the Salford and Trafford Research Ethics Committee, and was carried out in accordance with the Code of Ethics of the World Medical Association (Declaration of Helsinki) and the standards established by the author's ethics committee and granting agency. All participants provided informed, written consent after the nature of the experimental procedures provided in writing.

Materials
D-Glucose was purchased from Sigma-Aldrich (Gillingham, England). The CCK 1 receptor antagonist, dexloxiglumide (300 mg tablets), and matched placebo tablets were supplied at no cost by Rottapharm (Monza, Italy) as a scientific collaboration.

Gastric emptying protocol
As the medulla houses the sensory and motor nuclei responsible for relaying vago-vagal enteric circuits, as well as signalling to higher centres, we used the measurement of changes in gastric emptying induced by glucose as a convenient proxy measure to identify a dose of glucose that would have effects on brainstem BOLD response (Little et al., 2010). Participants arrived at the laboratory at 09:00 h after an overnight fast from 22:00 h and orally ingested either 600 mg of dexloxiglumide (1 occasion), or matched placebo (2 occasions), tablets. Previous studies in our laboratory have demonstrated that this dose effectively blocks the effects of lipid (lauric acid (C12)) (Lal et al., 2004) and glucose (Little et al., 2010) on gastric emptying, and the effect of lipid on BOLD signal in the brainstem and hypothalamus . One hour later, a nasogastric feeding tube was positioned, with its distal tip in the stomach. Gastric delivery was employed to eliminate sensory or hedonic effects of oral sweet tasting, and for the MRI studies, movement artefacts during drinking and swallowing. A basal, end-expiratory, breath sample was collected; following which participants received a bolus intragastric infusion of 250 ml glucose (45 g, 180 kcal, 1000 mOsmol) or 250 ml of water (volumetric control). Each meal was labelled with 100 mg 13 Csodium acetate, used as a marker for gastric emptying (CK Gas Products Ltd., UK). For the assessment of gastric emptying rate, end-expiratory breath samples were collected at 5 minute intervals over a period of 45 min (Little et al., 2009(Little et al., , 2010. On each occasion, the subject exhaled through a mouth-piece to collect an end-expiratory breath sample into a 100 ml foil bag, which was then sealed with a plastic stopper, and stored for later analysis. Breath samples were analysed by non-dispersive infrared spectroscopy (IRIS, Wagner Analysen Technik, Bremen, Germany). This enabled evaluation of the isotopic composition of carbon in the expired carbon dioxide (CO 2 ) by determination of the ratio of 13 C: 12 C in the sample (Lee et al., 2000;Sanaka et al., 2006). 13 C-acetate is rapidly absorbed and metabolized after it is emptied from the stomach, hence the rate of appearance of 13 CO 2 in the systemic circulation closely corresponds to the rate of GE (Sanaka et al., 2006). physMRI protocol 12 participants were studied on three separate occasions at 08:30 h following an overnight fast from 22:00 h, to evaluate the spatiotemporal pattern of brain activation in response to intragastric infusion of: (i) glucose (250 ml, 1000 mOsmol (45 g glucose, 180 kcal)) following two placebo tablets, (ii) glucose (250 ml, 1000 mOsmol) following the CCK 1 receptor antagonist, dexloxiglumide (600 mg (two 300 mg tablets)) (glucose + dex), and (iii) 0.9% saline (control) following two placebo tablets. The dexloxiglumide (condition ii), or matched placebo, tablets (conditions i and iii), were orally ingested with 100 ml of water 1 h prior to the commencement of the intragastric infusion. All conditions were matched exactly to our previous studies (Jones et al., 2012;Lassman et al., 2010) to enable direct comparison between the glucose-and lipid-induced nutrient matrices. In the study design we purposefully omitted a saline + dex control arm since we had previously demonstrated, under the same study conditions (i.e. after an overnight fast and with identical imaging parameters), that there was no effect of dexloxiglumide in the absence of nutrient (i.e. during intragastric saline infusion) on BOLD signal across the whole brain . Furthermore, dexloxiglumide has no effect on the gastric emptying of water/saline (Little et al., 2010). Due to the large number of statistical comparisons to be performed, which do not require the placebo/placebo condition, and mindful of burden on experimental subjects, we considered that repeating this was unjustified.
Thirty minutes prior to scanning, participants were intubated with a nasogastric feeding tube and an intravenous cannula was positioned in an antecubital vein to enable repeated blood sampling. Participants were then positioned in the scanner (3.0 T Philips Achieva MR System, Magnetic Resonance Imaging Facility (MRIF), Salford Royal NHS Foundation Trust) and images of the gallbladder acquired. Immediately following this, participants were repositioned in the scanner for the 30 minute fMRI scan. After a baseline period of 4 min, the intragastric infusion (glucose or saline) was administered as a bolus, using a syringe via the nasogastric tube, over a period of 1 min. Subjective ratings of fullness, hunger and nausea were recorded every 5 min throughout the scan. Venous blood samples were taken to assess blood glucose, and plasma insulin, CCK and GLP-1 concentrations at the start of the scan protocol (t = 0 min), and then at t = 5, 10, 15, 20, and 25 min following the intragastric infusion. After the fMRI scan, a second gallbladder image was acquired. On the first visit, participants were positioned in the scanner an additional 10 min before the commencement of the scanning protocol to enable a high-resolution structural scan to be performed to more accurately normalise each individual's brain anatomy.

Gallbladder MR acquisition and volume measurement
Gallbladder volume, which is a classical in vivo bioassay for CCK activity, was determined to evaluate the functional significance of the CCK response to the glucose test meals and as a positive control for CCK 1 receptor blockade by dexloxiglumide (Lilja et al., 1982). Complete blockade of gallbladder contraction demonstrates effective CCK 1 receptor antagonism. Volumes were determined from single-shot, T2 weighted axial, coronal and sagittal 3 T anatomical MR images of the upper abdomen obtained at the beginning and end of each study using a SENSE torso coil triggered by respiration. Gallbladder volume was estimated using the ellipsoid method, whereby volume = length × depth × width × 0.523 (where π/6 = 0.523, depth = maximum antero-posterior diameter and width = maximum lateral diameter) (Dodds et al., 1985;Lassman et al., 2010).

Sensation scores
Subjects were asked to indicate their sensations of hunger, fullness and nausea on a 10 point scale, every 5 min during each scan. The subject rated each sensation by moving a pointer along the scale, via a response box held in their right hand .

Blood glucose, plasma insulin, CCK and GLP-1 measurements
Venous blood samples (10 ml) were collected into ice-chilled lithium heparin-treated tubes containing 400 kIU aprotinin per ml blood (Trasylol; Bayer Ltd., Berkshire, UK). Plasma was separated by centrifugation at 3000 rpm for 10 min at 4°C within 1 h of collection, and stored at −70°C until assayed.
Venous blood glucose concentrations (mmol/l) were determined at the time of collection by the glucose oxidase method using a portable glucose meter (Medisense Optium Xceed, Abbott, Alameda CA USA). The accuracy of this method has been confirmed using the hexokinase technique (Horowitz et al., 1991).
MRI acquisition fMRI brain volumes were acquired at 5 second intervals for 30 min using a multi-slice, single-shot EPI sequence to achieve whole brain coverage: 40 axial slices, TR/TE = 5000/35 ms, voxel size = 1.8 × 1.8 × 3.5 mm, flip angle = 79°, SENSE acceleration factor = 1.8, matrix = 128 × 128, FOV = 240 × 240 mm. Daily quality assurance was carried out to ensure a consistent high signal-to-ghost ratio using an automated quality control procedure. A SENSE 8 channel head coil was used for radio frequency transmission and reception.

MRI image analysis
The fMRI data were analysed using Statistical Parametric Mapping (SPM8, www.fil.ion.ucl.ac.uk/spm). Images were realigned, spatially normalised and smoothed to facilitate inter-subject averaging. The limits on head movement for inclusion were: 2 mm for x-and ytranslation and 4 mm for z-translation, both per volume and across the whole scan, and 3°for yaw and 1.5°for roll and pitch, again per volume and across the whole scan.
First-level analysis was performed, using the p-block ph/physMRI analysis technique (Jones et al., 2012;Lassman et al., 2010;McKie et al., 2005McKie et al., , 2011, on each subject for each study condition in the following ways: the pre-and post-infusion scans were split into two minute time-bins. The post-infusion scans were divided into 13 consecutive 2 minute time bins (T1-13) and the 24 scans from the 2 min immediately prior to the glucose/saline infusion formed the baseline time bin (T0). In each subject and condition, the signal averages for the 13 post-infusion time bins (T1-T13) were separately compared to the baseline average (T0) using regression within the general linear model framework. This resulted in 13 first level images corresponding to the BOLD signal change from baseline in each successive post-infusion time bin for each subject and condition. The realignment parameters for each session were included as regressors of no interest to correct for motion. Scanner drift was modelled using the scaling option for global normalisation in SPM. Contrast maps across time for glucose vs. saline and glucose vs. glucose + dex were calculated for each subject.
To determine whether statistically significant increments in the BOLD signal change from baseline across subjects occurred over time, two repeated-measures ANOVAs were used, one for the glucose vs. saline comparison and the other for the glucose vs. glucose + dex comparison, at a cluster-level statistical inference of p(False Discovery Rate; FDRc) b 0.05 at a height threshold of p = 0.005.
Correlations were performed to determine relationships between changes in BOLD signal with blood glucose, plasma GLP-1 and insulin concentrations, and the subjective ratings of nausea, hunger and fullness. These were performed using the average values per subject (area under the curve (AUC) and across time) for each significant cluster extracted from the glucose vs. saline and glucose + dex vs. saline comparisons. For the AUC correlations, all time bin BOLD signal change contrasts were averaged per cluster and compared to the baseline corrected AUC blood concentrations for each individual. For the correlations with time, each cluster's average BOLD signal at five time bins (T + 5, T + 10, T + 15, T + 20 and T + 25 min) was calculated and correlated to the each subject's blood concentration measures at the same time points. This resulted in a correlation coefficient, r, for each subject. Each subject's r-values were then entered into a one-sample t-test. All correlations are reported at a False Discovery Rate level (pFDR b 0.05) correcting for the number of cluster significant regions being tested.

Analysis of gastric emptying, blood glucose and GI hormone concentrations and appetite perceptions
Gastric emptying, blood glucose, plasma CCK, GLP-1 and insulin concentrations are displayed as raw data. VAS scores were analysed as change from baseline values. Areas under the curve were calculated for gastric emptying and blood glucose, CCK, GLP-1 and insulin concentrations using the trapezoidal rule. These data were analysed using repeated measures ANOVAs with treatment, and time (or for the AUC data, treatment) as factors. If ANOVAs revealed statistical significance, post-hoc paired comparisons, corrected for multiple comparisons using Bonferroni's correction, were performed. Statistical analysis was performed using SPSS Version 17.0. Data are presented as means (SEM).

Results
The study was well tolerated by all participants. Due to difficulty obtaining blood samples from 2 of the participants while in the scanner, complete data sets for glucose, insulin, CCK and GLP-1 are available from only 10 participants. Three participants self-reported smoking ≤ three cigarettes/day. There were no apparent differences in gastric emptying, BOLD response or gut peptide secretion between the smokers and nonsmokers, but this could not be formally evaluated due to the small numbers.

Gastric emptying
As in our previous study (Little et al., 2010), we demonstrated an effect of treatment on gastric emptying (p = 0.003). Intragastric glucose slowed gastric emptying when compared with water (p = 0.017), and dexloxiglumide abolished this glucose-induced inhibition (p = 0.031), with no difference between the emptying of water and glucose + dex. The slower gastric emptying observed after glucose consumption is therefore CCK 1 -receptor dependent (Fig. 1A).

Gallbladder volumes
There was an effect of intragastric glucose on gallbladder volume (Fig. 1B). Glucose decreased the volume of the gallbladder (p = 0.014) when compared with saline. As with gastric emptying, the effect of glucose was attenuated by dexloxiglumide pre-treatment (p = 0.008). This provides evidence of fully effective CCK 1 receptor blockade by dexloxiglumide.

Areas of glucose-induced BOLD signal change
Results of the one-way ANOVAs investigating the differences in BOLD signal responses over time following intragastric glucose (relative to saline) are presented in Table 1 and Figs. 2 and 4. A significant effect of time was observed in sections of the brainstem (Figs. 4A, B, C) and hypothalamus ( Fig. 4D) (regions previously reported to show a positive BOLD response between lipid and saline (Jones et al., 2012;Lassman et al., 2010)), as well as the cerebellum, right fusiform and lingual gyri, insula/putamen, left parahippocampal gyrus, right middle temporal and thalamic regions. Investigation of the average time courses for these significant clusters highlighted that the effect of time was generally driven by an initial increase in BOLD signal followed by sharp decrease at 5 min (Fig. 4).

Effect of dexloxiglumide on areas of BOLD signal change
Dexloxiglumide pre-treatment significantly modulated the BOLD signal response over time in the regions summarised in Table 2 and Fig. 3. Investigation of the average time courses for the significant clusters highlighted partial blockade of the effect of glucose to decrease BOLD signal in the hypothalamus (Fig. 4D). The supplementary motor area is of particular interest as this was the only region to show a positive BOLD response to glucose ingestion (glucose vs. saline) over time, Fig. 1. (A) Gastric emptying (n = 3/12) and (B) change in gallbladder volume (n = 12) following intragastric glucose (1000 mOsmol), glucose (1000 mOsmol) + dexloxiglumide (600 mg) or saline/water control. Data are mean (SEM). *glucose vs. water/saline,^glucose + dex vs. glucose, p b 0.05. an effect that was abolished by dexloxiglumide (Fig. 4E). Investigation of the average time course in the other brain regions in Table 2 showed that effect of time was driven in two separate ways: 1) for the cerebellum, left caudate, left and right cuneus, and left lingual gyrus, the effect of time was driven by the BOLD response for glucose + dex vs. saline being further reduced relative to glucose vs. saline, and 2) for the other regions the effect of time was driven by the initial glucoseinduced increase in BOLD signal and subsequent sharp decrease at 5 min being abolished by dexloxiglumide pretreatment.

Time course of changes in BOLD signal
There were significant correlations between the time course of BOLD signal changes with the time course of changes in circulating blood glucose and insulin concentrations in all of the brainstem, hypothalamic, cerebellar, fusiform and thalamic regions responding to glucose (Table 1). This temporal correlation was removed by pretreatment with dexloxiglumide in all of these regions, apart from the lateral cerebellum and the right occipital cortex (fusiform and lingual gyrus). A temporal correlation between the glucose + dex vs. saline BOLD signal with insulin was also observed in the left parahippocampal region. There were no significant correlations between BOLD signal changes with the AUC of blood glucose, GI hormones or subjective appetite measures. There were no significant temporal correlations between BOLD signal changes with GLP-1, or any of the subjective appetite measures. With respect to clusters that showed a significant modulation of the glucose-induced BOLD signal by dexloxiglumide, there were correlations between the time course of the BOLD signal change with the time course of blood glucose and insulin concentrations in the hypothalamus, cerebellum, right putamen, left lingual gyrus and left precuneus ( Table 2). There were no temporal correlations between the time course for blood glucose and the time course for the glucose + dex vs. saline BOLD signal in any region, however, there were temporal correlations observed with insulin and GLP-1 in the cerebellum, bilateral cuneus and left lingual gyrus i.e. the clusters that showed an the effect of time driven by the BOLD response for glucose + dex vs. saline being further reduced in comparison to glucose vs. saline. Interestingly, in the supplementary motor area, where BOLD signal increased in response to glucose (and where the effect was reversed by dexloxiglumide), there was no temporal correlation with circulating blood glucose, insulin or GLP-1 concentrations ( Table 2). There were no significant correlations between BOLD signal changes and the AUC of blood glucose, hormones or subjective appetite measures. There were no significant temporal correlations with BOLD for any of the subjective appetite measures.

Blood glucose, plasma insulin, CCK and GLP-1 concentrations Glucose
There was an effect of treatment on blood glucose concentrations (p = 0.002). Both glucose (p = 0.001) and glucose + dex (p = 0.009) increased blood glucose concentrations when compared with saline, with no difference between glucose and glucose + dex (p = 1.000) (Fig. 5A).

Insulin
There was an effect of treatment on the AUC for serum insulin concentrations (p b 0.001). Both glucose (p = 0.003) and glucose + dex (p = 0.001) increased serum insulin concentrations when compared with saline. There was a treatment * time interaction (p b 0.001). Insulin concentrations were greater for glucose and glucose + dex between t = 5-25 min, when compared with saline, and greater for glucose + dex when compared with glucose at t = 25 min (Fig. 5B).

CCK
There was an effect of treatment on the AUC for plasma CCK concentrations (p = 0.002). CCK was increased following glucose + dex when compared with glucose (p = 0.005) and saline (p = 0.007), with no differences between glucose and saline (p = 0.395) (Fig. 5C).

GLP-1
There was an effect of treatment on the AUC for plasma GLP-1 concentrations (p = 0.001) (Fig. 5D). Glucose increased plasma GLP-1 concentrations when compared with saline (p = 0.047), with no difference between glucose and glucose + dex, or glucose + dex and saline.

Perceptions of hunger, fullness and nausea
There were no differences in baseline ratings, nor any effect of any treatment, on perceptions of nausea (Fig. 6A) or hunger (Fig. 6B). There was an effect of treatment for fullness scores (p = 0.05). Glucose increased fullness when compared with saline (p = 0.009), with no differences between saline and glucose + dex (Fig. 6C).
Changes in fullness were not correlated to circulating levels of glucose, CCK, GLP-1 or insulin. Ratings of hunger were weakly related to plasma insulin (R = 0.36, P = 0.052), but not CCK, glucose or GLP-1, concentrations.

Discussion
The development of whole brain fMRI applied to intragastric infusion of nutrients (physiological MRI; physMRI) has provided an entirely new means to understand the gut-to-brain signalling mechanisms regulating gastrointestinal function, appetite and food intake in humans (Jones et al., 2012;Lassman et al., 2010). We have now, for the first time, demonstrated the whole brain responses to glucose when administered directly into the stomach to avoid the confounding effects of sight, taste or smell, and overcoming the technical issues associated with movement during swallowing. In the medulla, pons, and midbrain regions of the brainstem and in the hypothalamus (and other areas outlined in Table 1) glucose, compared to the saline, decreased the BOLD signal at 5 min, after a small initial increase. The brainstem and hypothalamic regions have previously been identified to respond to intragastric lipid, using identical methods (Jones et al., 2012;Lassman et al., 2010). However, in response to glucose, the decrease in BOLD Fig. 4. Changes in blood-oxygenation-level-dependent (BOLD) signal over time in the medulla (A), pons (B), midbrain (C), and hypothalamus (D), and motor cortex (E), following bolus intragastric glucose (1000 mOsmol) and glucose + dex, relative to saline. signal was temporally correlated to the increases in blood glucose and plasma insulin concentrations.
Since CCK clearly mediated gastric and gall bladder emptying following intragastric glucose, the role of CCK in the CNS response to intragastric infusion of glucose was investigated using a pretreatment with dexloxiglumide (a CCK 1 receptor antagonist). Investigation of the interaction between glucose and glucose + dex over time revealed several interesting results. There was a partial blocking of the effect of glucose to decrease BOLD signal in the hypothalamus by dexloxiglumide. Dexloxiglumide furthermore abolished the effect of glucose in the motor cortex, the only region to show a BOLD signal increase in response to glucose. In the cerebellum, left caudate, bilateral cuneus and left lingual gyrus, dexloxiglumide pre-treatment further reduced the glucose BOLD response. Interestingly, there was a significant temporal correlation with insulin and GLP-1 in all these regions apart from the left caudate. For the remaining regions dexloxiglumide changed the temporal characteristics of the glucose response by abolishing the initial increase in BOLD signal as observed for glucose vs. saline.
With regard to the areas showing decreased BOLD signal, our observations confirm previous reports of a decrease in BOLD signal within the hypothalamus following either oral or intravenous glucose in humans (Liu et al., 2000;Matsuda et al., 1999;Smeets et al., 2005aSmeets et al., , 2005bSmeets et al., , 2007Vidarsdottir et al., 2007), and rats Tsurugizawa et al., 2008), and for the first time define the brainstem response to the presence of glucose in the GI tract in humans. In seeking to explore candidate pathways for these effects of glucose to decrease BOLD signal in the CNS regions outlined in Table 1, our dexloxiglumide data indicate that, unlike the response to lipid , the response to glucose in these regions is not solely dependent on the activation of CCK 1 receptors on vagal afferents. Even though we demonstrated that glucose does activate processes known to be CCK-dependent (i.e. dexloxiglumide reversible gallbladder contraction and slowing of gastric emptying (Fried et al., 1991;Little et al., 2010;Moran et al., 1997)), there was no significant effect of dexloxiglumide on BOLD signal reduction in these regions. This suggests that although glucose is likely to have activated a subset of neurons via the CCK 1 receptor (as demonstrated by the dexloxiglumide sensitive increase in BOLD signal in cortical areas and possibly the small initial rise in BOLD signal observed prior to the decrease in the brainstem), there was also an additional, perhaps more direct, inhibitory action of circulating glucose and insulin, which may have occurred at the level of peripheral vagal afferents, and/or the brainstem and hypothalamus. This is supported by the observation that in rats, BOLD signal responses in the lateral hypothalamus, as well as other brain regions, to post-oral glucose were the same in both intact and vagotomised rats, whereas BOLD signal responses to glutamine were attenuated in vagotomised rats. This suggests that specific responses occur to different types of nutrients, with vagal activity being critical for the sensing of glutamine and lipid , and humoral factors, such as insulin, as well as changes in circulating glucose levels, being important for glucose detection (Tsurugizawa et al., 2009). To this effect a direct comparison of our independent studies using intragastric lipid and glucose has been provided as supplementary data.
It is recognised that elevated glucose levels can directly influence the activity of specific glucose-sensing neurons in hypothalamic and brainstem regions (Levin et al., 2004), and that there are subsets of neurons in which elevated glucose levels are either excitatory or inhibitory (Grabauskas et al., 2010;Levin et al., 2004). In addition, elevated blood glucose may have been signalling to the brain via activation (excitation or inhibition) of peripheral glucose-sensing neurons in the nodose ganglia (Grabauskas et al., 2010). Animal studies strongly indicate that vago-vagal pathways play an important role in the regulation of gastrointestinal motility (Zhou et al., 2008) and eating behaviour (Grabauskas et al., 2008) in response to acute changes in blood glucose levels. Additionally, acute increases in glucose levels modulate synaptic connections between vagal afferent nerve terminals and neurons in the nucleus tractus solitarius (Wan and Browning, 2008). However, while one study has demonstrated an effect of intravenous glucose on BOLD signal in the hypothalamus , this effect was not replicated by others, despite inducing comparable increases in blood glucose concentrations (Purnell et al., 2011). A number of studies have reported no relationship between blood glucose concentrations and glucose-induced changes in BOLD signal in the hypothalamus (Smeets et al., 2005a. Here, we have demonstrated a strong correlation between the timing of changes in blood glucose and plasma insulin levels, and the decrease in BOLD signal, suggesting that changes in circulating glucose and insulin concentrations are likely to be key mediators of the central response to glucose, rather than direct vagal afferent signalling from the gut. While we did demonstrate that dexloxiglumide accelerated gastric emptying of glucose, this was not of a sufficient magnitude to modulate circulating glucose or insulin levels, and indeed, in the CNS regions in which we observed a significant relationship with glucose and insulin (i.e. those regions in which BOLD signal decreased), there was no difference in effect between glucose and glucose + dex conditions. Hence, gastric emptying rate is unlikely to be a major determinant of the CNS response reported here.
It appears from our data that the onset of the BOLD signal rise in the motor cortex was rapid, occurring within 5 min post-infusion and before any measurable elevation of plasma glucose. Further analysis of the direct comparison between glucose and lipid BOLD responses, using a two-way repeated measures ANOVA with nutrient (vs. saline) as a between subject factor and time as a within subject factor, highlighted the spatial similarity and temporal interaction between glucose and lipid (Jones et al., 2012;Lassman et al., 2010) related changes in BOLD signal (Supplementary material). The data included in this comparison were collected in parallel on the same scanner with the same imaging parameters during the same time period. The motor cortex was not significant in this analysis due to a lack of direct overlap in the motor cortex for glucose and lipid however the time course for lipid from the motor cortex in Fig. 3D from Lassman et al., 2010 and in Fig. 4A in this paper are very similar (Supplementary Fig. 1A). Previous studies have also demonstrated that cortical areas (including the motor cortex) display an increase in BOLD signal during intravenous glucose (Purnell et al., 2011). That this response was blocked by dexloxiglumide suggests that the early changes in BOLD signal were mediated by a peripheral vagal CCK 1 receptor-dependent pathway. This is supported by a recent study that identified a positive relationship between changes in BOLD signal in the thalamus, amygdala, caudate, insula and orbitofrontal cortex with changes in plasma CCK concentrations following oral glucose ingestion (Li et al., 2012). The current study however went beyond pure observation and utilised the CCK 1 receptor antagonist dexloxiglumide, which we have previously shown to abolish the effect of lipid. Therefore this correlation cannot be causal: the fact that BOLD changes are in opposing directions in our study would make an identical signalling molecule driving both rather implausible. We validated that CCK activity was fully blockaded in our study by demonstrating the abolition of glucose-induced gallbladder contraction: measuring peripheral blood levels of CCK is clearly a poor assay of biological activity in this system, since much of the effects of CCK occur at a paracrine level within the gut. Where possible, physiology is better explored by using interventions such as receptor blockade.
It therefore appears that the effects of intestinal glucose are mediated by at least two nutrient sensing pathways in the CNS, both of which may modify appetite, eating behaviour and digestive function either directly (e.g. central detection of circulating insulin/glucose concentrations) or indirectly (e.g. activation of vagal afferents via gut peptides).
It should also be noted that intragastric infusion, while having benefits of ensuring consistent rate of nutrient delivery to the intestine and allowing a pure focus on gut-to-brain signals, does not entirely reflect normal meal ingestion due to the exclusion of oral and cephalic phases. For example, our analyses did not reveal relationships between BOLD signal changes with appetite perceptions. These analyses are likely to be underpowered since we were unable to collect blood from two participants while they were in the scanner. Additionally, subjective ratings of hunger and fullness in several subjects showed no change from preinfusion baseline and, therefore, could not be correlated with BOLD responses. We ended up with data for n = 6 for hunger, and n = 7 for fullness, in these models. In the included subjects, the increase in fullness and reduction in hunger were very small in magnitude and blood and VAS scores were collected at 5 min intervals, therefore there was less temporal resolution in comparison to the initial BOLD analysis which was done using 2 minute time bins. The small magnitude of change in VAS scores was not unexpected, since it is well established that intragastric infusion has less potent effects on appetite perceptions than oral consumption (Cecil et al., 1998). Future work using this paradigm should investigate the effects of these additional components during meal ingestion.
Of interest, it has recently been shown that GLP-1 in response to lipid is inhibited by dexloxiglumide indicating dependence on CCK receptor activation (Beglinger et al., 2010). This dependence does not appear to be the case in response to glucose stimulation and probably implicates a direct action of glucose on GLP-1 producing enteric mucosal cells.
The other brain regions responding to glucose, including the cerebellum, thalamus, precuneus and motor cortex have been consistently demonstrated in our previous studies to respond to nutrient ingestion and or gut hormones (i.e. ghrelin) (Jones et al., 2012;Lassman et al., 2010), suggesting that they play a role in mediating the gastrointestinal and appetite responses to ingested food. In particular, the cerebellum Fig. 6. Effects of saline, glucose and glucose + dexloxiglumide on perceptions of nausea (A), hunger (B) and fullness (C). There were no differences in baseline ratings, nor any effect of any treatment, on perceptions of nausea or hunger. There was an effect of treatment for fullness scores (p = 0.05). Glucose increased fullness when compared with saline (p = 0.009), with no differences between saline and glucose + dex. *P b 0.05 glucose vs. saline. has been shown to play an important role in food intake (Zhu and Wang, 2008).
In conclusion this study provides important new information about whole brain responses to intragastric glucose in man, and demonstrates that multiple mechanisms operate to modulate BOLD signal in the nutrient-induced brain matrix.
Disclosures TJL, SM, RBJ, RMC, JT and DGT have no conflict of interest to declare. MD is an employee of Rottapharm, Monza, Italy.
Contribution of the authors TJL: overall organisation and performance of the study, protocol design, data analysis, manuscript preparation.
SMc: physMRI protocol design and data analysis, drafting of the manuscript.
RBJ: conducting the study, data analysis and drafting of the manuscript.
MD: supply of study drug, protocol design, drafting of the manuscript. JMcL and DGT: secured grant, protocol design, data interpretation, and drafting of the manuscript.


extfieldsofstudy: Biology Medicine

text:
Genetic deficiency of neuronal RAGE protects against AGE-induced synaptic injury

Synaptic dysfunction and degeneration is an early pathological feature of aging and age-related diseases, including Alzheimer's disease (AD). Aging is associated with increased generation and deposition of advanced glycation endproducts (AGEs), resulting from nonenzymatic glycation (or oxidation) proteins and lipids. AGE formation is accelerated in diabetes and AD-affected brain, contributing to cellular perturbation. The extent of AGEs' involvement, if at all, in alterations in synaptic structure and function is currently unknown. Here we analyze the contribution of neuronal receptor of AGEs (RAGE) signaling to AGE-mediated synaptic injury using novel transgenic neuronal RAGE knockout mice specifically targeted to the forebrain and transgenic mice expressing neuronal dominant-negative RAGE (DN-RAGE). Addition of AGEs to brain slices impaired hippocampal long-term potentiation (LTP). Similarly, treatment of hippocampal neurons with AGEs significantly decreases synaptic density. Such detrimental effects are largely reversed by genetic RAGE depletion. Notably, brain slices from mice with neuronal RAGE deficiency or DN-RAGE are resistant to AGE-induced LTP deficit. Further, RAGE deficiency or DN-RAGE blocks AGE-induced activation of p38 signaling. Taken together, these data show that neuronal RAGE functions as a signal transducer for AGE-induced synaptic dysfunction, thereby providing new insights into a mechanism by which the AGEs–RAGE-dependent signaling cascade contributes to synaptic injury via the p38 MAP kinase signal transduction pathway. Thus, RAGE blockade may be a target for development of interventions aimed at preventing the progression of cognitive decline in aging and age-related neurodegenerative diseases.

Advanced glycation endproducts (AGEs) are members of a heterogeneous class of molecules, which modify cellular function by distinct mechanisms, including ligation and activation of signal transduction receptors. The products of non-enzymatic glycation (or oxidation) of proteins and lipids, AGEs contribute to the normal aging process and when accelerated have a causative role in the vasculature complications of diabetes mellitus and several neurodegenerative diseases, including Alzheimer's (AD), Parkinson's, and Huntington's diseases. [1][2][3][4][5] In diabetic patients, the concentration of circulating AGEs (serum AGE level) has been reported at 7.2-22 mU/ml (equivalent to 30-88 mg/ml AGE-BSA), which is significantly higher than that of non-diabetic patients (3 mU/ml, equivalent to 12 mg/ml AGE-BSA). [6][7][8] The brain AGE level was also increased to 5-6 mM (equivalent to 325-390 mg/ml AGE-BSA) in the diabetic animal model. 9 Excess AGE accumulation is detrimental to neurons and is believed to be a key to the pathogenesis of cognitive decline in normal aging and specific chronic diseases of aging. For example, in a recent clinical study, peripheral AGE levels were associated with cognitive decline in older adults with and without diabetes. 10 Diabetes complications affect the brain, increasing risk for depression, dementia, and AD. In fact, patients with type 2 diabetes are at twofold to threefold increased relative risk for AD [11][12][13][14][15][16][17][18] and accelerated cognitive dysfunction.
Long-lived proteins such as b-amyloid peptide (Ab) and hyperphosphorylated tau protein that accumulate in AD brain are highly susceptible to AGE modification. [19][20][21][22] AGEmodified Ab or tau protein results in increased oxidative stress and chronic inflammation, accelerating AD pathology and neuronal perturbation. 19,20,[22][23][24][25] Moreover, Ab or tau glycation results in increased aggregation and subsequent formation of senile plaques or neurofibrillary tangles, the major pathological feature of AD, 19,22 suggesting that AGE modification is an important risk factor for neurodegenerative diseases. 26 Although increased accumulation of AGEs in brain, as seen in aging, diabetes, or neurodegenerative diseases, speeds up oxidative damage to neurons contributing to synaptic dysfunction and cognitive decline, its underlying mechanisms are not well understood.
Receptor for advanced glycation endproduct (RAGE) was first identified as a cell surface receptor of the immunoglobulin superfamily for AGEs. 27,28 Increased expression of RAGE occurs in neuronal and non-neuronal cells in the peripheral and central nervous system in aging, diabetes, and ADaffected individuals, where RAGE ligands are upregulated. 29,30 Although it has been shown that AGEs-RAGE interaction contributes to cellular perturbation relevant to the pathogenesis of the cardiovascular disease and the diabetes vascular complications, [31][32][33] little is known about the role of AGEs and its interaction with RAGE on synaptic dysfunction. To understand the mechanisms involved in AGE-mediated synaptic damage, the following questions need to be addressed: (1) 'Do AGEs alter synaptic structure and function? If so, are these changes dependent on RAGE signaling?'; (2) 'Does RAGE blockage by genetic depletion protect from AGE-induced synaptic dysfunction and loss?'; and (3) 'What is the impact of neuronal RAGE in AGE-induced aberrant synaptic function?'. Thus it is important to evaluate the impact of AGEs-RAGE interaction on synaptic dysfunction and to explore the mechanism underlying AGE-RAGEdependent signal transduction and its contribution to synaptic damage.
Here we investigate neuronal RAGE signaling in AGEinduced synaptic injury using our novel conditional RAGE knockout mice targeted to cortical neurons as well as transgenic mice that overexpress signal transduction-deficient mutants of RAGE in neurons. Given that neuronal and non-neuronal cells in the brain may contribute to AGE-induced sustained neuronal and synaptic stress and dysfunction, we assessed the impact of global RAGE deletion in this setting and further delineated the mechanism by which RAGEdependent activation of p38 MAP kinase potentiates AGEinsulted synaptic injury.

Results
Generation and characterization of conditional neuronal RAGE selective knockout (nRKO) mice. We developed a model system in which neuronal expression of RAGE, in particular in the brain region responsible for learning and memory, is deleted so that consequences of receptor-ligand interaction could be assessed in an AGE-rich environment.
To this end, we have recently created Cre recombinase RAGE null mice, termed Tg nRKO mice, crossing neuronaltargeted and region-restricted Tg mice that express Cre recombinase in the entire forebrain under the control of the forebrain-specific calcium-calmodulin-dependent kinase II (CAMKIIa, CK2) promoter 34 to generate deletion of neuronal RAGE in the entire hippocampus and cortex. We used two Cre-loxP systems to generate the targeting vector and the RAGE/flox mice (performed by Ozgen Inc., Bentley DC, WA, Australia). Figure 1a demonstrates a targeting vector containing the 5 0 homology arm, the 3 0 homology arm, and loxP arm. Two loxP sites flanking RAGE exons 2-4 allow for Cre-mediated deletion using Cre recombinase (Figure 1b). Excision of exons 2-4 by Cre recombinase results in a frame shift and early stop codon from the mouse RAGE sequence, which block RAGE expression. We first made homozygous RAGE floxed allele mice (RAGE flox/flox ); then RAGE flox/flox mice were crossed with neuronal target CK2 mice to generate nRKO mice (RAGE flox/flox /CK2-Cre), which were then verified by analysis of tail DNA with PCR amplication using primers for flox (700 bp) and Cre transgene (300 bp) ( Figure 2a). Immunoblotting of cortical homogenates with anti-RAGE antibody showed significant reduction of RAGE expression levels in cerebral cortex of nRKO mice compared with non-Tg mice (B80% reduction versus non-Tg brain, Po0.01) (Figure 2b). Incomplete suppression of RAGE expression was due to brain homogenates, which contained non-neuronal cells. RAGE expression levels were not altered in cerebellum homogenates (Figure 2c). Confocal microscopy with double immunostaining of RAGE and MAP2 clearly demonstrated that there was virtually no RAGE signal in MAP2-positive neurons in cortex (Figures 2d-e) and hippocampus (Figure 2f) of nRKO mice, thereby verifying RAGE depletion in cortical neurons in the brain of nRKO mice.
Immunochemical characterization of Ne-carboxymethyllysine (CML) in AGE preparation. Of all forms of AGEs, CML is a major glycation product known to be stable and accumulate progressively in vivo in aging, AD, and  diabetes-associated complications. 21,35 To verify whether CML was a major component in our AGE preparation, we prepared AGEs by incubation of bovine serum albumin (BSA) with glucose 6-phosphate for 6-8 weeks at 37 1C and then performed immunochemical characterization using a monoclonal antibody targeting CML adducts. Using Coomassie blue dye for rapid reversible staining of protein bands in gel, we observed co-migration of BSA and AGE-modified BSA at B66 and B70 kD, respectively ( Figure 3a). The increase in molecular weight of glycated BSA is due to AGE modification, which results in intramolecular crosslinking. Immunoblotting of the same preparation of BSA and AGE-BSA using specific antibody to CML showed prominent CML immunoreactivity only in the presence of AGEs, suggesting that BSA was glycated. In contrast, such immunoreactivity was absent on BSA without glucose 6-phosphate ( Figure 3a). These results indicate that CML is a major component of our AGE preparation, used in all our experiments.
To determine direct effect of AGEs on synaptic function, we examined synaptic transmission under basal condition and during LTP, a form of synaptic plasticity that is widely studied as a cellular model for synaptic function, in hippocampal slices from non-Tg mice treated with AGEs or BSA as a control. In vehicle-treated slices, BSA alone did not alter LTP (228.8±17.04% versus 223.0±7.95% in BSA-treated slices, P40.05, Figures 3b and c) or basal synaptic transmission (BST) as measured by input-output measurement ( Figure 3d). AGE treatment significantly reduced LTP in a concentration-dependent manner (180.9 ± 15.36%, 139.1 ± 10.42%, and 124.9 ± 7.96% for 50, 100, and 200 mg/ml AGEs, respectively, versus 220.9±20.75% for 100 mg/ml BSA, Po0.01, Figures 3e and f). There were no significant differences of fieldexcitatory post-synaptic potential (fEPSPs) in CA1 stratum radiatum between AGE-and BSA-treated slices, indicating that AGEs did not affect normal BST in hippocampal synapses ( Figure 3g). These results demonstrate that AGEs impair long-term synaptic plasticity.

RAGE deficiency rescues AGE-induced LTP deficits.
To directly determine the impact of RAGE on AGE-mediated reduction of LTP, we first recorded LTP in hippocampal slices from global RAGE depletion (RKO) and non-Tg mice treated with AGEs. We found similar potentiation in RAGE-deficient   Figure 4d shows representative traces taken 1 min before (black line) and 60 min after LTP induction (gray line). BST was not affected in RAGE knockout or mutant mice ( Figure 4f). These results indicated that RAGEdependent signaling in neurons contributes to AGE-induced synaptic malfunction.
Mitogen-activated protein kinase 38 (MAPK p38) activation is involved in AGE-induced synaptic deficit. To determine which molecular players underlie AGE effects on hippocampal LTP, we evaluated the role of p38 and c-Jun   37 Using antibodies to phosphorylated forms of p38 and JNK with hippocampal extracts after exposure to AGEs and high frequency stimulation, we estimated activation of these two kinases. Addition of AGEs to non-Tg hippocampal slices significantly increased p38 phosphorylation compared with BSA addition, but treatment with AGEs did not affect total p38 levels ( Figure 5a). Densitometry analysis of the combined immunoreactive bands revealed a significant increase in p38 phosphorylation by 1.5-1.7-fold when normalized to total p38 versus that of BSA-treated controls (Figure 5b). AGEs treatment did not affect JNK phosphorylation ( Figure 5c). To further evaluate the effect of p38 activation on AGE-mediated LTP impairment, we perfused hippocampal slides with SB203580, a specific p38 inhibitor, along with AGEs. Clearly, SB203580 application significantly blocked induction of p38 phosphorylation in the presence of AGEs compared with the AGE treatment alone without p38 inhibitor (SB203580) (Figure 5d). Consequently, AGE-impaired hippocampal LTP was restored by inhibition of p38 with SB203580 perfusion (Figures 5e and f). These results suggest that AGE-mediated p38 activation contributes to synaptic injury.

RAGE deficiency blocks AGE-induced p38 activation.
To determine whether RAGE is a key player in AGEmediated p38 activation, we assessed the effect of RAGE deletion on p38 phosphorylation induced by AGEs. Slices from RKO mice showed completely suppressed p38 activation as shown by decreased p38 phosphorylation compared with non-Tg slices with AGEs (Figures 6a and b). Further, RAGE depletion specifically targeted to cortical/hippocampal neurons or neuron-targeted expression of a signal transduction-deficient mutant of RAGE abolished AGE-induced p38 phosphorylation. Quantification of all immunoreactive bands for phospho-p38 normalized to total p38 verified significantly reduced phospho-p38 immunoreactive bands in slices from RKO, nRKO, and DN-RAGE compared with non-Tg slices with AGE treatment (Figures 6a and b). These data indicate that RAGE deficiency prevents AGE-induced p38 activation. p38 activation of signal transduction is a consequence of AGEs-RAGE interaction on neurons, contributing to neuronal and synaptic perturbation in an AGE-enriched environment.  Effect of RAGE deficiency on AGE-induced p38 activation. Hippocampal slices from the indicated Tg mice and non-Tg mice were perfused with AGEs for 1 h and then subjected to immunoblotting for phospho-and total-p38. (a) Immunoblots for phospho-and total-p38 with tubulin used as protein-loading control. (b) Densitometry of phospho-p38 immunoreactive bands normalized by total p38 using the NIH Image J software. N ¼ 3-5 mice per group

Effect of AGEs-RAGE interaction on synaptic density.
To determine the extent of AGE-induced synaptic damage in AGE-insulted neurons, we first analyzed the effect of AGEs on synaptic density. Synapses were identified as synaptophysin-positive clusters attaching to dendrites labeled with MAP2 marker. Recruitment of the presynaptic protein, synaptophysin, to form synaptophysin-positive clusters is rapid and synaptic activity-dependent. 38 Two hours after AGEs treatment, synaptophysin density in hippocampal neurons was not altered compared with BSA-treated control cells, whereas hippocampal neurons exposed to AGEs for 48 h showed synaptophysin-positive cluster reduced by 50-60% (Figure 7a). Importantly, RAGE-deficient neurons largely reversed AGE-induced decrease in synaptic density (Figure 7b). These data indicate that RAGE depletion rescues AGE-induced synaptic loss.

Discussion
In addition to its ability to directly alter the structure and function of targeted proteins within cells that causes cell or tissue damage, emerging evidence has also demonstrated AGEs as a signaling ligand, interacting with RAGE; AGEs elicit signal transduction changes that adversely affect numerous peripheral organs. [39][40][41] Although AGE accumulation is increased in cortical neurons, hippocampal pyramidal neurons, astrocytes, and other glial cells in aging and AD brain, 4,42,43 the direct effect of AGEs-RAGE interaction on brain function, in particular on changes in synaptic structure and function, remains largely unknown. Using our novel transgenic mouse model with neuronal expression of RAGE signaling and lacking neuronal RAGE in the forebrain for evaluation of synaptic transmission and plasticity (almost every brain function relays on synaptic transmission), we provide convincing evidence to support a pivotal role of neuronal AGEs-RAGE interaction on MAPK P38 activation, hippocampal plasticity deficit, and synaptic injury.
We first demonstrated detrimental effect of AGEs on synaptic structure and function. The addition of AGEs significantly impaired synaptic plasticity by reducing LTP and synaptic morphology. LTP is important to long-term synaptic plasticity, and altered synaptic morphology (synaptic loss), which likely contributes to the learning and memory deficits seen in age-related cognitive decline. Aging is characterized by anatomical and functional decline across multiple organ systems. AGEs are found in the serum and accumulate in the tissue during aging; 5,44,45 this accumulation is accelerated in pathological conditions such as diabetes and neurodegenerative diseases, including AD. Increased levels of AGEs are reported in the cortical neurons of older adults and are positively correlated with the severity of cognition impairment. 46 The serum AGE levels were significantly increased in diabetic patients (equivalent to 30-88 mg/ml AGE-BSA versus 12 mg/ml AGE-BSA in non-diabetic patients). [6][7][8] Our previous studies demonstrated that AGE albumin (1 mM, equivalent to 66.5 mg/ml) resulted in oxidant stress and activation of transcription factor NF-kB in vitro and in vivo, a process involving the AGE-binding protein RAGE. Notably, AGE-induced cellular perturbation and oxidant stress were blocked by antibodies to RAGE. 47 Thus, increased levels of AGEs and of their receptor, RAGE, play an important role in AGE-mediated cellular perturbation during the pathogenesis of age-related processes and diabetes. The concentration of AGEs used in the present study correlates with that of the pathological condition found in human disease.
In our study, perfusion with AGEs in hippocampal slices impaired LTP in a concentration-dependent manner, consistent with findings observed in the rat model that exogenous AGE treatment by stereotaxic injection into the brain led to synapse and memory impairments. 48 However, the impact of neuronal RAGE on AGE-induced synaptic damage is unclear. Given that AGE formation and accumulation is accelerated in the brain of diabetic and AD subjects, AGEs-RAGE interaction could be an important target in the search for agents that halt disease acceleration or exacerbation relevant to diabetes-and AD-related cognitive decline and dementia. Next, using our genetic RAGE depletion mouse model, we investigated the effect of RAGE-dependent signaling on AGE-insulted synaptic injury. AGE perfusion-induced LTP reduction may be caused by varying factors; for example, one cause involves AGE binding to RAGE receptors, and another is via protein crosslinking-induced cellular malfunction. Genetic deletion of RAGE offers us the ability to identify AGEs-RAGE interaction in AGE-induced synaptic impairment. We recorded normal LTP readings in brain slices of our global RAGE null mice treated with vehicle compared with non-transgenic littermates, indicating that genetic RAGE deletion does not significantly alter synaptic structures important for LTP induction and expression under normal physiological conditions. Notably, RAGE deficiency protects against AGE-induced LTP deficits. Similarly, our data demonstrate the protective effect of RAGE depletion on AGE-induced synaptic loss. Although our results did not exclude other effects of AGEs such as protein crosslinking, we clearly demonstrated that AGEs-RAGE interaction is involved in AGE-induced synaptic injury.
As RAGE is expressed in neurons and non-neuronal cells (i.e., glial, endothelial cells, and pericytes of the blood-brain barrier), 49,50 the results obtained from global RAGE knockout mice do not delineate which cell type(s) is (are) important in AGE-induced synaptic plasticity impairment. Therefore, we created transgenic mice with nRKO and a signal transductiondeficient mutant of RAGE (DN-RAGE) specifically in the forebrain, including the hippocampus, to address the question of whether the activity of neuronal RAGE is responsible for AGE-induced hippocampal LTP reduction. Indeed, our findings clearly showed that nRKO and DN-RAGE brain slices were largely resistant to disruption of LTP via AGE exposure, implicating the involvement of neuronal RAGE in mechanisms underlying AGE-induced synaptic dysfunction. Further, introduction of neuronal DN-RAGE or RAGE deficiency in cortical neurons (nRKO) failed to completely prevent AGE-impaired LTP compared with perfusion with BSA. These results suggest that non-neuronal RAGE (i.e., RAGE expression in microglia, astrocyte, or endothelial cells) may be involved in AGE-induced synaptic dysfunction. Future studies will be required to investigate the role of non-neuronal RAGE in synaptic failure induced by AGEs. Regardless, our studies delineate the mechanisms involving neuronal RAGE-mediated signal transduction in AGEinduced synaptic dysfunction and degeneration. Neurons are susceptible to AGE insults via direct AGEs-RAGE interaction contributing to synaptic dysfunction and degeneration.
Finally, our results suggest that the MAPK P38 signaling pathway is critical for AGEs-RAGE-induced synaptic damage. AGE-induced LTP decline was attenuated in DN-RAGE slices with a signal transduction-deficient mutant of RAGE, indicating the contribution of RAGE-dependent signaling to AGE-induced synaptic impairment. Signaling cascades activated after ligand-RAGE interaction include pathways those of p21ras, extracellular signal-regulated kinase 1/2 (ERK1/2), p38, and JNK, Rho GTPases, phosphoinositol-3 kinase, GSK-3b, and the JAK/STAT (signal transducer and activator of transcription). 51,52 Activation of different protein kinase cascades may represent a principal target of RAGE activation for control of synaptic plasticity. 37,53 In particular, we considered the potential involvement of two different kinases, JNK and p38 in LTP inhibition based on our previous study showing that p38 and JNK activation are required for LTP inhibition induced by Ab-RAGE interaction. 37,54 Hippocampal slices perfused with AGEs showed increased p38 activity but no change in JNK, suggesting the involvement of p38 activation in p38 signaling and the AGE-induced synaptic plasticity deficits. Indeed, the addition of p38 inhibitor prevented AGE-induced p38 activation and LTP impairment. Note, RAGE depletion blocked induction of p38 phosphorylation and restored defects in synaptic plasticity and density from AGE exposure. Our results indicate that neuronal AGEs-RAGE interaction activates p38 MAK kinase, which in turn leads to synaptic dysfunction.
We further observed that if AGEs level remained chronically high, synapses become structurally damaged. Using hippocampal neuronal cultures, we showed that short-term AGEs exposure (for 2 h treatment) barely affected synaptic structure, whereas long-term treatment of AGEs (for 48 h) caused significant changes in synaptic density. Importantly, RAGE deletion rescues AGE-induced synapse loss. These data indicate that neuronal AGEs-RAGE interaction has functional (long-term plasticity) as well as structural effects on synapses, which may be important in the learning and memory deficit present in age-related dementia and age-associated neurodegenerative disease.
In summary, we provide substantial evidence of the protective effect of RAGE depletion on AGE-induced alterations in synaptic plasticity and density. Genetic depletion of RAGE and RAGE signaling in cortical neurons blocks AGEmediated activation of p38 MAP kinase signal transduction with LTP deficits. We propose that AGEs-RAGE-dependent activation of p38 MAP kinase is responsible for AGE-induced synaptic damage. Blockade of AGEs-RAGE axis may protect against the impaired cognitive function in aged subjects with and without neurodegenerative diseases, including those with diabetes and AD.

Materials and Methods
Transgenic mice Generation and characterization of conditional neuronal knockout of RAGE mice Targeted disruption of mouse RAGE gene. We used the Cre-loxP system to generate the targeting vector and the RAGE/flox mice (performed by Ozgen Inc.). The targeting vector consists of three fragments, namely, the 5 0 homology arm, 3 0 homology arm, and loxP arm. Two loxP sites flank RAGE exons 2-4 to allow for Cre-mediated deletion using Cre recombinase. Excision of exons 2-4 by Cre recombinase led to a frame shift and early stop codon from the mouse RAGE sequence. A PGK-neo selection cassette was inserted downstream of exon 1. The PGK-neomycin resistance selection cassette is flanked by FRT sites and was deleted using FLPe recombinase (Figure 1, targeting vector).
Generation of neuronal deletion of RAGE mice (nRKO). After deletion of the PKGneo cassette by crossing with FlpE-deletor C57BL/6 mice (Ozgen Inc.) to eliminate the neomycin resistance selection cassette (PKG-neo), the floxed RAGE allele was generated. To generate Cre recombinase RAGE null mice (RAGE flox/flox /CK2-Cre, namely, nRKO mice), selective for the cortical and hippocampal neuron under the control of the Camk2a (CK2) promoter, RAGE/flox mice were bred with neuronal-targeted and region-restricted transgenic mice that express Cre recombinase in the entire forebrain (L7Ag13 line) under the control of the forebrain-specific CAMK II promoter. 34 Cre mice (L7Ag13 line) were kindly provided by Dr. Eric Kandel (Columbia University, New York, NY, USA). Floxed RAGE mice were identified by PCR with primers (forward: 5 0 -TACCGAGTCC GAGTCTACCGTAAG-3 0 and reverse: 5 0 -TCTCCTGGTCTCTTCCTTCACGAG-3 0 ) to amplify a 551-bp fragment from the wild-type (WT) allele and a 700-bp fragment from the floxDneo allele. The Cre transgene was identified by PCR primers (forward: 5 0 -CTTCATCGTCGGTCCG-3 0 and reverse: 5 0 -ACATTCTCCCACCGTC-3 0 ) to amplify a 300-bp fragment. Specific neuronal RAGE knockout was further confirmed by western blotting of cortical and cerebellum homogenates using rabbit anti-RAGE antibody (generated in our Lab and used in our previous study 55 ). All transgenic mice were of the C57BL/6 strain. The Institutional Animal Care and Use Committee of the University of Kansas approved all protocols. Male transgenic mice and their littermate controls were used for the in vitro electrophysiology experiments.
Homozygous RAGE-null mice (RKO) and transgenic mice with cytosolic dominant-negative RAGE (DN-RAGE) We generated and characterized homozygous RAGE-null mice (RKO) as described previously. 33,56 In addition, we used transgenic mice with signal transduction-deficient mutants of RAGE in which the cytosolic domain of the receptor was deleted, thereby imparting a dominant-negative (DN)-RAGE effect, targeted to neurons (DN-RAGE) driven by the platelet-derived growth factor-B chain promoter. DN-RAGE mice were previously characterized, demonstrating localization of DN-RAGE in cortical neurons. 54

Preparation of AGEs
We generated AGEs by incubating BSA (25 mg/ml) with 250 mM glucose 6-phosphate at 37 1C for 6-8 weeks in 0.1 M phosphate buffer containing 1.5 mM PMSF, 0.5 mM EDTA, and 1 mM azide. 57 The control preparation (BSA) was treated identically with the exception that glucose 6-phosphate was omitted. We dialyzed preparations with phosphate buffer to remove free glucose, EDTA, PMSF, and azide after incubation. The degree of glycation was determined by immunoblotting using antibody to CML (TransGenic Inc., Kobe, Japan). Coomassie blue staining of gel was used as a protein-loading control.

Electrophysiological studies
Electrophysiological studies were performed as described. 54,58 Transverse hippocampal slices (400 mm) were cut from the mouse brain, maintained in an interface chamber at 29 1C, and perfused with artificial cerebrospinal fluid (aCSF) continuously bubbled with 95% O2 and 5% CO2. The aCSF composition (in mM) was: 124.0 NaCl, 4.4 KCl, 1.0 Na 2 HPO 4 , 25.0 NaHCO 3 , 2.0 CaCl 2 , 2.0 MgCl 2 , and 10.0 glucose. fEPSP was recorded for the CA1 region of the hippocampus by placing the stimulating electrode at the level of the Schaeffer collateral fibers and recording electrode in the CA1 stratum radiatum. BST was assayed by plotting the stimulus voltage (V) against slopes of fEPSP to generate input-output relations. For LTP experiments, a test pulse was applied every minute at an intensity evoking B35% of the maximum evoked response. LTP was induced using y-burst stimulation (4 pulses at 100 Hz, with bursts repeated at 5 Hz and each tetanus, including three 10-burst trains separated by 15 s). Responses were recorded for 60 min after tetanization and measured as fEPSP slope expressed as the percentage of baseline.
After blocking in TBST buffer (20 mM Tris-HCl, 150 mM sodium chloride, 0.1% Tween-20) containing 5% non-fat dry milk (Santa Cruz, Dallas, TX, USA) for 1 h at room temperature, we incubated the membrane, and gently shaking overnight (at 4 1C) with primary antibodies, followed by incubation with the corresponding secondary antibody for 1 h at room temperature. We used antibodies to phosphorylation and total of p38 and JNK (Cell Signaling, Boston, MA, USA) and tubulin (Sigma, St. Louis, MO, USA). ImageJ software (National Institutes of Health, Bethesda, MD, USA) was used for analysis of scanned blots and to quantify intensity of the immunoreactive bands.

Immunohistochemical analysis
Mice were anesthetized by an intraperitoneal injection of a mixture of ketamine (50 mg/kg) and xylazine (10 mg/kg) and perfused transcardially with 0.9% sodium chloride for 10 min. Mouse brains were fixed in 4% paraformaldehyde at 4 1C for 2 days. Floating coronal sections of 30-mm thicknesses were cut by vibratome (Leica VT 1000S, Wetzlar, Germany), collected, and immersed in wash buffer (0.1 M sodium phosphate, 0.5 M sodium chloride, Triton X-100, pH 7.4) for 30 min. After preincubation for 1 h in blocking solution (10% normal goat serum, 0.3% Triton X-100 in PBS), we incubated sections overnight at 4 1C with primary antibodies, using rabbit anti-RAGE IgG (3.5 mg/ml, generated in our Lab) and mouse anti-MAP2 (1 : 10 000, Chemicon, Billerica, MA, USA) for cortex and hippocampus staining. Sections were then incubated with Alexa Fluor 594-conjugated goat antirabbit IgG and 488 goat anti-mouse IgG secondary antibodies (1 : 1000, Invitrogen) for 1 h at room temperature. Nuclei were stained by DRAQ5 (5 mM, Cell Signaling) for 5 min at room temperature.

Measurement of synaptic density
Neuronal culture. We prepared hippocampal neurons from day 1 non-Tg or RKO mice in the C57/BL6 background as described previously, 59 culturing neurons in neurobasal medium supplemented with 1 X B27, 600 mM L-Glutamine and penicillin-streptomycin. At day 10 in vitro, neurons from both Tg mice were treated with 100 mg/ml of AGEs; BSA was used as a control in neurobasal medium supplemented with 0.5 X B27 for 2 and 48 h, respectively.
Immunostaining of cultured neurons. After the various treatments, neurons were fixed with 4% ice-cold paraformaldehyde for 5 min and then permeabilized with PBS containing 0.1% Triton and 5% goat serum for 1 h at room temperature followed by incubation with primary antibodies (rabbit anti-synaptophysin IgG (1 : 5000, Dako) and mouse anti-MAP2 IgG (1 : 10 000, Chemicon)) to visualize dendrites at 4 1C for 16 h. Neurons were incubated with both Alexa Fluor 488-conjugated goat anti-rabbit IgG and 594 goat anti-mouse IgG (1 : 1000, Invitrogen) for 1 h at room temperature. After washing with PBS, neurons were covered with Vectashield mounting medium (H-1000, Vector Laboratories, Burlingame, CA, USA). Images were taken at equal exposure for all the different groups at Â 63 oil lens under a confocal microscopy (Leica) and analyzed by the Universal Metamorp Image Program (Molecular Devices, Sunnyvale, CA, USA).
Synaptic density of cultured neurons was measured by counting the number of synaptophysin-positive clusters in secondary branches and puncta per micron of dendrite (presented as the number of synaptophysin clusters per micron of dendrite) and calculated by dividing the length of the dendrite. The experiments were performed by investigators blinded to the information about genotype and treatment. Codes for the ID of group cells were not disclosed until completion of image analysis.

Statistical analysis
One-way ANOVA was used for repeated measure analyses followed by Fisher's protected least significant difference for post hoc comparisons. We used StatView 5.0.1 Windows software (SAS Institute, Cary, NC, USA), and results are reported as mean ± S.E.M. Po0.05 was considered significant.