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. Author manuscript; available in PMC: 2017 Jan 12.
Published in final edited form as: Cell Metab. 2015 Nov 19;23(1):103–112. doi: 10.1016/j.cmet.2015.10.009

Striatal Dopamine Links Gastrointestinal Rerouting to Altered Sweet Appetite

Wenfei Han 1,2,3, Luis A Tellez 2,3, Jingjing Niu 2,3,6, Sara Medina 2, Tatiana L Ferreira 2,3,4, Xiaobing Zhang 5, Jiansheng Su 1, Jenny Tong 6, Gary J Schwartz 7, Anthony van den Pol 5, Ivan E de Araujo 2,3,8,*
PMCID: PMC4715689  NIHMSID: NIHMS739717  PMID: 26698915

Abstract

Reductions in calorie intake contribute significantly to the positive outcome of bariatric surgeries. However, the physiological mechanisms linking the rerouting of the gastrointestinal tract to reductions in sugar cravings remain uncertain. We show that a duodenal-jejunal bypass (DJB) intervention inhibits maladaptive sweet appetite by acting on dopamine-responsive striatal circuitries. DJB disrupted the ability of recurrent sugar exposure to promote sweet appetite in sated animals, thereby revealing a link between recurrent duodenal sugar influx and maladaptive sweet intake. Unlike ingestion of a low-calorie sweetener, ingestion of sugar was associated with significant dopamine effluxes in dorsal striatum, with glucose infusions into the duodenum inducing greater striatal dopamine release than equivalent jejunal infusions. Consistently, optogenetic activation of dopamine-excitable cells of dorsal striatum was sufficient to restore maladaptive sweet appetite in sated DJB mice. Our findings point to a causal link between striatal dopamine signaling and the outcomes of bariatric interventions.

Introduction

Substantial reductions in overall calorie intake greatly contribute to the positive outcome of bariatric surgeries (Laurenius et al., 2013; Mathes et al., 2015; Munzberg et al., 2015). Unfortunately, the neurophysiological mechanisms mediating these newly acquired food choices remain largely unknown (Arble et al., 2015; Miras et al., 2012; Munzberg et al., 2015; Seeley et al., 2015). While it remains to be determined if bariatric interventions are sufficient to induce substantial weight loss in high-sugar fed animals, current evidence indicates that the prevalent post-surgical decreases in sweet appetite are not due to faulty taste perception. Instead, they appear to result from alterations in post-ingestive nutrient sensing caused by the rerouting of the gastrointestinal tract (Mathes et al., 2015).

Such alterations in post-ingestive signaling are likely due to diversions in duodenal-jejunal nutrient influx. In fact, gastrointestinal rerouting, but not gastric banding, distinctly restrains excessive sugar intake (Ernst et al., 2009). Rodent models demonstrate in turn that exaggerated caloric intake is mediated by adaptations in dopamine receptor signaling within dorsal striatum (Furlong et al., 2014; Johnson and Kenny, 2010; Smith and Robbins, 2013). It is thus noteworthy that nutrient sensing in the gastrointestinal tract stimulates robust dopamine efflux in dorsal striatum (de Araujo et al., 2012; Tellez et al., 2013a; Tellez et al., 2013b). Building on such gastrointestinal-striatal dopamine link, we set out to confirm that intestinal rerouting curbs maladaptive sweet appetite, and hypothesized that i. Sugar-induced surges in striatal dopamine (Ren et al., 2010; Tellez et al., 2013c) depend on glucose sensing within specific segments of the intestine typically bypassed in bariatric interventions; and ii. Post-surgical restraint of hunger-independent sugar appetite can be overturned by artificially activating striatal dopamine-excitable cells.

Results

Recurrent exposure to sugar, but not to low-calorie sweetener, enhances sweet appetite in the absence of hunger

We established a behavioral protocol where mice are chronically exposed to either nutritive or non-nutritive sweeteners while orosensory differences between sweeteners are minimized. The overall objective of the assay is to measure sensitivity to sugar-induced satiation in both sugar-experienced and artificial sweetener-experienced mice. To induce satiation, an intra-gastric load of glucose was administered previous to a sweet-seeking task. Our goal was to determine the animals’ persistence in seeking the sweet reward after the sugar load was administered into the gut (see scheme in Figure 1A). The volumes of the intra-gastric preloads were equivalent to the animals’ baseline intake during 13 habituation days of exposure. By measuring sweet seeking after preloads, we thus quantified how much of the sweet rewards animals would ingest beyond their typical daily ad libitum intake.

Figure 1. Chronic exposure to sugar adjusts the sensitivity to satiating intra-gastric preloads, an effect mediated by the duodenal-jejunal route.

Figure 1

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A. Behavioral preparation. Mice are placed in a behavioral box where licks for the artificial sweetener sucralose trigger an infusion pump that delivers either glucose (blue) or more sucralose (red) directly into the gut. After 13 days of daily 1h exposure sessions, mice are tested for intake levels after administration of satiating intra-gastric preloads of either glucose or sucralose that correspond to average daily intake levels. B. The behavioral protocol was initially employed in mice with preserved gastrointestinal routing. C. Glucose-exposed mice (N=12) self-administer significantly more sucralose than sucralose-exposed mice (N=12) after an intra-gastric preload of glucose (t[22]=4.0, *p=0.001). Y-axis indicates final volume self-administered into the gut, such that the bottom half of the Y-axis (yellow area) shows the volume of the preload, and the upper half the amounts consumed after the preload. Note the preload volumes correspond to their baseline intake of each solution. D. Glucose-exposed mice (N=8) self-administer significantly more glucose than sucralose-exposed mice (N=8) after an intra-gastric preload of glucose (t[14]=2.6, *p<0.02). E. Glucose-exposed mice (N=6) self-administer significantly more glucose than sucralose-exposed mice (N=6) after an intra-gastric preload of sucralose (t[10]=6.3, *p=0.001). F. Glucose- and sucralose-exposed mice (N=6 each group) self-administer similar levels of sucralose after an intra-gastric preload of sucralose (t[10]=0.58, p=0.57). G. The behavioral protocol was also employed in mice sustaining a duodenal-jejunal bypass (DJB) intervention. Scheme representation of the DJB, which involves pylorus ligation (black line drawn under stomach) and a gastric-jejunal anastomose. Blue arrows indicate rerouting. H. DJB glucose-exposed mice (N=5) self-administer significantly less sucralose than glucose-exposed sham controls (N=5) after an intra-gastric preload of glucose (with respect to preload: SHAM=202.4% ± 47.4%; DJB=11.1% ± 8.5%, Paired T-test t[4]=4.2, *p=0.013). I. DJB glucose-exposed mice self-administer significantly less glucose than glucose-exposed sham controls after an intra-gastric preload of glucose (SHAM=93.7% ± 8.6%; DJB=50.3% ± 12.2%, t[4]=8.4, *p=0.001). J. DJB glucose-exposed mice showed a tendency to self-administer significantly less glucose than glucose-exposed sham controls after an intra-gastric preload of sucralose (SHAM=151.2% ± 15.8%; DJB=90.19% ± 8.7%, t[4]=6.8, *p=0.002). K. DJB glucose-exposed mice self-administered similar amounts of sucralose to glucose-exposed sham controls after an intra-gastric preload of sucralose (SHAM=237.3% ± 26.5%; DJB=204.4% ± 33.7%, t[4]=1.0, p=0.37). I = Intake after preload. P = Preload. n.s.=Statistically-non-significant. Data are represented as mean ± SEM.

During the thirteen habituation days, mice licked a spout containing a non-caloric sweetener (sucralose). In one group of mice, sucralose licking triggered intra-gastric infusions of D-glucose solutions (“glucose-exposed mice”). In a second group of mice sucralose licking triggered intra-gastric infusions of the same sucralose solution (“sucralose-exposed mice”, Figure 1A). We chose D-glucose to model sugar ingestion due to its unvarying presence in sugared products. The recorded baseline levels of intra-gastric glucose (Figure S1A–B) determined the glucose preload volume for both groups of mice. The same volumes were then used to determine the sucralose preloads for both groups of mice (so that all preloads differed in sugar content but resulted in comparable intra-gastric volume).

The rationale for this assay was to determine how much of the sweet rewards animals would ingest beyond their typical daily ad libitum intake (i.e. the daily baseline level)

After the 13 days of habituation, on day 14 we assessed sweet-seeking behavior in the absence of hunger by measuring sucralose self-administration immediately after the intra-gastric glucose preload. Strikingly, in animals in which gastrointestinal routing is preserved (Figure 1B), glucose-exposed mice self-administered significantly more sweetener than sucralose-exposed mice after preload administration (Figure 1C). This between-group effect also holds for other combinations of satiating preloads, including glucose self-administration after glucose preloads (Figure 1D), and glucose self-administration after sucralose preloads (Figure 1E). Interestingly, no between-group differences were observed in sucralose self-administration after sucralose preloads (Figure 1F), indicating that the non-homeostatic effects produced by gastrointestinal glucose exposure are observed only when sugar is present in the gastrointestinal tract. We also confirmed that glucose, but not sucralose, preloads significantly increased circulating glucose and insulin plasma levels (Figure S1C–D). In sum, daily exposure to a sugar, but not to an artificial sweetener, diminishes the satiating effects induced by intra-gastric caloric preloads.

A bariatric intervention that prevents sugar influx into upper duodenum suppresses sweet appetite in the absence of hunger

We then inquired whether this non-homeostatic pattern of sweet appetite could be suppressed in mice sustaining a duodenal-jejunal bypass (henceforth “DJB”) intervention. To divert food from its usual duodenal route, the pylorus was ligated while the mid-jejunum was side-by-side anastomosed to the stomach (thereby bypassing 6–8cm of duodenal tissue, Figure 1G and Figure S1E–H).

We applied the intra-gastric preload paradigm to both DJB and sham-operated mice. Importantly, both DJB and sham control mice were chronically exposed to sugar via intra-gastric glucose. As above, the preload volume was determined from the animals’ average volume of self-infused glucose. We calculated the relative intake during the sweet-seeking task with respect to each animal’s baseline level. After infusion of the sugar preload, sham-operated mice self-administered significantly more sweetener than DJB mice (compared to baseline levels, SHAM=~202% vs. DJB=~11%, Figure 1H). Similar effects were obtained for other types of preloads (Figure 1H–J), except when sucralose intake followed sucralose preloads (Figure 1K). This corroborates the view that sugar-promoted sweet appetite depends on duodenal glucose sensing. In sum, the sweet appetite-promoting actions of sugar exposure are attenuated in DJB mice.

Acute administration of glucose into duodenum does not alter sweet seeking We next asked if glucose duodenal sensing may produce acute stimulatory effects on sweet seeking in sugar-naïve mice (i.e. independently of previous exposures to glucose). Mice were implanted with duodenal catheters and infused glucose or sucralose duodenally immediately previous to the sweet seeking task described above. We have however found no significant differences between the sugar and sweetener conditions with this protocol (Figure S1I). It does therefore appear that repeated exposure to duodenal glucose is required for sugar intake to promote stimulatory effects on sweet seeking.

Intra-gastric glucose infusions induce significantly greater dorsal striatal dopamine release in sham compared to DJB mice

To investigate the neurophysiological bases of the behavioral patterns described above, we performed intra-gastric infusions of glucose or sucralose concomitantly to brain dopamine measurements (Figure 2A). Measurements were obtained from both dorsal and ventral striatal sectors (Figure 2B).

Figure 2. The Duodenal-jejunal route regulates gut-induced striatal dopamine.

Figure 2

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A. Assay for assessing gut-induced striatal dopamine in awake mice. Injections of sucralose or glucose into stomach are performed concomitantly to brain microdialysis in dorsal or ventral striatum. B. Representation of dialysis probe locations into dorsal (blue) and ventral (red) striatal sectors. C. Intra-gastric infusions of glucose induced significantly greater dopamine release in dorsal striatum of sham mice (N=5) compared to sucralose infusions (two-way repeated measures ANOVA, sampling time × intra-gastric stimulus interaction F[14,56]=4.39, *p<0.001. Y-axis represents percent change with respect to baseline levels. Mean absolute dopamine concentrations for baseline samples were 0.24±0.01pg/uL and 0.24±0.01pg/uL for glucose and sucralose sessions, respectively. D. This effect was completely abolished in DJB mice (N=5, F[14,70]=0.7, p=0.7). Mean absolute dopamine concentrations for baseline samples were 0.22±0.01pg/uL and 0.20± 0.01pg/uL for glucose and sucralose sessions, respectively. E. Intra-gastric infusions of glucose induced significantly greater dopamine release in ventral striatum of sham mice (N=5) compared to sucralose infusions (F[14,56]=3.9, *p<0.001). Mean absolute dopamine concentrations for baseline samples were 0.09±0.01pg/uL and 0.09±0.01pg/uL for glucose and sucralose sessions, respectively. F. Similar effects were observed in DJB mice (N=5, F[14,56]=7.3, *p<0.001). Mean absolute dopamine concentrations for baseline samples were 0.10±0.01pg/uL and 0.09± 0.004pg/uL for glucose and sucralose sessions, respectively. G. Injections of sucralose or glucose into different parts of intestine of awake mice are performed concomitantly to brain microdialysis in dorsal striatum of sham mice. H. Small 100mg/kg glucose infusions into duodenum induced significantly greater dopamine release than equivalent jejunal infusions (F[14,56]=4.1, *p<0.001). Mean absolute dopamine concentrations for baseline samples were 0.21±0.01pg/uL and 0.23±0.02pg/uL for duodenal and jejunal sessions, respectively. I. Similar experiments were performed in DJB mice. J. The effects were not preserved when infusions were made into the bypassed duodenum and compared to jejunum (between-subjects two-way ANOVA F[14,112]=1.390, p=0.17). Mean absolute dopamine concentrations for baseline samples were 0.19±0.01pg/uL and 0.22±0.03pg/uL for duodenal and jejunal sessions, respectively. K. Mice were implanted with portal (N=5) or jugular (N=5) intravenous catheters, through which the same dose of glucose as in intestine was infused. L. Portal infusions of glucose produced significant increases over baseline, and were significantly greater than equivalent infusions into the jugular vein (two-way mixed-model ANOVA, sampling time × vein interaction F[14,112]=9.2, *p<0.001. Mean absolute dopamine concentrations for baseline samples were 0.23±0.02pg/uL and 0.25±0.01pg/uL for portal and jugular sessions, respectively. Saline infusions did not produce vein-specific effects (F[14,112]=0.4 p=0.9). n.s. = statistically non-significant. Data are represented as mean ± SEM.

In order to mimic the intra-gastric administration patterns observed during the behavioral sessions, we used the infusion pumps timestamps recorded during behavioral sessions involving DJB mice to “replay” the intra-gastric infusions. We found that, in sham mice, glucose infusions induced significantly greater dopamine release in dorsal striatum compared to sucralose infusions (Figure 2C). In contrast, we found that neither glucose or sucralose infusions stimulated release in dorsal striatum of DJB mice (Figure 2D). Remarkably, intra-gastric infusions of glucose elicited greater dopamine release than sucralose infusions in the ventral striatum of both sham (Figure 2E) and DJB (Figure 2F) mice. These results indicate that dorsal, not ventral, striatum is the critical striatal site impacted by DJB interventions.

Duodenal glucose infusions induce significantly greater striatal dopamine release than equivalent jejunal infusions

To gain further insights into gut-stimulated dopamine effluxes in dorsal striatum, dopamine release was measured in the brain during small infusions of glucose or sucralose into different intestinal segments (Figure 2G). As expected, infusions of the artificial sweetener sucralose did not influence dopamine release when directly infused into either the duodenum or jejunum of awake mice (including in DJB mice, Figure S2A–B). In contrast, duodenal infusions of glucose induced significantly greater dopamine release than jejunal infusions in dorsal striatum of sham mice (Figure 2H). In contrast, this effect was greatly attenuated in DJB mice (Figures 2I–J). The duodenum thus emerges as the critical intestinal site mediating sugar-induced striatal dopamine release in dorsal striatum. Lower glucose concentrations failed to promote dopamine efflux upon duodenal or jejunal infusion (Figure S2C). Infusion parameters are shown in Table S1.

Because glucose absorption rates are the highest in the upper intestine, the above raises the possibility that a post-intestinal locus detects absorbed glucose to trigger dopamine efflux. We performed infusions of glucose into the portal-mesenteric system (Figure 2K), and observed a sustained increase in striatal dopamine levels that were not matched by equivalent jugular infusions (Figure 2L; see Table S2 for infusion parameters, and Figure S2D–E for dialysis probe placements). The outcome of the jugular infusions did not shift dopamine levels from baseline, thereby ruling out systemic effects potentially produced by portal infusions. In sum, glucose absorption through duodenum into the portal-hepatic system is sufficient for sugar-induced dopamine release.

Artificial activation of dorsal striatal dopamine-excitable neurons substitutes for sugar exposure and promotes sweet appetite in the absence of hunger

We then investigated the circuit basis of the above phenomena. Dopamine increases the excitability of striatal D1r-expressing neurons (Ericsson et al., 2013; Planert et al., 2013). Using optogenetc methods (Kravitz et al., 2012; Lobo et al., 2010), we first tested the hypothesis that artificial activation of D1r-neurones in dorsal striatum would substitute for sugar in its ability to promote sweet appetite in the absence of hunger. During optogenetic experiments, mice licked a spout containing sucralose such that detected licks triggered, concomitantly to intra-gastric infusions, light pulses to either dorsal or ventral striatum (Figure 3A). Cell-specific expression of the blue light-sensitive Channelrhodopsin (ChR2) in D1r-neurones was achieved by stereotaxically injecting the Cre-dependent viral construct AAV-EF1a-DIO-hChR2(H134R)-EYFP into the striatum of D1r-Cre mice (Figure 3B). Slice electrophysiological recordings confirmed robust excitation of D1r-neurones by 473nm laser pulses (Figure 3C). Cre-dependent expression of ChR2 was confirmed by hodological criteria (Figure S3A–E). Efficacy of laser stimulation in activating striatal neurons was confirmed in vivo (Figure S3F–H)

Figure 3. Optogenetic stimulation of D1r-neurones in dorsal striatum diminishes sensitivity to satiating intra-gastric preloads and counters the appetite-suppressing effects of duodenal-jejunal rerouting.

Figure 3

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A. Detected licks trigger 3s intra-gastric infusions concomitantly to 3s blue light laser pulse to either dorsal or ventral striatum. Animals were tested after administration of glucose or sucralose intra-gastric preloads. B. EYFP visualization confirms that ChR2 transfection was contained to DS. Note the dense bundle of axon terminals in entopenducular nucleus and Substantia nigra, pars reticulata and absence of labeled fibers in globus pallidus, confirming exclusive labeling of D1r-neurons. C. Whole-cell slice recordings reveal robust optogenetic activation of a ChR2+ D1r-neurone by 1/5/10/20 Hz blue light pulses (10ms) and by a continuous light train. D. Y-axis indicates final volume self-administered into the gut, such that the bottom half of the Y-axis (yellow area) shows the volume of the preload, and the upper half the amounts consumed after the preload. Laser pulses overrode the satiating effects of intra-gastric glucose preload in a region-dependent manner (two-way mixed model ANOVA, brain region × laser state F[2,15]=5.6, p=0.01). In animals transfected with ChR2 in D1r-neurons of dorsal striatum, laser pulses induced a drastic increase in sucralose intake despite the glucose preload (Laser ON vs. OFF, N=6, paired T-test t[5]=7.0, Bonferroni *p<0.01). However, no effect was observed when ventral striatum was activated (N=6, t[5]=0.5, p=0.5). Control mice (D1r-Cre mice implanted with optical fibers and transfected in dorsal striatum with a blue-light insensitive ion channel) showed no light-dependent effects as expected (N=6, t[5]=0.7, p=0.48). Laser activation in dorsal striatum significantly increased intake compared to both control (t[10]=−4.211, **p=0.002) and ventral striatum (t[10]=4.5, ***p=0.001) mice. E. In D1r-Cre DJB mice transfected with ChR2 in dorsal striatum, laser pulses induced a marked increase in sucralose intake despite the glucose preload (Laser ON vs. OFF, N=5, paired T-test t[4]=3.4, *p=0.027). F. Similar effects were observed for glucose intake after sucralose preload (t[4]=10.4, *p<0.001). I = Intake after preload. P = Preload. n.s. = statistically non-significant. Data are represented as mean ± SEM.

We found that, as predicted, optogenetic activation of dorsal striatal D1r-neurons overrode the satiating effects produced by intra-gastric glucose preloads (Figure 3D). This effect was region specific, as the ability to enhance sweet appetite after intra-gastric glucose preloads was not reproduced upon similar activation of ventral striatum D1r-neurons (Figure 3D). Importantly, all mice used in the optogenetic tests were chronically exposed to sucralose (Figure S3I), supporting the notion that artificial activation of dorsal striatal dopamine-excitable striatal neurons substitutes for chronic sugar exposure. Similar effects were observed when sucralose intra-gastric preloads were used instead (Figure S3J). Finally, we did not observe any sizeable effects of striatal optical stimulation on blood glucose (Figure S3K) or insulin levels (Figure S3L)

Artificial activation of dorsal striatal dopamine-excitable neurons mimics the influence of intra-gastric glucose on sweetener intake

We also found that pairing sucralose licking to intra-gastric administration of glucose produced the rather notable effect of suppressing sucralose licking during the infusion period (Figure S3M; note that licks detected during intra-gastric infusions had no programmed consequences, see Experimental Procedures). Remarkably, we observed the same pattern during optogenetic assays: Likewise licks detected during intra-gastric infusions, licks detected during laser pulses had no programmed consequence. Thus, dorsal, but not ventral, striatum stimulation of D1r-neurons significantly suppressed sucralose licking during the activation of the laser source (Figure S3N). Therefore, during intra-gastric self-administration, the animals’ primary goal appeared to consist in obtaining glucose into the gut. Analogously, during optogenetic self-stimulation, the animals’ primary goal appeared to consist in obtaining laser pulses delivered to dorsal striatum. Stimulation of D1r-neurons therefore substituted for sugar in its ability to modulate the reward value of sweet taste by prioritizing the administration of a post-ingestive conditioned reward.

Artificial activation of dorsal striatal dopamine-excitable neurons overturns DJB effects

Based on the above, we hypothesized that artificial activation of D1r-neurons in dorsal striatum would suffice to counter the inhibitory effects of DJB on hunger-independent sweet appetite. We thus performed the DJB bypass intervention on D1r-Cre mice and performed the optogenetic experiments as above. These animals were however exposed to intra-gastric glucose during the habituation phase. As predicted, optical stimulation of dorsal striatum produced a striking increase in sweetener intake after infusions of intra-gastric glucose preloads in DJB mice (Figure 3E). Also, upon D1r-activation, sucralose preloads failed to inhibit further glucose intake in DJB mice (Figure 3F), rendering DJB mice distinct from sham controls with respect to their reactions to D1r-neuronal artificial stimulation. Effects for other conditions are shown in Figure S3O–Q.

Cell-specific ablation of dorsal striatal dopamine-excitable neurons mimics the effects of duodenal diversions on sweetener intake

We next tested if the activation of dopamine-excitable cells in dorsal striatum are necessary for duodenal glucose sensing to control sweet seeking behavior. Cell-specific ablation of D1r-neurones in dorsal or ventral striatum was achieved by virally introducing a Cre-dependent construct allowing for caspase-mediated ablation of D1r-expressing cells in the striatum of D1r-Cre mice. Specifically, we stereotaxically injected the Cre-dependent viral construct AAV-flex-taCasp3-TEVp (Yang et al., 2013) into the striatum of D1r-Cre mice. Specificity of the lesions was confirmed via neuroanatomical retrograde tracing methods (Figures 4A–D; Figure S4A–C shows the results of sham lesions). Control groups included D1r-Cre mice transfected with AAV-GFP constructs, and non-Cre mice transfected with AAV-flex-taCasp3-TEVp.

Figure 4. Cell-specific ablation of D1r-neurones in dorsal striatum enhances sensitivity to satiating intra-gastric preloads and mimics the appetite-suppressing effects of duodenal-jejunal rerouting.

Figure 4

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A. Strong labeling throughout dorsal striatum is observed when retrograde fluorescent beads were injected into globus pallidus, the exclusive target of D2r-expressing neurons of dorsal striatum. B. Injection site associated with globus pallidus injections. C. Weaker labeling throughout dorsal striatum is observed when retrograde fluorescent beads were injected into Substantia Nigra, pars reticulata, the exclusive target of D1r-expressing neurons of dorsal striatum. D. Injection site associated with Substantia Nigra, pars reticulata injections. E. Ablation of D1r-expressing cells in dorsal, but not in ventral, striatum resulted in glucose preloads producing strong inhibitory effects on sucralose intake (Two-way ANOVA Between-Group effect F[3,26]=30.6, p<0.001). Post-hoc, Bonferroni corrected tests: DS-casp vs.VS-casp : t[13]=9.5 *p<0.004), DS-casp vs. D1-CTL : t[12]=10.9 **p<0.004, DS-casp vs. WT-casp : t[13]=6.4, ***p<0.004). The lower half of the Y-axis (yellow area) shows the volume of the preload, and the upper half the amounts consumed after the preload. Note the preload volumes correspond to their baseline intake of each solution. F. Similar effects were observed during glucose intake after glucose preloads (F[3,26]=35.3 p<0.001). Post-hoc, Bonferroni corrected tests: DS-casp vs.VS-casp : t[13]=8.4, *p<0.004), DS-casp vs. D1-CTL : t[12]=11.6, **p<0.004, DS-casp vs. WT-casp : t[13]=8.8, ***p<0.004). G. No effects were observed during glucose intake after sucralose preloads (F[3,26]=0.01 p=0.9) or H. during sucralose intake after sucralose preloads (F[3,26]=0.4 p=0.7). Data are represented as mean ± SEM.

All animals were sugar-exposed. Strikingly, we found that ablation of D1r-expressing cells in dorsal, but not in ventral, striatum greatly potentiated the satiating effects of glucose preloads in sugar-exposed mice; specifically glucose preloads produced strong inhibitory effects on sucralose intake in dorsal striatum-lesioned mice (Figure 4E). Importantly, this effect was independent of alterations in sweet intake during habituation sessions (Figure S4D). Similar effects were observed during glucose intake after glucose preloads (Figure 4F), but not in other conditions (Figure 4G–H). In sum, likewise the bypassing of duodenal segments, ablation of dorsal striatum D1r-expresisng cells is necessary for chronic sugar to exert an stimulatory effect of sweet seeking.

Discussion

Our data provide evidence for a dopamine-centered model linking gastrointestinal rerouting to restrained sweet appetite. We showed that daily exposure to sugar maintains sweet appetite even after the administration of hunger-suppressing intra-gastric preloads; however, these effects were abolished in sugar-exposed animals sustaining a duodenal-jejunal bypass (DJB). Consistent with a role for striatal dopamine in these effects, duodenal glucose infusions induced significantly greater striatal dopamine release than jejunal infusions. Moreover, artificial activation of striatal dopamine-excitable cells overturned the appetite-suppressing effects of DJB. Our data places bariatric surgeries in the context of current evidence favoring a central role for duodenal nutrient influx in dopamine-mediated feeding behaviors (Ackroff et al., 2010; de Araujo et al., 2012).

DJB procedures are known to directly affect glucose homeostasis. Specifically, the procedure rapidly lowers glucose concentrations in uncontrolled diabetes, an effect that requires jejunal nutrient sensing (Breen et al., 2012). Likewise, glucose jejunal infusions have been shown to be sufficient to alter systemic ghrelin levels (Overduin et al., 2005). It is therefore plausible to assume that the greater sensitivity of DJB mice to intra-gastric sugar preloads may result from systemic alterations in glucose metabolism or ghrelin levels. We note however that, independently of which specific mechanism acted peripherally to drive behavioral adaptations after DJB, our central finding states that activating dopamine-excitable cells in dorsal striatum can overturn such adaptations. Future research must identify the physiological pathways linking duodenal nutrient sensing to dopamine efflux. Hypothetical models include the action of gut-brain neural axes (Tellez et al., 2013b), as well as post-aborptive metabolic sensing (Ren et al., 2010). In fact, given our finding of portal-induced dopamine release in dorsal striatum, our data are consistent with the notion that glucose, upon duodenal transport, activates glucose sensors within the portal-mesenteric system (Bohland et al., 2014; Delaere et al., 2013).

Our duodenal-jejunal approach prompts two methodological observations. First, the approach did not involve the construction of a gastric pouch in addition to intestinal rerouting (as adopted in the Roux-en-Y procedure, Seeley et al., 2015). This concurs with our experimental design, as nutrient infusions into a small pouch would greatly complicate the interpretation of behavioral tests based on intra-gastric preloads. This approach also provides some conceptual clarity to the notion that intestinal rerouting is critical for restraining appetite after bariatric interventions. Second, we only performed the bypass surgery on normal-weight mice in order to avoid potential confounds associated with weight-loss per se. In fact our surgeries did not cause significant weight loss (Table S3). We thus rule out, in principle, any confounding factors associated with substantial DJB-induced weight loss or malnutrition.

Because dopamine release in striatum is critical for feeding (Palmiter, 2008), and for reinforcement in general (Gerfen and Surmeier, 2011; Wickens et al., 2007), it is somewhat natural to speculate that changes in striatal dopamine mediate the post-surgical restraint on compulsive intake. This is consistent with the notion that that disrupting dorsal, but not ventral, striatal dopamine signaling enhances sensitivity to reward devaluation (“habit formation”, Everitt and Robbins, 2005; Yin et al., 2008). Because satiating preloads represent a prototypical case of food reward devaluation, we speculate that gastrointestinal rerouting disrupts the expression of food habits by perturbing gut-stimulated striatal dopamine release. More specifically, we found that excitation of D1r-positive cells in dorsal (but not ventral) striatum suffices to overturn the satiating effects of glucose intra-gastric preloads. This is in line with recent reports that restricted access to sweet calories leads to impaired goal-directed responding, an effect attenuated by D1r antagonism in dorsolateral striatum (Furlong et al., 2014; Johnson and Kenny, 2010; Smith and Robbins, 2013) as inducers of addiction-like feeding patterns (Smith and Robbins, 2013).

Functional neuroimaging assessments of human dopamine signaling performed after Roux-en-Y interventions indicate a complex picture. Dorsal striatal dopamine signaling is altered in the obese (Guo et al., 2014), and different studies reported partial normalization after bariatric interventions; however, according to these studies, the direction of the effect appears to depend on how receptor binding potential was measured (de Weijer et al., 2014; Dunn et al., 2010; Steele et al., 2010). Rodent models have also provided fruitful information linking the Roux-en-Y intervention to alterations in dorsal striatum monoamine homeostasis (Reddy et al., 2014). In line with these latter findings, our study is the first to point to a mechanistic link between gastrointestinal rerouting and striatal dopamine. Our protocol, which combines behavioral assays, dopamine measurements, and optogenetic stimulation in bariatric mice, may additionally assist in resolving conflicting evidence regarding post-bariatric alterations in reward processing (Davis et al., 2012; Hajnal et al., 2012).

Recent rodent studies indicate that major neuronal reorganization takes place in lower brainstem after bariatric interventions (Ballsmider et al., 2015). Future research may reveal novel pathways linking brainstem function to duodenal-sensitive dopamine cells.

Experimental Procedures

Subjects A total of 135 adult male mice were used, including 84 C57BL6/J (Jackson Laboratory) and 48 D1-dopamine receptor Cre-recombinase male mice (Drd1a-cre+, strain EY262, Gensat). Three D2-dopamine receptor Cre-recombinase mice (B6.FVB(Cg)-Tg(Drd2-cre)ER44Gsat/Mmcd, Gensat) were used to assess Cre-dependent expression of viral constructs. At the time of experiments animals were 8–20 weeks old. All experiments were conducted in accordance with the J.B. Pierce Laboratory/Yale University and Albert Einstein College of Medicine/Yeshiva University regulations on usage of animals in research.

Stimuli Sucralose and glucose were obtained from Sigma and prepared fresh in tap water

Gastric catheter implantation Once animals had been anaesthetized with an I.P. injection of a ketamine/xylazine (100/15 mg kg-1), a midline incision was made into the abdomen. The stomach was exteriorized through the midline incision and in proximity to the pylorus a purse string suture was constructed, into which the tip of MicroRenathane tubing (Braintree Scientific Inc., Braintree, MA, USA) was inserted. The purse string was tightened around the tubing, which was then tunnelled subcutaneously to the dorsum via a small hole made into the abdominal muscle; a small incision to the dorsum between the shoulder plates was then made to allow for catheter exteriorization. Incisions were sutured and thoroughly disinfected and the exterior end of the catheter was plugged.

Duodenum-jejunum bypass To divert food from its natural course through the duodenum, the pylorus was ligated and, approximately 6–8cm below the pylorus, the jejunum was anastomosed to the body of the stomach.

Behavioral Protocol for Tasks Involving Intra-Gastric Infusions

Habituation phase

We designed the behavioral assay in such a way that one group of mice licked a spout containing the sweet stimulus (the artificial sweetener sucralose), with licks triggering intra-gastric infusions of glucose. This protocol was repeated daily, for one hour. For comparison, a second group of mice was similarly exposed to the taste of sucralose, but in this case sucralose licking triggered intra-gastric infusions of sucralose instead. Specifically, mice were trained to produce licks to a spout containing an artificial sweetener (sucralose 2mM) in order to receive intra-gastric infusions of either 2mM sucralose (sucralose-exposed groups) or 50% glucose. During the task, detected licks trigger intra-gastric infusions of one of the solutions lasting for 3 seconds at a rate of 0.6mL/min. Licks detected while an infusion is taking place have no programmed consequences. During habituation, mice were exposed to the task described above once a day, for 13 days, and were maintained on ad libitum water but food restricted (2.8g/day chow allowed, sufficient for maintaining ~90% of ad libitum body weight).

Rationale for pairing sweetener licking to intra-gastric infusions

We opted for this design (where animals lick for sucralose and receive intra-gastric infusions of either glucose or sucralose) because we were concerned that differences in the sensory qualities of sucralose and glucose could influence their responses to these compounds during the preload test.

Baseline levels and preload volumes

After the 1 hour-long protocol was employed for 13 consecutive days (the “habituation” phase above), mice in both groups were assessed in their persistence to seek the sweet reward upon being administered either glucose or sucralose intra-gastric preloads. The preload volume was determined, as explained above, by averaging the glucose volumes administered during the ad libitum habituation sessions (more precisely the average across days 9–13). The same volumes were then used to determine the sucralose preloads for both groups (so that all preloads differed in sugar content but resulted in comparable intra-gastric volume).

Sweet taste seeking after preload

After habituation, mice were tested in their sensitivity to satiating signals. Tests were performed daily (i.e. day 14 onwards). For intra-gastric preloads, volume is determined from the averaged infused volume across animals during habituation (~0.75mL glucose; same volume was used for sucralose preloads). Preload infusions were performed 10min previous to behavioral testing, at 100µL/min infusion rate. Importantly, in all tests glucose-exposed and sucralose-exposed mice were assessed in exactly the same conditions (including same preloads and post-preload tests). Sweet seeking behavior was then assessed in the two groups of mice (i.e. glucose-exposed and sucralose-exposed during habituation) after administering the preloads. As during habituation, sweet seeking after preload lasted for 1 hour. We tested all four possible combinations of preload and subsequent sweet seeking: 1) preload=glucose, intake=sucralose; 2) preload=glucose, intake=glucose; 3) preload=sucralose, intake=glucose; 4) preload=sucralose, intake=sucralose.

Acute administration of glucose to duodenum during sweet seeking We were also interested in determining if glucose duodenal sensing may produce acute stimulatory effects on sweet seeking. Mice were implanted with duodenal catheters and infused glucose or sucralose duodenally as animals licked the sucralose solution. 30µl of the glucose (100mg/kg) and sucralose (0.8mg/kg) solutions were injected at the rate of 5µl/min over 6 min immediately previous to the start of the 1-hour long behavioral sessions.

Dopamine measurements Microdialysis measurements were performed as in our previous studies (Ren et al., 2010; Tellez et al., 2013c). During the experimental sessions, microdialysate samples from the freely-moving mice are collected, separated and quantified by HPLC coupled to electrochemical detection methods (“HPLC-ECD”). Briefly, after recovery from surgery, a microdialysis probe (2mm CMA-7, cut off 6kDa, CMA Microdialysis, Stockholm, Sweden) is inserted into the striatum through the guide cannula (the corresponding CMA-7 model). After insertion, probes are connected to a syringe pump and perfused at 1.2µl/min with artificial CSF (Harvard Apparatus). After a 30 min washout period, dialysate samples are collected every 6 min and immediately manually injected into a HTEC-500 HPLC unit (Eicom, Japan). Analytes are then separated via an affinity column (PP-ODS, Eicom), and compounds subjected to redox reactions within an electrochemical detection unit (amperometric DC mode, applied potential range from 0 to ~2000 mV, 1mV steps). Resulting chromatograms will be analyzed using the software EPC-300 (Eicom, Japan), and actual sample concentrations will be computed based on peak areas obtained from 0.5pg/µl dopamine standards (Sigma) and expressed as % changes with respect to the mean dopamine concentration associated with baseline (i.e. previous to behavioral task) samples. Locations of microdialysis probes were confirmed histologically (Figure S2E–F).

Stereotaxic viral injections and optic fibers implantation For animals used in the optogenetics experiments, anaesthesia was induced with an I.P. injection of a ketamine/xylazine (100/15 mg kg-1) and the mouse was placed on a stereotaxic apparatus (David Kopf, Tujunga, CA, USA) under constant flow of ~1% isoflurane anaesthesia (1.5 l min−1). All viral injections were done bilateral, using modified microliter syringes (Hamilton) with a 22G needle. The tip of the needle was placed at the target regions and the injections were performed at a rate of 0.1µL min−1 (for coordinates and volumes see below). Once the injection was finished the needle was left in place for 10 min and then slowly removed. Immediately after the viral injection the optic fibers were implanted (200 µm core, 0.22NA, Doric Lenses). To allow time for viral expression, animals were housed for at least 2 weeks following injection before any experiments were initiated. Drd1a-Cre mice were used for injections at the dorsal and ventral striatal regions. A subgroup of animals sustained the duodenal-jejunal intervention. The coordinates used for dorsal striatum were AP=1.0 mm, ML=±1.7 and DV=−3.0 mm (from skull surface) and 1 µL of AAV-EF1a-DIO-hChR2(H134R)-EYFP virus (serotype 5, University of North Carolina Vector Core) was injected per hemisphere. The coordinates used for ventral striatum were AP=1.5 mm, ML=±0.6 and DV=−4.5 mm (from skull surface) and 0.5 µL of either the AAV-EF1a-DIO-hChR2(H134R)-EYFP was injected per hemisphere.

Behavior-coupled Optogenetics To couple consumption to laser activation, detected licks triggered a 3s-long 473nm blue laser pulse via TTL signals (see Figure 1A). Licks produced during stimulation had no programmed consequence. Intensity at tip of fibers was estimated at approximately 7mW. We chose to use 3s as stimulation length based on the fact that this is equivalent to the duration of intra-gastric infusions. The efficiency of these parameters was confirmed in slice electrophysiology experiments performed on 3 weeks old D1-Cre mice (see below). D1r-Cre mice used in optogenetics experiments were exposed to the Habituation and Intra-gastric satiating preloads tests above. DJB D1r-Cre mice were through two rounds of Habituation with intra-gastric glucose infusions, in one of them laser being activated during the last 4 Habituation sessions.

Slice electrophysiology On the day of the experiments, Drd1a-cre mice with selective ChR2 expression in striatal D1r neurons were anesthetized with isoflurane and decapitated for electrophysiological identification of striatal D1r-neurones. Brains were quickly removed and immersed in an ice-cold high-sucrose solution containing (in mM): 220 sucrose, 2.5 KCl, 6 MgCl2, 1 CaCl2, 1.23 NaH2PO4, 26 NaHCO3, and 10 glucose (gassed with 95% O2 / 5% CO2; 300–305 mOsm). Coronal brain slices 300 µm thick were sectioned using a vibratome. Brain slices were then transferred to an incubation chamber filled with an artificial CSF (ACSF) solution containing (in mM) 124 NaCl, 2.5 KCl, 2 MgCl2, 2 CaCl2, 1.23 NaH2PO4, 26 NaHCO3, and 10 glucose (gassed with 95% O2 / 5% CO2; 300–305 mOsm) at room temperature (22 °C). After a 1–2 hrs recovery period, slices containing striatum were selected and transferred to a recording chamber mounted on a BX51WI upright microscope (Olympus, Tokyo, Japan). The recording chamber was perfused with a continuous flow of gassed ACSF. A dual-channel heat controller (Warner Instruments, Hamden, CT) was used to control the temperature of recording solution at 33 ± 1 °C. Whole-cell patch-clamp recordings were performed on striatum D1r neurons that were visualized using an infrared-differential interference contrast (DIC) optical system combined with a monochrome CCD camera and a monitor. Pipettes were pulled from thin-walled borosilicate glass capillary tubes (length 75 mm, outer diameter 1.5 mm, inner diameter 1.1mm, World Precision Instruments, Sarasota, Fl) using a P-97 micropipette puller (Sutter Instruments, Novato, CA). Pipette solution containing (in mM) 145 K-gluconate, 1 MgCl2, 10 HEPES, 1.1 EGTA, 2 Mg-ATP, 0.5 Na2-GTP, and 5 Na2-phosphocreatine (pH 7.3 with KOH; 290–295 mOsm) were used for whole-cell recording. The pipettes of resistances ranging from 3 to 6 MΩ were used for experiment. EPC-10 patch-clamp amplifier (HEKA Instruments, Bellmore, NY) and PatchMaster 2.20 software (HEKA Elektronik, Lambrecht/Pfalz, Germany) were used to acquire and analyze data. Pipette and cell capacitance were compensated during experiment and neurons which the series resistance was >20 MΩ and changed >15% were excluded from the statistics. Traces were processed using Igor Pro 6.36 (Wavemetrics, Lake Oswego, OR). A LED array (BXRAC2002, Bridgelux, Livermore, CA) was used to evoke the stimulation for optogenetic activation of channels in brain slices. Continuous stimulation and stimulation of 10 ms duration with different frequencies (1, 5, 10 or 20 Hz) were used in the experiment to test photostimulation-evoked responses.

Caspase-mediated ablation of dopamine-excitable cells To achieve Cre-dependent caspase-mediated ablation of dopamine-excitable neurons in striatum, the Cre-dependent viral construct AAV-flex-taCasp3-TEVp (serotype 5, University of North Carolina Vector Core) was stereotaxically injected into the dorsal or ventral striatum of D1r-Cre mice at same coordinates used for optogenetic experiments. The two control groups included D1r-Cre mice transfected with AAV5-GFP constructs, and non-Cre mice transfected with AAV-flex-taCasp3-TEVp.

Cell-specificity of the ablations was confirmed via neuroanatomical retrograde tracing methods. The fluorescent retrograde tracer Red Retrobeads (LumaFluor) was injected into the Substantia Nigra, pars reticulata (0.1µl), the exclusive ipsilateral target of D1r-expressing neurons of dorsal striatum on one hemisphere, and into the Globus Pallidus (0.1µl), the exclusive ipsilateral target of D2r-expressing neurons of dorsal striatum, on the other hemisphere. It was then confirmed, using fluorescent microscopy, the strong labeling throughout dorsal striatum on the same hemisphere in which the retrograde fluorescent beads were injected into globus pallidus, and weaker labeling throughout dorsal striatum on the same hemisphere in which the retrograde fluorescent beads were injected into Substantia Nigra pars reticulata. Equivalent Retrobead injections were made into the ventral pallidum, the preferential target of ventral striatal D1r-expressing neurons.

cFos quantification and Histological procedures Mice were deeply anesthetized with a dose of ketamine and xylazine (400 mg ketamine + 20 mg xylazine kg body weight−1 I.P.). All animals were transcardially perfused with filtered saline, followed by 4% paraformaldehyde. Following perfusion, brains were left in 4% paraformaldehyde for 24 hours and then moved to a 20% sucrose solution in PBS for 2 days. Brains were then frozen and cut into four series 40 µm sections (either coronal or sagittal) with a sliding microtome equipped with a freezing stage. To identify fibre and electrode locations, relevant sections were identified and mounted on slides. Sections were then photographed under bright field and fluorescence. For c-fos measurements, unilateral stimulation was performed using 1 minute ON/1 minute OFF cycles during 10 minutes. For striatal stimulation 0.5s on/0.5s off cycles were used during the ON cycles. 90 minutes after the stimulation protocols mice were sacrificed and perfused as described before. To visualize Fos immunoreactivity, the ABC/DAB procedure was used. Briefly, brain sections were first rinsed with 0.02 M potassium phosphate buffer (KPBS, pH 7.4), then immersed into a incubating solution [2% normal goat serum (NGS) and 0.3% Triton X-100 in KPBS] containing a rabbit polyclonal antiserum directed against the N-terminal region of the Fos gene (Ab-5, Calbiochem; dilution 1:10000) and incubated at 4 °C for 36 h. After primary antibody incubation, tissues were washed in 0.02 M KPBS and incubated at room temperature for 2h with goat anti-rabbit, biotinylated secondary IgG (anti-rabbit IgG, Vector Laboratories, 1:200), washed again in 0.02 M KPBS and subsequently reacted for 1 h with avidin-biotin-peroxidase complex (“ABC” method, Vectastain Elite ABC kit, Vector Co) at room temperature. A nickel diaminobenzidine (Nickel-DAB) glucose oxidase reaction was used to visualize Fos-like immunoreactive cells. Finally, sections were washed in KPBS and mounted, air-dried, dehydrated in alcohol, cleared in xylene, and coverslipped. Fos expression was analysed and quantified as follows: Coronal sections at ~160µm intervals throughout the rostral-caudal extent of the dorsal striatum were photographed at 10× magnification and montaged with Adobe Photoshop to preserve anatomical landmarks. Fos+ neurons were counted manually on each slice (5 slices per animal) and expressed as the cumulative sum of Fos+ neurons within the relevant regions for each animal.

Blood Glucose and Insulin measurements Wild-type mice were habituated to intra-gastric infusions and to handling by experimenters for two days previous to blood collection. The mice were food-restricted for 8 hours previous to the start of the experiment. Blood samples were collected from the tail vein. Blood glucose was measured immediately with a handheld glucometer (OneTouch), and data are presented as milligrams per deciliters. Blood samples used for insulin measurement were collected into vials containing EDTA to reach a final concentration of 1.735mg/mL, and centrifuged immediately after collection. The freshly prepared serum was stored at −20C for later analysis. Insulin concentrations were determined detected using standard ELISA methods (Rat/Mouse Insulin ELISA kit, Cat.# EZRMI-13K" EMD Millipore).

Statistical Analyses Data analyses were performed using SPSS (PASW Statistics Release 18.0.0) and made use of linear model analyses as well as two-way (repeated measures) ANOVAs and post-hoc tests as relevant. Data are reported as the mean ± standard error of the mean (SEM).

Supplementary Material

Acknowledgments

This work was supported by NIH grants R01DC014859 and R01CA180030 (to I.E.A.); R01 DK103176, DK084052, and NS48476 (to A.P.); R01DK105441, 5P60DK020541, and P30DK026687 (to G.J.S.,); China Scholarship Council 201206260072 (to W.H.); and FAPESP (Sao Paulo) 2013/09405-3 (to T.L.F.).

Abbreviations

DS-Casp

Caspase-driven D1r-dependent lesions in dorsal striatum of D1r-Cre mice (N=7)

VS-Casp

Caspase-driven D1r-depedent lesions in ventral striatum of D1r-Cre mice (N=7)

WT-Casp

Caspase-driven lesions in dorsal striatum of non-Cre mice (N=7)

D1-CTL

D1r-dependent viral GFP expression in dorsal striatum of D1r-Cre mice (N=6). n.s. = non-statistically significant

Footnotes

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