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Vgat-ires-Cre31, Sst-Cre32 and PV-Cre33 knock-in mice (The Jackson Laboratory, Bar Harbour) and C57BL/6 male mice, 10–25 weeks old, were used. Mice were housed under standard conditions in the animal facility and kept on a 12 h light/dark cycle. All procedures were performed in accordance with national and international guidelines, and were approved by the local health authority (Landesamt für Gesundheit und Soziales, Berlin) and the Stanford University Institutional Animal Care and Use Committee. Injections were performed as described previously22, 34. Mice were anaesthetized with isoflurane and placed in a stereotaxic head frame (David Kopf Instruments). A 34-gauge bevelled metal needle connected via a tube with a microsyringe pump (PHD Ultra, Harvard Apparatus) was used to infuse viruses at a rate of 100 nl min−1. After infusion, the needle was kept at the injection site for 10 min and then slowly withdrawn before the incision was sutured. Optogenetic constructs from K. Deisseroth, purchased from Penn Vector Core, UNC Gene Therapy Center Vector Core, or provided by K. Deisseroth, were used. For manipulations of the LS–LH pathway, Vgat-Cre mice were injected bilaterally in the LS (right: anterior-posterior (AP) 0.74, lateral (L) 0.38, ventral (V) 3.3; 3.1 mm; left: AP 0.62, L 0.33, V 3.45; 3.0 mm) with 0.3 μl per injection site of AAV2/5-Ef1a-DIO-ChETA(E123T/H13R)-eYFP-WPRE-hGH (Penn Vector Core, titre 1.75 × 1012 viral genomes (vg) per ml). Sst-Cre mice were injected bilaterally in the LS (right: AP 0.38, L 0.4, DV 3.0, 2.7 mm; left: AP 0.26, L 0.4, V 3.0, 2.5 mm) with 0.125–0.25 μl per injection site of AAV2/5-Ef1a-DIO-ChETA(E123T/H13R)-eYFP-WPRE-hGH, titre 1.75 × 1012 vg ml−1 (Penn Vector Core) or 0.2 μl per injection site of AAVdj-nEF-DIO-NpHR-TS-p2A-hChR2(H134R)-eYFP (eNPAC2.0, titre 6.1 × 1012 vg ml−1) or 0.125–0.2 μl per injection site of AAV2-EF1a-DIO-eYFP-WPRE-hGH (Penn Vector Core, titre 2 × 1012 vg ml−1). For manipulation of LH cells, Vgat-Cre mice were injected bilaterally in the LH (AP −1.5, L ±1, V 5.4 mm) with 0.3 μl per injection site of AAV2/5-Ef1a-DIO-ChETA(E123T/H13R)-eYFP-WPRE-hGH (Penn Vector Core) or 0.3 μl per injection site of AAVdj-nEF-DIO-NpHR-TS-p2A-hChR2(H134R)-eYFP (eNPAC2.0, titre 6.1 × 1012 vg ml−1) or 0.3 μl per injection site of AAV2-EF1a-DIO-eYFP-WPRE-hGH (Penn Vector Core, titre 2 × 1012 vg ml−1). For manipulations of the mPFC–LH pathway, mice were bilaterally injected in the mPFC (AP 1.70, L ±0.35, V 2.85 mm) with 0.25–0.5 μl per injection site of AAV2-CaMKIIa-hChR2(H134R)-eYFP (Penn Vector Core, titre 2.55 × 1012 vg ml−1) or 0.25–0.5 μl per injection site of AAV5-CaMKIIa-ChETA(E123T/H134R)-eYFP-WPRE-hGH (Penn Vector Core, titre 1.26 × 1013 vg ml−1), or 0.25–0.5 μl per injection site (AAVdj-hSyn-NpHR-TS-p2A-hChR2(H134R)-eYFP (eNPAC2.0, titre 2.9 × 1013 vg ml−1) or 0.5 μl per injection site of AAV2-CaMKIIa-eYFP (University of North Carolina Vector Core, titre 5 × 1012 vg ml−1). For CLARITY experiments, mice were injected in the mPFC (AP 2.0, L 0.3, V 2.6 mm) with 1 μl AAV8-CaMKIIa-eYFP-NRN. For synaptophysin imaging, 1 μl AAV8-CaMKIIa-synaptophysin-mCherry (7 × 1013), was injected in the mPFC (AP 2.0, L 0.3, V 2.6 mm). Optic fibre implants were fabricated from 100 μm diameter fibre (0.22 numerical aperture (NA), Thorlabs) and zirconia ferrules (Precision Fibre Products). For optogenetic manipulations of mPFC–LS pathway, mice were implanted with optic fibre implants on top of the LS (right, AP 0.1, L 0.25, V 2.25 mm, left, AP 0.5, L 0.3, V 2.7 mm). For optogenetic manipulations in the LH, optical fibres were bilaterally (for LS–LH and LH stimulation or inhibition) or unilaterally (LH stimulation combined with the LH silicon probe recordings) implanted above the LH (AP −1.6, L 1, V 4.8 mm). Arrays of single tungsten wires (40 μm, California Fine Wire Company), stationary implanted linear silicon probes (CM32, NeuroNexus Technologies), or movable probes (B32 or B64, NeuroNexus Technologies) mounted on a microdrive35 were implanted as described previously22, 26. The following coordinates were used for electrode implantations in the LS: AP 0–0.5, L 0.2–0.45, V 2.3–3.4 mm (B32 probes, B64 probes (mPFC–LS co-implantations), CM32, wire arrays); LH: AP −1.6, L 1, V 4.7 mm (B32 probes, B64 probe, wire arrays); mPFC: AP 1.4–1.9, L 0.3, V 3.0 mm (B64 probes (mPFC–LS co-implantations), wire arrays); dorsal hip: AP −2.1, L 1.6, V 1.5 mm (wire arrays), ventral hip: AP −3.16, L 2.5–3.5, V 4.0 mm (wire arrays). Reference and ground electrodes were miniature stainless-steel screws in the skull above the cerebellum. The implants were secured on the skull with dental acrylic. Electrodes were connected to operational amplifiers (HS-8, Neuralynx, or Noted BT) to eliminate cable movement artefacts. Electrophysiological signals were differentially amplified, band-pass filtered (1 Hz–10 kHz, Digital Lynx, Neuralynx) and acquired continuously at 32 kHz. A light-emitting diode was attached to the headset to track the mouse’s position (at 25 Hz). Timestamps of laser pulses were recorded together with electrophysiological signals. A 3-m-long fibreoptic patch cord with protective tubing (Thorlabs) was connected to a chronically implanted optical fibre with a zirconia sleeve (Precision Fibre Products), which allowed the mice to explore an enclosure freely or perform a behavioural task during optogenetic manipulations. Subjects were randomly assigned to the experimental conditions. For optogenetic stimulation, the patch cord was connected to a 473-nm diode-pumped solid-state laser (R471005FX, Laserglow Technologies) with an FC/PC adaptor. The laser output was controlled using a stimulus generator and MC_Stimulus software (Multichannel Systems). Optogenetic stimulation of LS–LH and mPFC–LS projections consisted of 5 ms blue (473 nm) light pulses, at 66.7 Hz or a control, non-gamma (theta) intensity-matched stimulation (167 Hz bursts of 4 ms pulses repeated at 9 Hz) with the light power output (during light-on parts of illumination cycles) of 10–25 mW from the tip of the patch cord measured with a power meter (PM100D, Thorlabs). Optogenetic stimulation of LH somata consisted of 5 ms blue (473 nm) light pulses, at 20 Hz, with light power output (during light-on parts of illumination cycles) of 10–25 mW from the tip of the patch cord. For bilateral optogenetic inhibition, optic fibre implants were connected via patch cords to a 593-nm diode-pumped solid-state laser (R591005FX, Laserglow Technologies) using a multimode fibre optic coupler (FCMM50-50A-FC, Thorlabs), continuous yellow light, approximately 20 mW from the tip of each patch cord. Duration of light delivery is described below for each type of behavioural experiment. For control experiments, mice expressing YFP in the same brain regions (mPFC or LS) were used, and optostimulation was performed as described above. For within-animal comparisons in sessions in which food intake was measured during LH stimulation (Extended Data Figs 2i, 8c, d, f), optic patch cords were connected to dummy ferrules, attached to the headset, and light of the same wavelength and power as during opsin-activating stimulation was delivered. Free-access feeding model. This was performed in a chamber similar to that described previously36 (Fig. 1a). Mice freely explored a custom two-chamber (30 × 50 × 20 cm) enclosure, which contained food and water in designated areas (each area 10 × 10 cm; see Fig. 1a). Food (Dustless Precision Pellets, 20 mg, Rodent Purified Diet, Bio Serv) was provided either in a food cup or in a pellet feeder (Coulbourn Instruments Pellet Feeder H14-23M; sampling rate 10 Hz, one nose poke led to the delivery of one food pellet). Before experiments, mice received these pellets in the homecage for at least 2 days, and were habituated to the behavioural setup for at least 3 days. Coordinates of 10 × 10 cm food, drinking, non-food corner zones and a control zone located in the non-food compartment were defined. Times of entering and leaving each zone were extracted from the mouse’s position-tracking data. An approach rate was defined as the distance between a position of the mouse and the centre of the food zone, the drinking zone or the control zone, divided by the time it took to enter a respective zone. For each experiment, a corner zone, one of which was a food zone, visited first after the onset of stimulation was detected. Latency to enter each zone was defined as the time between the beginning of optogenetic stimulation and the first entry of the mouse into a zone, with the mouse staying in the zone for at least 1 s. To account for differences in distances to a zone after the stimulation onset, in each experiment we have normalized the latency after stimulation onset to the average latency of entering the same zone from the same distance during the baseline. Duration of experimental sessions and optogenetic manipulations are described below. Optogenetic activation of LS –LH projections. Mice explored the enclosure for 30 min: 10 min before stimulation, 10 min during optogenetic stimulation, 10 min after stimulation. Blue light (473 nm) was bilaterally delivered over LS–LH projections, in 5 ms pulses at 66.7 Hz or using a control, non-gamma (theta) intensity-matched stimulation (167 Hz bursts of 4 ms pulses repeated at 9 Hz), with light power output of 10–25 mW. For brief gamma stimulation, 5 ms pulses at 66.7 Hz were delivered for 30 s, followed by a break of 2 min, during a 10 min period. Optogenetic inhibition of LS –LH projections during food approach. Mice explored the enclosure for 30 min. Each time the mouse crossed the border of a food-approach area (20 × 20 cm, marked as an orange dotted line on Extended Data Fig. 6e), continuous yellow (593 nm) light was bilaterally delivered over LS–LH projections. Light delivery stopped each time a mouse left the approach zone. Optogenetic activation of LS –LH projections during free-access to high-fat food. Mice explored the enclosure for 20 min. Blue light (473 nm) was bilaterally delivered over LS–LH projections, in 5 ms pulses at 66.7 Hz, with light power output of 10–25 mW. High-fat food pellets (Testdiet, 60% energy from fat) were weighted before and after the experiment, to calculate the amount of food (>5 mg) consumed per session. Optogenetic stimulation of LH cells. Mice explored the enclosure for 30 or 60 min: 10 or 20 min before stimulation, 10 or 20 min during optogenetic or control light stimulation, 10 or 20 min after stimulation, Dustless precision pellets (BioServ) were counted to measure the amount of food (>1 pellet) consumed per session. Blue light (473 nm) was delivered bilaterally, in 5 ms pulses at 20 Hz. Optogenetic inhibition of LH cells in food-deprived mice. Mice received approximately 2.5–3.0 g of standard chow daily; the mouse weight was controlled and weight loss did not exceed 10%. Dustless precision pellets (BioServ) were counted to measure the amount of food consumed by hungry mice (>3 pellets in baseline) per session. The experiments consisted of four epochs: 10 min light-on (optogenetic or control stimulation), 10 min light-off, 10 min light-on, and 10 min light-off. This was performed in a custom two-chamber enclosure similar to the one used for the free-feeding model (30 × 50 × 20 cm). One of the chambers contained a familiar object, whereas the other contained a new object. Before experiments, mice were habituated to the enclosure containing two objects, then for each experimental session one of the objects (new object) was replaced, whereas the object in the other chamber (familiar object) remained the same. Optogenetic stimulation started as the mouse was put in the enclosure. Mice freely explored the enclosure maximally for 2 min, otherwise a session was finished once the mouse visited both objects. Blue light (473 nm) was bilaterally delivered over LS–LH projections, in 5 ms pulses at 66.7 Hz, with light power output of 10–25 mW. Spatial non-matching to place testing on the T-maze was performed as described elsewhere37. The T-maze (start arm: 46 × 11 × 10 cm, choice arm: 80 × 11 × 10 cm; see Fig. 4p) was made of pieces of wood painted dark-grey. For spatial non-matching to place testing, each trial consisted of a sample run and a choice run. During the sample run, mice could run only to one arm (left or right, according to a pseudorandom sequence with equal numbers of left and right turns per session) because another arm was blocked by a wooden block. A reward (0.1 ml condensed milk or a 20 mg food pellet) was available in the food well at the end of the arm. After the sample run, mice stayed in another, familiar, enclosure for 10–15 s. The block was then removed, and mice were placed at the end of the start arm to perform the test run. Mice were rewarded for choosing the previously unvisited arm (that is, for alternating). For this test and all subsequent experiments, entry into an arm was defined as when a mouse had placed all four paws into the arm. Mice ran one trial at a time with inter-trial intervals of 3–5 min. Each mouse conducted 20–40 trials in total (10 trials per day). For a subset of experiments, mice were water-restricted and water was used as a reward instead of food. A number of slow (30–60 Hz) or fast (60–90 Hz) gamma-oscillation episodes (detected as described below) in the start or choice arm was normalized by dividing by the mean number of gamma events in each arm during the whole experiment. Optogenetic activation or inhibition of mPFC–LS projections in the T-maze. Optogenetic stimulation started as the mouse was put at the end of the start arm, and finished when the mouse reached the reward. Blue light was delivered on mPFC–LS projections in 5-ms pulses at 66.7 Hz, or in a non-gamma (theta) intensity-matched stimulation protocol, described above for free-feeding model. For inhibition, yellow light was delivered onto mPFC–LS projections continuously during the run. LFP was obtained by down-sampling of the wide-band signal to 1,250 Hz using Neurophysiological Data Manager38 (http://neurosuite.sourceforge.net/). Gamma oscillations were detected at 30–60 Hz, 60–90 Hz and, for the analysis shown in Extended Data Fig. 1i, 90–120 Hz, bandpass filtered, rectified and smoothed with a 15-ms window LFP signals. Events with amplitudes exceeding 2 s.d. above mean for at least 25 ms were detected13. The beginning and the end of oscillatory epochs were marked at points at which the amplitude fell below 1 s.d. Power spectral density and coherence were computed using the multitaper method (NW = 3). For the analysis of association between gamma power and approach rate, the cumulative power in the 30–60 and 60–90 Hz bands as well as the approach rate (see ‘Behavioural assays’) was computed, and for each 1-s recording epoch, gamma power was z-transformed. Values within 10 s before entry in the food or drinking zones were statistically evaluated. Current source density (CSD) maps (versus time and depth) were computed as previously described37, 39. LFP depth profiles, recorded using CM32 probes with the spatial sampling of 100 μm, were averaged using peak gamma oscillations detected in an LS channel as triggers. The second spatial derivative of the obtained voltage traces, that is, CSD, indicates locations of current sinks and sources in the extracellular space40. For the analysis of mPFC–LS and hippocampus–LS coherence, normalized current flow density in the LS was computed by subtraction of gamma-band filtered LFP signals, recorded by a pair of wire electrodes in the LS against a common screw-reference above cerebellum40, 41. Action potentials were detected in a high-pass filtered signal using NDManager16 (http://neurosuite.sourceforge.net/). Spike waveforms were extracted and represented by the first three principle components and by amplitudes of action potentials. Spike sorting was performed automatically using KlustaKwik42 (http://klusta-team.github.io/klustakwik/) followed by manual clusters adjustment using Klusters38. Isolation distance42 was computed for sorted units (LH: 101.5 ± 8.0, LS: 66.3 ± 4.6, mPFC: 56.3 ± 3.6). Phase of gamma oscillations was computed for signal epochs within detected gamma episodes as described elsewhere37, 43. In brief, 0° and 360° were assigned to troughs of each gamma cycle and 180° to a cycle peak, phases for each data sample between these points were computed using linear interpolation13, 37. Subsequently, gamma phases were obtained for data samples when action potentials were emitted, for each recorded neuron, and firing phase histograms were computed. A possible asymmetry of oscillation cycles leads to a different number of phase samples composing ascending and descending parts of the cycle and can bias firing phase histograms39. To prevent this, we tested uniformity of grand gamma phase distributions for each recording using the Rayleigh test and, if significantly non-uniform, computed a deviation of a grand phase histogram from uniformity, via division by the average across all bins. In such recordings, firing histograms were normalized by the corrected grand phase histogram37, 44, 45. Each firing phase histogram was normalized by its total number of spikes. Circular uniformity, mean phase and the resultant vector length were estimated for each histogram. Before averaging, individual histograms were convolved with the Gaussian kernel46 of size 0.65 s.d. Putative LH neurons were optogenetically identified based on rapid (<10 ms lags in laser pulse onset-triggered cross-correlations, computed with 1-ms bins) increase of firing after onset of laser pulses. Reliability of light-induced responses was estimated as a probability of the maximal light-induced spike count in a Poisson distribution computed for cross-correlogram (CCG) delays in the pre-pulse baseline26, 47. To estimate gamma-rhythmic responses of LH cells to LS–LH stimulation, a cross-correlation (CCG) with the times of LS–LH light stimulation was computed for each cell. To avoid spurious CCG peaks at the stimulation frequency, every second time stamp of light was used as a trigger. A reshuffled CCG was computed using light times shifted to a baseline, light-off, recording epoch. A power spectrum of the response of a cell was then obtained by subtracting the power spectrum of the reshuffled CCG from the power spectrum of the stimulation CCG. Firing of LH neurons in the free-access feeding model was evaluated as described elsewhere for quantification of positional firing37, 48. Firing maps were computed by dividing the number of spikes in a given spatial pixel (2 × 2 cm) by the time spent in this pixel. Periods of immobility (speed <3 cm s−1) were excluded from the analysis. Peak firing rate was defined as the maximum firing rate over all pixels in the environment. For calculation of food-zone preference (FZ-match index), the average firing rate of a cell in the food zone was divided by the average firing rate in a control zone of the same size (10 × 10 cm), located in the non-food compartment of the enclosure. For the analysis of firing during gamma oscillations, cells were split in ‘FZ-match’ or ‘FZ-mismatch’ groups based on an FZ-match index higher or lower than 1, respectively. For identification of LH cells, excited in response to LH or LS –LH optogenetic stimulation, the number of spikes (x) was computed for each 100-ms bin during the baseline (10 min) and within 3 s after stimulation onset. A distribution derived from the baseline was fitted to a Poisson distribution, and rate parameter λ was estimated. A bin with maximal count of spikes during stimulation was assigned to observed value x . P value was defined as P(x ≥ x ), in which x follows Pois(λ). The firing rate ratio (R) between 3 s baseline and stimulation epochs was computed for each stimulation epoch and then averaged across stimulation epochs. A cell was classified as excited if P < 0.05 and R > 1. Code is available from the corresponding authors upon request. Each statistical test was used according to the design of the experiment and the structure of the data. Two-group comparisons were performed using t-test, Mann–Whitney or Wilcoxon matched-pairs tests depending on the normality of a distribution. Assessment of effects in experiments involving several conditions was performed using ANOVA, followed, when appropriate, by Bonferroni (for pre-selected contrasts) or Tukey tests, adjusting for multiple comparisons. Grubbs’ test was used to exclude outlier points from behavioural datasets. Depending on the normality of a distribution, Pearson’s or Spearman’s correlations were computed. For group comparisons, two-tailed statistical tests were applied. Sample size was determined according to the accepted practice for the applied assays, no statistical methods were used to predetermine sample size. Conditions of the experiments were accounted during design of analysis algorithms, computations were subsequently performed blindly using automatic selection of data from a database. A detailed description of statistical analysis is provided in the statistical section of the Supplementary Information. Descriptive statistics are reported as mean ± s.e.m. Primary cultured neurons were prepared from the hippocampi of P Sprague–Dawley rat pups (Charles River Laboratories), as described previously49. CA1 and CA3 hippocampal regions were taken out and digested with 0.4 mg ml−1 papain (Worthington), and plated onto 12-mm glass coverslips that were pre-coated with 1:30 Matrigel (Beckton Dickinson Labware). Cultures were kept under neurobasal-A medium (Invitrogen) containing 1.25% FBS (HyClone), 4% B-27 supplement (Gibco), 2 mM glutamax (Gibco) and 2 mg ml−1 fluorodeoxyuridine (FUDR, Sigma) and plated at a density of 65,000 cells per well in 24-well plates. The plates were incubated at 37 °C in a humid incubator with a constant level of 5% CO . Cultured neurons were transfected at 6–10 days in vitro (DIV). A DNA–CaCl mix composed of the following was prepared for transfection per well: 1 μg of endotoxin-free DNA for recordings, 1.875 μl 2 M CaCl , and sterile H O for a total volume of 15 μl. Another 15 μl of twice-filtered HEPES-buffered saline (HBS, in mM: 50 HEPES, 1.5 Na HPO , 280 NaCl, pH 7.05 with NaOH) for each DNA–CaCl mix. This mix was incubated at room temperature for 20 min. In the meantime, the neuronal growth medium was removed from the wells and kept at 37 °C, and replaced with 400 μl pre-warmed minimal essential medium (MEM). After incubation of the DNA–CaCl –HBS mix was complete, the mix was then added dropwise into each well, and plates were incubated for 45–60 min at 37 °C. Once the transfection was complete, each well was washed three times with 1 ml of pre-warmed MEM. The MEM was then replaced with the original neuronal growth medium, and plates were placed into the culture incubator at 37 °C. Whole-cell patch-clamp recordings of cultured hippocampal neurons were performed 3–5 days after transfection with the construct AAVdj-hSyn-NpHR-TS-p2A-hChR2(H134R)-eYFP (eNPAC2.0). Expression of the construct was identified by eYFP fluorescence. The external recording medium was composed of the following (in mM): 125 NaCl, 2 KCl, 25 HEPES, 2 CaCl , 2 MgCl , 30 d-(+)-glucose. pH 7.3, with synaptic transmission blockers d-2-amino-5-phosphonovaleric acid (AP5; 25 μM), 2,3-dihydroxy-6-nitro-7-sulfamoyl-benzo[f]quinoxaline-2,3-dione (NBQX; 10 μM), and gabazine (10 μM). The intracellular recording solution contained (in mM): 130 potassium gluconate, 10 KCl, 10 HEPES, 10 EGTA and 2 MgCl . An upright microscope (BX61WI, Olympus) with infrared differential interference contrast (IR-DIC) was used for visualization and recording of the expressing neurons. A Spectra X Light engine (Lumencor) attached to the fluorescent port of the microscope was used for light application, for detecting eYFP expression and for blue or yellow light delivery for opsin activation. A 475/28 nm and a 586/20 nm filter (Chroma) were used for blue and yellow light respectively (Chroma). A power meter (ThorLabs) was used to measure the light power through the microscope objective, and light power density was set at 5 mW mm−1 (ref. 49). Recordings were obtained using a MultiClamp700B amplifier, 1440A Digidata digitizer, and pClamp10.3 software (Molecular Devices). Data were analysed with pClamp10.3 and SigmaPlot (SPSS). Photocurrent amplitudes at blue and yellow were measured at steady-state at the end of a 1-s light stimulation protocol. To measure spike inhibition probability at 586 nm, we first applied a 50–200 pA electrical current injection (depending on spike threshold of the recorded cell) to induce spiking in the expressing neurons. Spike inhibition probability was calculated as the percentage in which yellow light application inhibited spiking during the electrical current injection. To measure spike generation probability with blue light, we applied 5-ms width pulses of 475 nm light at 5 or 20 Hz frequency, and calculated the percentage of action potentials generated by the blue light pulse train. Series resistance was carefully monitored for stability throughout the recordings. To ensure accurate measurements of voltage-clamp recordings, data were incorporated for analysis only if the series resistance was below 25 MΩ and changed less than 20% throughout the recording. Standard whole-cell slice patch-clamp recordings were undertaken after slice preparation of at least 2-month-old mice. In brief, after gluing a block of brain with cyanoacrylate glue to the stage of a Campden Vibroslice, coronal brain slices of 250-μm thickness containing the LH were cut while immersed in ice-cold slicing solution. Slices were incubated for 1 h in artificial cerebrospinal fluid (ACSF) at 35 °C then transferred to a submerged-type recording chamber. Living neurons containing fluorescent markers were visualized in acute brain slices with an upright Olympus BX61WI microscope equipped with an oblique condenser and appropriate fluorescence filters. After identifying appropriate neurons by their fluorescence, oscillatory currents of 10 pA amplitude (30, 50, 70 and 100 Hz) were injected for 5 s to the cell during whole-cell patch-clamp recordings. Recordings of membrane potentials were analysed in MatLab. To record selectively from Vgat neurons, a cross between Vgat-ires-Cre and CAG-tdTomato mice33 was used. To target MCH-Cre neurons selectively, MCH-Cre mice were injected into the LH (1.3 mm caudal from bregma; ±0.95 mm lateral from midline; and 5.25 and 5.15 mm ventral from brain surface) with a Cre-dependent ChR-mCherry. For brain slice recordings, ACSF and ice-cold slicing solution were gassed with 95% O and 5% CO , and contained the following (in mM) ACSF: 125 NaCl, 2.5 KCl, 1 MgCl , 2 CaCl , 1.2 NaH PO , 21 NaHCO , 2 d-(+)-glucose, 0.1 Na+-pyruvate and 0.4 ascorbic acid. Slicing solution: 2.5 KCl, 1.3 NaH PO.H 0, 26.0 NaHCO , 213.3 sucrose, 10.0 d-(+)-glucose, 2.0 MgCl and 2.0 CaCl . For standard whole-cell recordings, pipettes were filled with intracellular solution containing the following (in mM): 120 K-gluconate, 10 KCl, 10 HEPES, 0.1 EGTA, 4 K ATP, 2 Na ATP, 0.3 Na GTP and 2 MgCl , pH 7.3 with KOH. Multitaper power spectra of voltage traces and of injected current traces were computed and divided, resulting in impedance spectra. Mean impedance at ±1.5 Hz around stimulation frequency was computed. Brain hemispheres were clarified using the CLARITY procedure as described elsewhere27. In brief, a brain hemisphere was fixed in hydrogel solution (4% PFA, 1% acrylamide/bis) for 72 h at 4 °C. After polymerization (37 °C, 4 h), the brain hemispheres were clarified in 8% SDS for 8 days (at 40 °C), then washed three times with PBST (0.2% Triton X-100) for a total of 24 h at 37 °C. Hemisphere images were acquired with the Ultramicroscope II (Lavision Biotec)27. Samples were mounted to a custom 3D printed holder using RapidClear Mounting Gel (Sunjin laboratory). Brains were imaged using a 2×/0.5 NA objective at 0.8× zoom using a single light sheet illuminating from the dorsal side of the sample. Z-step was set to 4 μm. Fourteen horizontal focal points were set to each imaging plane to create a homogeneous field of view. For synapsin staining the brains were cut, after CLARITY processing, into 1-mm-thick sections. Primary antibody: rabbit anti-synapsin (Cell Signaling, 5297), 1:400, in 0.3% PBST, room temperature, 24 h; secondary antibody: donkey anti-rabbit (Alexa 594, Jacksonimmuno), 1:200, in 0.3% PBST, room temperature, 24 h; then sections were refractive index-matched and mounted in RapidClear CLARITY-specific gel (Sunjin Laboratory). Sections were imaged at bregma = 0.5 using Olympus FV1200 confocal, 40×, 1.3 NA, oil objective, at 4× zoom. For synaptophysin imaging, brains expressing CaMKIIa-synaptophysin-mCherry in the mPFC were cut into 0.5-mm-thick section for CLARITY clearing and imaging. The sections were imaged at bregma = 0.5 using Olympus FV1200 confocal, 40×, 1.3 NA, oil objective, at 4× zoom. After completion of the experiments, mice were deeply anaesthetized and electrolytic lesions at selected recording sites were performed. Subsequently, the mice were perfused intracardially with saline followed by 4% paraformaldehyde in PBS and decapitated. Brains were fixed overnight in 4% paraformaldehyde, equilibrated in 1% PBS for an additional night and finally cut into 40 or 50 μm slices using an oscillating tissue slicer (EMS 4500, Electron Microscopy Science). Brain slices were mounted (Flouromount Aqueous Mounting Medium, Sigma-Aldrich). Images were taken using an Olympus BX 61 microscope (×2/0.06 NA, ×10/0.3 NA and ×20/0.5 NA, dry) or using a Leica DMI 6000 microscope (×20/0.7 NA, ×63/1.4 NA; oil-immersion objectives). All data generated or analysed during this study are either included in this published article or are available from the corresponding authors on reasonable request.


News Article | March 9, 2016
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LAP-tTA and TRE-MYC mice were previously described and MYC expression in the liver was activated by removing doxycycline treatment (100 μg ml−1) from the drinking water of 4-week-old double transgenic mice for both TRE-MYC and LAP-tTA as previously described9, 13. C57BL/6 mice were obtained from NCI Frederick. Chemically induced HCC was established by intraperitoneal injection of diethylnitrosoamine (DEN) (Sigma) into 2-week-old male pups at a dose of 20 μg g−1 body weight13. Twelve-week-old male B6.Cg-Lepob/J (ob/ob) mice or wild-type control mice were obtained from Charles River. Foxp3–GFP mice were previously described31. NAFLD was induced by feeding mice with a methionine–choline-deficient (MCD) diet (catalogue number 960439, MP biomedical), a choline-deficient and amino-acid-defined (CDAA) diet (catalogue number 518753, Dyets) or a high-fat diet (catalogue number F3282, Bio Serv) for the indicated time10, 11, 32. The MCD diet was supplied with corn oil (10%, w/w), and no fish oil was added. Control diet was purchased from MP Biomedical (catalogue number 960441). Custom-made high- or low-linoleic-acid mouse diets were purchased from Research Diets. The modified diets were based on AIN-76A standard mouse diet, and are isocaloric (4.45 kcal g−1) and contained the same high-fat content (23%, w/w). Linoleic-acid-rich safflower oil and saturated fatty-acid-containing coconut oil were supplied at different ratios to yield 2% (w/w) for the low-linoleic-acid diet or 12% (w/w) for the high-linoleic-acid diet. C57BL/6 mice were fed with the high- or low-linoleic-acid diet for 4 weeks. MYC mice were injected i.p. with 50 μg CD4 antibody (clone GK1.5, BioXcell) every week for the indicated time period to deplete CD4+ T cells33. N-acetylcysteine (NAC) was given in drinking water (10 mg ml−1)34 for the indicated time period to prevent excess ROS production. Mitochondrial-specific antioxidant mitoTEMPO was purchased from Sigma. Mice received mitoTEMPO at a dose of 0.7 mg kg−1 per day25 by osmotic minipumps (ALZET). At the experimental end points, mice were killed. For flow cytometry analysis, single-cell suspensions were prepared from spleen, liver and blood as described previously. Red blood cells were lysed by ACK Lysis Buffer (Quality Biologicals). Parts of live tissue were fixed by 10% formaldehyde and subjected to H&E staining. Free fatty acids were purchased from Sigma. Lipid accumulation was detected by Oil Red O staining in frozen liver sections using the custom service of Histo Serv. Cells were surface-labelled with the indicated antibodies for 15 min at 4 °C. Flow cytometry was performed on BD FACSCalibur or BD LSRII platforms and results were analysed using FlowJo software version 9.3.1.2 (TreeStar). The following antibodies were used for flow cytometry analysis: anti-CD3-FITC (clone 17A2, BD Pharmingen), anti-CD4-PE (clone RM4–4, Biolegend), anti-CD4-APC (clone RM4–5, eBioscience), anti-CD8-Alexa Fluor 700 (clone 53–6.7 Biolegend), anti-CD45, anti-CD44-PE (clone IM7, eBioscience), anti-CD62L-PerCP/Cy5.5 (MEL-14, Biolegend), anti-CD69-Pacific blue (clone H1.2F3, Biolegend), PBS57/CD1d-tetramer-APC (NIH core facility). To determine cytokine production, cells were stimulated with PMA and ionomycine for 30 min, and then were fixed and permeabilized using cytofix/cytoperm kit (BD Pharmingen) followed by anti-IFN-γ-PE (clone XMG1.2, BD Pharmingen), anti-IL-17-PerCP/Cy5.5 (clone TC11-18H10.1, Biolegend) staining. Cell death and apoptosis were detected with annexin V-PE (BD Pharmingen) and 7-AAD (BD Pharmingen) staining according to the manufacturer’s instructions. Intrahepatic CD4+ lymphocytes were gated on the CD3hiCD4+ population from total live hepatic infiltrating mononuclear cells. Absolute numbers were calculated by multiplying frequencies obtained from flow by total live mononuclear cell count, then divided by liver weight. The antibodies used for human peripheral blood mononuclear cell (PBMC) staining are the following: anti-CD3-PE (clone SK7, BD Pharmingen), anti-CD4-FITC (clone RPA-T4, BD Pharmingen), anti-CD8-APC (clone RPA-T8, BD Pharmingen). Murine T assays were performed as described31. Briefly, liver T cells were isolated as CD4+GFP+ by flow-cytometry-assisted cell sorting from Foxp3–GFP mice kept on an MCD or control diet for 4 weeks. CD4+GFP− T effector (T ) cells (5 × 104) were stimulated for 72 h in the presence of irradiated T-depleted splenocytes (5 × 104) plus CD3ε monoclonal antibody (1 μg ml−1), with or without T cells added at different ratios. 3H-Thymidine was added to the culture for the last 6 h and incorporated radioactivity was measured. Freshly isolated splenocytes from MYC-ON MCD mice were incubated with 5 μg ml−1 of mouse α-fetoprotein protein (MyBioSource) for 24 h. Golgiplug was added for the last 6 h. Then, cells were fixed and permeabilized using cytofix/cytoperm kit (BD Pharmingen) followed by anti-IFN-γ-PE (clone XMG1.2, BD Pharmingen) staining. Primary mouse hepatocytes were isolated from MYC mice and cultured according to a previous report35. Briefly, mice were anaesthetized and the portal vein was cannulated under aseptic conditions. The livers were perfused with EGTA solution (5.4 mM KCl, 0.44 mM KH PO , 140 mM NaCl, 0.34 mM Na HPO , 0.5 mM EGTA, 25 mM Tricine, pH 7.2) and Gey’s balanced salt solution (Sigma), and digested with 0.075% collagenase solution. The isolated mouse hepatocytes were then cultured with complete RPMI media in collagen-I-coated plates. Hepatic fatty acid composition was measured at LIPID MAPS lipidomics core at the University of California (San Diego) using an esterified and non-esterified (total) fatty acid panel. Briefly, liver tissues were homogenized and lipid fraction was extracted using a modified Bligh Dyer liquid/liquid extraction method. The lipids were saponified and the hydrolysed fatty acids were extracted using a liquid/liquid method. The extracted fatty acids were derivatized using pentaflourylbenzylbromine (PFBB) and analysed by gas chromatography (GC) using an Agilent GC/mass spectrometry (MS) ChemStation. Individual analytes were monitored using selective ion monitoring (SIM). Analytes were monitored by peak area and quantified using the isotope dilution method using a deuterated internal standard and a standard curve. Isolated primary hepatocytes from MYC mice fed with MCD or control diet were cultured in complete RPMI for 24 h. Supernatant were harvested and FFAs were identified by GC/MS. Splenocytes from MYC mice were cultured with or without 50 μM C18:2 for 24 h. CD4+ and CD8+ T lymphocytes were sorted and total RNA was extracted using miRNeasy mini kit (Qiagen). Array analysis was performed in the Department of Transfusion Medicine, clinical centre at NIH. Mouse gene 2.0 ST array (Affymetrix) was used and performed according to the manufacturer’s instruction. Data were log-transformed (base 2) for subsequent statistical analysis. The Partek Genomic Suite 6.4 was used for the identification of differentially expressed transcripts. The Ingenuity Pathway Analysis tool (http://www.ingenuity.com) was used for analysis of functional pathways. RNA was extracted from frozen tissues with RNeasyMini Kit (Qiagen). Complementary DNA was synthesized by iScriptcDNA synthesis kit (BioRad). Sequence of primers used for quantitative RT–PCR can be obtained from the authors. The reactions were run in triplicates using iQSYBR green supermix kit (BioRad). The results were normalized to endogenous GAPDH expression levels. CD4+ T lymphocytes were isolated from the spleen of MYC mice by negative autoMACS selection using a CD4+ T lymphocytes isolation kit (Miltenyi Biotec) or flow cytometry cell sorting. Human CD4+ T lymphocytes were prepared from PBMCs by autoMACS using a CD4+ T lymphocytes isolation kit (Miltenyi Biotec). The purity of CD4+ T lymphocytes was above 90% after autoMACS separation and above 95% after flow cytometry cell sorting. C16:0, C18:0, C18:1,and C18:2 were purchased from Sigma. Fatty acids were dissolved in DMEM with 2% fatty-acid-free bovine serum albumin (BSA; Sigma, catalogue number A8806) after solvent was evaporated, then followed by two rounds of vortexing and 30 s of sonication. Isolated CD4+ T lymphocytes or splenocytes were incubated with different fatty acids or conditioned medium from hepatocyte culture for 3 days. Unless specifically described, fatty acids were used at 50 μM concentration. For fatty acid depletion, active charcoal (catalogue number C-170, Fisher) was used as described before36. Briefly, 0.5 g of active charcoal was added into every 10 ml of conditioned medium. Then pH was lowered to 3.0 by addition of 0.2 N HCl. The solution was rotated at 4 °C for 2 h. Charcoal was then removed by centrifugation, and the clarified solution was brought back to pH 7.0 by addition of 0.2 N NaOH. NAC (10 mM), catalase (1,000 U ml−1) or mitoTEMPO (10 μM) was used to inhibit ROS production, mitochondrial ROS levels were determined by mitoSOX staining 24 h after treatment, cell death and apoptosis were measured by annexin V and 7-AAD staining 3 days after treatment. Caspase activity assay was measured by caspase-Glo 3/7 assay kit (Promega) according to the manufacturer’s protocol. Fresh prepared liver-infiltrating mononuclear cells were washed and resuspended in 500 μl of BODIPY 493/503 at 0.5 μg ml−1 in PBS. Cells were stained for 15 min at room temperature. Then cells were subjected to flow cytometry analysis. Two pZIP lentiviral shRNA vectors targeting human CPT1a and a control vector (NT#4) were purchased from TransOMIC Technologies. Lentivirus was packed in 293T cells. Jurkat cells were purchased from the German Collection of Microorganisms and Cell Cultures (DSMZ), and no authentication test was performed by us. Cells were cultured in complete RPMI medium and were tested to be mycoplasma free. Jurkat cells were infected with shRNA lentivirus. Puromycin was added to eliminate non-transduced cells. Doxycycline (100 ng ml−1) was added to induce shRNA and GFP expression for 3 days. Efficiency of shRNAs was confirmed by western blot. Jurkat cells with CPT1a knockdown were treated with 200 μM C18:2 for 24 h. Mitochondrial ROS production and cell survival were measured in GFP+-transduced cells. Fatty acid oxidation was measured according to a previous publication37. 1-14C-C18:2 and 1-14C-C16:0 were purchased from PerkinElmer. Briefly, isolated CD4+ or CD8+ T lymphocytes were pretreated with C18:2 or kept in regular media. After 24 h, cell media was changed to media containing 50 μM cold C18:2 plus 1 μCi 1-14C-C18:2 per ml or 50 μM cold C16:0 plus 1 μCi 1-14C-C16:0 per ml. After 2 h, medium was removed and mixed with concentrated perchloric acid (final concentration 0.3 M) plus BSA (final concentration 2%) to precipitate the radiolabelled fatty acids. Samples were vortexed and centrifuged (10,000g for 10 min). Radioactivity was determined in the supernatant to measure water-soluble β-oxidation products. Mitochondrial membrane potential was measured by TMRM (ImmunoChemistry Technologies) staining according to the manufacturer’s protocol. Briefly, cells were kept in culture medium with 100 nM of TMRM for 20 min in a CO incubator at 37 °C. After washing twice, cells were processed to flow cytometry analysis. Mitochondria-associated superoxide was detected by mitoSOX (Life Technologies) staining according to the manufacturer’s protocol. Briefly, cells were first subjected to surface marker staining. Then cells were stained with 2.5 μM mitoSOX for 30 min in a CO incubator at 37 °C. After washing twice, cells were processed for flow cytometry analysis. OCR was measured using an XFe96 Extracellular Flux Analyzer (Seahorse Bioscience) as previously described38. AutoMACS-sorted mouse CD4+ and CD8+ T lymphocytes were attached to XFe96 cell culture plates using Cell-Tak (BD Bioscience) in RPMI media with 11 mM glucose. Cells were activated with 1:1 CD3:CD28 beads (Miltenyi BioTech) and vehicle or 50 μM C18:2 was added. Twenty-four hours after activation, cells were incubated in serum-free XF Base Media (Seahorse Bioscience) supplemented with 10 mM glucose, 2 mM pyruvate and 2 μM glutamine, pH 7.4, along with 50 μM C18:2 if previously present, for 30 min at 37 °C in a CO -free cell culture incubator before beginning the assay. Five consecutive measurements, each representing the mean of 8 wells, were obtained at baseline and after sequential addition of 1.25 μM oligomycin, 0.25 μM trifluorocarbonylcyanide phenylhydrazone (FCCP), and 1 μM each of rotenone and antimycin A (all drugs from Seahorse Bioscience). OCR values were normalized to cell number as measured by the CyQUANT Cell Proliferation Assay Kit (Life Technologies). Human liver samples were stained as previously described8. For immunostaining, formalin-fixed, paraffin-embedded human liver tissue samples were retrieved from the archives of the Institute of Surgical Pathology, University Hospital Zurich. Fibrosis grade was analysed for NASH according to NAFLD activity score (NAS)39 and for others according to METAVIR score40. The study was approved by the local ethics committee (Kantonale Ethikkommission Zürich, application number KEK-ZH-Nr. 2013-0382). Human PBMCs from healthy donors were obtained on an NIH-approved protocol and prepared as described previously41. Informed consent was obtained from all subjects. The sample sizes for animal studies were guided by a previous study in our laboratory in which the same MYC transgenic mouse stain was used. No animals were excluded. Neither randomization nor blinding were done during the in vivo study. However, mice from the same littermates were evenly distributed into control or treatment groups whenever possible. The sample size for the patient studies was guided by a recent publication also studying NASH-induced HCC, but focused on different aspects8. Statistical analysis was performed with GraphPad Prism 6 (GraphPad Software). Significance of the difference between groups was calculated by Student’s unpaired t-test, one-way or two-way ANOVA (Tukey’s and Bonferroni’s multiple comparison test). Welch’s corrections were used when variances between groups were unequal. P < 0.05 was considered as statistically significant.


Froberg-Fejko K.M.,Bio Serv
Lab animal | Year: 2012

Environmental enrichment can be defined as altering the living environment of captive animals in order to provide them with opportunities to express more of their natural behavioral repertoire. The challenge of providing effective enrichment in laboratory species is to ensure that it allows for normal behavioral opportunities. For many animals, these behaviors include foraging, sheltering, exploring, nest building and gnawing. In the wild, many species use wood and bark to satisfy these behaviors, thereby maintaining physiological and behavioral health. For laboratory animals, various wood enrichment products are available that will provide appropriate environmental enrichment and satisfy those same needs.


Husbandry conditions in a laboratory environment can be barren and monotonous. Improving those conditions by providing opportunities for laboratory mice to engage in species-specific behavior can improve their mental and physical well-being. Giving the animals choices and control over their environment is key to reducing stress. Nesting is a normal behavior of mice, and giving mice nesting opportunities allows them to choose how and where to create their nest and provides them a means of thermoregulation in their microenvironment.


Froberg-Fejko K.M.,Bio Serv | Lecker J.L.,Bio Serv
Lab Animal | Year: 2013

Environmental enrichment can be defined as altering the living environment of captive animals in order to provide them with opportunities to express their natural behavioral repertoire. As important as offering an enriched environment is assuring lab animals are housed in the safest conditions possible. Cage flooding events are an unfortunate reality; however, technology is advancing to minimize these events. Bio-Serv, in collaboration with Allentown, Inc., has developed an innovative and economical shelter called the Safe Harbor Mouse Retreat (Fig. 1). This shelter offers a life-saving refuge for mice during these occasional, but devastating cage-flooding accidents. Mice will not be lost due to chilling or drowning caused by water exposure. Breeding mice can save their litters by moving their pups to the second level, and all mice can escape to the higher level where they can remain warm and dry until they are rescued. This clever shelter is not only life-saving for mice but offers several other significant benefits as well.


Froberg-Fejko K.M.,Bio Serv | Lecker J.,Bio Serv
Lab Animal | Year: 2011

Certain animal and protocol conditions, including genetic alteration, post-operative intervention, dehydration and small weanling size, create a requirement for ‘special needs’ products to ensure animal survival and to encourage reliable research outcomes. Bio-Serv was the first to create a product to meet these special needs. Nutra-Gel is a highly palatable, nutritionally complete food and water gelled diet. © 2011 Nature Publishing Group. All rights reserved.


PubMed | Bio Serv
Type: Journal Article | Journal: Lab animal | Year: 2012

Environmental enrichment can be defined as altering the living environment of captive animals in order to provide them with opportunities to express more of their natural behavioral repertoire. The challenge of providing effective enrichment in laboratory species is to ensure that it allows for normal behavioral opportunities. For many animals, these behaviors include foraging, sheltering, exploring, nest building and gnawing. In the wild, many species use wood and bark to satisfy these behaviors, thereby maintaining physiological and behavioral health. For laboratory animals, various wood enrichment products are available that will provide appropriate environmental enrichment and satisfy those same needs.


PubMed | Bio Serv
Type: Journal Article | Journal: Lab animal | Year: 2015

Idiopathic chronic diarrhea of nonhuman primates is a major gastrointestinal disorder and a leading cause of serious morbidity in nonhuman primates kept in captivity. Many animals are not responsive to traditional treatments. Millions of dollars are spent annually on diagnosis and supportive care of these animals. Probiotics like Bio-Servs PrimiOtic and PrimiOtic Plus can help to reduce the incidence of diarrhea in captive nonhuman primates by supporting the natural microflora in the gut.


PubMed | Bio Serv
Type: Journal Article | Journal: Lab animal | Year: 2013

Environmental enrichment can be defined as altering the living environment of captive animals in order to provide them with opportunities to express their natural behavioral repertoire. As important as offering an enriched environment is assuring lab animals are housed in the safest conditions possible. Cage flooding events are an unfortunate reality; however, technology is advancing to minimize these events. Bio-Serv, in collaboration with Allentown, Inc., has developed an innovative and economical shelter called the Safe Harbor Mouse Retreat (Fig. 1). This shelter offers a life-saving refuge for mice during these occasional, but devastating cage-flooding accidents. Mice will not be lost due to chilling or drowning caused by water exposure. Breeding mice can save their litters by moving their pups to the second level, and all mice can escape to the higher level where they can remain warm and dry until they are rescued. This clever shelter is not only life-saving for mice but offers several other significant benefits as well.


PubMed | Bio Serv
Type: Journal Article | Journal: Lab animal | Year: 2010

Husbandry conditions in a laboratory environment can be barren and monotonous. Improving those conditions by providing opportunities for laboratory mice to engage in species-specific behavior can improve their mental and physical well-being. Giving the animals choices and control over their environment is key to reducing stress. Nesting is a normal behavior of mice, and giving mice nesting opportunities allows them to choose how and where to create their nest and provides them a means of thermoregulation in their microenvironment.

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