Fasting protocol

Mice to be included in ‘fasted’ experimental groups were always collectively housed to minimize stress. Food was removed from the cage hopper and cage floor 18 hours prior to culling. The fast period was always kept constant with respect to the light/dark cycle. Fasting always began three hours prior to the dark period. Water was always available to animals ad libitum. Animals fasted but not culled were not subjected to another fast for at least one week to ensure recovery.

2.4 Physiology

2.4.1 Diet

At 4-6 weeks of age mice were weaned and placed on solid food diets. Mice were given a chow diet (Specialty Feeds, meat-free rat and mouse diet, Western Australia, Australia) containing 3.1 kcal/g, 4.8% fat. In some experiments mice were given a high fat diet (Specialty Feeds, meat-free rat and mouse diet, Western Australia, Australia) containing 4.75 kcal/6 and 42% fat.

2.4.2 Food Intake

Food intake was quantified in multiple studies outlined here. Food intake was always measured post-surgery in order to monitor the health and recovery progress of mice. Food intake measurements were always taken at the same time each day, immediately prior to the onset of the dark period. Mice whose food intake was monitored were always individually housed. To check intake, food from the cage hopper was transferred to the scale, with care taken to look for scraps of food within the cage itself. All food intake measurements were taken on a UME 600 ± 0.1g Z series scale.

2.4.3 Body Weight

Body weight was taken as a measurement throughout multiple studies, and was always taken alongside food intake measurements. Body weight was always measured at the same time each day, immediately prior to the onset of the dark period. Mice subjected to these experiments were always individually housed and all measurements were taken on a UME 600 ± 0.1g Z series scale.

2.5 Electrophysiology

2.5.1 Slice Preparation

Rats and mice were euthanized using terminal inhalation anesthesia (Isoflurane, Baxter Healthcare, NSW, Australia) followed by decapitation. Following decapitation, the brain was swiftly removed from the cranial cavity and transferred into freshly prepared, ice-cold (<4°C), carbogen (95% O2, 5% CO2)

gassed artificial cerebrospinal fluid (aCSF). pH and osmolarity of aCSF was regularly checked and maintained at 7.4 and between 310-315 mOsm.kg-1_,

respectively. For experiments requiring the use of aCSF containing <10 mM glucose, osmolarity was compensated with the addition of D-mannitol at levels equimolar to the glucose levels removed.

Dura, arachnoid and pia matter were removed from the surface of the brain using fine forceps (Fine Science Tools). The cerebellum was removed using a razorblade to provide a flat edge for the remaining brain to be adhered to a tissue plate of a vibratome (Leica VTS 1000S, Leica Microsystems Nussloch, Germany) using cyanoacrylate glue. Coronal slices (200-300 μm) containing the dorsomedial hypothalamus (DMH), arcuate nucleus (ARH), and/or cortex were cut using the vibratome. The vibratome cutting chamber was filled with ice-cold aCSF, and was regularly replaced to keep temperature as constant as possible during slicing. The arcuate nucleus was identified as: the area immediately dorsal to the median eminence and immediately lateral to the third ventricle. The dorsomedial hypothalamus was identified as the area immediately lateral to the dorsal end of the third ventricle, and immediately posterior to the paraventricular nucleus, both in accordance with coordinates determined from a brain atlas (Paxinos, 2008). Once brain slices were cut, they were immediately transferred from the cutting

bath to a 200 ml beaker containing aCSF warmed to 34°C in a water bath where they were incubated for 20 minutes. After incubation the beaker was removed from the water bath and placed on the lab bench to acclimatize to room temperature for at least an hour before the slices were used for recording. Prior to recording slices were transferred to a recording chamber for electrophysiological experiments. The slices were secured to the cover slip-bottomed recording chamber with a custom-made nylon grid, weighted with a platinum arc to prevent any movement of the tissue during recording. The recording chamber was constantly perfused with room temperature, carbogen-bubbled aCSF, and the flow rate of aCSF was maintained between 2-10 ml/min.

2.5.2 Recording Procedure

Brain slices and individual cells were visualized using a Zeiss Axioskop 2 microscope (Carl Zeiss Ltd., Welwyn Garden City, UK) equipped with a Zeiss Axiocam MRc camera (Carl Zeiss Ltd., Welwyn Garden City, UK) connected to a computer running Axiovision 4.1 imaging software (Carl Zeiss Vision). The microscope was fitted with two immersion lenses (20X and 63X) and a 2.5X booster lens mounted in the microscopes rotational magazine allowing for slice visualization at 20X, 50X and 63X and 157.5X. The microscope was also fitted with a fiber optic fluorescent light source (Carl Zeiss Vision) and filter sets 37 and 46 allowing for the visualization neurons expressing eGFP and YFP, respectively.

Whole-cell current-clamp and voltage-clamp recordings were performed using recording electrodes pulled from borosilicate, filamented, thin-walled glass capillaries (GC150TF-10) using a horizontal electrode puller (P1000 or P-97,

Sutter Instrument Co, Novato, CA, USA) and had resistances of 4-9 MΩ when back- filled with intracellular pipette solution.

Electrophysiological recordings were made with an Axon 700B amplifier (Axon Instruments, Foster City, CA, USA) connected to a personal computer running multiclamp, pClamp 9 (Axon Instruments) and Axiovision 4.1 (Carl Zeiss Vision) software. Experimental data was monitored online using Clampex 9 (Axon instruments) and Multiclamp (Axon instruments) software. Data was filtered at 1- 5 KHz (for voltage-clamp experiments), digitized at 2-100 KHz (Digidata 1322A, Axon instruments) and was saved directly to the PC hard drive for analysis using Clampfit 9 (Axon instruments) software.

Resting membrane potential and firing rate was established from traces in Clampfit software (Axon Instruments) and value offsets due to liquid junction potentials were not accounted for. Input resistance was calculated from Ohm’s Law (R=V/I) using an experimental protocol in which square-wave current steps (- 5 to -100 pA, 500 ms to 1000 ms) were injected and the amplitude of the evoked membrane voltage deflections subsequently measured.

All electrophysiological recordings were made at room temperature as it was deemed unrealistic to perform long-term recordings at physiological temperature. It must be noted that this is a caveat of research described here, not least because the hypothalamus contains populations of temperature-sensitive neurons.

In experiments demanding recordings of fluorescent neurons, cell phenotype was confirmed after whole-cell access was attained. This was achieved under fluorescent illumination, observing that the flurophore had been partially drawn-up into the recording electrode. In these experiments, any neuron in which

this was not observed was deemed to not contain the flurophore and was discarded.

Electrotonic coupling between pairs of cortical interneurons was confirmed when the injection of negative current into only one of two simultaneously recorded neurons, evoked a voltage change in both neurons. Neuronal input resistance and coupling coefficients were determined from membrane potential responses to injection of a range of depolarizing and hyperpolarizing current pulses into either neuron. Coupling coefficients were determined as the ratio of the postsynaptic to presynaptic membrane potential change elicited in response to current injection. The junctional conductance (gj) was estimated from the experimentally determined input resistance and coupling coefficients using a model system whereby each neuron was represented by a single compartment connected via a resistor constituting the electrical synapse. Thus gj between two cells was estimated from:

gj1-2 = R1k1-2/((R1R2)-R1k1-2)2)

In which R1 and R2 represent the input resistances of the pre-and

postsynaptic neurons respectively and k1-2 represents the coupling coefficient

2.6 Solutions

2.6.1 aCSF

The composition of aCSF used in this study is shown in table 2.2, below:

1 mM Glucose aCSF (mM) 2 mM Glucose aCSF (mM) 3 mM Glucose aCSF (mM) 5 mM Glucose aCSF (mM) 10 mM Glucose aCSF (mM) NaCL 127 127 127 127 127 KCL 1.9 1.9 1.9 1.9 1.9 KH2PO4 1.2 1.2 1.2 1.2 1.2 NaHCO3 26 26 26 26 26 D-Glucose 1 2 3 5 10 D-Mannitol 9 8 7 5 0 Ascorbic Acid 0.34 0.34 0.34 0.34 0.34 CaCl2 2.4 2.4 2.4 2.4 2.4 MgCl2 1.3 1.3 1.3 1.3 1.3

2.6.2 Pipette solution

Intracellular pipette solution was made daily from stock solutions kept at 4°C except adenosine triphosphate (ATP) which was stored at -20°C. The compositions of pipette solutions used in this study are shown in table 2.3, below:

2mM ATP pipette solution (mM) 4mM ATP pipette solution (mM) K-gluconate 140 140 HEPES 10 10 KCL 10 10 EGTA 1 1 Na2ATP 2 4 Biocytin 5 5

Table 2.3: Pipette solution composition

The pH was adjusted to 7.4 with KOH and osmolarity adjusted to within 10 mOsm.kg-1 _{below that of the aCSF (typically 290 -310) with sucrose.}

2.6.3 Solutions used in thesis chapters

aCSF glucose concentration and pipette solution ATP concentration was tailored to the requirements of the experiment on the day. During tissue preparation, brain slices were maintained in aCSF of identical glucose concentration to that used in experimental control conditions. aCSF flow rate was kept between 2 – 5 ml/min and system volume was around 15 ml meaning that upon drug/ glucose application, the intended concentration of drug/glucose was present in the recording chamber after a maximum of 7.5 minutes. aCSF glucose and pipette solution ATP concentration used in experiments presented in this thesis are shown in table 2.4, below:

Thesis Chapter aCSF Glucose (mM) aCSF Mannitol (mM) Pipette Solution ATP (mM) Chapter 3 1 or 5 9 or 5 2 Chapter 4 1 or 5 9 or 5 2 Chapter 5 3 7 2 Chapter 6 2 8 2 Chapter 7 5 5 2 Chapter 8 10 0 4

Table 2.4: Composition of aCSF and pipette solution