Tissue collection and analysis

3.8.1 Liver lipid content

The current protocol for lipid extraction is the Folch procedure (Folch et al., 1957). Liver was dissected into finely sliced 3g pieces and placed in a solution containing 50 mL of chloroform (LiChrosolv, Merch, Damstadt, Germany): methanol (LiChrosolv, Merch, Damstadt, Germany) 2:1 ratio, containing approximately 10 mg/L butylated hydroxytoluene (Kosher, SAFC, Sigma-Aldrich, Louise, USA) (antioxidant used for longevity of samples) and were kept in the cool room.

For lipid extraction, the samples were returned back to room temperature. The extraction solution was filtered into a separation funnel equipped with filtration paper. Furthermore, the sample bottles were rinsed with 10 mL of chloroform: methanol (2:1 ratio) twice and were poured through the funnel with the filtration paper into the separation funnel. Then the bottles were rinsed with 10 mL of chloroform once. By rinsing the bottles this permitted maximum collection of the extraction solution. Then the funnel and filter paper was removed when the greatest amount of the extraction solution was in the separation funnel and the filter paper was dry. Next, 16 mL of 0.6 % NaCl (2.4g sodium chloride powder and 400 mL of distilled water H2O) was poured into the separation funnel. The separation funnels were capped, mixed well and left overnight at room temperature to allow the solutions to partition.

Once the upper and lower phase of the solutions was reached, the bottom phase containing the lipid was collected in a round bottomed flask (pre rinsed with chloroform) and the upper phase was discarded. In addition, rotary evaporation (Heidolph VV 2000, waterbath type WB 2000, Schwabach, Germany) at 40 °C, speed 120-150 with warm water of the bottom phase

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occurred. The rotary evaporator rotates the round flask which causes the solvent to evaporate and therefore the lipid is left in the round flask.

Once dry, the lipid was reconstituted with 2.5 mL of chloroform (CHCI3) three times to ensure the maximum amount of lipid extract was obtained. Then the lipid extract was transferred into a 10 mL volumetric flask up to the mark and mixed well with the pipette. Then 1 mL of the lipid extract was transferred into a pre-weighed 4 mL vial, dried off under N2 gas via the nitrogen evaporator (N-EVAP111, Organomation Associates Inc., Berlin, MA, USA) (leaving only the lipid) and left in the desiccator overnight (lidless) to prevent moisture exposure. The 4 mL vials placed in the desiccator were re-weighed to calculate the lipid (g/mL) content. The total lipid as percentage to the liver wet tissue was calculated with the following formula:

Lipid in 1 mL solution x (10 / 3*) x 100

10 = volume of lipid extract. *= mass of tissue sample acquired from dissection.

3.8.2 Histological analysis of liver

Liver tissue was sectioned, weighed and finely diced using a sharp scalpel blade. Liver samples were fixed in 10 % neutral-buffered formalin (NBF). The formalin solution was prepared by mixing 100 mL of formaldehyde (Ajax Finchem Pty Ltd, NSW, Australia), 900 mL of distilled water, 4.0 g of NaH2P04 (Sodium dihydrogen orthophosphate dehydrate, BDH Laboratory Supplies, England) and 6.5 g of Na2HPO4 (Sodium phosphate, Sigma Chemical Co, St Louise, USA) together. Liver tissue was then dehydrated via a series of graded ethanol baths in order to displace the water and was then infiltrated with wax. This was achieved by firstly submerging liver tissue in 70 % ethanol to begin the processing

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schedule using the automated tissue processor for a total of 14 hours. The following paraffin processing cycle was used: 70 % ethanol for 1 hour, 90 % ethanol for 1 hour, 100 % ethanol for 1 hour, 100 % ethanol for 1 hour and 30 minutes, 100 % ethanol for 1 hour and 30 minutes, 50/50 % ethanol/xylene for 1 hour and 30 minutes, xylene for 1 hour and 30 minutes, paraffin wax for 1 hour, paraffin wax for 1 hour, paraffin wax for 1 hour, and paraffin wax for 30 minutes.

Following embedding of tissue into wax blocks (stored at room temperature), microtome sectioning was then undertaken. Firstly, the water bath was set at 47-48 °C with fresh ionized water and the slide warmer was set at 60 °C. The blocks that were sectioned were first placed face down on an ice block for at least 20 minutes to keep the tissue cool. A fresh blade was placed on the microtome after sectioning up to 10 blocks. The rotary handle was locked and the guard was then put over the blade. The alignment of the blade was set at 10. To begin, the paraffin block (cassette) was put on the clamp (specimen holder) with the block facing the blade and aligned in a vertical plane. The microtome was then unlocked. The distance between the paraffin block and the blade was adjusted so that the blade was only touching the block slightly by turning both wheels. The microtome dial was set at 20-30 microns (thickness of trim) in order to plane the block. Once the blade was cutting smoothly, the dial was set at 2 microns. Once the desired section of thickness was produced, the ribbon was transported to the water bath (floating on the surface of the water bath). A microscope slide was put at a 90 °C angle next to the section and then pulled up (the section adhered to the glass slide). The slides were put in a pre-labelled rack and put in the oven (37 °C) to bond the tissue to the glass ready for Haematoxylin and Eosin (H & E) staining.

H & E staining was performed following a standard protocol (Ross and Pawlina, 2011). Sections were cleared in xylene for five (3 and 2 minutes) minutes, then immersed in absolute

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alcohol for two minutes (1 and 1 minute) and rinsed in 70 % alcohol and then rinsed in 30 % alcohol followed by a thorough wash with tap water. Sections were then counterstained with Mayer’s haematoxylin for two minutes, then returned to tap water and then immersed in Scott’s tap water (blueing solution) for one minute and then rinsed in tap water again for one minute. The sections were then examined microscopically. Following this, sections were then stained with Eosin (1 % Sigma Aldrich) for two minutes followed by a brief rinse in tap water. Sections were then immersed in 30 % alcohol (x 10 dips), then 70 % alcohol (x 10 dips) and then absolute alcohol for 30 dips (10 dips and 20 dips) and then cleared with xylene for 3 minutes (1 and 2 minutes) and mounted with DPX mounting media (Thermo Scientific) on coverslips. Liver sections were imaged at 400 X magnification (Carl Zeiss microscope) from two rats in each group for the analysis of liver lipid droplets using Axio Vision 4.8 software.

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