Trehalase

A medium containing 0.5 g of casamino acid hydrolysate, 0.5 g of yeast extract, and 0.5 g of peptone per liter was developed to produce trehalose (Difco™ R2A agar, Fisher scientific). The carbon source to produce trehalose in this medium was added as 2% glucose. The culture cell pellets were boiled at 95°C for 20 min to remove the trehalose from the pellets. After centrifuging the resulting mixes, the supernatant was collected. By continuing to boil the supernatant at 95°C until it lost half of its volume, the supernatant containing the extracted trehalose was concentrated. Furthermore, the trehalose extract sample was mixed with 495 mL of 50 mM sodium citrate buffer (pH 6.0) to create a total volume of 1,000 mL for the trehalose assay. Purified trehalase (Megazyme), with an enzyme concentration of 0.0009 UmL⁻¹, was added in a volume of 5 μL to start the reaction. The 3,5-dinitro salicylic acid colorimetric method (DNS method) (Miller et al., 1960 (link)) was used to evaluate the degradation of trehalose to glucose molecules. In this procedure, 1,000 μL of the reacted sample and 300 μL of the DNS solution were mixed and boiled for 5 min. The mixture was allowed to cool at room temperature, and the absorbance was measured at 540 nm using a spectrophotometer. Subsequently, by measuring the absorbance of a standard solution that contained 20 nmol of glucose instead of trehalose, glucose production was ascertained. A control sample devoid of trehalose was used in the measurement to gauge the level of pre-existing glucose in the tested sample. Additionally, a blank sample without trehalose extract was included as a reference.

For trehalose determinations, cells (50 mg wet weight) were collected and frozen in liquid nitrogen. After thaw on ice, samples were resuspended in 0.5 ml 0.25 M Na₂CO₃, incubated at 95°C for 20 minutes and centrifuged at maximal speed. 5 μl 1 N acetic acid and 5 μl buffer T (300 mM sodium acetate, 30 mM CaCl₂ buffer pH 5.5) and 20 μl trehalase (aprox. 10 units) were added to 10 μl of the supernatant. Then, the reaction mix was incubated at 40°C for 2 hours, centrifuged and glucose was determined in the supernatant. The Wiener laboratory kit was used and the manufacturer´s guidelines were followed to quantify glucose.
Glycogen accumulation was detected by the brown colour produced by staining with iodine. Cells were collected and resuspended in a solution of 0.2% iodine/0.4% potassium iodide, incubated 3 min and then spotted onto YPD plates. The darker the color, the higher the amount of glycogen that was intracellularly accumulated [37 (link)].
Ethanol quantification was performed by a coupled reaction of 2 purified enzymes: Pichia pastoris alcohol oxidase (AOX) and radish peroxidase (HRP). Ethanol is a substrate for the AOX enzyme, which in the presence of O₂ generates formaldehyde and H₂O₂. Hydrogen peroxide is a substrate for the HRP enzyme which, together with a colorless substrate, ABTS (2,2’-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid)), generates a colored complex quantifiable by spectrometry [38 ]. To determine the standard curve, 30 μl of known concentrations of ethanol (0 mM, 2 mM, 4 mM, 8 mM and 10 mM) were plated in duplicate in a 96-well plate. On the other hand, serial dilutions were made to the medium of the supernatants obtained from the samples to be analyzed and 30 μl of each dilution were placed in duplicate in the 96-well plate. Then, 120 μl of the reaction mixture (5 mg/ml ABTS, 1.2 U/ml HRP and 0.3 U/ml AOX in 20 mM phosphate buffer pH 7) were placed in each well and the plate was incubated in the dark for 35 minutes at 30°C. To stop the reaction, 100 μl of 1% SDS were added and final product was measured by spectrometry at 412 nm.