Metabolism is mainly coordinated by cellular energy availability and environmental conditions. Assays for knowing how cells adapt energetic metabolism to different nutritional and environmental conditions give valuable information to elucidate molecular mechanisms. Oxidative phosphorylation is the primary source of ATP in most cells, and mitochondrial respiration activity is a key component of oxidative phosphorylation, maintaining mitochondrial membrane potential for ATP synthesis. Mitochondrial respiration is often studied in isolated mitochondria that are missing the cellular context. Here, we present a method for quantifying mitochondrial respiration in yeast-intact cells. This method applies to any yeast species, although it has been generally used for Saccharomyces cerevisiae cells. First, the yeast growth in specific conditions is tested. Then, cells are washed and resuspended in deionized water with a 1:1 ratio (w/v). Cells are then placed in an oximeter chamber with constant stirring, and a Clark electrode is used to quantify oxygen consumption. Some molecules are sequentially placed into the chamber and selected according to this effect on the electron transport chain or ATP synthesis. ATPase inhibitor oligomycin is first added to measure respiration coupled to ATP synthesis. Afterward, an uncoupler is used to measure the maximal respiratory capacity. Finally, a mix of electron transport chain inhibitors is added to discard oxygen consumption unrelated to mitochondrial respiration. Data are analyzed using a linear regression to obtain the slope, representing the oxygen consumption rate. The advantage of this method is that it is specific for yeast mitochondrial respiration, maintaining the cellular context. It is essential to highlight that inhibitors used in mitochondrial respiration quantification could vary between yeast species.