Testing Substrate Specificity in Yeast Fermentation

As Buchner discovered at the turn of the 20th century, the process of fermentation is a multistep, enzyme-catalyzed reaction. Inherent to maximal enzyme action is a defined set of optimal conditions and substrates. Therefore, the enzymes responsible for glycolysis and subsequent fermentation reactions will exhibit optimal reaction rates in an environment that mimics physiological conditions. To demonstrate this point, we will use “active dry” yeast packets from the supermarket (such as Fleischmann’s or Red Star brands). This common product is a freeze dried collection of Saccharomyces cerevisiae, also known as “baker’s yeast.”

Naturally found on ripe fruits, like grapes, as well as on and in the human body, S. cerevisiae is a facultative anaerobe, and its biodiversity and carbon utilization is dictated by the carbon and energy sources available in its specific habitat. This demonstrates the incredible metabolic flexibility housed in these tiny eukaryotes, however, some substrates are more efficiently metabolized than others. In the absence of oxygen, S. cerevisiae will switch on its fermentation pathway as a mechanism to maintain a favorable cellular redox status (fermentation regenerates NAD⁺, which is essential for glycolysis), generating ethanol and carbon dioxide as byproducts. 

To understand how different sugar substrates are utilized by S. Cerevisiae, we can measure the amount of CO₂ produced. If you recall the stoichiometry for fermentation, for every mole of glucose, yeast cells will produce two moles of CO₂, which makes a quantification of sugar metabolism fairly straightforward. While scientists have invented a number of devices to quantifiably measure the rate CO₂ production resulting from fermentation in yeast, these devices are not practical for classroom settings. Here we will use a basic 15ml conical tube (such as Falcon or Corning brands) with gradation markings as a device to measure CO₂ production in response to a variety of carbon sources.

• Prepare sugar stock solutions at 40% w/v (40g per 100ml of H₂O)
• Prepare dilute sugar solutions for each sample tested
  • For a recommended starting concentration of 0.5%, v/v, you’ll need 0.5 ml stock in 10 ml final volume
• Heat water baths and/or hot plates to 40 °C
• Warm all solutions to be used in the 40 °C bath
• Prepare a gas-collection setup (see below)
• Immediately before the experiment, prepare the 7% yeast solution in H₂O

Setup

Set up your tubes and water bath in such a way that the gas produced in each fermentation will rise to the tops of the tubes and volumes of gas can be marked with a permanent marker or waxed pencil.

In the example at right, plastic tubes were used with a few holes poked into each plastic cap using a needle. The tubes were filled with yeast solution and food source (e.g. sucrose), covered, and inverted quickly into the water bath. The samples should be held inside the tubes and displaced into the water bath as gas is produced.

A similar setup can be used with test tubes and stoppers with holes (cover with your finger when inverting) or using a glass slide to invert each tube into the bath before sliding off the slide and allowing the fermentation gas produced to displace the yeast solution. You may also be able to use a typical gas collection apparatus from a chemistry laboratory at your school.

Finally, volumes of gas can be determined by emptying a marked tube and filling it with water exactly to the mark. Then that water can be poured into a graduated cylinder of appropriate size to determine the volume of gas produced.