In Vivo Monitoring of Transcriptional Activity During Metabolic Transition Using a Bioluminescent Reporter in Yeast

Summary

This protocol employs a bioluminescent reporter, allowing measurements of transcriptional activity in Saccharomyces eubayanus to monitor the glucose-to-maltose transition, enabling real-time analysis of metabolic adaptations and supporting strain optimization for industrial fermentation under diverse conditions.

Abstract

Sequential sugar consumption, from a preferred sugar source to a less preferred one, represents a critical metabolic adaptation in yeast, which is particularly relevant for survival in fluctuating environments such as those found in beer fermentation. However, sugar transitions are an environmental variable that is challenging to predict and detect, impacting the outcome of beer fermentations. This protocol describes an in vivo system to monitor transcriptional activation associated with the glucose-to-maltose metabolic shift in Saccharomyces eubayanus that applies to different wild Saccharomyces yeast strains.

The system employs an episomal bioluminescent transcriptional reporter for maltose metabolism, focusing on MAL32, since it provides a good readout for metabolic shifts, as studied in S. cerevisiae. For this, yeast strains were transformed with plasmids containing the MAL32 regulatory region from S. eubayanus, controlling the expression of a gene encoding for a destabilized version of firefly luciferase¹, and a hygromycin resistance gene used exclusively during transformation to ensure plasmid acquisition. Following selection, transformed yeast cells can be cultured under non-selective conditions, as the episomal plasmid remains stable in culture conditions for up to 7 days.

This system was validated under a complex sugar environment in microfermentation assays, confirming the effectiveness of the luciferase reporter in informing metabolic transitions. Samples were collected regularly and analyzed with a luminometer, providing continuous insights into yeast responses. While broadly applicable, this protocol is particularly valuable for assessing yeast performance under fermentation conditions, where metabolic changes pose a significant challenge. Additionally, this methodology can be adapted by selecting alternative promoters to explore a broader range of responses to environmental changes, allowing characterization as well as optimization of wild yeast strains for diverse industrial applications.

Introduction

Microorganisms such as yeasts must constantly adapt to dynamic environmental conditions to maintain fitness and survive¹. These adaptations often involve complex gene regulatory circuits integrating multiple extracellular signals to orchestrate precise metabolic responses^2,3. In industrial settings, the efficiency of these metabolic transitions is critical, particularly in fermentation processes where disruptions can lead to suboptimal yields or incomplete fermentations³. A key metabolic challenge to overcome is when cells transition from a preferred to a secondary carbon source, such as the glucose-to-maltose shift. This process introduces a lag phase during which genes required for the metabolism of the secondary carbon sources are derepressed, enabling growth resumption^4,5.

In brewing, Saccharomyces yeasts must efficiently transition from glucose to maltose metabolism. In particular, S. eubayanus, the cold-tolerant parental species of lager yeasts, displays substantial phenotypic variability in its ability to adapt to such transitions⁶. Wild isolates, such as those from Patagonia, often exhibit prolonged lag phases and slower maltose consumption compared to domesticated strains, which have been selected for their optimized fermentative capacities^7,8. While domesticated strains have adapted to ferment mixed sugar environments efficiently, wild strains often display a slower metabolic transition, potentially due to stronger glucose repression and variable regulation of the MAL locus^6,9.

This study utilizes the natural variability of S. eubayanus as a model to investigate metabolic adaptations under glucose-to-maltose conditions, leveraging an episomal destabilized luciferase reporter to monitor gene expression in vivo, by tracking luminescence¹. The selected reporter MAL32 encodes a maltase protein, a pivotal enzyme for maltose catabolism during glucose-to-maltose transitions^10,11. Remarkably, the MAL32 promoter represents a successful marker for assessing maltose metabolism induction after glucose depletion¹². By incorporating this reporter system, we aimed to elucidate strain-specific adaptive mechanisms and identify potential targets for optimizing fermentation performance. Furthermore, this protocol can be expanded beyond brewing, offering applications in biotechnology and environmental studies where complex sugar environments play a significant role. Understanding the genetic and regulatory determinants of fluctuant environment responses in S. eubayanus enhances our knowledge of yeast physiology, supporting the development of robust strains for diverse industrial and research applications.

Protocol

1. Construction of episomal reporters

NOTE: We selected a reporter regulatory region based on yeast literature to construct the episomal plasmid for monitoring maltose consumption^6,11,12. The promoter of the candidate reporter gene was defined as the regulatory sequence immediately upstream from the candidate ORF up to the nucleotide flanking the adjacent upstream ORF. This region was amplified from the genomic DNA of S. eubayanus CBS12357^T reference strain¹⁰. This approach ensures the high-fidelity construction of episomal plasmids suitable for downstream applications, including studying other interesting conditions in yeast.

1. Design the DNA fragments and overlapping primers to include 30 nucleotide pairing sequences between fragments for a seamless assembly via yeast recombinational cloning¹³ (Figure 1). See Supplemental Table S1 for primer sequences.
2. To avoid mutations, amplify the promoter region (see above), as well as the destabilized luciferase (LUC-PEST) and hphMX cassettes¹ using a high-fidelity DNA polymerase (refer to the polymerase manufacturer for protocol details). Set the thermocycler to 98 °C for 10 min for initial denaturation, followed by 30 cycles of 98 °C denaturation, 60 °C annealing, and 72 °C extension; complete the protocol with a final extension at 72 °C for 10 min.
   NOTE: When using yeast shuttle plasmids as templates for PCR, it is recommended to subject amplicon reactions to a DpnI digestion (see the enzyme manufacturer's protocol) to eliminate any traces of template plasmid that may interfere with the yeast transformation.
3. Digest the pRS426 plasmid¹⁴ backbone with EcoRI and XhoI restriction enzymes to linearize the vector, cutting it at the multiple cloning site, and prepare it for recombination (refer to the enzyme manufacturer's protocol).
4. Mix the linearized pRS426 backbone with the amplified DNA fragments and transform the mixture into Saccharomyces cerevisiae BY4741 using yeast recombinational cloning^13,15 to allow the yeast's homologous recombination machinery to assemble the plasmid construct in vivo.
5. Plate the transformed yeast on a synthetic complete (SC) medium lacking uracil (SC-URA) to select for auxotrophic transformants containing the novel constructed vector pRS426- MAL32-LUC-hphMX plasmid.
6. Extract plasmids from yeast using a yeast mini-prep kit according to the manufacturer's instructions and transform the extracted plasmid into Escherichia coli DH5α to amplify the construct for further analyses.
7. Screen bacterial colonies via PCR to confirm the correct assembly of the constructs.
8. Purify the plasmid DNA using a standard plasmid purification kit and perform Sanger sequencing to verify the integrity and sequence of the assembled plasmids.

2. Transformation of yeast strains

NOTE: The yeast transformation protocol was adapted from a previously established method for S. eubayanus¹⁶ and applied successfully to other Saccharomyces species and fermentative yeast strains. This protocol was derived from the traditional yeast transformation method from the Gietz Lab¹⁵. This approach enables efficient plasmid integration and selection under diverse experimental conditions, offering a robust and flexible method for transforming different yeast strains. It ensures reliable selection and maintenance of the episomal plasmid under hygromycin pressure.

1. Preculture a single colony of freshly grown yeast in 5 mL of YPD (1% w/v yeast extract, 2% w/v peptone, 2% w/v glucose) at 20 °C at 200 rpm of agitation overnight.
2. Dilute the culture to an OD₆₂₀ of 0.2 in fresh YPD medium and incubate at 20 °C with agitation at 200 rpm until mid-log phase (OD₆₂₀ 0.4-0.6). Depending on the yeast strain, this typically requires 3-4 h.
3. Harvest the cells by centrifugation at 21,380 × g for 1 min at room temperature, discard the supernatant, and wash the pellet 3x with sterile water.
4. Resuspend the washed cells in 100 µL of 0.1 M lithium acetate, centrifuge as described in step 2.3, and repeat the process 2x.
5. Add the following components to the cells in this order: 240 µL of 50% w/v PEG-4000, 35 µL of 1 M LiAc, 5 µL of plasmid DNA (100 ng/µL), 20 µL of single-stranded carrier DNA (ssDNA) previously heated at 98 °C for 10 min. Mix gently to homogenize the transformation mixture.
6. Incubate the mixture at 20 °C for 30 min, followed by heat shock at 34 °C for 55 min. After heat shock, add 38 µL of 100% ethanol to a final concentration of almost 0.1% and incubate for an additional 5 min at 34 °C.
   NOTE: The temperatures used in this protocol are optimized for Saccharomyces eubayanus, a cryotolerant strain isolated from temperate forests.
   The use of 100% ethanol is required to induce cellular stress, enhancing the transformation protocol's efficiency.
7. Add 600 µL of YPD to the transformed cells, centrifuge as in step 2.3, and discard the supernatant.
8. Resuspend the cell pellet in 600 µL of fresh YPD and incubate the suspension without agitation overnight at 4 °C to allow recovery.
9. Plate the transformed cells on YPD agar supplemented with 200 µg/mL hygromycin and incubate at 20 °C-30 °C for 48-72 h.
10. Select 10-12 colonies from the transformation plates and re-streak them onto YPD agar containing 200 µg/mL hygromycin to ensure the presence of the plasmid and generate a mixed patch of the selected colonies for further use.

3. Validation of luminescence

NOTE: To validate the functionality of the luminescent reporters, transformed strains were tested under conditions designed to induce differential expression of the luciferase reporter. This stepwise validation allows for the assessment of reporter functionality under fluctuating sugar conditions, leveraging the robustness of luminescent assays to capture real-time metabolic responses.

1. Pick a sample from a transformed yeast strain patch and inoculate it into YP medium (1% w/v yeast extract, 2% w/v peptone) containing 5% w/v glucose and 200 µM hygromycin. Incubate at 25 °C without shaking for 24 h. Refresh the cultures in fresh medium at a 1:10 dilution and incubate under the same conditions for another 24 h.
2. Wash the cells with YP medium without sugar and inoculate them into the test media at a 1:10 dilution. The testing media is supplemented with luciferin to a final concentration of 3 mM and contains 200 µM hygromycin to maintain plasmid stability.
3. For mixed-sugar growth testing, use a glucose-maltose matrix with the following concentrations (% w/v): glucose (0%, 0.5%, 1%, 2%) and maltose (0%, 2%, 5%, 10%, 20%, 30%) in YP medium. Each condition is supplemented with luciferin (3 mM final concentration) and 200 µM hygromycin. Dispense 200 µL of culture into each well of a 96-well plate (Figure 2).
4. Measure luminescence and OD₆₂₀ every 30 min for 72 h using a luminometer plate reader, without attenuation and 1 s of integration time, ensuring continuous reporter activity and cell density monitoring.

4. Fermentation sampling and luminescence monitoring

NOTE: Transformed strains were subjected to controlled micro-fermentation conditions to evaluate luminescence activation during fermentation. This enables the comparison of luminescence activation across different fermentation conditions, providing insights into yeast metabolic responses during extended fermentation periods.

1. Prepare malt extract medium (12 °Plato) by dissolving malt extract in water and sterilize at 100 °C for 20 min. Allow the medium to cool to room temperature before inoculation.
2. Preculture transformed yeast strains in 5 mL of YPD (1% w/v yeast extract, 2% w/v peptone, 2% w/v glucose) at 20 °C with agitation at 200 rpm for 24 h. Transfer the preculture to 50 mL of malt extract medium. Incubate the culture at 20 °C with agitation at 200 rpm for 24 h.
3. Harvest the cells by centrifugation at 21,380 × g for 1 min at room temperature and resuspend them to prepare triplicate 50 mL micro fermentations in 12 °Plato malt extract medium⁹. To enhance fermentation performance, supplement the medium with 0.3 mg/L ZnCl₂.
4. To ensure consistent cell densities across replicates, calculate the inoculum volume using the formula:
   Inoculum volume = (1.5 × 10⁶) × Volume (mL) × Degrees Plato
5. Inoculate the prepared medium and incubate the cultures at 20 °C without shaking for 14 days. Monitor the fermentation progress by recording CO₂ loss daily. Weigh the vessels simultaneously each day to measure cumulative weight loss as an indicator of CO₂ production.
6. For luminescence monitoring, periodically sample 200 µL from each microfermentation. Add luciferin to the samples to achieve a final concentration of 3 mM and measure luminescence and OD₆₂₀ using a luminometer plate reader without attenuation and 1 s of integration time. Perform measurements at defined intervals throughout the fermentation period (Figure 3).

Results

The following results demonstrate the usability of the newly constructed luminescent reporter to monitor the glucose-to-maltose transition in yeast cells in a fermentative process. The reporter plasmids are initially assembled using yeast recombinational cloning¹³ to generate episomal reporter constructs. This process requires nucleotide sequence overlapping of at least 30 nucleotides between the different amplicons, all depicted in Figure 1. The regulatory regions co...

Discussion

This study demonstrates the effectiveness of an episomal bioluminescent reporter for monitoring transcriptional activation in S. eubayanus under metabolic transitions. By employing MAL32 as a transcriptional reporter¹¹, we could track key metabolic transitions in real time, providing a robust framework for understanding strain-specific adaptations. This reporter, selected for their role in maltose metabolism, offers distinct advantages in evaluating metabolic flexibility in yeast...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Ampicillin, sodium salt	ThermoFisher Scientific	11593027
D-Glucose	Sigma-Aldrich	G8270
DpnI	New England Biolabs	R0176S
EcoRI	New England Biolabs	R0101S
Hygromycine B	Gold Biotechnology	H-270-1
L-Luciferine	Gold Biotechnology	L-127-10
Maltose monohydrate	Sigma-Aldrich	47288
Phusion Plus PCR Master Mix	ThermoFisher Scientific	F631S
Tecan Infinite 200 PRO M	Tecan
Wizard Plus SV Minipreps DNA Purirfication System	Promega	A1330
XhoI	New England Biolabs	R0146S
Zymoprep Yeast Plasmid Miniprep I	Zymo Research	D2001