Engineering

Design

Exoelectrogenic bacteria are the most commonly used and studied for the purposes of creating MFC [4]. However, yeast has certain advantages over bacterial cells: yeast’s metabolism is well studied [5]; multiple efficient tools for genetic engineering exist [6]; yeast can utilize many different substrates for its growth that makes them good candidates for efficient waste-water treatment; yeast-based devices are eco-friendly; yeast can grow under low pH [7], high ethanol level [8] and can tolerate osmotic stress [9]. All mentioned above makes yeast attractive organisms for MFC.

However, in wild-type yeast, a lot of the reducing power is consumed internally in fermentation pathways (ethanol, glycerol production) or channeled into the mitochondrial respiratory chain. NADH molecules, as key intracellular electron carriers, can serve as donors of electrons to external circuits in microbial fuel cells (MFCs), enabling the conversion of chemical energy into electrical energy. Increasing the cytosolic NADH pool could therefore enhance electron transfer efficiency and overall MFC performance.

To increase cytosolic NADH pool, several gene-deletion strains of S. cerevisiae were engineered (Figure 2):

• Glycerol-3-phosphate dehydrogenase (GPD1, GPD2) : key enzyme of glycerol synthesis.It reduces dihydroxyacetone phosphate to glycerol-3-phosphate using NADH [10];
• Alcohol dehydrogenase (ADH1, ADH5) : converts acetaldehyde into ethanol, consuming NADH [11];
• NADH dehydrogenase (NDE1, NDE2) : catalyzes the oxidation of cytosolic NADH, providing cytosolic NADH to the mitochondrial respiratory chain [12].

However, expected increase in NADH pool might not result in the enhanced electron transfer from the cells toward anode. In order to improve yeast capacity for electron transfer, we decided to employ another strategy: express cellobiose dehydrogenase (CDH) from fungus Phanerochaete chrysosporium in S. cerevisiae [13]. This enzyme is an extracellular oxidoreductase responsible for cellobiose utilization that can enhance the yeast’s capacity for wastewater treatment and to further facilitate electron transfer to the anode. CDH overexpression broadens the range of substrates that the cell can metabolize. In the context of microbial fuel cells, expressing CDH enables S. cerevisiae to catalyze the oxidation of various organic compounds while directly transferring electrons to the electrode, either through direct electron transfer or via redox mediators. This should significantly improve the efficiency of bioelectricity generation and enhance the degradation of organic pollutants in wastewater [14]. Extracellular expression of CDH might enhance electron transfer toward anode in the presence of cellobiose. In addition, yeast cells displaying fully functional CDH on their surface may eliminate the need for electron transfer mediators such as methylene blue, or at least significantly reduce their required amount, making the entire system more eco-friendly.

Initially, the yeast codon-optimized sequence of WT CDH was designed as synthetic DNA and contained overhangs, compatible with cloning using MoClo yeast toolkit [15]. However, after further literature review, we decided to revise our design because the native CDH activity in S. cerevisiae appears to be very low/undetectable. The new design was based on [16]. In this case, CDH was fused with Aga2 protein for cell surface expression and three mutations were introduced into the CDH coding sequence (D20N, A64T, V592M). The authors claimed that it resulted in much higher activity of the mutated chimeric protein.

Together, all genetic modifications are intended to increase the cytosolic pool of NADH, enhancing the availability of this key electron donor to boost current generation in MFCs. In addition, the surface expression of CDH is expected to increase the yeast cells’ capacity for electron transfer.

Build

In order to use the CRISPR/Cas9 tool for gene deletion, our first objective was to introduce Cas9 plasmid (pCfB2312) into the DOM0090 yeast strain. This was the primary step for the majority of further lab work. We checked colonies after transformation by PCR for the presence of Cas9 plasmid. For further work, strains containing Cas9 plasmid were maintained on the plates with G-418 antibiotic.

gRNA plasmid design

At the initial stage of the project, we planned to generate multiple deletion mutants. To better understand the impact of each target gene on MFC efficiency, single deletion strains Δgpd1, Δgpd2, Δnde1, and Δnde2 were created. To improve the efficiency of gene deletions and facilitate subsequent strain engineering, CRISPR/Cas9 was selected as the primary tool.

We started with the design of gRNA (single guide RNA) and assembly of gRNA plasmids. The purpose of gRNA is to provide the guiding sequence for Cas9 protein so that it can be directed to the target gene site.

gRNA part of the plasmid consists of:

• Promoter – controls gRNA expression.
• gRNA - single RNA molecule consisting of a 20 bp spacer sequence (located adjacent to the Protospacer Adjacent Motif, or PAM, in the genome) and a structural RNA region that activates Cas9 and guides it to the specific genomic site defined by the spacer sequence.
• Terminator – signals gRNA transcription termination [17].

Since initially we wanted to study the effect of single gene deletions, four plasmids containing single gRNAs for GPD1, GPD2, NDE1, NDE2 genes were designed as following:

1. Benchling.com CRISPR/Cas9 design tool was used for screening target genes for the presence of potential gRNA sequences.
2. Two gRNA oligonucleotides were designed and ordered for each of the four target genes (forward and reverse strands, each was 24 bp in length, including cloning overhangs). A list of all designed oligonucleotide pairs used for gRNA plasmid construction is provided in Table 1. The forward and reverse oligos were then phosphorylated and annealed prior to cloning.
3. The annealed oligo pairs were cloned into the pSBME133 backbone using the Golden Gate (GG) cloning and the Eco31I restriction enzyme (Figure 3).
4. Turbo competent E. coli cells were transformed with the GG reaction mixture.
5. Colonies carrying plasmids were selected on LB plates containing ampicillin. An additional color-based selection was used to identify colonies containing gRNA plasmids. During the Golden Gate (GG) assembly, the gRNA sequence replaces the mCherry dropout cassette; therefore, desired colonies appear white, while those carrying the original plasmid without insertion remain pink. Several colonies from each transformation were screened by colony PCR across the gRNA insertion site to confirm the expected amplicon size.
6. Positive colonies carrying desired plasmids were used to grow cultures for miniprep.

We also aimed to investigate the cumulative effect of multiple gene deletions on MFC performance. To this end, we constructed plasmids carrying two gRNAs for simultaneous deletion of two genes. Two sets of multi-gRNA plasmids were designed to target NDE1 + NDE2; GPD1 + GPD2 (Figure 4). For the triple gene deletion (ADH1, ADH2, ADH5) we used ready plasmid that had gRNA for all three ADH genes. Although ADH2 catalyzes the reverse reaction—the oxidation of ethanol to acetaldehyde—it should remain inactive under MFC conditions, as it is glucose-repressible and becomes active only when sugars are depleted. Therefore, including ADH2 in the triple gRNA plasmid (targeting ADH1/2/5) should not compromise our system.

For the multiple gRNA plasmids, the gRNA oligos for each target gene were designed as described above. The only difference was the presence of different overhangs, compatible with other target plasmids. “Intermediate” plasmids each carrying one gRNA expression cassette were first created using pSBME134 and pSBME136 as backbone vectors (pSBME134-GPD1_gRNA, pSBME136-GPD2_gRNA, pSBME134-NDE1_gRNA, and pSBME136-NDE2_gRNA). Assembly was performed using GG with the Eco31I restriction enzyme. For the second GG reaction with Esp3I enzyme, plasmids containing gRNA for NDE1 and NDE1, or GPD1 and GPD2 genes (products of the first assembly) were mixed with pSBME121 (contains assembly connector sequence) and pSBME106 to get double-gRNA plasmid vectors. Transformation was performed in E. coli, followed by selection on ampicillin-containing LB plates.

Moving forward, our focus shifted to designing synthetic donor DNA fragments to repair the DNA break that the Cas9 introduces. These donor DNAs provide templates for homology-directed repair in order to eliminate mutations and preserve genome structural organisation. The synthetic DNA fragments were designed with appropriate homology arms (around 100 bp) and cloned into a plasmid containing AmilCP chromoprotein (was used as a reporter for color selection of the colonies carrying the insert). Finally the donor DNA fragments were PCR-amplified from pAmilCP plasmids and used for transformation of DOM90 + pCfB2312 (Cas9-containing plasmid) yeast strains along with the corresponding gRNA plasmids. The transformed cultures were grown on plates, which contained selection markers for both the Cas9 and gRNA plasmid. To verify the success and efficiency of gene deletions, colony PCR was performed.

To verify successful deletions, we designed primers binding approximately 100 bp upstream and 100 bp downstream of each target locus. In the DOM90 control strain, where no deletion occurred, this setup amplified a fragment corresponding to the intact gene sequence in addition to the 200 bp flanking regions. In contrast, in correctly edited strains, the amplification product was reduced to the ~200 bp flanking sequence only, confirming that the intervening gene had been removed. In addition, for single-gene deletions, several colonies were chosen to perform sequencing and thereby ensure that the donor DNA was correctly inserted in a place of the deleted genes.

For heterologous expression of CDH, synthetic DNA with overhangs compatible with cloning into the plasmids of the MoClo kit was designed [15]. Two versions of CDH plasmids were created (WT CDH version and Aga2-CDHmut) (Figure 5). The rationale behind the use of every variant is explained above in the Design section.

Test

To test the effect of our gene deletion on the growth, glycerol and ethanol production, a time course sample collection experiment was performed. Two colonies from each of the single gene deletion strains along with the DOM90 + Cas9 as a control strain were grown in CSM + 2% glucose. OD600 measurements were performed and samples for HPLC and NADH/NAD⁺ measurement assay were collected at the following time points: 0, 1.5, 3, 4.5, 6.5, 8.5, 10.5, 12, 24, and 48 hours.

Growth experiment and HPLC analysis

OD measurements indicated that most of the engineered strains exhibited impaired growth, with the slowest growth observed in the Δgpd1 and Δgpd2 strains (Figure 6). After 48h, OD600 in Δgpd1_7, Δgpd2_1, and Δgpd2_1 strains was about 3 times lower than in one of the controls. However, the Δnde1 and Δnde2 strains, as well as the Δgpd1_8 mutant, did not show such substantial differences compared to the control.

HPLC analysis of glycerol production over a 48-hour time course demonstrated the impact of our gene deletions. As shown in the graph, both Δgpd1 strains exhibited complete suppression of glycerol accumulation. The Δgpd2 and Δnde2 strains accumulated glycerol more slowly during the initial stages of growth, but by the end of the experiment, their glycerol levels were similar to those of the control strain. In contrast, the Δnde1 mutant showed enhanced glycerol accumulation compared to the control, likely reflecting the need to reduce cytosolic NADH due to impaired mitochondrial uptake—directly indicating the intended effect of the mutation (Figure 7).

However, under our experimental conditions, complete suppression of glycerol biosynthesis had also an adverse effect on growth as it was revealed by both OD measurements and HPLC analysis. Consistent with their lower growth rates, Δgpd1_7, Δgpd2_1, and Δgpd2_1 strains exhibited reduced glucose consumption compared to the DOM0090 control. Glucose was not fully depleted from the culture medium even after 48 hours of growth, with approximately half of the initial glucose amount remaining (Figure 8).

All the Δgpd1_7, Δgpd2_1, and Δgpd2_1 strains also exhibited decreased ethanol production during growth, whereas other deletion mutants showed ethanol synthesis rates similar to the DOM0090 control. However, differences were observed in ethanol consumption after 24 hours, with the DOM0090 strain displaying more intensive ethanol utilization (Figure 9).

Ethanol accumulation in the culture medium was measured throughout the growth of the parental DOM90 strain and deletion strains (Δgpd1, Δgpd2, Δnde1, Δnde2). The graph depicts the fermentative activity of each strain and the impact of specific gene deletions on ethanol production over time.

NADH/NAD⁺ assay

Since the ultimate goal of our gene deletions was to increase the cytosolic NADH pool in yeast cells, we measured NADH/NAD⁺ levels using Amplite® Colorimetric NADH/NAD⁺ Ratio Assay Kit. Samples for the assay were also collected during the growth experiment.

However, the sample measurements did not yield meaningful results. In some cases, the absorbance measured for total NADH + NAD⁺ samples was lower than that of NADH alone (Table 3). We therefore concluded that these results are unreliable and that further optimization of the protocol is required.

Although the NADH/NAD⁺ measurements did not provide any reliable data, we can still conclude that the cytosolic NADH pool in our engineered yeast strains increased, as evidenced by the suppression of glycerol production.

To evaluate the performance of our engineered yeast strains as biocatalysts for electron transfer in MFCs, we designed custom-built hardware and tested multiple yeast strains under these conditions. The results of testing our engineered strains in the MFC hardware showed that the Δgpd1 strain performed slightly better than the DOM90 control as a catalyst for electron transfer.

Hardware

To evaluate the performance of our engineered yeast strains as biocatalysts for electron transfer in MFCs, we designed custom-built hardware and tested multiple yeast strains under these conditions. The results of testing our engineered strains in the MFC hardware showed that the Δgpd1 strain performed slightly better than the DOM90 control as a catalyst for electron transfer. These results confirm that suppressing glycerol biosynthesis following GPD1 deletion increases the cytosolic NADH pool, thereby enhancing electron transfer to the electrodes, and validate our hypothesis that boosting NADH levels is an effective strategy for engineering yeast as a high-performance cell factory. Thus, the measurement results obtained from our hardware directly validate our genetic design by linking the metabolic modifications in yeast to measurable improvements in MFC performance.

Modeling

Microbial fuel cell (MFC) design has traditionally relied on trial-and-error strain screening, lacking a predictive framework to link genetics with power output. Here, we introduce a mechanistic multi-scale model that addresses this gap. Cellular NADH flux models predict electron surplus resulting from specific gene deletions, biofilm kinetics translate single-cell metabolism into collective current dynamics, and reactor optimization maximizes power across biological and electrochemical timescales.

The key innovation is the integration of redox balance constraints at the molecular level with population dynamics and electrochemical processes. By measuring only five strains, our framework can predict the performance of novel genotypes and validate these predictions using bootstrap-based uncertainty quantification. We identified statistically significant electron surplus, demonstrating that the model accurately captures real metabolic constraints. Most critically, the model prevented generation of double mutants Δgpd1Δnde1 and Δgpd2Δnde2, which had been planned initially.

This approach transforms MFCs from an empirical biotechnology into a predictive bioengineering platform, providing a genotype-to-watts pipeline for systematic optimization Modeling.