Compartmentalized Sesquiterpenoid Biosynthesis and Functionalization in the Chlamydomonas reinhardtii Plastid

Abstract

Terpenoids play key roles in cellular metabolism and can have specialized functions. Their heterologous production in microbial hosts offers an alternative to natural extraction. Here, we developed a subcellular engineering approach in the model green alga Chlamydomonas reinhardtii by targeting both sesquiterpenoid synthases and cytochrome P450s (CYPs) to the plastid, exploiting its photosynthetic electron transport chain to drive CYP‐mediated oxidation without reductase partners. Nuclear‐encoded sesquiterpenoid synthases were expressed with farnesyl pyrophosphate synthase fusions and targeted to the plastid, while CYPs were modified for soluble localization in the plastid stroma by removing transmembrane domains. The plastid environment supported hydroxylation, epoxidation, and oxidation reactions, with functionalization efficiencies reaching 80 % of accumulated products. Carbon source availability influenced product ratios, revealing metabolic flexibility in the engineered pathways. Overall sesquiterpenoid yields ranged between 250–2500 μg L⁻¹ under screening conditions, establishing proof‐of‐concept for using plastid biochemistry in complex terpenoid biosynthesis. Living two‐phase terpenoid extractions with different perfluorinated solvents revealed variable performances based on sesquiterpenoid functionalization and solvent type. This work demonstrates that photosynthetic electron transport can drive CYP‐mediated functionalization in engineered subcellular compartments. However, improvements in photobioreactor cultivation concepts will be required to facilitate the use of algal chassis for scaled production.

Photosynthetic microalgae were engineered for compartmentalized sesquiterpenoid biosynthesis by targeting synthases and cytochrome P450s to the chloroplast. The plastid's electron transport chain drives P450‐mediated oxidation without reductase partners, enabling production of complex functionalized terpenes. This metabolic engineering approach demonstrates how photosynthetic biochemistry can be harnessed for sustainable production of valuable compounds from CO₂.

Introduction

Terpenoids are one of the most diverse classes of natural organic compounds, playing crucial roles in biological processes across all domains of life.[ ¹ , ² ] These molecules function in photoprotection, photosynthesis, electron transport, defense, and signaling.[ ¹ , ³ , ⁴ ] Terpenoids have wide‐ranging applications in medicine, flavoring and fragrances.[ ⁴ , ⁵ ] However, their structural complexity poses challenges for chemical synthesis.[ ² , ⁶ ] Current specialty terpenoid production relies on extraction from plant sources, resulting in low yields and impurities.[ ³ , ⁶ ] Natural sources of some specialty terpenoids face increasing pressure due to slow growth rates of their native hosts[ ² , ³ ] and sustainability concerns, driving the need for alternative production strategies. ^[4] Microbial production of terpenoid biosimilars through metabolic engineering is an alternative to source these valuable chemicals. Photosynthetic microalgae provide potential advantages through their native capacity for high flux to terpenoid pigments and potential sustainable cultivation using carbon dioxide as a carbon source.[ ⁶ , ⁷ , ⁸ ]

Terpenoid biosynthesis begins with the formation of five‐carbon (C₅) isoprene units: isopentenyl pyrophosphate (IPP, C₅) and its isomer dimethylallyl pyrophosphate (DMAPP, C₅). ^[2] These units are generated through the mevalonate (MVA) pathway or the 2‐C‐methyl‐D‐erythritol 4‐phosphate (MEP) pathway.[ ¹ , ² ] Prenyltransferases catalyze the sequential addition of IPP and DMAPP to produce linear precursors, including geranyl pyrophosphate (GPP, C₁₀), farnesyl pyrophosphate (FPP, C₁₅), and geranylgeranyl pyrophosphate (GGPP, C₂₀). ^[2] Terpenoid synthases (TPS) convert these precursors into cyclic chemical skeletons through complex reactions. ^[6] This process yields various terpenoid structures: monoterpenoids (C₁₀), sesquiterpenoids (C₁₅), diterpenoids (C₂₀), triterpenoids (C₃₀), and tetraterpenoids (C₄₀). ^[9] The resulting skeletons often undergo further modification through functionalization catalyzed by enzymes like acetyltransferases, carboxylases, and cytochrome P450 (CYP) monooxygenases.[ ² , ⁸ ] CYPs are found overexpressed in tissues and cell types where terpenoids accumulate in higher organisms. ^[10]

Here, we investigated an engineering approach exploiting the biochemical environment of the model green alga Chlamydomonas reinhardtii plastid for complex terpenoid synthesis. We leverage advances in transgene design for robust expression of heterologous enzymes from the alga's nuclear genome, investigate subcellular precursor availability in the cytoplasm and plastid, and by localizing both sesquiterpene synthases and CYPs in the plastid, we leverage its photosynthetic electrons to drive CYP‐mediated oxidation without requiring cytochrome P450 reductases (CPRs). This strategy enables the production of sesquiterpenoids (STP, C₁₅) with application in fragrance, where minimal chemical modifications to the terpenoid backbone can expand scent profiles. We also evaluated carbon source effects on product profiles and examined extraction efficiency using various biocompatible solvents. This work demonstrates a proof‐of‐concept for using the algal plastid to facilitate complex terpenoid biosynthesis.

Results and Discussion

Sesquiterpenoid Production from Terpene Synthases Located in the Cytoplasm or Chloroplast

Engineering compartmentalized sesquiterpenoid biosynthesis in C. reinhardtii required probing the alga's capacity for either cytoplasmic or plastid‐targeted heterologous terpenoid production.[ ⁷ , ¹¹ , ¹² , ¹³ , ¹⁴ , ¹⁵ , ¹⁶ ] C. reinhardtii produces all terpenoid precursors for photosynthesis and other cellular processes using only the MEP pathway in its plastid. ^[6] Recent advances in transgene design have improved the expression of nuclear‐encoded transgenes, facilitating metabolic engineering efforts for non‐native terpene production and elucidating the metabolic flexibility of the photosynthetic cell.[ ¹⁶ , ¹⁷ ] Although C. reinhardtii lacks endogenous sesquiterpenoid (STP) synthases, it can be engineered to produce diverse STPs by redirecting carbon flux from cytosolic farnesyl pyrophosphate (FPP).[ ¹² , ¹⁴ , ¹⁵ , ¹⁷ , ¹⁸ , ¹⁹ , ²⁰ ] In contrast, FPP levels in the plastid are natively low but can be increased to produce heterologous STPs by localizing overexpressed native or heterologous FPP synthases (FPPS) to this subcellular compartment.[ ¹² , ¹⁵ , ¹⁸ , ²⁰ ]

We compared the yields of terpenoids produced from 10 sesquiterpenoid synthases expressed from the algal nuclear genome and localized to either the cytoplasm or the plastid. Plastid‐targeted constructs were designed with C‐terminal FPPS fusions, as previous studies have shown that free FPP levels in the plastid are minimal and FPPS fusion to sesquiterpenoid synthases does not enhance productivity in the cytoplasm.[ ¹⁴ , ²⁰ ] The cell line employed features a constitutive knockdown of the native squalene synthase, which competes directly for cytoplasmic FPP with the introduced sesquiterpenoid synthases. ^[12] This modification allowed us to compare yield variations under best‐case production scenarios between the cytoplasmic and plastid‐localized sesquiterpenoid synthases fused to FPPS. We evaluated three different construct designs (Figure 1A–C, Supplementary information, Table S1, File S1, and Figures S1–S3): cytoplasmic targeted sesquiterpenoid synthases alone and two different plastid‐targeted FPPS fusions to sesquiterpenoid synthases, either the from Saccharomyces cerevisiae (Erg20) or Escherichia coli (ispA). Multiple STPs were successfully produced, including aristolochene, valencene, selinene, santalene, bisabolol, τ‐cadinol, α‐cadinene, β‐cadinene, γ‐cadinene, murolene, α‐guaiene, β‐guaiene, δ‐guaiene, α‐humulene, alloaromadendrene, valerianol, and patchoulol in both compartments, with yields ranging from ~250 to 2500 μg L⁻¹ (Figure 1D, Supplementary information, Files S2–S4). In the cytoplasmic compartment, highest yields were achieved for patchoulol (2004±432 μg L⁻¹), guaiene (1707±296 μg L⁻¹), valerianol (1563±83 μg L⁻¹), cadinol (1199±234 μg L⁻¹), and bisabolol (1172±130 μg L⁻¹)(Figure 1E, Supplementary information, Table S2, and Figure S5). The yields of sesquiterpenoid synthases were comparable for each STPs product from both cytoplasm (sesquiterpenoid synthase alone) and plastid‐targeted (sesquiterpenoid synthases ‐FPPS fusion) containing transformants (Figure 1E, Supplementary information, Table S2, and Figure S5). Cytoplasmic bisabolol and cadinol synthases achieved higher levels of production than their plastid‐localized counterparts, though yields were comparable when considering variability (two‐way ANOVA, F(6, 2)=158, p<0.05). Bisabolol production reached 1172±130 μg L⁻¹ in the cytoplasm (construct A07) and 1024±187 μg L⁻¹ in the plastid (construct B07). Cadinol synthesis yielded 1199±234 μg L⁻¹ in the cytoplasm (construct A08) and 977±305 μg L⁻¹ in the plastid (construct B08) (Figure 1E, Supplementary information, Table S2, and Figure S5).

Figure 1.

Genetic constructs and sesquiterpenoid production in engineered C. reinhardtii strains. (A‐C) Schematic representation of genetic constructs for sesquiterpenoid production in C. reinhardtii. Constructs include sesquiterpene synthases (STPS) fused to fluorescent protein reporters (FP) and farnesyl pyrophosphate synthases (FPPS) targeted to: (A) cytoplasm, (B) plastid with Erg20 FPPS (S. cerevisiae), and (C) plastid with ispA FPPS (E. coli). Promoters: H‐ß (heat‐shock protein/beta‐tubulin), pPsaD (photosystem I subunit II promoter), H (heat‐shock protein 70S promoter), R (RuBisCO small subunit 2 promoter). CTP: chloroplast transit peptide (PsaD). RBCS intron 1 (i1) and intron 2 (i2) are spread throughout the coding sequences of optimized genes. FDX: ferredoxin 1 terminator. Inset: Engineered C. reinhardtii strain with chloroplast (brown), indicating modified carotenoid synthesis; letters indicate intended localization of recombinant enzyme products. (D) Representative GC‐MS chromatograms of sesquiterpenoid products from each STPS expression, compared to a parental strain negative control extract. Black dots indicate intended sesquiterpenoid products. (E) Sesquiterpenoid yields (μg/L culture) for each construct. Data represent means ± SD from four independent transformants, each analyzed in biological triplicate (n=12). Error bars show SD. Open circles indicate individual data points. Constructs are labeled as cytoplasmic (A, blue), plastid‐Erg20 (B, yellow), and plastid‐ispA (C, red). Genetic constructs and GC‐MS/FID data can be found in Supplementary information Tables S1–S3 and Files S1–S4.

The choice of FPPS influenced production efficiency in an unpredictable manner for each sesquiterpenoid synthase. We found that ispA fusion constructs (C02‐C11) yielded comparable or slightly higher titers for some sesquiterpenoids compared to their Erg20 counterparts (B02‐B11), though differences were not consistent across all constructs (Figure 1E; Supplementary information, Table S2, Figure S3, and S5). For instance, ispA fusions showed higher production for guaiene (C09: 1294±186 μg L⁻¹ vs. B09: 893±149 μg L⁻¹) and valerianol (C10: 954±354 μg L⁻¹ vs B10: 587±218 μg L⁻¹). However, other sesquiterpenoids including aristolochene (C02: 196±208 μg L⁻¹ vs. B02: 165±91 μg L⁻¹), valencene (C03: 315±160 μg L⁻¹ vs B03: 215±120 μg L⁻¹), and selinene (C04: 290±125 μg L⁻¹ vs B04: 253±100 μg L⁻¹) showed no significant differences between ispA and Erg20 fusions due to high variability in transformant populations (Supplementary Table S3). These results suggest that selecting the appropriate FPPS can enhance the production of specific STPs; however, iterative empirical testing for each target sesquiterpenoid synthase‐FPPS fusion is required.[ ²¹ , ²² ] Next, we investigated whether the reducing environment of the plastid could mediate the chemical functionalization of heterologous STP products by co‐expressing CYPs specific to these compounds.

Functionalization of Heterologous Sesquiterpenoids Mediated by Co‐Expression of Plastid‐targeted P450s

The plastid environment provides unique opportunities for terpenoid functionalization through its native redox machinery. In native hosts, terpenoids undergo chemical modifications that enhance their complexity and biological activities.[ ¹⁴ , ²³ ] Metabolic engineering can recapitulate these reactions by expressing the corresponding metabolic pathway in a foreign host.[ ² , ⁴ , ⁹ , ²⁴ , ²⁵ ] CYPs commonly catalyze reactions that add hydroxyl or other functional groups to terpenoids, receiving electrons from CPRs typically on the cytoplasmic side of the endoplasmic reticulum (ER) membrane.[ ¹⁰ , ¹⁹ , ²⁶ ] CYPs and CPRs contain transmembrane (TM) anchors and have been successfully expressed and shown to function in non‐native hosts such as yeasts and tobacco.[ ¹⁰ , ¹¹ , ²⁷ ] Recent studies have demonstrated that when CYPs are expressed and localized in cyanobacteria or plant plastids, photosynthesis‐derived electrons can replace CPRs in driving CYP reactions.[ ²⁷ , ²⁸ , ²⁹ , ³⁰ , ³¹ ]

As we produced reasonable amounts of STPs in the C. reinhardtii plastid by introducing STPS‐FPPS fusions (ranging from 165±91 to 1562±187 μg L⁻¹) (Figure 1E, Supplementary information, Table S2, and Figures S2–S3), we aimed to use its redox potential for sesquiterpenoid chemical functionalization.[ ¹⁰ , ¹¹ , ¹⁹ ] For each STP studied, we co‐expressed and targeted CYPs to the algal plastid that had predicted or were previously shown to be responsible for mediating their functionalization (Supplementary information, Table S1, File S1, Figures S2 and S4). Transformants confirmed to express both sesquiterpenoid synthase‐FPPS fusion and heterologous CYPs were cultivated with solvent overlays, and the products were analyzed by GC‐MS/FID. Specific changes in chromatographic profiles for strains co‐expressing CYPs versus those with sesquiterpenoid synthases alone were observed as shifts in retention times and alterations in mass spectra (Figure 2A, Supplementary information, Table S3, and Files S5–S6). These changes indicate the formation of STPs derivatives, confirming the modification of base STPs in the plastid through CYP co‐expression. For example, strains expressing santalene synthase (B06) with CYP06 to CYP09 showed new peaks consistent with santalol and bergamotol, while valencene synthase (B03) with CYP04 and CYP05 produced a peak matching nootkatone (Figure 2A, Supplementary information, Table S3, and Files S5–S6). Functionalization efficacy varied among CYPs, with some producing the expected product for a specific STP, while others generated numerous side products or inefficiently synthesized target compounds. (Figure 2A and B, Supplementary information, Tables S7–S8, and Files S5–S6). However, it is important to note that some newly formed compounds had low probability matches against the NIST library (Supplementary information, Tables S7–S8, and Files S5–S6). These changes nonetheless provide evidence for successful CYP‐mediated modifications of STPs in the plastid. As no CPR was co‐expressed in these strains, the results indicate that electrons present in the plastid can drive these targeted chemical modifications of heterologous terpenoids.[ ¹⁰ , ¹¹ , ¹⁴ , ¹⁹ , ²⁵ , ²⁷ ] The exact electron donor is unknown, but it could be ferredoxin or simply the reducing environment of the plastid itself in illuminated conditions.

Figure 2.

Plastid‐targeted sesquiterpenoid biosynthesis and functionalization by cytochrome P450 enzymes in C. reinhardtii. Sesquiterpene synthases (STPS) and cytochrome P450 enzymes (CYPs) were targeted to the algal chloroplast. Each STPS construct included C‐terminal ScErg20 FPPS, while CYPs were co‐expressed through plastid targeting and without transmembrane domains. (A) Representative GC‐MS chromatograms of dodecane extracts from C. reinhardtii strains expressing plastid localized STPS and CYP combinations. Numbers indicate specific compounds identified by MS. Dotted lines mark retention times of identified compounds (B) Quantitative analysis of sesquiterpenoid production based on GC‐FID peak areas data. Circles: sum of peak areas for sesquiterpenoids; triangles: sum of peak areas for modified sesquiterpenoids. Data represent means ± SD from four independent transformants, each analyzed in biological triplicate (n=12). Error bars show SD. (C) Functionalization efficiency (%) of each CYP, calculated as the fraction of functionalized sesquiterpenoids relative to total sesquiterpenoids. Bars represent means from four independent transformants, with individual data points shown as open circles. Error bars show standard deviation (n=4). Compounds: [1] Aristolene, [2] Aristolochene, [3] Aristolochone, [4] Valencene, [5] Nootkatone, [6] α‐Santalene, [7] Bergamotol, [8] Santalol, [9] α‐Cadinene, [10] δ‐Cadinene, [11] β‐Cadinene, [12] Muurolene, [13] Muurolol, [14] τ‐Cadinol, [15] α‐Guaiene, [16] β‐Guaiene, [17] α‐Humulene, [18] δ‐Guaiene, [19] Alloaromadendrene, [20] α‐Guaiol, [21] β‐Guaiol, [22] Globulol, [23] Rotundone, [24] Alloaromadendrene oxide. GC‐MS/FID data in Supplementary information File S5, Tables S3, and Figures S6–S7.

For each sesquiterpenoid synthase‐CYP combination, we identified one CYP which generated the target functionalized STP as predicted (Figure 2B and C, Supplementary information, Tables S4, and S7–S8). Functionalization efficiency, calculated as the total sum of new‐STPs relative to the original STPs abundance, varied among each (Figure 2C, Supplementary information, Figure S7, Tables S3, and S5). We observed the following conversions: (1) Aristolochene to aristolochone by CYP02 (UniProt: W6QP06) at 45±21 % efficiency. (2) Valencene to nootkatone by CYP04 (UniProt: E1B2Z9) at 62±18 % efficiency. (3) Santalene to bergamotol and santalol by CYP09 (UniProt: VR5EU4) at 55±13 % efficiency. (4) α‐Cadinene, β‐cadinene, γ‐cadinene, and muurolene to muurolol and α‐cadinol by CYP10 (UniProt: A0 A0 N9H930) at 51±8 % efficiency. (5) α‐guaiene, β‐guaiene, δ‐guaiene, α‐humulene, and alloaromadendrene to α‐guaiol, β‐guaiol, globulol, rotundone, and alloaromadendrene oxide by CYP12 (UniProt: E3 W9 C4) at 66±19 % efficiency.

Selecting appropriate CYPs to ensure product formation remains challenging, and our screening suggests that empirical testing is necessary to determine the correct combination (Figure 3).[ ¹⁰ , ¹² , ¹⁹ , ²⁷ ] Each CYP also led to the formation of off‐target products (Figure 3, Supplementary information, Tables S3–S4, and S7–S8). A heatmap of the major identifiable products illustrates the chemical diversity obtained with each combination of CYP and sesquiterpenoid synthases (Figure 3, 4, Supplementary information, Figure S6, and Table S7–S8). Functionalization efficiency consistently remained below 80 % (Figure 2C, Supplementary information, Table S4, and Figure S7), indicating room for improvement, which may be attained in the future through the formation of metabolons or artificial STPS‐CYP associations via enzyme engineering strategies.[ ¹⁰ , ¹¹ , ²⁴ , ²⁵ , ²⁸ , ²⁹ , ³⁰ , ³¹ ]

Figure 3.

Relative abundance of sesquiterpenoids produced in C. reinhardtii with different cytochrome P450s (CYPs). Heat map showing the relative abundance of sesquiterpenoid compounds based on GC‐FID response. Columns represent sesquiterpene synthase and CYP combinations. Color intensity indicates relative abundance from 0 % (blue) to 80 % (red) of GC‐FID peak area response, normalized to the highest peak area detected for each compound across all samples. Data represent mean values from four independent transformants (n=4), each analyzed in biological triplicate. Statistical significance was determined by one‐way ANOVA followed by Tukey's post‐hoc test (p<0.05). Sesquiterpenoid compounds identified by MS are listed on the right with corresponding numbers [01–24]. Detailed GC‐MS/FID data and statistical analyses are provided in Supplementary information File S5, Table S3, and Figure S6.

Figure 4.

Sesquiterpenoid compounds produced from farnesyl pyrophosphate (FPP) from the C. reinhardtii plastid. Radial schematic showing sesquiterpenoid diversity. The central grey circle represents FPP, the universal precursor. Colored sections illustrate distinct sesquiterpenoid classes: aristolochene‐type (pink, [01‐03]), valencene‐type (orange, [04‐05]), santalene‐type (purple, [06‐08]), cadinene‐type (blue, [09‐14]), and guaiene‐type (green, [15‐24]). Within each class, darker shading indicates non‐functionalized compounds while lighter shading represents oxidized derivatives. Compounds are numbered [01‐24]: Aristolene [01], Aristolochene [02], Aristolochone [03], Valencene [04], Nootkatone [05], α‐Santalene [06], Bergamotol [07], Santalol [08], α‐Cadinene [09], δ‐Cadinene [10], β‐Cadinene [11], Muurolene [12], Muurolol [13], τ‐Cadinol [14], α‐Guaiene [15], β‐Guaiene [16], α‐Humulene [17], δ‐Guaiene [18], Alloaromadendrene [19], α‐Guaiol [20], β‐Guaiol [21], Globulol [22], Rotundone [23], and Alloaromadendrene oxide [24]. All structures were confirmed by GC‐MS analysis (match factor >80 %) through comparison with authentic standards or mass spectral database matching. Detailed analytical data available in Supplementary information File S5, Tables S7–S8.

Carbon Source Effects on Plastid Sesquiterpenoid Biosynthesis and Functionalization

Chlamydomonas can grow on organic acetic acid, inorganic CO₂, or both as carbon sources. The trophic mode of cultivation induces major rearrangements in cell architecture, prompting us to investigate the effects of these changes on the product profiles of our engineered strains. ^[32] For a detailed analysis of carbon source effects, we selected the five sesquiterpenoid synthases and CYP combinations that demonstrated the highest activities in our initial screening (Figure 2B and C). These pairs – B02+CYP02, B03+CYP05, B06+CYP09, B08+CYP10, and B09+CYP12 – showed functionalization efficiencies ranging from 45–66 % and produced clearly identifiable products (Figure 2C, Supplementary information, Table S4). We employed two‐phase cultivation (also known as ′solvent milking′), where cells grow in the presence of a biocompatible organic solvent that continuously extracts produced compounds while maintaining cell viability. This approach allows in situ extraction of STPs as they are produced, preventing potential product toxicity and enabling continuous production. We analyzed products from strains grown under three illuminated conditions: CO₂ alone, acetate alone, or combined CO₂+acetate (Figure 5). Chromatograms revealed distinct STPs and derivative profiles under these cultivation modes (Figure 5, Supplementary information, File S6). Relative abundance data showed variations in sesquiterpenoid production and functionalization efficiency depending on the carbon source (Figure 5, Supplementary information, File S6).

Figure 5.

Carbon source effects on plastid‐targeted sesquiterpenoid biosynthesis and functionalization in C. reinhardtii. GC‐MS/FID analysis of dodecane overlay samples from C. reinhardtii strains expressing different STPSs and CYPs under three carbon source conditions: CO₂, acetate, or CO₂+acetate. Left panels show chromatograms with black dots indicating non‐functionalized sesquiterpenoids and black triangles marking functionalized derivatives. Right panels show relative abundance (%) with error bars representing SD from four independent transformants, each analyzed in biological triplicate (n=12, p<0.05 by one‐way ANOVA with Tukey's post‐hoc test). (A) Aristolochene synthase (B02) + CYP02, (B) Valencene synthase (B03) + CYP05, (C) Santalene synthase (B06) + CYP09, (D) Cadinol synthase (B08) + CYP10, and (E) Guaiene synthase (B09) + CYP12. Bars show relative abundance of non‐functionalized (circles) and functionalized (triangles) products. Chromatogram peaks are normalized to the highest peak area detected across all conditions for each strain. Complete GC‐MS/FID data and statistical analyses available in Supplementary information File S6, Tables S6–S8.

Aristolochene synthase (B02)+CYP02 (Figure 5A, Supplementary information, File S6) exhibited three major peaks across all conditions. CO₂ alone and CO₂+acetate conditions yielded higher relative abundances of the functionalized product than acetate alone. Valencene synthase +CYP05 (Figure 5B, Supplementary information, File S6) primarily produced nootkatone, with two minor peaks under CO₂ and CO2+acetate conditions. Nootkatone abundance was highest with CO₂ alone, followed by CO₂+acetate, and lowest with acetate alone.

Santalene synthase +CYP09 (Figure 5C, Supplementary information, File S6) produced multiple peaks, representing α/β‐santol, bergamotol, and precursors. CO₂ alone yielded the highest relative abundance of functionalized products, while acetate alone showed the lowest. Cadinol synthase +CYP10 (Figure 5D, Supplementary information, File S6) showed multiple peaks across all conditions. Acetate and CO₂+acetate conditions produced higher relative abundances of functionalized products compared to CO₂ alone. The most diverse compound array was generated by guaiene synthase +CYP12 (Figure 5E, Supplementary information, File S6). CO₂ and CO₂+acetate conditions displayed greater peak diversity than acetate alone, with CO₂+acetate showing the highest relative abundance of functionalized products.

These results indicate that a mixed carbon source strategy enhances sesquiterpenoid production and functionalization in C. reinhardtii. Each condition was tested with four independent transformants analyzed in biological triplicate (n=12). Product distributions showed consistent patterns with relative standard deviations below 15 % for major peaks. (Supplementary information, File S6). The combination of CO₂+acetate generally resulted in higher relative abundances of functionalized products in growth conditions tested, likely due to higher cell densities (Figure 5, Supplementary information, File S6).[ ⁸ , ¹¹ , ²⁵ , ²⁶ , ²⁷ , ³¹ ] This metabolic flexibility suggests complex regulation of precursor availability and enzyme activity under different growth conditions. The variation in product profiles adds complexity to predicting functionalized STP outputs from engineered strains. Whether product profiles can be consistently tailored during scaled cultivations is still unknown. Chlamydomonas is not routinely cultivated phototrophically at scale, and research is ongoing to test methods of scaled extraction of engineered terpenoids through solvent ‘milking’.[ ³³ , ³⁴ ] Future studies should address these limitations through production scale‐up and long‐term cultivation experiments with variable light regimes to assess production feasibility.[ ⁶ , ¹⁴ , ³⁵ ]

Culture‐Solvent Extraction Efficiencies

Extracting non‐native STPs from C. reinhardtii requires the culture to grow in contact with a biocompatible solvent.[ ⁵ , ²¹ , ²³ , ²⁶ , ³¹ , ³³ , ³⁴ , ³⁶ , ³⁷ , ³⁸ ] While current yields remain in the mg/L range, limiting immediate industrial application, optimizing extraction methods is crucial for future scale‐up efforts. The choice of extraction solvent has environmental and economic implications for bioprocess designs. ^[39] While dodecane is a standard biocompatible solvent for lab‐scale terpenoid extraction and quantification, perfluorinated solvents offer advantages in safety, stability, and reusability.[ ¹² , ³³ , ³⁴ , ³⁷ , ³⁸ , ⁴⁰ , ⁴¹ ] We recently reported a method to use perfluorinated solvents to extract algal‐produced terpenoids. ^[12] This method allowed the subsequent transfer of terpenes from perfluorinated solvent to ethanol through liquid‐liquid separation, enabling the recycling of the clean perfluorinated solvent to algal culture and direct use of the ethanol‐terpene mixture for fragrance applications.

Here, we evaluated a larger pool of perfluorinated solvents for their capacity to accumulate heterologous and chemically complex sesquiterpenoids produced by C. reinhardtii compared to dodecane (Figure 6, Supplementary information, Table S5). Using dodecane extraction as a reference point (set to 100 %), we found that terpenoid accumulation was lower in all fluorinated solvents, with extraction efficiencies varying across solvents and sesquiterpenoid compounds (Figure 6, Supplementary information, Table S5). This variability is likely due to individual differences in solvent and sesquiterpenoid properties, as each solvent is unique, and the sesquiterpenoids have structural differences.[ ¹² , ²¹ , ³³ , ³⁴ ] FC‐40 and FC‐770 demonstrated higher extraction capacities for bisabolol (32 % and 31 % compared to dodecane, respectively), while CFL7160 extracted aristolochene (15 %) and selinene (16 %) more effectively (Figure 6, Supplementary information, Table S5).[ ¹² , ¹³ , ¹⁴ , ¹⁸ , ³³ , ³⁴ ] We observed enhanced accumulation of ‐OH group–containing STPs in all perfluorinated solvents tested (Figure 6, Supplementary information, Table S5), with bisabolol, candinol, valerianol, and patchoulol accumulating at higher levels than aristolochene, valencene, selinene, vetispiradiene, and santalene.

Figure 6.

Sesquiterpenoid milking efficiencies using different perfluorinated solvents on engineered C. reinhardtii. Heat map comparing relative extraction capacities of fluorinated solvents to dodecane (set as 100 %) for various sesquiterpenoid compounds. Solvents tested: FC‐40, FC‐770, FC‐3284 (perfluoro‐n‐dibutylmethylamine), CFL7160 (perfluoro nonyl trifluoroethyl ether), CXFL‐3288 (perfluorotripropylamine), CXFL‐68 (perfluorotributylamine), CFL3000 A (hexafluoropropene trimer), FC‐43, FC‐72 (perfluoro‐n‐dibutylmethylamine), and FC‐3283 (perfluorotripropylamine). Color intensity indicates extraction capacity (blue: 0 %, red: 40 %). Error bars represent SD from three biological replicates. GC‐MS/FID data in Supplementary information Table S5.

GC‐MS/FID analysis of extracted compounds showed high product purity (>90 %) for most STPs in both dodecane and fluorinated solvent extracts. The main impurities consisted of minor enzymatic side products rather than cellular contaminants, suggesting that current extraction methods provide relatively clean fractionation of target compounds. These results indicate that solvent choice will need to be tailored to individual products if scaled processes for algal‐produced terpenes are to be implemented. Our screening was conducted in small volume well‐plates, in which the contact surface area of perfluorinated solvents is reduced by forming liquid beads under the aqueous phase. In scaled cultivation, the surface area for solvent can be improved, although chemical partitioning variability is likely to persist. Current yields across our engineered strains range from 250–2500 μg L⁻¹, depending on the specific STP product. While these yields enable detailed analytical characterization and demonstrate proof‐of‐concept for our engineering approach, they remain significantly below the g L⁻¹ threshold typically required for industrial implementation. Major improvements in both strain engineering and cultivation strategies will be needed to achieve commercially viable production levels. Scaling these extraction methods for industrial applications presents challenges for which photobioreactors have not yet been optimized.[ ¹² , ³³ , ³⁶ , ³⁸ ] To utilize engineered algae for heterologous terpene production, novel reactor designs that enable scalable culture‐solvent interaction while maintaining optimal light‐driven algae growth will be necessary.

Conclusions

This study demonstrates the feasibility of producing a diversity of chemically complex functionalized terpene products from the engineered algal cell. We show that functionalization of heterologous terpenoids can be achieved using soluble CYPs targeted to the plastid without the need for partner CPRs. While our investigation focused on STPs, this strategy could be applied to higher‐value diterpene products as precursors for pharmaceutical production. The algal system offers a potential platform for rapid investigation of CYP activity, given the affordability of gene synthesis, quick generation of transformants within weeks, and straightforward analysis of terpenoid products. However, challenges and uncertainties remain in whether this can be scaled as a valuable production chassis for complex terpenoid compounds.

Experimental Section

Algae Cultivation, Plasmid Design, Transformation, and Screening

Experiments used a C. reinhardtii strain derived from UPN22, modified for enhanced terpenoid biosynthesis through squalene synthase knockdown and β‐carotene ketolase overexpression.[ ¹² , ¹⁵ ] Cultures were maintained in TAPhi‐NO₃ medium under LED illumination (150 μmol m⁻² s⁻¹). We selected ten sesquiterpene synthases (STPSs) based on documented activity and complete sequence availability, including selinene synthase (UniProt: O64404) and vetispiradiene synthase (NCBI: QBB02271.1, UniProt: A0 A411G8 M5). ^[12] All genes underwent codon optimization and intron spreading for nuclear genome expression.[ ¹⁸ , ¹⁹ , ²⁰ , ⁴² , ⁴³ , ⁴⁴ ] We developed three pOpt3‐based STPS construct designs: cytoplasmic expression with paromomycin selection (APHVIII), and two chloroplast‐targeted expression with hygromycin selection (APHVII).[ ¹⁹ , ²⁰ ] Chloroplast‐targeted constructs utilized the PsaD promoter and chloroplast targeting peptide (CTP), fused to mKOk fluorescent protein and either S. cerevisiae (Erg20) or E. coli (ispA) farnesyl diphosphate synthase (FPPS). FPPS sequences included C‐terminal stop codons to preserve activity.[ ¹⁴ , ¹⁸ ] CYPs were selected based on three criteria: (1) previous biochemical characterization, (2) documented activity on target sesquiterpenoid scaffolds, and (3) availability of complete sequence information. We modified CYPs for plastid targeting by removing transmembrane domains, identified through TMHMM‐2.0 server analysis and AlphaFold structural modeling. ^[46] The modified sequences were subcloned into pOpt3‐based constructs containing the PsaD promoter, CTP, mTFP1 reporter, and zeocin resistance marker (shBle). ^[45] All constructs were synthesized by Genscript (Piscataway, NJ, USA) (Supplementary information, Table S1, File S1).

For transformation, we linearized plasmid DNA using XbaI and KpnI restriction enzymes and introduced 10 μg DNA per transformation via glass‐bead protocol. ^[47] Following 8‐hour recovery in liquid TAPhi‐NO₃ medium ^[15] under low light, cells were plated on selective media containing appropriate antibiotics: spectinomycin (200 μg mL⁻¹), paromomycin (10 μg mL⁻¹), hygromycin B (15 μg mL⁻¹), or zeocin (15 μg mL⁻¹). After 7 days under continuous illumination, we employed a PIXL colony‐picking robot (Singer Instruments, UK) to transfer up to 384 colonies onto fresh TAPhi‐NO₃ plates. A ROTOR robot (Singer Instruments, UK) duplicated colonies after 3 days onto screening plates containing amido black (150 μg mL⁻¹).[ ¹² , ¹⁵ ] We selected four independent transformants per construct based on fluorescent protein signal intensity (Supplementary information, Figure S1–S4). Each transformant underwent analysis in biological triplicate (n=12) to account for expression variability due to random nuclear integration. Selected colonies were cultured in 12‐well plates containing 2 mL TAPhi‐NO₃ medium at 160 rpm for subsequent two‐phase cultivation and solvent analysis.[ ⁵ , ¹² , ¹⁵ , ³³ , ³⁴ ]

Capture and Analysis of Algal‐produced Sesquiterpenoids by GC‐MS/FID

We employed a two‐phase cultivation system for sesquiterpenoid extraction and quantification.[ ⁵ , ¹² , ³³ ] For each construct, four independent transformants showing strong fluorescence signals were analyzed in biological triplicate (n=12 total samples). Cultures were grown in 6‐well microtiter plates containing 4.5 mL TAPhi‐NO₃ medium with a 500 μL dodecane overlay (10 % total volume) for 7 days.[ ¹² , ³³ ] To systematically evaluate extraction efficiency, we compared ten perfluorocarbon solvents (FCs) against dodecane: CFL7160, CXFL‐68, CFL3000 A, CXFL‐3288, FC‐770, FC‐3284, FC‐43, FC‐72, FC‐40, and FC‐3283 (sourced from Sigma‐Aldrich, Germany; Acros Organics, Belgium; Hunan Chemfish Pharmaceutical Co., Ltd, China). FC extractions used 1000 μL solvent (20 % total volume) with 4 mL cultures, with dodecane forming an upper phase and FCs forming lower phases (Supplementary information, Table S5). For these extractions, we used 1000 μL of solvent (20 % of total volume) with 4 mL cultures. Dodecane formed an upper ′overlay’ while FCs formed ′underlays′. We monitored culture volumes throughout cultivation to account for evaporation effects. Phases were separated by centrifugation (3500×g, 5 min), and both solvent fractions were collected for GC analysis. Cell density was determined using flow cytometry (Supplementary information, File S7). ^[33]

GC‐MS/FID analysis followed established protocols[ ¹² , ¹³ , ³³ ] and processed chromatograms using MassHunter software (Agilent, Germany, version B.08.00). Compound identification involved three complementary approaches: (1) Comparison of mass spectra against the NIST Mass Spectral Library (National Institute of Standards and Technology, USA); (2) Analysis of retention indices; (3) Matching against authenticated standards. For absolute quantification, we generated calibration curves (1–500 μM) using purified standards in both dodecane and FCs: δ‐guaiene, patchoulol, α‐santalene, valerianol, α‐bisabolol, valencene, and cedrene (Toronto Research Chemicals, Canada) (Supplementary information, Figure S5). Compound identification reliability was evaluated using three metrics: (1) mass spectral match factor (“P%” in Supplementary information, Tables S7 and S8), calculated as the mathematical similarity between sample and reference spectra on a scale of 0–100; (2) retention index comparison with literature values; and (3) comparison against a standard terpene mixture containing 98 compounds at 1 mM in methanol (MetaSci, Canada) (Figure 6, Supplementary information, Table S6). Mass spectral matches below 50 % were considered tentative identifications. For functionalized products without available standards, identification relied on the presence of expected mass shifts corresponding to specific chemical modifications (e. g., +16 m/z for hydroxylation) and fragmentation patterns characteristic of the predicted structures. These identifications are in the data tables (Supplementary information, Tables S7–S9).

Data Analysis

The experimental design included four independent transformants per construct, each analyzed in biological triplicate (n=12 total samples per construct). We included two types of controls: the parental non‐transformed C. reinhardtii strain and vector‐only constructs with matched fluorescent reporters. GC‐MS/FID measurements followed a rigorous quality control protocol including: (1) Manual review of all chromatograms for peak quality and integration accuracy; (2) Triplicate technical measurements of each biological sample; (3) Monitoring of internal standards for instrument performance; (4) Regular blank injections to detect potential carryover. Compound identification utilized three criteria: (1) Retention index comparison with reference standards; (2) Mass spectral match factors (reported as P% in Supplementary information, Tables S7 and S8); (3) NIST library comparison with match quality threshold >50 %. For quantitative analysis, we calculated means and standard deviations of sesquiterpenoid yields from biological replicates. Due to random nuclear transgene integration in C. reinhardtii, expression levels can vary substantially between transformants.[ ¹² , ¹⁴ , ¹⁹ ] Our multi‐transformant approach captures this inherent variability, providing a realistic assessment of production potential. High standard deviations in yield data reflect this biological variation rather than technical measurement uncertainty. Statistical analysis employed two‐way ANOVA with Tukey's post‐hoc test to evaluate differences between subcellular localization strategies (Supplementary information, Table S3). We used JMP v.16 (SAS Institute, NC) and R v.3.6.2 (R Foundation for Statistical Computing, Austria) for statistical computations. Data visualization utilized JMP v.16 and GraphPad Prism v.10.3 (GraphPad Software, USA). Figures were prepared using Affinity Designer v.2.5.3 (Serif Ltd., UK) for diagrams, ChemDraw v.20.1 (PerkinElmer, MA, USA) for chemical structures, and Affinity Publisher v.2.5.3 (Serif Ltd., WB, UK) for layout integration.

Author Contributions

SG, SO, GBW, and KJL conceptualized and planned the project. SG and KJL designed genes and plasmids for this study. SG undertook cloning, transformation, screening, in situ extraction experiments and analyzed the results. SG and SO interpreted the GC–MS/FID data. SG, SO, GWB, and KJL analyzed data and wrote the manuscript. KJL supervised the project and secured funding. All authors reviewed and discussed the manuscript.

Conflict of Interests

The authors declare no conflict of interest.

Supporting information

As a service to our authors and readers, this journal provides supporting information supplied by the authors. Such materials are peer reviewed and may be re‐organized for online delivery, but are not copy‐edited or typeset. Technical support issues arising from supporting information (other than missing files) should be addressed to the authors.

Supporting Information

Gutiérrez S., Overmans S., Wellman G. B., Lauersen K. J., ChemBioChem 2025, 26, e202400902. 10.1002/cbic.202400902

Data Availability Statement

All experimental information and data supporting the findings of this study are included in the Supplementary information. Source data and genetic files are available in DRYAD (https://doi.org/10.5061/dryad.zgmsbccmz).