Synthetic Biology of Sclareol: From the Plant Biosynthetic Pathway to Engineered Microbial and Photosynthetic Chassis

ABSTRACT

Sclareol is a highly valued bicyclic diterpene widely used as a precursor for the fragrance ambroxide and has also attracted interest as a bioactive natural product. Traditional production relies heavily on plant extraction, primarily from Salvia sclarea, an approach constrained by environmental variability, long cultivation cycles, and costly downstream purification. To overcome these supply bottlenecks, synthetic biology‐enabled biomanufacturing has emerged as a sustainable and scalable alternative. Recent advances in synthetic biology and metabolic engineering have enabled the elucidation and reconstruction of the sclareol biosynthetic pathway in heterologous hosts. This review summarizes current knowledge of sclareol biosynthesis, highlights representative engineering strategies for its production in microbial cell factories, while briefly introducing emerging photosynthetic and plant‐based platforms as complementary green production systems. Finally, we outline the current challenges and future perspectives for the industrial biomanufacturing of sclareol and other high‐value terpenoids.

Graphical Abstract and Lay Summary

Sclareol is a valuable diterpene precursor for ambroxide with diverse bioactivities. Traditional plant extraction suffers from low efficiency and high cost. Synthetic biology enables sustainable biomanufacturing. This review summarizes sclareol biosynthesis, engineering strategies in microbial cell factories, and emerging photosynthetic and plant‐based platforms. Key challenges and future perspectives for industrial production of sclareol and related terpenoids are also discussed.

Abbreviations

BTS1

geranylgeranyl pyrophosphate synthase

DMAPP

dimethylallyl pyrophosphate

ERG20^F96C

farnesyl pyrophosphate synthase variant

ERG9

squalene synthase

FPP

farnesyl diphosphate

GGPP

geranylgeranyl diphosphate

GGPPS

geranylgeranyl diphosphate synthase

GPP

geranyl diphosphate

HMGR

hydroxymethylglutaryl‐CoA reductase

IPP

isopentenyl pyrophosphate

LDPP

labda‐13‐en‐8‐ol diphosphate

SsLPPS

Salvia sclarea labda‐13‐en‐8‐ol diphosphate synthase

SsSS

Salvia sclarea sclareol synthase

tHMG1

truncated HMG‐CoA reductase 1

1. Introduction

Sclareol, a labdane‐type diterpene diol, is a high‐value natural product with extensive applications in the fragrance, pharmaceutical, and food industries owing to its amber‐like aroma and diverse biological activities [1]. As an important precursor for the synthesis of the valuable fragrance compound ambroxide, sclareol holds significant economic importance [2, 3]. In addition, its pharmacological properties, such as antitumor, anti‐inflammatory, and antibacterial activities, highlight its potential as a lead molecule in drug discovery and development [4, 5, 6, 7, 8]. Traditionally, sclareol has been obtained by extraction from plants such as S. sclarea (clary sage). However, this approach is hindered by constraints including limited plant resources, long cultivation cycles, low natural abundance, and challenging purification processes [9]. These limitations result in unstable supply chains and high production costs, restricting the broader industrial application of sclareol.

Recent progress in synthetic biology and metabolic engineering has paved new avenues for sustainable sclareol production through biomanufacturing. Constructing microbial cell factories offers distinct advantages, including shorter production cycles, controllable yields, and environmentally friendly processes, making them an attractive alternative to plant extraction. Crucially, the progression of sclareol biomanufacturing is increasingly driven by a paradigm shift from isolated pathway modifications to multi‐layered, interdependent systems engineering. Rather than simply introducing heterologous enzymes, current efforts focus on holistically addressing specific metabolic and physiological bottlenecks—spanning from maximizing precursor supply and blocking competing pathways, to alleviating product toxicity and extending cellular lifespan. These integrated advances across diverse chassis, ranging from heterotrophic microorganisms like Saccharomyces cerevisiae and Escherichia coli to microalgae and higher plants, are accelerating the shift from traditional plant‐based extraction to scalable biomanufacturing.

This review first introduces the physicochemical properties and industrial value of sclareol, then systematically summarizes the elucidation of its native biosynthetic pathway, which serves as the foundation for heterologous reconstruction. Special emphasis is placed on recent metabolic engineering strategies that enhance sclareol biosynthesis in microbial cell factories, followed by an overview of emerging photosynthetic and plant‐based production platforms. Finally, we discuss the remaining challenges and future directions in the field. Through this comprehensive overview, we aim to provide insights that may guide the sustainable biomanufacturing of sclareol and related high‐value terpenoids.

2. Sources and Applications of Sclareol

Sclareol (C₂₀H₃₆O₂, CAS: 515‐03‐7), chemically named labd‐14‐ene‐8α,13β‐diol, is a labdane‐type diterpene diol primarily found in plants such as clary sage. In 1928, Volmar first isolated this compound from clary sage [10]. Subsequently, researchers discovered its presence in various plants, including tobacco (Nicotiana spp.) [11], acacia (Acacia sp.) [12], Palestine sage (Salvia palaestina) [13], and candy leaf (Stevia monardaefolia) [14]. In nature, sclareol is most abundant in the flower calyces of clary sage and the leaves of sticky tobacco (N. glutinosa) [15]. Leveraging its characteristic physicochemical properties and diverse biological activities, sclareol has attracted considerable interest for applications in the fragrance, pharmaceutical, agricultural, and related industries.

2.1. Application in Flavors and Fragrances

In the fragrance industry, sclareol serves as the core precursor for synthesizing the valuable fragrance ambroxide [16]. Due to the scarcity of natural ambergris and associated ecological concerns [2], finding high‐quality substitutes has long been an urgent industrial demand. In 1950, Firmenich first reported that sclareol could be converted into ambroxide through reactions like oxidation and cyclization [17]. Classical multistep chemical synthesis routes from sclareol to ambroxide are well established. These routes proceed via intermediates such as sclareolide and ambradiol. However, they typically involve expensive or hazardous reagents and generate considerable waste. This has motivated the development of greener and more concise processes. Recent work has demonstrated a one‐pot synthesis of ambroxide from sclareol using hydrogen peroxide and a quaternary ammonium phosphomolybdate catalyst, providing a more environmentally benign alternative to traditional procedures [18]. Building on this shift toward greener production, He et al. [19] designed an artificial microbial consortium composed of an engineered sclareol‐producing S. cerevisiae strain and a natural transforming fungus, Hyphozyma roseonigra. By modularly dividing the pathway from glucose to ambradiol, an ambradiol titer of 644.2 mg/L was achieved under optimized conditions, thereby opening a promising route for sustainable ambroxide production. More recently, co‐culture of sclareol‐producing S. cerevisiae with another natural transforming fungus Cryptococcus albidus ATCC 20918 enabled sclareolide (626.3 mg/L) production from glucose [20]. Additionally, sclareol‑containing essential oils show antimicrobial and sedative effects [5]. Clinical studies show sclareol‑containing essential oils reduce pulse rate and blood pressure, providing stress relief and mood regulation [21], similar to the lavender‑based anxiolytic Silexan [22], providing a pharmacological basis for the incorporation of sclareol‐containing essential oils into high‐end personal care and aromatherapeutic products.

2.2. Pharmaceutical and Biological Activities

Sclareol exhibits broad pharmacological potential, with research evolving from early cytotoxicity evaluations to more integrated studies on systemic inflammatory and metabolic regulation [23, 24]. It induces apoptosis and sensitizes cancer cell lines to chemotherapy [25], exhibiting synergistic therapeutic effects when combined with cisplatin and adriamycin, alleviating side effects induced by chemotherapy [26]. Additionally, sclareol and its labdane‐type derivatives demonstrate prominent anti‐inflammatory and antimicrobial properties, effectively modulating immune cytokine profiles to alleviate systemic inflammation and inhibit foodborne pathogens [13, 27]. However, poor aqueous solubility and low bioavailability limit its clinical translation, so sclareol is increasingly valued as a privileged bioactive scaffold for structural modification. Derivatives generated via oxidation or esterification show enhanced antifungal and anti‑herpes activity with low cytotoxicity [28], and oxidative Heck coupling at C15 gives potent anticancer derivatives [29]. More recently, the incorporation of greener methodologies such as photocatalysis, electrocatalysis, and biocatalysis has further broadened the toolbox for the late‐stage functionalization of sclareol and its derivatives [30]. Notably, sclareol also demonstrates growing commercial value as a cosmetic agent by protecting against UVB‐induced photoaging to alleviate facial wrinkles [31].

2.3. Agricultural Applications

In the realm of crop protection, sclareol functions as an important component of the plant secondary metabolic defense system and acts as a natural pesticide. Studies have shown that sclareol effectively inhibits spore germination of fungal pathogens such as wheat rust and soybean rust, thereby significantly reducing crop infection rates [11, 32]. Mechanistically, it can induce lignin accumulation in plants, reinforcing physical barriers against pathogens such as root‐knot nematodes [33, 34]. Sclareol induces the expression of the ABC transporter protein NpABC1 in tobacco (Nicotiana plumbaginifolia) cells [35]. This plasma membrane‐localized transporter mediates the polar secretion of antifungal diterpenoids, underscoring the central role of sclareol in eliciting induced defense responses in plants.

With the rapid advancement of modern biotechnology, significant progress has been made in heterologous sclareol production through enzyme engineering, metabolic pathway compartmentalization, and rational design guided by genome‐scale metabolic models, laying a solid foundation for its broader industrial application in pharmaceuticals, agrochemicals, and functional foods [36, 37].

3. Biosynthetic Basis and Production Methods of Sclareol

At present, sclareol production primarily relies on botanical extraction and microbial biosynthesis. Owing to its intricate molecular architecture, characterized by multiple chiral centers and a rigid cyclic framework, chemical synthesis remains highly challenging, and no economically viable industrial route has been established so far. Consequently, the development of efficient and sustainable biomanufacturing strategies has emerged as a significant research hotspot.

3.1. Plant Extraction

Sclareol predominantly accumulates within the glandular trichomes of the inflorescences and leaves of clary sage [38], and its concentration is significantly influenced by cultivars, cultivation regions, and harvest periods. Since the successful isolation of this compound from the stems and leaves of S. sclarea by Stoll and Hinder in 1950 [17, 39], the efficient recovery of this diterpene has remained a focal point of research. Both the yield and the composition of the resulting extracts are strongly affected by the specific plant tissues selected and the extraction techniques employed. Studies have shown that essential oils obtained via hydro‐distillation are rich in light compounds such as linalyl acetate, with sclareol content limited to approximately 5.9%. In contrast, extraction using organic solvents like dichloromethane can elevate the sclareol concentration to 73.6% [15]. Although distillation yields are extremely low (approximately 0.55%), solvent extraction using n‐hexane or ethanol can achieve a maximum yield of up to 98.3% [40].

In industrial practice, solvent‐based processes typically employ organic solvents such as petroleum ether [41] and n‐hexane [42] to produce clary sage concretes. However, these solvent‐intensive approaches present significant drawbacks: the solvents are often toxic and environmentally hazardous, and require costly downstream operations to ensure complete removal from the final product. In addition, non‐polar solvents tend to co‐extract plant waxes and pigments together with the diterpene fraction [42], further complicating purification.

To overcome the limitations of traditional processes, supercritical CO₂ (SC‐CO₂) extraction has emerged as an attractive alternative owing to its non‐toxic nature, high permeability, and low viscosity. Operating under low‐temperature and high‐pressure conditions, SC‐CO₂ behaves as a tunable solvent that enables the selective extraction of target compounds, minimizes oxidation of thermolabile components, and yields products essentially free of organic solvent residues [43, 44, 45]. Experimental data have shown that, under optimized conditions, the sclareol content in SC‐CO₂ extracts can reach up to 50% of the recovered fraction [46]. Because the solvent power of supercritical CO₂ increases with its density, adjusting temperature and pressure allows fine‐tuning of selectivity. However, excessively high densities favor the co‐extraction of high‐molecular‐weight impurities such as waxes and long‐chain alcohols [45]. In practice, to balance extraction efficiency, selectivity, and process economics, an increasingly adopted strategy is to first concentrate lipophilic components into a clary sage concrete by conventional solvent extraction and then subject this semi‐solid feed to SC‐CO₂ fractionation to enrich sclareol [47, 48, 49].

Nevertheless, plant‐based extraction is inherently constrained by biological and environmental factors. It has been reported that a clary sage field yields approximately 10–15 kg of inflorescences per hectare every two years, and the overall recovery of purified sclareol is only about 35% [50, 51]. The long cultivation cycles, sensitivity to geographical and climatic conditions, and the high cost associated with complex purification steps severely limit the stable and large‐scale supply of sclareol.

3.2. Elucidation of the Biosynthetic Pathway

Plant extraction currently remains the main industrial source of sclareol, but its intrinsic limitations constrain supply stability and increase production costs. In contrast, elucidation of the native biosynthetic pathway has provided the necessary biochemical foundation for transitioning toward sustainable and scalable microbial production.

The precursors of sclareol are IPP and DMAPP, which can be interconverted by isopentenyl pyrophosphate isomerase. In plants, these precursors are synthesized via two spatially separated routes: the mevalonate (MVA) pathway in the cytosol and endoplasmic reticulum, and the 2‐C‐methyl‐D‐erythritol 4‐phosphate (MEP) pathway in the plastids [52, 53]. Despite this compartmentalization, isotopic labeling studies indicate physiologically relevant crosstalk and exchange of C₅ intermediates between the MVA and MEP‐derived isoprenoid pools [54]. However, specific pathway‐inhibitor and 1–¹³C‐glucose tracer experiments conducted in 2021 confirmed that sclareol biosynthesis in clary sage glandular trichomes relies almost exclusively on the plastidial MEP pathway [38]. This finding is crucial as it clarifies the physiological origin of carbon flux directed toward sclareol in the native producer.

The biosynthesis of bicyclic labdane‐type diterpenes proceeds through the stepwise ionization and cycloisomerization of GGPP. In angiosperms, this cascade requires the sequential action of two monofunctional diterpene synthases: a class II diTPS and a class I diTPS [50, 55, 56]. Early in vitro studies using tobacco cell cultures established the presence of such enzymatic activities [57, 58], laying the groundwork for identifying the specific genes involved.

In 2012, Caniard et al. [50] successfully cloned and characterized the two key diterpene synthases from S. sclarea. The class II enzyme, designated SsLPPS, catalyzes the protonation‐initiated cyclization of GGPP to produce labda‐13‐en‐8‐ol diphosphate (LPP, also referred to as LDPP) as its major product. Subsequently, the class I enzyme SsSS triggers ionization of the diphosphate group of LPP, orchestrating its final conversion into sclareol (Figure 1). By combining these two recombinant diterpene synthases with GGPP in vitro, the two‐step conversion to sclareol was reconstituted, validating the enzymatic basis of sclareol biosynthesis [59].

FIGURE 1.

Biosynthetic pathway of sclareol. Arrows represent reactions. Double‐headed arrows represent reversible reactions. Multiple arrows represent multiple enzymatic steps. DXS, 1‐deoxy‐D‐xylulose‐5‐phosphate synthase; DXR, 1‐deoxy‐D‐xylulose‐5‐phosphate reductoisomerase; CMS, 2‐C‐methyl‐D‐erythritol 4‐phosphate cytidylyltransferase; CMK, 4‐diphosphocytidyl‐2‐C‐methyl‐D‐erythritol kinase; MDS, 2‐C‐methyl‐D‐erythritol 2,4‐cyclodiphosphate synthase; HDS, (E)‐4‐hydroxy‐3‐methylbut‐2‐enyl‐diphosphate synthase; HDR, 4‐hydroxy‐3‐methylbut‐2‐en‐1‐yl diphosphate reductase; ACAT, acetyl‐CoA C‐acetyltransferase; HMGS, hydroxymethylglutaryl‐CoA synthase; MK, mevalonate kinase; PMK, phosphomevalonate kinase; MVD, diphosphomevalonate decarboxylase; IDI, isopentenyl‐diphosphate Δ‐isomerase; CDP‐ME, 4‐(cytidine 5′‐diphospho)‐2‐C‐methyl‐D‐erythritol; CDP‐ME2P, 2‐phospho‐4‐(cytidine 5′‐diphospho)‐2‐C‐methyl‐D‐erythritol; ME‐cPP, 2‐C‐methyl‐D‐erythritol 2,4‐cyclodiphosphate; HMBPP, 1‐hydroxy‐2‐methyl‐2‐butenyl 4‐diphosphate; HMG‐CoA, hydroxymethylglutaryl coenzyme A; MVP, 5‐phosphomevalonate; MVPP, 5‐diphosphomevalonate.

Following the discovery of SsLPPS and SsSS, researchers have identified functionally analogous enzymes in other plant species. For example, the copal‐8‐ol diphosphate synthase CcCLS from Cistus creticus [60] and NtCPS2 from Nicotiana tabacum [61] both catalyze the formation of LPP from GGPP. Although these homologous enzymes share similar cyclization mechanisms with SsLPPS, they offer distinct catalytic efficiencies and host compatibilities, thereby providing a versatile enzyme toolbox for pathway engineering.

Overall, the complete elucidation of the native biosynthetic route has not only explained how sclareol is formed in nature but also established the direct genetic blueprint required for its de novo reconstruction in heterologous hosts, as detailed in the following sections.

4. Advances in Microbial Production of Sclareol

With the rapid advancement of synthetic biology and metabolic engineering, extensive work has demonstrated that microbial cell factories can efficiently produce a wide range of terpenoids derived from the universal isoprenoid precursors IPP and DMAPP [62, 63]. Building on the elucidated plant biosynthetic pathway, researchers have transferred SsLPPS and SsSS and engineered isoprenoid precursor supply in a variety of chassis, achieving gram‐per‐liter sclareol titers in several systems (Table 1).

TABLE 1.

Microbial production of sclareol.

Host	Publication year	Engineering strategies	Fermentation mode	Titer	Ref.
E. coli	2012	Overexpression of mvaA, mvaS, mvaK1, mvaK2, mvaD, fni, ERG20, CrtE, atoB; truncated SsLPPS, SsSS	3.7 L fed‐batch	1.5 g/L	[59]
S. cerevisiae	2013	Overexpression of tHMG1 and fusion proteins LPPS‐SS, BTS1‐ERG20	Shake flask	8.96 mg/L	[64]
S. cerevisiae	2015	Overexpression of CD‐HMG2; iterative carotenoid screening, knockout of 6 genes (ROX1, DOS2, VBA5, YER134C, YNR063W, YGR259C); fusion expression of CcCLS with ERG20F96C	Shake flask	750 mg/L	[65]
S. cerevisiae	2015	Overexpression of SsSS, fusion protein CcCLS–ERG20F96C; chromosomal integration of three copies of HMG2(K6R), heterozygous deletion of ERG9; replacement of ERG20 with ERG20F96C .	Shake flask	403 mg/L	[66]
S. cerevisiae	2015	Diploid strain (BY4741L x BY4742) overexpressing tHMG1 and fusion proteins LPPS‐SS, BTS1‐ERG20; high‐cell‐density fed‐batch (ethanol/glucose)	3 L Fed‐batch	408 mg/L	[67]
S. cerevisiae	2023	Enhanced central metabolism; overexpression of fusion proteins MBP‐SS‐LPPS, BTS1‐PaGGPPS; overexpression of ERG20F96C, tHMG1, ERG10, three copies of HMG2(K6R), LAC1, OYE3; knockout of ROX1, DOS2, VBA5, YER134C, YNR063W, YGR259C	1 L Fed‐batch	11.4 g/L	[36]
S. cerevisiae	2024	Overexpression of two copies of tSsLPPS, two copies of tSsSS, CrtE03M, ERG20F96C ; knockout of ROX1; P HXT1 ‐mediated ERG9 downregulation.	Shake flask	357.3 mg/L	[19]
S. cerevisiae	2024	Overexpression of fusion protein TPS‐LPPS, fusion protein BTS1‐ERG20F96C, SaGGPPS, tHMG1, and ZWF1; knockout of ROX1; P HXT1 ‐mediated ERG9 downregulation.	Shake flask	536.2 mg/L	[20]
S. cerevisiae	2025	Chronological lifespan engineering (TOR1‐CLN2, ERG9‐CLN2, FAR8Δ) in precursor‐enhanced strain SCX42; overexpression of ACS1, ACS2	1 L Fed‐batch	25.9 g/L	[68]
Y. lipolytica	2024	Scaffold‐free multienzyme complex via short peptide tags (RIDD/RIAD: tSsLPPS‐RIDD, SsSs‐RIAD); overexpression of tHMG1, ERG20, SsGGPPS, PaGGPPS	5 L Fed‐batch	12.9 g/L	[37]
Y. lipolytica	2025	Overexpression of fusion protein SsSS‐SsLPPS (four copies), GGS1 tPaGGPPS, tHMG1, IDI, ERG20, ERG19, ERG13, ERG12, ERG8, and CAT2; knockout of DGA2; The expression of the SL gene was driven by the P TEFin promoter.	Shake flask	2656.2 mg/L	[69]
Y. lipolytica	2026	Overexpression of BTS1 and ERG20F88C ; four‑copy integration of tSsSSV325I and tSsLPPS; co‑expression of PaGGPPS and SsGGPPS; knockout of DPP1 and LPP1; overexpression of the transporter YALI1_C28310; knockout of CEX1	5 L Fed‑batch	13.9 g/L	[70]
P. pastoris	2026	Overexpression of fusion protein SsSS‑SsLPPS (three copies), ERG20F98C ; PaGGPPS‑BTS1, tHMG1, ERG10‑ERG13, ERG12, ERG8, ERG19, ERG9‐CLN2, ECM33; knockout of VBA5, YNR034W, YGL203C and knockdown of YNL096C‐CLN2	1.5 L Fed‐batch	27.8 g/L	[71]

4.1. Engineered E. coli Strains

E. coli, as a classic heterologous expression host, offers a well‐characterized genetic background, ease of manipulation, and rapid growth, and was therefore the first microbial platform in which de novo sclareol biosynthesis was demonstrated. E. coli generates IPP and DMAPP through its endogenous MEP pathway [72, 73]. However, due to the extremely low intracellular content of GGPP in E. coli, overexpression of an exogenous GGPP synthase is essential to establish a functional diterpene pathway.

Building on the functional characterization of the clary sage diterpene synthases, Schalk et al. [59] engineered E. coli to heterologously express a GGPP synthase from Pantoea agglomerans together with the plant enzymes SsLPPS and SsSS, thereby reconstituting the complete sclareol biosynthetic pathway and achieving de novo production of sclareol in this host. To enhance precursor supply, a heterologous MVA pathway was introduced. To overcome product volatility and feedback inhibition, a liquid–liquid two‐phase cultivation strategy was employed, with the addition of 10% dodecane as an organic phase for in situ product extraction. Coupled with high‐cell‐density fed‐batch fermentation technology, a final sclareol titer of 1.5 g/L was achieved. Subsequent efforts have largely shifted away from E. coli toward eukaryotic hosts, because the absence of an endogenous MVA pathway, limited membrane system, and lower compatibility with plant‐derived enzymes pose significant bottlenecks for further improving sclareol yields.

4.2. Engineered S. cerevisiae Strains

S. cerevisiae, favored for its generally recognized as safe (GRAS) status, active cytosolic MVA pathway, mature genetic toolbox, and high compatibility with plant‐derived enzymes, has become the core host for microbial sclareol synthesis. In 2012, Caniard et al. [50] used heterologous expression in yeast to validate the functions of SsLPPS and SsSS in vivo, thereby demonstrating the suitability of engineered yeast for sclareol production. Over the past decade, engineering strategies have evolved from single‐gene modifications to system‐level metabolic rewiring, resulting in orders‐of‐magnitude increases in sclareol titers in yeast. Following the construction of the sclareol biosynthetic pathway in S. cerevisiae, subsequent studies have mainly focused on five optimization strategies to improve production performance: precursor supply engineering, competitive pathway blocking, biosynthetic enzyme engineering, cellular longevity engineering, and fermentation and process optimization (Figure 2).

FIGURE 2.

Metabolic engineering strategies for enhancing microbial production of sclareol. The figure is created in https://BioRender.com. (A) Precursor supply engineering; (B) Competing pathway blocking; (C) Engineering of sclareol synthases; (D) Cellular longevity engineering; and (E) Fermentation and process optimization. In panels (A–D), genes shown in red are typically overexpressed, whereas genes shown in gray are typically knocked out or downregulated. In panel (B), downregulation of ERG9 reduces the formation of the byproduct squalene. In panel (C), MBP denotes maltose‐binding protein. In panel (D), the specific regulatory strategies for extending yeast chronological lifespan are highlighted within the red dashed box, including: attaching the degradation peptide CLN2 at the C‐terminus of TOR1 (TOR1‐CLN2) to enable conditional degradation; and deletion of FAR8 to enhance mitophagy. "P" in the diagram denotes phosphorylation. Acetyl‐CoA, acetyl coenzyme A; α‐KG, α‐ketoglutarate; Acetoacetyl‐CoA, acetoacetyl coenzyme A; NADPH, reduced nicotinamide adenine dinucleotide phosphate; NADP⁺, oxidized nicotinamide adenine dinucleotide phosphate; DGA2, diacylglycerol acyltransferase 2; CAT2, carnitine acetyltransferase 2; ACS1/2, acetyl‐CoA synthetase 1/2; MPC, mitochondrial pyruvate carrier; RtCIT1; Rhodotorula toruloides citrate synthase 1; CTP1, citrate transport protein 1; YHM2, Citrate/oxoglutarate carrier protein; AnACL, Aspergillus nidulans‐ATP‐citrate lyase; MmACL, Mus musculus ATP‐citrate lyase; ERG10, acetoacetyl‐CoA thiolase; HMG2, 3‐hydroxy‐3‐methylglutaryl‐CoA reductase isozyme 2; SpHMGR, Streptococcus pneumoniae‐derived HMG‐CoA reductase; CD‐HMG2, catalytic domain of HMG‐CoA reductase; TOR1, target of rapamycin; CLN2, cyclin 2 (the PEST degron is derived from the C‐terminal PEST‐rich domain of CLN2); Far3‐11, the negative regulators of the Far complex.

4.2.1. Precursor Supply Engineering

Efficient heterologous synthesis of sclareol critically depends on the high‐flux allocation of carbon from central metabolism to the isoprenoid pathway. In S. cerevisiae, this involves the systematic reconstruction of three core modules: cytosolic acetyl‐CoA supply, MVA pathway reinforcement, and GGPP precursor provision (Figure 2A). As the starting point of the entire pathway, the cytosolic acetyl‐CoA level directly determines the theoretical upper limit of downstream yield. Traditional acetyl‐CoA synthetase overexpression strategies, such as upregulating ACS1/ACS2 within the ethanol utilization pathway, have been widely adopted to boost cytosolic acetyl‐CoA pools. More recently, integrating this type of precursor supply enhancement with chronological lifespan engineering enabled a sclareol titer of 25.9 g/L with a yield of 0.051 g/g glucose [68]. An alternative and conceptually appealing approach is to redirect metabolic flux from fatty acid metabolism. For instance, Cao et al. [36] utilized a modular metabolic reconstruction strategy to partition global metabolism into three modules: central metabolism, the isoprenoid pathway, and regulatory factors. By restoring FAA1/4 while simultaneously deleting FAS1/2 and TESA, they leveraged the high‐flux background of fatty acid synthesis to provide an abundant supply of acetyl‐CoA and NADPH for terpenoid synthesis, saving 18 genetic operations (including 8 genes for acetyl‐CoA supply and 10 genes for NADPH supply). This “chassis‐borrowing” strategy bypasses the growth inhibition typically associated with extensive genetic manipulation of central metabolism, achieving an efficient switch of carbon flux.

Regarding the reinforcement of the MVA pathway, research focus has shifted from targeting single rate‐limiting enzymes to constructing global metabolic pumps. HMGR is a pivotal enzyme in the GGPP biosynthetic pathway. Due to its complex feedback inhibition and degradation regulation, it has become a primary engineering target. Among the endogenous HMGR isoenzymes, HMG1 is more stable and less prone to degradation than HMG2 [74]. Given that endogenous HMG1 is stringently regulated by sterol levels, employing a truncated version (tHMG1) has become a universal strategy to relieve negative feedback inhibition [19, 20, 64, 67, 68]. By truncating the N‐terminal transmembrane domain—which senses degradation signals—the enzyme tHMG1 is released from the endoplasmic reticulum into the cytosol. This modification maintains high stability while ensuring a continuous influx of metabolic flux toward isoprenoid synthesis. Yang et al. [64] initially achieved a yield of 8.96 mg/L by overexpressing tHMG1 and introducing the fusion protein GGPPS‐FPPS (ERG20‐BTS1) to exploit substrate channeling. Subsequently, the team significantly expanded the precursor pool through multi‐gene synergy by co‐expressing tHMG1, ERG20^F96C , ERG10, and a stable HMG2(K6R) mutant [36]. Subcellular compartmentalization has also shown great potential in mitigating metabolic interference. Targeting isoprenoid biosynthesis modules to specific compartments such as mitochondria can increase local FPP concentrations, creating an optimized microenvironment for downstream synthases [66].

In terms of enhancing GGPP supply, the core challenge lies in intercepting the FPP carbon flow directed toward the sterol branch and redirecting it toward GGPP. By engineering FPP synthase into the mutant ERG20^F96C , the enzyme acquires the capability to synthesize GGPP directly. Ignea et al. [66] successfully redirected metabolic flux from sterol synthesis to diterpene production using this strategy, achieving a titer of 403 mg/L. To balance cell growth and product synthesis, a dynamic induction system was constructed by deleting GAL80, so that genes controlled by GAL promoters are repressed at high glucose concentrations and induced under low‐glucose or non‐fermentable carbon sources. Combined with the downregulation of competing pathways, controlled carbon flux redirection can thus be achieved. Researchers have overexpressed genes such as ERG20^F96C, tHMG1, and CrtE03M and deleted ROX1 on a GAL80Δ background to strengthen precursor supply, increased the copy number of downstream synthases to pull the metabolic flux, and simultaneously downregulated ERG9 to block competing pathways. This comprehensive strategy resulted in a sclareol yield of 357.3 mg/L [19]. Additionally, optimizing GGPP synthesis and cofactor balance is also an important direction for improving production. Coordinated overexpression of GGPP‐forming enzymes and modification of the HMGR catalytic domain can increase GGPP production, thereby enhancing sclareol titers [75]. Introduction of SaGGPPS from Sulfolobus acidocaldarius, enhancement of MVA flux (overexpression of tHMG1), and upregulation of ZWF1 to increase NADPH generation achieved the high‐yielding strain S9 (536.2 mg/L) [20]. During large‐scale screening and iterative strain improvement, fusion expression of CcCLS with ERG20^F96C combined with overexpression of the stabilized HMG2 variant (CD‐HMG2) further pushed the yield to 743 mg/L [65].

At the transcriptional regulation level, global repressors that negatively regulate the MVA pathway inherently limit precursor supply. To address this, Trikka et al. [65] employed an iterative carotenoid‐screening strategy on a library of 4700 yeast deletion mutants to systematically identify targets that enhance GGPP flux. Stacking the deletions of six endogenous genes (ROX1, DOS2, YER134C, VBA5, YNR063W, and YGR259C) yielded a shake‐flask sclareol titer of 750 mg/L, a 12‐fold increase over the initial strain. Notably, ROX1 is a known transcriptional repressor of MVA pathway genes [76], and its deletion is now widely adopted to relieve bottlenecks in isoprenoid biosynthesis [77]. The benefits of ROX1 knockout are highly reproducible: it increased sclareol production by 30.4% in a modularly engineered strain [36] and by 24.7% in a separate MVA‐optimized shake‐flask culture [19]. Moreover, ROX1 knockout remains highly effective in complex co‐culture systems [20]. Together, these consistent improvements across various setups confirm ROX1 as a key target for optimizing metabolic flux in high‐level diterpenoid production.

4.2.2. Competing Pathway Blocking

Heterologous pathways inevitably compete with the host's endogenous metabolic network for shared precursors, severely limiting sclareol production. The sterol pathway is the primary competitor, as squalene synthase ERG9 diverts FPP toward squalene (Figure 2B). To address this, researchers frequently replace the native ERG9 promoter with the glucose‐inducible promoter P _HXT1 . Because P _HXT1 is repressed upon glucose depletion, this strategy effectively decouples cell growth from product synthesis and minimizes competition during the production phase [78]. Consequently, dynamic downregulation of ERG9 significantly decreased FPP flux toward squalene, increasing sclareol titers by 24.3%–27.2% [19, 36]. Beyond transcriptional control, fusing the CLN2 degradation tag to ERG9 has enabled more precise regulation of enzyme levels by reducing its intracellular half‐life [68]. This progression from promoter replacement to targeted protein degradation highlights the shift toward fine‐tuned control of competing metabolic pathways.

4.2.3. Engineering of Sclareol Synthases

Plant‐derived diterpene synthases often display low expression levels, modest catalytic efficiency, and suboptimal compatibility with yeast metabolism. Therefore, enzyme engineering targeting these bottlenecks is central to optimizing the sclareol synthesis pathway. The heterologous enzymes for sclareol biosynthesis, the class II labdane‐related diterpene synthase and the class I sclareol synthase, directly determine pathway efficiency through their activity, stability and expression level. Through prediction and experimental validation, Yang et al. [64] removed the N‐terminal signal peptides of SsLPPS and SsSS, thereby improving their expression in yeast and increasing sclareol production by 2.4‐fold.

Enzyme fusion is an effective strategy to enhance substrate channeling and reduce intermediate leakage (Figure 2C). Fusion expression of SS and LPPS resulted in a 6.7‐fold increase in sclareol titer [36]. Subsequent incorporation of a maltose‐binding protein (MBP) tag at the N‐terminus of the fusion protein to enhance enzyme stability further boosted the yield by 43%. A similar upstream‐downstream fusion strategies were also validated. N‐terminal fusion of the Cistus creticus‐derived CcCLS with engineered ERG20^F96C facilitated efficient conversion of DMAPP to the intermediate 8OH‐CPP, ultimately achieving a sclareol titer of 403 mg/L in shake‐flask fermentation [66]. Collectively, signal peptide truncation, enzyme fusion, and introduction of stabilizing tags have significantly enhanced the efficiency of sclareol biosynthesis in yeast.

Future gains in production are expected from in‐depth engineering of these heterologous enzymes. For instance, rational design guided by crystal structures or artificial intelligence‐based predictive models may be used to fine‐tune active‐site residues and access tunnels, thereby enhancing substrate specificity and catalytic efficiency [79, 80]. Alternatively, scaffold‐free multienzyme complex strategies enable the precise construction of metabolic channels by modulating the spatial organization of enzymes, thereby further reducing intermediate leakage [81, 82]. Furthermore, systematic screening of fusion orientations and optimization of linker length and composition will provide additional opportunities to synergize enzymes originating from different origins within the same pathway.

4.2.4. Cellular Longevity Engineering

In addition to pathway‐ and process‐level optimizations, the physiological robustness and lifespan of production strains can become limiting factors during prolonged fed‐batch fermentations. Wu et al. [68] combined chronological lifespan engineering with classical metabolic engineering, thereby expanding the strategies available for improving sclareol production (Figure 2D). By attenuating nutrient‐sensing pathways (TOR1‐CLN2) and enhancing mitophagy (FAR8Δ), they significantly prolonged cell survival during extended fermentation, increasing sclareol production by 70.3% compared to the base metabolic engineering strain. Combining this with overexpression of acetyl‐CoA synthetase (ACS1/2) and precise regulation of the competing sterol pathway using a degradation tag (ERG9‐CLN2) ultimately led to a fed‐batch titer of 25.9 g/L with a yield of 0.051 g/g glucose, representing the highest titer reported to date for microbial sclareol production in S. cerevisiae. This study was among the first to link cellular longevity to sclareol biosynthetic capacity. It provides an alternative strategy for improving production. Specifically, remodeling mitophagy and nutrient‐sensing pathways established a positive correlation between cellular longevity and biosynthetic capacity in high‐density sclareol fermentations.

4.2.5. Fermentation and Process Optimization

Building upon efficient sclareol synthesis modules, fermentation and process optimization are essential to fully realize the production potential of engineered strains. Key operational parameters such as medium composition, cultivation temperature, induction regime, and fermentation strategy must be tuned to balance biomass formation with product synthesis, minimize intermediate accumulation and side reactions, and ultimately enhance titer, productivity, and yield. In addition, accumulation of hydrophobic products such as sclareol can exert cytotoxicity and impose membrane stress, thereby limiting both cell growth and productivity. Establishing a two‐phase cultivation system for in situ product extraction into an overlay organic phase helps alleviate metabolic burden (Figure 2E). For example, the use of a synthetic medium that improved plasmid stability increased the retention rate of both plasmids from 58.0% in YPD medium to 72.8%. Combined with in situ extraction with dodecane, a glucose/ethanol mixed carbon source, and a high cell density fed‐batch process with dissolved oxygen‐stat (DO‐stat) feeding, the sclareol titer reached 408 mg/L in S. cerevisiae [67]. Adding 10% dodecane after 24 h of fermentation effectively captured sclareol in the organic phase [64]. He et al. [19] systematically optimized fermentation in strain YN42. Upon addition of 10% dodecane, 92.9% of sclareol was captured in the organic phase, and the titer increased by 23.8% compared with monophasic culture. Alternatively, adding 10 mM hydroxypropyl‐β‐cyclodextrin (HP‐β‐CD) dramatically increased the extracellular sclareol proportion from 10.6% to 98.6%, highlighting the value of employing solubilizing agents to facilitate product export and enabling efficient de novo ambradiol production in a related co‐culture system. Consistent with this concept, enhanced inter‐strain mass transfer by adding 0.5% Triton X‐100 in a similar co‐culture system boosted sclareolide production by 2.87‐fold [20].

Complementary to extraction and transport engineering, feeding strategies and cultivation modes are critical levers for maximizing space‐time yields. Cao et al. [36] achieved a significant sclareol titer of 11.4 g/L in a 1 L monophasic fed‐batch bioreactor by employing an engineered S. cerevisiae strain and an exponential feeding strategy (μ = 0.05 h⁻¹) at pH 5.6. Building on this success, the same team further engineered the strain, extended the fermentation period, and optimized the feeding profile under identical monophasic conditions, ultimately achieving an elevated titer of 25.9 g/L with a yield of 0.051 g/g glucose [68]. Notably, both studies employed a streamlined monophasic cultivation approach. During these high‐titer fermentations, sclareol spontaneously precipitated and adhered to the bioreactor walls. This in situ phase separation effectively reduced the dissolved product concentration in the culture broth, thereby alleviating potential cytotoxicity. Compared to traditional biphasic fermentation, this monophasic fermentation eliminates the need for organic overlays (such as dodecane). This not only reduces operational costs and toxicity risks but also prevents solvent‐induced cell damage, ensuring prolonged cell viability and production stability. Furthermore, the final product can be efficiently recovered from the whole broth via simple n‐hexane extraction and centrifugation. Ultimately, the combination of simplified reactor operation and straightforward downstream processing makes the monophasic fermentation strategy potentially attractive for industrial sclareol production.

4.3. Engineered Y. lipolytica Strains

Y. lipolytica, an emerging non‐conventional oleaginous yeast, has become a popular chassis for terpenoid synthesis due to its abundant cytosolic acetyl‐CoA supply, strong robust tricarboxylic acid (TCA) cycle, and high tolerance to hydrophobic products [83]. The rapid development of genetic tools and lipid‐metabolism engineering strategies has laid a solid foundation for high‐level sclareol production. In 2024, Sun et al. [38] constructed the de novo sclareol biosynthesis pathway in Y. lipolytica for the first time. The study screened and truncated the signal peptide of SsLPPS and identified the more active CcCLS. By constructing a two‐layer GGPP accumulation pathway and co‐expressing the FPP‐preferring SsGGPPS with the direct IPP/DMAPP‐utilizing PaGGPPS (Pantoea agglomerans‐derived geranylgeranyl pyrophosphate synthase), they maximized GGPP supply, increasing sclareol titer by 34.4 folds. Innovatively, they used RIDD/RIAD short peptide tags to assemble tSsLPPS and SsSS into a scaffold‐free multienzyme complex with a 2:1 stoichiometry (Figure 2C), significantly promoting substrate channeling and increasing yield by a further 58.25%. Ultimately, fed‐batch fermentation in a 5 L bioreactor yielded 12.9 g/L of sclareol, representing the highest titer achieved in this host at that time.

The following year, Chen et al. [69] further elevated sclareol production in Y. lipolytica through a combinatorial metabolic engineering approach. They first fused SsSS and SsLPPS via a GGG linker, which resulted in a 2.94‐fold improvement in yield. To optimize precursor supply, they introduced the endogenous GGPP synthase GGS1 and systematically evaluated GGPP synthases (GGPPSs) from different sources. They found that a truncated GGPPS from Phomopsis amygdali (designated tPaGGPPS) [37] outperformed other variants in boosting sclareol production. Next, building upon a chassis that already overexpressed tHMG1, IDI, and ERG20, they integrated ERG19, ERG13, ERG12, and ERG8 to systematically strengthen the MVA pathway and balance metabolic flux, thereby increasing the titer to 858.11 mg/L. Innovatively, they redirected the central acetyl‐CoA pool from lipid accumulation toward terpenoid synthesis by enhancing lipid degradation (overexpressing CAT2) and disrupting lipid synthesis (knocking out DGA2) (Figure 2A), which further raised the titer to 1113.17 mg/L. Finally, integrating four additional copies of the fusion gene yielded the final strain YAs24. This strain produced 2656.20 mg/L of sclareol in shake flasks, representing a 746‐fold increase over the initial strain.

In a recent study, Huang et al. [70] first co‑expressed the truncated sclareol synthase genes tSsLPPS and tSsSS in Y. lipolytica, and subsequently engineered tSsSS through semi‑rational mutagenesis, obtaining the tSsSS^V325I mutant, which improved the sclareol titer by 70%. Subsequently, they optimized the expression of these two key genes via multi‑copy integration. To maximize precursor availability and minimize byproduct formation, they introduced heterologous GGPP synthases (PaGGPPS and SsGGPPS) and simultaneously knocked out DPP1 and LPP1 to prevent the diversion of GGPP. Furthermore, they harnessed the ABC transporter YALI1_C28310 to facilitate product efflux and knocked out the citrate exporter CEX1 to eliminate competitive citrate overflow metabolism. As a result, the engineered strain achieved a sclareol titer of 3307.3 mg/L in shake flasks and 13.9 g/L in a 5‑L bioreactor, establishing the highest sclareol production level reported to date in Y. lipolytica.

4.4. Engineered Pichia pastoris Strains

Pichia pastoris has been progressively developed as a chassis cell for the synthesis of terpenoids and other secondary metabolites, owing to its high‐cell‐density fermentation capability and strictly respiratory metabolism. Recently, Zhang et al. [71] established a host‐tailored engineering framework in this yeast, which helped alleviate metabolic constraints and enabled efficient sclareol biosynthesis. At the pathway level, they constructed a SS‐LPPS fusion protein, strengthened the mevalonate pathway by introducing the ERG20^F98C mutation, overexpressing PaGGPPS‐BTS1 fusion protein, and employing truncated tHMG1. Additionally, they attached a CLN2PEST degradation tag to Erg9 to attenuate the competing sterol pathway, while increasing the copy number of key pathway genes, thereby significantly enhancing precursor supply. At the regulatory level, multi‐omics‑guided target mining identified several novel regulatory targets, including overexpression of the cell wall protein Ecm33, deletion of VBA5, YNR034W, and YGL203C, and CLN2PEST‑mediated knockdown of YNL096C. Moreover, deletion of genes encoding the global transcriptional regulators Cra1 and Gln3 minimized ethanol overflow and reduced carbon diversion into nitrogen metabolism, further improving carbon utilization toward sclareol. At the process level, glycerol was used as the main carbon source instead of glucose and combined with a pH‑coupled ammonia feeding strategy that simultaneously controlled nitrogen supply and culture pH, thereby maintaining osmotic and redox balance. Ultimately, under monophasic fermentation conditions, the engineered strain PFM63H achieved a sclareol titer of 27.8 g/L in minimal medium in a 1.5 L bioreactor, representing the highest titer reported to date for microbial sclareol production.

5. Photosynthetic and Plant‐Based Production Platforms

While reconstructing the sclareol pathway in heterotrophic microbes is the mainstream strategy for industrial scale‐up, researchers are also exploring photosynthetic organisms and engineered plants as alternative, solar‐driven production methods. These platforms aim to bridge the gap between traditional agricultural extraction and industrial fermentation by directly converting CO₂ into high‐value diterpenes.

Chlamydomonas reinhardtii, a unicellular photosynthetic microalga, utilizes only the MEP pathway for terpene synthesis and harbors abundant GGPP in its chloroplasts. In 2019, Papaefthimiou et al. [84] first established a platform for producing labdane‐type diterpenes in the C. reinhardtii chloroplast. By codon‐optimizing the CcCLS gene and driving its expression with strong algal promoters, they achieved a maximum sclareol yield of 0.038 µg/mg dry weight. Later, Einhaus et al. [85] demonstrated that translational fusion of endogenous CrGGPPS with SsLPPS, combined with overexpression of the MEP pathway rate‐limiting enzyme SpDXS2, boosted sclareol production 8‐fold. Using photoautotrophic high cell density cultivation in a 2.5 L photobioreactor, the titer reached 656 mg/L after 19 days, demonstrating the potential of microalgae.

Similarly, Physcomitrella patens, a model moss characterized by highly efficient homologous recombination, can be easily cultivated in liquid bioreactors. In 2015, Pan et al. [86] established a sclareol production platform in P. patens by constructing a single expression cassette linking SsLPPS and SsSS with a 2A peptide, leading to a maximum production of 2.84 mg/g dry weight. Notably, by quantifying sclareol in both the biomass and the culture medium, the study revealed that a larger proportion of the synthesized sclareol was secreted extracellularly. This finding not only validates the moss as a viable photosynthetic platform but also suggests the presence of endogenous transporters capable of facilitating sclareol efflux to alleviate intracellular toxicity.

Beyond lower plants and algae, higher crops like tobacco (Nicotiana tabacum) are increasingly engineered as “green external factories”. Tobacco offers a short growth cycle, high biomass, and well‐established genetic manipulation toolkits. More importantly, its leaf surface is densely covered with glandular trichomes that naturally synthesize and store massive amounts of terpenoids [87, 88, 89]. Ma et al. [90] recently introduced the LPPS and SS genes from S. sclarea into tobacco. Transgenic lines driven by the trichome‐specific P _eCBTS promoter achieved sclareol accumulation at 4.1 µg/cm², comparable to the natural producer Nicotiana glutinosa, with an estimated yield of nearly 1.5 kg per hectare. The suitability of tobacco is intrinsically linked to its native LPP precursor pool [61] and the presence of sclareol‐induced ABC transporters (e.g., NpPDR1) that actively facilitate product export [35, 91].

Overall, the development of diverse biological chassis—ranging from established heterotrophic cell factories such as S. cerevisiae and Y. lipolytica with gram‐per‐liter titers, to emerging solar‐driven platforms including microalgae and engineered plants—provides a rich and highly complementary technological portfolio. To systematically evaluate the suitability of these different production platforms, this article presents a comprehensive cross‐platform comparison of microbial chassis, microalgae, moss, and tobacco across six key dimensions: carbon efficiency, growth rate, containment, genetic stability, extraction burden, and scalability (Table 2). In terms of absolute titer, the heterotrophic microbial system (27.8 g/L, fed‐batch fermentation) outpaces the liquid suspension culture of moss (2.28 mg/L) by over four orders of magnitude and exceeds the photoautotrophic production of microalgae (656 mg/L, 2.5 L photobioreactor) by approximately 42‐fold. However, while modern monophasic or biphasic fermentation systems successfully drive sclareol to accumulate extracellularly—thus avoiding energy‐intensive cell disruption—microbes still face a moderate extraction burden due to the reliance on organic solvents for downstream polishing. Additionally, they exhibit a lower carbon efficiency stemming from their inherent reliance on exogenous organic sugar feedstocks.

TABLE 2.

Comparative analysis of bioproduction platforms for sclareol.

Comparison dimension	Microbial chassis	Microalgae	Moss	Tobacco
Carbon efficiency	Moderate; relies on exogenous sugars with inherent carbon loss (CO2 release) during pyruvate decarboxylation and competitive biomass synthesis.	High; photoautotrophic, directly fixes CO2 via chloroplast MEP pathway; however, MEP carbon flux must simultaneously satisfy chlorophyll/carotenoid biosynthesis	Moderate; photoautotrophic, but non‐photosynthetic tissues and cell wall polysaccharide synthesis cause significant carbon partitioning away from product	Low product‐specific carbon flux; while overall whole‐plant carbon fixation is efficient, most carbon is partitioned to vegetative growth (roots, stems) rather than the localized glandular trichomes
Growth rate	Fastest; industrial scale‐up achieves g/L‐level titers within days	Moderate; cultivation period 14–30 days in photobioreactors	Slow; liquid suspension culture period typically 2–4 weeks	Slowest; obtaining stable transgenic lines and completing field/greenhouse cultivation requires 3–6 months
Containment (Biosafety)	High; fully enclosed stainless‐steel fermenters; hosts predominantly GRAS strains	High; enclosed photobioreactor cultivation; no environmental release risk	High; enclosed liquid bioreactor suspension cultivation	Low; open‐field cultivation; risks of pollen drift, seed dispersal, and transgene escape
Genetic stability	High; mature gene‐editing tools enable precise chromosomal integration, avoiding plasmid loss and ensuring stable expression	Moderate; chloroplast transformation relatively stable; high‐yield strains require continuous maintenance through selection	High; efficient homologous recombination enables precise, stable single‐copy integration	Moderate; obtaining homozygous lines requires lengthy multi‐generational screening, and long‐term cultivation faces risks of transgene silencing
Extraction Burden	Moderate; products often precipitate in monophasic systems or partition in biphasic systems, however, downstream polishing still typically requires large volumes of organic solvents	High; intracellular accumulation necessitates energy‐intensive physical disruption (e.g., sonication, bead milling) to release products, followed by solvent extraction and concentration	Low; a substantial proportion of sclareol is secreted extracellularly, simplifying recovery directly from the culture medium	Low (leaf trichomes); ∼95% of the surface‐accumulated product can be recovered by simple solvent rinsing
Scale‐up limitations	Highly scalable; stainless‐steel fermenters seamlessly scale to tonnage; process control parameters are fully quantified and standardized	Limited by light penetration and land footprint; open ponds are susceptible to contamination and light limitation	Limited by bioreactor scale‐up; plant cells face challenges of high broth viscosity and shear sensitivity; lack of industrial experience	Highly scalable via agricultural cultivation, but strictly constrained by field GMO regulations and seasonal/climatic dependencies

In contrast, photosynthetic platforms boast superior carbon efficiency by directly fixing CO₂, driving a highly sustainable carbon economy. Among them, microalgae and moss cultivated in liquid bioreactors offer strict physical containment (biosafety) and manageable genetic stability. Their unique capacity for extracellular secretion (as demonstrated in P. patens) significantly minimizes the downstream extraction burden. However, their scalability is hindered by distinct platform‐specific bottlenecks: while microalgae are primarily constrained by light‐penetration limits in large‐scale photobioreactors, moss suspension cultures face severe bioreactor engineering challenges related to high broth viscosity and the inherent shear sensitivity of plant cells. Conversely, tobacco operates as an agricultural platform, offering virtually unlimited scalability and a drastically reduced extraction burden by sequestering sclareol in easily accessible surface trichomes. Nevertheless, it is heavily constrained by prolonged growth cycles, inherent seasonal and climatic dependencies, and the stringent containment regulations governing field‐grown genetically modified organisms (GMOs). Collectively, these distinct profiles dictate that no single chassis is universally optimal. Instead, they represent a coherent strategy: future biomanufacturing strategies for sclareol and other high‐value terpenoids should balance titer against sustainability, and flexibly integrate these platforms to meet different regulatory and economic demands across the supply chain.

6. Challenges and Perspectives

After over two decades of intensive research, remarkable achievements have been made in the microbial synthesis of sclareol. Titers have advanced from initial gram‐scale production in E. coli to more than 25 g/L in engineered S. cerevisiae and P. pastoris, largely driven by successive efforts of pathway, chassis, and process engineering. Despite these achievements, several critical challenges must still be addressed to achieve economically viable, industrial‐scale production.

Metabolic network rigidity and pathway compatibility remain primary obstacles. Heterologous plant‐derived diterpene synthases often suffer from low expression, poor catalytic activity, or misfolding in microbial hosts, severely limiting carbon flux through the sclareol biosynthetic pathway. Although strategies such as codon optimization, signal peptide truncation, and enzyme fusion have yielded notable success, seamless compatibility between plant enzymes and microbial host metabolism is still far from realized. Systematic comparison of SsLPPS/SsSS orthologs across S. cerevisiae, Y. lipolytica, and P. pastoris to identify host‐specific codon adaptation and folding requirements has not been performed. Future advances in enzyme and pathway engineering are likely to draw on strategies already validated in structurally related diterpenoids. AlphaFold‐enabled structural modeling, for instance, has identified activity hotspots in class I terpene synthases with low template homology and guided mutagenesis that improved catalytic efficiency [70, 79, 80], suggesting that the same approach could be extended to resolve the expression and folding problems that currently limit SsLPPS and SsSS. In Y. lipolytica, the RIDD and RIAD short‐peptide tag system improved sclareol yield by 58.25% through proximity channeling of pathway enzymes [37], demonstrating that scaffold‐free multienzyme assemblies can overcome diffusion losses in diterpenoid metabolism. Rational organelle targeting to the endoplasmic reticulum or lipid droplets has similarly increased flux through competing terpenoid pathways by concentrating enzymes and their hydrophobic substrates [81, 82]. These precedents indicate that integrating structure‐guided enzyme redesign with spatial organization strategies offers a concrete route toward precise carbon allocation in sclareol biosynthesis.

During prolonged, high‐intensity fermentation, product‐induced membrane toxicity and metabolic burden‐associated cellular aging can severely compromise strain productivity. Thus, improving cell viability and long‐term robustness is as important as enhancing instantaneous metabolic flux. The underlying cytotoxic mechanism is closely tied to the physicochemical properties of sclareol. As a hydrophobic diterpene diol, sclareol partitions readily into lipid bilayers, and molecular dynamics simulations indicate that its hydrophobic core inserts into and disorders the phospholipid bilayer despite its two hydroxyl groups [92, 93]. Differential scanning calorimetry (DSC) studies have also confirmed this. When sclareol is incorporated into model membranes (liposomes) composed of dipalmitoylphosphatidylcholine (DPPC), it significantly alters the membrane properties [94]. These changes include elimination of the pretransition, a decrease in the main phase transition temperature, and a reduction in the corresponding enthalpy change (ΔH), all of which indicate that the insertion of sclareol disrupts the ordered packing of the lipid bilayer and increases its fluidity. Furthermore, maintaining genetic and phenotypic strain stability over multiple generations in large‐scale bioreactors remains a critical prerequisite for commercialization. The concept of chronological lifespan engineering, as demonstrated by Wu et al. [68], provides an additional strategy for improving sclareol production by remodeling mitophagy and nutrient‐sensing pathways. Specifically, attenuation of TOR1‐CLN2 signaling combined with enhancement of FAR8‐dependent mitophagy extended cell survival during prolonged fed‐batch fermentation and contributed to the current 25.9 g/L record. Future research should further elucidate the principles of cellular homeostasis under industrial conditions, aiming to construct robust cell factories capable of maintaining high viability and stable expression profiles over extended fermentation periods through dynamic sensing and regulatory networks, thereby ensuring stable operation in fluctuating industrial environments.

Product toxicity and transport bottlenecks are equally pressing issues. The high hydrophobicity of sclareol renders its transmembrane transport heavily dependent on non‐specific diffusion or inefficient endogenous transporters. Notably, the plant ABC transporter NpABC1 is induced by sclareol [35], but subsequent biochemical studies found no direct evidence that this transporter physically translocates sclareol itself, leaving the precise efflux mechanism unresolved. In microbial chassis, no dedicated sclareol exporter has been validated beyond the single report of YALI1_C28310 in Y. lipolytica, which increased extracellular sclareol by 8.6‐fold [70]. Nevertheless, transporter engineering has proven effective for structurally analogous hydrophobic terpenoids in yeast. Endogenous PDR‐family transporters of S. cerevisiae including Pdr5, Pdr10, Snq2, and Yor1 have been successfully redirected to export β‐carotene [95], and heterologous expression of Y. lipolytica‐derived ABC2/ABC3 transporters improved S. cerevisiae tolerance to long‐chain alkanes by approximately 80‐fold [96]. These precedents suggest that systematic screening of PDR‐family and oleaginous‐yeast‐derived transporters, combined with site‐directed mutagenesis of substrate‐binding residues, represents a promising but largely untested avenue for dedicated sclareol efflux engineering. Although process‐level solutions such as biphasic cultivation, cyclodextrin supplementation, and surfactant‐assisted export have proven effective, a more sustainable, long‐term solution requires the mining and engineering of dedicated terpene efflux pumps. Concurrently, tuning membrane composition to mitigate endocytic and envelope stress will be essential, in line with recent advances in transporter engineering for other lipophilic natural products [97, 98].

Addressing downstream separation and industrial scalability constraints is equally critical. On the bioprocessing front, while recent advances in monophasic fermentation have greatly simplified initial product recovery via spontaneous precipitation and crystallization, large‐scale downstream processing often still relies on massive volumes of organic solvents (e.g., n‐hexane) for final extraction. This constitutes a heavy cost and environmental burden: solvent consumption, recovery, and waste treatment are recurring contributors to operating expenditure in comparable microbial terpene processes [99, 100, 101], where downstream separation has been identified as a major techno‐economic bottleneck. Specifically, fragrance‐grade sclareol as an ambroxide precursor demands purity exceeding 98% [102], requiring the complete removal of co‐extracted host lipids, sterols, and pigments. Achieving this stringent purity at an industrial scale necessitates additional chromatographic or crystallization steps that have not been systematically benchmarked across platforms. Concrete directions include maximizing the benefits of spontaneous product precipitation to minimize solvent usage, the development of solvent‑minimized in situ product‑capture resins that selectively bind sclareol over host lipids, and conducting rigorous techno‑economic analysis and life‑cycle assessments of monophasic versus biphasic processes. Direct comparisons of crystallization‑based and chromatographic polishing routes are also required to determine the most cost‐effective path to achieve fragrance‑grade purity. Ultimately, completely substituting traditional organic solvent extraction with sustainable alternatives—such as the development of solvent‐free in situ product‐capture resins or advanced membrane‐based separation technologies—remains a major open challenge for achieving truly green biomanufacturing. Furthermore, complementary production concepts such as exploiting crop leaf‐surface trichomes as “external green factories” in parallel with microbial fermentation, may further strengthen the techno‐economic feasibility of biobased sclareol.

Looking ahead, the integration of genome‐scale metabolic models with multi‐omics data offers a powerful route to systematically identify hidden competing sinks and regulatory constraints, thereby prioritizing engineering targets with quantitative rigor. Simultaneously, the emergence of oleaginous hosts like Y. lipolytica and carbon‐neutral platforms like photosynthetic microalgae and Pichia pastoris [103] provides a diverse array of chassis, each offering distinct carbon‐partitioning advantages for diterpenoid production. Additionally, successfully navigating the complex regulatory frameworks and safety certifications governing the commercialization of genetically modified organism (GMO)‐derived fragrance molecules will be an unavoidable final hurdle for market entry. Overcoming these specific translational bottlenecks through the systematic integration of diversified host platforms, advanced design tools, integrated bioprocessing strategies, and proactive regulatory compliance is expected to advance the development of sustainable biomanufacturing for sclareol and increasingly complex natural terpenoids, facilitating broader access to high‐value molecules for the fragrance, pharmaceutical, and agrochemical industries.

Author Contributions

Yichen Li: investigation, visualization, writing – original draft. Lidan Ye: conceptualization, supervision, funding acquisition, writing – review and editing. Hongwei Yu: project administration, funding acquisition.

Funding

This work was supported by the National Natural Science Foundation of China (Grant Nos. 22478340 and 32471481), and the Fundamental Research Funds for the Central Universities (Grant No. 226‐2025‐00043).

Conflicts of Interest

The authors declare no conflicts of interest.

Declaration of Generative AI Use

During the preparation of this work the authors used DeepSeek and Gemini 3.1 Pro in order to polish the manuscript. After using this tool service, the authors reviewed and edited the content as needed and takes full responsibility for the content of the published article.

Data Availability Statement

No data was used for the research described in the article.