Heterologous expression of AtLEC1 and AtLEC1-LIKE transcription factors redirects carbon flux toward lipid accumulation in diatom

Abstract

Background

Microalgal biodiesel is a key fossil fuel alternative, but enhancing lipid accumulation via single metabolic gene overexpression is often ineffective. Transcription factor engineering overcomes this by coordinating multiple metabolic pathways. To address the unexplored role of LEC1-type transcription factors in diatoms, we engineered the euryhaline and psychrotolerant biodiesel candidate diatom Phaeodactylum tricornutum through heterologous expression of the key plant lipid regulators AtLEC1 and AtL1L.

Results

Codon-optimized genes driven by the endogenous fcpA promoter were integrated into the nuclear genome, with regulators localization confirmed in the nucleus. Crucially, AtL1L transformants exhibited significant redirection of carbon flux from carbohydrates toward lipids, evidenced by lipid content increasing to 29.8%-33.9% of dry weight compared to 20.9% in wild-type controls while carbohydrates decreased to 13.3%-16.5% from 23.1%. AtL1L transformants accumulated 42–64% more neutral lipids and 48–68% higher total fatty acids without compromising biomass yield or photosynthetic efficiency (Fv/Fm). Molecular analyses revealed coordinated upregulation of key lipogenic, glycolytic and pyruvate metabolism genes such as acetyl-CoA carboxylase, pyruvate kinase and malic enzyme, which were corroborated by significant increases in corresponding enzyme activities and NADPH levels. Metabolite profiling confirmed accumulation of lipid precursors including acetyl-CoA (1.7-fold elevation) concurrent with reduction of sugars like glucose to less than 39% of wild-type levels.

Conclusions

This study demonstrates the first functional transfer of plant transcription factors to diatoms, providing a transformative strategy for high-productivity microalgal biodiesel.

Graphical abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s12934-025-02893-9.

Background

The ever-growing global energy demand, coupled with the depletion of fossil fuel reserves and their detrimental environmental impacts, necessitates the urgent development of sustainable alternatives [1]. Biodiesel derived from renewable biomass represents a crucial solution, with microalgae emerging as one of the most promising feedstocks due to their rapid growth, high photosynthetic efficiency, CO₂ consumption, and substantial capacity for intracellular lipid accumulation [2]. Compared to traditional oil crops, microalgae offer significant advantages by utilizing non-arable land and wastewater/seawater, avoiding competition with food production [3].

However, achieving high lipid productivity in microalgae remains challenging. While environmental stresses like nitrogen starvation can induce lipid accumulation, they invariably suppress cell growth and photosynthetic efficiency, ultimately limiting overall productivity [4, 5]. Advances in microalgal genomics and genetic tools have enabled both forward and reverse genetic approaches for strain improvement. While traditional mutagenesis (e.g., using UV or chemicals) has yielded lipid-overproducing mutants, it is often laborious and prone to reversion [6, 7]. Genetic engineering, particularly the targeted single key metabolic enzymes (e.g., acetyl-CoA carboxylase, glycerol-3-phosphate acyltransferase), has shown promise but frequently faces limitations due to cellular homeostasis and complex feedback regulation, resulting in suboptimal or negligible increases in lipid content [8, 9].

These limitations underscore the need for strategies that coordinately regulate entire metabolic pathways. Transcription factor (TF) engineering, which leverages the ability of TFs to modulate the expression of multiple genes within a network, has emerged as a powerful approach to overcome the bottlenecks of single metabolic gene manipulation [10]. This strategy has successfully enhanced the production of valuable metabolites, including lipids, in various organisms [11–13]. For instance, heterologous expression of the soybean (Glycine max) TF GmDof4 or manipulation of algal-specific TFs (e.g., NsbZIP1 in Nannochloropsis, PSR1 in Chlamydomonas) has demonstrated the potential to enhance microalgae lipid accumulation, sometimes even improving growth [14–17]. This highlights the potential of TF engineering for systemic metabolic rewiring.

Among the key regulators of plant seed oil biosynthesis, the NF-Y-type LEAFY COTYLEDON1 (LEC1) and its paralog LEC1-LIKE (L1L) stand out as master regulators coordinating fatty acid synthesis and storage [18]. They are crucial for embryogenesis and lipid accumulation in Arabidopsis thaliana and their overexpression significantly increases seed oil content in plants like Brassica napus [19, 20]. Strikingly, mammalian CCAAT-binding factors (homologous to NF-Y), directly regulate fatty acid synthase expression [21–23], suggesting deep evolutionary conservation in lipid metabolic regulation mediated by this TF family. Critically, despite BLASTX analysis identified approximately 20 putative LEC1 homologs in diatom genome sharing >60% amino acid similarity to AtLEC1 (Supplementary Figure S1), the functional role of LEC1-type TFs in these ecologically and industrially important microalgae remains completely unexplored.

Diatoms, responsible for approximately 20% of global carbon fixation [24], are prime candidates for biodiesel production due to their high lipid productivity and hypothesized contribution as major progenitor of petroleum deposits [25]. The model diatom Phaeodactylum tricornutum is particularly attractive as a biodiesel feedstock. It exhibits rapid growth, accumulates lipids up to 20–30% of its dry weight [26], possesses a fully sequenced genome and extensive genetic resources [27–30], and benefits from a well-developed molecular toolkit for genetic manipulation [31–33]. Its euryhaline and psychrotolerant nature further enhances its suitability for large-scale cultivation [34].

Therefore, this study aimed to bridge this significant knowledge gap and exploit the potential of LEC1-type regulators by heterologously expressing Arabidopsis AtLEC1 and AtL1L in P. tricornutum. We sought to determine whether these plant master regulators could enhance lipid biosynthesis in diatom, assess their impact on growth and photosynthesis, elucidate the molecular mechanisms underlying the observed phenotypes, and ultimately develop a transformative strategy for high-productivity microalgal biodiesel without the growth penalty associated with conventional stress induction.

Materials and methods

Total RNA extraction and cDNA synthesis in Arabidopsis Thaliana

Fresh Arabidopsis leaves were cut into small pieces and ground in liquid nitrogen. Total RNA was extracted using RNAiso Plus (TaKaRa, Cat. No. 9108). Residual genomic DNA was removed using Recombinant DNase I (TaKaRa, Cat. No. 2270 A). cDNA was synthesized using the PrimeScript™ II 1st Strand cDNA Synthesis Kit (TaKaRa, Cat. No. 6210 A).

Cloning of AtLEC1 and AtL1L genes

The coding sequences of AtLEC1 (AT1G21970) and AtL1L (AT5G47670) were retrieved from the Arabidopsis genome database (https://www.arabidopsis.org/). Primers were designed using Primer Premier 5.0 and listed in Table S1.

PCR amplification was performed using Q5^® High-Fidelity DNA Polymerase (NEB, Cat. No. M0494S). The PCR products were A-tailed using the DNA A-Tailing Kit (TaKaRa, Cat. No. 6109), purified with the TaKaRa MiniBEST DNA Fragment Purification Kit Ver.4.0 (TaKaRa, Cat. No. 9761), and ligated into the pMD19T cloning vector (TaKaRa, Cat. No. 6013). The constructs were transformed into DH5α competent cells (TaKaRa, Cat. No. 9057). After sequencing verification, plasmids were extracted using the TIANprep Mini Plasmid Kit (TIANGEN, Cat. No. DP103).

Codon optimization

The coding sequences of AtLEC1 and AtL1L genes from Arabidopsis thaliana were codon-optimized according to the nuclear genome codon usage bias of P. tricornutum [35]. The optimized gene sequences were subsequently synthesized commercially by Synbio Technologies (Suzhou, China).

Construction of AtLEC1 and AtL1L overexpression vectors

Using the synthetic gene plasmids as templates, the codon-optimized AtLEC1 and AtL1L open reading frames (ORFs) were amplified with Q5^® High-Fidelity DNA Polymerase (NEB, Cat. No. M0494S). KpnI restriction sites were introduced, and the stop codons were removed. The amplified DNA fragments were purified by gel extraction, digested with KpnI, and ligated into the N-terminus of the eGFP (enhanced Green Fluorescent Protein) gene in the pPha-T1-eGFP expression vector, generating the pPha-LEC1-eGFP and pPha-L1L-eGFP constructs. The recombinant plasmids were transformed into DH5α competent cells. Colonies harboring expression vectors with the correct insert orientation were screened using the vector-specific upstream primer Sp6 and gene-specific downstream primers. The primer sequences employed in this experimental phase were documented in ‌Table S2.

Transformation of P. tricornutum and screening of positive transformants

Biolistic transformation and transformants screening were performed as described by Falciatore et al. [36]. The primers designed for direct PCR screening were tabulated in Table S3.

Quantitative Real-Time PCR (qPCR) analysis

Total RNA from P. tricornutum was extracted, and RNA concentration was measured using a NanoDrop 2000 C spectrophotometer (Thermo Scientific). cDNA was synthesized using the PrimeScript™ RT Reagent Kit with gDNA Eraser (TaKaRa, Cat. No. RR047A). The relative expression levels of target genes in transformants were quantified using ACTIN as the internal reference. The primer sequences for qPCR were designed as listed in Table S4. SYBR Green fluorescence was detected using the FAM channel on a TP800 Thermal Cycler (TaKaRa). Relative gene expression was calculated using the 2^−△△CT method.

Subcellular localization

Fluorescence microscopy: Blue light excitation filter was selected to observe and capture the expression of exogenous gene fused with eGFP in algal cells using a BX53 fluorescence microscope (OLYMPUS).

Confocal laser scanning microscopy: Algal cells were pelleted via centrifugation and subjected to fixation in 3.7% formaldehyde for a duration of 30 min. Subsequent to the removal of the fixative, the cells were subjected to three washing cycles with 0.1 M phosphate-buffered saline (PBS, pH 7.4), with the supernatant being discarded after each wash. The cell pellet was then treated with 150 µl of DAPI stain (Beyotime, C1006) and incubated for 10 min at ambient temperature. Following the removal of the DAPI solution, three additional washes with PBS were performed. For microscopic observation, a 5 µl aliquot of the algal suspension was mounted onto a confocal dish, followed by the application of one drop of anti-fade mounting medium (Beyotime, P0126). An agar pad was subsequently applied to immobilize the specimen prior to imaging with a Leica TCS SP5 laser scanning confocal microscope.

Southern blotting

A digoxigenin (DIG)-labeled probe targeting the Shble gene was generated by PCR using specific primers (Shble-S-F: 5’ TCGAGTTCTGGACCGACCGGCT 3’; Shble-S-R: 5’ ACGAAGTGCACGCAGTTGCCGG 3’), DIG-dUTP/dNTP mix, and plasmid template, followed by purification with TaKaRa MiniBEST kit (Cat. No. 9761). Genomic DNA from P. tricornutum, extracted using the TIANGEN DP305 kit, was digested with HindIII at 37 °C for approximately 16 h. Digestion efficiency was verified by electrophoresis, followed by purification via phenol: chloroform. The digested DNA was separated overnight on a 0.7% agarose gel at 25 V under low-temperature conditions (4 °C). Gel DNA underwent denaturation, neutralization, equilibration in 2× SSC, and was transferred to a nylon membrane via upward capillary transfer for 20 h. The membrane was then baked at 80 °C for 2 h. Pre-hybridization occurred in Hyb-100 buffer at 37 °C for 2 h, followed by overnight hybridization at 37 °C with the denatured DIG-probe in Hyb-100 buffer. Post-hybridization washes consisted of two 5-minute washes with 2× SSC/0.1% SDS at room temperature and two 15-minute washes with 1× SSC/0.1% SDS at 65 °C. After blocking, the membrane was incubated with Anti-DIG-AP antibody diluted 1:5000, and signals were detected using CSPD chemiluminescent substrate.

Western blotting

Total protein was extracted using a plant protein extraction kit (Sangon, Cat. No. C510003). A 12% separating gel and 5% stacking gel were prepared. Protein samples were mixed with 5× loading buffer, denatured by boiling for 10 min, and separated by SDS-PAGE (90 V for stacking gel, 150 V for separating gel). Proteins were transferred to a PVDF membrane (0.45 μm pore size) using a Mini Trans-Blot^® Electrophoretic Transfer Cell (Bio-Rad) at 300 mA for 40 min. The membrane was blocked with 5% BSA-TBST for 60 min and incubated with primary antibodies (GFP, Beyotime AF0159 or ACTIN, Sangon D110007) at 4 °C overnight. After washing, HRP-conjugated secondary antibodies (1:5000 dilution in 5% NFDM-TBST) were applied for 40 min. Chemiluminescent signals were developed using ECL and subsequently captured.

Physiological and biochemistry parameters analysis of algal cells

Growth curve and chlorophyll fluorescence assays

Transformed and wild-type P. tricornutum cells in logarithmic growth phase were cultured in fresh f/2 medium at 20 ± 2 °C under 70 µmol photons/m²·s light intensity and a 12:12 light: dark cycle. Cultures were manually shaken three times daily. Growth curves were generated using a hemocytometer starting from day 5. Aliquots (2 mL) of algal cells were subjected to dark adaptation for 30 min, after which chlorophyll fluorescence parameters in P. tricornutum were measured using a Multicolor-PAM system (Walz, Germany).

Relative neutral lipid content assay

1.8 mL of algal cell suspension was mixed with 200 µL of Nile red stock solution (10 µg mL⁻¹) to achieve a final Nile red concentration of 1 µg mL⁻¹. The mixture was vortexed at 1400 rpm for 30 s. Fluorescence intensity was then measured using a spectrofluorometer (Cary Eclipse, Varian) with excitation (Ex) at 530 nm and emission (Em) at 585 nm. Both the Ex and Em slit widths were set to 10 nm. Nile fluorescence intensity was normalized by Fo, which was measured using the Multicolor-PAM system. Cells stained with Nile red were observed and imaged using a fluorescence microscope (BX53, Olympus) equipped with a green excitation filter.

Enzyme activity and NADPH content assays

Microalgal cells were harvested, flash-frozen in liquid nitrogen, and ground to a fine powder. The resulting powder was then thoroughly homogenized with ice-cold extraction buffer. Enzyme activities of malic enzyme (ME), pyruvate kinase (PK), pyruvate dehydrogenase (PDH), and phosphoglycerate kinase (PGK) were determined using specific assay kits from Suzhou Comin Biotechnology Co. (China). Acetyl-CoA carboxylase (ACC) activity was measured using a kit from Jianglai Biological Technology Co. (China). NADPH content was quantified using a kit from Nanjing Jiancheng Bioengineering Institute (China).

Analysis of cellular component content in algal cells

Biomass measurement

A 50 mL aliquot of algal suspension was centrifuged at 8000 × g for 15 min at 4 °C. Following supernatant removal, the pellet was washed twice with ice-cold PBS buffer. The resulting wet algal paste was transferred to a pre-weighed aluminum foil weighing boat (W₀) and lyophilized to constant weight. The total weight (W_t) was recorded, and the algal dry weight (DW) was calculated as DW = W_t - W₀.

Carbohydrate content measurement

Algal cells harvested from 10 mL of culture by centrifugation were resuspended in 1 mL of distilled water. To this resuspension, 1 mL of 5% (w/v) aqueous phenol solution was added, followed by the addition of 5 mL of concentrated sulfuric acid. The mixture was vigorously vortexed and then allowed to stand at room temperature for 30 min before measuring its absorbance at 485 nm using a spectrophotometer. A standard curve was generated using a series of D-glucose solutions, and the total carbohydrate content in the sample was calculated based on this curve.

Lipid content measurement

Algal cells harvested from 50 mL of culture by centrifugation were extracted with 15 mL of ice-cold chloroform/methanol mixture (2:1, v/v) by vortexing for 15 min. After centrifugation, the supernatant was collected. The pellet was subjected to a second extraction with fresh solvent, and the supernatants were combined. To the combined supernatant, 0.2 volumes of 0.88% KCl solution were added. The mixture was gently inverted for mixing and allowed to stand for 20 min to achieve phase separation. The lower chloroform phase was carefully transferred to a pre-weighed glass Petri dish. The solvent was evaporated under a stream of nitrogen at low temperature, and the residue was further dried under vacuum for 1 h to constant weight. The total lipid content was determined by calculating the weight difference of the Petri dish before and after lipid extraction.

Protein content measurement

Algal cells harvested from 10 mL of culture by centrifugation were resuspended in 1 mL of RIPA lysis buffer (Beyotime, Cat. No. P0045). Cell disruption was performed using an ultrasonic homogenizer on ice. The lysate was centrifuged at 12,000 × g for 20 min at 4 °C, and the supernatant was collected. The protein concentration in the supernatant was determined using a BCA protein assay kit (Beyotime, Cat. No. P0010S) according to the manufacturer’s instructions.

Isolation, purification, and structural identification of total soluble carbohydrate

Total soluble carbohydrate was extracted from the freeze-dried biomass of P. tricornutum using a modified warm water extraction method. Briefly, algal powder was suspended in deionized water at a solid-to-liquid ratio of 1:30 (w/v). The extraction was conducted at 50 °C for 30 min under constant agitation. The resulting slurry was centrifuged at 8,000 × g for 20 min at 4 °C to collect the supernatant. To remove proteins, the supernatant was treated with the Sevag reagent (chloroform: n-butanol = 4:1, v/v) repeatedly until no protein interface was visible. For additional purification, enzymatic treatment with DNase, RNase, and Proteinase K was performed sequentially to eliminate nucleic acids and residual proteins. Decolorization was then carried out using 2% (w/v) activated carbon at 60 °C for 30 min, followed by filtration. The carbohydrate was precipitated by adding four volumes of absolute ethanol and incubating at 4 °C overnight. The precipitate was recovered by centrifugation, redissolved in deionized water, and dialyzed against deionized water for 72 h. Following dialysis, the retentate was lyophilized and further purified by DEAE anion-exchange chromatography. The crude carbohydrate was dissolved in deionized water and loaded onto a DEAE column equilibrated with the same solvent. The column was eluted with a linear gradient of 0 to 1 M NaCl, and carbohydrate-rich fractions were monitored by the phenol-sulfuric acid method. The carbohydrate-containing fractions were pooled, dialyzed, and lyophilized to obtain the purified carbohydrate.

For subsequent NMR analysis, the lyophilized powder was subjected to deuterium exchange by dissolving in D₂O and lyophilizing again. The resulting material was finally dissolved in high-purity D₂O (99.96%) for acquisition of ¹H (600 MHz) and ¹³C (150 MHz) NMR spectra at 30 °C. For FTIR analysis, the purified sample was thoroughly ground with dry KBr powder and pressed into a transparent pellet. FTIR spectra were then acquired in the frequency range of 4000 to 400 cm⁻¹.

Analysis of total fatty acids in algal cells

Lyophilized algal cells (10 mg) were subjected to acid-catalyzed transesterification with 2 mL of 5% H₂SO₄ (v/v) in methanol containing 0.2% BHT (w/v) at 90–95 °C for 1.5 h in PTFE-lined sealed vials. After cooling, 2 mL saturated NaCl and 1 mL n-hexane were added, followed by vigorous vortexing for 1 min. The mixture was centrifuged at 4 °C, 3500 rpm for 5 min, and 60 µL of the n-hexane supernatant was collected for GC-FID analysis.

Metabolites analysis

Four biological replicates each of the AtL1L transformed line experimental group and the wild-type (WT) control group were processed. Samples were ground in liquid nitrogen, followed by the addition of 1 ml of methanol/acetonitrile/water solution (2:2:1, v/v). Samples were vortexed for 60 s, subjected to low-temperature ultrasonication (30 min, repeated twice), and incubated at − 20 °C for 1 h to precipitate proteins. After filtration, supernatants were collected by centrifugation at 14,000 g for 20 min at 4 °C, lyophilized, and stored. Sample separation was performed using an Agilent 1290 Infinity LC UHPLC system equipped with a HILIC column maintained at 25 °C. The mobile phase comprised solvent A (water containing 25 mM ammonium acetate and 25 mM ammonium hydroxide) and solvent B (acetonitrile). The column effluent was directed to an Agilent 6550 Q-TOF mass spectrometer for detection.

Results and discussion

Cloning and sequencing results of AtLEC1 and AtL1L‌

As shown in Figure S2A, PCR amplification of the AtLEC1 and AtL1L genes yielded fragments of 905 bp and 953 bp, respectively. A single band was amplified for each gene. Subsequent sequencing and alignment revealed that the full-length cDNA sequences of AtLEC1 and AtL1L were 717 bp and 705 bp, encoding proteins of 238 and 234 amino acids, respectively (Figure S2B, C), which matched the sequences in the database. BLASTP analysis of the amino acid sequences indicated that both proteins possess a CBFD_NFYB_HMF domain (Figure S2D, E).

Codon optimization results for AtLEC1 and AtL1L

Transcription and translation represent distinct regulatory stages influencing gene expression levels. Codon optimization primarily alters gene expression by modulating the efficiency of protein translation. Optimized genes exhibit higher expression levels and are more amenable to genetic manipulation. The optimization of AtLEC1 and AtL1L involved replacing rare codons for protein translation in P. tricornutum, such as ATA (Ile); AGG, AGA, CGG (Arg); TTA, CTA (Leu); TCA (Ser); GTA (Val); GGG (Gly), and others with usage frequencies below 10% (Figure S3A, Figure S4A).

The Codon Adaptation Index (CAI) represents the ratio of the relative usage frequency of codons in a target gene to the relative usage frequency if only the optimal codons were used. Optimal codons are typically the most frequently used synonymous codons for a given amino acid. For heterologous gene expression, CAI serves as a measure of the consistency between the codon usage of the target gene and that of the host cell. The CAI value ranges from 0 to 1. Highly expressed genes in humans and Escherichia coli typically exhibit CAI values between 0.75 and 1.0. We elevated the CAI of both AtLEC1 and AtL1L from approximately 0.5 to around 0.8. Additionally, we optimized the GC content profile to achieve a smoother curve and minimized stable hairpin structures (Figure S3B-E; Figure S4B-E). These optimizations enable more efficient translation by ribosomes, and the codon usage and distribution are now more consistent with the characteristics of P. tricornutum.

Construction of expression vectors for heterologous genes

The optimized heterologous genes were excised from the cloning vector pUC57 and subsequently ligated into the intermediate vector pPha-T1-eGFP. This process resulted in the successful construction of the final expression vectors pPha-LEC1-eGFP and pPha-L1L-eGFP (Figure S5, Figure S6). The heterologous genes were fused to the N-terminus of the eGFP gene. Expression of these fusion genes is driven by the endogenous fcpA promoter to ensure high efficiency. A Kozak sequence (GGTACC) was incorporated upstream of the initiation codon ATG of the heterologous genes to enhance translation efficiency in eukaryotic systems and prevent ribosome scanning leakage. Fusion with the eGFP gene facilitates the selection of transformants and enables subcellular localization studies of the target proteins.

Restriction enzyme digestion verification of Recombinant overexpression vectors and screening of positive transformants

Following successful construction of the recombinant expression vectors, plasmid DNA was extracted for each. As these vectors contain two KpnI restriction sites, digestion with KpnI alone yielded specific fragments of 717 bp and 705 bp for the pPha-LEC1-eGFP and pPha-L1L-eGFP vectors, respectively (Figure S7A, B). Both expression plasmids were subjected to sequencing, and subsequent sequence alignment confirmed the absence of any base mutations, verifying successful vector construction.

For the pPha-LEC1-eGFP and pPha-L1L-eGFP plasmids, multiple transformant colonies were randomly picked from each transformation group for direct PCR screening using algal cells. As shown in Figure S7C, all tested transformants contained a 298-bp fragment corresponding to the Shble zeocin resistance gene.

To confirm the integration of the target genes, PCR amplification was performed on both types of transformants using the universal vector primer Sp6 paired with gene-specific downstream primers. The results (Figure S7D) showed that transformants harboring the AtLEC1 gene produced a specific 794-bp fragment, while those harboring the AtL1L gene produced a specific 1114-bp fragment. This demonstrates the successful integration of the target genes into the P. tricornutum cells.

Expression of the heterologous transcription factors in the transformed algal strains was detected by qPCR. Based on this analysis, two strains exhibiting relatively higher expression levels for each gene (LEC1-2, LEC1-3; L1L-1, L1L-4) were selected for further investigation (Figure S7E).

Subcellular localization results of the transformants

As shown in Figure S8, fluorescence microscopy examination of the transformant cells revealed stable green fluorescence expression specifically localized to the nucleus under blue light excitation in all transformants harboring either the AtLEC1 or AtL1L gene. As shown in Fig. 1, the merged confocal microscopy image showed that the fluorescence from the nuclear dye DAPI overlapped with the green fluorescence from eGFP. Given that AtLEC1 and AtL1L are transcription factors in Arabidopsis thaliana, their nuclear localization within this heterologous expression system (P. tricornutum) suggest that they may also function as transcription factors in the diatom cells.

Fig. 1.

Confocal microscopy observations of different transformants. A L1L-1 transformant; B L1L-4 transformant; C LEC1-2 transformant; D LEC1-3 transformant. Red: chlorophyll autofluorescence, Blue: DAPI fluorescence, Green: eGFP fluorescence. Scale bar = 7.5 μm

Analysis of transformants: Southern blot, Western blot, chlorophyll fluorescence and physiological parameters

Following hybridization with digoxigenin-labeled PCR probe, a single-copy band corresponding to the expected size of 4806 bp was observed in the positive control plasmid lane. Transformant LEC1-2 harbored more than 5 copies of the exogenous gene, while transformants LEC1-3, L1L-1, and L1L-4 contained between 4 and 5 copies (Fig. 2A). The Southern blot results demonstrate successful integration of the heterologous genes into the nuclear genome of P. tricornutum.

Fig. 2.

Molecular characterization and physiological analysis of transformants. A Southern blot analysis of transformants. Lane M: DNA Molecular-Weight Marker II, DIG-labeled; Lane 1: Positive control plasmid pPha-T1-eGFP; Lane 2: Wild-type P. tricornutum; Lane 3: LEC1-2 transformant; Lane 4: LEC1-3 transformant; Lane 5: L1L-1 transformant; Lane 6: L1L-4 transformant. B Western blot analysis of transformants. Lane WT corresponds to wild-type P. tricornutum; Lane P represents the positive control transformed with the empty vector pPha-T1-eGFP. C Growth curve analysis of transformants. D Maximum photosynthetic efficiency (Fv/Fm) analysis of transformants. E Relative neutral lipid content analysis of transformants. Statistically significant differences versus wild-type (WT) are denoted as follows: *p < 0.05, **p < 0.01

Western blot analysis is presented in Fig. 2B. The empty vector control strain (pPha-T1-eGFP) displayed a specific band at approximately 27 kDa corresponding to the green fluorescent protein. In contrast, transformants LEC1-2, LEC1-3, L1L-1, and L1L-4 exhibited bands at approximately 53 kDa, consistent with the expected size of the fusion proteins from the heterologous genes and eGFP. Both wild-type (WT) and transformed algae expressed the 42 kDa β-actin internal control protein. These Western blot results confirm the translation and expression of the AtLEC1 and AtL1L transcription factors in P. tricornutum cells.

As shown in Fig. 2C, all of the transformants displayed comparable growth dynamics to wild-type algal cells throughout the cultivation period. Although the cell density of LEC1-2 and LEC1-3 transformants was slightly lower than that of the wild-type in the late growth phase (Day 20), no significant difference was observed in their maximum cell densities.

Figure 2D demonstrates that the maximum photochemical efficiency (Fv/Fm) of LEC1-2 and LEC1-3 transformants showed no significant difference from the wild-type control (p > 0.05). However, the L1L-1 and L1L-4 transformants exhibited significantly higher Fv/Fm values compared to wild-type algal cells during most of the cultivation period (p < 0.05).

All transformants exhibited a significant increase in neutral lipid content compared to the WT algal strain (Fig. 2E). Specifically, neutral lipid content increased by 24% and 15% in LEC1-2 and LEC1-3 transformants, respectively. More substantial increases of 64% and 42% were observed in L1L-1 and L1L-4 transformants, respectively.

Nile red staining of neutral lipids in algal cells

Fluorometric analysis revealed that the AtL1L transcription factor significantly increased neutral lipid accumulation in P. tricornutum (Fig. 2E). To validate this finding, we visualized intracellular lipid synthesis via fluorescence microscopy. As shown in Fig. 3, Nile Red-stained neutral lipids exhibited golden-yellow fluorescence upon blue-light excitation (450–490 nm). The transformants L1L-1 and L1L-4 exhibited significantly higher intensity of fluorescent lipid droplets compared to the wild-type control.

Fig. 3.

Nile red staining micrographs of neutral lipids in P. tricornutum. A1 L1L-1 transformant under fluorescence-field. A2 L1L-1 transformant under bright-field. B1 L1L-4 transformant under fluorescence-field. B2 L1L-4 transformant under bright-field. C1 Wild-type algal cells under fluorescence-field. C2 Wild-type algal cells under bright-field. Scale bar = 20 μm

Analysis of algal cellular composition

During the late logarithmic growth phase, we detected significant alterations in cellular carbon partitioning within the engineered diatoms compared to the wild type (Fig. 4). Specifically, transformants L1L-1 and L1L-4 exhibited a substantial increase in total lipid content, reaching 29.8% to 33.9% of dry cell weight, significantly higher than the wild-type level of 20.9% at p < 0.01. Concomitantly, total carbohydrate content markedly decreased to 13.3%–16.5% from the wild-type value of 23.1%, also significant at p < 0.01. In contrast, protein content showed a modest, statistically non-significant decrease, ranging from 30.3% to 30.6% versus the wild-type 33.4% at p > 0.05.

Fig. 4.

Allocation of cellular carbon to the main cellular components

The FTIR spectrum of the total soluble sugars extracted from wild-type algal cells (Fig. 5D) shows characteristic absorption peaks: the band at 3406 cm⁻¹ is attributed to O–H stretching vibration, the peak at 2923 cm⁻¹ corresponds to C–H stretching vibration, and the absorption at 1621 cm⁻¹ is assigned to the asymmetric stretching vibration of C = O [37]. The broad absorption between 1140 and 1043 cm⁻¹ arises from C–O–C stretching vibrations of glycosidic linkages in the sugar ring, indicating the presence of a pyranose form. The peak at 892 cm⁻¹ is characteristic of β-D-glycosidic bond vibration [38]. The FTIR spectrum of total soluble sugars from the AtL1L-transformed algae (Fig. 5A) is nearly identical to that of the wild-type.

Fig. 5.

Structural characterization of total soluble carbohydrate isolated from the AtL1L transformant and wild-type P. tricornutum. A Fourier-transform infrared spectroscopy of total soluble carbohydrate isolated from the AtL1L transformant; B ¹H spectra of nuclear magnetic resonance of total soluble carbohydrate isolated from the AtL1L transformant; C ¹³C spectra of nuclear magnetic resonance of total soluble carbohydrate isolated from the AtL1L transformant; D Fourier-transform infrared spectroscopy of total soluble carbohydrate isolated from wild-type P. tricornutum; E ¹H spectra of nuclear magnetic resonance of total soluble carbohydrate isolated from wild-type P. tricornutum; F ¹³C spectra of nuclear magnetic resonance of total soluble carbohydrate isolated from wild-type P. tricornutum. NA, not assigned

In the ¹H NMR spectrum of the total soluble sugars extracted from wild-type algal cells (Fig. 5E), the two anomeric proton singlets at δ 4.48 and 4.16 ppm confirm the presence of β-glycosidic linkages, corresponding to the H-1 of β-1,3- and β-1,6-linkages, respectively. This is consistent with the FT-IR results mentioned above. The ¹³C NMR spectrum (Fig. 5F) showed major signals at δ 102.7, 84.4, 76.2, 75.8, 73.4, 68.3, 67.7, and 60.9 ppm. Notably, the anomeric carbon signal at δ 102.7 ppm indicates a β-configuration for the glycosidic linkage. These results align with chrysolaminarin isolated from Tribonema utriculosum [39] and Tribonema aequale [40]. Therefore, the major component of the purified total soluble carbohydrate is unambiguously identified as chrysolaminarin, which is composed of pyranose glucose units with β-D-(1→3) (backbone) and β-D-(1→6) (branch) linkages. The ¹H and ¹³C NMR spectra of the total soluble carbohydrate from the AtL1L-transformant (Figs. 5B, C) were nearly identical to those of the wild-type, indicating that the heterologous expression of the AtL1L gene did not alter the primary composition of the major soluble carbohydrate in P tricornutum.

Analysis of total fatty acid content in algal cells

Figure S9A shows the gas chromatogram of a mixed standard containing 38 fatty acid methyl esters (FAMEs). The chromatogram demonstrates that all 38 FAME peaks are well-resolved with good peak shape, indicating suitability for quantitative analysis. Figure S9B presents a representative GC-FID chromatogram of an experimental P. tricornutum sample.

As presented in Fig. 6, fatty acids in P. tricornutum primarily consist of saturated fatty acids such as myristic acid (C14:0) and palmitic acid (C16:0) (Fig. 6B), as well as unsaturated fatty acids including monounsaturated fatty acid palmitoleic acid (C16:1n7) (Fig. 6C) and polyunsaturated fatty acid eicosapentaenoic acid (C20:5n3) (Fig. 6D). Overexpression of the AtL1L transcription factor significantly enhanced individual and total fatty acid content in engineered algal strains L1L-1 and L1L-4. For instance, myristic acid (C14:0) increased by 57.6% and 33.7%, palmitic acid (C16:0) increased by 75.8% and 46.6%, and the unsaturated fatty acid palmitoleic acid (C16:1n7) increased by 59.8% and 32.0%, eicosapentaenoic acid (C20:5n3) increased by 55.8% and 57.8%, respectively. Critically, total fatty acid content was significantly increased by 68.0% and 48.1% in transformed algal strains L1L-1 and L1L-4 (p < 0.01) (Fig. 6A). The observation that AtL1L overexpression broadly enhances lipid accumulation without substantially altering fatty acid composition profiles strongly suggests that AtL1L orchestrates lipid biosynthesis at the pathway level, promoting overall flux through the fatty acid synthesis and assembly pathways, rather than exerting fine-tuned regulation on individual fatty acid synthase genes.

Fig. 6.

Fatty acid composition analysis. A Total fatty acid content (µg/mg DW). B Saturated fatty acid compounds (µg/mg DW) including myristic acid (C14:0), pentadecanoic acid (C15:0), palmitic acid (C16:0), stearic acid (C18:0), arachidic acid (C20:0), behenic acid (C22:0) and lignoceric acid (C24:0). C Monounsaturated fatty acid compounds (µg/mg DW) including palmitoleic acid (C16:1n7), cis-10-heptadecenoic acid (C17:1n7), cis-9-oleic acid (C18:1n9) and nervonic acid (C24:1n9). D Polyunsaturated fatty acid compounds (µg/mg DW) including linoleic acid (C18:2n6), γ-linolenic acid (C18:3n6), α-linolenic acid (C18:3n3), cis-11,14-eicosadienoic acid (C20:2n6), arachidonic acid (C20:4n6), cis-5,8,11,14,17-eicosapentaenoic acid (C20:5n3), cis-13,16-docosadienoic acid (C22:2n6), cis-7,10,13,16,19-docosapentaenoic acid (C22:5n3) and cis-4,7,10,13,16,19-docosahexaenoic acid (C22:6n3). The values represent the mean ± SD of three independent experiments. DW, dry weight. WT, wild-type P. tricornutum. Statistically significant differences compared with WT are indicated by asterisks where * denotes p < 0.05 and ** denotes p < 0.01

AtL1L target candidates involved in lipid biosynthesis and carbohydrate metabolism

Microarray analysis by Mu et al. [19]. revealed that AtLEC1 (or AtL1L) overexpression in Arabidopsis upregulated 48 and 33 genes functionally annotated to lipid biosynthesis and carbohydrate metabolism, respectively. Subsequent qPCR validations by Tan et al.. in BnLEC1/BnL1L-expressing rapeseed (Brassica napus) [20] and Tang et al.. in AtLEC1-transformed peanut (Arachis hypogaea) [41] further demonstrated coordinated upregulation of lipogenic and glycolytic genes. To assess functional conservation in diatoms, we quantified transcript levels of AtL1L putative target genes in P. tricornutum via qPCR, using primers detailed in Table S4.

As quantified in Fig. 7 and Table S5, five genes in the chloroplast fatty acid biosynthesis pathway exhibited significant upregulation: acetyl-CoA carboxylase (ACC), malonyl-CoA: ACP transacylase (MCAT), 3-Ketoacyl-ACP synthase II (KAS II), long-chain acyl-CoA synthetase (ACSL) and stearoyl-CoA 9-desaturase (SCD). In plants, ACC functions as a multienzyme complex comprising biotin carboxylase, biotin carboxyl carrier protein, and carboxyltransferase. This complex catalyzes the committed step of de novo fatty acid biosynthesis by converting acetyl-CoA to malonyl-CoA. As the first and rate-limiting enzyme in this pathway, ACC plays a pivotal role in regulating carbon flux toward de novo lipogenesis [42]. Zhang et al. [43]. demonstrated that GmDof4-overexpressing Chlorella showed 46.4–52.9% higher lipid content with concomitant upregulation of seven ACC genes and enhanced enzymatic activity, confirming ACC’s pivotal role in lipid accumulation. ACSL activates C₁₂–C₂₀ free fatty acids into long-chain acyl-CoAs, which serve as essential substrates for triacylglycerol (TAG) biosynthesis in the endoplasmic reticulum. Significant upregulation of SCD gene, which encodes a Δ9-desaturase catalyzing the conversion of stearoyl-CoA to oleoyl-CoA, was observed. As shown in Fig. 6C, the oleic acid (C18:1n9) content in L1L-1 and L1L-4 reached 4.8- and 4.3-fold of wild-type level, respectively, indicating C18:1n9 as one of the most dramatically elevated fatty acids in transformants. This metabolic shift likely stems from the enhanced SCD expression.

Fig. 7.

Expression of lipogenesis and carbohydrate metabolism-related genes in transformant L1L-1 (grey), L1L-4 (black) and wild-type strain (white). Statistically significant differences versus wild-type were denoted as follows: *p < 0.05, **p < 0.01. A-E Fatty acid biosynthesis related genes including acetyl-CoA carboxylase (ACC), malonyl-CoA: ACP transacylase (MCAT), 3-oxoacyl-[acyl-carrier protein] synthase II (KAS II), long-chain acyl-CoA synthetase (ACSL) and stearoyl-CoA 9-desaturase (SCD); F-G Glycerolipid metabolism related genes including diacylglycerol O-acyltransferase (DGAT1) and glycerol‑3‑phosphate dehydrogenase (GPDH); H-K Glycolysis related genes including phosphofructokinase (PFK), phosphoglycerate kinase (PGK) and pyruvate kinase (PK2, PK3); L PMI, a mannose-6-phosphate isomerase gene; M-O Pyruvate metabolism related genes including pyruvate dehydrogenase E1 component (PDH), malic enzyme (ME) and fumarate hydratase (FUM); P LDP1, a lipid droplet-associated protein gene. All values are expressed as mean ± SD (n = 3). Asterisks denote statistically significant differences, with * indicating p < 0.05 and ** indicating p < 0.01

Significant upregulation was observed in two genes encoding diacylglycerol O-acyltransferase (DGAT1) and glycerol‑3‑phosphate dehydrogenase (GPDH) within the glycerolipid metabolism pathway. DGAT—a key enzyme mediating the terminal acylation step in TAG assembly—was substantiated in its metabolic role by Niu et al. [44], who achieved a 35% lipid increase in P. tricornutum through DGAT2 overexpression, and Dinamarca et al. [45], who reported doubled lipid content (100% increase) via DGAT2D expression. This collective evidence underscores DGAT’s capacity to channel carbon into TAG storage pools. In higher plants and algae, GPDH catalyzes the conversion of dihydroxyacetone phosphate, derived from the glycolytic pathway, to glycerol-3-phosphate, the essential backbone for lipid synthesis [46–48]. Supporting this metabolic role, P. tricornutum overexpressing GPDH showed 60% higher neutral lipid content and 6.8-fold increased glycerol production [49]. Thus, GPDH upregulation likely elevates glycerol-3-phosphate flux toward lipid assembly.

In the glycolytic pathway, four genes showed significant upregulation: phosphofructokinase (PFK), phosphoglycerate kinase (PGK), pyruvate kinase 2 (PK2), and pyruvate kinase 3 (PK3). PFK catalyzes the thermodynamically irreversible phosphorylation of fructose-6-phosphate to fructose-1,6-bisphosphate, while PK catalyzes the irreversible, ATP-generating step converting phosphoenolpyruvate to pyruvate, a key precursor for acetyl-CoA production. Both enzymes serve as major flux-controlling steps in glycolysis. PGK catalyzes the conversion of 1,3-bisphosphoglycerate to 3-phosphoglycerate with concomitant ATP generation, thereby fueling energy demands for lipid biosynthesis. Additionally, significant induction of mannose-6-phosphate isomerase (PMI) was detected. This enzyme isomerizes mannose-6-phosphate to fructose-6-phosphate, thereby channeling carbon flux into the glycolytic pathway via PFK-catalyzed formation of fructose-1,6-bisphosphate.

In the pyruvate metabolic pathway, three genes exhibited significant upregulation, including pyruvate dehydrogenase E1 component (PDH), malic enzyme (ME), and fumarate hydratase (FUM). ME catalyzes the irreversible oxidative decarboxylation of malate to generate pyruvate, NADH and CO₂. In multiple heterotrophic microorganisms, overexpression of ME increases cellular lipid content [50–52], whereas inhibition of ME activity significantly suppresses lipid metabolism [53], indicating a strong correlation between ME activity and lipid accumulation. Specifically, overexpression of this enzyme in P. tricornutum elevated total lipid content by 2.5-fold, reaching 57.8% of cellular dry weight [54]. FUM reversibly catalyzes the hydration of fumarate to malate, replenishing intracellular malate pools. Malate generated primarily in mitochondria likely enters chloroplasts via putative interorganellar shuttle systems to serve as a fatty acid precursor [55]. The catalytic component E1 of the pyruvate dehydrogenase complex was significantly upregulated. This multienzyme complex catalyzes the irreversible oxidative decarboxylation of pyruvate to acetyl-CoA, a critical precursor for fatty acid biosynthesis. Supporting its regulatory significance, Arabidopsis overexpressing Brassica napus PDK (pyruvate dehydrogenase kinase) exhibited reduced pyruvate dehydrogenase activity and diminished seed oil content [56]. Thus, enhanced expression of PDH components likely exerts positive control over algal lipid yield.

Furthermore, we quantified the expression of LDP1—a well characterized ‌oleosin gene in P. tricornutum [57] —via qPCR analysis. The engineered strains L1L-1 and L1L-4 exhibited 7.62-fold and 6.84-fold upregulation of LDP1 transcription, respectively. Oil bodies are phospholipid monolayer-coated organelles comprising a hydrophobic lipid core surrounded by oleosin-family proteins, primarily serving as cytoplasmic TAG reservoirs. This pronounced LDP1 induction correlates directly with the observed phenotypic changes in transformants: significantly increased oil body number and expanded total storage volume (Fig. 3), demonstrating its pivotal function in expanding cellular lipid storage capacity.

Enzyme activity validation and NADPH content analysis

To validate that the transcriptional upregulation of key metabolic genes observed in AtL1L transformants translated into enhanced catalytic capacity, we measured the activities of representative enzymes: acetyl-CoA carboxylase (ACC), phosphoglycerate kinase (PGK), pyruvate kinase (PK), pyruvate dehydrogenase E1 component (PDH), and malic enzyme (ME). Additionally, given the critical role of NADPH as a reducing power source required for multiple steps in de novo fatty acid biosynthesis, we quantified cellular NADPH levels. Strikingly, the measured enzymatic activities closely mirrored the transcriptional upregulation patterns of their corresponding genes (ACC, PGK, PK, PDH, ME) quantified by qPCR (Fig. 8A-E), demonstrating that overexpression of the AtL1L transcription factor not only elevated mRNA expression of key lipid synthesis, glycolytic and pyruvate metabolism genes but also robustly enhanced their functional enzymatic capacities in P. tricornutum.

Fig. 8.

Enzymatic activities of key metabolic enzymes and NADPH content in AtL1L transformants and wild-type P. tricornutum. A acetyl-CoA carboxylase (ACC) activity. B phosphoglycerate kinase (PGK) activity. C pyruvate kinase (PK) activity. D pyruvate dehydrogenase (PDH) E1 component activity. E malic enzyme (ME) activity. F Cellular NADPH content. Data are presented as mean ± SD from three independent experiments. Statistically significant differences compared to the wild-type control are indicated by asterisks (*p < 0.05, **p < 0.01)

Malic enzyme, a pivotal enzyme in this pathway, is categorized into NAD-dependent ME (NAD-ME: EC 1.1.1.38) and NADP-dependent ME (NADP-ME: EC 1.1.1.40) according to coenzyme specificity and substrate preference. Here, the ME gene (Phatr2_51970) was annotated as NADP-dependent, with WoLF PSORT predicting chloroplast localization. ME activity, catalyzing the decarboxylation of malate to generate pyruvate, CO₂ and NADPH, was significantly elevated in transformants (L1L-1: 2390.3, L1L-4: 2159.4 nmol NADPH/min/g DW) versus wild-type (1542.1 nmol NADPH/min/g DW), corresponding to 1.55-fold and 1.40-fold increases, respectively (Fig. 8E). Concomitantly, cellular NADPH content was significantly higher in AtL1L transformants (L1L-1: 274.7 µmol/g DW; L1L-4: 252.6 µmol/g DW) compared to the wild-type control (215.5 µmol/g DW) (Fig. 8F). Prior studies in diatoms demonstrated that NADP-ME overexpression in P. tricornutum boosts NADPH levels and drives cellular lipid accumulation [58, 59]. In this study, coordinated enhancement of NADP-ME activity and NADPH pool size likely contributes substantially to the dramatically enhanced lipid accumulation phenotype in the AtL1L transformants.

Results of differential metabolites analysis

Both the principal component analysis (PCA) score plot and orthogonal partial least squares-discriminant analysis (OPLS-DA) score plot demonstrate low intra-group variation and significant inter-group differences among the samples (Figure S10A, B). Permutation validation (n = 200 iterations) confirmed the reliability of OPLS-DA models, where R² and Q² values derived from randomly shuffled Y-matrix exhibited no overfitting (Figure S10C).

Comparative metabolites analysis uncovered a metabolic reprogramming signature in AtL1L-engineered strains (Table 1), characterized by concerted depletion of sugar pools—including glucose and mannobiose—alongside significant accumulation of organic acid intermediates such as pyruvate, malate and fumarate. This substrate reallocation phenomenon directly corroborates the reduced total carbohydrate content observed in Fig. 4. Concurrently, we detected significant enrichment of lipogenic precursors, with acetyl-CoA pools expanding by 69%, thereby providing abundant carbon flux for de novo lipogenesis. Notably, the cellular concentration of glycerol-3-phosphate—the essential glycerol backbone for TAG biogenesis—surged 1.63-fold, further substantiating enhanced neutral lipid production capacity. This coordinated metabolite shift illustrates channeling of carbon skeletons from carbohydrate pools toward lipogenic pathways. Furthermore, in the AtL1L transformed algal strains, the contents of major fatty acids in P. tricornutum—including myristic acid, palmitic acid, cis-9-palmitoleic acid, and eicosapentaenoic acid—increased by 1.48- to 1.64-fold compared to the wild-type, consistent with the observations in Fig. 6.

Table 1.

Differential metabolites involved in lipid biosynthesis and carbohydrate metabolism between AtL1L transformants and wild-type strains

Metabolites	Fold change	p
Glucose	0.61	<0.01
Mannobiose	0.67	<0.01
Glycerol-3-phosphate	1.63	<0.01
Pyruvate	1.76	<0.01
Malate	1.54	<0.01
Fumarate	1.42	<0.01
Acetyl-CoA	1.69	<0.01
Myristic acid	1.48	<0.01
Palmitic acid	1.64	<0.01
cis-9-Palmitoleic acid	1.47	<0.01
Eicosapentaenoic acid	1.57	<0.01

Codon optimization of heterologous genes

The significant impact of codon bias is well-established across all protein expression systems, where it can lead to differences in protein expression levels spanning several orders of magnitude. The essence of codon bias lies in the substantial variation in tRNA abundance for identical codons across different species, which consequently affects the translation efficiency of heterologous proteins. Therefore, the consideration of codon bias is imperative for the heterologous expression of genes from other species in P. tricornutum. Notably, distinct codon usage frequencies exist not only between different species but also between the nuclear genome and the chloroplast genome within the same species. The nuclear genome of P. tricornutum exhibits a GC content of 48%, while its chloroplast genome has a GC content of 33% [28, 60]. During the codon-optimization process for exogenous transcription factors AtLEC1 and AtL1L, all identified rare codons were strategically excluded from the synthetic nucleotide sequences. Multilayer molecular characterization via qPCR-based transcript profiling (Figure S7E), eGFP epifluorescence tracking (Fig. 1, Figure S8), and Southern/Western blotting detection (Fig. 2A, B) collectively confirmed robust and stable expression of these engineered transcription factors in P. tricornutum. Critically, serial subculturing beyond 10 generations under antibiotic selection-free conditions revealed undiminished exogenous gene expression, demonstrating exceptional genetic stability.

Promotion of lipid synthesis in P. tricornutum by AtLEC1 and AtL1L

In Arabidopsis thaliana and rapeseed (Brassica napus), certain studies indicate that overexpression of transcription factors LEC1 or L1L can enhance lipid accumulation in transformed plants, but concurrently causes severe growth inhibition [19, 20, 61] and even lethal effects in seedlings [62]. In this study, the expression of AtLEC1 and AtL1L in the diatom P. tricornutum exhibited minimal impact on algal growth (Fig. 2C). This observation aligns with Tang et al.‘s report that peanut (Arachis hypogaea) plants overexpressing LEC1 transcription factor developed normally without adverse effects on major agronomic traits [41, 63]. This apparent discrepancy likely stems from inherent differences between the recipient species and the strength of the promoters employed for exogenous gene expression. Consistent with unimpaired growth, transformants exhibited stable total protein quotas comparable to wild-type levels (Fig. 4). In addition, transformants L1L1-1 and L1L1-4 exhibited a significantly higher maximum photochemical efficiency (Fv/Fm) compared to wild-type controls (Fig. 2D). This enhancement may be attributed to the substantial accumulation of fatty acids potentially promoting photosynthetic activity in microalgae [64].

In this study, both the AtLEC1 and AtL1L transformed algal lines exhibited a significant increase in neutral lipid content compared to the wild-type P. tricornutum. Specifically, AtL1L-engineered strains achieved 42–64% higher neutral lipids (Fig. 2E) coupled with 48–68% increases in total fatty acid content (Fig. 6A). Photosynthetically fixed carbon allocation shifted significantly in AtL1L transformants, where the lipid proportion of total biomass rose from 20.9% to 29.8–33.9% while total carbohydrate content decreased from 23.1% to 13.3–16.5%, alongside a mild reduction in protein levels (Fig. 4). These compositional analyses indicated that overexpression of AtL1L restricts the carbon flux derived from photosynthetic fixation into carbohydrate synthesis. Instead, a greater proportion of the fixed carbon was siphoned towards lipid production.

The FTIR and NMR (¹H and ¹³C) spectroscopic analysis of the purified soluble sugar revealed no difference in the primary sugar type between the transformed and wild-type algal cells, which was specifically identified as chrysolaminarin. Chrysolaminarin primarily functions as a temporary carbon and energy reserve. During the light period, when photosynthetic carbon fixation exceeds immediate demands, diatoms synthesize and store it as chrysolaminarin; this reserve is then mobilized during darkness or periods of high energy demand. Consequently, reducing chrysolaminarin content is analogous to diminishing a readily accessible “savings account”. Redirecting carbon flux toward lipid (TAG) synthesis represents an alternative, yet effective, energy storage strategy. Although lipid mobilization might be less immediate than that of water-soluble chrysolaminarin, TAG remains a highly energy-dense molecule. Thus, from a macroscopic energy storage perspective, the total cellular energy reserve (lipids + remaining chrysolaminarin) may not be substantially reduced. Instead, a shift in the form of storage occurs, maintaining overall energy homeostasis and thereby supporting continued cell growth. Furthermore, an earlier study in P. tricornutum involving the suppression of UDP-glucose pyrophosphorylase (UGPase) gene expression demonstrated a significant reduction in chrysolaminarin content alongside enhanced cellular lipid synthesis [65]. The phenotype of the mutant in that study is similar to ours, exhibiting only a minor decrease in growth rate (maximum reduction of approximately 5%). This independent report corroborates that a reduction in chrysolaminarin content has a relatively limited overall impact on the growth of P. tricornutum.

Similar to these findings in Arabidopsis thaliana overexpressing AtLEC1 (AtL1L) [19], Brassica napus overexpressing BnLEC1 or BnL1L [20], Camelina overexpressing ZmLEC1 [66] and peanut overexpressing AtLEC1 [41], where numerous genes involved in lipid synthesis and carbohydrate metabolism were induced transcriptional activation, this study observed analogous results in AtL1L-overexpressing P. tricornutum (Fig. 7). These collective findings affirm that LEC1/L1L transcription factors as phylogenetically ancient master regulators of carbon rechanneling toward storage lipids, even across phylogenetically distant vascular plants and diatoms. Critically, this transcriptional change was functionally validated by our enzyme activity assays, which demonstrated concomitant and significant increases in the catalytic activities of ACC, PK, PDH, ME, and PGK in the AtL1L transformants (Fig. 8A-E). This establishes a direct link between AtL1L-mediated gene expression and enhanced enzyme activity.

Analysis of differential metabolites revealed significantly elevated levels of acetyl-CoA in transformants (Table 1), demonstrating that AtL1L-mediated lipid enhancement initiates the process from de novo fatty acid synthesis. This process involves coordinated upregulation of key regulatory enzymes—including the gatekeeping acetyl-CoA carboxylase (ACC) (Figs. 7A and 8A) and acyl carrier protein (ACP)- centered fatty acid synthase (KAS II) (Fig. 7C). Conserved regulatory logic of ACC was evidenced by direct AtLEC1/AtL1L-driven control in Arabidopsis thaliana and peanut [19, 41], suggesting that ACC is an evolutionarily conserved target of LEC1-type transcription factors. Another major category of genes orchestrated by the AtL1L transcription factor in P. tricornutum was predominantly enriched in glycolysis metabolic pathways. This included two key rate-limiting enzymes of glycolysis pathway: phosphofructokinase (PFK) and pyruvate kinase (PK). Notably, two distinct pyruvate kinase transcript variants were significantly upregulated (Fig. 7J, K). Conserved functional evidence from Arabidopsis demonstrates that PK loss-of-function mutants develop wrinkled seeds with 60% reduced oil content [67, 68], mechanistically linking PK activity to lipid deposition. Corroborating this regulatory axis, ‌oil-enriched AtL1L transformants displayed concurrent upregulation of pyruvate kinase gene expression and enzymatic activity (Fig. 8C), along with a 1.76-fold increase in its metabolic product pyruvate relative to wild-type strains (Table 1). Collectively, these results indicate that the PK gene likely represents another crucial target for AtL1L-mediated transcriptional regulation of lipid accumulation. Furthermore, the approximately 2000 bp sequences upstream of the start codon (ATG) of the upregulated genes involved in lipid synthesis and glycolysis were downloaded. These sequences were then scanned for the AtLEC1 or AtL1L transcription factor motif using the JASPAR database, with the similarity threshold set to a stringent 90% (compared to the default 80%). It was revealed that, with the exception of the mannose-6-phosphate isomerase (PMI) gene, all other upregulated genes contained potential AtLEC1 or AtL1L binding sites (see Appendix for the corresponding analysis). This finding provides direct bioinformatic evidence that these pathways are regulated by the AtLEC1/AtL1L transcription factors in P. tricornutum.

In addition, coordinated upregulation of NADP-ME activity and NADPH levels (Fig. 8E, F) provides crucial biochemical evidence for lipid boost. ME catalyzes the production of both pyruvate (a direct precursor for acetyl-CoA) and NADPH. The significantly elevated NADPH pool serves as the indispensable reducing equivalent for multiple reductive reactions in fatty acid biosynthesis, particularly those catalyzed by fatty acid synthase complexes. This dual role of ME in providing carbon skeletons and reducing power positions it as a pivotal node in AtL1L-driven metabolic rewiring towards lipid overproduction in P. tricornutum.

Analysis of differential metabolites revealed that the transformants of AtL1L exhibited a 33%-39% decrease in the content of saccharides such as glucose and mannobiose. Conversely, the levels of precursor substances for lipid synthesis, including pyruvate, malate, acetyl-CoA and glycerol-3-phosphate, were increased by 54%-76% (Table 1). This metabolic shift—coupled with transcriptional induction of glycolytic and lipogenic genes, reduced carbohydrate quotas, and enhanced lipid stores—establishes that AtL1L redirects photosynthetic carbon partitioning from carbohydrate biosynthesis toward fatty acid assembly. Concurrently, a portion of the cellular carbohydrates enters the glycolytic pathway, generating substantial amounts of lipid precursors (such as malate and pyruvate). These precursors are subsequently channeled through de novo fatty acid synthesis and Kennedy pathway to produce abundant TAG (Fig. 9).

Fig. 9.

Putative model illustrating TAG accumulation in AtL1L transformants of P. tricornutum. Up-regulated genes or metabolites are indicated in red font, while down-regulated metabolites are shown in blue. ACC acetyl-CoA carboxylase, ACSL long-chain acyl-CoA synthetase, DGAT1 diacylglycerol O-acyltransferase, FUM fumarate hydratase, GPDH glycerol‑3‑phosphate dehydrogenase, KAS II 3-oxoacyl-[acyl-carrier protein] synthase II, LDP1 lipid droplet-associated protein, MCAT malonyl-CoA: ACP transacylase, ME malic enzyme, PDH pyruvate dehydrogenase E1 component, PFK phosphofructokinase, PGK phosphoglycerate kinase, PMI mannose-6-phosphate isomerase, PK2, PK3 pyruvate kinase, SCD stearoyl-CoA 9-desaturase. (Created with BioRender.com)

Overexpression of the ZmWRI1 transcription factor in maize (Zea mays) increased seed oil content by 46% without altering protein levels, but reduced starch accumulation by approximately 60% compared to wild-type controls [61]. This suggests that ZmWRI1 likely enhances oil biosynthesis by redirecting carbon flux from carbohydrate biosynthesis toward lipid production. Crucially, the AtL1L transcription factor in this study induced analogous metabolic phenotypes to ZmWRI1, characterized by suppressed carbohydrate synthesis and elevated lipid production. Previous studies have demonstrated that the WRI1 transcription factor is a downstream regulatory target of LEC1-class transcription factors [18, 19]. However, whether a functional ortholog of WRI1 exists in diatom P. tricornutum remains to be elucidated and warrants further investigation.

Finally, a comprehensive bioinformatic analysis of the twenty potential LEC1 homologs in P. tricornutum was performed (See Appendix for JGI hit sequence analysis). Subsequent analysis revealed that the first 18 sequences are all located within the same region on chromosome 1 (2504797–2505521, (-) strand), indicating their representation of a single gene differing only in exon splicing junctions. The remaining two sequences were both located on chromosome 2 (1304551–1305504, (-) strand), confirming the presence of substantial sequence redundancy in the JGI database. In future work, focus will be placed on these two newly identified endogenous genes to determine their precise transcript sequences and validate their functions in P. tricornutum.

Conclusions

Heterologous expression of Arabidopsis thaliana transcription factors AtLEC1/AtL1L in P. tricornutum significantly enhanced lipid production while simultaneously elucidating LEC1-type transcription factors regulatory mechanisms in diatoms. Key achievements include: (1) Establishment of an efficient expression system via codon optimization (CAI 0.5→0.8), confirmed by nuclear localization and stable protein expression; (2) Achieving lipid synthesis without compromising growth, with AtL1L transformants exhibiting 42–64% increased neutral lipid accumulation while maintaining maximum cell densities and elevated maximum photochemical efficiency (Fv/Fm) comparable to the wild type. This result provides an effective strategy to circumvent the traditional dilemma of “high-lipid but low-growth” typically induced by nitrogen stress; (3) Systematic elucidation of the mechanism by which AtL1L redirects carbon flux through dual regulatory actions. It promotes the synthesis of lipid precursors (pyruvate, acetyl-CoA) and reducing power NADPH by activating glycolysis (upregulation of rate-limiting enzymes including PFK, PK) and pyruvate metabolism (upregulation of PDH, ME). Concurrently, it enhances de novo fatty acid synthesis (upregulation of ACC, KAS II) and TAG assembly (upregulation of DGAT1), ultimately leading to significant TAG accumulation. This study provides fundamental insights into diatom lipid regulation and establishes a transformative platform for renewable energy microalgal biotechnology, with the engineered strain combining substantially improved lipid productivity with inherent salt tolerance and low-temperature adaptability for promising application in coastal arid bioenergy production.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

YC and HH conceived the research project. YC, LG, ZH, ND and JD performed experiments and data analysis. YC and LG drafted the manuscript. LZ and HW revised the manuscript. YC and HW acquired the financial support. All authors read and approved the final manuscript.

Funding

This work was supported by the Liaoning Provincial Science and Technology Joint Funds Program (Grant No. 2023-BSBA-145) and the Basic Scientific Research Project of Higher Education Institutions in Liaoning Province (Grant No. LJ212510160003).

Data availability

Data is provided within the manuscript and supplementary information files.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

All authors approved the manuscript.

Competing interests

The authors declare no competing interests.