The parental strain of A. fumigatus employed in this study was A1160 and Af293 (FGSC). Another strain, A1160^C, which was labeled as WT in the text, was constructed by reintroducing the pyrG gene into A1160 (Jiang et al., 2014). Itraconazole-resistant strains R1 and R2 were mutated from Af293. For A. fumigatus cultivation, YAG (containing 2% glucose, trace elements, 0.5% yeast extract and 2% agar) and YUU (YAG+5 mM uridine, 10 mM uracil) were used. For colony growth tests, A. fumigatus strains were grown on YAG or YUU supplemented with the indicated reagent.

To evaluate CA’s antifungal activity, YAG plates containing 0, 30, 45, or 60 μg/mL CA were inoculated with 3 × 10⁴ A. fumigatus WT spores (A1160 background). After incubation at 37 °C for 2.5 d, colonies were photographed and their diameters measured. Experiments were performed in triplicate for each CA concentration, with growth inhibition assessed by calculating mean colony diameters (±SD).

To assess the antifungal activity of CA against A. fumigatus strain Af293 and its drug-resistant variants R1 and R2, 3 × 10⁴, 3 × 10³, and 3 × 10² spores were plated on YAG agar containing CA at concentrations of 0, 30 and 45 μg/mL, respectively. After 48-h incubation at 37 °C, colony formation was evaluated to determine growth inhibition.

Biomass quantification was conducted by introducing 5 × 10⁷ A. fumigatus WT spores into 100 mL YAG liquid medium containing 30, 40, or 60 μg/mL CA. Biomass was harvested after 20 h of cultivation using sterile gauze filtration and oven-dried at 80 °C to a constant weight. Each sample was assayed in triplicate to ensure statistical reliability.

To elucidate whether CA exhibits fungistatic or fungicidal activity, a quantitative viability assay was performed. Initially, 1 × 10⁸ A. fumigatus WT spores were inoculated into YAG liquid medium. The experimental groups were treated with 200 and 400 μg/mL of CA for 3 h, while the control group remained untreated. Post-incubation, samples were centrifuged at 8000 rpm for 3 min to pellet the spores and remove the CA-containing supernatant. The pelleted spores were then washed twice with sterile water to eliminate residual CA, followed by serial dilution for spread plating. Specifically, the experimental samples were diluted 10³-fold, and the control samples were diluted 10⁵-fold to ensure countable colonies. Each treatment group was replicated three times, and colony-forming units (CFUs) were enumerated to statistically assess the impact of CA on fungal viability.

For E-test susceptibility analysis, A. fumigatus WT spores (4 × 10⁶) were uniformly suspended in 20 mL of molten agar medium, achieving a final concentration of 2 × 10⁵ spores/mL. Following solidification, ITZ E-test strips were aseptically placed at the center of each plate. After a 24-h incubation at 37 °C, drug resistance was assessed by determining the inhibition zone.

For the minimum inhibitory concentration (MIC) assay, the procedure was performed according to the EUCAST DEFINITIVE DOCUMENT E. DEF 9.3. In brief, double-strength RPMI 2% G medium buffered with MOPS was prepared. Serial dilutions of cinnamaldehyde (CA) were prepared to achieve final concentrations of 80, 40, 20, 10 and 0 μg/mL in the test wells, with 1 × RPMI glucose medium serving as the negative control. Each well was inoculated with 2 × 10⁵ A. fumigatus WT spores/mL and incubated at 37 °C for 48 h. The MIC was defined as the lowest CA concentration that completely inhibited visible fungal growth after the incubation.

The ergosterol extraction protocol was performed as previously described (Song et al., 2016). Briefly, 5 × 10⁷ WT spores were cultured in 100 mL YAG liquid medium supplemented with or without 45 μg/mL CA at 37 °C with shaking (220 rpm) for 24 h. The harvested mycelia were washed with distilled water, lyophilized, and ground to a powder. For each sample, 100 mg of mycelial powder was saponified with 3 mL of 25% KOH alcoholic solution (methanol: ethanol, 3:2, v/v) at 85 °C for 1 h. The mixture was then extracted with 3 mL of n-pentane. The upper organic phase was collected, evaporated to dryness, and redissolved in 2 mL methanol. The solution was filtered (0.45 μm) before HPLC analysis using a C18 column with 100% methanol mobile phase (flow rate 1 mL/min) and detection at 282 nm.

To knock out pth1 in A. fumigatus, the method of homologous recombination was adopted in this study. Firstly, the fragments of approximately 1.5 kb upstream and downstream of the pth1 gene were amplified by primer pairs pth1 P1/3 and pth1 P4/6, respectively. Subsequently, the screening marker hph fragment was amplified using the primer pair hygF/R. Finally, the three fragments were homologously recombined into the vector pBARGPE1 at the ClaI cloning site using pEASY®-Basic Seamless Cloning and Assembly Kit (TransGen Biotech, Beijing). The fused fragment, amplified by the primer pair pth1 P2/5, was introduced into the protoplasts of A1160, yielding the pth1 transformants. Subsequently, verification of these transformants was carried out using the primer pair pth1 S1/2, which culminated in the successful acquisition of the pth1 deletion mutant.

For genetic complementation, the pth1 gene fragment, including its upstream and downstream flanking regions, was amplified using primers pth1com F and pth1com R. The PCR product was then cloned into the SpeI site of the pZero-uu vector, which carries the NcpyrG selectable marker. The recombinant plasmid was subsequently transformed into protoplasts of the Δpth1 knockout strain via polyethylene glycol (PEG)-mediated transformation. Transformants were selected on medium lacking uridine/uracil. The oligonucleotides used in this study are displayed in Supplementary Table 1.

Metabonomic experiments were performed by Shanghai Applied Protein Technology Co., Ltd. Briefly, the conidia of A. fumigatus of the WT strains were inoculated into YAG liquid medium and incubated on a rotary shaker at 220 rpm and 37 °C for 20 h. Six biological replicates were exposed to 75 μg/mL CA for 0, 90 and 180 min, respectively. The samples were then filtered through gauze, rinsed with deionized water, and promptly frozen in liquid nitrogen. For metabolite extraction, the samples were homogenized into a powdered form using liquid nitrogen. For each sample, 80 mg was added to 1 mL of a cold extraction solvent mixture of methanol/acetonitrile/H₂O (2:2:1, v/v/v), followed by thorough vortexing and incubation on ice for 20 min. Subsequently, the samples were centrifuged at 13,000 g for 20 min at 4 °C. The supernatant was collected and passed through a 96-well protein precipitation plate. The elution was collected and dried in a vacuum centrifuge at 4 °C. For LC–MS analysis, samples were redissolved in 100 μL of 1:1 (v/v) acetonitrile/water and analyzed using a Sciex TripleTOF 6,600 quadrupole time-of-flight mass spectrometer coupled to hydrophilic interaction chromatography (HILIC) via electrospray ionization (ESI). Chromatographic separation was performed on an ACQUITY UPLC BEH Amide column (2.1 mm × 100 mm, 1.7 μm; Waters, Ireland) using Solvent A (25 mM ammonium acetate and 25 mM ammonium hydroxide in water) and Solvent B (acetonitrile). The gradient profile was as follows: 85% B for 1 min, linearly reduced to 65% over 11 min, further reduced to 40% in 0.1 min and held for 4 min, then increased back to 85% in 0.1 min, followed by a 5-min re-equilibration period. Flow rate: 0.4 mL/min; column temperature: 25 °C; injection volume: 2 μL. The mass spectrometer was operated in both positive and negative ionization modes. Raw MS data (wiff.scan files) were converted to MzXML format using ProteoWizard MSConvert prior to import into the open-source XCMS software. For peak picking: centWave m/z = 25 ppm, peakwidth = c (10, 60), prefilter = c (10, 100). For peak grouping: bw = 5, mzwid = 0.025, minfrac = 0.5. Differentially expressed metabolites (DEMs) were identified using FC > 2 or <0.5, p < 0.05 and VIP > 1 as cutoffs.

Proteomic experiments were conducted by Shanghai Applied Protein Technology Co., Ltd. The strain cultivation and CA treatment procedures were identical to those of the aforementioned metabolomics treatment methods. A total of three biological replicates were performed for the proteomics analysis. Proteins were extracted using SDT buffer (4% SDS, 100 mM Tris–HCl, 1 mM DTT, pH 7.6) and quantified using the BCA Protein Assay Kit (Bio-Rad, USA). 20 μg of protein per sample were separated by SDS-PAGE, and protein bands were visualized with Coomassie Blue R-250 staining. SDS-PAGE analysis confirmed no evidence of protein degradation, and the protein yield was deemed sufficient for subsequent experiments, thus enabling the progression to formal assays. Subsequently, proteins were digested with trypsin. LC–MS analysis was carried out on a timsTOF Pro mass spectrometer (Bruker) coupled to a Nanoelute (Bruker Daltonics). Digested A. fumigatus peptides were loaded onto a reversed-phase trap column (Thermo Scientific Acclaim PepMap100, 100 μm × 2 cm, nanoViper C18) connected to a reversed-phase C18 analytical column (Thermo Scientific Easy Column, 10 cm × 75 μm, 3 μm C18 resin). The peptides were equilibrated in Buffer A (0.1% formic acid) and separated using a linear gradient of Buffer B (84% acetonitrile, 0.1% formic acid) at a flow rate of 300 nL/min, controlled by IntelliFlow technology. The MS raw data of each sample were analyzed using MaxQuant 1.6.14 software against the A. fumigatus UniProt database for identification and quantification. FDR was set to 1% at both peptide and protein levels. Differentially expressed proteins (DEPs) were identified using FC > 2 or <0.5 and p < 0.05 as cutoffs.

Fungal cultures were initiated by inoculating 5 × 10⁷ WT spores into YAG medium incubating for 18 h at 37 °C, 220 rpm. Experimental groups were treated with 75 μg/mL CA for either 15, 30 or 60 min, with three biological replicates per treatment condition. Mycelia were harvested by filtration through sterile gauze, washed three times with deionized water, blotted dry, and flash-frozen in liquid nitrogen. For RNA extraction, frozen mycelia were ground in liquid nitrogen, and 100 mg of mycelial powder was added to 1 mL TRIzol reagent. After vortexing and incubation for 5 min at room temperature, 200 μL chloroform was added, followed by vigorous vortexing for 30 s and an additional 5 min incubation. The mixture was centrifuged at 13,000 rpm for 10 min at 4 °C, and 400 μL of the aqueous phase was carefully transferred to a new RNase-free tube. RNA was precipitated by adding an equal volume of isopropanol, incubating for 5 min, and centrifuging at 13,000 rpm for 5 min. The resulting RNA pellet was washed with 75% ethanol, air-dried, and finally dissolved in 300 μL DEPC-treated water. RNA integrity was confirmed by agarose gel electrophoresis. For cDNA synthesis, 0.8 μg of total RNA was reverse transcribed, and the resulting cDNA was diluted fivefold for subsequent qPCR analysis.

Previous studies have demonstrated that CA possesses potent antifungal activity (Shreaz et al., 2016; OuYang et al., 2019; Long et al., 2024). In this study, we have tested its antifungal activity on the wild-type strain of A. fumigatus. Our antifungal assays demonstrated that cinnamaldehyde (CA) exhibits concentration-dependent anti-fungal activity against A. fumigatus. In the solid YAG medium supplemented with 30, 40, and 60 μg/mL CA, the mycelial growth of A. fumigatus was progressively inhibited. At 30 μg/mL, partial inhibition (23% reduction in colony diameter vs. control) was observed, while 45 μg/mL caused significant suppression (68% reduction). When the concentration was increased to 60 μg/mL, the growth of mycelium was completely inhibited (Figure 1A). Minimum inhibitory concentration (MIC) assays confirmed that the MIC of CA against the WT strain fell within the range of 40–80 μg/mL. Similarly, in the liquid medium, the biomass of A. fumigatus decreased in a dose-dependent manner with increasing concentration of CA (Figure 1B), which further confirmed the antifungal potential of CA. To validate whether CA exerts fungistatic or fungicidal effects, A. fumigatus spores were exposed to escalating concentrations of CA (200 and 400 μg/mL), followed by quantitative colony-forming unit (CFU) enumeration. Statistical analysis revealed a dose-dependent reduction in CFUs, relative to the untreated control (Figure 1C), strongly indicate that CA exhibits fungicidal activity against A. fumigatus. These data indicate that CA shows good antifungal activity against A. fumigatus.

Previous study has demonstrated that the antifungal effect of CA on F. sambucinum is achieved through the inhibition of ergosterol biosynthesis (Wei et al., 2020). To further investigate whether CA inhibits ergosterol biosynthesis in A. fumigatus, the intracellular ergosterol levels were analyzed in the presence or absence of CA. The results showed no significant difference in ergosterol levels between the CA-treated and untreated groups (Supplementary Figure S1). Since azole resistance in fungi, particularly resistance to ergosterol synthesis inhibition, poses a significant challenge, we aimed to test whether CA exhibits similar antifungal effects against azole-resistant strains of A. fumigatus. To evaluate this, we first tested the susceptibility of two itraconazole-resistant strains (R1 and R2) derived from A. fumigatus strain Af293 to itraconazole (ITZ). The MICs of R1 and R2 were 8 μg/mL each, which were significantly higher than the MIC of Af293 (1.5 μg/mL) (Figure 2A). Interestingly, the susceptibility of the two itraconazole-resistant strains (R1 and R2) to CA was similar to that of Af293 (Figure 2B). This indicating that CA might not rely on Cyp51A mediated ergosterol biosynthesis inhibition but also has an alternative mechanism of action with the same inhibitory effect on azole-resistant strains.

Although some of previous studies have explored the inhibition of fungi by CA, identifying further specific molecular mechanisms will undoubtedly support the development of therapeutic strategies to combat invasive fungal infections (OuYang et al., 2019; Wei et al., 2020; Wang et al., 2024). Here, we employed proteomic and metabolomic analyses to investigate the molecular effects of CA on A. fumigatus. The quality control was validated by metabolomic PCA analysis or protein ratio distribution analysis (Supplementary Data Sheet 1 and 2). Proteomic analysis revealed that 53 and 167 differentially expressed proteins (FC > 2 or <0.5, p < 0.05) were identified following treatment with CA for 90 min and 180 min, respectively (Figure 3A; Supplementary Table 2). Metabolomic analysis revealed that 145 metabolites showed significant alterations (fold change >2 or <0.5, p < 0.05, VIP > 1) after 90 min of CA treatment. Upon extending the treatment duration to 180 min, the number of differential metabolites increased to 350 (Figure 3B; Supplementary Table 3).

The TCA cycle is a fundamental metabolic pathway represented in most organisms, playing a crucial role in the oxidation of nutrients and the process of energy production. Malate dehydrogenase (MDH) is an essential enzyme in the TCA cycle that catalyzes the conversion of malate to oxaloacetate. CA has been shown to inhibit the activity of malate dehydrogenase in the TCA cycle of A. niger (Wang et al., 2024). Our metabolomic analysis demonstrated that CA treatment induced significant perturbations in tricarboxylic acid (TCA) cycle intermediates of A. fumigatus. Specifically, results showed a significant accumulation of malate and acetyl-CoA, while observed a significant decrease in alpha-ketoisovaleric acid following CA treatment, respectively (Figure 4). These metabolic alterations indicate that CA inhibits the growth of A. fumigatus by interfering with the tricarboxylic acid (TCA) cycle.

Protein group analysis demonstrated that CA treatment led to a significant down-regulation of proteins associated with protein metabolism. Specifically, under the condition of 180-min CA treatment, the putative translation initiation factor eIF4E3 (AFUB_051690), leucyl-tRNA synthetase LeuRS (AFUB_093380), prolyl-tRNA synthetase ProRS (AFUB_0101700), and peptidyl-tRNA hydrolase Pth1 (AFUB_053480) were down-regulated by 2.25, 2.32, 3.13, and 8.33-fold, respectively (Figure 5A). Further, we validated the expression of eIF4E3, leuRS, proRS, and pth1 at the mRNA level. RT-qPCR analysis revealed that the expression levels of eIF4E3, leuRS, and proRS decreased to varying degrees after 15, 30 or 60 min of CA treatment (Figure 5B). However, the transcription of pth1 exhibited only a slight decrease following 30 min of CA treatment; conversely, it showed an increase after 60 min of treatment (Figure 5B). Moreover, metabolomic analysis revealed a significant increase in the intracellular accumulation of various short peptides following CA treatment (Figure 5C), supporting the disruption of protein metabolism by CA.

Pth1 plays a crucial role in disengaging the nascent polypeptide chain from the tRNA molecule during the process of protein synthesis, which is conserved in both prokaryotes and eukaryotes (Mundra and Kabra, 2024). Although proteomic analysis revealed a significant reduction in Pth1 protein expression following CA treatment (Figure 5A), transcriptional validation of pth1 via RT-qPCR showed only a minor decrease in mRNA levels after 30 min of CA treatment (Figure 5B). Furthermore, pth1 transcription was increased after 60 min of CA treatment. To further investigate the function of Pth1 in A. fumigatus during its response to CA, we conducted gene knockout analyses of pth1using the A. fumigatus A1160 strain. Diagnostic PCR analysis showed that pth1 was completely replaced by the hygromycin resistance gene hyg, indicating successful knockout of the pth1 gene (Supplementary Figure S2). The empirical findings divulged that upon the knockout of pth1 significantly reduced the growth (Figure 6A), and increased its susceptibility to CA at the concentration of 30 μg/mL CA (Figure 6B). To validate that the growth defect phenotype of Δpth1 was indeed caused by pth1 deletion, the pth1 gene was complemented in the Δpth1 strain. The complemented strain exhibited growth patterns comparable to the WT, thus confirming that the growth defect in Δpth1 is attributed to loss of the pth1 gene (Figures 6A,B). These results suggest that pth1 is essential for the growth and CA resistance of A. fumigatus, and provide valuable insights for further investigations into the molecular mechanism underlying its antifungal effects.