Exploring the cellular and ultrastructural impact of Cymbopogon martinii essential oil against postharvest fungal pathogens of papaya

Abstract

Postharvest fungal infections significantly reduce the quality and marketability of papaya fruit, leading to substantial economic losses. Plant-derived essential oils (EOs) have gained attention as eco-friendly alternatives to synthetic fungicides due to their bioactivity and biodegradability. In this study, the antifungal potential of Cymbopogon martinii essential oil (CMEO) was evaluated against two major postharvest pathogens, Colletotrichum gloeosporioides and Fusarium longifundum. Gas chromatography–mass spectrometry (GC–MS) analysis identified 51 compounds representing 98.06% of the total oil, predominantly oxygenated monoterpenes. CMEO exhibited strong concentration-dependent antifungal activity, achieving complete mycelial growth inhibition at 1500 µg/mL for C. gloeosporioides and 1200 µg/mL for F. longifundum. Although its efficacy was lower than that of Bavistin fungicide, CMEO demonstrated significant antifungal potential. Mechanistic investigations revealed disruption of fungal membrane integrity, evidenced by reduced ergosterol content, increased leakage of cellular components (OD₂₆₀), extracellular acidification, and severe morphological and ultrastructural damage observed through scanning and transmission electron microscopy. Overall, this study highlights the potential of CMEO as a natural antifungal agent and provides mechanistic insights supporting its application for sustainable postharvest disease management of papaya.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-026-51293-5.

Introduction

Papaya (Carica papaya L.) stands out as one of the most economically vital tropical fruits worldwide, largely due to its rich nutritional value and numerous medicinal benefits. However, its postharvest quality is severely compromised by fungal infections, resulting in substantial economic losses and reduced shelf life^1,2. Among the major pathogens, species of Colletotrichum and Fusarium are predominant causes of postharvest diseases such as Anthracnose and Fusarium rot in papaya fruits^3,4.

Traditionally, synthetic fungicides have been widely used to control these infections. Despite their effectiveness, their extensive use has led to serious concerns, including the development of resistant fungal strains, residual toxicity on fruits, and adverse environmental impacts. Consequently, there is an increasing demand for safer, sustainable, and eco-friendly alternatives for postharvest disease management. In this context, plant-derived natural products, particularly essential oils (EOs), have emerged as promising candidates^5–7.

Essential oils are volatile secondary metabolites obtained from aromatic plants and are well known for their broad-spectrum antimicrobial, antioxidant, and bio-preservative properties⁸. Among these, Cymbopogon martinii (Palmarosa), a commercially important aromatic grass native to India, has attracted attention for its high content of bioactive constituents. Its major components, including geraniol, geranyl acetate, linalool, and limonene, are reported to exhibit strong antifungal activity against a wide range of phytopathogens^9,10.

Although several studies have reported the antifungal efficacy of Cymbopogon martinii essential oil (CMEO), its specific mechanism of action against papaya-associated postharvest pathogens remains insufficiently explored^11,12. Essential oils generally exert antifungal effects by targeting multiple cellular and physiological processes. These include disruption of membrane integrity, alteration of permeability, interference with enzymatic activity, and induction of oxidative stress in fungal cells^13–16.

Geraniol, a key component of CMEO, has been shown to interact with fungal cell membranes by altering lipid bilayer structure, leading to increased permeability and leakage of intracellular constituents^17,18. Additionally, essential oils can induce pronounced morphological and ultrastructural alterations, such as cytoplasmic disintegration, cell wall deformation, and damage to organelles, including mitochondria and the endoplasmic reticulum^13,19. However, these mechanisms have not been comprehensively investigated in the context of Colletotrichum gloeosporioides and Fusarium longifundum, the primary pathogens responsible for postharvest deterioration of papaya.

Therefore, the present study aims to evaluate the antifungal efficacy of C. martinii essential oil against these major postharvest pathogens and to elucidate its underlying mechanisms of action. Particular emphasis is placed on assessing membrane integrity, ergosterol depletion, leakage of cellular components, extracellular pH changes, and structural as well as ultrastructural alterations using SEM and TEM analyzes. This study provides novel mechanistic insights into CMEO activity and highlights its potential as a sustainable alternative to synthetic fungicides for postharvest disease management in papaya.

Results

Chemical composition

GC‒MS analysis of the CMEO led to the identification of 51 compounds, 98.06% of the total identified oil composition. The dominant compounds included cis-p-mentha-1(7),8-dien-2-ol (15.7%), trans-p-mentha-1(7),8-dien-2-ol (10.39%), and β-ocimene (7.62%). Other key compounds included trans-piperitol, (1 S)-trans-isopiperitenol, and carvone. These compounds may contribute to the oil’s aroma and bioactivity. A small fraction of sesquiterpenes and miscellaneous compounds were also identified. The essential oil yield was 5.33 ± 0.25 mL/kg (0.53%, w/w), which may vary depending on plant developmental stage and collection conditions. The compound identities were confirmed by their retention indices (RI) and retention times (RT) against standard libraries, as shown in Fig. 1; Table 1.

Fig. 1.

GC‒MS chromatogram of Cymbopogon martinii essential oil showing identified chemical constituents with their corresponding retention times and relative percentage areas.

Table 1.

GC‒MS analysis of Cymbopogon martinii essential oil showing identified compounds, CAS numbers, molecular weights, molecular formulas, retention indices, retention time (RT), and relative percentage area.

S. No.	Name of Compounds	CAS NO.	Molecular Weight	Molecular Formula	RIa	RIb	RT	Area (%)
1.	cis-p-Mentha-1(7),8-dien-2-ol	14055-33-0	202.31	C₁₄H₂₂O₂	1231	1232	7.85	15.70 ± 1.57
2.	trans-p-Mentha-1(7),8-dien-2-ol	20822-58-8	136.22	C₁₀H₁₆O	1185	1187	7.2	10.39 ± 1.04
3.	β-Ocimene	3338-55-4	136.23	C₁₀H₁₆	1043	1045	5.85	7.62 ± 0.76
4.	trans-Piperitol	3729-83-4	152.25	C₁₀H₁₆O	1205	1207	5.1	7.00 ± 0.70
5.	(1 S)-trans-Isopiperitenol	13248-42-5	152.25	C₁₀H₁₆O	1210	1212	7.41	6.40 ± 0.64
6.	cis-Piperitol	3729-82-3	152.25	C₁₀H₁₆O	1197	1199	7.05	5.26 ± 0.53
7.	1,3,8-p-Menthatriene	634-67-4	136.22	C₁₀H₁₄	1078	1079	7.7	5.16 ± 0.52
8.	Carvone	60-33-3	218.34	C₁₈H₃₂O₂	1243	1245	8.27	4.64 ± 0.46
9.	Carveol	97-63-3	152.25	C₁₀H₁₆O	1203	1204	7.25	4.41 ± 0.44
10.	Linalool oxide	120-81-0	152.25	C₁₀H₁₆O	1102	1103	5.06	3.30 ± 0.33
11.	D-Limonene	138-86-3	136.24	C₁₀H₁₆	1024	1025	5.31	3.05 ± 0.30
12.	cis-Dihydrocarvone	15058-73-5	152.25	C₁₀H₁₄O	1220	1223	8.00	2.77 ± 0.28
13.	trans-Carveyl acetate	67435-59-4	176.27	C₁₃H₂₀	1238	1239	8.18	2.68 ± 0.27
14.	β-Myrcene	123-35-3	136.22	C₁₀H₁₆	990	992	8.03	2.64 ± 0.26
15.	Linalool	78-70-6	154.25	C₁₀H₁₈O	1098	1100	7.57	2.53 ± 0.25
16.	cis-Verbenone	88-86-5	152.25	C₁₀H₁₄O	1218	1218	8.77	1.81 ± 0.18
17.	Pulegone	89-82-8	152.25	C₁₀H₁₆O	1201	1202	8.42	1.04 ± 0.10
18.	Oxycymol	499-75-2	150.22	C₁₀H₁₄O	1298	1297	9.01	0.65 ± 0.07
19.	Cyclohexene, 4-isopropenyl-1-methoxymethoxymethyl-	51115-67-4	196.29	C₁₂H₂₀O₂	1227	1228	8.89	0.65 ± 0.07
20.	trans-Decalin	87-46-4	148.25	C₁₀H₁₈	973	972	8.96	0.62 ± 0.06
21.	Terpineol	98-55-5	154.25	C₁₀H₁₈O	1177	1180	7.93	0.60 ± 0.06
22.	(S)-(-)-Perillyl alcohol	1408-95-3	136.22	C₁₀H₁₆O	1297	1299	7.36	0.57 ± 0.06
23.	Limonene oxide	75-64-7	136.22	C₁₀H₁₆O	1123	1123	7.34	0.54 ± 0.05
24.	Methyl linoleate	97-53-0	164.2	C₁₀H₁₂O₂	2086	2087	8.39	0.48 ± 0.05
25.	Norbornane	498-66-8*	96.17	C₇H₁₂	756	758	6.78	0.48 ± 0.05
26.	Eugenol	102-99-4	136.22	C₁₀H₁₆O	1362	1364	8.32	0.40 ± 0.04
27.	Ethanone, 1-(3-methylphenyl)-	123-91-1	136.21	C₉H₁₀O	1157	1156	8.15	0.36 ± 0.04
28.	1,6-Dihydrocarveol	10428-17-5	152.25	C₁₀H₁₆O	1204	1203	7.96	0.35 ± 0.03
29.	cis-p-Mentha-2,8-dien-1-ol	13224-60-6	152.25	C₁₀H₁₆O	1138	1136	7.29	0.35 ± 0.03
30.	γ-Terpinene	99-85-4	136.24	C₁₀H₁₆	1054	1055	7.66	0.32 ± 0.03
31.	o-Cymene	95-63-6	134.22	C₉H₁₂	1021	1023	7.11	0.31 ± 0.03
32.	Humulene	6753-98-6	204.35	C₁₅H₂₄	1467	1469	8.53	0.30 ± 0.03
33.	Carvomenthenal	1195-33-8	152.25	C₁₀H₁₄O	1232	1233	8.11	0.28 ± 0.03
34.	L-Perillaldehyde	495-75-8	136.22	C₁₀H₁₆O	1237	1236	9.17	0.26 ± 0.03
35.	6,9,12,15-Docosatetraenoic acid, methyl ester	108-55-5	342.52	C₂₂H₃₄O₂	2234	2235	17.64	0.25 ± 0.03
36.	Ageratriol	15401-90-2	228.35	C₁₅H₂₄O	1513	1514	14.75	0.25 ± 0.03
37.	Myrtenol	86-02-8	136.22	C₁₀H₁₄O	1192	1194	8.69	0.25 ± 0.03
38.	Menthol	2216-51-5	156.27	C₁₀H₂₀O	1171	1174	7.5	0.24 ± 0.02
39.	β-Cyclocitral	93-83-6	136.21	C₁₀H₁₄O	1152	1153	14.44	0.23 ± 0.02
40.	p-Cymene	99-87-7	134.22	C₉H₁₂	1015	1017	6.59	0.22 ± 0.02
41.	Styrene	100-42-5	104.15	C₈H₈	898	897	9.07	0.18 ± 0.02
42.	Isoledene	593-84-0	204.36	C₁₅H₂₄	1377	1377	6.47	0.16 ± 0.02
43.	Isopinocarveol	139-10-3	152.25	C₁₀H₁₄O	1150	1149	8.46	0.11 ± 0.01
44.	10,13-Octadecadiynoic acid, methyl ester	2184-43-5	296.42	C₂₀H₂₆O₂	2085	2086	14.01	0.10 ± 0.01
45.	Methyl 8,10-octadecadiynoate	2184-39-2	296.42	C₂₀H₂₆O₂	1967	1969	9.27	0.10 ± 0.01
46.	Piperitone	88-73-1	152.25	C₁₀H₁₆O	1233	1235	9.39	0.09 ± 0.01
47.	trans-Verbenol	15596-27-9	152.25	C₁₀H₁₆O	1166	1169	6.79	0.09 ± 0.01
48.	Camphene	78-81-4	136.23	C₁₀H₁₆	928	931	3.09	0.07 ± 0.01
49.	Bergamotol, Z-a-trans-	88034-74-6	220.35	C₁₅H₂₄O	1693	1694	9.57	0.06 ± 0.01
50.	Myrtenal	86-02-8	136.22	C₁₀H₁₄O	1170	1172	6.73	0.05 ± 0.01
51.	2,5-Octadecadiynoic acid, methyl ester	20822-59-9	136.22	C₁₀H₁₆O	2078	2076	8.61	0.03 ± 0.01
Total compound identified (%)		98.06 ± 2.55
Yield (mL/kg)		5.33 ± 0.25
Grouped compounds (%)
Oxygenated monoterpenes		77.9 ± 2.03
Monoterpene hydrocarbons		14.77 ± 0.38
Miscellaneous compounds		4.38 ± 0.11
Oxygenated sesquiterpenes		0.55 ± 0.01
Sesquiterpene hydrocarbons		0.46 ± 0.01

a: Literature Retention index (Adams, 2007); b: Calculated retention indices (DB-5 column).

Fungal isolation and molecular identification

Two predominant postharvest fungal pathogens from naturally infected papaya fruits were isolated: Colletotrichum gloeosporioides and Fusarium longifundum. These were chosen from several morphologically distinct fungal isolates initially recovered during the isolation process. C. gloeosporioides formed fast-growing gray-white colonies with orange conidial masses (Supplementary Fig. S1a, b), while F. longifundum produced cottony white colonies with pink pigmentation (Supplementary Fig. S1c, d). Microscopic examination showed that C. gloeosporioides produced straight conidia, whereas F. longifundum formed sickle-shaped septate macroconidia. BLAST analysis of ITS-PCR products confirmed 99–100% similarity with the respective reference sequences (NCBI database, accessed on [10-04-2025]). The sequences were deposited in GenBank under accession numbers PP594940 (F. longifundum) and PP594941 (C. gloeosporioides).

Antifungal assay

The antifungal activity of CMEO against C. gloeosporioides and F. longifundum was evaluated based on percentage mycelial growth inhibition, along with MIC and MFC (Fig. 2, Supplementary Figs. S2‒S3 and Tables 2‒4). CMEO exhibited clear, concentration-dependent inhibitory effects against both fungi. CMEO achieved 100% inhibition of F. longifundum at 1200 µg/mL, whereas Bavistin produced complete inhibition at a lower concentration of 300 µg/mL (Fig. 2a and Supplementary Fig. S2a). For C. gloeosporioides, inhibition progressively increased with concentration and reached complete (100%) growth suppression at 1500 µg/mL, comparable to Bavistin at 600 µg/mL (Supplementary Fig. S2a). For C. gloeosporioides, inhibition progressively increased with concentration and reached complete (100%) growth suppression at 1500 µg/mL, comparable to Bavistin at 600 µg/mL (Fig. 2b and Supplementary Fig. S2b).

Fig. 2.

Antifungal activity of Cymbopogon martinii essential oil (CMEO). Effect of CMEO and positive control (Bavistin) on mycelial growth inhibition of (a) Fusarium longifundum, (b) Colletotrichum gloeosporioides, and (c) Minimum inhibitory concentration (MIC) and Minimum fungicidal concentration (MFC) of CMEO and positive control against both fungi. Different letters indicate significant differences based on Tukey’s test (p < 0.05).

Table 2.

Effect of Cymbopogon martinii essential oil (CMEO) and positive control (Bavistin) on mycelial growth and growth inhibition of Fusarium longifundum at different concentrations.

CMEO Concentration (µg/mL)	Mycelial Growth (mm)	Growth Inhibition (%)	Positive Control (µg/mL)	Mycelial Growth (mm)	Growth Inhibition (%)
0	66.89 ± 1.34a	0.00 ± 0.00g	0	69.00 ± 1.40a	0.00 ± 0.00f
200	56.07 ± 1.73b	17.91 ± 2.83f	50	54.20 ± 1.50b	15.30 ± 1.30e
400	48.79 ± 1.60c	29.85 ± 2.62e	100	40.50 ± 1.30c	32.80 ± 1.50d
600	32.40 ± 0.74d	56.72 ± 1.21d	150	27.10 ± 1.10d	61.50 ± 1.20c
800	22.39 ± 0.69e	73.13 ± 1.13c	200	13.50 ± 0.90e	87.40 ± 1.00b
1000	12.37 ± 0.87f	89.55 ± 1.43b	250	6.00 ± 0.00f	100.00 ± 0.00a
1200	6.00 ± 0.00g	100 ± 0.00a	300	6.00 ± 0.00g	100.00 ± 0.00a

Values are expressed as mean ± standard deviation.

Table 4.

Minimum inhibitory concentration (MIC) and minimum fungicidal concentration (MFC) of Cymbopogon martinii essential oil (CMEO) and positive control (Bavistin) against Colletotrichum gloeosporioides and Fusarium longifundum.

Fungal Strains	CMEO		Positive Control
	MIC	MFC	MIC	MFC
Colletotrichum gloeosporioides (PP594941)	1583.3 ± 28.8a	1883.3 ± 76.4a	516.7 ± 28.8c	833.3 ± 28.8d
Fusarium longifundum (PP594940)	1250.0 ± 50.0e	1550.0 ± 50.0f	216.7 ± 28.8g	466.7 ± 28.8b

Values are expressed as mean ± standard deviation.

Table 3.

Effect of Cymbopogon martinii essential oil (CMEO) and positive control (Bavistin) on mycelial growth and growth inhibition of Colletotrichum gloeosporioides at varying concentrations.

CMEO Concentration (µg/mL)	Mycelial Growth (mm)	Growth Inhibition (%)	Positive Control (µg/mL)	Mycelial Growth (mm)	Growth Inhibition (%)
0	74.95 ± 1.57a	0.00 ± 0.00g	0	74.80 ± 1.50a	0.00 ± 0.00f
250	58.71 ± 2.77b	23.61 ± 2.21f	100	59.50 ± 1.60b	20.45 ± 1.99e
500	49.12 ± 2.14c	37.50 ± 1.94e	200	45.30 ± 1.40c	39.44 ± 1.26d
750	39.54 ± 1.30d	51.39 ± 1.88d	300	31.40 ± 1.20d	58.02 ± 1.17c
1000	23.25 ± 1.16e	75.00 ± 1.68c	400	18.20 ± 1.10e	75.67 ± 0.85b
1250	12.71 ± 1.64f	90.28 ± 2.38b	500	6.00 ± 0.00f	100.00 ± 0.00a
1500	6.00 ± 0.00g	100.00 ± 0.00a	600	6.00 ± 0.00g	100.00 ± 0.00a

Data are presented as mean ± standard deviation.

Representative Petri plate images demonstrating the concentration-dependent inhibition of mycelial growth in C. gloeosporioides and F. longifundum are shown in Supplementary Fig. S3a and S3b, respectively. CMEO demonstrated significant inhibitory effects against both fungi, showing a clear concentration-dependent response. Furthermore, MIC and MFC values supported these findings. For C. gloeosporioides, the MIC and MFC values of CMEO were recorded at 1583 µg/mL and 1883.3 µg/mL, respectively. In contrast, Bavistin exhibited significantly lower values (MIC: 516 µg/mL and MFC: 833 µg/mL). In the case of F. longifundum, the MIC and MFC values for CMEO were lower compared to C. gloeosporioides, at 1250 µg/mL and 1550 µg/mL, respectively, but remained notably higher than those of Bavistin (MIC: 216 µg/mL, MFC: 466 µg/mL) (Fig. 2c). These differences were statistically significant as indicated by Tukey’s test (p < 0.05).

Intracellular ergosterol content analysis

The effect of CMEO on the ergosterol content of F. longifundum and C. gloeosporioides was evaluated to understand its impact on fungal membrane integrity. The results, as presented in Fig. 3; Table 5, showed a concentration-dependent decrease in ergosterol levels in both fungal species when exposed to CMEO at sub-MIC, MIC, and MFC concentrations. Table 5 shows the decrease in ergosterol content in F. longifundum and C. gloeosporioides when treated with different concentrations of CMEO. In F. longifundum, ergosterol content decreased by 45% at sub-MIC, 72% at MIC, and was completely depleted (100% reduction) at the MFC concentration compared to the control. Similarly, in C. gloeosporioides, ergosterol levels were reduced by 40% at sub-MIC, 64% at MIC, and 100% at MFC. The Bavistin also significantly reduced ergosterol content, with reductions of 91% in F. longifundum and 93% in C. gloeosporioides, though it did not result in complete depletion as seen with CMEO at the MFC concentration. In addition to ergosterol depletion, a consistent decline in fungal biomass was observed with increasing CMEO concentrations in both fungi, indicating that the reduction in ergosterol directly affected cell growth and viability. Notably, F. longifundum exhibited slightly higher sensitivity to CMEO than C. gloeosporioides, as evidenced by greater ergosterol reduction at sub-MIC levels. Overall, these findings suggest that CMEO disrupts fungal cell membrane integrity by significantly inhibiting ergosterol biosynthesis, thereby contributing to its strong antifungal effect.

Fig. 3.

Effect of Cymbopogon martinii essential oil (CMEO) on ergosterol content in fungal cell membranes. (a) Reduction of ergosterol in Fusarium longifundum; (b) Reduction in Colletotrichum gloeosporioides. Treatments include control, sub-MIC (MIC/2), MIC, positive control (Bavistin), and MFC concentrations. A concentration-dependent decrease in ergosterol content was observed with increasing CMEO concentration.

Table 5.

Intracellular ergosterol content and membrane integrity of Colletotrichum gloeosporioides and Fusarium longifundum treated with Cymbopogon martinii essential oil (CMEO).

Fungal Strain	Essential Oil Concentration (mg/mL)	Abs (230 nm)	Abs (282 nm)	Fungal Cell Weight (mg)	Ergosterol Content (%)	Percent Reduction (%)
Fusarium longifundum	Control	2.3	1.2	9.8 ± 0.4	5.5 ± 0.3	0 ± 0
	MIC	1.5	0.8	8.2 ± 0.4	2.5 ± 0.2	45 ± 2.3
	MIC/2	0.9	0.5	6.5 ± 0.3	1.0 ± 0.1	72 ± 3.1
	MFC	0	0	5.8 ± 0.2	0 ± 0	100 ± 0
	Positive Control	0.3	0.15	7.1 ± 0.3	0.4 ± 0.1	93 ± 2.0
Colletotrichum gloeosporioides	Control	1.8	0.9	14.5 ± 0.6	6.5 ± 0.4	0 ± 0
	MIC	1.2	0.6	13.2 ± 0.5	3.9 ± 0.3	40 ± 2.5
	MIC/2	0.8	0.4	12.4 ± 0.4	2.3 ± 0.2	64 ± 2.8
	MFC	0	0	11.1 ± 0.3	0 ± 0	100 ± 0
	Positive Control	0.2	0.1	12.0 ± 0.3	0.6 ± 0.2	91 ± 2.2

Values are expressed as mean ± standard deviation. MIC/2 (Sub-MIC or Half the MIC).

Release of cellular components in fungi

The release of intracellular components from F. longifundum and C. gloeosporioides in response to CMEO was quantified by measuring the absorbance at 260 nm (OD₂₆₀) over a period of 120 min (Fig. 4a, b). The treatments included CMEO at sub-MIC, MIC, MFC concentrations, Bavistin, and control. In both fungal species, CMEO treatment led to a time- and concentration-dependent increase in OD₂₆₀. In F. longifundum, the OD₂₆₀ value at MFC increased sharply, reaching 0.750 ± 0.018 at 120 min. This was higher than the absorbance induced by Bavistin (0.705 ± 0.011), MIC (0.622 ± 0.014), and sub-MIC (0.526 ± 0.015), whereas the control remained constant at 0.351 ± 0.00. Similarly, for C. gloeosporioides, the OD₂₆₀ value at MFC was 0.714 ± 0.016 at 120 min, followed by Bavistin (0.621 ± 0.012), MIC (0.552 ± 0.019), sub-MIC (0.413 ± 0.016), and the control (0.319 ± 0.018). The Bavistin induced considerable cellular leakage, surpassing MIC levels but remaining lower than MFC levels in both fungi. Importantly, when comparing the two species, F. longifundum consistently exhibited higher OD₂₆₀ values across all treatment concentrations and time points compared to C. gloeosporioides, indicating that F. longifundum is more sensitive to CMEO-induced membrane disruption.

Fig. 4.

Effect of Cymbopogon martinii essential oil (CMEO) on membrane integrity and extracellular pH of postharvest fungal pathogens. (a, b) Release of cellular components from Fusarium longifundum and Colletotrichum gloeosporioides, respectively, was measured by absorbance over time. (c, d) Changes in extracellular pH for Fusarium longifundum and Colletotrichum gloeosporioides. Treatments include control, sub-MIC (MIC/2), MIC, MFC, and Bavistin. CMEO induced a concentration-dependent increase in membrane permeability and acidification of the extracellular environment.

Changes in extracellular pH

The extracellular pH values in F. longifundum and C. gloeosporioides exposed to CMEO treatments were observed to change over time (Fig. 4c, d). Treatment with CMEO at various concentrations (MIC, sub-MIC, MFC) and Bavistin caused a concentration-dependent decrease in extracellular pH. At the highest concentration (MFC), pH declined rapidly, whereas at lower concentrations (MIC and sub-MIC), the decrease was more gradual. The reduction in pH at all CMEO concentrations was statistically significant when compared to the control (p < 0.05). In contrast, CMEO treatments, including MIC, sub-MIC, and MFC, resulted in a statistically significant decrease in pH over the 120-minute exposure period. The pH values of the MFC treatment at 120 min were notably lower than those of the control, demonstrating a stronger acidification effect, with values of 5.96 ± 0.01 and 6.07 ± 0.02 (mean ± SD) for F. longifundum and C. gloeosporioides, respectively. The pH values at MIC and sub-MIC concentrations were also significantly reduced compared to the control (p < 0.05), confirming that CMEO-induced acidification followed a clear concentration-dependent trend.

SEM analysis of fungal morphology

SEM analysis (Fig. 5) revealed clear morphological alterations in F. longifundum and C. gloeosporioides treated with CMEO. Untreated controls (control: Fig. 5a, b, p, q) showed smooth, cylindrical, and intact hyphae. At sub-MIC concentrations (Fig. 5c, r), CMEO caused shrinkage, deformation, and surface irregularities, while at MIC concentrations (Fig. 5d, s) led to severe fragmentation and complete structural collapse of hyphae. The extent of damage increased with concentration, demonstrating a clear concentration-dependent effect of CMEO on fungal morphology. Where applicable, this concentration-dependent effect may be supported by quantitative observations, such as the proportion of deformed or collapsed hyphae per microscopic field.

Fig. 5.

Scanning electron microscopy (SEM) analysis of fungal morphological alterations induced by Cymbopogon martinii essential oil (CMEO). (a, b) Untreated control Fusarium longifundum; (c) treated with sub-MIC(MIC/2); (d) treated with MIC. (p, q) Untreated control Colletotrichum gloeosporioides; (r) treated with sub-MIC; (s) treated with MIC. CMEO treatment caused visible deformation, collapse, and surface disruption of fungal hyphae, whereas controls exhibited a smooth, intact morphology.

TEM analysis of fungal ultrastructure

TEM observations (Fig. 6) revealed marked ultrastructural changes in F. longifundum and C. gloeosporioides after treatment with CMEO. Control cells (control: Fig. 6a, p) displayed intact cell walls (CW) and membranes, with uniformly distributed cytoplasm and normal organelles. In contrast, at sub-MIC concentrations (Fig. 6b, q) exhibited early stress responses, including vacuolization (V) and cytoplasmic disorganization. MIC concentrations (Fig. 6c, r) showed severe cytoplasmic degradation, blurred cell wall boundaries, and vacuole fusion, while MFC concentrations (Fig. 6d, s) exhibited extreme damage, such as complete cytoplasmic collapse, organelle degeneration (OG), and empty cell walls. These alterations indicate progressive, concentration-dependent structural damage, culminating in the total loss of cellular integrity at higher CMEO concentrations.

Fig. 6.

Transmission electron microscopy (TEM) analysis of fungal ultrastructural changes after treatment with Cymbopogon martinii essential oil (CMEO). (a–d) Fusarium longifundum: (a) control, (b) sub-MIC (MIC/2), (c) MIC, (d) MFC. (p–s) Colletotrichum gloeosporioides: (p) control, (q) sub-MIC, (r) MIC, (s) MFC. CMEO treatment caused concentration-dependent damage, including cytoplasmic disorganization, vacuolization (V), organelle degeneration (OG), and disruption of the cell wall (CW). [ER: Endoplasmic reticulum; N: Nucleus].

Discussion

The present study demonstrates the significant antifungal properties of CMEO against C. gloeosporioides and F. longifundum, with clear evidence of concentration-dependent inhibition of fungal growth, disruption of membrane integrity, and marked morphological alterations. GC‒MS analysis revealed that CMEO is primarily composed of oxygenated monoterpenes (77.9%), which play a central role in its bioactivity. This composition differs from earlier reports in which geraniol is typically the major constituent²⁰, reflecting the considerable chemotypic variability of C. martinii influenced by environmental and genetic factors²¹. Such variability may arise from differences in climate, soil conditions, geographical origin, plant developmental stage, and genetic background, all of which can influence essential oil composition and, consequently, antifungal efficacy. The major components identified, cis-p-Mentha-1(7),8-dien-2-ol, trans-p-Mentha-1(7),8-dien-2-ol, and β-ocimene, align with findings from previous studies on essential oils that highlight the antifungal potential of monoterpenes^22,23.

The results from the antifungal assay support the hypothesis that CMEO exerts its fungicidal activity via direct contact with fungal membranes, leading to growth inhibition in a concentration-dependent manner^24,25. The inhibition rates observed for C. gloeosporioides and F. longifundum are consistent with studies on other essential oils, which have shown that monoterpenes can reduce fungal growth by disrupting cellular processes^26,27. However, CMEO demonstrated higher MIC and MFC values compared to Bavistin, suggesting that although CMEO is effective, it requires higher concentrations to match the efficacy of synthetic fungicides. From a practical perspective, this indicates that CMEO may be less potent on a concentration basis than the synthetic fungicide, but its antifungal relevance remains high because it exerts activity through multiple cellular targets rather than a single specific site of action. This multi-target behavior may offer an important advantage by reducing the likelihood of resistance development and supporting its potential use as a natural alternative in integrated disease management. The Bavistin exhibited different effective concentrations against C. gloeosporioides and F. longifundum, reflecting species-specific variability in fungicide susceptibility. Such differential sensitivity among fungal pathogens is commonly reported and necessitates the use of optimized concentrations to achieve comparable inhibitory effects. Similar concentration-dependent antifungal responses and variation in MIC/MFC values among different fungi have been documented for essential oils, including C. martinii, where inhibitory efficacy depended on the intrinsic tolerance of each species^28–30. These findings support the use of species-specific control concentrations in the present study to ensure biologically relevant and accurate comparisons.

A critical finding in this study was the significant reduction in ergosterol content, a major component of fungal cell membranes, upon treatment with CMEO. The observed depletion of ergosterol, especially at MFC concentrations, indicates that CMEO disrupts fungal cell membrane integrity, leading to loss of membrane function and eventual cell death. These results are in agreement with previous research showing that essential oils and their components can interfere with ergosterol biosynthesis, ultimately destabilizing the fungal cell membrane^31,32. Interestingly, Bavistin reduced ergosterol content by 91% and 93% in F. longifundum and C. gloeosporioides, respectively, but did not achieve complete depletion. This suggests that CMEO may exhibit comparable or complementary effects on membrane permeability and integrity due to its multi-target mode of action, rather than acting through a single specific pathway like synthetic fungicides^33,34. Analysis of cellular leakage, measured by OD₂₆₀, further corroborated CMEO’s antifungal effects. Both fungal species exhibited increased absorbance, indicative of leakage of intracellular components, with F. longifundum showing higher OD₂₆₀ values across all treatment concentrations. This finding suggests that F. longifundum is more susceptible to CMEO-induced membrane disruption, which is consistent with the broader literature showing that the composition and structure of fungal membranes can influence their susceptibility to plant-derived compounds^35,36. Additionally, extracellular pH measurements indicated that CMEO treatment acidified the fungal environment, which could further exacerbate disruption of cellular functions, as previous studies have suggested that pH changes can impair enzyme activity and cellular homeostasis in fungi^37–40. Taken together, the reduction in ergosterol content, increased leakage of intracellular materials, and extracellular acidification indicate a linked mechanism in which CMEO first compromises membrane structure, then disturbs ionic and metabolic balance, and ultimately leads to severe cellular dysfunction. These biochemical disturbances are consistent with the observed morphological and ultrastructural damage, thereby providing a stronger mechanistic explanation for CMEO-induced fungal death.

SEM and TEM analyses provided direct visual evidence of the morphological and ultrastructural alterations induced by CMEO. Treatment with CMEO at higher concentrations led to severe damage to fungal hyphae and cell walls, including shrinkage, deformation, collapse, and fragmentation, which are consistent with observations of other essential oils, such as those from Thymus vulgaris and Origanum vulgare, which also cause morphological disruptions in fungal cells^41,42. TEM analysis revealed that the damage was progressive, with more severe ultrastructural changes, including vacuolization and cytoplasmic degradation, at higher CMEO concentrations. These structural alterations likely arise as downstream consequences of membrane destabilization, ergosterol depletion, and intracellular imbalance, reinforcing the view that CMEO primarily acts through membrane-targeted and multi-site antifungal mechanisms.

These findings highlight the potential of CMEO not only as a fungicide but also as an alternative to synthetic fungicides, particularly due to its multi-target action and lower risk of resistance development, making it a promising candidate for environmentally sustainable disease management strategies.

Experimental methodology

Plant collection and study site

The aerial parts of C. martinii were collected from Kailashpuri, a mountainous region in Udaipur, Rajasthan, India, over a two-year period (2021–2022). Plant collection was conducted in accordance with institutional and ethical guidelines, and formal permission was obtained from the relevant department. Plant samples were collected in accordance with ethical and institutional guidelines, with prior oral consent from the local community and assistance from forest officials. The species was authenticated and identified by Dr. Amit Kotiya, Department of Botany, University of Rajasthan, Jaipur, India. A voucher specimen (RUBL21430) has been deposited in the University of Rajasthan Herbarium (RUBL) for future reference. The collection site is located at 24°44’46"N latitude and 73°43’20"E longitude. Environmental conditions during the collection period included relative humidity ranging from 33 to 72%, ambient temperatures between 24 and 37 °C, and cumulative rainfall of 76.8–235.6 mm, reflecting the seasonal variability of the region.

Hydro-distillation of essential oil

Essential oils from the aerial parts of C. martinii were obtained using a modified hydro-distillation protocol adapted from Rathore et al.⁴³. Fresh plant samples, collected during three distinct developmental phases (vegetative, reproductive, and post-reproductive), were processed through a 3 L round-bottom flask coupled with a Clevenger apparatus for 4 h. For essential oil extraction, approximately 500 g of fresh aerial plant material was used per hydro-distillation cycle, and the process was repeated multiple times to obtain sufficient oil for experimental analyzes. The extracted oils were isolated, dehydrated using anhydrous sodium sulfate to remove residual moisture, and preserved in light-protective amber vials at 4 °C until analysis. The oil yield was determined as a percentage of the fresh plant biomass, calculated using Eq. (1). To evaluate variations in oil production, the procedure was replicated for samples harvested at each growth phase, facilitating a quantitative comparison of yields across developmental stages.

Essential oils analysis by gas chromatography–mass spectrometry

The chemical composition of the essential oil was characterized using gas chromatography–mass spectrometry (GC–MS) with a Thermo Fisher Scientific TRACE™ 1300 GC system coupled to a TSQ 9000 triple quadrupole mass spectrometer. Separation was performed on a dimethyl polysiloxane-coated capillary column (TG-1MS, 30 m × 0.25 mm ID, 0.25 µM film thickness). The oven temperature was programmed from 60 °C (held for 5 min) to 250 °C (held for 15 min). Samples diluted in methanol (1:100, w/v) were injected in split mode (1:50). Helium was used as the carrier gas at a constant flow rate of 1.0 mL/min. The ion source temperature was maintained at 300 °C with electron ionization at 70 eV. Compound identification was based on retention indices (relative to C₉–C₄₀ n-alkanes) and comparison of mass spectra with Wiley 7 and NIST libraries, supported by published literature⁴⁴. Relative abundance (%) was calculated from GC–MS peak areas. Data acquisition and processing were performed using Chromeleon™ 7.3 software.

Sample collection, fungal isolation, and identification

Mature, naturally ripened papaya (Carica papaya L.) fruits with visible fungal infections (lesions, discoloration, or decay) were collected from local markets and farms. Infected tissue was excised aseptically, surface-sterilized in 0.5% sodium hypochlorite (NaOCl) for 1 min, and rinsed twice with sterile distilled water. Tissue segments (5 × 5 mm) from the infection margins were placed on potato dextrose agar (PDA) plates with Bavistin (20 µg/mL) as a positive control, to inhibit bacterial growth, and incubated at 25 ± 2 °C for 5–7 days. Fungal colonies were isolated and purified by sub-culturing onto fresh PDA medium⁴⁵. The selected isolates were representative of the predominant postharvest pathogens and were confirmed through both morphological and molecular identification. Genomic DNA was extracted and PCR amplification of the internal transcribed spacer (ITS) region of rDNA was performed using primers ITS1 and ITS4. PCR products were confirmed on 1.5% agarose gels⁴⁶ and sequenced. Sequence data were analyzed using BLAST-N against GenBank entries and aligned using Clustal Omega for accurate molecular identification^47,48.

In vitro antifungal assay of cCymbopogon martinii essential oil

The antifungal efficacy of CMEO was assessed using a direct contact assay, as described by Yexiao et al.⁴⁹. Stock solutions of CMEO were prepared in 0.5% acetone and incorporated into molten PDA (~ 45 °C) at concentrations ranging from 0.25 to 2 mg/mL. This concentration range was selected based on preliminary experiments and previously reported antifungal studies to evaluate concentration-dependent effects. Negative controls contained 0.5% acetone without CMEO, while Bavistin (positive controls) included PDA with 20 µg/mL Bavistin. A 6-mm mycelial plug from actively growing three-day-old fungal cultures was placed centrally on each plate. Plates were sealed with parafilm and incubated at 26 ± 2 °C in darkness for 4–7 days, depending on the fungal species. Each treatment was performed in triplicate, and the experiment was repeated independently to ensure reproducibility. Fungal growth inhibition was determined by measuring colony diameters and calculating the percentage inhibition using the formula Eq. (2). All treatments were performed in triplicate to ensure statistical reliability.

Determination of minimum inhibitory concentration (MIC) and minimum fungicidal concentration (MFC)

The fungistatic and fungicidal activities of CMEO were assessed using the macro-dilution method adapted from Balouiri et al.⁵⁰. Stock solutions were prepared in 0.5% acetone, and working concentrations ranging from 0.25 to 2 mg/mL were obtained based on preliminary screening to capture both sub-inhibitory and inhibitory effects. Each dilution was added to sterile test tubes containing 5 mL of potato dextrose broth (PDB), followed by inoculation with standardized fungal spore suspensions. Spore suspensions were prepared from 7-day-old fungal cultures grown on PDA. The colony surfaces were gently scraped using a sterile loop, and the spores were collected in sterile distilled water. The suspension was homogenized to ensure even spore distribution, and spore density was determined using a hemocytometer. The final inoculum was adjusted to 0.4–5.0 × 10⁵ CFU/mL. The Bavistin (10–50 µg/mL) was used as a positive control. Untreated tubes served as controls, while Bavistin (10–50 µg/mL) was used as the positive control. Tubes were incubated at 26 ± 2 °C in darkness for 4–7 days, depending on the fungal species. The MIC was defined as the lowest concentration of the essential oil that completely inhibited visible fungal growth. To determine the MFC, 100 µL from MIC-negative tubes was plated on PDA and incubated for 24–48 h. The MFC was recorded as the lowest concentration at which no fungal colonies were observed, indicating > 99.9% reduction in viability. All experiments were performed in triplicate to ensure accuracy and reproducibility.

Determination of intracellular ergosterol content

Sterol extraction was adapted from the standardized protocol of Arthington-Skaggs et al.⁵¹. Fungal mycelia, cultured in PDB containing CMEO at varying concentrations (0, MIC, MFC), were incubated for 5 days at 28 ± 2 °C. These concentrations were selected based on preliminary antifungal assays to evaluate sub-inhibitory and inhibitory effects. Untreated cultures served as controls, while Bavistin-treated cultures were used as the positive control. A total of two selected fungal isolates (C. gloeosporioides and F. longifundum) were analyzed, and all experiments were performed in three independent biological replicates. Harvested mycelia were collected by centrifugation (1223 × g for 5 min), washed, and weighed. Sterol extraction was then carried out as follows. The pellets were homogenized with 25% ethanolic KOH (25 g KOH, 35 mL dH₂O, ethanol to 100 mL), vortexed (1 min), and incubated at 85 °C (1 h). After cooling, 3 mL n-heptane and 1 mL dH₂O were added, followed by vigorous vortexing (3 min). The upper heptane layer was carefully collected and stored at − 20 °C for 24 h. The extract was then diluted five-fold in ethanol for UV analysis (220–300 nm) using a Shimadzu UV-1900i spectrophotometer. Ergosterol and 24(28)-dehydroergosterol (DHE) were identified via characteristic absorption spectra. Ergosterol content was calculated as a percentage of wet biomass weight using the formula Eq. (3), and the reduction in ergosterol content was determined using the equation Eq. (4).

Quantifying cellular component release in fungi

The release of cellular components from fungal mycelia was quantified following an adapted protocol based on Paul et al.⁵². Briefly, mycelia were cultivated in 100 mL PDB for six days, harvested by centrifugation (4000 × g for 20 min), and washed three times with phosphate-buffered saline (PBS, pH 7.0). The pelleted mycelia were resuspended in 100 mL PBS. CMEO was added to achieve final concentrations corresponding to MIC and MFC values, while untreated suspensions served as controls. The suspensions were incubated at 28 ± 2 °C under shaking conditions (200 rpm), which were maintained constant throughout the experiment to ensure reproducibility. Samples were collected at defined time points (0, 30, 60, and 120 min) to monitor temporal changes in cellular leakage. At each time interval, 2 mL aliquots were centrifuged at 12,000 × g for 2 min. The supernatant (1 mL) was then analyzed for absorbance at 260 nm using a Shimadzu UV-1900i spectrophotometer to assess nucleic acid release. Absorbance values were expressed directly without normalization to biomass, as equal amounts of mycelial mass were used across all treatments. All experiments were conducted in three independent biological replicates. The selected time points and CMEO concentrations were chosen to capture both immediate and progressive membrane damage responses.

Determination of extracellular pH

The extracellular pH of fungal cultures was monitored using a Delta 320 pH meter following a modified protocol based on Zhang et al.⁵³. The pH meter was calibrated prior to measurements using standard buffer solutions (pH 4.0, 7.0, and 9.0). Fungal suspensions (10⁵ CFU/mL) were inoculated into 20 mL PDB and incubated for 48 h at 28 ± 2 °C under controlled humidity. After incubation, cultures were centrifuged (4000 × g for 20 min), and the biomass was washed three times with sterile water before being resuspended in 20 mL sterile water. CMEO was added to achieve final concentrations corresponding to sub-MIC, MIC and MFC values, while untreated suspensions served as controls. Extracellular pH was recorded at 0, 30, 60, and 120 min post-treatment. All measurements were performed using three independent biological replicates. pH measurements were used to monitor temporal changes in the growth medium during antifungal treatment.

Morphological analysis by scanning electron microscopy (SEM)

The selected postharvest fungal pathogens were cultured on PDA with MIC/2 (sub-MIC), MIC, and without (control) essential oil treatment at 28 °C for 5 days. Agar segments (5 × 10 mm) were excised for SEM preparation. Initially, the samples were fixed overnight at 4 °C in 2.5% glutaraldehyde, followed by three washes in 0.1 M sodium phosphate buffer (pH 7.2) for 20 min each. Gradual dehydration was carried out using an ethanol series of 30%, 50%, 70%, 80%, 90% and 100% concentrations for 20 min at each step. Samples were then subjected to final dehydration in absolute ethanol for 45 min. Critical point drying was performed using liquid carbon dioxide. The dried samples were mounted on silver stubs and coated with a ~ 10 nm gold layer using a Polaron gold sputter coater. At least three to five independent samples were prepared, and five to ten representative fields/images per sample were examined. The fungal morphology was examined using a Zeiss EVO 18 SEM operated at an accelerating voltage of 20 kV⁵⁴.

Ultrastructural observations by transmission electron microscopy (TEM)

Transmission electron microscopy (TEM) was performed following the modified procedure of Xing et al.⁵⁵. Small pieces (2 mm in diameter) of selected postharvest fungal pathogens, cultured for 5 days in PDB with sub-MIC, MIC and without (control) CMEO treatment, fixed overnight in 2.5% glutaraldehyde at 4 °C. After two washes with 0.2 M PBS buffer (pH 7.4) for 30 min each, the samples were post-fixed in 1.0% osmium tetroxide for 2 h, followed by another 30-min wash with PBS buffer. All fixation steps were conducted at 4 °C. The mycelia were then dehydrated using a gradient ethanol series (50%, 60%, 70%, 80%, 90%, and 100%). Subsequently, samples were transitioned through acetone: ethanol mixtures (1:2, 1:1, and 2:1) for 10 min each before embedding in epoxy resin. Blocks were sectioned into ultrathin slices (~ 70 nm) using a diamond knife. At least three to five sections per sample were examined across three independent replicates. These sections were contrasted with uranyl acetate and lead citrate for 30 min each, then examined under a Talos HR TEM operated at 200 kV.

Statistical analysis

All numerical values were expressed as mean ± standard deviation (SD) from three independent biological replicates per treatment group. Prior to analysis, data were tested for normality and homogeneity of variance. Statistical analysis was performed using one-way analysis of variance (ANOVA) for all datasets, including mycelial growth inhibition, ergosterol content, extracellular pH, and OD₂₆₀ measurements, using OriginPro 2021 (OriginLab, Northampton, MA, USA). Significant differences between means were determined using the Tukey test at the significance level of p < 0.05.

Mathematical formulas

Oil yield (%) was calculated as

Where: Weight of essential oil: Mass of oil extracted, expressed in grams (g).

Weight of fresh plant material: Mass of plant material used for extraction, expressed in grams (g).

1

Mycelial growth inhibition (%) was expressed as

2

Where: D_c: Colony diameter in the control plate, expressed in millimeters (mm).

D_t: Colony diameter in the treatment plate, expressed in millimeters (mm).

Ergosterol content was quantified as

3

Where: F dilution factor of the sterol extract in ethanol (–).

𝐴_282/290: Absorbance at 282 nanometers (nm) divided by extinction coefficient 290 for crystalline ergosterol.

𝐴_230/518: Absorbance at 230 nanometers (nm) divided by extinction coefficient 518 for 24(28)-dehydroergosterol (DHE).

Wet biomass: Mass of fungal cells in grams.

Percentage reduction was determined by

4

Where: Initial ergosterol content: Ergosterol content measured in the control sample.

Current ergosterol content: Ergosterol content measured in the treated sample.

Conclusion

In the present study, Cymbopogon martinii essential oil exhibited strong antifungal activity against C. gloeosporioides and F. longifundum, two postharvest pathogens of papaya. CMEO disrupted fungal membrane integrity, resulting in ergosterol depletion, leakage of cellular contents, and significant morphological and ultrastructural damage. The results indicate that CMEO is an effective natural antifungal agent with potential for postharvest disease control. Its use may offer a safer, more sustainable alternative to chemical fungicides for managing postharvest spoilage of papaya.

Supplementary Information

Below is the link to the electronic supplementary material.

Abbreviations

ANOVA

Analysis of variance

CFU

Colony-forming unit

CMEO

Cymbopogon martinii essential oil

dH₂O

Distilled water

DHE

Dehydroergosterol

EOs

Essential oils

FID

Flame ionization detector

GC‒MS

Gas chromatography–mass spectrometry

ITS

Internal transcribed spacer

KOH

Potassium hydroxide

MFC

Minimum fungicidal concentration

MIC

Minimum inhibitory concentration

mL

Milliliter

mm

Millimeters

NaOCl

Sodium hypochlorite

nm

Nanometers

µL

Microliter

OD

Optical density

PCR

Polymerase chain reaction

PDA

Potato dextrose agar

PDB

Potato dextrose broth

RI

Retention indices

rpm

Revolutions per minute

RT

Retention time

SD

Standard deviation

SEM

Scanning electron microscopy

TEM

Transmission electron microscopy

Author contributions

PR and MM contributed to the conceptualization of the study. PR and GY performed the experiments. PR, GY, and MM conducted the statistical analysis. PR prepared the original draft of the manuscript. MM contributed to visualization, validation, and manuscript editing, and supervised the entire study. All authors reviewed and approved the final version of the manuscript.

Funding

The authors declare that no funding was received for this study.

Data availability

The nucleotide sequences of the fungal isolates identified in this study have been deposited in the GenBank database under accession numbers [Fusarium longifundum (PP594940); Colletotrichum gloeosporioides (PP594941)] at NCBI (https://www.ncbi.nlm.nih.gov/). Other datasets generated during the current study are related to ongoing research and are therefore confidential and not publicly available. All other data generated or analysed during this study are included in this published article and its supplementary information files.

Declarations

Competing interests

Dr Mukesh Meena is an Editorial Board Member of Scientific Reports. This submission was handled independently to ensure an unbiased peer-review process. The authors declare no other competing interests.

Ethics declarations

Cymbopogon martinii is a naturally occurring ephemeral grass that grows during the rainy season and dies off in winter. The plant material used in this study was collected from naturally growing populations in non-protected areas with permission granted by the Department of Botany, Mohanlal Sukhadia University, Udaipur, Rajasthan, India. The species is not listed as endangered under the IUCN Red List or CITES. All research and field studies on this plant complied with institutional, national, and international guidelines and legislation.