Characterization and Antifungal Activity of a Hevein‐Like Peptide‐Containing Extract From Capsicum annuum Leaves Against Phytopathogenic Fungi

Abstract

This study is aimed at isolating and characterizing antimicrobial peptides (AMPs) from the leaves of Capsicum annuum (L.) cv. Carioquinha and investigating their antifungal activity and mechanism of action against phytopathogenic fungi. An initial extract was obtained from leaves through hydromethanolic extraction followed by partitioning with dichloromethane. The extract was resuspended in aqueous acetonitrile and centrifuged to obtain the soluble supernatant fraction (ExS). The ExS fraction was purified by reverse‐phase high‐performance liquid chromatography (RP‐HPLC). Protein bands from the ExS fraction were excised from Tricine‐SDS‐PAGE gels and analyzed by LC–MS/MS. The obtained peptide sequences were compared with protein databases. Antifungal activity of the ExS fraction was evaluated through quantitative fungal growth inhibition assays against the phytopathogenic fungi Fusarium solani, Fusarium oxysporum, Colletotrichum gloeosporioides, and Colletotrichum scovillei, and tests were conducted to investigate the mechanism of action of the peptides present in the ExS fraction. RP‐HPLC analysis of the ExS fraction revealed a single peak (P1) corresponding to a protein band of approximately 6.5 kDa, and all fragments obtained by LC–MS/MS were similar to peptides belonging to the hevein‐like family. The highest inhibition was observed for C. gloeosporioides, with 78.3% growth inhibition at 200 μg mL⁻¹. Mechanistic studies revealed that the ExS fraction caused membrane permeabilization in C. gloeosporioides, induced reactive oxygen species (ROS) production, and reduced cell viability. The ExS fraction also caused vacuolization of hyphae in F. oxysporum. This study highlights the potential of AMPs isolated from C. annuum leaves to inhibit phytopathogenic fungi.

1. Introduction

The application of chemical pesticides has significantly improved agricultural productivity. Considering the global demand for food, this will be even more critical in the future. Almost 20 million tons of these chemicals are used every year across the globe to control pests and plant diseases [1]. However, there is great concern regarding the intensive use of these pesticides because of their significant health risks and environmental damage. The intensive use of chemical pesticides over a prolonged period has been associated with serious health conditions such as chronic illness, diabetes, and cancer, along with neurological and respiratory disorders [2–4]. Family farmers in Brazil are quite exposed and have been reported to suffer sustained exposure, with clinical symptoms consistent with pesticide toxicity [5]. Eliminating useful pest strains decreases soil quality and beneficial microbes while causing resistant pests to evolve, which leads to more complicated disease management strategies owing to an uncontrolled rise in superbugs [3].

One of the main challenges is the control of phytopathogenic fungi, which are responsible for 7%−24% of crop loss, especially because of their increased resistance to traditional fungicides [6, 7]. These challenges have given rise to a growing interest in exploring natural, plant‐based compounds as alternatives to chemical pesticides. These compounds are therefore considered promising sources for the development of biomolecules aimed at controlling phytopathogens. Plant antimicrobial peptides (AMPs) are promising alternatives for sustainable pathogen control. AMPs are small bioactive molecules present in various parts of plants that play crucial roles in defense against biotic and abiotic stresses [8, 9]. Because of their direct action on pathogenic microorganisms and low toxicity to human cells, they are excellent candidates for the development of new therapeutic strategies [10].

Filamentous fungi from the Colletotrichum and Fusarium genera are phytopathogenic organisms of high economic and environmental importance because of their wide geographic distribution, which can cause diseases such as anthracnose and Fusarium wilt. Their widespread occurrence and species diversity, coupled with mycotoxin production, complicate control efforts, which are further exacerbated by the development of resistance to standard fungicides [11–14]. Anthracnose is a fungal disease caused by species of the genus Colletotrichum, which produces dark spots and necrotic lesions on fruits, leaves, and shoots, potentially leading to leaf drop and reduced crop productivity [13, 14]. Therefore, the development of AMPs targeting these pathogens is a promising strategy.

Within the scope of potential sources of AMPs, the genus Capsicum (Solanaceae) is recognized for producing several active compounds, including AMPs, with antimicrobial and therapeutic activities. Capsicum peppers encompass species cultivated and consumed globally, such as Capsicum annuum, Capsicum chinense, and Capsicum frutescens [15–17]. Studies have demonstrated the ability of these plants to produce peptides with antimicrobial activity against a wide range of pathogens, emphasizing the need for biotechnological exploration [18, 19]. Hevein‐like peptides (HLPs) constitute an important group of plant AMPs. These peptides are characterized by high cysteine and glycine contents, molecular masses ranging from 3 to 5 kDa, and a high number of disulfide bonds [10, 20]. HLPs have a conserved domain capable of binding to chitin, which is the main component of the fungal cell wall. This structural feature confers remarkable stability to these peptides and contributes to their antifungal activity, playing an important role in plant defense against pathogenic microorganisms [10, 21, 22].

AMPs of the hevein group and HLPs have been reported in several plant species, including the rubber tree (Hevea brasiliensis), moringa (Moringa spp.), European hawthorn (Euonymus europaeus), and sweet peas (Stellaria media). These peptides show great promise in phytopathological research because of their antifungal and antibacterial properties. However, there is a lack of literature concerning the antimicrobial activity of HLPs from C. annuum, especially regarding their action on fungi, such as Candida and resistant phytopathogenic fungi. Heveins and HLPs have only recently been classified as emerging plant AMPs, and their biological activities are less understood than those of other well‐studied AMPs [10].

The potential usefulness of Capsicum AMPs stems from their strong antimicrobial activity, low tendency to foster resistant microbial strains, lower toxicity to human cells, and weaker chances of developing microbial resistance compared with other products [10]. This study is aimed at purifying and characterizing the extract supernatant (ExS) fraction from the leaves of C. annuum (L.) cv. Carioquinha and analyzing its antifungal activity against several phytopathogenic fungi.

2. Materials and Methods

2.1. Seeds

Pepper seeds were provided by Laboratório de Melhoramento Genético Vegetal (LMGV) of the Centro de Ciências e Tecnologias Agropecuárias (CCTA), Universidade Estadual do Norte Fluminense (UENF), Campos dos Goytacazes, Rio de Janeiro, Brazil. The plants were grown in commercial substrate in seedling trays for 30 days under controlled conditions (30°C and a 12‐h photoperiod). After this period, seedlings that exceeded 10 cm in height were transplanted into 5‐L pots and cultivated with irrigation twice a day for 60–90 days.

2.2. Microorganisms

The fungal species Colletotrichum gloeosporioides, Colletotrichum scovillei, Fusarium solani, and Fusarium oxysporum were obtained from the Microorganism Collection (Micoteca) of the Universidade Federal de Pernambuco, Recife, Pernambuco, Brazil. The filamentous fungi were maintained on Sabouraud agar (1% peptone, 2% glucose, and 1.7% agar‐agar) (Merck) at the Laboratório de Fisiologia e Bioquímica de Microrganismos, Centro de Biociências e Biotecnologia (CBB), UENF, Campos dos Goytacazes, Rio de Janeiro, Brazil.

2.3. Extraction and Partial Purification of Peptides

Protein extraction of leaves was carried out according to the methodology described by Cherene et al. (2023). Fresh leaves (40 g) were macerated and extracted with 120 mL of 60% methanol (v/v) at room temperature for 24 h. The extract was filtered to remove plant debris and partitioned with dichloromethane (1:1, v/v) for 24 h at room temperature. The hydromethanolic phase was separated, concentrated in a fume hood for 18 h, and lyophilized. The lyophilized extract (0.1 g) was dissolved in 10 mL of 20% acetonitrile (ACN) (v/v) and centrifuged at 6000 rpm for 3 min, and the supernatant (ExS) was collected and lyophilized.

The ExS fraction was purified by reverse‐phase high‐performance liquid chromatography (HPLC) using a C₁₈ VP‐ODS column (250 × 4.6 mm; Shim‐pack, Shimadzu) with a C₈ guard column (Pelliguard, Sigma). The column was equilibrated with solution A (0.1% trifluoroacetic acid, TFA, in ultrapure water). The ExS sample was loaded onto the pre‐equilibrated column and eluted with a gradient of solution B (90% propanol, 0.1% TFA): 0% B (0–2 min), 0%–10% B (2–10 min), 10%–32% B (10–60 min), 32%–50% B (60–74 min), and 50% B (74–75 min), followed by a return to 0% B (75–85 min). Separation was performed at 37°C with a flow rate of 0.5 mL min⁻¹, and peptide detection was monitored at 220 and 280 nm. Target fractions were collected and lyophilized for further analysis [23].

2.4. Tricine Gel Electrophoresis

The molecular weight profile of ExS and HPLC peak were analyzed by polyacrylamide gel electrophoresis containing tricine and SDS (sodium dodecyl sulfate), according to the method described by Schägger and von Jagow (1987). Samples were denatured in SDS buffer at 95°C for 5 min, and 10 μL of each was loaded onto the gel. Electrophoresis was conducted at 120 V for 18 h. Gels were stained with Coomassie Blue, and bands of interest were excised for subsequent peptide characterization.

2.5. Analysis of Amino Acid Residues

2.5.1. In‐Gel Enzymatic Digestion

Protein bands visualized by SDS–PAGE were excised with a sterile scalpel and cut into ~1‐mm³ gel pieces. Each sample was transferred to a 1.5‐mL microcentrifuge tube containing 1 mL of destaining solution: 50 mM ammonium bicarbonate (AmBic, Sigma–Aldrich) and ACN (Merck) in a 1:1 (v/v) ratio and incubated overnight at room temperature with gentle agitation in a thermomixer. The solution was replaced with 200 μL of fresh destaining solution for 1 h and then completely removed. The gel pieces were dehydrated twice with 500 μL of 100% ACN for 1 min each.

For protein reduction, 200 μL of 10 mM dithiothreitol (DTT, GE Healthcare) in 100 mM AmBic was added, and the mixture was incubated at 55°C for 30 min with gentle mixing. After the reducing solution was removed, the gel pieces were dehydrated again with ACN, alkylated with 200 μL of 55 mM iodoacetamide (IAA, Merck) in 100 mM AmBic, and incubated in the dark at room temperature for 30 min. Digestion was performed by adding 200 μL of cold trypsin solution (10 mM AmBic with 10% ACN, 1:100 enzyme‐to‐protein ratio; V5111, Promega) to each sample. The tubes were incubated at 4°C for 30 min, followed by incubation at 37°C for 18 h. After digestion, 200 μL of extraction buffer (5% formic acid in 100% ACN, 1:2, v/v) was added to each sample, which was subsequently incubated at 37°C for 30 min in a thermomixer. The extracts were then completely dried via a SpeedVac and resuspended in 50 μL of 0.1% formic acid prepared in 50 mM AmBic prior to mass spectrometry analysis [24].

2.5.2. Mass Spectrometry Analysis

Peptides were characterized using a high‐resolution NanoAcquity UPLC system coupled with a Synapt G2‐Si HDMS mass spectrometer (Waters, Manchester, United Kingdom). For each run, 2 μg of peptide sample was injected. The samples were first loaded onto a C₁₈ trap column (180 μm × 20 mm, 5 μm; Waters) at a flow rate of 5 μL min⁻¹ for 3 min. Separation was performed on a nanoAcquity HSS T3 C₁₈ analytical column (75 μm × 150 mm, 1.8 μm) at 400 nL min⁻¹ and 45°C. A binary solvent system was used with mobile phases A (water + 0.1% formic acid) and B (ACN + 0.1% formic acid). The gradient increased from 7% to 40% B over 92.72 min, reached 99.9% at 106 min, returned to 7% at 106.1 min, and remained stable until the end of the 120‐min run.

Data acquisition was performed in positive ion mode in high‐resolution V mode with ion mobility and data‐independent acquisition. The ion mobility wave velocity was set to 600 m s⁻¹, and the transfer collision energy ranged from 19 to 55 V in high‐energy mode. The source temperature was maintained at 70°C, with cone and capillary voltages set to 30 and 2750 V, respectively. Time‐of‐flight (TOF) scans were collected every 0.5 s across a mass range of 50–2000 Da. Glu1‐fibrinopeptide B (Sigma–Aldrich) at 100 fmol μL⁻¹ was used as an external calibrant, with lock mass acquired every 30 s. Data acquisition and processing were performed using ProteinLynx Global Server (PLGS v.3.0.2; Waters). Sequences obtained via MS/MS were analyzed using similarity searches against the Protein Data Bank (PDB) database [25]. Selected sequences were aligned using ClustalW [26].

2.6. In Silico Peptide Structure

Three‐dimensional structures of the identified proteins were retrieved from the AlphaFold protein structure database (https://www.alphafold.ebi.ac.uk/) based on their respective FASTA sequences [27]. AlphaFold is an artificial intelligence system capable of predicting protein folding and spatial conformation with high accuracy, using only the primary amino acid sequence as input. The amino acid sequence obtained via mass spectrometry was subjected to the AlphaFold pipeline, and the resulting model was visualized and analyzed using PyMOL (Schrödinger, LLC) to assess secondary and tertiary structural features.

2.7. Antifungal Assay

To verify the effect of ExS on filamentous fungi (the species C. gloeosporioides, C. scovillei, F. solani, and F. oxysporum), fungal growth inhibition assays were conducted according to a previously described protocol, with some modifications [28]. Conidia at a concentration of 1 × 10⁴ conidia mL⁻¹ were incubated at 30°C in a final volume of 100 μL in the presence of ExS at different concentrations (1.56; 3.12; 6.25; 12.5; 25; 50; 100; and 200 μg mL⁻¹). Fungal growth was also determined in the absence of peptides. The assay was performed in cell culture plates (96 wells) at 30°C. Optical density was determined after 24 h using a microplate reader at a wavelength of 620 nm, allowing quantitative evaluation of fungal growth inhibition. The percentage of fungal growth inhibition was calculated relative to the untreated control using the formula: inhibition (%) = [(ODcontrol − ODsample)/ODcontrol] × 100. The entire test procedure was performed under aseptic conditions in a laminar flow hood, according to the methodology adapted from Broekaert et al. (1990) [29].

2.8. Cell Viability Assay

To analyze the effects of the peptides on filamentous fungi C. gloeosporioides, a concentration of 1 × 10⁵ conidia mL⁻¹ in sterile distilled water was used and incubated at 30°C in a final volume of 100 μL in the presence of ExS (200 μg mL⁻¹) and 20% sucrose. For counting, cells treated or untreated with ExS were diluted 1000 times after 6 h of incubation. An aliquot was spread over the surface of a Petri dish containing Sabouraud agar via a Drigalski spatula and cultured at 30°C for 36 h. Colony‐forming units (CFUs) were generated after incubation [30]. Results were expressed as CFU mL⁻¹ and relative viability compared to the untreated control. The viability of C. gloeosporioides was analyzed under the effects of ExS after the cells were incubated for 6 h at a concentration of 200 μg mL⁻¹, and the CFUs were counted after 36 h. The cell viability assay was performed only with ExS owing to the quantity required to perform the assay. All assays were performed in triplicate.

2.9. Membrane Permeabilization Assay

The membrane permeabilization of filamentous fungal cells treated with ExS (200 μg mL⁻¹) was investigated via the Sytox Green fluorescent probe according to the methodology described by Taveira et al. (2018), with some adaptations. Twenty‐four hours after the inhibition test, the filamentous fungi were incubated with 0.2‐μM SYTOX Green fluorescent probe for 15 min at 30°C. They were then analyzed for differential interference contrast (DIC) using an optical microscope (Axioplan. A2, Zeiss, Version 4.0) equipped with a set of fluorescence filters (excitation wavelength 450–490 nm; emission wavelength: 500 nm) [31]. Untreated cells were used as negative controls, whereas the positive control consisted of cells subjected to stress by dry heat exposure. Negative control images from the membrane permeabilization assay showed little or no green fluorescence, indicating intact fungal cell membranes. DIC images should show normal hyphal growth with typical hyphal morphologies and branching patterns that are characteristic of filamentous fungi. Positive control images should exhibit intense green fluorescence, indicating penetration of the dye into the fungal cells.

2.10. ROS Induction Determination Assay

In order to evaluate the potential for ExS to induce oxidative stress, the fluorescent probe H₂DCFDA (2 ^′,7 ^′ dichlorodihydrofluorescein diacetate) was utilized to measure the increase in reactive oxygen species (ROS). This methodology was adapted from the approach described by Mello et al. (2011) with certain modifications. Initially, cells from different species of filamentous fungi were incubated with the samples (200 μg mL⁻¹). Following a 24‐h period of incubation, the cells were once more incubated with the specific probe for ROS detection (H₂DCFDA, 20 μM) [32]. Following a 30‐min incubation period at ambient temperature, the cells were analyzed using a fluorescence microscope. The cells exhibit permeability to H₂DCFDA, which, when oxidized by ROS, forms dichlorofluorescein (DCF), a highly fluorescent compound. Increased fluorescence signals indicate higher levels of ROS, whereas lower or absent fluorescence signals suggest the presence of these species. Employing this methodology, the negative control (untreated cells) should manifest fungal hyphae with a normal morphology under DIC optical microscopy, whereas fluorescence microscopy should demonstrate low fluorescence signals, consistent with reduced levels of ROS. Hydrogen peroxide (30%) was used as a positive control, which resulted in the presence of dead fungal hyphae with evident morphological changes in bright‐field images, accompanied by strong fluorescence signals, indicating high levels of ROS in the cells.

2.11. Metacaspase Activity Detection Assay

Metacaspase activity was detected using CaspACE FITC‐VAD‐FMK marker (Promega). Following a 24‐h incubation with ExS (200 μg mL⁻¹), the fungal cells were resuspended and washed once in 500 μL of PBS and then resuspended in 50 μL of staining solution containing 50 μM of the FITC‐VAD‐FMK marker, as described by Taveira et al. (2018). To validate the assay, 1 mM acetic acid was used as the negative control. The samples were observed under a fluorescence microscope. The CaspACE FITC–VAD–FMK probe is a fluorescent marker utilized for detecting the activity of caspases, which are pivotal enzymes in the process of apoptosis.

2.12. Vacuolar Mapping of Fungi

Vacuolar mapping of filamentous fungi was performed using an FM4‐64 probe. Fungal cells were incubated with ExS (200 μg mL⁻¹) for 24 h and then treated with FM4‐64 for 1 h at 30°C. The cells were subsequently washed and centrifuged twice with PBS to remove the free FM4‐64. The control group was treated with only FM4‐64 [33]. The cells were analyzed by DIC under an optical microscope (Axioplan. A2, Zeiss) equipped with a fluorescence filter set to absorption/emission wavelengths of 585/590 nm.

2.13. Mitochondrial Functionality

Mitochondrial functionality was assessed using the fluorescent probe JC‐1 according to the methodology described by Taveira et al. (2018). After the growth inhibition assay, the cells treated with the ExS (200 μg mL⁻¹) were incubated with 2 μL of the fluorescent probe JC‐1 (3.85 mM) for 40 min in the dark at 30°C. After this period, the cells were observed under an optical microscope (Axioplan Version 4.0) equipped with fluorescence filter sets for excitation/emission at 450–490 nm and 585/590 nm [34].

Cell proliferation reagent WST‐1 (Roche) was used to assess mitochondrial activity of the fungal cells treated with ExS (200 μg mL⁻¹) [35]. The cells treated were incubated with 10 μL of WST‐1 reagent and 2 μL of 2,3,5,6‐tetramethyl‐1,4‐benzoquinone (duroquinone; final concentration, 0.24 mM of a stock of 12.24 mM in dimethyl sulfoxide [DMSO]), which was used as an electron couple reagent. Positive control used 10‐μL acetic acid 100% incubated for 10 min. Mitochondrial activity percentage was calculated according to [(ABS450nm of test samples × 100)/ABS450nm of control]. After this period, the optical density was monitored at 450 nm for 2 h in the dark at 30°C.

2.14. Fluorescence Analysis

Cellular fluorescence was initially assessed both qualitatively and quantitatively via fluorescence microscopy, which uses specific markers, such as SYTOX Green (for membrane integrity assessment) and fluorescent probes for ROS. To complement these qualitative observations, a quantitative analysis of fluorescence was performed from representative images using ImageJ software (NIH, Bethesda, Maryland, United States), expressed as a percentage of the fluorescent area (area fluorescent/area total) × 100. Analyses were performed using a standardized threshold‐based fluorescence segmentation method. The parameters that were extracted from each image included the fluorescent area (μm²), the surface area occupied by fluorescence, and the percentage of the fluorescent area. In addition, the proportion of fluorescent area relative to the total area of the image was determined. The images were obtained with the same objective and exposure parameters for all samples, with a defined scale of 20 μm, without zoom application. As the resolution varied between experiments (2584 × 1936 pixels for Colletotrichum spp. and 1292 × 968 pixels for Fusarium spp.), the pixel values were converted to μm² on the basis of the scale calibrated in the software. This ensured comparability of the data.

2.15. Statistical Analysis

Growth inhibition assays for filamentous fungi were performed in triplicate, and the results are presented as mean ± standard deviation. Statistical analysis was performed using one‐way analysis of variance (ANOVA) followed by Tukey′s post hoc test. ANOVA with p < 0.05 between the means of the control and treatment groups was considered significant. All statistical analyses were performed using GraphPad Prism software (Version 8.0, for Windows).

3. Results

3.1. Peptide Purification and Partial Characterization From the ExS Fraction

The ExS fraction was subjected to a purification process by HPLC. The resulting chromatogram exhibited a unique peak with a retention time ranging from 7 to 10 min (Figure 1A). The electrophoretic profile corresponding to the peak obtained by HPLC also revealed a single band with a molecular mass close to the 6.5‐kDa marker (Figure 1B).

Figure 1.

(A) Chromatogram of the ExS protein extract leaves of C. annuum. The column preparation involved the use of 0.1% trifluoroacetic acid (TFA) in solution A, and elution was performed using a gradient from 90% propanol in 0.1% TFA to solution B. (B) SDS‐Tricine gel electrophoresis of the corresponding band present in ExS of approximately 6.5 kDa; MBP: molecular mass markers (kDa). The molecular weights of the substances in question are 26.6, 17.0, 14.2, 6.5, 3.3, and 1.0 kDa.

3.2. Mass Spectrometry

The analysis of amino acid residues by mass spectrometry (LC–ESI–MS/MS) was performed on two distinct samples: the band of approximately 6.5 kDa present in ExS, and the P1 peak obtained after reversed‐phase chromatography (HPLC). The sequence alignment was performed using the CLUSTALW algorithm and compared to the NCBI database. The analysis of the mass spectrum of the band from the ExS fraction revealed a sequence fragment (TREYCGPGCQSNCRR) with high similarity to HLPs. The protein band of the P1 peak obtained from the purified fraction (LCCSQFGFCGTTREYCGAGCQSNCR) also exhibited high similarity to HLPs, including regions with 100% identity to prohevein sequences (Figure 2), hereafter referred to as CaCHev (C. annuum HLP), a nomenclature that reflects both its provenance and its sequence similarity with the hevein family. A sequence comparison of CaCHev reveals the presence of a highly conserved region, which is also found in antimicrobial proteins such as PNAMP1 (PHT87891.1) and antifungal protein (AAL73184.1), as well as proheveins from Capsicum species, including prohevein from C. chinense Jacq. (PHU23618.1) and prohevein from C. annuum (KAF3628901.1). In addition, amino acid sequence similarity with a chitin‐binding peptide isolated from seeds of C. chinense Jacq. was found.

Figure 2.

Alignment of the 15 amino acid residues present in the ExS fraction and the 25 amino acid residues present in P1 indicated compatibility with sequences similar to hevein‐like peptides. Partial antimicrobial protein PN‐AMP1 (1) (sequence ID: PHT87891.1), antifungal protein (2) (sequence ID: AAL73184.1), prohevein from C. chinense (3) (sequence ID: PHU23618.1), prohevein from C. annuum (4) (sequence ID: KAF3628901.1), and chitin‐binding peptide from C. chinense (5). P % indicates the percentage of positive residues with the same physicochemical characteristics. Bold letters represent amino acid residues in the sequence. Spaces (‐) were introduced for better alignment. I % indicates the percentage of identical residues and amino acids are highlighted in bold. The red colors indicate amino acid residues of cysteine present in the sequence.

3.3. Physicochemical Properties and Structural Prediction of the CaCHev Peptide

The three‐dimensional structure of the CaCHev peptide was predicted based on similar modeling using a prohevein template. The resulting model revealed a compact and organized structure, typical of HLPs, composed of two antiparallel β‐strands and one α‐helix (Figure 3A). These secondary structure elements are indicated with arrows, highlighting the β‐sheets and α‐helix arrangement. A chitin‐binding domain (CBD) was identified based on sequence similarity and structural prediction. This predicted fold reinforces the conserved architecture of hevein‐like proteins involved in carbohydrate recognition and antifungal activity.

Figure 3.

(A) Predicted three‐dimensional structure of prohevein from C. chinense (AlphaFold ID: A0A2G3CY09), visualized in PyMOL. The protein is shown in cartoon representation (light blue), with the fragment identified by mass spectrometry (LCCSQFGFCGTTREYCGAGCQSNCR) highlighted in pink. (B) Helix projection of the CaCHev peptide is presented. Basic polar residues are indicated in red, acidic polar residues are indicated in blue, charged polar residues are indicated in green, and nonpolar residues are indicated in yellow.

The CaCHev peptide, composed of 25 amino acid residues, was analyzed for its physicochemical properties and residue distribution. According to the classification of side‐chain characteristics, polar amino acids—including both basic (R, H, K) and neutral residues (S, T, N, Q, C)—were predominant. These residues are associated with potential hydrogen bonding and electrostatic interactions. Acidic residues (D, E) were also present and may contribute to structural stabilization through salt bridge formation. Nonpolar residues (A, G, V, I, L, M, F, Y, W, P), distributed along the sequence, suggest the presence of hydrophobic regions that may interact with cellular membranes—a feature commonly observed in AMPs. These characteristics were visualized using the NetWheels peptide helical wheel projection, which revealed a distinct separation between hydrophobic and hydrophilic residues along the helical structure (Figure 3B).

3.4. Effects of Proteins on Fungal Growth

A quantitative assay was performed to evaluate the effect of the ExS fraction on the growth of phytopathogenic fungi (Figure 4). The growth of C. gloeosporioides was reduced by 78.3% and 64.7% following incubation with ExS at 200 and 100 μ g mL⁻¹, respectively. At 200 μ g mL⁻¹, ExS reduced the growth of F. solani by 17.4%. For F. oxysporum, growth inhibition reached 26.9% and 24.2% at 200 and 100 μ g mL⁻¹, respectively. No inhibitory effect was observed against C. scovillei at any tested concentration of ExS.

Figure 4.

The present study investigates the effect of ExS at varying concentrations (1.56, 3.12, 6.25, 12.5, 25, 50, 100, and 200 μg mL⁻¹) on the growth of fungi belonging to the genera (A and C) Colletotrichum and (B and D) Fusarium over 24 h. The number of asterisks corresponds to different levels of significance: p < 0.05 ( ^∗), p < 0.01 ( ^∗∗), p < 0.001 ( ^∗∗∗), and p < 0.0001 ( ^∗∗∗∗).

3.5. Effect of ExS on Growth and Viability of C. gloeosporioides

The results demonstrated that ExS induced a continuous and significant inhibition of C. gloeosporioides growth over the 48 h. In comparison with the control group (untreated cells), treatment with ExS resulted in a significant difference in fungal growth during the 48‐h evaluation (Figure 5A). The findings of this study indicate that ExS not only initiates growth inhibition but sustains its effect over time. The results showed a decrease in colonies in the ExS‐treated group compared to the control group (untreated cells) (Figure 5B). In addition, the reduction in cell viability in the treated group was 36.7%, indicating that ExS treatment induces a decrease in fungal cell viability. These results complement the optical density measurements by providing a direct assessment of fungal viability. After performing these tests, we proceeded to analyze the potential mechanisms of action of ExS that interfered with fungal viability.

Figure 5.

(A) The growth curve of the plant pathogen C. gloeosporioides was analyzed at different concentrations (μg mL⁻¹) of C. annuum ExS over a period of 48 h. ● Control (0 μg mL⁻¹), ■ 200 μg mL⁻¹, ▲ 100 μg mL⁻¹, ▼ 50 μg mL⁻¹, ◆ 25 μg mL⁻¹, ○ 12.5 μg mL⁻¹, ▢ 6.25 μg mL⁻¹, △ 3.12 μg mL⁻¹, and ▽ 1.56 μg mL⁻¹. The analysis was performed at intervals of 12, 24, 30, 36, and 48 h. The values represent the mean (±SD) of triplicates. Asterisks are used to denote significant differences (p < 0.05) between treatments and the control. (B) The cell viability of C. gloeosporioides was examined following treatment with ExS (200 μg mL⁻¹) for a period of 6 h, with subsequent colony forming unit (CFU) counts being conducted after a 36‐h incubation. The data presented herein correspond to the mean values (±SD) derived from triplicate experiments. Asterisks are used to denote significant differences (p < 0.01) between treatments and the control.

3.6. Effect of ExS on Plasma Membrane Permeabilization

The results demonstrated the presence of fluorescence signals in C. gloeosporioides and C. scovillei treated with ExS (200 μg mL⁻¹), suggesting membrane permeabilization after 24 h (Figure 6). It has been demonstrated that this mechanism is associated with a loss of cellular integrity and probable cell death. No evidence of membrane permeabilization was detected in either F. oxysporum or F. solani. In C. gloeosporioides, the negative control exhibited no fluorescent area (< 0.01%), whereas the positive control registered 79.89%, and the ExS treatment resulted in 53.62% of the fluorescent area. The data suggest that membrane permeabilization was significantly induced only in C. gloeosporioides, being minimal or absent in the other fungal species under the evaluated conditions.

Figure 6.

Analysis of membrane permeabilization of C. gloeosporioides, C. scovillei, F. solani, and F. oxysporum in the presence of ExS (200 μg mL⁻¹) with the fluorescent marker Sytox Green (0.2 μM). The analysis was performed after 24 h in the fungal species tested. Negative control: untreated cells. Positive control: cells induced to stress by dry heating. Bar = 20 μm.

3.7. Determining the Induction of Intracellular ROS in Fungal Cells

The results demonstrate a significant increase in fluorescence in C. gloeosporioides, F. oxysporum, and F. solani treated with ExS (200 μg mL⁻¹) in comparison to the negative control (Figure 7), whereas C. scovillei exhibited minimal fluorescence, similar to the negative control. Fluorescent signals were particularly prominent in the hyphae and conidia of C. gloeosporioides, suggesting oxidative stress mediated by ROS. Quantitative fluorescence analysis demonstrated that, in C. gloeosporioides, the negative control exhibited minimal fluorescence (0.01%), whereas the positive control reached 467.67%, and treatment with ExS resulted in 31.39%. For C. scovillei, the fluorescence levels were found to be low, with 0.00% recorded in the negative control, 18.89% in the positive control, and 0.41% following ExS treatment. In F. solani, no fluorescence was observed in the negative control (0.01%), but a substantial increase was detected in the positive control and ExS treatment, reaching 62.35% and 60.68%, respectively. In addition, F. oxysporum exhibited no fluorescence in the negative control (0.00%), moderate fluorescence in the positive control (24.70%), and a marked increase to 59.64% with ExS treatment. The findings indicate that ExS induces ROS accumulation to varying degrees across the fungal species analyzed.

Figure 7.

Analysis of ROS induction in C. gloeosporioides, C. scovillei, F. solani, and F. oxysporum in the presence of ExS (200 μg mL⁻¹), using the fluorescent probe H₂DCFDA (20 μM). The analysis was performed after 24 h for all fungal species tested. Negative control: cells treated only with the 2,7‐dichlorofluorescein diacetate probe. Positive control: 30% hydrogen peroxide (H₂O₂) solution was used to validate ROS induction. Bar = 20 μm.

3.8. Vacuolar Structure Analysis

The results demonstrated that treatment with ExS induced significant morphological alterations in F. oxysporum and C. gloeosporioides compared to the negative control. After 24 h of incubation with ExS (200 μg mL⁻¹), F. oxysporum exhibited notable changes in cellular morphology, including enlarged vacuoles and disorganized hyphal structures, as evidenced by the microscopy images. In F. oxysporum, FM4‐64 labeling revealed that vacuoles became smaller and more numerous after treatment, suggesting a significant reorganization of vacuolar structures. In C. gloeosporioides, ExS induced the formation of smaller vacuoles; however, no significant morphological changes were detected in comparison to the control (Figure 8).

Figure 8.

Vacuolar structure analysis of C. gloeosporioides and F. oxysporum after treatment with ExS (200 μg mL⁻¹), visualized using FM4‐64 fluorescent staining. The analysis was performed 24 h after treatment for both fungal species. White arrows indicate vacuolar compartments showing altered size and distribution, whereas black arrows highlight morphological alterations in fungal cells. Negative controls represent untreated cells. Bar = 10 μm.

3.9. Mitochondrial Functionality Analysis

Mitochondrial functionality in C. gloeosporioides was evaluated using JC‐1 staining. Some fungal cells treated with the ExS fraction exhibited increased green fluorescence, indicating a loss of mitochondrial membrane potential compared to the negative control. After 24 h of incubation with ExS (200 μg mL⁻¹), the overlap of images red and green channels confirmed that the treatment induced mitochondrial depolarization, suggesting impaired mitochondrial functionality in the fungal cells (Figure 9).

Figure 9.

Mitochondrial functionality analysis of C. gloeosporioides after treatment with ExS (200 μg mL⁻¹) for 24 h, using JC‐1 (3.85 μM) fluorescent staining. Increased green fluorescence in treated cells indicates mitochondrial membrane depolarization, whereas negative controls (untreated cells) display predominant red fluorescence corresponding to polarized mitochondria. Bar = 10 μm.

Mitochondrial metabolic activity of C. gloeosporioides cells treated with ExS (200 μg mL⁻¹) was evaluated using the WST‐1 reduction assay. The results showed a significant increase in metabolic activity compared to the negative control, which may reflect a compensatory metabolic response to mitochondrial stress rather than increased mitochondrial efficiency (Figure 10).

Figure 10.

After treatment with the extract (200 μg mL⁻¹), the WST‐1 reduction assay was performed to evaluate the metabolic activity of Colletotrichum gloeosporioides cells. The bars represent absorbance values measured at 450 nm for the negative control (untreated cells), the positive control (cells treated with acetic acid), and the extract treatment. The treated group exhibited an increased WST‐1 reduction, indicating preserved mitochondrial and overall cellular dehydrogenase activity, as evidenced by a significant increase in absorbance compared to the negative control. Data represent the mean ± standard deviation of three independent replicates.

4. Discussion

Mass spectrometric analysis of the ExS fraction of C. annuum cv. Carioquinha leaves revealed a predominant peptide band around 6.5 kDa (Figure 1A–B), which is consistent with previous reports on low‐molecular‐weight AMPs [36]. Sequence analysis indicated that CaCHev shares high similarity with HLPs. Comparison with sequences available in the literature revealed that CaCHev shares a highly conserved region with cysteine residues (LCCSQFGFCGTTREYCGAGCQSNCR), consistent with regions typically associated with the CBD found in HLPs. This finding was supported by analysis of other AMPs and proteins. Comparative analysis included the antifungal protein (AAL73184.1), PN‐AMP1 (PHT87891.1), and prohevein sequences from C. chinense (PHU23618.1) and C. annuum (KAF3628901.1). In addition, a chitin‐binding peptide previously isolated from C. chinense Jacq. seeds was used as a reference for sequence comparison with CaCHev (Figure 2).

These results demonstrate that the fragment identified in the ExS fraction (TREYCGPGCQSNCR) was closely aligned with a region of the CaCHev sequence (TREYCGAGCQSNCR), differing by a single amino acid substitution: Alanine (A) in CaCHev was replaced by proline (P) in the ExS fragment. Although both amino acids are nonpolar, proline has a unique cyclic structure that can introduce conformational constraints. Despite this variation, the strong similarity supports the hypothesis that both sequences originate from processing of the same precursor. The presence of HLPs in ExS and in the purified fraction suggests a functional role for these molecules in plant defense. Sequence similarity analysis revealed that the CaCHev peptide exhibited 100% identity with the C‐terminal region of prohevein from C. annuum and C. chinense. Proheveins are high‐molecular‐weight proteins that release peptides following proteolytic processing involving the enzymatic cleavage of prohevein, resulting in the release of specific domains, including the C‐terminal region where CaCHev fits [37].

Consequently, the CaCHev peptide and band present in ExS may represent products derived from the proteolytic maturation of prohevein. These results suggest that the peptides present in ExS and CaCHev can be processed from prohevein, generating smaller fragments that maintain their biological activity. Mass spectrometry confirmed the presence of 15 amino acid residues in ExS and 25 amino acid residues in CaCHev, which correspond to the functionally active region of the peptide. This region is rich in cysteine residues, consistent with the formation of disulfide bridges, a hallmark of HLPs. Furthermore, CaCHev has a conserved sequence, SXXG and GXXXX, associated with the CBD, thus supporting its classification as a chitin‐binding hevein‐like peptide (CB‐HLP). This finding suggests that CaCHev may have a mechanism of action involving interactions with chitin, thereby contributing to the maintenance of the structural stability and biological activity of HLPs [21, 38].

HLPs have previously been described for their antifungal and antibacterial properties in the seeds and leaves of C. chinense and C. annuum. Gonçalves et al. (2024) reported the peptide Cc‐Hev from C. chinense. This protein exhibits high levels of disulfide bonds and high chitin binding affinity, leading to the inhibition of Candida species growth. In C. annuum, HEV‐CANN HLP recovered from leaf tissues were used for the bioinformatics prediction of stable structures with activity against Alternaria solani and some phytopathogenic bacteria [39]. Taken together, these data reinforce the biotechnological prospects of employing HLPs from Capsicum species. Thus, this study focused on expanding current knowledge by analyzing the antifungal activity and possible mechanisms of action of the ExS fraction cultivated from the leaves of C. annuum cv. Carioquinha. Fungal growth inhibition assays were performed to monitor the antifungal potential of ExS in vitro. It is important to note that the biological assays were performed using the ExS fraction rather than the isolated peptide. Therefore, the observed antifungal effects cannot be exclusively attributed to CaCHev, as other bioactive components present in the extract may contribute to the activity.

Structural analysis via AlphaFold prediction (Figure 3A) confirmed that CaCHev has typical features of hevein‐like AMPs, including regions with organized β‐sheets and an α‐helix near the CBD. This primary structure is likely reinforced and globally stabilized by disulfide bridges between the cysteine residues situated at conserved positions. The β‐sheets probably constitute a region that allows functional interaction with chitin, as the peptide′s core is supported by the α‐helix. Notably, the presence of the CBD correlates with antifungal activity, since these domains help anchor chitinous fungal cell walls, which may disrupt membrane integrity. Overall, these features strengthen the proposed mechanism of action of CaCHev and further substantiate its role in targeting fungal pathogens through direct interactions with their cell walls, as observed for other HLPs [10, 21]. The interaction between CBD‐containing peptides and fungal cell walls is not limited to passive binding. It has been proposed that these peptides can disrupt the spatial organization of chitin microfibrils and their association with β‐1,3‐glucans, leading to weakened structural integrity. Additionally, interference with chitin synthase activity or chitin deposition during cell wall expansion has been suggested as a potential antifungal mechanism [40]. Although these mechanisms have been described for purified HLPs, their involvement in the activity of the ExS fraction remains speculative, as other compounds present in the extract may also contribute to the observed effects. Nevertheless, such mechanisms provide a plausible framework for understanding how components of ExS may affect fungi such as C. gloeosporioides.

These additional features of CaCHev add to the belief that it has antimicrobial properties. There was a greater percentage of polar and charged residues, which may favor electrostatic interactions with microbial membranes (Figure 3B). Charge separation of the hydrophobic and hydrophilic amino acids observed in the helical projection indicated an amphipathic profile. This allows for interactions with membranes and possible insertion into membrane dynamics, which is commonly observed in AMPs. Cationic amino acids, such as Arg and His, increase binding to anionic phospholipids, whereas nonpolar Ans may lead to membrane destabilization associated with pore or micelle formation [41]. Conversely, less abundant acid residues can contribute to internal stabilization, likely by forming salt bridges in standard physiological environments.

These findings show that ExS has a strong inhibitory effect, particularly on C. gloeosporioides, where fungal cell growth was reduced by 78%. However, the other fungi tested did not exhibit considerable inhibition (Figure 4). The differential susceptibility observed among fungal species may be associated with variations in cell wall composition and architecture [42]. In filamentous fungi, the relative proportions of chitin, β‐glucans, and associated glycoproteins can vary significantly depending on species and developmental stage [43]. For instance, hemibiotrophic pathogens such as C. gloeosporioides undergo dynamic remodeling of their cell wall during host infection, including modulation of β‐glucan exposure and chitin deacetylation to evade plant immune recognition. These changes may transiently increase the accessibility of chitin or alter wall porosity, potentially enhancing the binding and activity of chitin‐targeting peptides [43, 44]. In contrast, species such as F. oxysporum may exhibit different wall organization, including higher incorporation of glycoproteins and structural polysaccharides that can mask chitin or reduce peptide accessibility.

This aligns with the results of previous studies on the antimicrobial activity of peptides derived from plants, particularly those belonging to the Capsicum genus. For example, Cherene et al. (2023) described the extraction of defensin‐like peptides, lipid transfer proteins (LTPs), and protease inhibitors from C. annuum leaves, which showed biological activity against several Candida species [45]. Aguieiras et al. (2021) reported that C. chinense Jacq. produces AMPs in immature fruits that inhibit C. albicans and C. tropicalis growth [46].

Cell viability analysis revealed that ExS had a pronounced effect on C. gloeosporioides, resulting in reduced fungal proliferation over 48 h, indicating its fungistatic properties (Figure 5A). ExS also reduced cell viability by 36.7%, illustrating its effect on cellular processes (Figure 5B). Cell viability assays measure the operational balance between cellular function and natural reproduction and thus describe the quantitative and qualitative aspects of cell behavior [47]. These measurements are critical for understanding the mechanisms underlying the actions of these cells, with a focus on determining the pathways responsible for living processes or death [48].

Treatment with ExS at 200 μg mL⁻¹ induced significant membrane permeabilization in C. gloeosporioides, resulting in compromised cell integrity and suggesting that membrane‐compromising effects potentially lead to cell death (Figure 6). This finding aligns with the known mechanisms of AMPs, which often target membrane structures. Importantly, membrane susceptibility to AMPs is also influenced by lipid composition. Fungal plasma membranes are rich in ergosterol and negatively charged phospholipids, which facilitate electrostatic interactions with cationic peptides. However, variations in lipid composition, membrane fluidity, and sterol content among fungal species may affect peptide insertion and pore formation. Thus, differences in membrane organization between species such as C. gloeosporioides and F. oxysporum could partially explain the variability in membrane permeabilization and antifungal activity observed in this study [49, 50]. Additionally, the involvement of subcellular organelles such as mitochondria, endoplasmic reticulum, and vacuoles may influence membrane dynamics and contribute to fungal susceptibility. Organelle dynamics and membrane trafficking are critical for the integrity and function of fungal cells [51].

Thus, ExS may target either the structure or a component of the C. gloeosporioides plasma membrane. The positive charge and structure of the peptide likely allow it to interact with the negatively charged surface of the membrane through electrostatic interactions. Therefore, the peptide can bind to the membrane and induce changes that assist in membrane insertion, and may cause the formation of transmembrane pores. Moreover, the peptide may bind to specific membrane components that are necessary for maintaining membrane integrity, such as lipids and membrane‐associated components [51–53]. The current findings align with earlier work that identified various plant peptides, including defensin‐like peptides, thionins, heveins, and protease inhibitors, that show antifungal activity through their membrane‐permeabilizing capacity [54–57]. Our findings indicate the potential of AMPs, particularly those derived from C. annuum. These findings revealed how compounds from C. annuum disrupt fungal membranes, leading to the development of new antifungal strategies.

AMPs may also act on intracellular targets, including induction of oxidative stress. Therefore, we evaluated whether ExS could trigger ROS production as a potential antifungal mechanism. The results demonstrated that ExS (200 μg mL⁻¹) induced oxidative stress in C. gloeosporioides, C. scovillei, F. oxysporum, and F. solani, as indicated by increased fluorescence in hyphae and conidia, suggesting the occurrence of ROS‐mediated cellular damage (Figure 7). Disruption of cell wall integrity or membrane perturbation can activate stress response pathways, leading to mitochondrial dysfunction and ROS accumulation. In fungi, cell wall stress is closely linked to signaling pathways that regulate redox balance and survival. Given the complexity of the ExS fraction, it is likely that multiple components contribute to these effects, either through direct intracellular targeting or secondary responses triggered by membrane or wall‐associated disturbances. Therefore, ROS production observed in this study may reflect consequence of multifactorial stress rather than a single, specific mode of action.

We investigated whether ExS causes apoptosis in C. gloeosporioides, particularly through metacaspases, which are often associated with programmed cell death (PCD). However, the absence of significant activation of the metacaspase pathway suggests that apoptosis may not be the main mechanism underlying ExS‐induced antifungal activity (data not published). Failure to trigger metacaspase activation could imply an increased fungicide specificity of ExS, leading to decreased toxicity to nonfungal cells. Biochemical markers of apoptosis, such as the accumulation of ROS and metacaspase activity, have been employed to test the apoptotic process in fungi [58]. Apoptosis is an automatic cell death mechanism that can be triggered by various factors such as hypoxia, toxins, radiation, ROS, viruses, or other stressors. The cells shrank, and small pieces of chromatin started to group together. In contrast, necrosis is a form of death that is uncontrolled by the cell as a result of external injury and is usually severe enough to rupture the plasma membrane and leak out of the cytosol and/or organelle contents. Characteristics of necrosis include swelling of cells, dilation of organelles, increased membrane permeability, vacuolization, and membrane disruption [59].

Fluorescence microscopy analyses of vacuolar structures revealed significant changes in F. oxysporum and C. gloeosporioides after treatment with ExS (200 μg mL⁻¹) (Figure 8). In both fungi, the presence of smaller and more numerous vacuoles suggests stress‐induced reorganization of vacuolar dynamics, potentially indicating early autophagic responses [60]. A high level of vacuoles was more pronounced in F. oxysporum infected with ExS, which may signal a reactive stress response related to autophagy, a cellular coping mechanism that helps exile exhausted filaments and reprocess damaged structures under adverse conditions. The fungal autophagy described here is classified as PCD induced by stress. This process involves vacuolization at an advanced stage and an increase in the number of vacuoles. Autophagy isolates and recycles damaged cellular parts resulting from stress, and prolonged or chronic stress overrides this system [60, 61]. Fungal vacuoles serve as multifunctional organelles that store ions, such as amino acids, phosphate, and calcium, along with certain metabolic processes, such as the turnover and hydrolysis of glycoproteins, and provide ionic homeostasis, regulation of pH levels, and maintenance of osmotic balance. Recent studies have demonstrated that vacuoles play a central role within fungi, resulting in many changes that can be perceived from outside or inside the cells [62].

Under this framework, it appears that both F. oxysporum and C. gloeosporioides, despite showing limited growth inhibition, hinted at some oxidative processes through vacuole formation induced by ExS, suggesting that the peptides may influence the vacuolar or endocytic pathways. Moreover, the changes associated with vacuolar topology treatment observed via FM4‐64 staining reinforce this theory. Endocytosis is essential for polarized hyphal proliferation and pathogenicity in filamentous fungi [60]. Thus, ExS likely affects one or more major controllers of vesicular trafficking and/or membrane remodeling. Moreover, alongside the oxidative processes that stem from ExS, which were confirmed by ROS assays, there are suggestions of reorganizational impacts as vacuoles exposed by F. oxysporum.

To assess mitochondrial functionality in C. gloeosporioides treated with ExS, JC‐1 staining was performed. JC‐1 is a cationic dye that accumulates in mitochondria depending on the membrane potential (ΔΨm): Healthy, polarized mitochondria display red fluorescence, whereas depolarized mitochondria emit green fluorescence [63]. Mitochondrial dysfunction was observed in hyphae treated with ExS (200 μg mL⁻¹), indicating that it affects intracellular targets (Figure 9). The increase in green fluorescence observed in JC‐1–stained cells suggests port depolarization of the mitochondrial membrane, which is indicative of probable mitochondrial dysfunction and cellular stress. Disruption of mitochondrial potential can impair ATP production and cause elevated levels of ROS, which together lead to a decrease in cell viability and growth inhibition [64, 65]. Similar mitochondrial alterations have been reported for other AMPs that target fungal cells. For example, Zhou et al. (2019) demonstrated that AMP‐induced mitochondrial depolarization in Candida albicans correlated with elevated ROS production and loss of metabolic balance [66].

The WST‐1 assay was also performed to evaluate the metabolic activity of the mitochondria in ExS‐treated cells (200 μg mL⁻¹). WST‐1 reduction is attributed to the action of mitochondrially and cytosol localized NAD(P)H‐dependent oxidoreductases [65, 67]. The ExS was also able to augment the metabolic activity of the sample compared to the negative control, and, therefore, metabolic activity compensation may have occurred. The activity of the dehydrogenases in energy‐producing pathways may trigger the formation of ROS, which leads to decompensated cell homeostasis and potentially expedites the excess activity of metabolism under stressed condition or adaptation to the action of bioactive peptides [64,68,69].

In fungi, the mitochondrial electron transport chain is highly flexible and differs significantly from that of mammalian systems, as it includes alternative pathways such as external and internal NADH dehydrogenases and the alternative oxidase (AOX), which operate in parallel to the classical cytochrome pathway. These alternative components allow electron flow to be maintained even when proton‐pumping complexes are impaired, but without contributing to the generation of a proton gradient across the inner mitochondrial membrane [70].

As a result, cells may exhibit reduced mitochondrial membrane potential (ΔΨm), as observed with JC‐1 staining, while simultaneously maintaining or even increasing electron transport activity, which is detected by WST‐1 through NAD(P)H‐dependent oxidoreductases. This phenomenon is consistent with a state of metabolic uncoupling, in which dissipation of the electrochemical gradient relieves thermodynamic constraints on electron transport, leading to increased respiratory rates and redox activity. Similar mechanisms have been described in fungal species such as Candida, where branched respiratory chains enable metabolic adaptation under stress conditions [71, 72].

In addition, activation of these alternative pathways is often associated with increased production of ROS, either due to electron leakage from impaired complexes or as a consequence of elevated electron flux through the respiratory chain. Therefore, the simultaneous observation of mitochondrial depolarization, increased metabolic activity, and ROS accumulation suggests a stress‐induced metabolic imbalance, in which fungal cells attempt to compensate for mitochondrial dysfunction by enhancing electron transport through alternative routes [72, 73]. Considering that the assays were performed using the ExS fraction, it is likely that multiple components contribute to this effect, potentially acting as mitochondrial stressors or uncoupling agents.

Taken together, these findings suggest that the antifungal activity of ExS is likely driven by a combination of factors, including potential interactions with cell wall components, membrane perturbation, and induction of intracellular stress responses. The presence of a HLP containing a CBD supports a possible contribution to targeting structural components such as chitin. The selective sensitivity observed for C. gloeosporioides highlights the importance of species‐specific differences in cell wall composition, organization, and membrane properties. These factors likely influence the accessibility and efficacy of bioactive compounds present in ExS, reinforcing the need for further studies using isolated molecules to dissect their mechanisms of action.

5. Conclusions

Research into plant peptides may significantly improve the management of phytopathogenic fungi. The ExS fraction obtained from the leaves of C. annuum (L.) cv. Carioquinha possesses a 25‐amino acid long hevein‐like peptide (CaCHev) which may contribute to the observed antifungal activity against C. gloeosporioides and F. oxysporum. AMPs have several ways to prevent damage, including the permeabilization of membranes, oxidative damage, destruction or alteration of vacuolar structures and functions, and reduced mitochondrial functionality. CaCHev is predicted to act similarly to other HLPs as it contains a conserved cysteine‐rich domain associated with chitin binding and structural stability. Peptides such as CaCHev may contribute to the natural defense of plants against pests and diseases and represent promising, nontoxic alternatives for antifungal applications in agriculture and biotechnology. These findings highlight the importance of further studies exploring their potential as integrated disease management strategies.

Author Contributions

The study was conceived by V.M.G. and A.C.O. Experimental procedures were carried out by S.S.S.O., M.B.C., G.B.T., V.B.P., M.P.C.M.S., R.R., and E.O.M. Data analyses were performed by A.C.O., G.B.T., E.O.M., and S.S.S.O. The paper was written by S.S.S.O., M.B.C., and V.M.G. All authors reviewed the manuscript.

Funding

This study was supported by Conselho Nacional de Desenvolvimento Científico e Tecnológico, 10.13039/501100003593, 307590/2021‐6; Fundação Carlos Chagas Filho de Amparo à Pesquisa do Estado do Rio de Janeiro, 10.13039/501100004586, E‐26/200567/2023, E‐26/210353/2022, E‐26/200.127/2023; and Coordenação de Aperfeiçoamento de Pessoal de Nível Superior, 10.13039/501100002322, Finance Code 001.

Ethics Statement

This article does not contain any studies with human or animal subjects.

Conflicts of Interest

The authors declare no conflicts of interest.

de Oliveira, Samuel Salomão Silva , Cherene, Milena Bellei , Taveira, Gabriel Bonan , Pinto, Vitor Batista , da Conceição Moura da Silva, Marcos Paulo , Rodrigues, Rosana , de Oliveira Mello, Érica , de Oliveira Carvalho, André , Gomes, Valdirene Moreira , Characterization and Antifungal Activity of a Hevein‐Like Peptide‐Containing Extract From Capsicum annuum Leaves Against Phytopathogenic Fungi, International Journal of Microbiology, 2026, 7301018, 17 pages, 2026. 10.1155/ijm/7301018

Data Availability Statement

All the data generated or analyzed during this study are included in this published article.