Abstract
Introduction:
The genus Bifidobacterium is a key component of the honey bee gut microbiota, playing a fundamental role in maintaining host health and colony well-being. Alongside other core genera such as Bombilactobacillus, Gilliamella, Lactobacillus, and Snodgrassella, Bifidobacterium contributes to essential functions including nutrient digestion, immune modulation, and protection against pathogens. Among threats to honey bee health, Chalkbrood disease, caused by fungus Ascosphaera apis, remains a major concern due to detrimental effects on colony strength and honey yield.
Materials and methods:
We characterized enzymatic activity and carbohydrate assimilation of nine Bifidobacterium strains isolated from the honey bee intestinal tract. In parallel, we assessed antifungal potential against A. apis strains, focusing on volatile organic compounds (VOCs).
Results and discussion:
Notably, Bifidobacterium asteroides 3CP-2B exhibited enzymatic capabilities supporting digestive functions and metabolism of sugars potentially harmful to honey bees. This strain showed marked antifungal activity against A. apis, mediated by volatile and non-volatile bioactive metabolites. Among VOCs identified, propanoic acid, ethanol, acetic acid, ethyl propionate, and 1-propanol were the most prominent compounds associated with the antifungal effect.
1 Introduction
In recent years, honey bees (Apis mellifera L.) have emerged as an important model system for understanding the functional roles of bacteria within the gut microbiome (, ). However, it remains unclear how specific members of the gut microbiota influence bee health and physiological state (). Honey bee gut is primarily dominated by nine bacterial taxa, which together comprise more than 95% of the total gut microbial community. Among these, five phylogenetic lineages are consistently present in every individual and are defined as the core members of the honey bee gut microbiota. These core lineages represent genus-level taxa from distinct bacterial classes: Gilliamella (Gammaproteobacteria), Snodgrassella (Betaproteobacteria), Lactobacillus Firm-4 (including Bombilactobacillus), Lactobacillus Firm-5 (including Apilactobacillus), and Bifidobacterium (Actinobacteria).
This characteristic taxonomic composition of the microbiota, comprising largely species exclusive to social honey bees, along with their essential biochemical contributions to the host, suggests a highly specialized and co-evolved relationship between microbes and honey bees (, ). A gut microbiota with a balanced composition plays a crucial role in defending against pathogens and parasites, detoxifying foodborne toxins, and regulating the immunity, metabolism, behavior, and development of honey bees. Conversely, dysbiosis of this community can lead to altered gene expression related to these key functions, potentially compromising overall health and well-being ().
The genus Bifidobacterium encompasses Gram-positive bacteria belonging to the family Bifidobacteriaceae within the phylum Actinomycetota (). Bifidobacterium spp. are symbiotic microorganisms that contribute to gastrointestinal homeostasis in humans, animals, and insects; in honey bees, they colonize the gut throughout development, with maximal abundance in the adult hindgut (, –). Although typically less abundant than other core gut taxa, they play a critical role in host metabolism, immune regulation, disease resilience, and adaptation to environmental stressors (, ). To date, multiple Bifidobacterium species have been identified and characterized from the gut microbiota of various honey bee species within the family Apidae (Table 1). Recently, Bifidobacterium favimelis, a novel species isolated from black comb honey of A. mellifera, was identified by Li et al. (). The presence and divergence of Bifidobacterium strains in honey bees is attributed to a long-term coevolutionary process, reflecting their adaptation to various microenvironments within the bee gut and hive, as well as to hive-mediated vertical transmission across generations (–). Populations of bifidobacteria in honey bees have been observed to remain relatively stable over time, suggesting that these microorganisms play a consistent and essential role in host physiology (, ). Strains of Bifidobacterium inhabiting the honey bee gut are of particular interest due to their potential probiotic properties. For example, B. asteroides has been shown to stimulate the production of host-derived hormones, such as prostaglandins and juvenile hormone derivatives, which are known to influence honey bee development (). Comprehensive genomic analyses have revealed that Bifidobacterium species harbor a substantial repertoire of genes involved in carbohydrate metabolism, underscoring their functional role in insect physiology (, , ). Recent studies on pollinator gut microbiota have further elucidated the involvement of bifidobacteria in maintaining immune function, enhancing disease tolerance, and improving resistance to environmental stressors (). The genus Bifidobacterium supports honey bee health through polysaccharide degradation and immune modulation; however, its abundance and overall gut microbiota stability are influenced by factors such as diet, seasonal changes, caste roles, geography, and exposure to xenobiotics like herbicides and antibiotics, which can disrupt microbial balance and lead to dysbiosis, impairing metabolism and vitamin biosynthesis (, , –). Moreover, a disrupted gut microbiota may increase honey bee susceptibility to parasitic infections, including those caused by Nosema spp. and Ascosphaera apis (, –40).
Table 1
| Bifidobacterium species | Host insect | Reference |
|---|---|---|
| Bifidobacterium asteroides | Honeybee (Apis mellifera, Apis cerana) | () |
| Bifidobacterium coryneforme | Honeybee (Apis mellifera, Apis cerana) | () |
| Bifidobacterium indicum | Honeybee (Apis mellifera, Apis cerana) | () |
| Bifidobacterium kimbladii | Honeybee (Apis mellifera) | () |
| Bifidobacterium apicola | Honeybee (Apis mellifera) | () |
| Bifidobacterium apis | Honeybee (Apis mellifera) | () |
| Bifidobacterium apousia | Honeybee (Apis mellifera) | () |
| Bifidobacterium choladohabitans | Honeybee (Apis mellifera) | () |
| Bifidobacterium polysaccharolyticum | Honeybee (Apis mellifera) | () |
| Bifidobacterium mellis | Honeybee (Apis mellifera) | () |
| Bifidobacterium mizhiense | Honeybee (Apis mellifera) | () |
| Bifidobacterium actinocoloniiforme | Bumble bees (Bombus lucorum) | () |
| Bifidobacterium bohemicum | Bumble bees (Bombus lapidarius) | () |
| Bifidobacterium bombi | Bumble bees (Bombus terrestris) | () |
| Bifidobacterium commune | Bumble bees (Bombus hypnorum) | () |
| Bifidobacterium xylocopae | Carpenter bees (Xylocopa violacea) | () |
| Bifidobacterium aemilianum | Carpenter bees (Xylocopa violacea) | () |
Bifidobacterium species isolated from the gut microbiota of different bee species.
Chalkbrood, caused by the fungus A. apis, is a widespread fungal disease that primarily affects developing honey bee brood, especially in A. mellifera colonies, although it can also impact various other bee taxa (41–44). Recent evidence indicates an increasing global incidence of chalkbrood, which is contributing to honey bee population declines and significant reductions in colony productivity (45–48); moreover, it has been shown that chalkbrood infection alters the honey bee gut bacteriome and increases the host’s vulnerability to other pests and pathogens (49–52). A. apis is generally regarded as an opportunistic pathogen that is efficiently dispersed and highly prevalent; however, its mere presence in the hive does not necessarily lead to disease manifestation. Rather, one or more predisposing factors must coincide for a clinical outbreak to occur. These include environmental stressors such as damp and cold weather, colony health status, genetic susceptibility, and developmental stress within the brood (53, 54). Infection is initiated when larvae orally ingest fungal ascospores, which subsequently germinate in the posterior midgut. The resulting hyphae invade the epithelial cells and basement membrane, ultimately leading to larval death. Fungal development continues in a necrotrophic phase even after the host’s demise (55).
Over the years, various chemotherapeutic agents have been investigated for their efficacy against A. apis (53, 56), but, none have proven effective in preventing chalkbrood, despite their antifungal activity. Moreover, the presence of antifungal residues in honey represents a potential health hazard for consumers (57). Consequently, there is a growing demand for eco-friendly, and sustainable alternatives for disease control (58–63). In this context, the use of microbial resources as biocontrol agents against honey bee pathogens, including A. apis, offers promising opportunities (64–66). Several studies have demonstrated that Apilactobacillus kunkeei and Lactiplantibacillus plantarum, isolated from the honey bee gut, can inhibit A. apis mycelial growth in vitro, suggesting their potential as prophylactic agents to restore and maintain gut microbial balance (56, 67). Similarly, other beneficial microbes have shown effectiveness in the biocontrol of chalkbrood (68, 69). Notably, Daisley et al. (70) demonstrated that hive treatments with a probiotic formulation containing L. plantarum, Lacticaseibacillus rhamnosus, and A. kunkeei exerted strong antifungal effects against A. apis while also promoting the recovery of symbiotic gut communities. These microbiome shifts were positively correlated with enhanced brood production and colony development (70).
To date, there is limited research on the use of B. asteroides as an anti-fungal or probiotic agent in beekeeping (71). In a study by Alberoni et al. (), symbiotic species including B. asteroides, B. coryneforme, and B. indicum were shown to enhance colony productivity when administered in sugar syrup as a dietary probiotic. Additionally, Bifidobacterium spp. supplementation led to reduced Nosema infection rates in honey bee colonies (). More recently, Dengiz et al. (72) reported significant antimicrobial activity by B. asteroides, B. choladohabitans, and B. polysaccharolyticum against key bee pathogens such as Paenibacillus larvae, Melissococcus plutonius, and Serratia marcescens. Conversely, a Bifidobacterium bifidum strain, isolated from human feces, did not exhibit inhibitory effect on A. apis mycelial growth (73). This finding supports the growing consensus that exogenous probiotics, not derived from the honey bee microbiota, may lack beneficial effects, or even pose risks, to health (, 74–77). Therefore, the identification and characterization of autochthonous Bifidobacterium strains with honey bee-specific probiotic properties is essential for developing effective, safe, and sustainable tools for disease prevention in apiculture.
In the present study, a preliminary characterization of Bifidobacterium strains isolated from the gastrointestinal tract of A. mellifera (collected from apiaries in central-southern Italy) was conducted, including analyses of enzymatic activity and carbohydrate assimilation profiles. Furthermore, the antifungal activity of these strains against multiple A. apis isolates was evaluated, with particular focus on the production of volatile organic compounds (VOCs). This is the first report describing the potential of B. asteroides as a biocontrol agent against chalkbrood disease.
2 Materials and methods
2.1 Fungal cultures
Table 2 provides a detailed overview of the A. apis strains employed in the present study.
Table 2
| Fungal strain ID | Taxonomical identification | Accession number |
|---|---|---|
| 1A1R 2.2 | Ascosphaera apis | PV056024 |
| 1B3R (1) | Ascosphaera apis | PV056025 |
| 1A1R 1.1 | Ascosphaera apis | PV056026 |
| 1A3R (2) | Ascosphaera apis | PV056027 |
| 1B1R (1) | Ascosphaera apis | PV056028 |
| 1A3R 1.1 | Ascosphaera apis | PV056029 |
| CB2 | Ascosphaera apis | PV056030 |
| CB3 | Ascosphaera apis | PV056031 |
| 1A1R 1.2 | Ascosphaera apis | PV056032 |
| AA | Ascosphaera apis | PV056033 |
| CB1 | Ascosphaera apis | PV056034 |
| 1A2R 1.2 | Ascosphaera apis | PV056035 |
| 1B2R 2.2 | Ascosphaera apis | PV056036 |
| 1B2R 2.1 | Ascosphaera apis | PV056037 |
| CB4 | Ascosphaera apis | PV056038 |
Catalogue of A. apis strains utilized in this study, accompanied by their GenBank accession numbers from the National Center for Biotechnology Information (NCBI).
2.2 Isolation of bifidobacteria
Worker bees (A. mellifera subsp. mellifera) were collected from managed apiaries in the Molise and Campania regions (central-southern Italy). After euthanization by rapid cooling on ice, bees were transported under refrigeration to the laboratory on the same day and stored at −80 °C until DNA extraction. Dissection of the intestinal tract was performed under sterile conditions using stainless-steel scissors. Entire guts were placed in sterile glass Petri dishes with physiological saline solution (0.9% NaCl) and homogenized. Serial dilutions of the homogenates were plated on Bifidobacterium Selective Medium agar (BSM; Sigma-Aldrich) and incubated at 37 °C for 72 h under anaerobic conditions (Anaerogen system, Oxoid, Milan, Italy). Colonies showing Gram-positive staining with characteristic bifurcated (Y- or V-shaped), club-shaped, or spatula morphologies were presumptively identified as Bifidobacterium.
2.3 Genotypic characterization
Genomic DNA (gDNA) was extracted using the Bacterial Genomic DNA Isolation Kit (Norgen Biotek, Thorold, ON, Canada) according to the manufacturer’s instructions. The 16S rRNA gene was amplified by PCR using universal primers 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1492R (5′-TACGGTTACCTTGTTACGACTT-3′). Each 20 µL reaction included 1× Master Mix (Norgen Biotek), 2.5 µM of each primer, and 10 ng of template DNA. Negative controls using Milli-Q water were included. PCR was performed using a Mastercycler Nexus (Eppendorf, Hamburg, Germany) with cycling parameters: 95 °C for 5 min; 35 cycles of 95 °C for 30 s, primer-specific annealing temperature for 1 min, 72 °C for 1.5 min; followed by a final extension at 72 °C for 5 min. Products were resolved on a 1% (w/v) agarose gel in 1× TAE buffer, visualized under UV light (Bio-Rad, Hercules, CA, USA), and compared against a 1 kb DNA ladder (Norgen Biotek). Amplicons were purified using the QIAquick PCR Purification Kit (QIAGEN GmbH, Hilden, Germany) and sequenced (Eurofins MWG Biotech, Ebersberg, Germany). Sequences were analyzed using BLAST (78) against the NCBI nucleotide database (NCBI 79). Strains with ≥98% identity were assigned species-level taxonomy (80).
2.4 Biochemical characterization
2.4.1 Carbohydrate assimilation patterns
Carbohydrate utilization was tested using Fermentation Broth Base (FBB; Biolife, Milan, Italy) supplemented with bromocresol purple as a pH indicator. Prior to testing, bacterial strains were cultured in BSM broth at 37 °C for 48 hours under anaerobic conditions. Cultures were centrifuged, and the cell pellet washed with 0.9% NaCl solution to remove the residual medium. The pellet was then resuspended in saline to reach a standard turbidity of 0.5 McFarland (approx. 1.5 × 108 CFU/mL) (81), and used as inoculum. Thirteen carbohydrates were tested: D-arabinose, fructose, galactose, glucose, lactose, maltose, mannose, melezitose, melibiose, raffinose, rhamnose, sucrose, and xylose. For each assay, 4.5 mL of FBB were mixed with 500 μL of carbohydrate solution and 100 μL of bacterial suspension. Negative controls were prepared identically but without inoculum. Sugar solutions were sterilized using 0.22 μm syringe filters. Assays were incubated at 37 °C for 48 hours under anaerobic conditions. The change of color from purple to yellow is due to the production of acids during fermentation. All tests were performed in triplicate for each strain–carbohydrate combination.
2.4.2 Enzymatic profile
Enzymatic activity was evaluated using the API ZYM system (BioMérieux, Lyon, France). The cell pellet (CP), prepared as above, was resuspended in 0.9% NaCl solution to achieve a turbidity of 5 McFarland. Wells of the API ZYM strip were inoculated with 65 µL of this suspension and incubated at 37 °C. After 4 hours, enzymatic activity was assessed based on color change, according to the manufacturer’s guidelines.
2.4.3 Biogenic amine production
Biogenic amine production by Bifidobacterium strains was qualitatively assessed using method from Torracca et al. (82), with slight modifications. The Bifidobacterium strains were grown in BSM broth at 37 °C for 48 h anaerobically. Subsequently, the cultures, in a volume of 50 µL, were inoculated using a spot inoculation method onto solid media formulated with the following components: 0.5% tryp-tone, 0.5% yeast extract, 0.5% meat extract, 0.25% NaCl, 0.05% glucose, 0.1% Tween 80, 0.02% MgSO4, 0.005% MnSO4, 0.004% FeSO4, 0.2% ammonium citrate, 0.001% thia-mine, 0.2% K2HPO4, 0.01% CaCO3, 0.005% pyridoxal-5-phosphate, and 1.5% agar. The medium was supplemented with 1% of each amino acid precursor: L-histidine, L-tyrosine, L-lysine monohydrate, and L-ornithine monohydrochloride. Bromocresol purple (0.006%) was incorporated as a pH indicator, and the medium pH was adjusted to 5.3 prior to inoculation. Petri dishes were incubated at 37 °C for 72 hours under anaerobic conditions. The decarboxylation of the amino acids to the corresponding biogenic amines results in an increase in pH, detected by the culture medium color change. A purple coloration indicated the production of histamine, cadaverine, or putrescine, while medium de-colorization suggested tyramine production. Negative controls lacking amino acid precursors were included to confirm the specificity of the reactions. All tests were conducted in triplicate and all reagents were purchased from Merck KGaA (Darmstadt, Germany).
2.5 Antifungal activity assessment
2.5.1 Preliminary evaluation of antifungal activity
Antifungal activity of Bifidobacterium strains was assessed using a method adapted from Iorizzo et al. (69). Three matrices were tested: broth culture (BC), cell-free supernatant (CFS), and CP. Bifidobacteria were grown in BSM broth at 37 °C for 48 hours under anaerobic conditions to a final cell density of 108 CFU/mL. BC was collected without further treatment. For CFS, 5 mL of bacterial culture was centrifuged at 8000 rpm for 15 min at 4 °C, and the supernatant filtered through a 0.22 μm cellulose acetate membrane. CP was prepared by washing and resuspending the pellet in 5 mL of sterile distilled water. Antifungal assays were conducted by transferring a 6 mm mycelial disc of A. apis, pre-cultured on Sabouraud Dextrose Agar (SDA) at 30 °C for 3 days, to the center of 90 mm Petri dishes containing fresh SDA. In different plates containing the pathogenic fungus, 5 mL of each matrix (BC, CFS, or CP) was added alternately. An SDA plate containing only the pathogenic fungus was used as a control. All plates were incubated aerobically at 30 °C, and each experimental condition was tested in triplicate. Following six days of incubation, the radial growth of A. apis mycelium was measured using a digital caliper. The percentage of mycelium radial growth inhibition (% I) was calculated according to the formula: % I = [(C – T)/C] × 100 (83), where C represent the radial growth in the control, and T represents the radial growth in the presence of different matrices obtained from the Bifidobacterium cultures.
2.5.2 Antifungal activity of the VOCs produced by Bifidobacterium
The antifungal activity of VOCs produced by Bifidobacterium strains was evaluated using a modified double-dish system (DDS) based on Ruiz-Moyano et al. (84). A 100 μL aliquot of a 48-hour culture (108 CFU/mL) was spread on BSM agar in 90 mm Petri dishes. Simultaneously, a 6 mm disc of A. apis mycelium (grown on SDA) was placed in the center of a separate plate. Lids of both plates were removed and the dishes sealed together in an inverted DDS configuration using Parafilm (Pechiney Plastic Packaging Co., Milwaukee, WI, USA), with the fungal plate on the bottom. This setup allowed VOCs to diffuse freely. Control DDS setups (no bacteria) were included. After 6 days at 30 °C, radial growth inhibition was measured as previously described. All experiments were performed in triplicate.
2.6 Volatile organic compounds profiling
2.6.1 VOCs extraction
VOCs were extracted using headspace solid-phase microextraction (HS-SPME) (Figure 1). B. asteroids 3CP-2B was cultured for 72 h at 30°C in BSM medium (15 mL) directly in 30 mL screw capped SPME vials (Agilent Technologies, Santa Clara, CA, USA). The vials were sealed with a magnetic screw capped PTFE/silicone liner septum. After 72 hours, the vial was equilibrated to 40 °C for 15 minutes to allow equilibration of the headspace. A 2 cm DVB/CAR/PDMS fiber (50/30 µm; Supelco, Bellefonte, PA, USA) was then inserted into the vial headspace and exposed at 40 °C for 30 minutes to adsorb volatile compounds. Following adsorption, the fiber was immediately transferred to the GC injector port, where desorption was performed at 240 °C for 10 minutes in splitless mode. To distinguish bacterial VOCs from background volatiles, control vials containing non-inoculated media were processed in parallel. Blank runs were also conducted between samples to confirm the absence of carryover or contamination throughout the extraction and analytical procedures. The experiment was done in triplicate.
Figure 1
2.6.2 Gas chromatography–mass spectrometry analysis
Analysis of volatile organic compounds was conducted using a GC–MS system consisting of an Agilent 7890A gas chromatograph coupled with a 5975A mass selective detector (Agilent Technologies, Santa Clara, CA, USA). Chromatographic separation was achieved using a polar HP-Innowax capillary column (30 m × 0.25 mm i.d., 0.50 μm film thickness; Agilent Technologies). The method was adapted from Serradilla et al. (85), with the oven temperature program set as follows: initial hold at 40 °C for 3 minutes; ramp to 150 °C at 4 °C/min, held for 1 minute; then increased to 220 °C at 3 °C/min with a final hold for 2 minutes. Helium was employed as the carrier gas at a constant flow rate of 1.0 mL/min. The injector was operated in splitless mode, and the desorbed VOCs were introduced directly into the ion source. Electron impact (EI) ionization was performed at 70 eV. The ion source and quadrupole temperatures were set to 230 °C and 150 °C, respectively. Mass spectra were acquired in full-scan mode over a range of m/z 30–300. Compound identification was performed by comparing the obtained mass spectra and retention indices (linear retention indices, LRI) with entries in the NIST05 and Wiley07 spectral libraries. When available, identification was further confirmed using authentic standards. Semi-quantitative analysis of each VOC was expressed as the relative peak area (RPA%), calculated as the ratio of the individual compound’s area to the total area of all detected VOCs in the total ion chromatogram (TIC).
2.7 Statistical analysis
Statistical analyses were conducted in Rstudio (R version 4.3.0). Results from triplicate experiments were expressed as mean ± standard deviation (SD). One-way ANOVA followed by Tukey’s post hoc test was used to determine significant differences (p < 0.05).
3 Results
3.1 Taxonomical identification
The nine isolates have been identified as members of the species B. asteroides, B. apousia, B. mizhiense and B. choladohabitans as reported in Table 3.
Table 3
| Bacterial strain ID | Taxonomical identification | % identity | Accession number |
|---|---|---|---|
| 1CP-1B-B | B. apousia | 99.67 | PV053127 |
| 3CP-6B | B. apousia | 99.04 | PV053134 |
| 3CP-10B1 | B. asteroides | 99.47 | PV053124 |
| 3CP-2B | B. asteroides | 99.73 | PV053131 |
| 3CP-8BG1 | B. asteroides | 99.38 | PV053133 |
| 3CP-3B | B. asteroides | 98.99 | PV053126 |
| 3CP-1G | B. choladohabitans | 99.41 | PV053144 |
| 1CP-3BG | B. mizhiense | 99.83 | PV053125 |
| 1CP-10B | B. mizhiense | 99.43 | PV053128 |
List of Bifidobacterium strains isolated in this study with their taxonomic assignment and NCBI GenBank accession number.
3.2 Biochemical characterization
The enzymatic activities of the bacterial isolates were comprehensively assessed using the qualitative API ZYM kit, with results summarized in Table 4. The enzymatic profiles of the Bifidobacterium isolates, including B. asteroides, B. apousia, B. mizhiense, and B. choladohabitans, revealed activities for several enzymes, notably leucine arylamidase, naphthol-AS-BI-phosphohydrolase, β-galactosidase, and β-glucosidase. Enzymatic activities varied within the B. asteroides species. For example, B. asteroides 3CP-3B exhibited α-galactosidase activity, which was absent in other strains of the same species. Additionally, strains 3CP-10B1S and 3CP-8BG1B were the only ones to display α-glucosidase activity. Strains 3CP-10B1, 3CP-3B, and 3CP-8BG1B also showed α-mannosidase activity, while strain 3CP-2B was unique in exhibiting α-fucosidase activity. Among B. apousia strains, 1CP-1B-B was distinct in showing both esterase and α-glucosidase activities. In contrast, B. apousia 3CP-6B-B was the only strain within its species to demonstrate α-mannosidase activity. No enzymatic differences were observed between the two B. mizhiense strains.
Table 4
| Enzyme | Bifidobacterium asteroides strains | Bifidobacterium apousia strains | Bifidobacterium mizhiense strains | Bifidobacterium choladohabitans | |||||
|---|---|---|---|---|---|---|---|---|---|
| 3CP-10B1S | 3CP-3B | 3CP-2B | 3CP-8BG1B | 3CP-6B-B | 1CP-1B-B | 1CP-3BGS | 1CP-10B | 3CP-1G | |
| Alkaline phosphatase | – | – | – | – | – | – | – | – | – |
| Esterase (C4) | – | – | – | – | – | + | – | – | + |
| Esterase lipase (C8) | – | – | – | – | – | – | – | – | – |
| Lipase (C14) | – | – | – | – | – | – | – | – | – |
| Leucine arylamidase | + | + | + | + | + | + | + | + | + |
| Valine arylamidase | – | – | – | – | – | – | – | – | – |
| Cystine amyralidase | – | – | – | – | – | – | – | – | + |
| Trypsin | – | – | – | – | – | – | – | – | – |
| α-chymotrypsin | – | – | – | – | – | – | – | – | – |
| Acid phosphatase | + | + | + | + | + | + | + | + | – |
| Naphthol-AS-BI-phosphohydrolase | + | + | + | + | + | + | + | + | + |
| α-galactosidase | – | + | – | – | + | + | + | + | + |
| β-galactosidase | + | + | + | + | + | + | + | + | + |
| β-glucuronidase | – | – | – | – | – | – | – | – | – |
| α-glucosidase | + | – | – | + | – | + | – | – | + |
| β-glucosidase | + | + | + | + | + | + | + | + | + |
| N-acetil-β-glucosaminidase | – | – | – | – | – | – | – | – | – |
| α-mannosidase | + | + | – | + | + | – | + | + | + |
| α-fucosidase | – | – | + | – | – | – | + | + | + |
Enzymatic profiles of the nine Bifidobacterium strains assessed using the API ZYM system.
3.3 Carbohydrate assimilation profiles and biogenic amines production
The carbohydrate assimilation abilities of the nine Bifidobacterium strains are detailed in Table 5. All strains were capable of metabolizing fructose, glucose, maltose, melibiose, raffinose, and sucrose. In contrast, none of the strains metabolized D-arabinose or rhamnose. Assimilation of lactose was strain-specific and limited to B. apousia 1CP-1B-B, B. mizhiense 1CP-10B, and B. choladohabitans 3CP-1G. Regarding the production of biogenic amines, none of the strains were able to synthesize these compounds from the tested amino acid precursors.
Table 5
| Carbohydrate | Bifidobacterium asteroides | Bifidobacteria apousia | Bifidobacteria mizhiense | Bifidobacterium choladohabitans | |||||
|---|---|---|---|---|---|---|---|---|---|
| 3CP-10B1S | 3CP-3B | 3CP-2B | 3CP-1G | 3CP-6B-B | 1CP-1B-B | 1CP-3BGS | 1CP-10B | 3CP-1G | |
| D-Arabinose | - | - | - | - | - | - | - | - | – |
| Fructose | + | + | + | + | + | + | + | + | + |
| Galactose | + | + | + | + | + | + | + | + | + |
| Glucose | + | + | + | + | + | + | + | + | + |
| Lactose | - | - | - | - | - | + | - | + | + |
| Maltose | + | + | + | + | + | + | + | + | + |
| Mannose | + | + | + | + | + | - | + | - | + |
| Melezitose | + | + | + | + | + | + | + | + | – |
| Melibiose | + | + | + | + | + | + | + | + | + |
| Raffinose | + | + | + | + | + | + | + | + | + |
| Rhamnose | - | - | - | - | - | - | - | - | – |
| Sucrose | + | + | + | + | + | + | + | + | + |
| Xylose | + | + | + | + | + | + | + | + | + |
Carbohydrate assimilation profiles of the nine Bifidobacterium strains.
(+ positive; – negative).
3.4 Screening of antifungal activity by Bifidobacterium strains
Table 6 presents the antifungal activity of the nine Bifidobacterium isolates against A. apis CB3. In most cases, the use of unprocessed culture broth BC resulted in complete inhibition (100%) of fungal growth. Notable exceptions were isolates 3CP-8BG1B and 3CP-6B-B, which exhibited inhibition rates of 76.7% and 98.9%, respectively. The CFS also showed strong antifungal activity, with inhibition percentages ranging from 65.9% (B. asteroides 3CP-8BG1B) to 100% (B. asteroides 3CP-2B). Regarding the CP fraction, B. mizhiense 1CP-10B exhibited the lowest inhibition (35%), whereas strains 3CP-2B and 1CP-3BGS achieved full inhibition (100%). In terms of VOCs, overall inhibition levels were modest. However, strains 3CP-2B and 3CP-8BG1B showed moderate activity, with inhibition values of 54.2% and 47.2%, respectively.
Table 6
| Bifidobacterium strains | |||||||||
|---|---|---|---|---|---|---|---|---|---|
| Matrices | 3CP-10B1S | 3CP-3B | 3CP-2B | 3CP-8BG1B | 3CP-6B-B | 1CP-1B-B | 1CP-3BGS | 1CP-10B | 3CP-1G |
| BC | 100.0 ± 0.0a | 100.0 ± 0.0a | 100.0 ± 0.0a | 76.7 ± 0.1c | 98.9 ± 0.1b | 100.0 ± 0.0a | 100.0 ± 0.0a | 100.0 ± 0.0a | 100.0 ± 0.0a |
| CFS | 75.0 ± 0.6c | 77.2 ± 0.6b | 100.0 ± 0.0a | 65.9 ± 0.7e | 71.7 ± 0.6d | 75.3 ± 0.7c | 70.9 ± 0.9d | 67.5 ± 0.3e | 67.3 ± 0.5e |
| CP | 52.8 ± 0.6c | 47.5 ± 0.5e | 100.0 ± 0.0a | 54.7 ± 0.8b | 46.1 ± 0.5e | 49.2 ± 0.4d | 100.0 ± 0.0a | 35.0 ± 0.6f | 49.1 ± 0.3d |
| VOCs | 11.9 ± 0.7f | 38.2 ± 0.6c | 54.2 ± 0.3a | 47.2 ± 0.6b | 6.1 ± 0.5h | 24.9 ± 0.4e | 8.3 ± 0.5g | 33.9 ± 0.6d | 6.1 ± 0.5h |
Inhibitory effects (%) of Bifidobacterium strains against A. apis CB3 using different matrices: broth culture (BC), cell-free supernatant (CFS), cell pellet (CP), and volatile organic compounds (VOCs).
Subsequently, B. asteroides 3CP-2B, the most effective strain, was selected as the reference bacterium for further antifungal assays against all A. apis strains listed in Table 2.
The inhibitory activity of B. asteroides 3CP-2B against A. apis strains was assessed under four different treatments: BC, CFS, CP, and VOCs (Figure 2). Overall, the BC treatment exhibited the highest and most consistent antifungal efficacy, with most strains achieving complete inhibition. The CFS treatment also resulted in high inhibition levels, ranging from 88.0% (1A1R 1.2) to 100.0% (1A3R 1.1, 1A2R 1.2, 1A3R (2) and CB1). The CP treatment led to moderately reduced inhibition (from 85.6% to 93.3%). VOCs were the least effective treatment, with inhibition ranging widely from 6.2% (CB1) to 72.8% (CB3), and statistically significant differences among nearly all strains.
Figure 2
3.5 Profiling of volatile organic compounds
Supplementary Table 1 lists all 37 VOCs detected by GC–MS analysis, along with their semi-quantitative relative peak area (RPA%) data. Based on peak areas, the major compounds detected were propanoic acid (45.8%), ethanol (25.0%), acetic acid (17.3%), ethyl propionate (3.1%), 1-propanol (2.3%), isoamyl alcohol (1.7%), propyl propionate (0.7%), ethyl acetate (0.4%), butanoic acid (0.3%), benzaldehyde (0.2%), and 2-methyl-propanoic acid (0.2%) (Table 7).
Table 7
| Volatile compounds | Code | %RPA | AKIsp/KIt | BID |
|---|---|---|---|---|
| Aldehydes | ||||
| Benzaldehyde | Ald2 | 0.2 | 1518/1520 | RI/MS/S |
| Esters | ||||
| Ethyl acetate | E1 | 0.4 | 860/863 | RI/MS/S |
| Ethyl propionate | E2 | 3.1 | 942/946 | RI/MS/S |
| Propyl propionate | E3 | 0.7 | 1045/1047 | RI/MS |
| Alcohols | ||||
| Ethanol | Alc1 | 25.0 | 933/934 | RI/MS/S |
| 1-Propanol | Alc3 | 2.3 | 1033/1037 | RI/MS/S |
| Isoamyl alcohol | Alc6 | 1.7 | 1212/1215 | RI/MS/S |
| Acids | ||||
| Acetic acid | A1 | 17.3 | 1448/1445 | RI/MS/S |
| Propanoic acid | A2 | 45.8 | 1530/1534 | RI/MS |
| 2-Methyl-propanoic acid | A3 | 0.2 | 1581/1581 | RI/MS/S |
| Butanoic acid | A4 | 0.3 | 1632/1630 | RI/MS/S |
Main Volatile Organic Compounds (VOCs) produced by Bifidobacterium asteroides 3CP-2B detected by HS-SPME/GC-MS with corresponding %RPA.
Mean values of 3 samples are calculated as RPA (%). aRIsp: Relative retention indices calculated against n-alkanes (C8–C20) on HP-Innowax column; RIt: Relative retention indices on polar column reported in literature bIdentification method as indicated by the following: RI: Kovats retention index on a on HP-Innowax column; MS: NIST and Wiley libraries spectra; S: co-injection with authentic standard compounds, where commercially available, on the HP-Innowax column. For each metabolite the coefficient of variability of determinations, evaluated as relative standard deviation, was in all cases <10%.
4 Discussion
The isolated microbial cultures were identified as members of the species B. asteroides, B. apousia, B. mizhiense, and B. choladohabitans.
Among these, B. asteroides is of particular interest due to its previously reported oxygen tolerance and its role in carbohydrate metabolism (86). This species metabolizes dietary sugars, including glucose and fructose, and utilizes the malolactic fermentation pathway to convert malic acid into lactate, thereby contributing to the host’s energy metabolism (86). However, functional data on B. apousia and B. mizhiense remain still scarce. Based on the known metabolic capabilities of other Bifidobacterium species, these isolates are hypothesized to play a significant role in sugar degradation, with B. apousia potentially involved in hemicellulose breakdown (). The balance of the honey bee gut microbiota is crucial for host health, with microbial enzymatic activity directly supporting digestive function (87, 88). Notably, the B. apousia strain 1CP-1B-B and B. choladohabitans 3CP-1G showed esterase activity; a function involved in lipid digestion, and playing a role in detoxification by hydrolyzing or degrading various compounds, including drugs, pesticides, and other xenobiotics (89–92). Honey bees employ a multifaceted detoxification strategy, including enzymatic processes such as those involving cytochrome P450 monooxygenases, glutathione S-transferases, and carboxylesterases. These enzymatic defenses are complemented by behaviors forming a “social detoxification system,” which includes forager discrimination, dilution through pollen mixing, and colony-level food processing via microbial fermentation, reducing the intake of harmful chemicals (93–96, 46). Given the widespread use of insecticides in agriculture, supplementing the honey bee diet with appropriate probiotics, capable of degrading such compounds, may benefit bee health (64, 97, 98).
All tested isolates exhibited leucine arylamidase activity, suggesting a common ability to participate in protein hydrolysis, consistent with other Bifidobacterium species (99). This enzymatic function complements the proteolytic capabilities of other core honey bee gut microbes, such as Snodgrassella alvi and Gilliamella apis (, 100). Positive activities were also recorded for acid phosphatase, naphthol-AS-BI-phosphohydrolase, β-galactosidase, and β-glucosidase. Among these, β-glucosidase is particularly important for degrading plant-derived polysaccharides such as cellulose and hemicellulose (, 101), while β-galactosidase catalyzes the hydrolysis of β-D-galactosides, contributing to the digestion of galactose-containing nectar compounds (). α-Glucosidase activity was detected in B. asteroides 3CP-10B1S and 3CP-8BG1B, B. apousia 1CP-1B-B, and B. choladohabitans 3CP-1G. This enzyme, also secreted by the hypopharyngeal glands of honey bees (52, 102), plays a critical role in maltose hydrolysis and starch degradation (, 103, 104), thus contributing to the conversion of nectar into honey.
Additional enzymatic functions, including α-galactosidase, α-mannosidase, and α-fucosidase were detected in certain isolates. Although less studied in honey bee-associated Bifidobacterium, these enzymes are involved in degrading complex plant oligosaccharides and polysaccharides. They may contribute to digestion in insects and produce prebiotic compounds that support immune modulation in mammals, including humans (105–107). For example, α-galactosidase breaks down complex carbohydrates such as raffinose and stachyose, important for nutrient absorption in insects (108). α-Fucosidase releases terminal fucose residues, which are key to cell–cell communication and host–microbe interactions in mammals (109, 110). α-Mannosidase hydrolyzes mannose-containing carbohydrates and helps produce prebiotic mannooligosaccharides, which promote the growth of beneficial gut bacteria (111).
Previous studies have suggested that the honey bee gut microbiome may facilitate the metabolism of toxic sugars (112–114). In our study, carbohydrate assimilation profiles revealed both intra- and inter-species variability. All isolates effectively utilized a range of mono- and oligosaccharides commonly found in the honey bee gut, including fructose, glucose, maltose, melezitose, melibiose, raffinose, and sucrose. These results align with earlier findings showing that Bifidobacterium species are well adapted to the bees’ carbohydrate-rich diet (, 86). Some sugars present in the honey bee diet, such as galactose, mannose, lactose, raffinose, and xylose, can be toxic due to the absence of necessary host enzymes for their degradation (, 114–116). Gut symbionts enhance the honey bee’s ability to process complex polysaccharides and detoxify harmful sugars, improving dietary efficiency and resistance to diseases (101, 117).
Regarding antifungal activity, our results demonstrated that the different matrices (BC, CFS, CP, and VOCs) derived from the evaluated Bifidobacterium strains were effective (Table 6). Variability in antifungal activity likely reflects differences in the types and quantities of antifungal metabolites produced (118). These metabolites, such as lactic acid, acetic acid, phenyl lactic acid (PLA), short-chain fatty acids (SCFAs), proteins, and others, can disrupt fungal cell membranes, causing damage and inhibiting growth (119; 72, 120–122). The antifungal effects of VOCs are mainly attributed to cell wall and membrane disruption, leakage of intracellular contents, and the induction of oxidative stress (123). Bifidobacteria degrade hexose sugars via the “bifid shunt” pathway, in which fructose-6-phosphoketolase (EC 4.1.2.2) plays a key role and serves as a taxonomic marker for the Bifidobacteriaceae family. This pathway typically yields 3 moles of acetate and 2 moles of lactate per 2 moles of glucose, though other byproducts, such as ethanol, can also be produced (124). Ethanol and acetic acid are known for their antimicrobial properties, including antifungal activity, through mechanisms such as membrane disruption, protein denaturation, and interference with fungal DNA and protein synthesis (125, 126). Similarly, 1-propanol and other alcohols (e.g., isoamyl alcohol, 1-butanol) exhibit antifungal effects likely through membrane disruption, inhibition of spore germination, and interference with transcription and translation processes (127, 128).
In our study, B. asteroides 3CP-2B produced abundant propanoic and butanoic acids, confirming that Bifidobacteria are effective SCFA producers (129, 130). These compounds increase membrane fluidity, causing leakage of intracellular contents and ultimately cell death (131). Propionic acid, in particular, generates reactive oxygen species (ROS), reduces ATP levels, and activates metacaspases, leading to mitochondrial-mediated apoptosis in fungal cells (132). Notably, propionic acid has been identified as a natural constituent of honey, contributing to its flavor and preservation (133).
Esters such as ethyl and propyl propionate have demonstrated antifungal activity (84), while other VOCs, like dimethyl disulfide and limonene, were also detected. Limonene damages fungal hyphae, causing cytoplasmic granulation, membrane detachment, and vacuole formation, ultimately leading to cell death (134, 135). Dimethyl disulfide exhibits antifungal activity by damaging membranes and inhibiting spore germination and hyphal growth (136, 137). B. asteroides 3CP-2B also produces methylpyrazines, aromatic hydrocarbons commonly found in foods and considered safe (138, 139). Pyrazine derivatives have broad biological activity, including antifungal effects (138, 140, 141). Gong et al. (142) demonstrated that methylpyrazine and dimethyl disulfide significantly inhibit fungal growth and spore germination. Transcriptome analysis showed that these VOCs downregulate ribosomal synthesis genes, activate the proteasome system, and suppress genes related to spore development, membrane synthesis, mitochondrial function, and toxin production. Exploring natural antifungal strategies may offer sustainable options for improving bee health. Microbial VOCs can be delivered using formulations designed to overcome their volatility and short lifespan. Recent studies have investigated the use of hydrogels and sprays containing microbial VOCs to control plant pathogenic fungi (123, 143). Similarly, antifungal hydrogel or spray formulations based on symbiotic bacteria like B. asteroides could represent a promising, eco-friendly strategy to manage fungal diseases such as Chalkbrood in honey bee colonies.
5 Conclusions
This study contributes to our understanding of the intricate relationship between honey bees and their gut microbiota. Through a preliminary characterization of Bifidobacterium strains isolated from the honey bee gut, we have demonstrated that certain isolates possess enzymatic activities involved in the detoxification of xenobiotics through hydrolysis or breakdown of harmful compounds. Additionally, several strains exhibited enzymatic capabilities that enhance nutrient bioavailability and facilitate the metabolism of specific sugars, such as mannose, lactose, raffinose, and xylose, that can otherwise be toxic to bees. Notably, B. asteroides 3CP-2B exhibited strong antifungal activity, suggesting its potential application as a probiotic supplement in honey bee diets or as an environmentally friendly biocontrol agent to reduce the incidence of fungal diseases such as chalkbrood. These findings lay a solid foundation for future biocontrol strategies based on honey bee-associated symbionts. However, further studies are essential to evaluate the safety of the VOCs produced by B. asteroides 3CP-2B, particularly their effects on healthy brood development. This will be crucial for developing safe and effective application strategies that do not disrupt the hive’s environmental balance or compromise colony productivity.
Statements
Data availability statement
The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found below: https://www.ncbi.nlm.nih.gov/genbank/, PV053127 https://www.ncbi.nlm.nih.gov/genbank/, PV053134 https://www.ncbi.nlm.nih.gov/genbank/, PV053124 https://www.ncbi.nlm.nih.gov/genbank/, PV053131 https://www.ncbi.nlm.nih.gov/genbank/, PV053133 https://www.ncbi.nlm.nih.gov/genbank/, PV053126 https://www.ncbi.nlm.nih.gov/genbank/, PV053144 https://www.ncbi.nlm.nih.gov/genbank/, PV053125 https://www.ncbi.nlm.nih.gov/genbank/, PV053128 https://www.ncbi.nlm.nih.gov/genbank/, From PV056024 to PV056038.
Ethics statement
The manuscript presents research on animals that do not require ethical approval for their study.
Author contributions
MI: Resources, Investigation, Funding acquisition, Project administration, Conceptualization, Writing – review & editing, Writing – original draft. SG: Visualization, Validation, Writing – review & editing. BT: Writing – review & editing, Formal Analysis. LD: Formal Analysis, Writing – review & editing. GA: Writing – review & editing, Software, Writing – original draft, Data curation, Formal Analysis. MS: Investigation, Writing – review & editing. FC: Validation, Writing – review & editing, Visualization. RC: Writing – review & editing, Data curation. CM: Writing – review & editing, Formal Analysis. DC: Writing – review & editing, Validation, Visualization. CT: Writing – review & editing, Validation, Visualization, Investigation. AC: Project administration, Conceptualization, Resources, Investigation, Writing – review & editing, Funding acquisition.
Funding
The author(s) declare financial support was received for the research and/or publication of this article. This research was funded by the University of Molise, under the project BIOMOXE, CUP H33C23003280005.
Conflict of interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Generative AI statement
The author(s) declare that no Generative AI was used in the creation of this manuscript.
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Supplementary material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/finsc.2025.1669013/full#supplementary-material
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Summary
Keywords
honey bee, gut microbiota, Bifidobacterium asteroides, chalkbrood disease, Ascosphaera apis
Citation
Iorizzo M, Ganassi S, Testa B, Di Donato LM, Albanese G, Succi M, Coppola F, Cozzolino R, Matarazzo C, Di Criscio D, Tedino C and De Cristofaro A (2025) Ascosphaera apis as a target for the antifungal activity of symbiotic Bifidobacteria in honey bees. Front. Insect Sci. 5:1669013. doi: 10.3389/finsc.2025.1669013
Received
18 July 2025
Accepted
17 September 2025
Published
01 October 2025
Volume
5 - 2025
Edited by
Maristella Mastore, University of Insubria, Italy
Reviewed by
Walaa Ahmed Elsayeh, Newcastle University, United Kingdom
Chun Tao Gu, Northeast Agricultural University, China
Updates
Copyright
© 2025 Iorizzo, Ganassi, Testa, Di Donato, Albanese, Succi, Coppola, Cozzolino, Matarazzo, Di Criscio, Tedino and De Cristofaro.
This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Sonia Ganassi, sonia.ganassi@unimol.it; Gianluca Albanese, gianluca.albanese@unimol.it
Disclaimer
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.