Antifungal Activities of Propolis and its Main Components with an Emphasis Against Phytopathogenic Fungi

The first accounts of propolis's benefits date back to the ancient Egyptians. They used it to embalm corpses due to its antiputrefactive properties. The Incas used propolis as an antipyretic agent to treat fevers (Silva-Carvalho et al., 2015).

Many ancient Greek and Roman manuscripts mention its preparation and use as a traditional medicine in various diseases and treatments (Ghedira et al., 2009). The first authors to describe its medicinal effects were Aristotle, Dioscorides and Hippocrates among the Greeks, and Pliny and Galen for the Romans (de Funari et al., 2007; Toreti et al., 2013). Propolis was used as for cicatrising and an antiseptic to treat wounds and, as a mouthwash. The texts spoke of propolis as a “cure for contusions and suppurating ailments”. In Rome, doctors applied it for poultices (Garedew et al., 2004), and it was highly sought after and more expensive than honey. In battle, Roman legionaries carried with themselves a small quantity to treat any wounds. These curative uses were perpetuated in the Middle Ages and via Islamic medicine (Ferhoum, 2010; Fokt et al., 2010). In the 12^th century, propolis was used in Europe to treat mouth and throat infections (Ozcan et al., 2004), and as early as the 17^th century, it was listed as an official medicine of the London Pharmacopoeia and became very popular in Europe from that time, due to its antimicrobial activity.

In France, propolis first appeared in written documents only at the beginning of the 18^th century. It reached its zenith during the Boer War (1899–1902) in South Africa at the end of the 19^th century, when it was promoting as a disinfectant and antiseptic for wound healing and tissue regeneration (Ferhoum, 2010; Mora et al., 2011). In World War II, several Soviet clinics treated tuberculosis with propolis; the results were conclusive, with a reduction in lung problems and a notable recovery of appetite.

Its use has been well known for centuries, but the enthusiasm for it from a scientific point of view has considerably increased since the 1980s. For instance, a great deal of work has focused on gradually testing new therapeutic possibilities for this substance. The latest techniques have provided a better understanding of the mechanisms involved and the molecular signalling pathways at work. These discoveries suggest that propolis might be used in increasingly vast fields, including treating cancer, diabetes, cardiovascular or neurodegenerative diseases, but also in veterinary medicine, or agronomy.

Propolis arised from an assembly between a plant resin and beeswax. The plant resin is harvested by worker bees (Apis mellifera L.) from the buds of certain trees, bushes, but also from young branches, leaf petioles and cracks in the bark of certain plants found in the ecosystem surrounding the hive. There are considered to be as many types of different propolis as there are ecosystems on the planet. Tab. 1 and 2 present a non-exhaustive list of the different propolis types existing in the world. Propolis in Europe is harvested almost exclusively from the buds of poplar trees (genus Populus) whereas in Brazil from Baccharis dracunculifolia and in Tunisia from Cistus sp.

This plant resin consists of wax secreted by bees and their saliva. The enzyme 13-glycosidase existing in bee saliva hydrolyses flavonoid glycosides into flavonoid aglycones (Ramos & Miranda, 2007; Farooqui & Farooqui, 2012) thereby giving rise to propolis. Its different physicochemical characteristics mainly depend on its botanical origin. Its colour varies from yellow to green and from red to dark brown and depends on the type and age of the plant foraged by bees (Ghedira et al., 2009). The typical balsamic odour of propolis makes it easily recognisable. Poplar propolis can be hard and brittle in low temperatures and soft, rubbery and sticky in hot temperatures, while Baccharis propolis hard and friable at room temperature (Fokt et al., 2010). The materials available for “making” propolis are lipophilic substances produced by the surrounding plants in the bee ecosystem (Bankova, 2005).

In its ecology, the bee utilizes this malleable propolis as “cement”, which is deposited inside the hive. In order to be perfectly viable, the hive must ensure a degree of imperviousness compatible with their various tasks (e.g. aeration of honey). Propolis is therefore used to construct and repair the hive, plug holes and ensure it is sealed. However, paradoxically this imperviousness results in a confined space in which the thermal conditions (34±2°C) and relative humidity (around 80%) are conducive to the development of such pathogenic organisms as bacteria or fungi (Ota et al., 2001). However, bees know how to harvest a substance that has the physical characteristics of “cement” and the chemical characteristics of an antimicrobial weapon. Indeed, propolis deposited in the hive has shown to prevent the decomposition of organic matter inside the hive by inhibiting microbial growth, as in mummifying the carcasses of invaders (Quiroga et al., 2006; Pietta et al., 2012).

The biological activities and antimicrobial properties of propolis are attributed to molecules that come exclusively from the foraged plant that provides the resin (Vardar-Ünlü et al., 2007). The survival of the species is ensured because bees adapt to their ecosystem by harvesting a resin which ensures the wholesomeness of the hive while protecting it from harmful microbes specific to the same ecosystem. The great natural variability in propolis depends on the type of plants foraged by the workers present at the collection site but also on climatic conditions, the season, and bee type.

Now popular thanks to much available information, Propolis is considered as an alternative natural therapeutic, and propolis-based preparations as much surer and much less harmful tofor the health than numerous medicines (Castaldo & Capasso, 2002). This renewed enthusiasm has motivated both researchers and the industry to continue investigating and broadening the fields of application for propolis (Temiz et al., 2013).

Over the last thirty years, intense research on propolis has confirmed or discovered numerous pharmaceutical and biological activities including antibacterial, antiviral, antifungal, antioxidant, anti-inflammatory, antitumoral and immunomodulating properties. The range of its medical applications has therefore been widely expanded. Current research covers medicine with some new therapeutic targets, as well as work in the cosmetics, food and the animal-care industries (Tosi et al., 2006).

Today, propolis can be administered either orally or systemically in the form of ampoules, infusions, syrups, tablets and capsules, or in topical applications such as toothpastes, pastilles-lozenges, mouth and nasal sprays (Almas et al., 2001; Ghedira et al., 2009). Mouthwashes and other dentifrices prevent tooth decay, gingivitis and stomatitis, and many formulations exist for treating allergies, inflammatory diseases, asthma, diabetes and hypertension and for use as a dietary supplement (Marcucci, 1995; Farooqui & Farooqui, 2012). In Chile, propolis is consumed in very popular alcoholic and non-alcoholic beverages (Herrera et al., 2010). Almost all products available on the market are based on ethanol extracts of propolis, but industries also use glycerine and propylene glycol in product preparation (Ramos & Miranda, 2007).

Despite the exteninsive and varied scientific literature on both the chemical composition and biological activities of propolis, most therapeutic applications remain limited primarily to human pathologies. Its application for food preservation in such fields as the agrifood sector or agriculture has seen little development. Despite the need for scientific knowledge on propolis, researchers or publishers encounter difficulties in ensuring that results published are repeatable and allow comparison, so as to extract usable and reproducible interpretations essential for application in the field.

Many authors have claimed that the general composition of raw propolis (Tosi et al., 2006; Sawaya et al., 2011; Haile & Dekebo, 2013; Nedji & Loucif-Ayad, 2014) consists of around 50% resin (containing the polyphenol fraction), 30% beeswax (wax and fatty acid), 10% essential oils (volatile compounds), 5% pollen (pollen protein and free amino acids) and 5% other substances (vitamins, enzymes, steroids, etc.), referring to old articles, some dating back over seventy years. More recently, Bankova et al. (2014) has criticized those data and downwardly revised the volatile compound percentage, citing 1% and occasionally 2 to 3%. Differences in appearance and textures between raw propolis samples from poplar Baccharis and Dalbergia suggest that these substances do not share the same general composition in percentage terms. An internal study (data not shown) showed that the distribution of resin/wax/others in propolis from poplar Baccharis and Dalbergia was 60-60-40/20-10-20/20-30-40, respectively. As with any natural substance, raw propolis is variable. Factors of variability in the chemical composition of propolis include source plants, their availability at the time of collection, a hive's geographical origin, bee type, climate and season (Ghedira et al., 2009; Haile & Dekebo, 2013). How raw propolis is harvested by humans could also be added to these natural elements.

Walker & Crane (1987) first published a work inventorying plants in the world that supply propolis; however, authors working on its composition and consequently on its biological activities have rarely specified the botanical origin, and geographical origin is not enough to effectively describe a propolis sample. In Brazil for example, Park et al. (2004) demonstrated the existence of thirteen different chromatographic profiles from propolis samples taken throughout the Brazilian territory. Of all the molecules listed on those chromatograms, only two flavonoids and one phenolic acid were common to the thirteen types of Brazilian propolis, a new one displayed a bright red colour and its chemical composition differed from the other twelve types of propolis already inventoried. Silva et al. (2008) identified the botanical origin of this new propolis by comparing the chemical profile of ethanol extracts of the red propolis to those of resins collected from twenty different Brazilian plants. Only one plant, Dalbergia ecastaphyllum, had a virtually similar profile rich in isoflavonoids.

Tab. 1–2 give a list that is not exhaustive but matches information given in articles about propolis. Each article mentions the geographical origin of the propolis, while only 40% of them specify the botanical origin. For example, the table shows us that the majority of compounds found and identified by Mohammadzadeh et al. (2007) as coming from poplars in Iran were the same as those found in propolis from Chile (Herrera et al., 2010; Curifuta et al., 2012) and China (Yang et al., 2011). Although the botanical origin of those propolis samples was not mentioned, it is likely to have been poplars.

Markham et al. (1996) presented a propolis specific to New Zealand which was slightly similar to Manuka honey also from New Zealand. However, the authors stated that New Zealand had largely reintroduced the European poplar to its soil. Catchpole et al. (2004) qualified New Zealand propolis as a poplar propolis identical to European propolis. The composition of the red propolis from north-eastern Brazil, derived from Dalbergia, is similar to that of a red propolis rich in isoflavonoids, such as medicarpin and 3-hydroxy-8.9-dimethoxypterocarpan, found in Cuba's Pinar Del Rio province, proving the existence of the species Dalbergia in that region of Cuba (Meneses et al., 2009). In terms of the chemical composition of raw propolis, these different examples show how little importance the geographical origin carries when compared to the botanical origin.

Several authors have worked on seasonal impacts affecting propolis composition. One study compared the chemical composition of propolis samples from the same region of Sonora in Mexico over 4 consecutive seasons. The results of the chromatographic profiles obtained, and the relative abundance of the main peaks, did not reveal any significant difference (Valencia et al., 2012). Similar work had already been undertaken five years earlier at the same site, and again the comparison of the chromatograms did not reveal any significant difference. Nonetheless, a few differences were found for the relative abundance of certain non-majority compounds, which confirms that seasonality does not significantly influence quality but maybe quantitatively a few compounds, hence maybe biological activity.

To determine how bee type plays a role in the variability of propolis composition, some work was undertaken to analyse the volatile compounds of propolis samples harvested from various regions of Brazil with two different subspecies of Meliponinae bees. The percentages of different volatile compound families were compared using samples from the same place but with six different bee subspecies, including genus Plebia spp. and Tetragonisca angustula. In quantitative terms, two samples had diterpenic acids as did the majority family, three had triterpenic acids and the last sample had derivatives of benzoic acids. However, no qualitative analysis was undertaken, and instead bees of the subspecies Melipona quadrifasciata anthidioides were analysed in different geographical zones, for which some differences appeared. The author was unable to reach any conclusion regarding the species of bee or geographical origin, for determining the chemical composition of a propolis sample (Velikova et al., 2000). The enzyme and glandular system of bees were shown to not vary in terms of chemical composition, ans consequently the variations encountered in propolis composition are solely due to botanical origin (Bankova, 2005).

The chemical composition of raw propolis varies from a region with a tropical climate to a region with a temperate climate. In fact, in a temperate zone such as Europe propolis mainly comes from poplars, whereas in a country with a tropical climate such as Brazil, most propolis comes from Baccharis dracunculifolia or Auraucaria spp. Whilst not being a natural element, the propolis-harvesting method since the human domestication of bees has added a source of variation. For instance, Stan et al. (2011) showed that when poplar propolis is harvested with the screen method, the proportion of wax tends to not exceed 21%, unlike the standard results obtained with the frame scraping method where the proportion largely exceeds 30%. This difference in wax percentage is important because, physically, the drop in wax means a larger quantity of resin. As the latter is the sole source of the active ingredients for propolis, this suggests that the larger the proportion of resin, the greater will be the active ingredient contents.

Lastly, although as many sorts of propolis exists as there are different ecosystems on earth, only propolis from Baccharis, from poplars and more recently from Dalbergia have been widely studied. The botanical origin of these types of propolis was identified by the presence of molecular markers from the source plant. For instance, the main families of compounds encountered in poplar propolis are flavones (chrysin), flavanones (pinocembrin) and phenolic acids (caffeic acid), while a propolis derived from Baccharis is mostly composed of terpenoids and prenylated derivatives of p-cinnamic acid (artepillin C) and one derived from Dalbergia is composed of isoflavonoids (Bankova et al., 2002; Fokt et al., 2010).

We have seen that a number of natural and human factors exist which both govern and influence the composition of propolis in terms of its active ingredient. Given that the pharmaceutical-biological activities of propolis can be attributed to those same active ingredients, the botanical origin must be specified and no longer the geographical origin, which is of little interest. As shown in Tab. 1 and 2, many authors do not mention the botanical origin of propolis; the majority compounds are often listed but rarely quantified. Without that information, the repeatability of the work undertaken cannot be guaranteed. From now on, all studies will be more complete and precise as regards botanical origin, bee type and harvesting method, as these factors are decisive in the composition of propolis and thereby in the possibility of comparing certain biological activities of the same raw propolis.

Propolis has been known for its antimicrobial activity, and recently, a great deal of work on this natural substance has confirmed that property, while fine-tuning it for its antibacterial, antiviral and antifungal virtues. Even though many articles on human pathogens have reported on propolis as potential antifungal candidate (Dobrowolski et al., 1991; Ota et al., 2001; Dota et al., 2011), propolis is not officially recognized for its antifungal activity in the context of increasing antibiotic resistance. Some natural substances rich in flavonoids and phenolic acids have been shown to display an antifungal activity against such plant pathogenic fungi as Botrytis cinerea, Aspergillus niger and Alternaria alternata (Mohammadzadeh et al., 2007; Castro et al., 2009), yet there is little work with propolis on plant pathogens.

Despite this, those studies are representative of the type of study undertaken as their design and the conclusions presented for human pathogens. We shall therefore study the different techniques used to assess the antifungal activity of propolis applied to plant pathogens. A critical analysis will be conducted on the inconsistencies, bias and lack of readability by certain authors regarding their expression of results, the protocols published and the interpretations made. The antifungal activity of propolis extracts is generally assessed through the study of mycelium growth or spore germination. The assessment of plant pathogen mycelium growth is most widely described in literature (Ghaly et al., 1998; Ozcan et al., 2004; Meneses et al., 2009; Yang et al., 2011; Temiz et al., 2013; Dudoit et al., 2020; Hosseini et al., 2020).

The dilution method and the diffusion method are mainly used to assess the effect of propolis extract on mycelium growth. The dilution method is widely used for assessing the antifungal activity of propolis and carried out in an agar medium or a liquid medium.

In an agar medium, the different concentrations of propolis liquid extract are incorporated into the agar prior to inoculation on the mycelium plug surface (Quiroga et al., 2006; Curifuta et al., 2012; Agüero et al., 2014). Mycelium growth inhibition is determined through the comparison of the mycelium radial growth diameter of the negative control (without propolis) with that of the tested extract (Bosio et al., 2000). In a liquid medium, the different concentrations of propolis liquid extract to be tested are incorporated into a spore suspension in a nutrient broth. After incubation, both mycelium growth and inhibition are determined with spectrophotometry (Hegazi et al., 2000). this method, also called the broth microdilution method, is most often used to determine the minimal inhibitory concentration (MIC) which corresponds to the lowest propolis extract concentration for which no microorganism growth is visible (fungistatic effect) (Quiroga et al., 2006; Mohammadzadeh et al., 2007; Agüero et al., 2014).

Although less common, the diffusion method can be carried out with the disc or well technique. The disc diffusion method consists of placing a paper disc imbibed with the propolis extract to be tested in different concentrations on the surface of the agar inoculated uniformly beforehand with a suspension of microorganisms. Following appropriate incubation conditions, antifungal activity is assessed by measuring the diameter of the growth inhibition zone surrounding the disc (Bosio et al., 2000; Sawaya et al., 2011).

The well technique consists of introducing the propolis extract into wells made in the agar after its inoculation with the microorganism to be tested. The inhibition diameter around the wells is measured to determine the mycelium growth of the pathogen in question. The dilution method is more widely used than the diffusion method to determine mycelium growth. Indeed, incorporation of the extract directly into the culture medium facilitates the dispersal of the active molecules and enables clearer visualization of the extract's activity.

Spore germination can be assessed with the dilution method in an agar or liquid medium as seen. After incubation, the number of spores is counted with the use of a Malassez cell under a light microscope, and the percentage of spore germination inhibition is determined (Ghaly et al., 1998).

All of these techniques require a negative control without propolis extract so that there is no solvent effect on the inhibition of the tested pathogen. In addition, the presence of positive controls, i.e. such chemical fungicides as amphotericin B, ketoconazole or terbinafine, is an asset as they can be used to compare the activity of the extract to that of some reference molecules (Quiroga et al., 2006; Agüero et al., 2014). Researchers do not often take this parameter into account.

The thin-layer chromatography (TLC)-bioautography method is a more recent technique (Ndjolo, 2012). TLC plates are used to visualize the inhibition zones (representative of activity) generated by one or a group of molecules separated beforehand with this method (Moreno et al., 1999; Quiroga et al., 2006; Kasote et al., 2015). The chemical families present in the active fractions are then visualized through fluorescence and/or characterized through HPLC/MS.

A list not exhaustive but representative of the studies on the antifungal activity of propolis extract against plant pathogens is presented in Tab. 2, which sums up the results, experimental conditions and methods used. In these studies, antifungal activity was assessed through mycelium growth inhibition (Ozcan et al., 2004; Quiroga et al., 2006; Mohammadzadeh et al., 2007; Meneses et al., 2009; Yang et al., 2011; Curifuta et al., 2012; Haile, & Dekebo, 2013; Temiz et al., 2013; Ali et al., 2014; Mattiuz et al., 2015; Dudoit et al., 2020), inhibition of spore germination (Ghaly et al., 1998; Mattiuz et al., 2015) or MIC determination (Quiroga et al., 2006; Mohammadzadeh et al., 2007; Agüero et al., 2014; Xu et al., 2019; Hosseini et al. 2020).

Such various plant pathogenic fungi as Fusarium sp., Aspergillus sp., Penicillium sp., Colletotrichum sp., Botrytis cinerea and Aternaria sp. used in these studies to investigate the antifungal activity of propolis extracts came from collections or were isolated from a food matrix (Ozcan et al., 2004; Quiroga et al., 2006; Meneses et al., 2009; Curifuta et al., 2012). Mycelium growth inhibition is usually determined through dilution in an agar medium. Ethanol extracts from Turkey, Ethiopia, Chile and Brazil in studies by Curifuta et al. (2012), Haile & Dekebo (2013), Mattiuz et al. (2015) and Temiz et al. (2013) showed total inhibition of mycelium growth for the different plant pathogens studied at a concentration of between 2.5 and 5% (v/v).

Some works have tested the dose-response effect of propolis ethanol extracts on mycelium growth. Ozcan et al. (2004) and Mattiuz et al. (2015) found that inhibition increased partially and, then completely when the concentration of their Brazilian propolis extract rose from 0.5 to 2.5 % (v/v), while an increase from 66 to 89% inhibition was observed with concentrations of 0.25 and 0.75% for Chinese propolis. The inhibition percentage rose from 15 to 100% when the concentration of Turkish propolis rose from 1 to 10%. Another study by Ozcan et al. (2004) tested the effectiveness of methanol extracts of propolis from five different regions of Turkey at concentrations of 2% and 5%. Only two propolis extracts displayed total inhibition from a concentration of 5%. The results for these two methanol extracts were similar to those obtained previously with ethanol extracts. However, the methanol extracts of the other three regions showed a substantial inhibiting effect of 52% at a concentration of 2%.

Four sub-fractions were made up from the same ethanol extract of Chinese propolis with different solvents (ethanol, water, petroleum ether, n-butanol, ethyl acetate) and compared. At an identical concentration (200 mg mL⁻¹), only the ethyl acetate sub-fraction completely inhibited the growth of Penicillium italicum. The inhibition percentages obtained, in decreasing order, were 100, 93, 35, 25 and 6% for the ethyl acetate, ethanol, petroleum ether, n-butanol and water fractions, respectively (Yang et al., 2011). Interestingly, the initial ethanol extract concentrated at 1200 mg mL⁻¹ did not lead to complete inhibition, but unlike its ethyl acetate sub-fraction though it was six times less concentrated. Meneses et al. (2009) tested the inhibiting power of different fractions of Colombian propolis on: Colletotrichum gloeosporioides strains isolated from mango and papaya and Botryodiplodia theobroma strain isolated from avocado. The four fractions tested were the n-hexane/methanol fraction (EPEM), dichloromethane (CH₂Cl₂), ethyl acetate (EtOAc) and methanol (MeOH). The (CH₂Cl₂) fraction gave the best results with 47 and 38% of inhibition for the two strains of C. gloeosporioides, while the EPEM fraction led to a better inhibition of the B. theobromae strain. Unfortunately, the authors did not specify what concentrations were used to obtain those results.

Curifuta et al. (2012) and Temiz et al. (2013) chemical fungicides as positive controls (sodium benzoate and benzimidazole) in their myceliumgrowth inhibition studies and concluded that the chemical fungicides had a weaker inhibiting activity than the tested propolis extracts. Liquid medium dilution is most frequently used to determine the MIC value and the germination inhibition percentage. Some ethanol extracts of Brazilian propolis were tested against the germination of a Colletotrichum gloeosporioides, the main pathogen of mango anthracnose. The inhibition of C. gloeosporioides spore germination is dose-dependent, inhibition of this strain being total beyond a concentration of 2% (v/v) propolis extract (Mattiuz et al., 2015). However, worth noting is the exponential effect of the propolis concentration on that inhibition, since the fact of increasing the concentration from 1 to 1.5% increased inhibition from 47 to 96%. Ghaly et al. (1998) found that their ethanol extract of Egyptian propolis only reduced the spore germination of the 2 Aspergillus isolates by 56 to 76% for concentrations of between 3 and 4 g L⁻¹. In that study, the authors expressed their result as the concentration of initial raw propolis prior to extraction, which in reality amounts to the extraction ratio.

Quiroga et al. (2006) and Agüero et al. (2014) determined similar MIC value (250 vs 232 μg mL⁻¹) for their propolis extract originating from different regions of Argentina, extracted through different processes. With a MIC > 250 μg mL⁻¹, both judged that the Aspergillus strain was not susceptible to the propolis extract used, while Mohammadzadeh et al. (2007) using an ethanol extract of Iranian poplar propolis determined a MIC value of 500 μg mL⁻¹ against the same Aspergillus strain. Although their MIC was twice as high as the previous value, Agüero et al. (2014) and Quiroga et al. (2016) concluded that this fungal strain was susceptible to their propolis extract.

Quiroga et al. (2006) also determined the MIC values of the chemical fungicides ketoconazole and clortrimazole and of the isolated and purified flavonoids, pinocembrin and galangin, which they compared to the MIC of the propolis extract on five fungus strains (a strain of Aspergillus, a strain of Penicillium and 3 strains of Fusarium). The results for the five tested strains indicated MIC values of between 195 and 350 μg mL⁻¹, between 1 and 8 μg mL⁻¹ and between 25 and 45 μg mL⁻¹ for the propolis extracts, the chemical fungicides and the isolated flavonoids, respectively. These results lead to the conclusion that the chemical fungicides better inhibited than the isolated compounds, which performed better than the tested propolis extract. The studies undertaken with the TLC-bioautography method confirmed that the active molecules responsible for at least some of the antifungal activity belonged to the family of phenolic compounds (Quiroga et al., 2006; Yang et al., 2011).

Due to the recent use of biologically active natural products to treat fruits and vegetables there has been a real research challenge to replace propolis synthetic fungicides for controlling pre and postharvest decay. Propolis has been found to inhibit antifungal postharvest pathogens in vitro. The efficacy of propolis extract on mycelial growth (Ozcan et al., 2004; Yang et al., 2011; Mattiuz et al., 2015) and spore germination (Ghaly et al., 1998; Mattiuz et al., 2015) on phytopathogen strains such as Colletotrichum gloeosporioides, Aspergillus flavus, Alternaria alternata, Fusarium oxysporum, Penicillium italicum has been widely described in literature. Recent studies have been conducted on the application of ethanolic extract of propolis alone or in combination with edible coating on postharvest biopreservation of fruits (Ali et al., 2014; Barrera et al., 2015; Mattiuz et al., 2015). Interesting results have been obtained in vivo on the efficacy of propolis as an alternative process for food preservation. However, no request or approval documents have been made to introduce propolis on the market in plants and crops protection. The use of a natural extract or active substance must be approved as a basic substance by European Commission and listed in Regulation (EU) N° 540/2011.

Mattiuz et al. (2015) reported that the ethanolic extract of propolis (EEP) did not limit the growth of C. gloeosporioides on mango the first fourteen days of storage, but on day 21 the treatment significantly reduced lesion areas on mango compared to control. Matny et al. (2015) also reported the efficacy of Iraqi EEP to control and reduce apple fruit decay. EEP treatment reduced mold area by 66.8% compared with untreated fruit against P. expansum. Giovanelli (2008) showed the potential of South African EEP to inhibit post-harvest diseases and prevent infection caused by Colletotrichum on avocado fruit.

In these two studies, fruits were first inoculated with 15 μL of Colletotrichum sp. conidia (1 × 105 conidia mL⁻¹). To understand the preventive action of EEP against Colletotrichum, fruits were immediately sprayed with 5 mg mL⁻¹ (MIC value) and 10 mg mL⁻¹ EEP. Whereas in the study on the efficacy of EEP to inhibit disease symptoms of infected fruit, these fruit were treated twenty-four hours after infection. In both cases, the use of 5 mg mL⁻¹ and 10 mg mL⁻¹ EEP on the post-harvest treatment of avocado fruit inoculated with Colletotrichum conidia evidenced a similar potential to inhibit disease symptoms compared with untreated fruit. Similarly, Moraes et al. (2011) has proved the efficiency of 10% propolis extract concentration for the control of powdery mildew in tomato plants. Lastly, Matny (2015) soaked oranges with concentrations of 1, 2 and 3% EEP from Baghdad, Iraq, for the control of Penicillium digitatum pathogen causeing gray mold, and found that 3% more efficiently inhibited disease severity. All EEP treatment still showed a fungicidal activity compared with pathogen treatment only. The effect of 1%, 2% and 3% EEP on disease incidence was respectively 66.67%, 33.34% and 22.23%.

Few studies deal with the efficacy of propolis incorporated into edible films. Ali et al. (2014) reported the use of EEP and edible coating with gum arabic (GA) and EEP to control anthracnose of papaya caused by C. gloeosporioides. After a four-week storage, 1.5% EEP treatment alone or in combination with 10% GA concentration provided a disease incidence reduction of 80% compared with control fruit. Barrera et al. (2015) investigated the application of an edible propolis coating on fruit to minimize postharvest decay. Compared to water-coating control, a coating of chitosan and EEP treatment on papaya during storage against C. gloeosporioides reduced lesion diameter In vivo results revealed that propolis could be a natural antifungal substitute to the actual chemical fungicide for control post-harvest diseases of fruits and vegetables.