Efficacy of Botanicals on Parasitoids of Mango Fruit Flies in Burkina Faso ()
1. Introduction
In Burkina Faso, mango is subject to fruit fly attacks, which in 2017 led to a 15% drop in production on a volume of around 200,000 tonnes [1] . Ceratitis cosyra (Hendel) (Diptera: Tephritidae) and Bactrocera dorsalis (Walker) (Diptera: Tephritidae) are the two main fruit fly species responsible for significant damage to the mango industry and food security in Africa, Asia, the Pacific and parts of South America [2] . They can cause losses ranging from 50 to 85% if no appropriate phytosanitary controls are put in place [3] .
In response to the threat posed by fruit flies, several control methods have been developed, including prophylactic control, chemical control, biological control, the sterile insect technique (SIT), mass trapping and the use of protein baits [4] [5] . Unfortunately, some methods (those using synthetic insecticides or fumigants) have drawbacks that limit their use. In the case of chemical control, the overuse of pesticides leads to resistance in fruit flies (in pulp and on mango), the presence of pesticide residues causing food poisoning and environmental pollution. This situation is exacerbated by the relatively high prices of good quality pest control products on local markets. To remedy this, the search is on for an effective control alternative that is more respectful of the environment and human and animal health, and that meets market requirements (where traceability and quality control standards are increasingly stringent) [6] . Combinations of integrated fruit fly management have been used to protect mango orchards in Côte d’Ivoire [7] and Burkina Faso [8] . In Burkina Faso, IPM has been implemented in 12 mango orchards in three provinces. The combination was sanitation + M3 food bait (SM), sanitation + GF-120 protein bait (SG), sanitation + Timaye + M3 food bait (STM) and sanitation + Timaye + GF-120 bait (STG). From this integrated pest management approach, it appears that the GF-120 STG bait was the most effective in reducing both the B. dorsalis and C. cosyra populations in these orchards. This technique has the advantage of prolonging the mango season, by protecting mangoes left on the trees after the export season. The sterile insect technique (TIS) [9] has proved effective against fruit flies in Montreal. Indeed, fruit fly control technologies based on the use of plant substance extracts have particularly attracted our attention due to their availability, biodegradability thus preserving human and animal health and the environment. This work was carried out with a view to finding alternatives to synthetic pesticides that would solve these thorny problems and contribute to sustainable environmental management. We evaluated the effect of hydroalcoholic formulations of Capsicum annuum and Strophantus hispidus on the mango fruit fly parasitoid Diachasmimorpha longicaudata (Hymenoptera: Braconidae). These two substances are generally selective insecticides, non-hazardous to humans and the environment, biodegradable and less costly than synthetic insecticides [5] [7] [8] [9] .
The general aim of this study is to evaluate the sensitivity of the parasitoid D. longicaudata to hydroalcoholic formulations of C. annuum and S. hispidus under laboratory conditions. Before using hydroalcoholic formulations of C. annuum and S. hispidus in an agricultural or ecologically sensitive ecosystem, it is necessary to measure their effects on beneficial organisms (parasitoids and predators). Our study measured the effects of hydroalcoholic formulations of C. annuum and S. hispidus on the parasitoid D. longicaudata used worldwide in biological control programs against fruit Tephritidae [10] . The following hypothesis of research is stated: there is an optimum effective concentration of each hydroa coholic formulation (Capsicum annum and Strophantus hispidus) that is less harmful or tolerated by D. longicaudata.
2. Methodology
2.1. Location of Study Area
The study was conducted at the eco-toxicology laboratory and the biological control laboratory of the Centre National de Spécialisation en Fruits et Légumes (CNS-FL) of the Institut de l’Environnement et de Recherches Agricoles (INERA) in Farako-Bâ, near Bobo-Dioulasso in western Burkina Faso. The rearing room conditions were 12 h/12h photoperiod (dark/light) with a temperature of 25˚C - 28˚C and a relative humidiy of 60% - 70%.
2.2. Composition of the Different Formulations Used in the Laboratory
The material used to apply the different formulations in the laboratory consisted of formulations based on Strophantus hispidus, Capsicum annuum and Success bait (GF120). The yellow pepper Capsicum annuum is available from market gardeners and Strophantus hispidus is a plant available in Burkina Faso in the south-west region and in Africa.
Strophantus hispidus belongs to the Apocynaceae family. This plant has been used to preserve stored foodstuffs [11] . Capsicum annuum belongs to the Solanaceae family and comprises up to 30 species, of which the five main cultivated species are Capsicum annuum L., C. chinense Jacq., C. frutescens L., C. baccatum L. and C. pubescens Ruiz et Pav. Capsicum annuum was used by who tested the insecticidal activity of its extracts on the fall armyworm, Spodoptera frugiperda J.E Smith (Lepidoptera: Noctuidae) in maize cultivation and their effects on the microorganisms of a ferruginous soil, in Burkina Faso [12] . C. annuum extracts are rich in sterols, triterpenes and alkaloids, giving them insecticidal properties.
Strophantus hispidus DC belongs to the Apocynaceae family. Both root and leaf extracts have been shown to inhibit Escherichia coli, Klebsiella pneumoniae, Neisseria gonorrhoeae, Proteus mirabilis, Pseudomonas aeruginosa, Staphylococcus aureus and Streptococcus pyogenes in vitro. Strains of these pathogens to which resistance against certain antibiotics has been induced in the laboratory were also sensitive to aqueous and ethanol extracts of Strophanthus hispidus [13] . S. hispidus extracts are not rich in sterols and triterpenes, but contain many anthraquinones, coumarins and cardenolide derivatives, alkaloids, tannins, flavonoids and anthocyanocides. Certain compounds such as anthraquinones, saponins and flavonoids confer insecticidal properties.
Success bait (GF-120) is a concentrated, water-based suspension of 0.24 g/L spinosad and food attractants (proteins, sugar, fruit flavoring).
Success Appat (S.A.) is a mixture of food attractants and a spinosad-based insecticide (0.24 g/L) capable of attracting and killing fruit fly species present in the intervention zone. The natural insecticide spinosad is obtained by fermentation of a soil bacterium, Saccharopolyspora spinosa.
Animal material: The animal material consisted of adults of the fruit fly B. dorsalis and its parasitoid, D. longicaudata.
2.3. Preparation of Crude Total Extracts of C. annuum and S. hispidus
The various total extracts were obtained from plant powders [14] . Total extracts of C. annuum, were obtained from 3.1 kg of C. annuum ripe fruit powder and those of S. hispidus were obtained from 3.1 kg of S. hispidus leaf powder.
Ripe C. annuum fruits and S. hispidus leaves were shade-dried in a greenhouse, then finely ground using a microgrinder. Extractions were carried out using the Nair method [15] , based on the ability of bioactive substances of plant origin to be carried away by solvents of different polarities. For this study, the crushed C. annuum fruit (3.1 kg) was macerated in 2 liters of ethyl acetate, stirred well with a spatula and left for 24 hours at room temperature. The crushed S. hispidus leaves (3.1 kg) were macerated in 7 liters of ethanol and left for 24 hours at room temperature, after thorough stirring. After 24 hours, the different macerates were then filtered through 5 mm fine mesh muslin placed on a bucket and held in place by an elastic band, then each substrate was rinsed several times with the solvents concerned in reserve to extract the maximum amount of active ingredient.
The filtrates obtained were then concentrated under reduced pressure using a rotavapor. Pressure reduction is facilitated by a device associated with the Büchi R 110 R steam rotavapor, regulated at an appropriate temperature for each extraction solvent, enabling a vacuum to be created at the surface of the liquids (solvent plus solute) to bring them to boiling point at a temperature where the molecules are not destroyed by heat. Thus, for our study, the boiling temperature thanks to the pressure reduction system was located around 50˚C to 60˚C. This enabled evaporation of solvents and concentration of solutes in the rotating flask in the Marie bath.
At the end of the processes, the total crude extract obtained was 200 mL for C. annuum from the starting volume. For S. hispidus, the volume of crude extract obtained was 720 mL. The extracts were then dried in a ventilated Cole Parmer oven, set at 45˚C. The masses of the dry extracts obtained were determined by weighing using a Radwag AS 110.R1 precision analytical balance.
Success bait preparation: Success bait (GF-120) is a concentrated, water-based suspension of 0.24 g/L spinosad and food attractants (proteins, sugar, fruit flavoring).
It is used to control fruit flies in organic production.
For the tests, it was recommended to dilute 1 L of Success bait in 5 L of water.
The preparation and production of hydroalcoholic formulations based on the extracts obtained enabled us to move on to the next stage of the study, which involved testing the effect of the formulations in the laboratory on the parasitoid D. longicaudata. The aim of this stage was to determine the effect of the hydroalcoholic formulations on the parasitoid in a controlled environment, thus ensuring their application in orchards.
A personal protective equipment (overalls + gloves + boots + masks + hats) was used. The maceration time was 24 h.
2.4. Efficacy Testing of Hydroalcoholic Extracts on the Parasitoid
B. dorsalis was reared to obtain male and female adults for the test. Rearing room conditions were 12 h/12h photoperiod (dark/light), temperature 25˚C - 28˚C, relative humidity 60% - 70%.
Rearing consisted in preparing egg-laying trays (height 11 cm; diameter 10 cm), which are yellow funnels perforated at regular intervals and lined with a black cloth soaked in mango juice. The prepared nests were introduced into the various 25 × 25 × 25 cm breeding cages containing sexually mature females and males (15 days old). The scent of mango juice diffused throughout the cage, attracting female B. dorsalis to deposit their eggs through the egg-laying holes. After 24 hours’ exposure to the females, the nesting boxes were removed from the various cages and the eggs were collected by rinsing the funnels in water using a soft brush. The collected eggs were placed on toilet paper and deposited in small bowls containing B. dorsalis larval nutrient medium.
The whole was placed in a large transparent dish (height 11 cm; diameter 15 cm) containing sterilized sand, from which the L3 stage larvae of B. dorsalis would jump and fall to become pupae. The contents were covered with 5 mm fine-mesh muslin to allow air to circulate and prevent the larvae from emerging. The nutrient medium for B. dorsalis larvae was watered every two days. After 12 to 15 days of incubation, the sand was sieved to recover the pupae, which were then placed in the cages for emergence. After emergence of B. dorsalis, they were kept in rearing cages and fed with a mixture of yeast hydrolysate enzymatic (3 measures of cane sugar and 1 measure of yeast hydrolysate enzymatic) and drinkers (water-filled bottles with a piece of water-soaked cotton in the lid).
Diachasmimorpha longicaudata was reared in the laboratory to test the effect of hydroalcoholic formulations on it. It was reared under the same conditions as B. dorsalis. The rearing was carried out using B. dorsalis L3 stage larvae (in their third developmental stage) previously obtained from the rearing of the aforementioned pest.
These larvae were placed in oviposition units containing a nutrient medium for future D. longicaudata larvae. The egg-laying units were then exposed in a cage to sexually mature females of the parasitoid. The egg-laying units were probed by D. longicaudata females, who positioned their ovipositors in the B. dorsalis larvae to deposit their eggs. D. longicaudata eggs hatch inside B. dorsalis larvae and feed on the latter, which then become parasitoid larvae. After 24 h of exposure, the egg-laying units were removed from the D. longicaudata rearing cages. Parasitized B. dorsalis larvae were removed from the egg-laying units and transferred to tanks containing lightly-wetted sterilized sand, where pupation took place. After 7 days, the sand was sieved and the pupae collected were placed in cages in which water-soaked cotton was placed and honey droplets (on the upper wall of the cage) for the feeding of future parasitoids (they feed on the 2nd and 3rd larval development stages of B. dorsalis, then transform into parasitoid pupae and subsequently emerge as parasitoid adults).
Methods for obtaining crude extracts: Ripe C. annuum fruits and S. hispidus leaves were shade-dried in a greenhouse, and then finely ground using a microgrinder. Extractions were carried out using the Nair method [15] , based on the ability of bioactive substances of plant origin to be carried away by solvents of different polarities. The substances were extracted using solvents of increasing polarity (n-hexane, ethyl acetate, methanol). Phytochemical analysis of the active sub-fraction of C. annuum was carried out using the Romanian method of [14] [16] . These studies led to the development of the C. annuum-based hydroalcoholic formulation.
At the end of the processes, the total crude extract obtained was 200 mL for C. annuum from the starting volume. For S. hispidus, the volume of crude extract obtained was 720 mL. The extracts were then dried in a ventilated Cole Parmer oven, set at 45˚C. The masses of the dry extracts obtained were determined by weighing using a Radwag AS 110.R1 precision balance.
2.5. Method of Determining Test Concentrations
For success bait, the concentration considered in this study is that recommended by the manufacturer (0.24 g/L). For hydroalcoholic formulations, 3 different concentrations of total crude extracts were determined and considered for each plant substance. These were low, medium and high concentrations. The different concentrations of hydroalcoholic formulations were calculated based on the work of [14] . According to these authors, masses of dry extracts containing 1.5 mg of C. annuum and 1.5 mg of S. hispidus, i.e. concentrations of 1.5 mg/mL each, caused 18% and 15% mortality respectively in adult C. cosyra in the laboratory over 72 hrs. From these values it was possible to calculate the concentrations of these extracts that could cause 90% mortality in the pests (Table 1). We used the rule of 3 to calculate extracts capable of 90% pest mortality.
Method of preparing hydroalcoholic formulations: The various hydroalcoholic formulations were obtained from the crude total extracts of C. annuum and S. hispidus and the adjuvant (fruit fly attractant) (water, sugar, fresh bovine blood) previously developed. The adjuvant makes the hydroalcoholic extract suitable for practical and effective use [17] . According to these authors, it enhances the efficacy of chemical compounds. For example, the adjuvant ensures that the product adheres to plant leaves, thus improving the effect of product duration and safety. The formulation thus improves the efficacy, shelf life and ease of use of the hydroalcoholic extract.
The formulations were prepared according to the 3 different concentrations of hydroalcoholic extracts of each plant species (C. annuum (3.75; 7.5; 12.5 g/L); S. hispidus (4.5; 9; 15 g/L)). The formulations were based on low, medium and high concentrations.
The different formulations were prepared in 1000 mL of solution according to the following formula:
[18]
where:
Vi = volume of formulation to be pipetted;
Cf = concentration of the stock solution;
Vf = volume of stock solution;
Ved = volume of adjuvant.
Ci = concentration of the formulation to be used to prepare the stock solution;
The formulations obtained are shown in Table 1 and Table 2 [19] .
For packaging and storage of the formulations, empty cans of recovered water were used. As the maximum volume of these cans was 1.5 L, a 10-fold reduction factor was applied to the various hydroalcoholic formulations to obtain volumes of 100 mL. Experimental set-up and data analysis Bioassays were performed in a randomized Fisher block design with 8 treatments and 5 replicates. The treatments were:
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Table 1. Concentrations of hydroalcoholic formulations and success bait tested.
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Table 2. Hydroalcoholic formulations based on crude total extracts and adjuvant.
Capsicum annuum 3.75 (g/L) treatment.
Capsicum annuum treatment 7.5 (g/L).
Capsicum annuum treatment 12.5 (g/L).
Treatment Strophantus hispidus 4.5 (g/L).
Treatment Strophantus hispidus 9 (g/L).
Strophantus hispidus treatment 15 (g/L).
Success bait treatment 0.24 (g/L).
Untreated control treatment.
Testing the effects of different concentrations of hydroalcoholic formulations on the parasitoid D. longicaudata in the laboratory: The test consisted in syringing 1.5 mL of each concentration of hydroalcoholic formulation and pouring it into vials containing 0.25 g of cotton. Next, 20 parasitoid adults were sucked up with a mouth aspirator and placed in the vials, which were then covered with muslin and held in place with rubber bands. This operation was repeated 5 times for each concentration of the different hydroalcoholic formulations of C. annuum and S. hispidus, followed by Success bait and the control (water). All replicates were run simultaneously, and the vials containing the parasitoids were placed on shelves and kept in a room with the same facilities as the insect rearing room. Observations were made in 24 h and 72 h. considering that insects which did not respond to the touch of a fine brush were dead.
2.6. Data Processing and Statistical Analysis
Microsoft Office 2019 Excel spreadsheet software was used to enter and process the collected data and produce the various graphs. R software version 3.6.2 was used for statistical analysis. When the distribution of data did not follow the normal distribution, a non-parametric Kruskal-Wallis analysis was performed to detect differences between treatments. When there was a significant difference between treatments, pairwise comparison of means was performed using the pairwise t-test at the 5% threshold. The analyses concerned the following parameters:
Mortality rate was calculated using the following formula:
Mortality rate = (Number of dead individuals)/(total number of individuals) × 100
3. Results and Discussion
3.1. Results
Effects of different hydroalcoholic formulations on mortality of Diachasmimorpha longicaudata
After 24 h exposure of D. longicaudata adults to the different formulations, a significant difference was found between the 3 C. annuum treatments (Table 3). Parasitoid mortality rates obtained with the high (12.5 g/L) and low (3.75 g/L) doses of C. annuum were 13.80% and 12.88% respectively. With the medium dose (7.5 g/L) of hydroalcoholic concentration of C. annuum, the corresponding mortality was 4.60%. We also detected a significant difference between doses of S. hispidus in terms of the efficacy of this hydroalcoholic formulation against the parasitoid. An average mortality rate of 22.05% of D. longicaudata was obtained with the 15 g/L and 4.5 g/L doses of S. hispidus. The average parasitoid mortality rate recorded with the medium dose of S. hispidus (9 g/L) was 13.8%. Analysis of variance revealed a highly significant difference between the 8 treatments. The highest average parasitoid mortality rate was obtained with the high (15 g/L) and low (4.5 g/L) concentrations of S. hispidus, while the lowest average mortality rate (4.60%) of D. longicaudata was observed with the medium concentration (7.5 g/L) of C. annuum
Mortality effects of different formulations on Diachasmimorpha longicaudata after 72 h
A significant difference was detected between doses of C. annuum after 72 h of exposure of D. longicaudata adults (Table 4). Parasitoid mortality of 50.20% was observed with the 12.5 g/L dose of C. annuum (Table 4). Average mortality rates of 36.70% and 35.81% were recorded with doses of 7.5 g/L and 3.75 g/L of this hydroalcoholic concentration respectively. It was also possible to demonstrate a significant difference between the 3 S. hispidus treatments. Thus, a high average mortality rate for D. longicaudata (64.40%) was obtained with the 9 g/L dose, whereas the 4.5 g/L and 15 g/L doses of this hydroalcoholic concentration produced average mortality rates of 59.95% and 57.15% respectively. Analysis of variance showed a highly significant difference between the 8 hydroalcoholic concentration treatments. The highest mean mortality rate (64.40%) was obtained with the 9 g/L dose of S. hispidus, while the lowest mean mortality rate (35.81%) of D. longicaudata was observed with the 3.75 g/L concentration of C. annuum.
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Table 3. Average mortality rate of D. longicaudata to formulations of C. annuum and S. hispidus 24 hours after exposure, Farakô-Ba, Burkina Faso.
Numbers followed by the same letters (a, b, c etc.) are not significantly different at 5% treshhold.
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Table 4. Average mortality rate of D. longicaudata to formulations of C. annuum and S. hispidus 72 hours after exposure, Farakô-Ba, Burkina Faso.
Numbers followed by the same letters (a, b, c etc.) are not significantly different at 5% treshhold.
3.2. Discussion
The various mortality tests of D. longicaudata to different formulations of C. annuum and S. hispidus enabled us to identify which formulations were toxic to the parasitoids.
The S. hispidus formulations (15 g/L and 4.5 g/L) and the success bait were toxic to D. longicaudata adults after 24 h of exposure. This result could be explained by the fact that the S. hispidus formulations (15 g/L and 4.5 g/L) act through active ingredients such as flavonoids, cardenolides, sterols/triterpenes, alkaloids and saponosides, which are responsible for the observed toxicity towards parasitoids. These results are in agreement with [20] who showed that flavonoids were found in the leaves and nowhere else in the other plant organs. [14] showed the presence of sterols/triterpenes, flavonoids, anthraquinones, cardenolides, alkaloids and saponosides in S. hispidus extracts. This presence also depends on exogenous factors such as temperature, insect anatomy and morphology. The success bait was toxic to D. longicaudata adults. This result could be associated with the insecticidal effect of spinosad on parasitoid adults. These results are in agreement with those reported by [21] [22] who showed that spinosad, a bacterial insecticide derived from the actinomycete Saccharopolyspora spinosa contained in the bait was slightly harmful to D. longicaudata (IOBC class 2). However, the half-dose formulation of C. annuum (3.75 g/L) was less toxic to D. longicaudata adults. This concentration of C. annuum can be recommended for fruit fly control, while preserving the D. longicaudata parasitoid in the wild after its release in orchards.
After 72 h, the S. hispidus formulations (9 g/L; 15 g/L) and the success bait were highly toxic to D. longicaudata adults. These results could be explained by the presence of phytochemical compounds in S. hispidus that could have toxic effects on parasitoid adults through contact and/or ingestion. In contrast, C. annuum formulations proved less toxic to D. longicaudata. The phytochemicals in C. annuum may have had less toxic effects on adults of the parasitoid by contact and/or ingestion. This formulation of C. annuum can be recommended for fruit fly control, while preserving the D. longicaudata parasitoid in the wild after its release in orchards.
Successive baits were also found to be toxic to parasitoid adults. These results are in line with those of [23] , who carried out a spinosad contact test with Fopius arisanus (Sonan) and Pysttalia (Silvestri). In this test, spinosad was toxic to parasitoids at concentrations above 500 mg/liter after application of aerosol baits. In addition, spinosad (not in bait-sprays) has been shown to be toxic to some hymenopterans in previous studies [24] [25] [26] . [27] found that malathion in GF-120 aerosol bait formulations, which contain spinosad, was toxic to the aphid parasitoids Aphytis melinus DeBach and Lysiphlebus testaceipes Cresson. [28] showed the toxicity of three treatments of spheres with imidacloprid (2 and 4% active ingredient [MA] Provado 1.6 F) to D. longicaudata and an untreated (non-toxic) control sphere over a 24 h - 72 h exposure period. There was no significant difference in mortality of D. longicaudata exposed to spheres treated with imidacloprid at 2% and 4% (AI) for 2 or 4 weeks. Although higher mortality of D. longicaudata was recorded with 4% compared with 2% (AI) after 24 h, it is possible that the seasonal population dynamics of fruit fly parasitoids inhabiting the different niches of Anastrepha suspensa (for feeding) is so limited that populations would not be significantly reduced [29] . [30] [31] also reported that sprays of spinosad or phloxine B baits had little or no effect on the A. suspensa parasitoid, Fopius arisanus (Sonan), the main parasitoid of the Mediterranean fruit fly, Ceratitis capitata (Wiedemann), in Hawaii. Nevertheless, further research in real-life conditions would be needed to determine the toxicity of hydroalcoholic formulations of C. annuum and S. hispidus to D. longicaudata under natural conditions.