Introduction

Plant diseases are caused by infectious agents such as fungi, bacteria, nematodes, flagellated protozoa, viruses, viroids, or, even, abiotic factors such as edaphoclimatic alterations and the toxicity generated by pesticides or nutrients (Guzmán et al. 2012). Over the course of millions of years, the association between plants and nematodes has given rise to the evolution of phytoparasitic nematodes, pathogens widely distributed around the world in vascular plants which are often attributed to low crop yields and losses. The Food and Agriculture Organization (FAO) of the United Nations considers a pest any agent causing damage to plants or vegetable products (Armendáriz et al. 2015; Bernard et al. 2017). Phytoparasitic nematodes (PPNs) are a serious global problem for farmers, causing estimated annual crop losses of $118 to $157 billion dollars (Abad et al. 2008; Degenkolb and Vilcinskas 2016; Khan et al. 2020). Two types of PPNs, root-knot nematodes (RKNs) and cyst nematodes (CNs) are obligate parasites of a wide range of agricultural crops. Table 1 presents a summarize of the chemical control used for PPNs, based on previous information of Nicol et al. (2011) and according to Evans et al. (1993), Trudgill and Blok (2001), Luc et al. (2005), Nicol and Rivoal (2008),, Armendáriz et al. (2015), Rehman et al. (2016) and Ebone et al. (2019). Particularly in Mexico, PPNs are a problem for various crops of interest, as presented in Table S1. As a result, farmers have found chemical products (Table 1) as their preferred control method, since their rapid action and solubility in water ensure a uniform distribution on the upper soil layer (Degenkolb and Vilcinskas 2016). However, chemical control methods raise environmental and safety concerns. It has been shown that the application of all the chemical compounds with nematicidal activity potentially risks environmental contamination through mechanisms like direct ingress into waterways, the infiltration of groundwater, surface runoff into rivers, streams, lakes, and reservoirs from neighboring agricultural regions; aerial application and discharge of wastewater from industrial pesticide producers (del Puerto Rodríguez et al. 2014; Zisis 2018). Moreover, these compounds could become toxic to humans when come into contact through possible routes of exposure as: acute oral toxicity, dermal toxicity via absorption through the skin and oxidative stress, inhalation and chronic toxicity causing diseases such as cancer, diabetes mellitus, neurological system disorders, effects on the immune system, endocrine system disturbances and reproductive (sexual/genital) syndromes (Li et al. 2014; Joshi and Sukuraman 2019; Lata et al. 2021). Some of the main nematicides used are basamid, metam sodium, oxamyl, telone, carbofuran, methyl bromide, and thiodicarb (Armendáriz et al. 2015; Ebone et al. 2019).

Table 1 Chemical control method required for main nematode pest and crops described by Nicol et al. (2011) and according to Evans et al. (1993), Trudgill and Blok (2001), Luc et al. (2005), Nicol and Rivoal (2008), Armendáriz et al. (2015), Rehman et al. (2016) and Ebone et al. (2019)
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This has motivated research for the development of environmentally friendly PPN control alternatives to chemcal products currently on the market. Although there are control agents of this type available, not all of them are known to farmers due to the lack of information on both their mechanism of action and benefits. Alternative low impact methods for nematode control, such as genetic and induced resistance or the use of biological control agents, are highly desirable and actively sought (Molinari 2011; Stirling 2011). Within the Integrated Pest Management (IPM), biological control has become an environmentally safe alternative for reducing the use of chemical nematicides. In this context, diverse microbial biological control agents such as Purpureocillium lilacinum, Trichoderma spp., Pochonia chlamydosporia and Bacillus thuringiensis, among others, have been evaluated to reduce nematode infestations in susceptible crops (Nordbring-Hertz et al. 2006). Purpureocillium lilacinum was known previously as Paecilomyces lilacinus (Prasad et al. 2015) and Pochonia chlamydosporia var. chlamydosporia is also known as Verticillium chlamydosporium (Kerry et al. 1984). The present study focuses on the potential of these nematophagous fungi P. lilacinum and P. chlamydosporia, emphasizing the main challenges for their mass production and formulation.

Phytoparasitic nematodes

Nematodes are pluricellular worm-shaped organisms that generally measure between 0.2 and 2.5 mm in length, depending on the species, and constitute the phylum Nematoda. It is thought that this phylum emerged during the Cambrian explosion (550 million years ago) in marine habitats (Aguinaldo et al. 1997). Nematodes are found in land systems and marine and freshwater habitats (Bongers and Ferris 1999). Despite their small size, their biological organization is highly complex, wherein circulatory, and respiratory systems are not defined instead of, their body cavity contains a liquid that enables circulation and respiration functions in these organisms. The digestive system comprises a hollow tube extending from the mouth, through the esophagus, intestine, rectum, and anus (Talavera 2003). The large part of these organisms is generally elongated and cylindrical. The adult females of some phytoparasite species change from their cylindrical shape to either a pear or a bag-like appearance. Reproduction can be sexual, hermaphrodite or parthenogenic, namely a form of reproduction based on the development of unfertilized female sex cells (Ben-Ami and Heller 2005). As a result, the control of the species is even more difficult because, once the nematode hatches during stage J2, it is free-living and search for a host as a source of food. Phytoparasitic nematodes are generally classified according to their location in the vegetable tissue, known as the type of biotrophic relationship established with the host plant (Bello et al. 1994). Their persistence on tissue for long periods indicates the establishment of a very complex host-pathogen relationship, being subject of intense research (Stephen et al. 1998; Sanz-Alférez et al. 2000). Table 2 presents the classification of phytoparasitic nematodes (Guzmán et al. 2012; Armendáriz et al. 2015). Sedentary endoparasitic nematodes that form root knots and cysts currently cause the most damage to agricultural crops and are the most difficult to control causing as well significant crop losses (Trudgill and Blok 2001). The life cycle of these organisms is described in detail below.

Table 2 Classification of phytoparasitic nematodes and their characteristics
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Life cycle of sedentary endoparasitic nematodes

The lifecycle of endoparasitic nematodes comprises five stages (Mandal et al. 2021) (Fig. 1). The first is the juvenile stage J1, established with the molting or cuticle change that occurs inside the egg, wherein the female RKNs lay their eggs directly on the roots, while the CNs eggs remain inside the body of gravid females, forming a protective cyst (Perry 1989). The presence of root exudates of a host plant stimulates egg eclosion and emerges during the second juvenile stage J2, that is the free-living stage in the soil where nematode infects its host, and once J2-stage individuals have penetrated the root, they remain fixed to hosts and subterranean stems (Talavera 2003). The infection process occurs when the nematode reaches the host plant and perforates its cell walls, enzymatically softening them with its oral stylet. Cyst nematodes migrate intracellularly, via the cortex, directed to the vascular cylinder, where they induce specialized feeding structures (Fig. 1a), while J2-stage RKNs migrate intercellularly and feed on the giant cells that are formed via repeated cycles of mitosis without cytokinesis (Fig. 1b) (Jones et al. 2011; Davies et al. 2015; Rehman et al. 2016). Growth toward the interior of the cell wall occurs together with elements of xylem, which facilitates the absorption of nutrients in the developing syncytium (Fosu-Nyarko et al. 2016). Nematodes grow rapidly in size and undergo molting while being transformed into juvenile stages J3 and J4, while the last stage of the life cycle occurs when the nematodes enter the adult phase, becoming sexually dimorphic, with the females swelling and remaining sessile throughout the parasitic life cycle. On the contrary, adult males recover motility and are attracted by the females in order to inseminate and fertilize the eggs (Perry 1989). Life cycle, from egg to adult stage, may comprise three or four weeks under optimal environmental conditions, especially temperature, but may take longer in cold temperatures (Agrios 2005).

Fig. 1
figure 1

Life cycle of a a cyst-forming nematode and b a root-knot nematode at different stages of development

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Hence, to establish an effective biological control of PPNs, in particular cyst-forming and root-knot nematodes, it should be undertaken prior to the infection of the plant, when nematodes are in J2 stage, where they are free-living in the soil and searching for a host to feed. Meloidogyne spp. an RKN, is highly pathogenic and induces hypertrophy and hyperplasia in the root cells, forming root knots and damaging the plant, thus reducing its yield and predisposing it to infection via other pathogens such as bacteria and fungi (del Cid Prado et al. 2018). The CN of maize, Punctodera chalcoensis, is the second most important in Mexico after the golden nematode, given the impact of the damage it causes to maize crops, reducing maize quality and yield by up to 90% (Sosa-Moss 1987). The seriousness of this damage depends on the susceptibility of the crop, density of the nematode population, and the adequate levels of soil humidity (Nicol et al. 2011). The foliar symptoms consist of delayed growth, chlorosis, occasional leaf necrosis, and flaccidity in young plants, resulting in reduced yield and even, death (Van Gundy et al. 1974). Juvenile nematodes of the first J1 stage remain inactive in the soil for at least one year, until the eggs hatch in search for a host (Perry 1989). Microbial agents have been reported in the literature as methods for the biological control of nematodes. Below, we focus on the nematicidal potential of the nematophagous fungi Purpureocillium lilacinum and Pochonia chamydosporia.

Agents for the biological control of phytoparasitic nematodes

Biological control can be an environmentally friendly pest preventer option compared to that of synthetic chemical nematicides, since it harnesses the power of natural control provided by live organisms and/or their metabolites or subproducts. Biological control involves the use of non-native species derived from animals, plants, bacteria, viruses, and fungi to prevent, eliminate, or, even, reduce the damage to plants or their products (Lopez-y-Lopez et al. 2000; Armendáriz et al. 2015; Khan et al. 2020). Predatory organisms, such as parasitic fungi, nematodes, microarthropods, and pathogens, such as viruses and bacteria, are natural enemies of PPNs and reduce the size of their populations. While the control exerted by these organisms has been found to be effective under laboratory conditions, effective results have not been completely obtained via their application in the field, due to environmental circumstances (Galindo et al. 2013). Nematophagous bacteria and fungi are among the main microbial groups with potential as biological agents for the control of RKNs and CNs (Khan et al. 2020), given the need for environmentally beneficial and low-cost alternatives to chemical measures for the control of PPNs, without affecting vertebrates, crops, and other non-target organisms. Therefore, highly specific antagonists, preferably soil-borne, are the most appropriate for the above-described objective. Diverse entomopathogenic, nematophagous, and fungicides have been reported for the control of soil-transmitted insect pests and plant pathogens; from which nematophagous fungi may fulfil an analogous function for the biocontrol of PPNs (Degenkolb and Vilcinskas 2016). Table 3 shows the main characteristics of nematophagous fungi, which can be grouped as scavengers (predators), endoparasites, parasites of eggs and females or toxin producers, according to their mode of infecting nematodes.

Table 3 Classification of nematophagous fungi
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Although nematophagous fungi show a variable specificity regarding to the nematode species they infect, in general, they infect vermiform and/or nematode eggs. Purpureocillium lilacinum presents superior attributes for infecting nematode eggs, using its conidia to penetrate inside (Sagüés et al. 2011), while P. chlamydosporia var. chlamydosporia infects nematode eggs and J2-stage juveniles by means of its ability to produce secondary metabolites such as aurovertins and pochonins, among others (Shinonaga et al. 2009; Zhou et al. 2010). The mechanism of interaction between fungi and nematode begins during the immobile stage J1, when the nematode is yet to leave the egg. P. chlamydosporia hyphae grow towards the eggs, forming appressoria with its hyphae tips that penetrate the eggshell, then digesting the content of both immature and mature (containing juveniles) eggs (Nordbring-Hertz et al. 2006). Larriba et al. (2014) indicate that this parasitic mechanism comprises three stages: (1) adhesion, wherein the hyphae recognizes the surface and components of the chorion and secretes glycoproteins for egg adhesion; (2) an appressorium in the hyphae tip, via the secretion of proteases (P32, VCP1, and SCP1) which expose the chitin layer of the egg (in turn degraded by chitinases) that enables the penetration phase and finally, (3) the fungus colonization inside the egg affecting the first developmental stage of embryos as well as J1 to J2 juveniles actively developing. Subsequently, the fungus absorbs nutrients such as trehalose, a carbohydrate essential for the development and survival of the PPN and also for physiological processes, such as the hatching of the egg, development and growth during the different nematode developmental stages, sugar transport, energy accumulation and protection of somatic cells (Behm 1997; Sellito et al. 2016; Avelar et al. 2017; Silva-Valenzuela et al. 2020).

Figure 2 depicts some structural chemical forms related to a variety of metabolites produced by nematophagous fungi, for example, there are 139 compounds reported for P. chlamydosporia (Niu 2017; Silva-Valenzuela et al. 2020), from which aurovertins D, E, F, I and the phomalactones present nematicide activity against PPNs (Hellwing et al. 2003; Shinonaga et al. 2009; Niu et al. 2010; Zhou et al. 2010; Kumar et al. 2013; Wang et al. 2015; Bogner et al. 2017; Niu 2017). Similarly, Fig. 3 shows the main compounds of P. lilacinum with nematicide activity, among which 2-ethyl butyric acid, phenyl ethyl alcohol, benzoic acid, benzene acetic acid, 3,5-Di-t-butylphenol (Sharma et al. 2020) and ethyl acetate (Sharma et al. 2014) present an effective nematicide action against M. incognita, inhibiting the eclosion of the egg mass and the growth of juveniles during stage J2.

Fig. 2
figure 2

Structure of compounds that show nematicidal activity produced by P. chlamydosporia

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Fig. 3
figure 3

Structure of compounds that show nematicidal activity produced by P. lilacinum

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Challenges for the production of the nematophagous fungi Pochonia chlamydosporia and Purpureocillium lilacinum

According to the information described before about how nematophagous fungi exert their effect on PPNs, it is important to preserve their attributes of action, either in terms of their spores, chlamydospores or produced metabolites. However, one of the main challenges is to reduce the fermentation times reported, since the evidence shows that, metabolites of interest are produced in more than ten days, in specific culture media. This could complicate industrial production, due to the fact that longer times and greater scales for production implies higher energy costs of maintaining temperature, aeration, agitation and higher labor costs, given the duration of the process. Studies conducted mainly on submerged fermentation are presented below, along with a description of production conditions that could be applied to develop robust processes for mass production of nematophagous fungi.

Dube and Smart (1987) conducted experiments of P. lilacinum (Thorn) Samson at flask level (500 mL) with periodical agitation for 10 days at 25–30 °C in soaked and drained sterile wheat seeds, adding 4 g of wheat seed inoculated with 4 × 107 conidia. It was observed a suppression of the RKNs of M. incognita, at their egg mass, and hatching in greenhouse experiments conducted on tomato (Lycopersicon esculentum), tobacco (Nicotiana tabacum), and pepper (Capsicum annuum). Cabanillas and Barker (1989) produced P. lilacinum in plastic bags containing wheat kernels inoculated with a spore suspension (3.5 × 107 spores/mL), incubated for 21 days at 25 °C, obtaining 3.5 × 109 spores/g of wheat kernels. In the other hand, Cabanillas et al. (1989) tested the growth temperature (12, 16, 20, 24, 28, 30, 32, 34, and 36 °C) of 13 P. lilacinum isolates in a 125 Erlenmeyer flask containing 20 mL of potato dextrose broth inoculated with a 5 mm disc containing a 7-day-old PDA culture, agitated manually and then incubated in the dark. They found that optimum temperatures for P. lilacinum ranged from 24 to 30 °C, although they also observed similar growth patterns obtained varying levels of mycelium production. Interestingly, the best results for the control of M. incognita in tomato plants were obtained using an isolate from Peru and a mixture of P. lilacinum isolates, with the Peruvian strain PL 84 − 1 achieving moderate growth (3.23 mg/mL), lower than the 7.1 mg/mL obtained using one of the strains tested before.

Kerry et al. (1986) evaluated the conidia production in Czapek broth with the addition of trace elements from different P. chlamydosporia strains, using a 150 mL conical flask with 75 mL of medium incubated at 19 °C, 180 rpm for 28 days, obtaining large numbers of conidia (up to 8.4 × 107/mL) but not chlamydospores. Stirling and Smith (1998) developed granular formulations containing either P. chlamydosporia or Arthrobotrys dactyloides produced in glucose-peptone yeast or glucose-corn steep, respectively, in a two-liter Erlenmeyer flask. They harvested 10 g of dry mycelia from 1-liter liquid medium, using the biomass obtained to produce 1 kg of granules. They also reported that A. dactyloides was more effective in controlling RKNs in field trials conducted on a tomato plantation in Queensland, Australia. It was also reported the evaluation of media containing yeast extract, peptone, soybean meal, cotton seed meal, crushed maize meal, neopeptone, or malt extract in combination with 40 g/L glucose, which was used for the mass production of P. chlamydosporia at flask level, with 30 mL of medium incubated in 125 mL Erlenmeyer flasks at 25 °C and 200 rpm for 5 days (Stirling et al. 1998). They found that media containing either cottonseed meal or soybean flour were the most suitable for biomass production, obtaining 18 g/L and 15 g/L respectively, while conidia concentrations of approximately 6 × 108 conidia/mL were obtained for both media. Interestingly, the same study conducted fermentation with YPD medium in a 20-L bioreactor under initial conditions of 0.6 vvm, 200 rpm and 25 °C, making it one of the few studies conducted at bioreactor scale, producing 8–11 g/L biomass over four production runs. The authors describe that chlamydospores were not produced in this submerged culture (Stilring and Smith 1998).

Mo et al. (2005) evaluated the effect of 21 carbon sources and 15 nitrogen sources on the mycelial growth and sporulation of P. chlamydosporia, finding that sweet potato and l-tyrosine are the optimal carbon and nitrogen sources, respectively, for mycelial growth (5.1 g/L), whereas sweet potato and casein peptone were suitable for sporulation (1.71 × 108 conidia/mL). The authors also found different nutritional requirements for sporulation and growth, obtaining maximum conidia production (4.7 × 107 conidia/mL) with a carbon:nitrogen ratio (C:N) of 10 and initial pH of 3.7, while 9.25 g/L of biomass were produced at initial pH of 6.8 and C:N ratio of 40. The incubation conditions were 100 mL medium in a 250 mL conical flask at 170 rpm and 28 °C for seven days. Hernández and Hidalgo (2008) proposed a production method involving solid state fermentation in KlamiC® polypropylene bags using the P. chlamydosporia var. catenulata strain IMI SD 187, selected for its properties as a bioregulator for mass production, since it is able to grow in a pH range of 4–9 at soil temperatures among 9–38 °C. This method is recommended for the management of the Meloidogyne spp. populations that infect tomato, cucumber, capsicum, carrot, beet, and lettuce crops, among others.

The P. chlamydosporia strain YMF 1.00613, isolated from tobacco root infected with M. incognita, was used to isolate and identify four aurovertin-type metabolites, including a new compound, aurovertin I (A1), and three known metabolites, aurovertins E, F and D (shown in Fig. 2) (Niu et al. 2010). The authors obtained a production level of 20 L in 500 mL Erlenmeyer flasks containing 200 mL of production medium, 20% unpeeled potato, and 2% glucose, at a pH of 7, incubated at 28 °C, 180 rpm for 12 days. Both the mycelium and the fermentation broth were filtered and concentrated for characterization. After 48 h, aurovertins D and F presented a nematicide effect on the free-living nematode Panagrellus redivivus, with determined LC50 values of 41.7 and 88.6 µg/mL, respectively. However, the four aurovertins did not present inhibitory effects on the eclosion of M. incognita eggs (Niu et al. 2010). These results are important and reveal the need of producing not only mycelium and spores (conidia or chlamydospores) but also the metabolites and enzymes involved in PPNs inhibition. Wang et al. (2015) cultured different P. chlamydosporia strains, including the YMF 1.00613 strain isolated from tobacco nodules infected with M. incognita, in 250 mL PDB medium contained in 500-mL flasks incubated at 28 °C, 250 rpm for 12 days. The P. chlamydosporia strains that presented nematicide activity produced a distinctive yellow fermentation broth. Chemical studies have shown that yellow metabolites consist of polyketides from the aurovertins D, E, E, and I, with aurovertin D achieving the highest mortality for the RKN M. incognita, with a LC50 value of 16 µg/mL at their stage J2 and 33.50 µg/mL for C. elegans. It was also observed that aurovertin was produced from the fifth day of fermentation until the growth of the fungus concluded.

On the other hand, the addition of chitosan to the culture medium improved sporulation, as seen for the production of extracellular enzymes of P. chlamydosporia, and the parasitic effect on RKN eggs, observed in 50 mL culture medium containing chitosan concentrations of 0.1 mg/mL, 1 mg/mL, and 2 mg/mL as the nutrient source, mineral salts, yeast extract and 1% (w/v) glass wool (Escudero et al. 2016). Compared to the control (without chitosan), it was observed a 2-fold (0.1 mg/mL) to 4-fold (1 to 2 mg/mL) increase in proteolytic activity. The combinations were incubated in 250 mL flasks for 30 days, in darkness. It was also found that chitosan, at a concentration of 0.1 mg/mL, does not affect the viability and germination of chlamydospores and improves mycelial growth compared to cultures without chitosan (Escudero et al. 2017). Silva et al. (2017) grew P. chlamydosporia (var. catenulata and chlamydosporia) and P. lilacinum fungal strains in PDA medium + streptomycin (0.5 g/L) for a period of 18–21 days at 26 °C ± 0.5 °C in darkness. The cells (spores and mycelia) obtained from these cultures were suspended in distilled water + Tween 80 (0.05% v/v). The suspensions were then homogenized and filtered to facilitate the assays development with M. enterolobii nematode eggs, to evaluate their efficacy in reducing root infection and nematode reproduction in potted plants.

Another study cultured P. chlamydosporia in 250 mL Erlenmeyer flasks with dextrose and potato broth, incubated at 25 °C for ten days in a rotary shaker, with the medium then filtered at the end of the incubation. The filtered culture was used to simulate the effect of mortality during M. incognita egg eclosion and at their J2-stage, at 20, 40, 60, 80, and 100% filtrate concentrations diluted with water and taken from six isolates. The impact on mortality was within a range of 11.3–76.3% after 72 h of filtering and increased in line with the concentration. The fungus PC-6 inhibited nematode egg eclosion by 58.17%, with PC-1 achieving the second highest level of inhibition (Uddin et al. 2019).

Shirazi et al. (2019) cultured the fungi P. chlamydosporia and P. lilacinum on solid substrates for two months, using wheat, barley, rice husks, and rice bran, finding that P. chlamydosporia produced spores in wheat, barley, and rice bran after 30 days and after 60 days in rice husks, with an average of 1.5 × 108 and 7 × 107 spores/g, respectively. P. lilacinum colonized all the substrates, with higher spore yields obtained in barley grains and rice husks, with an average of 2.7 × 108 and 1.5 × 108 spores/g, respectively. The viability of both fungi decreased after 60 days of storage at 25 °C.

The production of P. lilacinum KU8 used five different agro-residues, such as wheat bran fine particles, beer waste, sugarcane bagasse, coffee husks and spent tea waste, with 10 g of each substrate placed in a 250 mL Erlenmeyer flask along with 4 mL of a mineral salt medium. The fermentation was carried out at a pH of 4.4 and a temperature of 30 °C for up to 12 h in order to obtain maximum biomass. The isolate produced a biomass of 107.46 mg/gdfs (mg of biomass per gram of dried fermented substrate) in rice bran. The experiment was conducted to evaluate the production of a bionematicide for the management of PPNs (Mousumi Das et al. 2020).

In other study, P. lilacinum 6029 spores were used in a medium based on karanja (a species of tree from the pea family Millettia pinnata) cake to inoculate Erlenmeyer flasks containing 100 mL Czapek-Dox broth and a karanja-cake based broth at a C:N ratio of 35.88 and a pH of 5.9. The fungal culture was incubated in darkness for 7 and 15 days at 27 °C, with the fermentation then filtered and used for in vitro bioassays to evaluate a possible nematicide effect. It was found that the karanja-cake-based culture medium killed 100% of J2 M. incognita nematodes, while a 78.28% mortality was observed for the filtered Czapek-Dox broth 12 h post exposure (Sharma et al. 2014). A recent study conducted by the same authors reported the identification of nematicide metabolites based on a directed fractionation produced using the fungus P. lilacinum grown in a defatted karanja cake-based liquid medium. The mortality rate of M. incognita during egg mass eclosion was 94.6% by the fifth day, while the maximum nematicide effect observed for J2 nematodes was 62% after 48 h of exposure (Sharma et al. 2020). The results reported by Ferreira et al. (2020) show that the application of ketamine, both in vitro and in vivo, confirms the nematicide potential of this molecule in the fungus P. chlamydosporia, which was cultured in a 250 mL Erlenmeyer flask containing a dextrose and potato liquid medium, incubated at 28 °C, 120 rpm for 20 days, filtrating after to separate the extract from the mycelial mass. Separated mass was used for the extraction process via maceration, while the ketamine compound was identified by nuclear magnetic resonance spectroscopy.

According to the information above mentioned, the conditions evaluated for cultures of nematophagous fungi at flask level were oriented mainly to the application against phytoparasitic nematodes instead of their massive production. However, for the case of submerged fermentation, such conditions and raw materials are useful for the process engineering in order to stablish the next approach in bioreactor studies at the laboratory scale and the subsequent steps for scaling-up the production process. In such approach, the effect of parameters such as agitation, aeration, dissolved oxygen tension, mixing, etc., on morphology, spores, conidia, and metabolite production must be elucidated.

Market opportunity

Pesticide market is predicted to reach USD $70.89 billion by 2025 (Industry ARC 2020). The biopesticides represent only 3% with a market of USD $2.2 billion globally and it was estimated to grow over 5% between 2020 and 2026 (Global Markets Insights 2019). For the case of bionematicides market, in 2015 it was valued over USD $143 million with an increment of 4% annual (Global Markets Insights 2015), whereas nematicide market in 2020 was valuated at USD $1.3 billion (Market and Market 2020). Hence, there is a huge market opportunity to replace synthetic pesticides with bionematicides. Table 4 presents the cost comparison between the principal raw materials reported in fermentation process for nematophagous fungi production, the price of biobased nematicides with nematophagous fungi, the cost of synthetic products commercialized in Mexico (local providers), and doses of application. Given these prices, the bionematicides can be competitive compared to synthetic products. It seems logical that agro-based low-cost raw materials can be used for large scale production of nematophagous fungi. However, there are intrinsic steps of pre-treatments for such raw materials and local availability that increases the total process cost. Additionally, the time required for many processes described for cultures of nematophagous fungi comprises several days, and some efforts must be made to reduce such time. A system for the production and formulation of the biological control agent should be developed to obtain a product with a suitable long shelf life, providing it certain competitive advantages in the environment in which it will be applied. Subsequently, the process should be escalated to pilot level to obtain enough product that allows to evaluate its activity in both the greenhouse and the field. Finally, if the product achieves the required attributes and its production is viable at both technical and economic terms, it will be registered and commercialized (Janisiewicz and Korsten 2002). Placing more biobased products with a high activity on the market will not only be an economic benefit but also a huge profit for the environment and human well-being.

Table 4 Cost comparison between some examples of raw materials reported for nematophagous fungi production, biobased products, and synthetic control agents
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Conclusions

Given the growing concerns on the environment, pollution, and health risks caused by many conventional agrochemicals, the demand for natural biological products is constantly increasing in all markets. It is expected that the use of chemical nematicides could be eliminated completely and substituted for biological alternatives, such as the use of fungi and bacteria which, further to killing nematodes, they promote plant growth without causing environmental damage. The aim is to raise awareness of bionematicides on the market in order to increase sustainable agriculture. The development of any biological control agent should take into consideration the life cycle of the target organism to develop the best formulation and application strategies. In order to achieve mass production, biomass, conidia or chlamydospore development is necessary, as it is the production of enzymes and metabolites from nematophagous fungi to evaluate which of them better inhibit PPNs in the short term to prevent the infection of the plant. The development of media for both submerged fermentation and solid-state strategies for production may be a significant step for achieving a robust production process. However, there is also a lack of research on the operating conditions for fermenters at a scale that could satisfy the needs for the application of this technology, which, as we have reported here, depends on the strains, the isolation source, and their ability to maintain its nematicidal activity.