Abstract
Background and goals Powdery mildew (Erysiphe necator), downy mildew (Plasmopara viticola), and botrytis bunch rot (Botrytis cinerea) are among the most destructive diseases affecting grapevines worldwide. These diseases can severely impact both foliage and fruit, leading to substantial yield and quality losses. While cultural practices contribute to disease suppression, chemical control—particularly fungicides—remains the primary method for managing these diseases. Both multisite and site-specific fungicides are widely used, however, the intensive and repeated application of site-specific fungicides increases the risk of resistance development in pathogen populations. This review aims to provide a comprehensive overview of known fungicide resistance in E. necator, P. viticola, and B. cinerea, and to examine tools available for resistance monitoring.
Methods and key findings We summarized findings from recent literature on the mechanisms and prevalence of fungicide resistance across major chemical classes used in viticulture. This review also evaluates diagnostic technologies ranging from traditional bioassays to emerging molecular tools which are currently used or have potential for detecting resistance in vineyard pathogens. Additionally, we highlight recent technological advances that could strengthen resistance monitoring and guide more sustainable fungicide use.
Conclusions and significance Fungicide resistance poses a critical challenge to effective disease management in grape production. This review aims to provide a comprehensive analysis of the current state of known fungicide resistance in E. necator, P. viticola, and B. cinerea, and explores current and emerging diagnostic tools for monitoring fungicide resistance. Practical insights are also provided for researchers, extension personnel, and growers seeking to integrate resistance management into long-term disease control planning.
Introduction
Vitis vinifera is the main grape species used globally for production of wine and blending juice, as well as for table and raisin grapes. While considered as nonpareil due to its superior fruit quality, it is susceptible to most major diseases (Wilcox et al. 2015). Three of these diseases include grapevine powdery mildew (GPM; caused by Erysiphe necator), grapevine downy mildew (GDM; caused by Plasmopara viticola), and botrytis bunch rot (BBR; caused by Botrytis cinerea) (Figure 1). The effective management of these three diseases in most grape production regions requires repeated prophylactic applications of fungicides (Gadoury et al. 2012). The objectives of this review are to provide a comprehensive evaluation of fungicide resistance in these major grape pathogens, and to summarize both the scope of fungicide resistance monitoring using existing and potential diagnostic tools, as well as the advancements in technology that can potentially assist in fungicide resistance management.
A) Typical grapevine powdery mildew (GPM) symptoms on upper side of leaf, B) typical grapevine downy mildew symptoms on upper side of leaf, C) typical botrytis bunch rot symptoms on cracked berries at initial infection and sporulation stages, D) colonies and chasmothecia on infected berries, E) Plasmopara viticola sporulation on lower side of leaf, and F) extensive grayish sporulation of Botrytis cinerea on grape berries during the later stages of infection.
Pathogen etiology and disease management of the Big Three: E. necator, P. viticola, and B. cinerea
E. necator infects all green tissues of the grapevine, with inflorescences and clusters suffering the most economic damage (Figure 1). Berries affected by powdery mildew, if they remain in the clusters until harvest, exhibit lower sugar and anthocyanin levels, higher acidity, and elevated phenylacetic and acetic acid concentrations, adversely affecting wine quality (Stummer et al. 2003, Calonnec et al. 2004). Germination, infection, and growth of the pathogen progress rapidly within the temperature range of 21 to 30°C, peaking at an optimal germination temperature of 25°C (Delp 1954). Management of powdery mildew relies primarily on intensive, preventive fungicide applications. The cost of managing this disease can reach up to 37% of gross production value in regions where it poses a significant threat (Sambucci et al. 2014). Effective management of E. necator on grape berries is most critical from prebloom through ~3 to 4 wk after fruit set (Gadoury et al. 2003). In general, six to 15 fungicide applications are usually applied to avoid losses due to powdery mildew outbreak, depending on the growing season and growing region. V. vinifera cultivars require greater fungicide inputs in regions of high disease pressure, which typically occurs under prolonged moderate temperatures combined with high humidity.
P. viticola infections can be severe in humid regions when average in-season temperatures are between 15 and 25°C, such as in eastern North America (Lalancette et al. 1988). Downy mildew symptoms include oily foliar spots that can merge into necrotic areas and cause premature defoliation and desiccation of clusters (Figure 1). The highly effective cycle of asexual reproduction by sporangia drives the rapid spread of P. viticola (Zachos 1959). Fruit clusters are most susceptible before and at bloom, requiring effective management strategies for grape downy mildew (Kennelly et al. 2005). In warm and humid conditions, seven to eight fungicide applications are generally applied to avoid losses caused by a downy mildew outbreak (as reported at https://extension.umd.edu/resource/downy-mildew-management/).
BBR is a significant threat in temperate climates. The fungus B. cinerea colonizes floral or other plant debris. It can also infect berries early in development, staying inactive in the developing berry until the sugar content reaches ~12 Brix. The fungus can also invade through damaged tissues and cause fruit rot that is characterized by conidial masses (Armijo et al. 2016) (Figure 1). Conidia, the primary inoculum of the disease, is dispersed by wind (McClellan and Hewitt 1973). Integrated pest management (including canopy management to reduce microclimate favorability) is critical for disease management, though chemical control at key developmental stages such as 100% bloom and bunch closure, and from veraison through harvest, is often necessary when temperatures are moderate and conditions are humid (15 to 20°C and >90% humidity) (McClellan and Hewitt 1973, Steel et al. 2011, Ciliberti et al. 2015). In some regions and years, no chemical management intervention is required for BBR control; in other instances, up to four fungicide applications are applied to avoid losses.
A brief history on fungicide use in V. vinifera
The history of routine fungicide on grapes dates back to 1882 in France, when it was discovered that grapevines sprayed with a mixture of copper sulfate and lime effectively controlled GPM and GDM (Millardet 1933). Until the 1940s, chemical disease control mainly relied on the use of inorganic chemicals that were mostly prepared by the users. These inorganic fungicides act on multiple sites by affecting multiple metabolic processes in the targeted pathogen’s cells. Today, use of various multisite fungicides is central to the management of powdery and downy mildews and BBR in grapes. These key multisite fungicides usually act at the contact site, have activity against multiple pathogens, and have a reduced likelihood of selecting for fungicide resistance in the target pathogens. The latter feature is a core component in strategies for management of fungicide resistance. Key multisite fungicides used in grape production include copper compounds (e.g., copper hydroxide and copper oxychloride) for downy mildew, mancozeb for BBR, and sulfur for powdery mildew. Sulfur is among the most widely used pesticides in viticulture and is applied across conventional, organic, and biodynamic farming practices (Neill et al. 2015). Frequency of application and sufficient coverage are the important factors for efficacy of multisite fungicides.
While essential for resistance management, multisite fungicides present several limitations. Due to their lack of systemic or translaminar movement, these fungicides provide protection only on the surfaces where they are directly applied, requiring thorough spray coverage and frequent reapplication to maintain efficacy (Dow et al. 2011). Some, such as copper-based products, carry a risk of phytotoxicity, particularly under conditions of high humidity or when applied to young, tender foliage (Pertot al. 2006). Their effectiveness is also highly weather-dependent, with heavy rainfall reducing residual activity and requiring earlier reapplication intervals (Dow et al. 2011).
Between 1940 and 1970, several important fungicide classes with and without systemic activity were introduced that had a lasting effect on grape disease management in the United States (Horsfall 1975, Morton and Staub 2008). These included dithiocarbamates (1941 to 1961, Fungicide Resistance Action Committee [FRAC] M03, protectant), phthalimide (1952, FRAC M04, protectant), fentin (1954, FRAC 30, systemic), benzimidazoles (1964 to 1970, FRAC 1, systemic), and phthalonitrile (1964, FRAC M05, protectant). Collectively, these fungicides expanded the range of tools available to control downy mildew, powdery mildew, and BBR. This trend was reflected by the drastic reduction in the amount of fungicides used in the U.S., from 136 million kg in 1944 to 68 million kg in 1971 (Gianessi and Reigner 2006, Morton and Staub 2008). Dithiocarbamates (key examples: mancozeb and zineb), phthalimides (key example: captan), and phthalonitriles (key example: chlorothalonil) were widely used as broad-spectrum protectants, though their effectiveness was limited to preventative applications due to their lack of systemic activity. Benzimidazoles represented a major advancement by providing systemic control, but fungicide resistance quickly developed in fungal populations (Yarden and Katan 1993). Fentin compounds were initially effective, particularly against powdery mildew, but their use declined due to concerns over phytotoxicity and environmental impact (Brent and Hollomon 2007, Morton and Staub 2008). Despite these limitations, the introduction of these chemistries laid the foundation for integrated fungicide programs and resistance management strategies in U.S. grape production.
Modern fungicide classes used in grape production that were introduced after the 1970s included dimethyl sterol biosynthesis inhibitors (also known as demethylation inhibitors [DMIs]) (late 1970s to 1980s, FRAC 3, systemic), phosphonates (1980s, FRAC P 07, systemic), strobilurins (quinone outside inhibitors [QoIs]) (1990s, FRAC 11, systemic), anilinopyrimidines (late 1990s, FRAC 9, systemic), carboxylic acid amides (CAAs) (early 2000s, FRAC 40, systemic), and succinate dehydrogenase inhibitors (SDHIs) (mid-2000s to 2010s, FRAC 7, systemic). These fungicides had lower use rates, and while they targeted a range of different pathogens, their fungicidal activity was specific to certain cellular mechanisms in pathogens, limiting their off-target effects. They also had systemic activity to varying degrees, including short distance (translaminar) or long distance (xylem or phloem mobile), which gave them longer efficacy windows, allowing for extended intervals between sprays. Although these qualities are popular among growers because of the flexibility they provide, these same properties are also the reason they can be misused. Fungicide misuse includes curative applications (i.e., applications made after pathogen infection), consecutive applications of the same fungicide class, or poor spray application techniques that reduce coverage. The latter, which may affect fungicide effectiveness in contact products, is initially compensated for in systemic products by improved plant absorption and distribution of the fungicide. Unfortunately, repeated misuse of these modern fungicides has resulted in the selection and build-up of fungicide-resistant pathogens.
Fungicide resistance: A modern production threat founded in evolution
Fungicide resistance refers to the heritable genetic change in a fungal or fungal-like pathogen that results in reduced sensitivity of that pathogen to a particular chemical (Delp and Dekker 1985). Under selection pressure (such as the repeated use of the same active ingredient or fungicide site-of-action), the proportion of the pathogen population that contains that genetic change can build. Over time, that can result in the loss of efficacy for that particular active ingredient or products containing the same fungicide site-of-action in controlling disease at the field-scale.
To date, occurrence of fungicide resistance in plant pathogens has been attributed to the four most common molecular mechanisms: mutations in the fungicide target that can reduce the fungicide’s ability to bind to its target protein; upregulation of adenosine triphosphate-binding cassette (ABC) or major facilitator superfamily (MFS) transporters, which drive efflux pumps; overexpression or duplication of the target gene that can result in an increased amount of the target protein; and epigenetic modifications, including chromatin or histone changes, that modify gene expression and help the pathogen adapt to fungicide stress (Deising et al. 2008) (Figure 2). Additionally, fungicide detoxification or metabolism and other unknown factors can also result in the development of fungicide resistance in pathogen populations (Deising et al. 2008) (Figure 2).
Overview of the interacting roles of pathogen biology, fungicide properties, and management strategies influencing fungicide efficacy and resistance development in Erysiphe necator, Plasmopara viticola, and Botrytis cinerea on grapevines. QoI, quinone outside inhibitors; DMI, demethylation inhibitors; FRAC, Fungicide Resistance Action Committee; SDHI, succinate dehydrogenase inhibitors.
Fungicide group names and FRAC groups registered for management
To mitigate fungicide resistance development, fungicide manufacturing industries have put major efforts toward understanding how fungicides work and the risk for pathogens accumulating resistance to certain fungicides. Companies have also focused on monitoring for baseline fungicide sensitivity in key pathogens under field settings (Morton and Staub 2008). Together with other research communities, the FRAC (www.frac.info) was formed, which focuses on developing fungicide use strategies to mitigate resistance accumulation and monitor situations such as fungicide cross-resistance in pathogens (Morton and Staub 2008).
Part of these efforts were to classify categories of fungicides based on their site-of-action, creating a simplified code by which fungicide users can quickly identify whether they are repeatedly using similar fungicides. In the following sections, we identify common fungicide groups used to control E. necator, P. viticola, and B. cinerea, and describe whether fungicide resistance has been identified to these different groups (Table 1 and Figure 3).
Overview of fungicide groups and active ingredients labeled for management of grapevine powdery mildew (GPM), grapevine downy mildew (GDM), and botrytis bunch rot (BBR) in the United States (U.S.), including known target sites, resistance mechanisms, and global resistance reports (“Europe” in the “Countries with resistance” column represents any European country). FRAC, Fungicide Resistance Action Committee; qPCR, quantitative PCR; LAMP, loop-mediated isothermal amplification; HRM, high-resolution melting; N/A, not available; rhAMP, RNase-H-dependent polymerase chain amplification; MAPK, mitogen-activated protein kinase.
Types of fungicide resistance: cross-resistance, multidrug resistance, and multiple resistance found in Erysiphe necator, Plasmopara viticola, and Botrytis cinerea in grapevines. FRAC, Fungicide Resistance Action Committee; AP, anilinopyrimidines; DMI, demethylation inhibitor; QoI, quinone outside inhibitors; ABC, ATP-binding cassette; MFS, major facilitator superfamily; PP, phenylpyrroles; SDHI, succinate dehydrogenase inhibitors; CAA, carboxylic acid amides.
Methyl benzimidazole carbamates (FRAC 1) – B. cinerea
Methyl benzimidazole carbamate (MBC) fungicides were first launched in the late 1960s. Currently, only thiophanate-methyl is labeled for grapes (Table 1). MBC fungicides work by binding to β-tubulin in the target pathogen. This causes proteins to unfold locally and inhibits polymerization of microtubules, leading to disruption in germ-tube elongation and mycelial growth (Leroux et al. 1999). The pathogen is considered to have a high risk for resistance selection to fungicides in FRAC group 1 (FRAC 2024). Resistance to MBCs has been attributed to multiple point mutations in the β-tubulin gene BctubA in B. cinerea field populations in grapes (Davidse and Ishii 1995, Avenot et al. 2020). Three amino acid changes, including substitution of glutamic acid with alanine (E198A), valine (E198V), or lysine (E198K) at codon 198, and another amino acid substitution where phenylalanine is replaced by tyrosine (F200Y) at codon 200, are considered to be responsible for fungicide resistance (Yarden and Katan 1993, Banno et al. 2008). Resistance to the MBC fungicide thiabendazole (FRAC 1) has been reported in B. cinerea in grapes in Argentina, the U.S., New Zealand, Iran, Sicily, and many European countries (Beever et al. 1989, Leroux et al. 2002, Panebianco et al. 2015, Reaei et al. 2019, Cosseboom and Hu 2021, Harper et al. 2022).
Dicarboximides (FRAC 2) – B. cinerea
Iprodione is the active ingredient in this FRAC group 2 that is commonly used on grapes (Table 1). The exact mode of action of dicarboximides is unclear, however, studies indicate that dicarboximide fungicides primarily act by disrupting the phosphorylation of the high osmolarity glycerol 1 (Hog1) mitogen-activated protein kinase (MAPK), which is regulated by group III HisK and its associated down-stream kinases (Fillinger et al. 2012). The pathogen is considered to have a medium-to-high risk for resistance selection to fungicides in FRAC group 2 (FRAC 2024). Resistance to FRAC 2 in B. cinerea has been attributed to point mutations in Bos1, with substitutions at codon I365 (I365S, I365N, or I365R) being the most common mutations (Oshima et al. 2006). Resistance to the dicarboximides has been reported in B. cinerea in grapes in the U.S., Africa, Canada, New Zealand, Iran, Sicily, and several European countries (Northover 1988, Beever et al. 1989, Fourie and Holz 1998, Leroux et al. 2002, Panebianco et al. 2015, Reaei et al. 2019, Cosseboom and Hu 2021).
Demethylation inhibitors (FRAC 3) – E. necator
DMIs, also known as sterol biosynthesis inhibitors, are considered one of the most successful fungicide classes for GPM control. Currently, six active ingredients are labeled for managing E. necator in grapes (Table 1). DMI fungicides target cytochrome P450 C14α-demethylase (CYP51) in the ergosterol production pathway. DMI fungicides bind to the heme iron of CYP51 via a nitrogen atom, blocking oxygen from binding and transferring to lanosterol’s C14-methyl group. This process stops lanosterol’s C14 demethylation process (Ziogas and Malandrakis 2015). DMIs were introduced in the 1970s, and currently six different DMI fungicides (tebuconazole, difenoconazole, tetraconazole, triflumizole, myclobutanil, and fenarimol) are widely used in viticulture (FRAC 2024). DMI fungicides significantly compromise membrane integrity and function, causing disruption in fungal growth (Jones et al. 2014). The pathogen is considered to have a medium risk for resistance selection to fungicides in FRAC group 3 (FRAC 2024). Resistance to DMI fungicides expresses as quantitative, and consequently, these fungicides have generally maintained their effectiveness even after being used in the market for years (Ziogas and Malandrakis 2015). However, reports of shift in sensitivity of E. necator to DMIs due to its repeated and extensive usage has been observed in various countries, including Australia, Chile, India, South Africa, the U.S., as well as several European countries (Gubler et al. 1996, Délye et al. 1997, Halleen et al. 2000, Frenkel et al. 2015, Hall et al. 2017).
Resistance mechanisms to DMIs include point mutations in the CYP51 gene, overexpression of efflux pumps and/or overexpression of the CYP51 gene, and copy-number variations of the CYP51 gene (Ziogas and Malandrakis 2015). The amino acid substitution of tyrosine by phenylalanine (Y136F) at codon 136 of the CYP51 gene, conferring DMI resistance in E. necator, was first reported in 1997 (Délye et al. 1997). Additionally, another synonymous mutation, A1119C, has been associated with CYP51 overexpression and azole resistance in E. necator. Although this mutation does not change amino acid sequence, it could possibly influence mRNA stability or be linked to another, unknown mutation in the promoter region that causes overexpression (Délye et al. 1997, Kunova et al. 2021). A combination of the Y136F mutation and CYP51 copy-number variation has been linked to DMI resistance in E. necator, suggesting that having extra copies of a fungicide-tolerant allele confers an advantage in resistance development (Jones et al. 2014), although this concept of the polygenic nature of DMI resistance has been controversial (Dyer et al. 2000). In other pathogens, other genetic factors have been hypothesized to play a role in increased DMI resistance, but only when the primary resistance gene is already present (Dyer et al. 2000, Kunova et al. 2021).
Phenylamides (FRAC 4) – P. viticola
Phenylamides (PA) were introduced to target oomycete pathogens in 1977 (Gisi and Sierotzki 2015). These fungicides provide long-lasting preventive effects, systemic movement within plants, and curative effects against pathogens (Gisi and Sierotzki 2015). Metalaxyl and metalaxyl-M (mefenoxam), available since 1970, are the active ingredients used against P. viticola in worldwide management of GDM (Wicks et al. 2005; Table 1). The mode of action of metalaxyl-M inhibits ribosomal RNA polymerase in P. viticola (FRAC 2024). Metalaxyl fungicide resistance in P. viticola was first reported in 1981 in France (Clerjeau and Simone 1982). In 2004, widespread metalaxyl resistance was observed in 92% of 813 vineyard samples collected in South Africa (Fourie 2004). Metalaxyl resistance in P. viticola has been reported in Australia, France, Italy, India, China, and Japan (Wicks et al. 2005, Sun et al. 2010, Corio-Costet 2015, Ghule et al. 2020). The exact resistance mechanism to this fungicide group in P. viticola is still unknown. A genome-wide association study conducted in Phytophthora capsici identified several candidate genes, including a homolog of yeast ribosome synthesis factor Rrp5, that may be linked to mefenoxam sensitivity (Vogel et al. 2021).
Succinate dehydrogenase inhibitors (FRAC 7) – E. necator and B. cinerea
SDHIs are a class of fungicides that block the activity of the succinate dehydrogenase enzyme, causing mitochondrial respiration to stop. Overall, SDHI fungicides are effective against plant pathogens by preventing germination of spores and development of germ tubes (Leroux et al. 2010). Currently, six different SDHI fungicides are widely used in viticulture to manage B. cinerea and E. necator (FRAC 2024; Table 1). These pathogens are considered to have a moderate-to-high risk for resistance selection to fungicides in FRAC group 7 (FRAC 2024). SDHI fungicide resistance in B. cinerea and E. necator has been reported in grapes in the U.S. and in several European countries (Leroch et al. 2011, De Miccolis Angelini et al. 2014, Cherrad et al. 2018, Alzohairy et al. 2021, Cosseboom and Hu 2021). Mutations occurring in the SdhB subunit of the SDHI gene are known to cause resistance to SDHI fungicides in B. cinerea and E. necator (Cherrad et al. 2018, Cosseboom and Hu 2021). SDHI fungicide resistance is primarily associated with single nucleotide polymorphisms (SNPs) occurring at codon 225, 230, or 272 at the SdhB subunit, leading to P225L/F/T, N230I, or H272L/R/Y amino acid shifts in B. cinerea and H242R and H242Y shifts in E. necator (Leroux et al. 2010, Cherrad et al. 2018). P225F/L and H272L substitutions are linked with the highest levels of boscalid fungicide resistance in B. cinerea (Lalève et al. 2014). Different H272 mutations in the SdhB subunit affect SDHI sensitivity of different active ingredients in distinct ways. For instance, the H272L amino acid substitution causes reduced sensitivity to all SDHI fungicides, while the H272Y amino acid substitution causes hypersensitivity (opposite of fungicide resistance) to fluopyram (Lalève et al. 2014).
Anilinopyrimidines (FRAC 9) – B. cinerea
Anilinopyrimidine (AP) fungicides inhibit methionine biosynthesis and suppress the release of cell wall-degrading enzymes that are essential for fungal infection. Two active ingredients are currently labeled for management of BBR in grapes (Table 1). The pathogen is considered to have a medium risk for resistance selection to fungicides in FRAC group 9 (FRAC 2024). Despite their effectiveness, resistance to AP fungicides has been reported in B. cinerea isolates across multiple crops, including grapes in the U.S., Chile, several European countries, and Australia (Leroux and Gredt 1995, Saito et al. 2019, Harper et al. 2022). Though resistance mechanisms remain unclear, mutations in nine distinct genes (Bcmix17, Bcdnm1, Bcafg3, Bcphb2, Bcmcr1, BcoliC, Bcatm1, Bcmdl1, and Bcpos5) that are associated with mitochondrial metabolism have been linked to AP fungicide resistance (Mosbach et al. 2017).
Quinone outside inhibitors (FRAC 11) – E. necator, P. viticola, and B. cinerea
QoIs, commonly known as strobilurin fungicides, are one of the most commonly used fungicides for grapes. These fungicides were originally derived from natural compounds like strobilurin A and oudemansin A (Hirooka and Ishii 2013, Kunova et al. 2021), which are produced by certain basidiomycete fungi and are optimized to improve light stability and reduce toxicity to mammals (Kunova et al. 2021). Azoxystrobin and kresoxim-methyl were the first QoI products introduced to the market in 1996 (FRAC 2024). Currently, six different active ingredients are registered in grapes for management of GPM, GDM, and BBR (FRAC 2024; Table 1).
QoI fungicides operate by targeting mitochondrial respiration in a broad range of fungi phyla, including Ascomycetes, Basidiomycetes, Deuteromycetes, and Oomycetes. QoI fungicides specifically bind to the quinol oxidation (Qo) site of cytochrome b in the inner mitochondrial membrane, blocking electron transfer to cytochrome c1 and ultimately stopping the production of adenosine-5′-triphosphate (Bartlett et al. 2002). As a result, these fungicides are particularly effective against plant pathogens by disturbing energy-dependent processes like spore germination and zoospore mobility (Bartlett et al. 2002). The pathogen is considered to have a high risk for resistance selection to fungicides in FRAC group 11 (FRAC 2024).
QoI resistance is reported in E. necator in the U.S. (Baudoin et al. 2008). The G143A mutation, an amino acid substitution from glycine to alanine in the cytochrome b (Cyt b) gene, is the most common cause of resistance in pathogens, including E. necator, B. cinerea, and P. viticola (Gisi et al. 2002, Grasso et al. 2006, Chen et al. 2007, Corio-Costet 2015). This mutation can cause a significant resistance factor greater than 1000. The primary mechanisms of resistance in the field isolates of E. necator and P. viticola are the mutations of G143A, F129L, and G137R, as well as the activation of the alternative oxidation pathway (Sierotzki et al. 2005, Miles et al. 2012). QoI resistance in E. necator has become widespread in grapegrowing regions across all U.S. states where QoI fungicides are used, as well as in multiple European countries, Australia, and Canada (Baudoin et al. 2008, Dufour et al. 2010, Miles et al. 2021). Similarly, QoI resistance in P. viticola has been reported in several European countries as well as in the U.S., Canada, Japan, Brazil, India, and China (Sierotzki et al. 2005, Toffolatti et al. 2007, Sharma et al. 2022, 2025). QoI resistance has also been reported in B. cinerea in various European countries, Australia, and the U.S. (De Miccolis Angelini et al. 2014, Saito et al. 2019, Alzohairy et al. 2021, Harper et al. 2022).
Phenylpyrroles (FRAC 12) – B. cinerea
Phenylpyrroles, represented by fludioxonil (Table 1), target the Bos1- and MAPK-dependent osmoregulation pathway, causing hyperosmolarity, glycerol accumulation, and inhibition of fungal growth (Vignutelli et al. 2002). Resistance to fludioxonil in B. cinerea has been linked to mutations in the mrr1 transcription factor. These mutations cause the overexpression of atrB, which in turn leads to increased activity of drug efflux transporters. Specifically, a point mutation at codon 632 of mrr1 and a 3-base pair deletion at codon 497, resulting in the loss of a leucine (L) in the Mrr1 protein, have been identified in fludioxonil-resistant grapes (Toffolati et al. 2020). Fludioxonil resistance in B. cinerea has been found in several European countries, Australia, Chile, Israel, and the U.S. (Korolev et al. 2011, Latorre and Torres 2012, Toffolatti et al. 2020, Cosseboom and Hu 2021, Harper et al. 2022).
Azanaphthalenes (FRAC 13) – E. necator
Azanaphthalenes are a group of fungicides with activity restricted to fungi from the Erysiphaceae family. Two active compounds, proquinazid and quinoxyfen, are approved for management of GPM (FRAC 2024; Table 1). While quinoxyfen was withdrawn from the European Union market in March 2020 (it is still used in the U.S.), proquinazid remains authorized for use (Kunova et al. 2021). Although the precise mechanism of action remains unclear, azanaphthalenes are known to disrupt signal transduction and serine esterase functions (Wheeler et al. 2003). These fungicides are mainly considered protectants and have localized systemic activity along with redistribution through vapor, and are known for their ability to stop early development of powdery mildew by halting spore germination and appressorium formation. Moreover, it has been observed that proquinazid can express defense-related genes in Arabidopsis thaliana by activating pathways related to ethylene signaling, phytoalexin production, reactive oxygen species generation, and pathogenesis-related genes (Genet and Jaworska 2009).
The pathogen is considered to have a medium risk for resistance selection to fungicides in FRAC group 13 (FRAC 2024). E. necator isolates resistant to quinoxyfen have been found in Europe (Genet and Jaworska 2009), and potential cross-resistance between resistance to quinoxyfen and proquinazid has also been identified across Europe (FRAC 2024). In field trials conducted in western New York during 2010 and 2011, a reduced efficacy of quinoxyfen in controlling E. necator was associated with suspected resistance to quinoxyfen (Wilcox and Riegel 2012a, 2012b). Furthermore, high quinoxyfen resistance in E. necator field isolates was found in leaf disc assays, but quinoxyfen still remained highly effective in 3-yr field trials conducted in Virginia (Feng and Baudoin 2018).
Keto reductase inhibitors (FRAC 17) – B. cinerea
Keto reductase inhibitors, represented by fenhexamid (Table 1), act by disrupting the activity of 3-ketoreductase, an enzyme involved in the C-4 demethylation process of ergosterol biosynthesis, which is critical for fungal membrane integrity. The pathogen is considered to have a low-to-medium risk for resistance selection to fungicides in FRAC group 17 (FRAC 2024). Resistance to fenhexamid in B. cinerea has been linked to multiple point mutations in the erg27 gene in grapes (Albertini and Leroux 2004, Fillinger et al. 2008, Esterio et al. 2011). Highly resistant HydR3+ isolates of B. cinerea in grapes exhibit amino acid substitutions at codons 63 (T63I), 412 (F412S/I/V/C), and 496 (T496R) (Albertini and Leroux 2004, Fillinger et al. 2008). In contrast, HydR3− isolates display weak-to-moderate resistance, with substitutions such as F26S, L195F, V309M, A314V, S336C, N369D, L400F, L400S, P238S, I199L, and Y408S, and a deletion at P298 (Fillinger et al. 2008, Esterio et al. 2011). In grapes, fenhexamid resistance in B. cinerea has been found in several European countries, Chile, and the U.S. (Albertini and Leroux 2004, Fillinger et al. 2008, Esterio et al. 2011, Alzohairy et al. 2021).
Polyoxins (FRAC 19) – E. necator and B. cinerea
Polyoxin (active ingredient polyoxin-D zinc salt [Table 1]) functions by inhibiting chitin synthesis, thereby disrupting the formation of fungal cell walls (Mamiev et al. 2013). Although polyoxin exhibits lower antifungal efficacy compared to many conventional fungicides, it may still serve an important function in integrated disease management programs, particularly as a rotation partner to help delay the development of fungicide resistance. Pathogens are considered to have a medium risk for resistance selection to fungicides in FRAC group 19 (FRAC 2024). Currently, resistance to E. necator and B. cinerea in grapes has not been reported; however, reduced sensitivity to polyoxin-D zinc salt has been observed in B. cinerea isolates collected from strawberry fields in South Carolina, North Carolina, Maryland, Virginia, and Ohio (Dowling et al. 2016).
Quinone inside inhibitors (FRAC 21) – P. viticola
Cyazofamid is an active ingredient in the quinone inside inhibitor (QiI) group that has activity against P. viticola (Table 1). The molecular mechanism of this fungicide class inhibits mitochondrial respiration of the pathogen. QiI fungicides bind to cytochrome bc1 at the quinone inside (Qi) site and disrupt complex III (Cherrad et al. 2023, Sharma et al. 2025). In general, QiI fungicides stop the quinol reduction in the QiI site (Cherrad et al. 2023). The pathogen is considered to have a moderate-to-high risk for resistance accumulation to fungicides in FRAC group 21 (FRAC 2024). Reports of resistance to QiI fungicides in P. viticola are rare but have been linked to specific mutations in the Cyt b gene (Cherrad et al. 2018, 2023). This involves an amino acid shift from leucine to serine at codon 201. Another change, two insertions of two amino acids, that occurs between codons 203 and 204 (E203-DE-V204 or E203-VE-V204) in the Qi site has also been associated with QiI fungicide resistance in P. viticola (Cherrad et al. 2018, 2023). To date, QiI resistance has only been reported in France. Furthermore, a recent survey found no evidence of resistance to QiI fungicides in the eastern U.S. and Canada (Cherrad et al. 2018, 2023, Sharma et al. 2025).
Carboxylic acid amides (FRAC 40) – P. viticola
Carboxylic acid amides (CAA) are one of the most extensively used fungicide groups to manage P. viticola worldwide (Gisi and Sierotzki 2015). Dimethomorph has been utilized to control grape and cucurbit downy mildew since the late 1980s and currently, mandipropamid is also used for grapes (Table 1). The molecular mechanism of CAA fungicides against targeted pathogens inhibits cell wall biosynthesis through targeting the cellulose synthase 3 (CesA3) gene (Gisi et al. 2007, Blum et al. 2010). The pathogen is considered to have a low-to-medium risk for resistance accumulation to fungicides in FRAC group 40 (FRAC 2024). Fungicide resistance to dimethomorph in P. viticola was first noticed in France in 1994 but did not affect field efficacy at that time (Sierotzki et al. 2011).
Resistance to CAA fungicides in P. viticola has been traced to a SNP that leads to an amino acid change from glycine to serine at codon 1105, and less frequently, to valine in the PvCesA3 gene (Blum et al. 2010). Interestingly, CAA fungicide resistance in P. viticola is controlled by a recessive nuclear gene, meaning both alleles are required to possess the resistant phenotype (Blum et al. 2010). Resistance to CAA fungicides in P. viticola populations has been reported in various grapegrowing regions, including Europe, India, Japan, China, and Brazil, as well as Virginia and North Carolina in the U.S. (Blum et al. 2010, Aoki et al. 2013, Sawant et al. 2016, Feng and Baudoin 2018, Santos et al. 2020). In a recent survey conducted in the eastern U.S. and Canada, CAA fungicide resistance was found in P. viticola from Michigan, Georgia, New York, Wisconsin, and Ontario (Sharma et al. 2025).
Quinone inside outside inhibitor (FRAC 45) – P. viticola
Ametoctradin, an active ingredient classified under quinone inside outside inhibitor (QioSI), is an effective fungicide used to control P. viticola populations (Table 1). The mode of action of this fungicide involves targeting both the Qi and Qo sites of complex III, interacting through a stigmatellin-like binding mechanism to disrupt mitochondrial respiration (Cherrad et al. 2023). The pathogen is considered to have a high risk for resistance selection to fungicides in FRAC group 45 (FRAC 2024). QioSI fungicide resistance has been observed in France in 2018 and in India in 2023 (Fontaine et al. 2019, Cherrad et al. 2023, Sagar et al. 2023). Resistance to QioSI fungicides has been attributed to an amino acid shift at codon 34 from serine to leucine (S34L) at the Cyt b gene. So far, this QioSI fungicide resistance is not widespread and has not been detected in the eastern U.S. or Canada (Sharma et al. 2025). However, the likelihood of resistance accumulation is high in this fungicide class; therefore, continuous monitoring is important to increase the longevity of fungicide efficacy in fields.
Aryl-phenyl ketones (FRAC 50) – E. necator
Aryl-phenyl ketones (FRAC 50) is a relatively new group of fungicides mainly used to control powdery mildews in multiple crops. The two main fungicides in this fungicide class are metrafenone and pyriofenone (Kunova et al. 2021; Table 1). The exact mechanism of this fungicide class is not entirely understood, however, studies on cereal powdery mildews suggest these fungicides may disrupt the actin cytoskeleton and affect hyphal morphogenesis, polarized growth, and cell polarity (Opalski 2005, Kunova et al. 2021). The pathogen is considered to have a medium risk for resistance selection to fungicides in FRAC group 50 (FRAC 2024). Resistance to these fungicides began emerging shortly after their introduction. Resistance to metrafenone was reported in E. necator in Europe in 2016 and in Serbia in 2023 (Kunova et al. 2016, Vojinović et al. 2023). Both moderate and high levels of resistance have been detected in E. necator (Kunova et al. 2016). Cross-resistance to pyriofenone has been observed in various European countries, though their prevalence and distribution can vary (Graf 2017).
Phenylacetamide (FRAC U06) – E. necator
Cyflufenamid, the active ingredient in this group of fungicides registered for use on grapes, was approved for use in Japan and Europe in 2002 and 2005, respectively (Kunova et al. 2021), and registered in 2012 for management of powdery mildew on grapes in the U.S. (Table 1). It has proven to be highly effective both as a preventive and curative treatment against powdery mildew on cereals, vegetables, hops, and grapes, as well as against brown rot in stone fruits (Dietz and Winter 2019). Cyflufenamid is also known for its ability to move in the vapor phase and penetrate plant tissue with translaminar activity (Dietz and Winter 2019).
Its specific biochemical mode of action has yet to be identified, which is why it is classified in an “unknown” group (i.e., “U”06) (FRAC 2024). Cyflufenamid does not prevent spore germination or appressorium development in Blumeria graminis f. sp. tritici, but instead strongly inhibits the formation of haustoria, thereby stopping further growth of the fungus (Haramoto et al. 2006). While no resistance to cyflufenamid has been reported in E. necator, it has been reported in Podosphaera xanthii (Pirondi et al. 2014, Vielba-Fernández et al. 2020).
Thiazolidine (FRAC U13) – E. necator
Flutianil, launched in 2013, controls powdery mildews in grapes and is the only fungicide classified in this group (Table 1). Flutianil’s exact mode of action is unknown, but studies show it does not affect early infection stages in B. graminis f. sp. hordei, while effectively blocking haustoria formation and inhibiting fungal progression. It also disrupts nutrient absorption, prevents secondary hyphae elongation, and alters the extra-haustorial matrix and fungal cell wall (Kimura et al. 2020). This fungicide does not significantly affect B. graminis survival genes, but alters the expression of multiple effector and three sugar transporter genes mainly active in haustoria (Kimura et al. 2021). E. necator resistance to flutianil has not been reported, but resistance has been found in P. xanthii in Japan (Miyamoto et al. 2020).
Fitness cost associated with fungicide resistance
The accumulation of fungicide resistance in a pathogen population poses a significant challenge in disease management. While fungicide resistance offers a selective advantage in the presence of fungicides, mutations that confer resistance may also have fitness costs, creating an evolutionary trade-off (Figure 4). These fitness costs may appear in traits like slower mycelial growth, increased abiotic stress sensitivity, reduced aggressiveness and spore production, stress sensitivity, fewer sexual fruiting bodies, and lower viability in the field (Hawkins and Fraaije 2018). For example, B. cinerea isolates resistant to phenylpyrroles fungicides (FRAC 12) have an unknown fitness disadvantage and are outcompeted by sensitive genotypes in field conditions if selection pressure is not applied (Chen et al. 2016). For some fungicides, resistance may not significantly compromise the overall fitness of the resistant population compared to the sensitive population. For example, in B. cinerea, populations with the E198A mutation have a very small fitness penalty following the discontinuation of MBC fungicides (FRAC 1) (Walker et al. 2013) (Figure 4).
A conceptual overview showing selection of fungicide resistance and the effect that the fungicide resistance fitness costs have on individuals in a pathogen population once fungicide selection pressure is removed. FRAC, Fungicide Resistance Action Committee.
A better understanding of how the fitness cost fungicide resistance may affect pathogen populations at the field level could aid in designing better fungicide programs that focus on resistance mitigation (Hawkins and Fraaije 2018). If resistance mutations impose a fitness cost, resistant isolates are likely to decline in frequency when fungicide pressure is relaxed, meaning that sensitive isolates can reestablish dominance. This provides an opportunity for integrated resistance management strategies such as rotating chemistries, implementing fungicide-free intervals, or reintroducing fungicides that had previously lost efficacy. Conversely, if resistance carries little or no fitness penalty, resistant populations can persist even without fungicide pressure, limiting management options and making resistance effectively irreversible. This can be done with both between-season and within-season population monitoring. Between-season monitoring involves evaluating the population dynamics of fungicide-resistant and -sensitive genotypes between seasons to determine whether there is a selective disadvantage in the absence of fungicide pressure and at a time when disease risk might be low, or whether the particular pathogen may be overwintering in a different reproductive state (asexual, sexual). Similarly, observing within-season dynamics in response to fungicide pressure or rotation can reveal whether resistant populations decline during periods without selection pressure (or are maintained with selection pressure), potentially allowing for the reintroduction of previously ineffective fungicides later in the same season. Fundamentally, the baseline approach to reducing the risk of selecting for field-scale fungicide resistance that might result in a disease control failure is monitoring: both for the pathogen and for molecular or genetic markers that can be associated with a resistance phenotype.
Diagnostics for monitoring pathogens and genotypes at the field scale
Detection of fungicide resistance in plant pathogens can be done using conventional methods, including testing spore germination or mycelial growth using discriminatory doses (Miles et al. 2012). While these methods are generally straightforward and allow for the tracking of fungicide sensitivity changes, they often do not provide information about the mechanism of resistance development. These methods are also time consuming, which restricts the number of field isolates that can be tested. Finally, traditional methods require isolating pathogens from the field and growing them in pure culture. That process brings complexities of poor germination or loss of pathogenicity, culture maintenance, and in-planta culturing of obligate pathogens, among other challenges (Massi et al. 2021).
To overcome these limitations, significant progress has been made in developing molecular techniques for detecting and tracking the fungicide resistance of plant pathogens. In cases where the resistance mechanism is known, several nucleic acid detection techniques have been developed in E. necator, B. cinerea, and P. viticola.
Ease of resistance monitoring using rapid molecular techniques depends on factors underlying the resistance mechanism of the fungicides. For example, molecular monitoring of fungicide resistance is considerably more tractable for QoI fungicides compared to DMI and SDHI classes. Resistance to QoIs is most commonly associated with a single, well-characterized point mutation (e.g., G143A in the cytb gene), enabling robust and high-throughput detection via allele-specific PCR and isothermal techniques. In contrast, resistance to DMIs is polygenic and involves multiple mutations in the CYP51 gene, promoter alterations leading to overexpression, and quantitative resistance patterns that complicate molecular detection (Miles et al. 2021). SDHI resistance also arises through multiple amino acid substitutions across subunits of the succinate dehydrogenase complex (SdhB, SdhC, and SdhD), necessitating multiplex quantitative PCR (qPCR) or sequencing-based approaches to comprehensively capture resistance allele diversity (Alzohairy et al. 2021). Consequently, while QoI resistance can be reliably diagnosed using relatively simple molecular tools, deploying similar tools for DMI and SDHI resistance is more technically demanding and less definitive without complementary phenotypic data. These constraints likely reduce their suitability for field-level diagnostics and emphasize the continued importance of centralized diagnostic laboratories for resistance monitoring and decision support.
The timing of molecular resistance diagnostics within the field season is essential to maximize their utility for disease management decision-making. Molecular resistance diagnostics are most effective when conducted early in the growing season, before or at the initiation of fungicide applications, to guide selection of effective active ingredients. Mid-season testing can be valuable for detecting shifts in resistance allele frequencies under ongoing fungicide pressure and for adjusting subsequent spray programs based on the pathogen’s genotypic composition in the vineyard. While late-season assays have limited in-season utility, they provide valuable information for planning future resistance management strategies. Targeted use of these tools at key time points can enhance resistance monitoring and support more informed fungicide deployment.
A brief description of the currently available molecular tools for detecting resistance to different fungicide classes in E. necator, B. cinerea, and P. viticola are described below.
Traditional PCR and qPCR
Three tests are available to monitor QoI resistance in P. viticola based on the G143A mutation described earlier (Gisi et al. 2002, Chen et al. 2007, Corio-Costet et al. 2011). Similarly, SYBR Green-based qPCR assays have been developed to detect G143A mutation in E. necator (Baudoin et al. 2008, Dufour et al. 2010, Miles et al. 2012). However, SYBR Green-based qPCR assays can generate false positives and cannot simultaneously distinguish between A-143 and G-143 alleles in a single reaction, increasing detection costs (Miles et al. 2012).
Furthermore, allele-specific TaqMan probe and digital-droplet PCR-based assays have also been designed to detect QoI resistance in E. necator (Miles et al. 2021). Both assays can be applied at any pathogen developmental stage. TaqMan probe-based qPCR tools are a popular choice due to their adaptability, rapidity, specificity, sensitivity, reliability, reproducibility, and ability to detect multiple alleles. In addition, TaqMan probe-based qPCR tools are successfully used by diagnostic laboratories for detecting the G143A mutation in E. necator.
RNase-H-dependent polymerase chain amplification
Another molecular diagnostic tool, RNase-H-dependent polymerase chain amplification (rhAMP) assays, are utilized for allelic discrimination of the P225F/H mutation associated with fluopyram-boscalid resistance in B. cinerea populations. The rhAMP assays were found to be more flexible in design, to have better allelic discrimination, and to be more affordable compared to TaqMan assays (Alzohairy et al. 2021). However, these assays require expensive equipment and trained operators, making them impractical for point-of-care use by growers.
Isothermal amplification
Isothermal amplification techniques (e.g., loop-mediated isothermal amplification [LAMP]) bypass the need for costly, complex equipment or intensive labor. Isothermal techniques like LAMP offer point-of-care detection for SNP-based fungicide resistance. LAMP assays are highly specific, efficient, and rapid, with amplification completed in under 30 min (Notomi et al. 2000). LAMP reactions can be monitored visually with affordable equipment like block heaters or water baths, or fluorometrically using portable devices such as Genie II (OptiGene Ltd.) or T16-ISO Axxin (Axxin), making it a strong candidate for point-of-care diagnostics (Notomi et al. 2000). LAMP has also been used to detect H272R mutation conferring SDHI-resistance of B. cinerea (Fan et al. 2018). Similarly, the peptide nucleic acid-locked nucleic acid-LAMP assay has been designed for QoI resistance in E. necator (Sharma et al. 2023). While isothermal assays are a promising choice for point-of-care detection, their sensitivity and specificity still require refinement for in-field use (Sharma et al. 2023).
Recently, several CRISPR-Cas-based detection platforms have been developed for detection of human and animal pathogens, offering high specificity, sensitivity, and accuracy. In medical science, a CRISPR-Cas12b system combined with recombinase polymerase amplification has also been shown to effectively detect the 3232A>G mutation in the BRCA1 gene associated with breast cancer (Teng et al. 2019). Incorporating CRISPR-Cas detection systems could be a viable strategy to enhance the specificity and sensitivity of the LAMP assay (Li et al. 2018, Sharma et al. 2023). Although these isothermal techniques can be a better option for rapid detection in the field, increasing grower adoption of isothermal diagnostic techniques would require extensive educational workshops to teach key skills such as pipetting, crude DNA extraction, operation of isothermal devices like the Genie II, and management of contamination risks. Given these technical constraints, it is likely that many growers would prefer to rely on diagnostic laboratories to perform these assays on their behalf.
Conclusion
Fungicides have been indispensable tools for managing grapevine diseases, but the intensive and repeated use of single-site chemistries has accelerated fungicide resistance selection in E. necator, P. viticola, and B. cinerea. Sustaining the effectiveness of these fungicides will require careful stewardship, particularly of high-risk compounds, and the implementation of integrated resistance management strategies. These include rotating fungicides with different modes of action, limiting the number of applications per season for high-risk groups, integrating non-chemical methods (e.g., canopy management, biological controls), and conducting in-season resistance monitoring to optimize spray programs. To prevent resistance buildup, continuous resistance surveillance should be prioritized for newly registered fungicides targeting E. necator, B. cinerea, and P. viticola. Additionally, identifying resistant pathogen genotypes and elucidating underlying resistance mechanisms will support the development of molecular diagnostic tools for rapid detection. Finally, adherence to FRAC-recommended resistance management guidelines for each fungicide group will help preserve their efficacy and ensure sustained control of grapevine diseases.
Data Availability
All data underlying this study are included in the article.
Footnotes
This work was partially supported by the USDA National Institute of Food and Health -Specialty Crop Research Initiative (titled: FRAME: Fungicide Resistance, Assessment, Mitigation and Extension Network for Wine, Table, and Raisin Grapes) code: 2018-03375, and by USDA-NIFA Hatch Project 7005262. Michigan State University is situated on the ancestral, traditional, and contemporary lands of the Anishinaabe, including the Three Fires Confederacy of the Ojibwe, Odawa, and Potawatomi peoples. These lands were ceded in the 1819 Treaty of Saginaw. We recognize the ongoing presence of Indigenous peoples and their enduring connection to this land.
Sharma N, Moyer MM and Miles T. 2025. Evolution of fungicide resistant pathogens in grapes: Erysiphe necator, Plasmopara viticola, and Botrytis cinerea. Am J Enol Vitic 76:0760028. DOI: 10.5344/ajev.2025.25031
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- Received June 2025.
- Accepted September 2025.
- Published online January 2026
This is an open access article distributed under the CC BY 4.0 license.
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