Trap Formation in the Nematode-Trapping Fungi: History
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Jinkui Yang
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Nematode-trapping (NT) fungi play a significant role in the biological control of plant- parasitic nematodes. NT fungi, as a predator, can differentiate into specialized structures called “traps” to capture, kill, and consume nematodes at a nutrient-deprived condition. Therefore, trap formation is also an important indicator that NT fungi transition from a saprophytic to a predacious lifestyle. With the development of gene knockout and multiple omics such as genomics, transcriptomics, and metabolomics, increasing studies have tried to investigate the regulation mechanism of trap formation in NT fungi. And elucidating the mechanism of trap formation will help to develop more effective anti-nematode agents by genetic modification.

  • traps
  • nematode-trapping fungi
  • signaling pathways
  • signal molecules
  • pathway collaboration
  • reverse genetics

1. Introduction

Plant-parasitic nematodes (PPNs) can cause direct damage to their host or act as virus vectors [1]. There are more than 4100 species of PPNs that have an impact on global agriculture and horticulture, causing an estimated annual loss of USD 173 billion [2][3]. Nematode-trapping (NT) fungi are potential biocontrol resources, which have the advantages of low toxicity, high efficiency, and environmental friendliness, so they have gradually become favored by people in recent years [3]. NT fungi can develop specialized structures called “traps”, an important indicator of the transition from saprophytic to predatory lifestyles, including constricting rings, adhesive networks, adhesive columns, and adhesive knobs, to capture, kill, and consume nematodes at a nutrient-deprived condition [4][5]. Therefore, trap formation is a crucial step in the lifestyle of NT fungi and indispensable for nematode predation. The factors involved in trap formation were variety, including multiple signal transduction pathways [6]; small molecular compounds [7][8][9][10][11][12]; intercellular communication [13]; adhesive protein [5]; nitrate assimilation [14]; woronin body [15]; peroxisome [16]; autophagy [17][18][19][20]; and pH-sensing receptors [21], according to the comprehensive research of genomics, transcriptomics, proteomics, metabolomics, and reverse genetics. Here, researchers review the recent progress in the regulatory mechanism of trap formation in the NT fungi based on phenotypes of various mutants and multi-omics analysis. Researchers especially focus on the latest studies in signal transduction pathways and small molecular compounds involved in trap formation. Elucidating the molecular mechanism of trap formation will not only provide a theoretical basis for the improvement of engineering biocontrol fungi, but contribute to understanding the adaptive evolution and lifestyle transition of NT fungi.

2. Multi-Omics Analysis Promotes Research on Trap Formation of Nematode-Trapping Fungi

Omics not only provides a macroscopic direction for the mechanism of trap formation but also provides specific targets. Comparative genomics studies have shown that many species-specific genes have expanded in the evolutionary process, and these genes may be related to the function specialization of NT fungi [5][6][22][23]. Arthrobotrys oligospora (teleomorph Orbilia auricolor), one of the typical NT fungi, was the first NT fungus whose genome and proteome were sequenced [6]. Comparative analysis showed that A. oligospora genome contains numerous pathogenicity-related genes, and a total of 398 homologous genes related to the pathogenicity of other fungi have been identified [6], while there were fewer lectin genes involved in fungus–nematode recognition in the Drechslerella stenobrocha genome [22]. This suggested that a different mechanism of trap formation may exist in various NT fungi. In addition, studies combined genome, proteome, and real-time PCR (RT-PCR) analyses to reveal that multiple signal transduction pathways have an integral part in trap formation [6]. Transcriptome sequencing and RT-PCR analysis showed that a large number of genes were significantly upregulated during the infection process of NT fungi, including genes involved in translation, amino acid metabolism, carbohydrate metabolism, cell wall and membrane biogenesis [6], secreted proteins [23], adhesion proteins [5], and the protein kinase C signal transduction pathway [22]. Comparative genomic analysis in the four representative trapping devices of NT fungi suggested that the simplification of the capture device was accompanied by the expansion of adhesion genes and the increase in adhesiveness on trap surfaces [5]. In conclusion, the above studies show that omics technologies have clarified the general direction for research on the regulation mechanism of traps formation. At the same time, the assembly of genomes in different fungi makes it possible to study the interaction between the NT fungi and nematodes at the molecular level. For instance, according to the genome sequence and annotation of Duddingtonia flagrans, a fluorescent protein system with native promoter was established, and a secretion protein PEFB was identified which was involved in the processes of infection against Caenorhabditis elegans [13]. Recently, nine species of NT fungi have been sequenced, such as adhesive-network-producing fungi A. oligospora and D. flagrans, adhesive-knob-producing fungi Monacrosporium haptotylum and Dactylellina entomopaga, constricting-ring-producing fungi D. stenobrocha and Drechslerella brochopaga, adhesive-columns-producing fungus Dactylellina cionopagum, and no trap producing fungus Dactylella cylindrospora (Table 1); these genomic information may help to elucidate the mechanism of trap formation of NT fungi.

Table 1. Genomic features of different NT fungi.

Trapping Devices

Fungi

Genome Size

GC Content (%)

Number of Genes

Reference

Adhesive network

A. oligospora ATCC24927

40.07 Mb

44.45

11,479

[6]

Adhesive network

A. oligospora TWF154

39.62 Mb

43.96

12,107

[24]

Adhesive network

D. flagrans

36.64 Mb

45.5

9927

[13]

Adhesive knob

M. haptotylum

40.40 Mb

45.24

10,959

[23]

Adhesive knob

D. entomopaga

38.39 Mb

44.9

11,130

[5]

Constricting ring

D. stenobrocha

29.02 Mb

52.5

5597

[22]

Constricting ring

D. brochopaga

35.43 Mb

49.42

10,234

[5]

Adhesive column

D. cionopagum

43.12 Mb

44.3

11,284

[5]

no trapping device

D. cylindrospora

37.71 Mb

46.02

10,785

[5]

3. Overview of Signaling Pathways Involved in Trap Formation

3.1. G-Protein Signaling Pathway Involved in Trap Formation

G-protein signaling is the most conserved signal transduction pathway in fungi, composed of heterotrimeric G-proteins (G-proteins), G-protein-coupled receptors (GPCRs), and regulators of G-protein signaling (RGSs), which plays a vital role in sensing the changes in various physical and chemical stimuli in the environment [25]. G-proteins have three subunits: Gα, Gβ, and Gγ. The latest research identified a single G-protein β-subunit gene (gpb1) in A. oligospora, and phenotypic analyses demonstrated that Δgpb1 mutants were strongly defective in the response to C. elegans and ascarosides, with the formation of few traps [24]. This means that GPCRs might be the receptor of ascarosides. In addition, G-protein signaling pathways are negatively regulated by RGSs. Recently, seven putative rgs genes were knocked out, and multi-phenotypic analyses shown that these genes affect the pathogenicity of A. oligospora to varying degrees. In particular, the ΔflbA deletion strains lost the ability to produce traps after being induced with nematodes [26]. In addition, resistance to inhibitors of cholinesterase 8 (RIC8), a conservative guanine nucleotide exchange factor, which is involved in the regulation of G-protein signaling in filamentous fungi. Recently, an orthologous RIC8 was characterized in A. oligospora, the Δric8 deletion mutants lost the ability to produce traps essential for nematode predation, accompanied by a marked reduction in cyclic adenosine monophosphate (cAMP) level. Further assay revealed that RIC8 interacted with G-protein subunit Gα1 and is involved in nematode predation through control of cell cycle, organelle, and secondary metabolism [27].
The superfamily of small GTPases comprises signal transducers that regulate multiple cellular functions. RAS, RHO/RAC, RAB, ARF, and RAN are conserved groups of the small GTPases family that cycle between GTP-bound (active) and GDP-bound (inactive) conformations as a switch in signal transduction. Recently, two RAB GTPases were identified in A. oligospora: The Δrab-7A deletion strains lost the ability to produce conidia and traps; however, the Δrab-2 mutants only slightly affected the conidiation but did not affect the trap formation [28]. In another research, three RAS GTPases were characterized by gene disruption and multi-omics analysis, the traps number of Δras2 and Δrheb deletion strains were significantly decreased, but Δras3 mutants had no significant changes compared with the wild-type (WT) strain [29]. Our latest research demonstrated that three RHO GTPases (RHO2, RAC, and CDC42) played an important role in trap formation and lifestyle transition of A. oligospora. The rac was significantly upregulated, and alternative splicing events occurred in rac and rho2 during the trap formation and infection process [30]. These studies indicated that the small GTPases have pleiotropic functions in the growth and development of A. oligospora; specifically, they play a very important role in trap formation and pathogenicity. Moreover, GTPase activating proteins (GAPs) are a family of proteins that induce the hydrolysis of GTP bound to small GTPases [31]. Recently, an ARF-GAP GLO3 was identified in A. oligospora. Trap formation was delayed; no nematodes were captured at nematode induction for 12 h in the Δglo3 mutants; and captured nematodes were significantly reduced at 24, 36, and 48 h compared with the WT strain [32]. Therefore, G-proteins and small GTPases play a pleiotropic role in the growth, development, trap formation, and pathogenicity in A. oligospora and other NT fungi.

3.2. Mitogen-Activated Protein Kinase (MAPK) Signaling Pathway Is Essential for Trap Formation

Accumulating evidences implicate that G-proteins can mediate signal transfer to MAPK-signaling cascade in filamentous fungi, which plays a significant role in pathogenicity [25]. Many NT fungi require very specific abiotic and biotic stimuli to form traps, the ability to sense and respond to environmental signals is essential during the trap formation and nematode predation [33]. MAPK signaling cascades are critical for pathogenic fungi to detect surrounding organisms in the environment [4][34]. There are three major MAPK cascades that have been well-studied in yeasts and filamentous fungi, including SLT2/MPK1 (cell wall integrity pathway), FUS3/KSS1 (pheromone-response and filamentous growth pathway), and HOG1 (hyperosmolarity pathway)[35][36]. Recent research showed that slt2 was required for trap formation in two nematode-trapping fungi A. oligospora (strain ATCC24927) and M. haptotylum [34]. However, another study demonstrated that slt2 is not essential for trap development in A. oligospora (strain TWF154), and its mutant was still able to develop traps after three days of nematode exposure [4]. Meanwhile, BCK1 and MKK1, two proteins of function upstream of SLT2, were mutants unable to produce spores and mycelial traps [37]. Moreover, the trap formation and predation efficiency were reduced in Δhog1 and Δmsb2 mutants, and those mutants were highly sensitive to high osmolarity [38]. Moreover, SSK1, as an upstream regulatory protein of HOG1 signaling pathway, played a negative regulatory role in trap formation, as manifested by significantly increased trap formation and predation efficiency in Δssk1 mutants [39]. In addition, deletion of ste7 and fus3 led to complete abolishment of conidiation and traps in mutants. In addition, STE12, a conserved transcription factor acting downstream of the pheromone-response pathway, induced disruption to defects in the response to nematode pheromone in A. oligospora [4]. Similarly, in Drechslerella dactyloides, the Ddaste12 deletion strain cannot form inflationary contraction ring, which make it unable to catch nematodes [40]. In addition, inducer of meiosis 2 (IME2), a non-classical MAPK-pathway molecule, was associated with mycelial growth and development, conidiation, osmolarity, and pathogenicity in A. oligospora. The Δime2 mutant cannot form mature traps containing multi-hyphal loop, and the electron-dense bodies in trap cells were less than the WT strain[41]. Together, these observations demonstrate that MAPK cascades are essential for trap formation of NT fungi.

3.3. cAMP-Dependent Protein Kinase A (cAMP/PKA) Signaling Pathway Is Indispensable for Trap Formation

The pathogenic process is related to infection-related formation and development in many pathogenic fungi, and the cAMP/PKA pathway plays an essential role in fungal pathogenesis [25]. In our latest research, the deletion of adenylate cyclase led to the abolishment of trap formation, and the number of traps in PKA subunits mutants were reduced in varying degrees (unpublished). In addition, the stunted protein STUA functions as the downstream of cAMP/PKA signaling pathway. The sporulation capacity of ΔstuA mutants were reduced 96%, and the ability to produce mycelial traps was lost [42]. Meanwhile, the transcriptional levels of several upstream genes of cAMP/PKA pathway were significantly reduced in ΔstuA mutants, such as gpr, gα, ras2, acyA, pkaC1, and pkaC2, and some downstream genes were also significantly downregulated, including nsdc, cdc28, and cdc6 [42]. Similarly, the cAMP levels in Δras2 and Δrheb mutants were significantly lower than in the WT strain, and the downstream genes of the cAMP/PKA were significantly downregulated [29]. The cAMP level of the Δric8 mutant was reduced to 3.0%–11.3% compared to the WT strain [27]. These findings indicate that the cAMP signaling pathway is indispensable for trap formation in A. oligospora and other NT fungi.

3.4. Ca2+-Related Signaling Pathway Regulates Trap Formation

Multifunctional Ca2+/calmodulin-dependent protein kinases (CaMKs) are necessary elements in the G-protein signaling pathway. Five CaMKs were identified in A. oligospora, the number of traps in the five CaMK-encoding genes deletion strains were significantly lower than that of the WT strain, and the trap formation was delayed in ΔcamkB mutant [43]. In addition, phospholipase C (PLC), a key enzyme in the inositol phospholipid signaling pathway, hydrolyzes phospholipids to produce inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (DAG) [44]. IP3 causes the release of intracellular calcium ions (Ca2+), and DAG triggers protein kinase C activation [45]. Recent research showed that the number of traps in Δplc2 mutants was reduced and the partial hyphal loop of the traps was irregular [46]. In addition, the low-affinity calcium uptake system (LACS) also played vital role in trap formation, and the deletion of two genes for the LACS transmembrane protein resulted in a 90% trap reduction and no trap formation, respectively [47].

4.Compounds as Signal Molecules to Regulate Trap Formation

There is an evolutionary arms race between predators and prey, and just as prey evolves specific strategies to avoid being hunted, predators also evolve stronger predation strategies, such as lure compounds, to ensure adequate food [48]. Normally, NT fungi are saprophytic, but they will become predators in order to maximize the chance of survival when nutrients are deficient, and their lifestyle transitions accordingly. However, NT fungi are non-motile predators, but nematodes can move at will. Hence, NT fungi evolved an ingenious way, “VOCs”, functioning as mimic pheromone, to lure nematodes. These volatile compounds (VOCs) include dimethyl disulfide, (±)2-methyl-1-butanol, 2,4-dithiapentane, S-methyl thioacetate, and methyl 3-methyl-2-butenoate, and especially, methyl 3-methyl-2-butenoate trigger strong sex- and stage-specific attraction in several Caenorhabditis species. Correspondingly, the olfactory neuron AWCs of C. elegans sensed the odors emanating from NT fungi and responded the attraction [49][7]. Interesting is that the ascarosides, an evolutionarily highly conserved family of small molecules produced by nematodes, downregulated the expression of polyketide synthase gene (artA), which in turn promoted the formation of new traps and then resulted in trapping networks. Therefore, the concentration of ascarosides increased as the nematodes approach, which in turn downregulated arthrosporol and 6-methyl-salicylic acid (6-MSA) formation, further causing the hunting ground to be covered with traps. The ascarosides disappear when the nematodes were completely digested, the contents of arthrosporol and 6-MSA returned to normal levels, and the mycelium switched to saprotrophic growth[50].

Trap formation is a highly energy-consuming process, and to conserve energy, NT fungi have to evolve a more effective strategy to regulate the triggering or closing of trap formation [8]: for example, triggering trap formation after sensing signals of nematodes during infection and terminating trap formation when nematodes were fully digested. Small molecular compounds played an important role in this conversion process, such as ascarosides from nematodes [8], and 6-MSA, oligosporons, arthrobotrisins, and arthrosporols isolated from A. oligospora and other NT fungi [50][51][52][53][54][55]. Recently, an increasing number of compounds and associated synthetic genes have been investigated. For instance, the latest research demonstrated that the chemical diversity of metabolites increased notably and exhibited species specificity in the process of changing lifestyle from saprophytic to predatory in NT fungi A. oligospora, A. thaumasia, and A. musiformis [56]. Volatile furanone and pyrone metabolites can help A. oligospora capture nematodes in the lifestyle transition[12], and abscisic acid was highly effective at enhancing trap formation of D. stenobrocha [57]. In addition, 6-MSA is a chemoattractant that can lure nematodes into the fungal mycelium. The artA expression can produce 6-MSA in hyphal tips, and was uncoupled from other enzymes required for the conversion of 6-MSA to arthrosporols; moreover, corresponding deletion strains produced more traps, suggesting a negative role of 6-MSA on trap formation in D. flagrans [50]. Furthermore, the gene cluster AOL_s00215 plays a key role in the production of arthrosporols in A. oligospora, and the number of traps was increased in deletion mutants of most genes in this gene cluster [10][11][58][59][60]. Arthrobotrisins were downregulated in Δric8, Δras2, and Δrheb mutants, indicating that G-proteins and small GTPases were involved in regulating the metabolism of arthrobotrisins [27][29]. Simultaneously, ammonia could function as a signaling molecule in NT fungi to trigger trap formation and kill nematodes, disrupting the gene involved in urea transport and metabolism, resulting in the abolition of urea-induced trap formation in A. oligospora [61]. Another study also demonstrated that ammonia can induce trap formation as a signal molecule in NT fungi A. oligospora, A. guizhouensis, D. phymatopaga, D. cionopaga, and D. brochopaga [62]. Furthermore, PKS−TPS hybrid pathway, for biosynthesis of sesquiterpenyl epoxy-cyclohexenoids, involved in trap formation via ammonia metabolism, deletion of most genes in the PKS−TPS hybrid pathway displayed significantly increase in trap formation [9]. Overall, the discovery of multiple compounds enriches our knowledge of the inducers in trap formation, which participate in trap formation as signaling molecules.

5. Multiple Cellular Processes Were Involved in Trap Formation

Trap formation of NT fungi was a sophisticated process and required the coordination and cooperation of diverse cellular processes, such as ubiquitin system [63][64], nitrate assimilation pathway [14], pH-sensing receptor [21], the velvet family proteins [65], scaffold proteins[66], lectins [67], actin [68], the striatin-interacting phosphatase and kinase (STRIPAK) [69], adhesin [70], reactive oxygen species [71], glycerol biosynthesis [72], milRNAs [73], woronin body [15], autophagy [17][18][19][20], and cell-to-cell communication and hyphal fusion [13]. Trap formation was affected to varying degrees by the deletion of genes associated with these cellular processes, such as reduction in number, morphological variation, and time delay in formation (Table 1). Interestingly, the organismic interaction between NT fungi and nematodes was just like a dramatic game of hunt or attack. Effectors played critical roles in regulating host cell physiology to promote virulence, biotrophic growth, or symbiosis [74][75]. In NT fungi, certain secreted proteins can be used to modulate the innate immune system of nematodes or target other intracellular processes. For instance, PEFB, a putative fungal virulence factor, was upregulated during nematodes infection and expressed in C. elegans, where it was localized to nuclei [13]. In addition, cell-to-cell communication was required for ring closure, the deletion of sofT inhibited the anastomosis of normal vegetative hyphae, resulting in spiral hyphae, while the mutant was still able to trap C. elegans [13]. The STRIPAK complex is a highly conserved signaling hub involved in the regulation of hyphal fusion. Deletion of the STRIPAK component SIPC resulted in failure to form complete loops and the formation of column-like trap structures with elongated compartments [69]. On the other hand, nitrogen plays a vital role in the growth of fungi, and autophagy was required for nitrogen homeostasis and recycling [19]. Deletion of atg8 not only abolished the autophagy induced by nematodes but also suppressed trap formation and reduces pathogenicity of A. oligospora [19]. Likewise, the formation of autophagosomes and traps were defective in Δatg1, Δatg4, and Δatg5 mutants [17][18][20].
Table 2. A list of characterized genes contributing significantly to trap formation in NT fungi.
Fungi Mutated
Genes		 Annotation Phenotypic Traits Reference
Traps Conidiation Mycelial Growth
A. oligospora gpb1 G-protein β subunit Y N N [24]
A. oligospora flbA Regulator of G-protein signaling

 Y Y Y [26]
A. oligospora rgsA Y N N [26]
A. oligospora rgsB Y Y Y [26]
A. oligospora rgsB2-1 Y Y Y [26]
A. oligospora rgsB2-2 Y N N [26]
A. oligospora rgsB2-3 Y Y N [26]
A. oligospora rgsC Y Y N [26]
A. oligospora gas1 GAS protein Y Y N [26]
A. oligospora ras2 RAS GTPase Y Y Y [29]
A. oligospora ras3 RAS GTPase N N N [29]
A. oligospora rheb RAS GTPase Y Y Y [29]
A. oligospora rab-7A RAB GTPase Y Y Y [28]
A. oligospora rab-2 RAB GTPase N Y N [28]
A. oligospora rho2 RHO GTPase N N N [30]
A. oligospora rac RHO GTPase Y Y Y [30]
A. oligospora cdc42 RHO GTPase Y Y Y [30]
A. oligospora pex1 Peroxisome biogenesis protein Y Y Y [16]
A. oligospora pex6 Peroxisome biogenesis protein Y Y Y [16]
A. oligospora mkk1 MAPK kinase MKK1 Y Y Y [37]
A. oligospora ste7 MAPK kinase STE7 Y Y Y [4]
A. oligospora fus3 MAPK FUS3 Y Y Y [4]
A. oligospora ste12 Tanscription factor Y N Y [4]
A. oligospora slt2 MAPK SLT2 Y Y Y [34]
A. oligospora hog1 MAPK HOG1 Y Y N [38]
A. oligospora msb2 Mucin protein Y N Y [38]
A. oligospora ime2 MAPK IME2 Y Y Y [41]
A. oligospora bck1 MAPK kinase kinase BCK1 Y Y Y [37]
A. oligospora ric8 Resistance to inhibitors of cholinesterase Y Y Y [27]
A. oligospora stuA Transcription factor Y Y Y [42]
A. oligospora glo3 ARF GTPase activator Y Y Y [32]
A. oligospora camk Ca2+/calmodulin-dependent protein kinases Y Y Y [43]
A. oligospora ssk1 Response regulator Y Y Y [39]
A. oligospora
A. oligospora		 atg1
atg13		 Autophagy protein
Autophagy protein		 Y
N		 Y
N		 Y
Y		 [20]
[20]
A. oligospora atg4 Autophagy protein Y Y Y [18]
A. oligospora atg5 Autophagy protein Y Y Y [17]
A. oligospora atg8 Autophagy protein Y Y Y [19]
A. oligospora hex1 Woronin body major protein Y Y Y [15]
A. oligospora gph1 Glycogen phosphorylase Y Y Y [72]
A. oligospora noxA NADPH oxidase Y Y Y [71]
A. oligospora niaD Nitrate reductase Y - Y [14]
A. oligospora niiA Nitrite reductase Y - Y [14]
A. oligospora nrtB Nitrate transporter Y - Y [14]
A. oligospora nirA nitrogen assimilation transcription factor Y - Y [14]
A. oligospora mad1 Adhesin protein Y - - [70]
A. oligospora crn1 Actin cytoskeleton and actin-associated protein Y Y N [68]
A. oligospora palH pH sensing receptor Y Y Y [21]
A. oligospora fig1 Low-affinity calcium system member Y Y Y [47]
A. oligospora ubr1 E3 ubiquitin-protein ligase Y - Y [63]
A. oligospora vosA Developmental regulator N Y N [65]
A. oligospora velB Developmental regulator Y Y Y [65]
A. oligospora g276 Fucose-specific lectin Y N N [67]
A. oligospora g207 F-box protein Y Y Y [64]
A. oligospora AOL_s00215g277 A putatively cupin-like family gene Y Y N [10]
A. oligospora AOL_s00215g278 Cytochrome P450 Y Y Y [60]
A. oligospora AOL_s00215g279 Oxidoreductase Y Y Y [10]
A. oligospora AOL_s00215g280 Cytochrome P450 Y Y Y [58]
A. oligospora AOL_s00215g281 Amidohydrolase Y N Y [11]
A. oligospora AOL_s00215g282 Cytochrome P450 oxidoreductase Y Y Y [11]
A. oligospora AOL_s00215g283 6-methylsalicylic acid synthase Y - N [59]
A. oligospora AOL_s00079g496 Polyketide synthase Y Y Y [12]
D. flagrans artA Polyketide synthase Y - N [50]
D. flagrans artB Cytochrome P450 Y - N [50]
D. flagrans artC Amidohydrolase N - N [50]
D. flagrans artD Cytochrome P450 Y - N [50]
D. flagrans sofT Hyphal anastomosis gene Y - Y [13]
D. flagrans sipC STRIPAK complex component Y Y Y [69]
D. dactyloides ste12 Transcription factor Y Y Y [40]
M. haptotylum slt2 MAPK SLT2 Y Y Y [34]
Y: Affect the corresponding phenotype; N: No effect on corresponding phenotype; -: Not mentioned.

This entry is adapted from the peer-reviewed paper 10.3390/jof8040406

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