Zoopagomycota

Zoopagomycota is an early-diverging phylum of terrestrial, non-flagellated fungi comprising approximately 900 to 1,000 described species that forms the earliest-branching lineage of the zygosporic (non-zoosporic) terrestrial fungi, diverging prior to the split between Mucoromycota and Dikarya (Ascomycota + Basidiomycota). Whereas Mucoromycota and Dikarya radiated primarily as plant-associated saprotrophs, mycorrhizal mutualists, and plant pathogens, Zoopagomycota is an overwhelmingly animal- and fungus-associated phylum.

Established by Joseph Spatafora, Jason Stajich, and colleagues in 2016 after phylogenomic resolution split the paraphyletic traditional phylum "Zygomycota," Zoopagomycota unites three subphyla that exhibit some of the most specialized predatory and neuro-manipulative lifestyles in the fungal kingdom:

1. Subphylum Zoopagomycotina (Order Zoopagales; ~200 spp.): Obligate predaceous fungi (Stylopage, Zoophagus, Acaulopage, Cochlonema) and mycoparasites (Piptocephalis, Syncephalis) that glue onto and digest living soil amoebae, testate rhizopods, nematodes, and rotifers.
2. Subphylum Kickxellomycotina (~350 spp., including the "Trichomycetes" orders Harpellales and Asellariales): Septate fungi bearing lens-shaped septal plugs that live as obligate gut-lining commensals attached to the chitinous hindgut cuticle of aquatic insect larvae (Ephemeroptera mayflies, Plecoptera stoneflies, and blackfly/midge Diptera).
3. Subphylum Entomophthoromycotina (Order Entomophthorales; ~300 spp. — Entomophthora muscae, Entomophaga maimaiga, Massospora cicadina, Pandora neoaphidis, Conidiobolus): Obligate insect pathogens ("fly destroyers") that replicate as wall-less protoplasts in the insect hemocoel, chemically hijack the host insect's central nervous system to trigger "summit disease" or hyper-sexual active transmission ("flying salt shakers of death"), and fire ballistic primary conidia at accelerations of >30,000 g.

More than 350 million years before nematode-trapping Ascomycota (Arthrobotrys oligospora) evolved constricting rings in the Mesozoic, Zoopagales evolved microscopic hunting traps for soil and freshwater microfauna:

• Amoeba and Nematode Adhesive Traps (Stylopage and Acaulopage): Hyphae of Stylopage hadra and Stylopage anomala snake across soil crumbs and herbivore dung, coated in golden droplets of a high-viscosity mucopolysaccharide adhesive that solidifies instantaneously ONLY upon contact with the glycocalyx of a passing naked amoeba (Amoeba, Thecamoeba) or rhabditid nematode. Once the prey is glued fast, the fungus extends a fine penetration peg through the cuticle/plasma membrane that branches inside the living victim into a nutrient-absorbing haustorial bush, fully draining the cytoplasm within hours.
• Zoophagus insidians — The Oral Lure Trap for Aquatic Rotifers: Growing on filamentous green algae in freshwater ponds, Zoophagus insidians erects short, perpendicular lateral peg-like branches spaced evenly along its main hypha. When a loricate or bdelloid rotifer (Lecane, Monostyla) mistakes the short hyphal peg for a filamentous algal food item and grasps the peg with its pharyngeal jaws (mastax / trophi), mechanical contact triggers immediate exocytosis of a fast-curing proteinaceous glue from vesicles stored in the peg tip—cementing the rotifer's mouth permanently onto the fungal peg while assimilative hyphae grow out of the peg down the rotifer's esophagus to digest the animal from the inside out!

Unlike hypocrealean zombie-ant fungi (Ophiocordyceps unilateralis in Ascomycota), which have compact ~22–30 Mb genomes, members of the order Entomophthorales (Entomophthora muscae, Entomophaga maimaiga, Massospora cicadina) possess the largest genomes in the fungal kingdom—ranging from 650 Mb to 1,500 Mb (1.5 Gb), expanded more than 50-fold by Ty3/gypsy and DNA transposons!

1. Immune Evasion as Wall-Less "Stealth Protoplasts"

When a ballistic conidium of Entomophthora muscae or Entomophaga maimaiga lands on the cuticle of a dipteran fly or lepidopteran caterpillar, it germinates a germ tube that slices through the chitinous exoskeleton via lipases, chitinases, and subtilisin-like proteases. The moment the hypha enters the insect's nutrient-rich hemocoel, it fully sheds its β-glucan/chitin cell wall and proliferates as motile, pleomorphic, wall-less protoplasts! Because insect innate immunity relies on β-1,3-glucan recognition proteins (βGRPs) and peptidoglycan recognition proteins to trigger hemocyte encapsulation and the prophenoloxidase (melanization) cascade, wall-less Entomophthora protoplasts multiply invisibly in the hemolymph while consuming the host's fat body while keeping its vital heart, flight muscles, and brain alive until the final hours.

2. "Summit Disease" in Entomophthora muscae: Circadian Clock and Neurosecretory Hijacking

As Entomophthora muscae protoplasts saturate the hemolymph of an infected housefly (Musca domestica) or fruit fly (Drosophila melanogaster), a dedicated subpopulation of fungal cells invades the fly's central brain and gnathal ganglion, specifically infiltrating the pars intercerebralis and surrounding clock neurons (Elya et al., 2018, 2023):

1. Circadian-Gated Climb (4–5 Hours Before Sunset): Specifically gated to late afternoon, the fungus triggers a burst of locomotor hyperactivity and negative gravitaxis, compelling the dying fly to climb to the highest available perch (summit disease).
2. Proboscis-Glueing and Wing-Raising Death Posture: Once at the summit, fungal invasion of the gnathal ganglion forces the fly to extend its proboscis onto the substrate, where fungal hyphae burst out of the labellum to glue the fly's mouthparts permanently to the surface. Minutes later, the fly's wings snap straight upward at 90° over its thorax (clearing an unobstructed firing arc for the abdomen!) and the fly dies.
3. Volatile Sesquiterpene Sex-Pheromone Lure of Healthy Males: Within 2 to 4 hours of death, thousands of white conidiophores erupt through the intersegmental membranes of the dead female fly's swollen abdomen. Simultaneously, the fungus synthesizes a volatile cocktail of sesquiterpenes and cuticular methyl-branched alkanes (Naundrup et al., 2022) that chemically amplify the female fly's sex pheromone, luring healthy male flies to copulate with the spore-covered corpse ("necrophilic attraction") while the fungus bombards them with sticky ballistic conidia!

3. Massospora cicadina: Amphetamine (Cathinone) and Psilocybin Dosing of Living Cicadas

Even more astonishing is the genus Massospora (Massospora cicadina on 13- and 17-year periodical cicadas [Magicicada in Hemiptera]; Massospora platypediae and M. levispora on annual cicadas), which skips killing its host altogether and instead turns the living, flying insect into an active venereal vector ("flying salt shakers of death"):

• As adult cicadas emerge from the soil, Massospora hyphae fully replace the posterior half of the cicada's abdomen—destroying its genitalia and sloughing off the rear abdominal tergites to expose a chalky white plug of infectious conidia.
• Despite having lost half its body, the infected cicada does not display sickness lethargy; instead, it flies vigorously and engages in hyper-sexual mating behaviors—with infected males even mimicking the wing-flicking receptivity signals of females to trick healthy males into attempting copulation with their spore-filled abdomens!
• Metabolomic analysis by Matt Kasson, Boyce, and colleagues (2019) discovered how Massospora achieves this behavioral manipulation:
  • Massospora cicadina synthesizes the plant-associated amphetamine alkaloid cathinone (the active central nervous system stimulant of the khat shrub Catha edulis!) directly inside the periodical cicada to suppress fatigue and sustain hyperactive flight.
  • Annual-cicada pathogens Massospora platypediae and M. levispora synthesize the tryptamine hallucinogen psilocybin (previously known only from agaricoid Psilocybe mushrooms in Basidiomycota) alongside erinacine-like terpenoids to alter monoaminergic neurotransmission in the cicada brain!

1. Spatafora, J. W., Chang, Y., Benny, G. L., Lazarus, K., Smith, M. E., Berbee, M. L., ... & Stajich, J. E. (2016). A phylum-level phylogenetic classification of zygomycete fungi based on genome-scale data. Mycologia, 108(5), 1028–1046. Google Scholar
2. Elya, C., Lok, T. C., Spencer, Q. E., McCausland, H., Martinez, C. C., & Eisen, M. (2018). Robust manipulation of the behavior of Drosophila melanogaster by a fungal pathogen in the laboratory. eLife, 7, e34414. Google Scholar
3. Boyce, G. R., Gluck-Thaler, E., Slot, J. C., Stajich, J. E., Davis, W. J., James, T. Y., ... & Kasson, M. T. (2019). Psychoactive plant- and mushroom-associated alkaloids from two behavior modifyingcicada pathogens. Fungal Ecology, 41, 147–164. Google Scholar
4. Naundrup, A., Bohman, B., Kwadha, C. A., Jensen, A. B., Becher, P. G., & De Fine Licht, H. H. (2022). Pathogenic fungus uses volatiles to entice male flies into fatal matings with infected female cadavers. ISME Journal, 16(10), 2388–2397. Google Scholar

• Fungi
• Mucoromycota
• Hemiptera
• Ephemeroptera
• Rotifera