Information

3.4: Glomeromycota (Endomycorrhizal Fungi) - Biology

3.4: Glomeromycota (Endomycorrhizal Fungi) - Biology


We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

Glomeromycota represent the endomycorrhizal fungi. Thus, these images contain primarily plant roots. They are a mysterious bunch!

Figure (PageIndex{1}): Glomus coremioides forming sporocarps, an unusual characteristic in this fungal phylum. The white to tan ones are still forming. Spores are produced within these structures. The ecology of this species is a bit mysterious. Photos by Damon Tighe, some rights reserved (CC-BY-NC).

Figure (PageIndex{2}): Endomycorrhizal fungi can form arbuscules (branching structures) and vesicles (globose structures). In this micrograph, a root has been stained and viewed at high magnification. The hyphal structures of the fungus can be seen within the root cells, particularly the vesicles, which stain darker. Msturmel, Public domain, via Wikimedia Commons.

Figure (PageIndex{3}): A root of a mycoheterotrophic plant, Corallorhiza. Arbuscules can be seen in many cells of the cortex of the plant root. Three of these arbuscules have been indicated with a note that says "cells filled with branching fungal hyphae". Photo by Melissa Ha, CC-BY-NC.


Endomycorrhiza

Chilekampalli A. Reddy , Ramu S. Saravanan , in Advances in Applied Microbiology , 2013

4.1 Arbuscular Mycorrhizae

Endomycorrhizas include arbuscular (AMs), ericoid, and orchid mycorrhizas. Arbuscular mycorrhizal fungi are soil fungi belonging to the phylum Glomeromycota ( Cairney, 2000 Castillo & Pawlowska, 2010 Sturmer, 2012 ). AM fungal association with plants dates back to 460 million years, predating the Rhizobium-legume root nodule symbiosis by >300 million years ( Cairney, 2000 Parniske, 2008 Selosse & Rousset, 2011 Shtark et al., 2011 Wilkinson, 2001 ). AM fungi are obligate symbionts and have limited saprobic ability. They are dependent on the plant for their carbon nutrition. AM fungi are the most commonest mycorrhizal group associated with plants and are found in association with angiosperms, gymnosperms, pteridophytes, and bryophytes ( Dermatsev et al., 2010 Hoffman & Arnold, 2010 Sturmer, 2012 ). AM fungi are able to develop a symbiotic association with most terrestrial plants throughout the world and play an important role in providing the plant with P, S, N, and various micronutrients from the soil. AM fungi mobilize N and P from organic polymers, release mineral nutrients from insoluble particulate matter, and mediate plant responses to stress factors and resistance to plant pathogens. Mycorrhiza also participate in a number of beneficial interactions with various groups of soil microorganisms ( Bianciotto & Bonfante, 2002 Krishna, 2005 Peterson, 2004 Tikhonovich & Provorov, 2007 Veresoglou, Menexes, & Rillig, 2012 ). Plants and their AM fungal microsymbionts interact in complex underground networks involving multiple partners ( Barea, Azcón, & Azcón-Aguilar, 2005 Bending et al., 2006 Rosendahl, 2008 ). How these partners maintain a fair, two-way transfer of resources between themselves was not well understood. Recently, however, Kiers et al. (2011) were able to show that plants can detect, discriminate, and reward the best fungal partners with more carbohydrates, while the fungal partners, in turn, enforce cooperation by increasing the nutrient transfer only to those host plant roots that provide more carbohydrates. Such mutualism between partners is evolutionarily stable because control is bidirectional and partners offering the best rate of exchange are rewarded.

The hyphae of AM fungi penetrate the plant root cells to establish an intracellular symbiosis, irrespective of the plant host. AM fungal colonization of plant root cells is complex. AM fungal spores germinate to form hyphopodia (or appressoria) at the root surface, followed by inter- and intracellular penetration by hyphae and the formation of characteristic branched intracellular structures called “arbuscules” ( Fig. 3.2 , panels c, d, and e) and produce bladder-like vesicles (hyphal swellings in the root cortex that contain lipids and cytoplasm) inside cortical cells. Arbuscules, which give their name to the symbiosis, are considered the main site of exchange of C, P, water, and other nutrients between the symbiotic partners ( Bonfante & Requena, 2011 Lee, Muneer, Avice, Jung, & Kim, 2012 Miransari, 2011 Parniske, 2008 Rillig & Mummey, 2006 ). The transfer of carbon from the plant to the fungi may also occur through the intraradical hyphae. After colonization, AM fungi produce runner hyphae (extraradicular hyphae) that grow from the plant root into the soil and take up P and micronutrients, which are transferred to the plant. AM fungal hyphae, which grow from the plant root, have a high surface-to-volume ratio, making their absorptive ability greater than that of plant roots ( Azcon, 2009 Bonfante & Anca, 2009 ). Also, these hyphae are finer than roots and can enter into pores of the soil that are inaccessible to roots. The size and the relative proportion of hyphae within the root and in the soil vary greatly between different AM species ( Abdel Latef, 2011 Dumas-Gaudot et al., 2004 Lumini, Orgiazzi, Borriello, Bonfante, & Bianciotto, 2010 ). Furthermore, AM fungi produce another morphologically distinct type of hyphae that grow from the roots and colonize other host plants in the proximity.

Figure 3.2 . Ectomycorrhizal and endomycorrhizal colonization of Acacia nilotica seedlings ( Saravanan, 1998 Saravanan &amp Natrajan, 2000 ). a) Morphology of a root colonized by the ectomycorrhizal fungus, Pisolithus tinctorius. b) Cross-section of a root colonized by P. tinctorius. Thick mycelial covering (mantle) on the surface of the root (yellow arrows) and intercellular hyphal network (“Hartig net”, shown by red arrows). 1. epidermal cells 2. cortical cells. c) Root surface showing germinated spores of Glomus mosseae (an AM fungus) d) Stained arbuscules of G. mosseae in the root (arrows) e) Stained vesicles of G. mosseae in the root (arrows). (For interpretation of the references to color in this figure legend, the reader is referred to the online version of this book.)

The magnitude of bacteria/mycorrhizal fungus/plant symbioses depends on a number of variables such as the extent of fungal colonization, C-transfer from the plant, the amount of P in the soil, and the relative amount of P transferred to the plant ( Lee et al., 2012 Sanon et al., 2012 Veresoglou et al., 2012 Zamioudis & Pieterse, 2012 ). The diversity of AM fungi in an ecosystem influences the diversity and productivity of plants in that environment ( Adesemoye & Kloepper, 2009 Andreas & Martin, 2006 Angeles, Ouyang, Aguirre, Lammers, & Song, 2009 Bago et al., 2003 Finlay, 2008 ). Symbiotic N-fixation between rhizobia (including Sinorhizobium, Bradyrhizobium, Mesorhizobium, and Azorhizobium) and a host plant involves signal molecules and pathways. This symbiosis may be favorably influenced by simultaneous AM fungal colonization with the same host plant ( Bonfante & Anca, 2009 Bonfante & Requena, 2011 Harrison, 2005 ).

Signal molecules and pathways of AM fungi and the host plant for the onset of symbiosis and the beneficial mutualism between the two have been reviewed ( Akiyama & Hayashi, 2006 Bonfante & Requena, 2011 Harrison, 2005 Kiers et al., 2011 ). Signals are exchanged between plant roots and AM fungi that are important for plant mycorrhizal symbiosis. Recent evidence suggests that phytohormones produced by the plant including strigolactones stimulate fungal metabolism as well as hyphal branching ( Bonfante & Requena, 2011 Castillo & Pawlowska, 2010 Dodd & Ruiz-Lozano, 2012 ). The symbiotic signals (“Myc factors”) are active in small doses and stimulate the growth of the root system, and the formation of mycorrhiza association. Biotechnological processes for synthesizing large amounts of these regulator molecules have been developed and will permit the study of their effects on crops, with the objective of improving their yield, without using chemical fertilizers ( Bonfante & Requena, 2011 Harrison, 2005 ). There are plant genes that are expressed by “Myc factors” at the onset of AM-host plant symbiosis and the fungal genes involved in structural and physiological alterations in the host plant have also been identified ( Dodd & Ruiz-Lozano, 2012 Hata et al., 2010 ). The physiological functions of the host plant need to be modified so that the host plant can provide the fungi with the required organic carbon compounds (through root exudate) in exchange for water and nutrients provided by the fungus. Some common plant genes are expressed during the AM symbiosis as well as N-fixation by rhizobium ( Akiyama & Hayashi, 2006 ).


Arbuscular Mycorrhizae

Endomycorrhizae have several functions, the major one being nutrient acquisition. Endomycorrhizae facilitate the exchange of nutrients between the host plant and the soil. Mycorrhizae aid in the uptake of water, inorganic phosphorus, mineral or organic nitrogen, and amino acids. In exchange for the mycorrhizae providing all of these nutrients, the plant in turn provides the mycorrhizae with carbon (1). This relationship benefits both organisms immensely. The mycorrhizae greatly increase the surface area of the plant’s root system which is hugely beneficial in areas where drought is common. This is also beneficial in areas where the soil is nutrient-poor. The larger surface area gives those plants an advantage over plants lacking this symbiotic relationship allowing plants with mycorrhizal relationships to out-compete for nutrients. Mycorrhizae also offer the roots of the plant a little more protection (3).


24.3 Ecology of Fungi

By the end of this section, you will be able to do the following:

  • Describe the role of fungi in various ecosystems
  • Describe mutualistic relationships of fungi with plant roots and photosynthetic organisms
  • Describe the beneficial relationship between some fungi and insects

Fungi play a crucial role in the constantly changing “balance” of ecosystems. They colonize most habitats on Earth, preferring dark, moist conditions. They can thrive in seemingly hostile environments, such as the tundra, thanks to a most successful symbiosis with photosynthetic organisms like algae to produce lichens. Within their communities, fungi are not as obvious as are large animals or tall treas. Like bacteria, they act behind the scene as major decomposers. With their versatile metabolism, fungi break down organic matter, which would otherwise not be recycled.

Habitats

Although fungi are primarily associated with humid and cool environments that provide a supply of organic matter, they colonize a surprising diversity of habitats, from seawater to human skin and mucous membranes. Chytrids are found primarily in aquatic environments. Other fungi, such as Coccidioides immitis, which causes pneumonia when its spores are inhaled, thrive in the dry and sandy soil of the southwestern United States. Fungi that parasitize coral reefs live in the ocean. However, most members of the Kingdom Fungi grow on the forest floor, where the dark and damp environment is rich in decaying debris from plants and animals. In these environments, fungi play a major role as decomposers and recyclers, making it possible for members of the other kingdoms to be supplied with nutrients and live.

Decomposers and Recyclers

The food web would be incomplete without organisms that decompose organic matter (Figure 24.19). Some elements—such as nitrogen and phosphorus—are required in large quantities by biological systems, and yet are not abundant in the environment. The action of fungi releases these elements from decaying matter, making them available to other living organisms. Trace elements present in low amounts in many habitats are essential for growth, and would remain tied up in rotting organic matter if fungi and bacteria did not return them to the environment via their metabolic activity.

The ability of fungi to degrade many large and insoluble molecules is due to their mode of nutrition. As seen earlier, digestion precedes ingestion. Fungi produce a variety of exoenzymes to digest nutrients. The enzymes are either released into the substrate or remain bound to the outside of the fungal cell wall. Large molecules are broken down into small molecules, which are transported into the cell by a system of protein carriers embedded in the cell membrane. Because the movement of small molecules and enzymes is dependent on the presence of water, active growth depends on a relatively high percentage of moisture in the environment.

As saprobes, fungi help maintain a sustainable ecosystem for the animals and plants that share the same habitat. In addition to replenishing the environment with nutrients, fungi interact directly with other organisms in beneficial, and sometimes damaging, ways (Figure 24.20).

Mutualistic Relationships

Symbiosis is the ecological interaction between two organisms that live together. This definition does not describe the type or quality of the interaction. When both members of the association benefit, the symbiotic relationship is called mutualistic. Fungi form mutualistic associations with many types of organisms, including cyanobacteria, algae, plants, and animals.

Fungus/Plant Mutualism

One of the most remarkable associations between fungi and plants is the establishment of mycorrhizae. Mycorrhiza , which is derived from the Greek words myco meaning fungus and rhizo meaning root, refers to the fungal partner of a mutualistic association between vascular plant roots and their symbiotic fungi. Nearly 90 percent of all vascular plant species have mycorrhizal partners. In a mycorrhizal association, the fungal mycelia use their extensive network of hyphae and large surface area in contact with the soil to channel water and minerals from the soil into the plant. In exchange, the plant supplies the products of photosynthesis to fuel the metabolism of the fungus.

There are several basic types of mycorrhizae. Ectomycorrhizae (“outside” mycorrhizae) depend on fungi enveloping the roots in a sheath (called a mantle). Hyphae grow from the mantle into the root and envelope the outer layers of the root cells in a network of hyphae called a Hartig net (Figure 24.21). The fungal partner can belong to the Ascomycota, Basidiomycota or Zygomycota. Endomycorrhizae ("inside" mycorrhizae), also called arbuscular mycorrhizae, are produced when the fungi grow inside the root in a branched structure called an arbuscule (from the Latin for “little trees”). The fungal partners of endomycorrhizal associates all belong to the Glomeromycota. The fungal arbuscules penetrate root cells between the cell wall and the plasma membrane and are the site of the metabolic exchanges between the fungus and the host plant (Figure 24.21b and Figure 24.22b). Orchids rely on a third type of mycorrhiza. Orchids are epiphytes that typically produce very small airborne seeds without much storage to sustain germination and growth. Their seeds will not germinate without a mycorrhizal partner (usually a Basidiomycete). After nutrients in the seed are depleted, fungal symbionts support the growth of the orchid by providing necessary carbohydrates and minerals. Some orchids continue to be mycorrhizal throughout their life cycle.

Visual Connection

If symbiotic fungi were absent from the soil, what impact do you think this would have on plant growth?

Other examples of fungus–plant mutualism include the endophytes: fungi that live inside tissue without damaging the host plant. Endophytes release toxins that repel herbivores, or confer resistance to environmental stress factors, such as infection by microorganisms, drought, or heavy metals in soil.

Evolution Connection

Coevolution of Land Plants and Mycorrhizae

As we have seen, mycorrhizae are the fungal partners of a mutually beneficial symbiotic association that coevolved between roots of vascular plants and fungi. A well-supported theory proposes that fungi were instrumental in the evolution of the root system in plants and contributed to the success of Angiosperms. The bryophytes (mosses and liverworts), which are considered the most ancestral plants and the first to survive and adapt on land, have simple underground rhizoids, rather than a true root system, and therefore cannot survive in dry areas. However, some bryophytes have arbuscular mycorrhizae and some do not.

True roots first appeared in the ancestral vascular plants: Vascular plants that developed a system of thin extensions from their roots would have had a selective advantage over nonvascular plants because they had a greater surface area of contact with the fungal partners than did the rhizoids of mosses and liverworts. The first true roots would have allowed vascular plants to obtain more water and nutrients in the ground.

Fossil records indicate that fungi actually preceded the invasion of ancestral freshwater plants onto dry land. The first association between fungi and photosynthetic organisms on land involved moss-like plants and endophytes. These early associations developed before roots appeared in plants. Slowly, the benefits of the endophyte and rhizoid interactions for both partners led to present-day mycorrhizae: About 90 percent of today’s vascular plants have associations with fungi in their rhizosphere.

The fungi involved in mycorrhizae display many characteristics of ancestral fungi they produce simple spores, show little diversification, do not have a sexual reproductive cycle, and cannot live outside of a mycorrhizal association. The plants benefited from the association because mycorrhizae allowed them to move into new habitats and allowed the increased uptake of nutrients, which gave them an enormous selective advantage over plants that did not establish symbiotic relationships.

Lichens

Lichens display a range of colors and textures (Figure 24.23) and can survive in the most unusual and hostile habitats. They cover rocks, gravestones, tree bark, and the ground in the tundra where plant roots cannot penetrate. Lichens can survive extended periods of drought, when they become completely desiccated, and then rapidly become active once water is available again.

Link to Learning

Explore the world of lichens using this site from Oregon State University.

It is important to note that lichens are not a single organism, but rather another wonderful example of a mutualism, in which a fungus (usually a member of the Ascomycota or Basidiomycota) lives in a physical and physiological relationship with a photosynthetic organism (a eukaryotic alga or a prokaryotic cyanobacterium) (Figure 24.24). Generally, neither the fungus nor the photosynthetic organism can survive alone outside of the symbiotic relationship. The body of a lichen, referred to as a thallus, is formed of hyphae wrapped around the photosynthetic partner. The photosynthetic organism provides carbon and energy in the form of carbohydrates. Some cyanobacteria additionally fix nitrogen from the atmosphere, contributing nitrogenous compounds to the association. In return, the fungus supplies minerals and protection from dryness and excessive light by encasing the algae in its mycelium. The fungus also attaches the lichen to its substrate.

The thallus of lichens grows very slowly, expanding its diameter a few millimeters per year. Both the fungus and the alga participate in the formation of dispersal units, called soredia—clusters of algal cells surrounded by mycelia. Soredia are dispersed by wind and water and form new lichens.

Lichens are extremely sensitive to air pollution, especially to abnormal levels of nitrogenous and sulfurous compounds. The U.S. Forest Service and National Park Service can monitor air quality by measuring the relative abundance and health of the lichen population in an area. Lichens fulfill many ecological roles. Caribou and reindeer eat lichens, and they provide cover for small invertebrates that hide in the mycelium. In the production of textiles, weavers used lichens to dye wool for many centuries until the advent of synthetic dyes. The pigments used in litmus paper are also extracted from lichens.

Link to Learning

Lichens are used to monitor the quality of air. Read more on this site from the United States Forest Service.

Fungus/Animal Mutualism

Fungi have evolved mutualisms with numerous insects in Phylum Arthropoda: joint-legged invertebrates with a chitinous exoskeleton. Arthropods depend on the fungus for protection from predators and pathogens, while the fungus obtains nutrients and a way to disseminate spores into new environments. The association between species of Basidiomycota and scale insects is one example. The fungal mycelium covers and protects the insect colonies. The scale insects foster a flow of nutrients from the parasitized plant to the fungus.

In a second example, leaf-cutter ants of Central and South America literally farm fungi. They cut disks of leaves from plants and pile them up in subterranean gardens (Figure 24.25). Fungi are cultivated in these disk gardens, digesting the cellulose in the leaves that the ants cannot break down. Once smaller sugar molecules are produced and consumed by the fungi, the fungi in turn become a meal for the ants. The insects also patrol their garden, preying on competing fungi. Both ants and fungi benefit from this mutualistic association. The fungus receives a steady supply of leaves and freedom from competition, while the ants feed on the fungi they cultivate.

Fungivores

Animal dispersal is important for some fungi because an animal may carry fungal spores considerable distances from the source. Fungal spores are rarely completely degraded in the gastrointestinal tract of an animal, and many are able to germinate when they are passed in the feces. Some “dung fungi” actually require passage through the digestive system of herbivores to complete their lifecycle. The black truffle—a prized gourmet delicacy—is the fruiting body of an underground ascomycete. Almost all truffles are ectomycorrhizal, and are usually found in close association with trees. Animals eat truffles and disperse the spores. In Italy and France, truffle hunters use female pigs to sniff out truffles (female pigs are attracted to truffles because the fungus releases a volatile compound closely related to a pheromone produced by male pigs.)


3.4: Glomeromycota (Endomycorrhizal Fungi) - Biology

Copyright © 2014 by authors and Scientific Research Publishing Inc.

This work is licensed under the Creative Commons Attribution International License (CC BY).

Received 20 June 2014 revised 10 July 2014 accepted 5 August 2014

Endomycorrhizal fungi play an important role in the survival of plants on poor soils. Planting seeds into lunar soil at a lunar colony will be a challenge for seeds of any plant. The seeds will need a special microbial “tool kit” that will help them germinate and the young seedlings establish themselves. In this study, seeds of the prickly pear cactus, Opuntia ficus-indica, were chosen to examine the presence of fungus spores in the soil, inside the seeds and after germination in the rhizosphere, roots and other tissues of the young seedlings. The nutrient poor lunar regolith simulant JSC-1A was used as autoclaved or untreated growth medium. The mycorrhizal fungus Trichoderma viride was predominantly identified on the roots of new seedlings. This fungus also demonstrated the strongest effect on the germination rate of the seeds in comparison with other fungi isolated from the rhizosphere of Opuntia plants. T. viride was not detected within seeds and also not within seedlings, besides the root tips, whereas an arbuscular mycorrhizal Glomus species was seed-borne and present throughout most of the seedling. A close association between T. viride and a Glomus species associated with O. ficus-indica is demonstrated through light microscopic and electron microscopic images of the outside and inside root tips of the seedlings.

Keywords:Endomycorrhizal Fungi, Seed Germination, Lunar Regolith Stimulant, Nutrient Poor Soil, Opuntia ficus-indica

The prickly pear cactus, Opuntia ficus-indica, is an important crop plant from Mexico that is well suited for hot, semiarid and arid environments and was therefore chosen by our team as a possible starter plant for an extraterrestrial colony. It establishes itself easily and is one of the most productive crop plants known [1] . The lunar regolith simulant JSC-1A is a volcanic, nutrient-poor and dusty soil that simulates the soil retrieved from the moon by NASA and is used to study the suitability of plants for agriculture in a lunar colony. It is also well suited for a study of associated microbes of plants that grow very successfully on nutrient-poor soils and the role that these microbes may play in their success. The symbiotic relationship of fungi with roots in the rhizosphere at the soil-root interface is a well-known phenomenon and assumed to be present in almost all plants, including the cacti of the genus Opuntia [2] . The mycorrhizae are formed by fungal hyphae growing either on the outside of plant roots (ectomycorrhizae) or into the roots (endomycorrhizae). Thus the mycorrhizal fungi increase the root surface and extend the plant’s reach for nutrients. They are known to play a major role in phosphate uptake of the plant and in accessing other nutrients in poor soils as well [3] -[6] . AM fungi reportedly also increase the production of dry matter in seedlings of the cactus Pachycereus pectin-aboriginum [7] . According to Pope (1993) [8] , the growth of endomycorrhizal fungi is promoted by high light intensity and poor soil conditions. Especially on desert soils they may therefore help plants acquire the minerals they need from the soil. The degree of root colonization, however, also varied and showed seasonal site variation on the prickly pear Opuntia humifusa in the study of Whitcomb (2000) [2] . Therefore the author suggests classifying a “plant’s mycorrhizal status based on the ecological condition in which they are found”. Dubrovsky and North (2002) [9] recognize three species of the genus Glomus as inhabitants of the rhizosphere of the Opuntia species they studied. These form arbuscular-mycorrhizal (AM) fungus associations between the zygomycete fungus with coenocytic hyphae and the plant. They are diagnosed by the presence of arbuscules in the cortical root cells and of terminal vesicles that contain oil droplets [8] . The hyphae extend from the infected root into the adjacent soil or may connect root to root. The spores of AM fungi remain in the soil in sporocarps, but may also become windborne. According to Barrow et al. (2008) [10] , endosymbiotic fungi even grow into developing seed embryos and are thus vertically transmitted by seeds, as reported from C3 forage grasses.

The seeds of the prickly pear cacti of the genus Opuntia are known for their strong physiological dormancy and a low growth potential of their embryos, which means that their seeds do not germinate easily, an adaptation to desert environments [11] . The seeds have a very hard endocarp so that the dormant seed embryo cannot break. Delgado-Sanchez et al. (2010 2011) [12] [13] found an effect of Penicillium chrysogenum, a Phoma species, and Trichoderma koningii on breaking germination resistance in Opuntia seeds. Since these fungi were observed growing on the testa of all germinated seeds and eroding the funiculus, the authors interpreted the resistance to germinate as a mechanical problem. This idea goes along with the commonly used methods of breaking Opuntia seed dormancy through scarification methods, which involve treatment with sulfuric acid, cold treatment and/or rubbing with sandpaper [14] .

In this study the effect of different soil fungi, including mycorrhizal fungi, on the germination rate of seeds of the prickly pear cactus, Opuntia ficus-indica, was evaluated, and the presence of endomycorrhizal fungi on and in the seeds and the seedlings growing on lunar regolith simulant JSC-1A was studied with the use of light and scanning electron microscopy (SEM). The importance of these fungi for the seed germination on a nutrient poor soil and the establishment of the seedlings are discussed.

Seeds of Opuntia ficus-indica were collected from mature fruits (tunas) obtained in supermarkets and stored in paper bags at room temperature to allow for a normal aging process to overcome their dormancy. At the time of usage, they were between 2 - 3 years old.

2.1. Determination of Fungi on and Inside Seeds

The presence of fungi on and inside the seeds was studied using untreated and surface-sterilized seeds. Alcohol, often together with hypochloride, is commonly used to sterilize the outer surface of seeds, including seeds of Opuntia species [12] [15] . To surface-sterilize the seeds for planting, they were washed in 70% ethanol for 10 min, rinsed four times in sterile, distilled water and dried before planting. Surface-sterilization of the seeds for fungus growth on agar included a bleach treatment. They were submerged in 70% ethanol for 30 sec, then for 10 min in a solution of 1% hypochlorite prepared of household bleach, followed by four rinses in distilled water. The seeds were then placed whole or crushed on sterile Sabouraud Dextrose Agar (Becton Dickinson) in Petri dishes (SDA plates). The plates were incubated in the dark at room temperature. Samples of the growing colonies were placed on microscopic slides and stained with Lactophenol cotton blue (LPCB) to check for presence of bacteria and fungi, and to identify the fungi microscopically. Bacteria were also Gram stained.

2.2. Mycorrhizal Fungi and Seed Germination

To study the fungus or fungi growing on the roots of seedlings, we designed several planting experiments. The lunar regolith simulant JSC-1A (Orbitec, LLC) was used as a planting soil for the seeds. Preliminary studies in our lab demonstrated that prickly pear cacti grow well on the regolith. First the native germination rate of our O. ficus-indica seeds on JSC-1A was determined. 20 seeds were placed into plain JSC-1A held within a coffee filter in a plastic planting pot and kept moist with distilled water (Table 1, #0). The roots of the freshly sprouted seedlings were swabbed with sterile cotton swabs to prepare streaks on SDA plates. The resulting fungal cultures were later used to study their effect on the germination rate of the O. ficus-indica seeds from the same batch.

In these germination experiments JSC-1A was used either untreated or autoclaved for 30 min at 121˚C. The presence of fungus spores in the untreated and autoclaved soil was determined through sprinkling a small sample onto sterile SDA plates. The SDA plates were kept in the dark at room temperature to promote fungus growth.

For the first series of germination experiments (Table 1) autoclaved soil was used to study the effect of a mycorrhizal fungus on the seed germination rate of O. ficus-indica seeds. The planting pots, coffee filters and plastic bags needed were first placed overnight under UV light in the laboratory hood for sterilization purposes. The planting procedure was performed under application of aseptic techniques. First the coffee filters were placed into the planting pots. Then autoclaved JSC-1A was added into the coffee filter and sterile distilled water added to moisten the soil well. The seeds were then placed onto the moist soil and pushed slightly into it, but care was taken to not cover them. The control pots were transferred at once into plastic Ziploc bags and the bags closed by 3/4 allowing for air exchange, but preventing airborne spores from falling onto the soil. To the other pots samples of the fungal culture, obtained from the root streak of seedlings described above (seedlings from experiment #0, Table 1), representing a green mycrorrhizal fungus (identified as Trichoderma viride), were streaked with a sterile loop onto the soil close to the seeds. For experiment #3 (Table 1) surface-sterilized seeds were used. These planting pots were then too placed into plastic bags and closed as described above. All germination experiments were kept at room temperature on a plant stand with artificial lighting. The humidity in the plastic bags was kept high by pouring sterile distilled water into the bags on demand. Twice a week the planting pots were checked for growth. The bags were opened under the laboratory hood when a seedling was visible and the root was swabbed to prepare a fungal culture.

In a second experiment the seed germination effect of the green mycorrhizal fungus (T. viride) (from experiment #0, Table 1) was compared to that of other soil fungi. Only untreated seeds were used in this experiment, and different soil fungi were added to untreated JSC-1A soil, in which also fungi originally present in the soil were allowed to interact with the introduced fungus. The fungi used for infection were isolated from swabs of JSC-1A soil, in which adult Opuntia plants were grown under carbon dioxide stress. We decided to use these

Table 1 . Seed germination results with and without T. viride addition.

fungi, since they were present around the roots of these extremely stressed plants that survived the stress treatment well. Five different fungus colonies and one bacterial culture were isolated from their soil previously in our lab. All experiments of this second seed germination series, including a control in which the soil was not infected, were prepared in parallel. The resulting germination rates (Table 2) were scored as positive (germination) or negative (no germination). SDA cultures were prepared from swabs of the roots of each seedling shortly after germination and of some seedlings again at a later time to determine the type of fungus growing on the root.

2.3. Fungus Staining and SEM Studies

From at least one seedling of each experiment with a positive result, roots were used for mycorrhizal staining followed by light microscopic studies. For the mycorrhizal fungus staining of the seedling roots the method described by Vierheilig et al. (1998) [16] was applied. The roots were rinsed well in tap water, boiled in 10% KOH for 5 min, stained for 3 min in a boiling hot 5% ink/vinegar solution (blue Pelican ink) and rinsed for 20 min with tap water containing a few drops of household vinegar. Other roots were used untreated for observation in the Scanning Electron Microscope (Hitachi TM-1000).

2.4. Mycorrhizal Fungi within Seedlings

Three seedlings from different pots were surface-sterilized, in order to look for evidence of fungi within the seedling itself. We followed the surface-sterilizing procedure described by Shekhawat et al. (2013) [17] for leaves and roots. Each plant was immersed into 75% ethanol for 1 min, then into an aqueous solution of sodium hypochloride (household bleach) for 15 min, followed by washing in 70% ethanol for 5 sec and rinsed in sterile distilled water. The seedling and its roots were then cut into segments, sliced open and placed on different SDA plates to examine fungus growth.

2.5. Interaction of an Endophytic with a Mycrorhizal Fungus

Finally, we also wanted to observe the interaction of the most prevalent green fungus obtained from root swabs of the seedlings in all our studies with one of the fungus species (Aspergillus terreus) identified from the crushed surface-sterilized seeds. In duplicate cultures (n = 2), both were placed on the same SDA plates near the edge and on opposite sides of each other and studied for their growth behavior over a period of three weeks.

On SDA plates with untreated JSC-1A many bacterial colonies grew first and a few green fungal colonies appeared after one week in culture. Autoclaved JSC-1A soil samples on SDA still produced a few bacterial, but no fungal colonies. Soil samples taken immediately after seed germination produced bacterial and fungal colonies on SDA. Microscopic analysis of samples from the colonies on the latter culture revealed a Penicillium sp., a Glomus sp., and cluster-forming, rod-shaped bacteria (Figure 1).

Table 2 . Seed germination after addition of different soil fungi and bacteria.

Figure 1 . Example Light microscopic images of fungus cultures from (previously autoclaved) JSC-1A soil sampled after seed germination—(a) Penicillium sp. (b) Glomus sp. and bacilli (Magnification: 1000×).

Untreated whole seeds placed on SDA produced no fungus growth at all, but some bacterial colonies, which were identified as small rods. From surface-sterilized crushed seeds a diversity of bacterial and fungal cultures grew on SDA. Among the fungi were: Aspergillus terreus, a Phoma sp., a Glomus sp., and a Penicillium sp., as demonstrated through light microscopic examination after LPCB staining (Figure 2). Bacteria were not identified. Not every seed produced the same fungi and bacteria. None of the tested surface sterilized seeds germinated during the experiment.

The regular (control) germination rate of non-surface-sterilized seeds on untreated JSC-1A was 55% (Table 1, exp. 0). Root swab cultures from the seedlings revealed predominantly a first white and later green colored fungus, which was identified as Trichoderma viride (Figure 3). The addition of the T. viride fungus from SDA cultures of the root swabs from control seedlings (exp. 0) had an obvious effect on the germination rate of the O. ficus-indica seeds in the autoclaved JSC-1A soil as recorded in Table1 Autoclaving the soil had reduced the total number of seed germination from 55% to 0% in the cultures without fungus addition. Adding T. viride to the seeds restored the germination rate to 29.6% and 42% respectively. In the exp. 1 and 2 (Table 1) the seeds were untreated, for exp. 3 (Table 1) seeds surface-sterilized with alcohol were used. Surface-sterilizing the seeds did affect germination negatively, with only one seed germinating very late in the culture where T. viride was added. Even an incubation period of six months did not improve this result.

Mainly T. viride was identified as the mycorrhizal fungus on the roots of all freshly germinated seedlings. Also present, but less frequent were Aspergillus nidulans, Penicillium sp., a Glomus sp., and a Phoma sp. Together with the T. viride fungus gram + bacilli and spore aggregates were regularly present (Figure 3).

In the second series of seed germination experiments the effect of different soil fungi and a bacterial colony on the seed germination rate of O. ficus-indica seeds was compared with the effect of the T. viride fungus. All seeds used in this series were untreated, planted into untreated JSC-1A and observed over a period of 3.5 months.

Figure 2 . Identified from surface sterilized crushed seed cultures were (a) Aspergillus terreus (b) Penicillium sp. (c) (d) Phoma sp., and (e) (f) Glomus sp. Magnification: 1000× (a)-(e), 400× (f).

Figure 3 . The green endomycorrhizal fungus Trichoderma viride (a) with pink colonies of gram + bacilli (b) on SDA (c) After Gram-staining and with white colonies of gram + bacilli and spore aggregates (d) on SDA (e) After Gram-staining. (Magnification 1000× (a, c, e)).

The germination time, germination rate and the fungi applied at the start of the experiment are listed in Table2 In addition the fungi present in the rhizosphere of the freshly sprouted seedlings are also listed in this table. The highest germination rate was obtained by infecting the soil with a mixture of T. viride and Glomus sp., which yielded a 47% germination rate. The second best germination rate (40%) was produced by adding Aspergillus glaucus to the seeds, followed by Penicillium bilaiae with the third highest germination rate (33%). Except for experiment #1, in which only one seed germinated, all swab cultures from seedling roots revealed the presence of T. viride and a Glomus sp., irrespective of the fungal culture originally used for infection. Some of the root swab cultures immediately showed strong presence of the T. viride fungus while it took some time to see it growing in others. We found that the earlier the green T. viride fungus appeared in the root swab cultures, the earlier the seeds had germinated in that experiment. Weak presence of T. viride thus resulted in poor seed germination.

Roots of seedlings from these infection experiments were studied in the SEM and the results compared to those obtained with light microscopy after mycorrhiza staining. Both examinations revealed that the roots were covered by mycorrhizal fungi—including the seedling of experiment #1 and that the Glomus sp. and T. viride fungi grew intertwined at the surface and into the inside of the roots (Figure 4). The hyphae of Glomus are coenocytic and wide, whereas those of T. viride are septate, thinner and branched at right angles. Glomus, a common AM fungus, forms vesicles inside the root, as depicted in the light-microscopic pictures (Figure 4). The vesicles inside the roots are clearly visible. Outside the root the characteristic sporangium of T. viride is recognizable (Figure 4(d) and Figure 4(e)).

A thin septate hypha extends from the sporangium into the root. Glomus sporangia are visible in the SEM picture from the root of a seedling (Figure 5(a)). In our SEM examination we found on several occasions penetration of Glomus hyphae into root cells (Figure 5(b)).

From sliced sections of the root tip of surface sterilized seedling roots we grew both T. viride and a Glomus sp. on SDA plates. Higher up within surface-sterilized roots and within the actual seedlings, we did not detect any T. viride, but a Glomus sp., Aspergillus sp. and a Phoma sp. were found to grow even high up inside the seedling.

The interaction of the endomycorrhizal fungi T. viride and Glomus with a pathogenic fungus, Aspergillus terreus, one of the fungi present inside the O. ficus-indica seeds, was demonstrated on SDA plates. Within two days the endomycorrhizal fungi produced a secretion that stained the agar yellow. Within a week the Aspergillus appeared contained in growth by the faster growing T. viride and Glomus sp. mix and a dense rim of T. viridesurrounded A. terreus.

On the nutrient poor soil JSC-1A good mycorrhizal fungus growth on and around the roots of O. ficus-indica seedlings could be documented through light and electron microscopic studies, confirming the results of Pope

Figure 4 . Mycorrhizae on roots of seedlings: septate hyphae of T. viride grow alongside the coenocytic Glomus hyphae outside of the root (a) Septate hyphae show penetration into the root (c). Vesicles formed by the AM fungus are visible (b) (d) (e) and on the root sporangia at the end of septate hyphae resemble those of T. viride (d) & (e) (Magnification 1000×).

Figure 5 . (a) Spores of the AM fungus on the root of a seedling (white arrow) (b) Penetration of the hyphae into the root cells are indicated by black arrows (SEM magnification 1500×).

(1993) [8] that poor soil conditions promote the presence of mycorrhizal fungi in the rhizosphere of plants. The endomycorrhizal fungus, T. viride, increased in our studies the germination rate of O. ficus-indica seeds by 29.6% and 42% respectively on the sterilized soil and under the conditions provided in this study (Table 1). On the roots of the young seedlings the presence of T. viride was regularly confirmed, even when other fungi or only bacteria were added to the soil together with the seeds (Table 2). The stronger the presence of T. viride was on the seedlings’ roots, the earlier the seeds had germinated. Though Aspergillus and Penicillium fungi also affected the germination rate of the seeds positively, the presence of T. viride in root swab cultures of the seedlings indicates that their role in the rhizosphere of the young seedling might be supportive of T. viride. Kumar (2012) [18] found the highest mycorrhizal root colonization in his studies with Menthaspicata when Glomus mossae and T. viride were inoculated together. The additional presence of an AM fungus, in our experiments an unidentified Glomus species, is demonstrated by the occurrence of vesicles and arbuscules in root tips of seedlings through light microscopic observation after mycorrhizal staining (Figure 4). AM fungi are Glomeromycota [4] . Their hyphae are coenocytic and the spores form terminal on them [19] , which we could demonstrate in SEM studies (Figure 5(a)). Endomycorrhizal fungi penetrate the root of the plant and enter the cells where they will live as intracellular symbionts. Penetration sites of the fungus into the root were recognized in our studies in the SEM (Figure 5(b)). De Jaeger et al. (2010) [20] describe that Trichoderma harzianum is penetrating its host plant through the arbuscular mycorrhizal fungus Glomus. In our studies with mycorrhizal fungus staining the septate and coenocytic hyphae seem to grow next to each other outside and inside the root (Figure 4). We could therefore not confirm a similar interaction of the two fungi in O. ficus-indica roots in our test system.

Glomus forms “Chlamydospores singly or in tight clusters (sporocarp), sometimes covered with hyphal peridium”, as described by Schenck and Smith (1982) [21] . Observations in the SEM (Figure 5(a)) revealed similar structures in our studies. Our combined data thus confirm also the presence of a Glomus sp. as the AM fungus in the rhizosphere of the O. ficus-indica seedlings on JSC-1A.

Mycorrhizal fungi reportedly protect the plant from pathogens by producing signaling compounds like abscisic acid [22] . Ellouze et al. (2012) [23] studied the effect of phytochemicals produced by the plant roots on fungus spore germination and concluded that the plants modify the microbial community of the soil to their advantage and the interaction is under genetic control. In our studies the T. viride—Glomus mixture produced a yellow secretion that prevented other fungi from growing on the same plate. Both endomycorrhizal fungi seem to be needed to secure the germination and establishment of the young O. ficus-indica seedling on the nutrient poor JSC-1A soil.

In this study we also demonstrate the presence of fungal spores and bacteria within the seeds of O. ficus-indica (Figure 2). Hijri et al. (2002) [19] found spores of pathogenic fungi inside the spores of an arbuscular mycorrhizal fungus without causing symptoms, which they interpreted as a possible weapon of the fungus against non-mycorrhizal competitors. The cultivation of a Glomus sp. among other fungi from sterilized seeds in our study (Figure 2) confirms the finding of Barrow et al. (2008) [10] that AM fungi can be transmitted through seeds. We speculate that the presence of fungal spores and bacteria in the seeds might give cacti an advantage on poor soils, which might also be poor of spores of mycorrhizal fungi that the plant could acquire. Thus the seeds of O. ficus-indica might be equipped with a “survival kit” to inhabit inhospitable areas. Dastager et al. (2010) [24] found that among phosphate solubilizing fungi, Aspergillus and Penicillium species were the most prominent. In our experiments both these two genera were represented in the rhizosphere of Opuntia seedlings (Table 2) and even within seeds themselves (Figure 2), another indication that their seeds are especially equipped for survival on nutrient poor soils.

AM fungi also interact with bacteria living on the roots and within the rhizosphere, but also incorporate bacteria as endosymbionts. The interaction between fungi and bacteria in the rhizosphere is another important finding of studies on plant fungus interactions. Dastager et al. (2010) [24] report on a plant growth promoting Micrococcus species. In our study the addition of a prevalent bacterial colony to O. ficus-indica seeds did not enhance the germination rate of the seeds. However, the presence of bacteria around and within the root tip and within the seeds seems to suggest a possible role in seed germination and/or establishment of the seedling. Puente et al. (2004) [5] found hardly any fungus spores in soil of arid areas they studied in Mexico. However, a large bacterial population was present in the rhizosphere of the plants, including cacti, which may indicate a stronger role of bacteria than fungi on dry soils.

The finding that alcohol sterilized seeds were negatively affected in their germination capability was a surprising result of our study, especially since alcohol is commonly used in seed sterilization procedures and was found to even enhance germination in some [15] . In their study these authors found that high concentrations of alcohol can be lethal to turfgrass seeds, which they studied. Alcohol seems to affect O. ficus-indica seedlings similarly however more studies are needed to confirm this finding.

In conclusion, the endomycorrhizal fungus T. viride demonstrates a close interaction with a Glomus species during and after germination of O. ficus-indica seeds on the nutrient poor lunar regolith simulant JSC-1A soil. They both grow in the root tip, however only the Glomus sp. was present within the rest of the seedling. The Glomus sp. is also passed on through the seed, together with bacteria and some pathogenic fungi, whose role still has to be evaluated.

Michelle J. Butcher, Jesus A. Castor-Macías, Ben Kohanloo, and Michelle Garcia and research reported in this publication were supported in part by the Research Initiative for Scientific Enhancement (RISE) Program funded by the National Institute of General Medical Sciences of the National Institutes of Health under Award Number R25GM060424. G. Konings-Dudin research was supported by the El Paso Community College (EPCC). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or EPCC.

We thank Dan Hawk, University of Wisconsin Green Bay, for providing the JSC-1A lunar regolith simulant.


Discussion

There have been numerous efforts to design PCR primers generally applicable for detection of the whole group of AMF ( Simon et al., 1992 Helgason et al., 1998 ), but later studies showed that they do not amplify DNA of all Glomeromycota or they also amplify ascomycetes, basidiomycetes or plant DNA ( Clapp et al., 1995 , 1999 Helgason et al., 1999 ). Other primers were successfully used for certain groups of the Glomeromycota ( Kjøller & Rosendahl, 2000 Redecker, 2000 Turnau et al., 2001 Wubet et al., 2003 , 2006 Gamper & Leuchtmann, 2007 ).

Many of the approaches require different primer pairs and independent PCR attempts for distinct target taxa. Comparison of such studies can be difficult since the distinct primer binding sites may behave very different in PCR and do not allow semiquantitative approaches. A single primer set for PCR amplification that covers all groups of the Glomeromycota and allows the identification of AMF at the species level was not available.

We have chosen the strategy of mixed primer sets to cover the defined sequence variability, instead of using fully degenerated primers. This reduces the degree of degeneration and results in a higher ratio of efficiently binding primers. The approach also allows adjustment of the concentrations of individual primers in future attempts. At the beginning of the study we speculated that the exonuclease activity of the proofreading DNA polymerase used could hamper discrimination by terminal 3′ primer mismatches, but no such problems were detected.

Primer specificity

The primers designed show some mismatches to AMF sequences at the 5′ end (Fig. 1), which do not hinder PCR amplification ( Bru et al., 2008 ). Primer mismatches such as C–T, T–C and T–G do not impair amplification strongly even when situated at the 3′ end of the primer ( Kwok et al., 1990 ). The forward primers SSUmAf as well as the reverse primers LSUmBr mismatched once with Ge. pyriformis, but did not hamper amplification. The LSU rDNA primers show sufficient sequence similarity to the target organisms, as the mismatches are either in the middle or at the 5′ end. LSUmAr primers displayed individual mismatches to sequences of Scutellospora spp., Gl. etunicatum, and one Acaulospora sp. (Fig. 1). Nevertheless, DNA of these species was successfully amplified from environmental samples and in the primer efficiency test (Fig. 3). Ambisporaceae and Archaeosporaceae species could not be included in the design of the LSU primers, but Ambispora fennica DNA from a single spore extraction (not shown) and Archaeospora sp. from single spores and roots of an Ecuadorian tree seedling (Table 3) could be amplified with the new primers, indicating well matching binding sites. Sequences from Otospora (Diversisporaceae Palenzuela et al., 2008 matching the SSU primers), Intraspora (closely related to Archaeospora), and Entrophospora (sensu Oehl & Sieverd. with two species only) are either not or only partly characterized and therefore could not be included in several aspects of primer design. Otospora and Intraspora are very closely related to their sister genera (maybe congeneric), so the lack of LSU rDNA sequences was therefore interpreted as a minor limitation.

We could successfully amplify all AMF tested with the new primers, but because of the lower number of LSU rDNA sequences available for AMF an optimization of the LSU primers might be reasonable in future. The discrimination against non-AMF and plant DNA is excellent, as shown on DNA extracts from environmental samples and spores from pot cultures. To discriminate against non-AMF, LSUmAr works much better than the nested primers LSUmBr. The cloned plant (Rumex) rDNA fragment that originated from root material can be interpreted as an ‘outlier’. The primer binding sites could not be investigated for Rumex, because of lacking sequence coverage. It should be indicated in this context that we did not use HPLC-purified primers. This means a certain fraction of primers may not be fully synthesized and could result in less specific amplification. All plasmids used in the plasmid test carried inserts that were originally amplified with SSUmAf. Therefore, the efficiency of this primer could not be validated, but because of the high number of SSU rDNA sequences known, it can be stated that the binding sites in the cloned fragments correspond to a realistic situation. The efficient amplification from spore DNA extracts was, moreover, confirmed in numerous former PCR.

Advantages over previously used PCR primer sets

In most former field studies SSU rDNA phylotypes were analysed for molecular detection of AMF. However, this region does not allow species resolution and each defined phylotype, irrespective of the used distance threshold value or phylogenetic analysis method, may hide a number of species ( Walker et al., 2007 ). In general, the LSU rDNA region allows species resolution, and thus the LSU primer pair FLR3–FLR4 ( Gollotte et al., 2004 ) was used for species-level community analyses. However, in particular, FLR4 is not phylogenetically inclusive ( Gamper et al., 2009 ) and discriminates many lineages, including Diversisporales, Archaeosporales and Paraglomerales, which results in a strong bias in community analyses towards the Glomeraceae. The primer FLR3 binds to DNA of many nontarget fungi as it shows no mismatch to > 1300 basidiomycete sequences and some ascomycete sequences in the public databases. Such problems obviously may bias tRFLP community analyses ( Mummey & Rillig, 2008 ) and seminested PCR approaches ( Pivato et al., 2007 ) using FLR3 and/or FLR4. The primer pair SSUGlom1–LSUGlom1 ( Renker et al., 2003 ) amplifies many non-AMF and plants. Combined with the primers ITS5–ITS4 in a nested PCR ( Hempel et al., 2007 ) this resulted in a 5.8S rDNA phylogenetic analysis, which resolved only the genus level. Even the ITS region does not always resolve species for AMF ( Stockinger et al., 2009 ).

In some cases, species-specific detection tools are available for individual species or certain well-defined and closely related species. The three closely related AM fungi Gl. mosseae, Gl. caledonium and Gl. geosporum were detected by using LSU primers in field studies ( Stukenbrock & Rosendahl, 2005 Rosendahl & Matzen, 2008 ), but these primers were designed to only amplify subgroups or certain taxa in the Glomeromycota. For the well-studied Gl. intraradices related AMF (e.g. DAOM197198), which are, however, not conspecific with Gl. intraradices ( Stockinger et al. 2009 ), microsatellite markers are available for their detection in the field ( Croll et al., 2008 Mathimaran et al., 2008 ). Some mtLSU region markers were also studied ( Börstler et al., 2008 ), but because of the high length variation observed (1070–3935 bp) and the difficulty in amplifying this region it is not very promising for community analyses. Thus, such markers cannot be used for general AMF community analyses.

The new primers described in the present study were used to amplify efficiently and specifically target rDNA from environmental samples of the main phylogenetic groups in the Glomeromycota. For the first time, this will allow molecular ecological studies covering all AMF lineages to be carried out with only one primer set. Furthermore, the long sequences allow robust phylogenetic analyses and species level resolution by inclusion of the variable ITS and LSU rDNA region ( Walker et al., 2007 Gamper et al., 2009 Stockinger et al. 2009 ), whereas formerly used primers mainly amplified rDNA fragments of up to 800 bp ( Helgason et al., 1999 Redecker, 2000 Lee et al., 2008 ).


Glomeromycota

Glomeromycota (informally glomeromycetes) is one of seven currently recognized phyla within the kingdom Fungi,[3] with approximately 200 described species.[4] Members of the Glomeromycota form arbuscular mycorrhizas (AMs) with the roots or thalli (e.g. in bryophytes) of land plants. Geosiphon pyriformis forms an endocytobiotic association with Nostoc cyanobacteria.[5] AM formation has not yet been shown for all species. The majority of evidence shows that the Glomeromycota are obligate biotrophs, dependent on symbiosis with land plants (Nostoc in the case of Geosiphon) for carbon and energy, but there is recent circumstantial evidence that some species may be able to lead an independent existence.[6] The arbuscular mycorrhizal species are terrestrial and widely distributed in soils worldwide where they form symbioses with the roots of the majority of plant species (>80%). They can also be found in wetlands, including salt-marshes, and associated with epiphytic plants.

The Glomeromycota have generally coenocytic (occasionally sparsely septate) mycelia and reproduce asexually through blastic development of the hyphal tip to produce spores[1] (Glomerospores) with diameters of 80-500μm.[7] In some, complex spores form within a terminal saccule.[1]

Initial studies of the Glomeromycota were based on the morphology of soil-borne sporocarps (spore clusters) found in or near colonized plant roots.[8] Distinguishing features such as wall morphologies, size, shape, color, hyphal attachment and reaction to staining compounds allowed a phylogeny to be constructed.[9] Superficial similarities led to the initial placement of genus Glomus in the unrelated family Endogonaceae.[10] Following broader reviews that cleared up the sporocarp confusion, the Glomeromycota were first proposed in the genera Acaulospora and Gigaspora[11] before being accorded their own order with the three families Glomaceae (now Glomeraceae), Acaulosporaceae and Gigasporaceae.[12]

With the advent of molecular techniques this classification has undergone major revision. An analysis of small subunit (SSU) rRNA sequences[13] indicated that they share a common ancestor with the Dikarya.[1]

Several species which produce glomoid spores (i.e. spores similar to Glomus) in fact belong to other deeply divergent lineages[14] and were placed in the orders, Paraglomerales and Archaeosporales.[1] This new classification includes the Geosiphonaceae, which presently contains one fungus (Geosiphon pyriformis) that forms endosymbiotic associations with the cyanobacterium Nostoc punctiforme[15] and produces spores typical to this phylum, in the Archaeosporales.

Work in this field is incomplete, and members of Glomus may be better suited to different genera[16] or families.[7]

The biochemical and genetic characterization of the Glomeromycota has been hindered by their biotrophic nature, which impedes laboratory culturing. This obstacle was eventually surpassed with the use of root cultures. The first mycorrhizal gene to be sequenced was the small-subunit ribosomal RNA (SSU rRNA).[17] This gene is highly conserved and commonly used in phylogenetic studies so was isolated from spores of each taxonomic group before amplification through the polymerase chain reaction (PCR). A molecular clock approach, based on the substitution rates of SSU sequences, was used to estimate the time of divergence of the fungi. The molecular analysis found that they are between 462 and 353 million years old.[7] The data enforces the long-held theory that they were instrumental in the colonization of land by plants.[18]

1. ^ a b c d e Schüßler, A. et al. (December 2001). "A new fungal phlyum, the Glomeromycota: phylogeny and evolution.". Mycol. Res. 105 (12): 1413–1421. doi:10.1017/S0953756201005196. http://journals.cambridge.org/action/displayAbstract?fromPage=online&aid=95091.
2. ^ Cavalier-Smith, T. (1998). "A revised six-kingdom system of Life". Biol. Rev. Camb. Philos. Soc. 73: 246. (as "Glomomycetes")
3. ^ Hibbett, D.S., et al. (March 2007). "A higher level phylogenetic classification of the Fungi". Mycol. Res. 111 (5): 509–547. doi:10.1016/j.mycres.2007.03.004. PMID 17572334.
4. ^ Neue Seite 1
5. ^ New Page 1
6. ^ Hempel, S., Renker, C. & Buscot, F. (2007). "Differences in the species composition of arbuscular mycorrhizal fungi in spore, root and soil communities in a grassland ecosystem". Environmental Microbiology 9 (8): 1930–1938. doi:10.1111/j.1462-2920.2007.01309.x. PMID 17635540.
7. ^ a b c Simon, L., Bousquet, J., Levesque, C., Lalonde, M. (1993). "Origin and diversification of endomycorrhizal fungi and coincidence with vascular land plants". Nature 363 (6424): 67–69. doi:10.1038/363067a0.
8. ^ Tulasne, L.R., & C. Tulasne (1844). "Fungi nonnulli hipogaei, novi v. minus cogniti auct". Giornale Botanico Italiano 2: 55–63.
9. ^ Wright, S.F. Management of Arbuscular Mycorrhizal Fungi. 2005. In Roots and Soil Management: Interactions between roots and the soil. Ed. Zobel, R.W., Wright, S.F. USA: American Society of Agronomy. Pp 183-197.
10. ^ Thaxter, R. (1922). "A revision of the Endogonaceae". Proc. Am. Acad. Arts Sci. 57 (12): 291–341. doi:10.2307/20025921. http://jstor.org/stable/20025921.
11. ^ J.W. Gerdemann & J.M. Trappe (1974). "The Endogonaceae in the Pacific Northwest". Mycologia Memoirs 5: 1–76.
12. ^ J.B. Morton & G.L. Benny (1990). "Revised classification of arbuscular mycorrhizal fungi (Zygomycetes): a new order, Glomales, two new suborders, Glomineae and Gigasporineae, and two new families, Acaulosporaceae and Gigasporaceae, with an emendation of Glomaceae". Mycotaxon 37: 471–491. http://www.cybertruffle.org.uk/cyberliber/59575/0037/0471.htm.
13. ^ Schüßler, A. et al. (January 2001). "Analysis of partial Glomales SSU rRNA gene sequences: implications for primer design and phylogeny". Mycol. Res. 105 (1): 5–15. doi:10.1017/S0953756200003725.
14. ^ Redeker, D. (2002). "Molecular identification and phylogeny of arbuscular mycorrhizal fungi". Plant and Soil 244: 67–73. doi:10.1023/A:1020283832275.
15. ^ Schüßler, A. (2002). "Molecular phylogeny, taxonomy, and evolution of Geosiphon pyriformis and arbuscular mycorrhizal fungi". Plant and Soil 224: 75–83. doi:10.1023/A:1020238728910.
16. ^ Walker, C. (1992). "Systematics and taxonomy of the arbuscular mycorrhizal fungi (Glomales) - a possible way forward". Agronomie 12: 887–897. doi:10.1051/agro:19921026.
17. ^ Simon, L. Lalonde, M. Bruns, T.D. (1992). "Specific Amplification of 18S Fungal Ribosomal Genes from Vesicular-Arbuscular Endomycorrhizal Fungi Colonizing Roots". American Society of Microbiology 58: 291–295.
18. ^ D.W. Malloch, K.A. Pirozynski & P.H. Raven (1980). "Ecological and evolutionary significance of mycorrhizal symbiosis in vascular plants (a review)". Proc. Natl Acad. Sci. USA 77 (4): 2113–2118. doi:10.1073/pnas.77.4.2113. PMC 348662. PMID 16592806. http://www.pnas.org/cgi/content/short/77/4/2113.

Source: Wikipedia, Wikispecies: All text is available under the terms of the GNU Free Documentation License


VISUAL CONNECTION

Figure 3: Two types of mycorrhizae. (a) Ectomycorrhizae and (b) arbuscular or endomycorrhizae have different mechanisms for interacting with the roots of plants. (credit b: MS Turmel, University of Manitoba, Plant Science Department)

If symbiotic fungi were absent from the soil, what impact do you think this would have on plant growth?


Answer:
Without mycorrhiza, plants cannot absorb adequate nutrients, which stunts their growth. Addition of fungal spores to sterile soil can alleviate this problem.

Figure 4: Mycorrhizae. The (a) infection of Pinus radiata (Monterey pine) roots by the hyphae of Amanita muscaria (fly amanita) causes the pine tree to produce many small, branched rootlets. The Amanita hyphae cover these small roots with a white mantle. (b) Spores (the round bodies) and hyphae (thread-like structures) are evident in this light micrograph of an arbuscular mycorrhiza by a fungus on the root of a corn plant. (credit a: modification of work by Randy Molina, USDA credit b: modification of work by Sara Wright, USDA-ARS scale-bar data from Matt Russell)

Other examples of fungus–plant mutualism include the endophytes: fungi that live inside tissue without damaging the host plant. Endophytes release toxins that repel herbivores, or confer resistance to environmental stress factors, such as infection by microorganisms, drought, or heavy metals in soil.


Link to Learning

Lichens are used to monitor the quality of air. Read more on this site from the United States Forest Service.

Fungus/Animal Mutualism

Fungi have evolved mutualisms with numerous insects in Phylum Arthropoda: joint-legged invertebrates with a chitinous exoskeleton. Arthropods depend on the fungus for protection from predators and pathogens, while the fungus obtains nutrients and a way to disseminate spores into new environments. The association between species of Basidiomycota and scale insects is one example. The fungal mycelium covers and protects the insect colonies. The scale insects foster a flow of nutrients from the parasitized plant to the fungus.

In a second example, leaf-cutter ants of Central and South America literally farm fungi. They cut disks of leaves from plants and pile them up in subterranean gardens ((Figure)). Fungi are cultivated in these disk gardens, digesting the cellulose in the leaves that the ants cannot break down. Once smaller sugar molecules are produced and consumed by the fungi, the fungi in turn become a meal for the ants. The insects also patrol their garden, preying on competing fungi. Both ants and fungi benefit from this mutualistic association. The fungus receives a steady supply of leaves and freedom from competition, while the ants feed on the fungi they cultivate.



Fungi, Bacteria, and Lichens

Thomas N. Taylor , . Michael Krings , in Paleobotany (Second Edition) , 2009

Glomeromycota

The Glomeromycota are a clade that was instituted based on rDNA phylogenies of living members (Schüssler et al., 2001 Redecker and Raab, 2006). The phylum, which includes the arbuscular mycorrhizal (AM) fungi, was formerly included within the Zygomycota, and is now considered to be the sister group of the clade formed by the Ascomycota+Basidio-mycota, based on molecular phylogenies (Blackwell et al., 2006 Redecker and Raab, 2006). Extant Glomeromycota are comprised of obligate symbionts that may form arbuscules in plant roots they produce large (40–800 μm), multilayered spores which are attached to non-septate hyphae. More than 90% of extant land plants have a symbiotic (mutualistic) relationship with mycorrhizal fungi in their roots. There are two basic types of extant mycorrhizae: ecto- and endomycorrhizae. Endomycorrhizae are formed by members of the glomeromycetes and are the most common form today. The fungal hyphae grow within the host root, and although they penetrate the host cell walls, they do not penetrate the plasma membranes. Most produce arbuscules, highly branched hyphal structures that provide for exchange between the fungal symbiont and its host. Some also produce storage organs called vesicles (the vesicular-arbuscular mycorrhizae). Ectomycorrhizae are formed by members of the Basidiomycota and a few ascomycetes they are less common today and occur primarily in woody plants of the temperate zone, including many conifers ( Chapter 21 ). In this case, the fungal hyphae form a net around the outside of the plant root, which penetrates between the cells of the root itself. Although there are many hypotheses for the establishment of this fungal–land plant association (e.g., resistance against drought, defense against root herbivory, etc.), the mutualistic association provides for increased mineral nutrient uptake by the plant in exchange for a source of carbon for the fungus.

The fossil record of Glomeromycota is believed to be ancient, extending well back into the Paleozoic. Spores and hyphae of a glomeromycotan type have been reported from rocks as old as the Cambrian (Pirozynski and Daplé, 1989), and from 460–455 Ma Upper Ordovician rocks (Redecker et al., 2000). Palaeoglomus grayi has aseptate (coenocytic) hyphae and spores that resemble living Glomus spores (Redecker et al., 2002). In these reports, however, there is no association with land plant remains, and thus the symbiotic association of these fungi remains unresolved. In addition, Butterfield (2005) noted that the very simplicity of these organisms does not provide enough diagnostic characters to separate them from other protists or parasites.

Palaeomyces is a name first used by Renault (1896a) ( FIG. 3.44 ) and later by Kidston and Lang (1921a) and others to describe large, isolated spores associated with Paleozoic plant remains some of these are now included in the morphotaxon Glomites ( FIG. 3.45 ). Although originally described from the Rhynie chert, they are also known from other sites. As well as being a source for beautifully preserved fossil fungi, the Rhynie chert ecosystem provides some of the best fossil evidence of fungi and land plants interacting in a symbiotic association. Glomites rhyniensis consists of aseptate hyphae and globose, multilayered spores, which occur in the tissues of several macroplants from this Devonian site (Taylor et al., 1995). Of special significance is the discovery of highly branched, intercellular arbuscules of this species within the cells of the land plant Aglaophyton major. In extant plants with AM fungi, arbuscules are confined to cells of the root, but in A. major the arbuscules occur in a narrow zone of cells inside the cortex, termed the mycorrhizal arbuscule zone ( FIGS. 3.46, 3.47 ), which extends throughout the rhizome and proximal portions of the aerial axes ( FIG. 3.48 ). More recently another land plant from the Rhynie chert, Rhynia gwynne-vaughanii, was described with Glomites fungi in the cortical tissues (Karatygin et al., 2006). In this taxon, G. sporocarpoides, arbuscules were not identified, but there were large, glomoid sporocarps containing numerous spores (20–24 μm in diameter) in the tissues. Rhizomatous and upright axes of Nothia aphylla, another land plant from the Rhynie chert, host a glomeromycotan fungus that closely resembles G. rhyniensis. Glomites rhyniensis is an intercellular endophyte, however, that becomes intracellular only within a well-defined region of the cortex where it forms arbuscules. The fungus in N. aphylla is initially intracellular, but later becomes intercellular in the cortex where it forms vesicles ( FIG. 3.49 ) and thick-walled spores (Krings et al., 2007b). If this fungus is functioning as an endomycorrhiza, N. aphylla displays an alternative mode of colonization by endomycorrhizal fungi, which may be related to the peculiar internal anatomy of the rhizomatous axis. In this part of the axis of N. aphylla, the cells are arranged in radial rows with virtually no intercellular spaces, so perhaps there is no intercellular infection pathway into the cortex (Krings et al., 2007a, b) this part of the axis of (see Chapter 8 for additional data on N. aphylla).


Watch the video: Episode 69 - Glomeromycota (September 2022).


  1. Gujar

    You are not right. I'm sure. Let's discuss this. Email me at PM.

  2. JoJozuru

    A very useful topic

  3. Adalwen

    I'm sorry, but in my opinion, you are wrong. I'm sure. I propose to discuss it.



Write a message