The protocol presents diverse microscopy techniques applied to understand the fungal colonization in mycoheterotrophic plants, from collecting to preparing samples in detail and include essential steps. Such techniques can be applied to various mycoheterotrophic plants, and even to a plant's materials other than mycoheterotrophics. This method is mainly related to structural botany and could also provide insights into mycorrhizal interactions, physiology, reproductive biology, evolution, and ecology of mycoheterotrophic plants.
To begin, collect the mycoheterotrophic plants by exploring the soil around the plant base, taking care not to damage the underground organs. Also, avoid pulling the plants from the ground to prevent disconnecting the aerial organs from the subterranean ones. Carefully, dig around aerial structures using a gardening trowel while exploring the subterranean organs like roots, stems, rhizomes and storage organs without damaging these structures.
Remove soil particles to preserve fragile structures and delicately wash these organs with tap water to rinse off the remaining soil particles before fixing the samples. Mycoheterotrophic plants associated with leaf litter demand extra attention. Hence, carefully collect the delicate organs connected to the decomposing material through their hyphae without pulling them from the connected structures.
Preserve structures with such connections and collect the litter for analysis. For transmission electron microscopy analysis, section the 3 to 4-millimeter-thick samples inside a drop of glutaraldehyde sodium cacodylate buffer into smaller sections of 1 to 2-millimeter thickness. Discard the cut edges outside the drop.
Ensure conducting the fixation process at the collection site immediately after the collection of plants. Immediately transfer the sections to a collection tube with a volume of fixative more than 10 times greater than the volume of the samples as it is an additive fixative. For surface analysis of the superficial hyphae in the organs, especially in subterranean ones and those in contact with leaf litter.
Observe fresh or fixed material under the dissecting microscope at a 7.5x or above magnification. Search for the areas of interest guided by the superficial hyphae and the rhizomorphs. Select the samples containing areas of superficial rhizomorphs as these can be sectioned to visualize pelotons and hyphae coils within cortical cells in roots and stems.
Prepare 0.2 milligrams per milliliter of wheat germ agglutinin fluorochrome conjugate and 1%calcofluor white solutions in 0.1 molar phosphate buffer as described in the text manuscript. Now, incubate the sections on glass slides by properly covering them with wheat germ agglutinin fluorochrome conjugate solution for 30 minutes. After the incubation, wash the sections with 0.1 molar phosphate buffer and incubate them in the calcofluor solution as a mounting medium.
Next, place cover slips on the slides and observe it under a confocal or fluorescence light microscope using the indicated filters. After fixing the samples, performing dehydration, and storing it in 70%ethanol, expose any desired surface for scanning electron microscopy analysis by using a sharp and new razor blade to make cuts with a one-way movement. If necessary, use a stereo microscope to select the samples and consider the metal stubs area while determining the sample sizes.
Further, dehydrate the samples in an ethanolic series. Maintain small and delicate samples for 30 minutes in each concentration and larger and denser samples for 1 hour. Next, fold small envelopes using tissue paper to organize samples for the next steps.
Larger samples can be handled without an envelope. Label the envelopes using a pencil and keep a log of the samples. Carefully, place the samples inside the envelopes using a paintbrush.
Now, keep these samples in absolute ethanol. Immediately proceed with critical point drying by operating a critical point dryer according to standard operating procedures. For this, place the samples in absolute ethanol in a sample holder and place it inside the critical point dryer's pressure chamber.
The intermediate fluid dissolves into the transition fluid at the critical carbon dioxide point, thus drying the samples. As reabsorption of atmospheric humidity can destroy the samples, store them in a desiccation container immediately after critical point drying. Before mounting the samples on metal stubs, put the gloves on to manipulate the stubs.
Immerse them in acetone for 5 minutes to eliminate any fat and let them dry. Under a stereo microscope, fix the samples on the stub using a conductive double-sided carbon adhesive tape and position them. The site from above is the only possible perspective in scanning electron microscopy images.
To manipulate the samples, use fine point tweezers. The sample part touched by the tweezers is usually damaged. So be careful and try touching parts positioned away from the areas of interest.
Maintain the stubs with samples in a sealed Petri dish with silica gel. Next, proceed with sputter coating to deposit a layer of gold or platinum on the surface of the samples in a low-pressure atmosphere of inert gas by following the standard operating procedures. The coating thickness depends on the sample's topography and is usually between 15 to 40 nanometers.
Finally, maintain the coated stubs in a sealed Petri dish with silica gel retaining humidity. The samples can be stored this way for weeks. Use a scanning electron microscope to analyze these samples.
An electron beam strikes the in vacuo sample during scanning electron microscopy and the signal emission from such interaction is interpreted as images. Ensure that the solutions and materials used in the symbiotic germination of seeds are sterile. to avoid contamination.
Start by autoclaving them for 20 minutes at 121 degrees Celsius. In a laminar airflow hood, superficially disinfect the fruits and seeds by immersing them in a sodium hypochlorite solution containing 2%active chlorine. Slim and fragile seeds can be immersed in a 1:1 diluted sodium hypochlorite solution.
After disinfection recuperate the seeds by filtering them using xerographic fabric. Wash the fruits and seeds thrice in autoclaved distilled water to remove the hypochlorite solution before proceeding with the germination tests. If necessary, store them in filter paper envelopes inside glass flasks with silica gel at 4 degrees Celsius and hermetically close the flasks and seal them with cling film, then evaluate the effectiveness of the washing process by transferring some drops of water from the last wash to potato dextrose agar.
For symbiotic germination of orchid seeds, incubate the seeds over 1 to 2-centimeter autoclaved filter paper discs placed in Petri dishes containing oatmeal agar culture medium. Now, inoculate the center of the Petri dish with the fragment of culture medium containing mycelium from the chosen isolated fungus. Seal the Petri dishes with cling film and incubate them in the dark at around 25 degrees Celsius or room temperature, depending on the fungal growth.
Prepare some dishes with seeds and without fungal inoculation as a negative control for the germination test. Analyze the germination results weekly by collecting quantitative and qualitative data and photographing protocorms and seedlings. The rhizomorphs can be easily recognized usually as dark, shoestring-like structures.
Freehand or other methods of obtaining sections thicker than 10 micrometers can better evince pelotons and provide more representative images of fungal patterns of colonization. Freehand sections can also be suitable for hyphae analysis in higher amplification. Although, details are better achieved in thinner sections.
The secondary cell walls in xylem elements can be easily identified by the light color toluidine acquires. Meanwhile, phloem elements composed only of primary cell walls are identified by their thinner and darker cell walls. Artifacts by autofluorescence can be seen in glycol methacrylate resin sections.
These artifacts are usually related to fluorochrome concentration and can be avoided by washing the samples a greater number of times with the buffer. The freehand section of the root shows internal and external hyphae. The same organ can be seen by scanning electron microscopy with an abundance of rhizomorphs and individual hyphae on its surface.
Most orchids species germinate within some weeks after being infected by the inoculated fungus or until nearly more than a month. Light in scanning electron microscopy can be applied to flowers, fruits, and seeds to investigate the reproductive biology or mycoheterotrophic plants. Symbiotic germination of seeds can also be tested in autotrophic orchids.
