This work was supported in part by an interagency agreement between NIOSH and NIEHS (AES12007001-1-0-6).

1. Preparing the NIOSH aerosol sampler
NOTE: The NIOSH aerosol sampler is a two-stage cyclone aerosol sampler that collects bioaerosols using two sampling tubes and a polytetrafluoroethylene (PTFE) filter.
<ol>
	<li>Before assembly, carefully inspect the sampler. Check the sampler to ensure it is not damaged, all screws are in place and snug, and the sealing tape around the seam formed by the two halves is intact. Inspect the sampler O-ring to ensure it does not have any nicks, cracks or tears and that it has a very light coating of silicone grease. 
	NOTE: The sampler includes a 37 mm 3 &#181;m PTFE filter in a polystyrene or polypropylene three-piece filter cassette.
	<ol>
		<li>Assemble the filter cassette by placing a cellulose or plastic filter support pad (included with the filters) on the gridded surface of the base piece (see Figure 1). Use filter forceps to put the filter on top of the filter support pad with the collection side facing up.</li>
		<li>Seal the filter cassettes using a manual or pneumatic press. Hand closure will allow air to leak around the filter and reduce the collection efficiency. Insert the ring-shaped piece (the extension cowl), and use the press to push it down tightly and evenly. Press down the extension cowl onto the filter tightly enough to ensure air does not leak around the filter. 
		NOTE: The top piece of the cassette is not used while sampling but should be saved to cover the filter once sampling is complete.</li>
		<li>Fit the assembled cassette onto the top of the sampler and push it down as far as possible. Wrap a piece of 19 mm (3/4 inch) tape around the outside of the filter cassette and sampler to hold the filter cassette in place and to act as a backup seal to prevent leaks.</li>
		<li>Screw each tube tightly and fully into the sampler until it bottoms out and wrap sealing tape around each tube to act as a secondary seal against leaks. 
		NOTE: The first tube, a 15 mL polypropylene tube, collects non-respirable particles with an aerodynamic diameter of &#62;4 &#181;m. The second tube, a 1.5 mL polypropylene microcentrifuge tube, collects respirable particles between 1 and 4 &#181;m. The remaining smaller particles, &#60;1 &#181;m, are collected on the PTFE filter (see Figure 2).</li>
	</ol>
	</li>
	<li>Calibrate the airflow through the NIOSH sampler with a calibration jar before each use as differences in the filters and samplers will cause the airflow to vary.
	<ol>
		<li>To use a calibration jar, insert the jar&#39;s Luer fitting into the top of the filter cassette, place the sampler in the jar, and seal the jar.</li>
		<li>Connect the calibration jar to a calibration flow meter and the sampling pump. Turn the flow meter on and allow it to warm up for a few minutes. Note that the flow meter must always have a filter attached to its inlet to avoid contaminating the flow meter and the sampler.</li>
		<li>Turn the sampling pump on and allow it to run for a few minutes to warm up and stabilize.</li>
		<li>Adjust the pump to set the correct flow rate at 3.5 L/min. 
		NOTE: The flow rate for the NIOSH aerosol sampler is usually set to 3.5 L/min. At this flow rate, the sampler conforms to the ACGIH/ISO criteria for respirable particle sampling16. Other flow rates can be used if different cut-off sizes are desired or to reduce the noise level, but be aware that if a different flow rate is used, then the sampler will no longer conform to the respirable sampling criteria.</li>
		<li>When calibration is finished, turn off the pump and flow meter.</li>
	</ol>
	</li>
</ol>
2. Static and personal aerosol sampling
<ol>
	<li>Set up the NIOSH aerosol sampler for either personal or static area sampling. 
	NOTE: Static sampling refers to using the aerosol sampler while it is attached to a tripod or other holding device (see Figure 3), as versus personal sampling when the sampler and pump are mounted on the person being studied (see Figure 4).
	<ol>
		<li>For static area sampling, keep the samplers as close as possible to the specific area being studied. Keep the samplers away from air inlets, room entrances and places that might be in the path of the airflow. 
		NOTE: Aerosol concentrations can vary considerably even over short distances, especially if the aerosol source is in the room17.</li>
		<li>For personal sampling, set up the sampler within the person&#39;s breathing zone. Attach the sampler to the lapel, shoulder or chest, and place the sampling pump at the waist or in a backpack. 
		NOTE: The breathing zone is commonly assumed to be a 30 cm hemisphere surrounding a person&#39;s nose from which the majority of air is drawn during inhalation17.</li>
	</ol>
	</li>
	<li>Make sure that the sampler tubing is well clear of the sampler inlets and that nothing is obstructing the inlets or interfering with the airflow into the sampler. The sampler tubing must not be pinched or kinked. Any blockage will cause the pump to stop.</li>
	<li>Turn on the pumps. After a minute or two, check if the pumps are still running. 
	NOTE: Conduct air sampling for time periods ranging from as little as 10 min to a full 8 h work shift, depending on the environment10,18,19. The results presented in Figure 6 are representative of a 60 min sampling period in an indoor environment.</li>
	<li>After air sampling is completed, collect the sample tubes and filter. Remove the sealing tape, unscrew the tubes, and cap them. Place the third piece of the filter cassette over the filter. Store samples at 4 &#176;C until ready for processing.</li>
</ol>
3. Extraction of genomic DNA from air samples
<ol>
	<li>Extract genomic DNA (gDNA) from each stage of the NIOSH BC251 sampler. Process the air sampling collection tubes and filter separately to allow for determination of respirable and non-respirable microbial aerosols in the sample. 
	NOTE: Alternatively, the three stages can be combined and extracted if sample inoculum is too small.
	<ol>
		<li>In a class II biological safety cabinet or laminar flow clean bench, aseptically remove the filter. Wipe the sampling cassette down using 70% ethanol and pry the sampling cassette open using a cassette-opening tool. Use a filter lifter to push the filter and support pad upwards. Grasp the filter using filter forceps and place it in a sterile petri dish. Cut the filter into 6 equal pieces and place them in a 2 mL reinforced tube containing 300 mg glass beads (0.2 - 0.5 mm).</li>
		<li>Place the tube containing the filter in liquid nitrogen for 30 s and immediately place it in a bead mill homogenizer set at 4.5 m/s for 30 s.</li>
		<li>Repeat step 3.1.2 once or twice until the filter is shredded. Small pieces of intact filter may remain. Add 0.5 mL of lysis buffer (4 M urea, 200 mM Tris, 20 mM NaCl, 200 mM ethylenediaminetetraacetic acid (EDTA), pH 7.4).</li>
		<li>Add 0.3 mL of lysis buffer to the 15 mL and 1.5 mL air sampler tubes. Vortex the tube for 10-&#160; 15 s while upright and then another 10 - 15 s inverted. Transfer the contents of each tube to 2 mL reinforced tubes containing 300 mg glass beads. 
		NOTE: Most of the material collected in each sampler tube will accumulate near the top of the tube.</li>
		<li>Process all tubes in the bead mill homogenizer at 4.5 m/s for 30 s and centrifuge them at 20,000 x g for 1 min at 22 &#176;C. Transfer supernatants to sterile 1.5 mL microcentrifuge tubes and centrifuge the tubes at 20,000 x g again for 1 min at 22 &#176;C. Repeat this step.</li>
	</ol>
	</li>
	<li>Add 30 &#181;L of lysis reagent (see Table of Materials) to each tube and incubate the tubes at 37 &#176;C for 15 min.</li>
	<li>Add 0.2 mL of binding buffer (10 M urea, 6 M guanidine-HCl, 10 mM Tris-HCl, 20% Triton X-100, pH 4.4) and proteinase K (100 &#181;g/mL) to each tube and incubate the tubes at 70 &#176;C for 10 min. Add 100 &#181;L of isopropanol to each tube.</li>
	<li>Transfer the extract solutions into glass fiber filter tubes (700 &#181;L capacity) placed in 2 mL collection tubes and centrifuge the collection tubes at 20,000 x g for 30 s at 22 &#176;C. Discard the collection tubes and place the filter tubes into new 2 mL collection tubes.</li>
	<li>Add 0.5 mL of inhibitor removal buffer (5 M guanidine-HCl, 20 mM Tris-HCl, pH 6.6, 38% ethanol) to each tube and centrifuge the collection tubes at 20,000 x g for 30 s at 22 &#176;C. Discard the collection tubes and place the filter tubes into new 2 mL collection tubes.</li>
	<li>Add 0.5 mL of wash buffer (20 mM NaCl, 2 mM Tris-HCl, pH 7.5, 80% ethanol) to each tube and centrifuge the collection tubes at 20,000 x g for 30 s at 22 &#176;C. Discard the collection tubes and place the filter tubes into new 2 mL collection tubes. Repeat this process.</li>
	<li>Centrifuge the collection tubes at 20,000 x g for an additional 1 min at 22 &#176;C to remove any residual wash buffer. Discard the collection tubes and place the filter tubes into new collection tubes.</li>
	<li>Add 100 &#181;L warm (&#8805;70 &#176;C) elution buffer (10 mM Tris-HCl, pH 8.5) to each tube and incubate them at room temperature for 1 - 2 min. Centrifuge the collection tubes at 20,000 x g for 30 s at 22 &#176;C.Transfer the empty filter tubes to a sterile 1.5 mL microcentrifuge tube and re-apply eluates from step 3.8. Incubate at room temperature for 1 - 2 min. Centrifuge at 20,000 x g for 30 s at 22 &#176;C. 
	NOTE: Eluates can be used immediately for step 4 or stored at -20 &#176;C until ready to use. Genomic DNA can be stored for up to a year at -20 &#176;C. It is recommended that DNA be stored at -80 &#176;C if long-term storage is required.</li>
</ol>
4. Amplification of fungal ribosomal DNA
<ol>
	<li>Use universal fungal primers to amplify ITS regions 1 and 2 from the extracted gDNA. 
	NOTE: For this protocol, the primer pair Fun18Sf (5&#39;-TTGCTCTTCAACGAGGAAT-3&#39;)/ITS4R (5&#39;-TCCTCCGCTTATTGATATGC-3&#39;) are used to provide the greatest coverage of the ITS regions4,5. Other primer sets, such as those that amplify ITS1 or ITS2 regions alone, can be used.
	<ol>
		<li>Set up polymerase chain reactions (PCR) for each sample in triplicate 50 &#181;L reactions in sterile 0.5 mL PCR tubes or 96-well PCR plates as follows: 5 &#181;L of extracted template DNA (from step 3), 33.3 &#181;L of PCR grade water, 5 &#181;L of 10x PCR buffer, 1.5 &#181;L of 50 mM MgCl2, 1 &#181;L of 10 mM 2&#39;-deoxynucleoside 5&#39;-triphosphates, 0.5 &#181;L of 20 &#181;M Fun18Sf forward primer, 0.5 &#181;L of 20 &#181;M ITS4R reverse primer, and 0.2 &#181;L of Taq DNA polymerase.</li>
		<li>Perform reactions in a thermal cycler: denaturation at 95 &#176;C for 3 min; 6 cycles of denaturation (96 &#176;C, 30 s) annealing (50 &#176;C, 45 s) and primer extension (72 &#176;C, 3 min); 20 cycles of denaturation (96 &#176;C, 30 s), annealing (50 &#176;C, 45 s), and primer extension (72 &#176;C,1 min); and primer extension at 72 &#176;C for 10 min. Keep the reaction mixtures at 4 &#176;C until next step.</li>
	</ol>
	</li>
	<li>Combine the triplicate PCR reactions purify using a silica membrane-based purification kit. 
	NOTE: This procedure allows for the removal of PCR components (primers, dNTPs, enzymes, salts, etc.) and other contaminants from the amplified DNA preparations.
	<ol>
		<li>Combine the three reactions for each sample (150 &#181;L total) in sterile 1.5 mL microcentrifuge tubes. Add binding buffer (5 M guanidine-HCl, 30% isopropanol) at a 5x volume (750 &#181;L) and mix the solution by pipetting up and down ten times.</li>
		<li>Add 450 &#181;L of the mixture to spin columns placed in collection tubes and centrifuge the tubes at 17,900 x g for 30 s. Discard the filtrates. Add the remaining 450 &#181;L to the columns and centrifuge at 17,900 x g for 30 s at 22 &#176;C. 
		NOTE: The spin columns contain a silica membrane that allows for adsorption of the DNA to the columns.</li>
		<li>Discard the filtrates and add 750 &#181;L washing buffer (10 mM Tris-HCl, pH 7.5, 80% ethanol) to the spin columns. Centrifuge the columns at 17,900 x g for 30 s at 22 &#176;C. 
		NOTE: This process removes the contaminants from the PCR reaction mentioned above.</li>
		<li>Discard the filtrate and centrifuge the empty columns at 17,900 x g for 1 min at 22 &#176;C. 
		NOTE: This step is to remove any residual washing buffer that may interfere with downstream applications.</li>
		<li>Transfer the spin columns to sterile 1.5 mL microcentrifuge tubes. Add 45 &#181;L of elution buffer (10 mM Tris-Cl, pH 8.5) and incubate the tubes at room temperature for 5 min. Centrifuge the spin columns at 17,900 x g for 1 min at 22 &#176;C.</li>
		<li>Use the eluted DNA immediately for steps 4 and 5 or store at -20 &#176;C until ready for use. 
		NOTE: The amplicons can be stored for up to a year at -20 &#176;C. It is recommended that DNA be stored at -80 &#176;C if long-term storage is required.</li>
	</ol>
	</li>
</ol>
5. Verification of fungal ITS amplification using agarose gel electrophoresis
<ol>
	<li>Cast a 1% agarose gel containing 1 &#181;g/mL ethidium bromide. Dissolve agarose in boiling 1x&#160;Tris-acetate-EDTA (TAE) buffer. Once the solution has cooled to approximately 50 &#176;C, add the ethidium bromide and pour into gel cast.</li>
	<li>After the agarose gel has solidified, immerse the gel in 1x&#160;TAE and prepare the amplicons to be loaded on the gel. To 8 &#181;L of amplified DNA, add 2 &#181;L of 5x loading buffer. 
	NOTE: These volumes can be adjusted depending on the concentration of the desired loading buffer.</li>
	<li>Load the samples, all 10 &#181;L, onto the gel along with a DNA ladder for size reference. Run the gel at 75 V (6 V/cm) for approximately 90 min and visualize bands, typically seen between 750 and 1,000 base pairs, using ultraviolet light (see Figure 5).</li>
</ol>
6. Sequencing and analysis of fungal ITS regions
<ol>
	<li>Sequence the extracted gDNA or amplified fungal ITS regions and analyze the sequences using current and appropriate methods.</li>
</ol>

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The findings and conclusions in this report are those of the authors and do not necessarily represent the official position of the National Institute for Occupational Safety and Health, Centers for Disease Control and Prevention.

Determining the fungal diversity within an occupational environment using sequencing-based approaches has improved fungal hazard identification and exposure assessment. Using this approach has allowed for the detection of many additional fungal species that are often not detected using culture or microscopy-based methods of assessment. A method for sampling bioaerosols from occupational and indoor environments and the extraction of genomic DNA from air samples for ITS amplification and sequencing is presented here. Determinations of fungal diversity using these methods is highly dependent on: (1) successful and complete extraction of genomic DNA from the air samples, (2) amplification of the gDNA with primers that allow for comparison of the amplified ITS sequences to banked sequences in the database and limit amplification biases, and (3) correct taxonomic identification of the sequences within the databases.
Successful genomic DNA extraction is reliant on the extraction methodologies and reagents employed in a study. As is mentioned in the protocol, a lot of the bioaerosols and other biological and non-biological particulate collected in the tube phases of the NIOSH two-stage cyclone sampler collect at the top of the tube. It is very important that the tubes be carefully opened when adding the lysis buffer as not to lose any particulate and that they be vortexed in a manner that allows for all the particulate to fall into the lysis buffer (right-side up and upside down). It is also important that the filter be carefully handled using aseptic methods as not to disrupt any bioaerosols that have collected on the surface prior to extraction or introduce contaminating DNA. It has been demonstrated that there is a large amount of variation between many of the commercially available genomic DNA extraction kits15. Some extraction procedures bias toward specific fungal species and result in varying DNA yields20. If the DNA yield following extraction is low, higher PCR reaction replicates can be performed. Not only do the extraction yields vary, but many of the kits contribute a substantial amount of microbial DNA contamination. For this reason, it is important to include proper controls throughout the procedure to identify and remove potential reagent contaminants from the analysis.
ITS amplification is another critical step of the protocol. The extraction methodologies used can vary in their ability to remove PCR inhibitors from the environmental samples. This is particularly concerning with environmental dust samples21. PCR amplification of samples extracted using the method presented here exhibited some PCR inhibition after the addition of 10 mg of dust15. Although PCR inhibition is of greatest concern in environmental dust samples, it should still be a consideration when amplifying DNA extracted from air samples for determining fungal diversity. The PCR primer set used in this method provides coverage of both the ITS 1 and ITS 2 regions. This allows for comparison to many of the sequences placed in the sequence databases. As with the extraction procedure, many amplification and primer biases have been previously described22. The fungal genome size and gene copy number could also influence ITS amplification23. These biases should be taken into consideration when interpreting the resultant sequence data. Taxonomic identification of ITS sequences is perhaps the most critical component of this method. It is important to keep identification criteria consistent. The ability to identify ITS sequences is dependent on the sequences banked in the databases. Many sequences cannot be identified to the species level. Having specific criteria when making taxonomic identifications based on banked sequences, like percentage identity cutoffs, is critical for keeping datasets consistent from study to study.
Compared to traditional assessment approaches, extracting and sequencing fungal DNA from occupational air samples provides a more complete representation of fungal diversity within an environment. In addition to the limitations discussed above, it must be noted that the results of these types of analyses are semi-quantitative unlike culture, spore counts, or quantitative PCR24 that can yield quantitative data. Sequencing data can be used to determine relative abundance per sample as well as calculate fungal diversity metrics. These methods can be used in conjunction with other methods, like qPCR, to get more quantifiable data. The datasets created using these approaches can be used in occupational health studies to gain a better understanding of fungal hazards in specific occupational environments. Developing more standardized approaches of extraction and primer selection are necessary to better characterize and compare sequencing-based studies of fungal diversity.

Fungal exposures in indoor and occupational environments can result in respiratory morbidities, including allergic sensitization and asthma1. Identification of fungal hazards is important for assessing risk and preventing worker exposure. These fungal hazards may be a result of indoor contamination, outdoor air intrusion, or environmental disturbances that result in the transport of fungal materials into areas where workers are present2. Methods to assess fungal exposure have included viable culture sampling as well as microscopic identification of fungal spores. These approaches have several limitations and often overlook many fungal species that could be contributing to the overall fungal burden3. Culture-based approaches can only differentiate those viable fungal organisms that can be cultivated on nutrient media. Identifying fungal spores to species level via microscopy can be confounded by spores sharing similar morphologies. Both methods are highly dependent on mycologists to analyze and identify the fungal species, with many remaining unidentified.
To improve upon existing methodologies used in occupational hazard identification and exposure assessments, many researchers have turned to molecular-based technologies. Sequencing-based approaches for assessing microbial diversity within indoor and occupational environments have revealed a broader spectrum of fungal species encountered compared to methods such as microscopy and viable culture3,4,5. The method presented here describes the air sampling of occupational environments and extraction of genomic DNA for the identification of potential fungal hazards. Hazard identification is accomplished by sequencing the nuclear ribosomal internal transcribed spacer, or ITS, regions that are highly variable among fungi and have been commonly used to differentiate fungal species6,7,8,9. Many species found in occupational settings, such as some species belonging to the phylum Basidiomycota, are not identifiable in viable culture and are difficult to differentiate microscopically. These fungi have been observed in high relative abundance within indoor and occupational environments assessed by sequencing fungal ITS regions3,4,10. ITS sequencing has provided greater knowledge into the diversity of fungi encountered within indoor and occupational environments.
The protocol described here details the methods used to collect, extract, and amplify fungal ITS regions from bioaerosols for sequence analysis. This approach utilizes the National Institute for Occupational Safety and Health (NIOSH) two-stage cyclone aerosol sampler to collect particulates in the air. This sampler was developed to collect bioaerosols and separate respirable (&#8804;4 &#181;m aerodynamic diameter) and non-respirable (&#62;4 &#181;m aerodynamic diameter) particles, which allows for identification of fungal organisms within indoor environments that are most likely to be inhaled by a worker11. Other air samplers, including cyclone samplers, are available on the market that have the ability to collect particles within the respirable range (&#60;4 &#181;m) using filters12,13. In contrast, the NIOSH two-stage cyclone aerosol sampler separates fungal species based on their aerodynamic diameter into disposable, polypropylene tubes that can be immediately processed for downstream applications14.
The processes of extracting genomic DNA and amplifying the fungal ITS regions are detailed in this protocol. The extraction methodologies presented have been developed specifically for the extraction of genomic DNA from fungi and bacteria, as many commercial kits target mammalian cells, bacteria, or specifically yeasts15. The primers used in this study are selected based on their overall coverage of both the fungal ITS 1 and ITS 2 regions4,5. Sequencing of these regions allows for the comparison of many banked ITS sequences, including those that sequence the ITS 1 region, the ITS 2 region, or both the ITS 1 and ITS 2 regions. The fungal diversity of air samples collected in an indoor setting using these methods are shown, revealing a substantial number of sequences placed in the phyla Ascomycota and Basidiomycota as well as other sequences belonging to less dominant fungal phyla, such as Zygomycota. The broad diversity of fungal sequences identified using this approach would not be captured using traditional hazard identification methodologies like cultivation or microscopy. Sequencing of fungal ITS regions provides an enhanced method to identify fungal hazards and allow for a better understanding of indoor and occupational fungal exposures.

<table><tbody><tr><td>NIOSH BC251 bioaerosol cyclone sampler</td><td>NIOSH</td><td>BC251</td><td>The NIOSH sampler is not yet commercially available. Please contact William Lindsley, PhD (wlindsley@cdc.gov) for information on obtaining the NIOSH sampler</td></tr><tr><td>Fisherbrand Sterile Microcentrifuge Tubes with Screw Caps</td><td>Fisher Scientific</td><td>02-681-373</td><td>1.5 mL polypropylene microcentrifuge tubes for air sampling; screw top threading must match the threading of the NIOSH sampler</td></tr><tr><td>Falcon 15 mL Conical Centrifuge Tubes</td><td>Corning</td><td>352096</td><td>15 mL polypropylene tubes for air sampling</td></tr><tr><td>Clean Room Vinyl Tape, Easy-Remove, 1/4&quot; Width</td><td>McMaster-Carr</td><td>76505A1</td><td>sealing tape</td></tr><tr><td>Filter Cassette, Clear Styrene, 37 mm</td><td>SKC Inc.</td><td>225-3LF</td><td>3-piece sampling cassette (no filter). Contains: cassette base, extension cowl, cassette cap and inlet/outlet plugs</td></tr><tr><td>PTFE hydrophobic fluoropore membrane filters, 3.0 &amp;micro;m, 37 mm</td><td>EMD Millipore</td><td>FSLW03700</td><td>Contains: 37 mm, 3.0 &amp;micro;m PTFE filters and support pads</td></tr><tr><td>Fisherbrand filter forceps</td><td>Fisher Scientific</td><td>09-753-50</td><td>filter forceps</td></tr><tr><td>Model 502 Precision PanaPress</td><td>PanaVise</td><td>502</td><td>pneumatic cassette press is constructed from this precision arbor press</td></tr><tr><td>Scotch Super 33+ vinyl electrical tape</td><td>McMaster-Carr</td><td>76455A21</td><td>19 mm tape</td></tr><tr><td>Multi-purpose Calibration Jar, Large</td><td>SKC Inc.</td><td>225-112</td><td>calibration jar</td></tr><tr><td>Universal PCXR4 Sample Pump</td><td>SKC Inc.</td><td>224-PCXR4</td><td>sampling pump</td></tr><tr><td>Mass Flowmeter 4140</td><td>TSI Inc.</td><td>4140</td><td>flow meter</td></tr><tr><td>Roche High Pure PCR Template Kit</td><td>Roche Diagnostics</td><td>11796828001</td><td>Kit used for genomic DNA extraction. Contains: Lysis buffer, Binding buffer, Proteinase K, Inhibitor removal buffer, Wash buffer, Elution buffer, Glass fiber filter tubes and 2 ml collection tubes</td></tr><tr><td>Fisherbrand 2 mL Reinforced Polypropylene Screw Cap Tubes with Caps</td><td>Fisher Scientific</td><td>15340162</td><td>2 mL reinforced tubes for bead homogenization</td></tr><tr><td>Glass beads, acid washed, 212-300 &amp;micro;m</td><td>Sigma-Aldrich</td><td>G1277</td><td>glass beads</td></tr><tr><td>Fisher Scientific Bead Mill 24 Homogenizer</td><td>Fisher Scientific</td><td>15-340-163</td><td>bead homogenizer</td></tr><tr><td>CelLytic B Cell Lysis Reagent, 10X</td><td>Sigma-Aldrich</td><td>C8740</td><td>lysis reagent</td></tr><tr><td>Platinum Taq polymerase</td><td>Invitrogen</td><td>10966-018</td><td>Contains: Platinum Taq polymerase, 10X PCR buffer (no MgCl2), 50 mM MgCl2, KB Extender</td></tr><tr><td>dNTP Mix</td><td>Invitrogen</td><td>18427-088</td><td>10 mM dNTP mix</td></tr><tr><td>QIAquick PCR Purification Kit</td><td>Qiagen</td><td>28106</td><td>Kit used to purify fungal amplicons. Contains: Buffer PB (binding buffer), Buffer PE (washing buffer), Buffer EB (elution buffer), pH Indicator dye (optional), and GelPilot loading dye</td></tr><tr><td>Owl EasyCast Mini Gel Electrophoresis System</td><td>Thermo Fisher</td><td>B1 or B2</td><td></td></tr><tr><td>TrackIt 1 KB Plus DNA Ladder</td><td>Thermo Fisher</td><td>10488-085</td><td>DNA ladder</td></tr></tbody></table>

collection and extraction of occupational air samples for analysis of fungal dna

Traditional methods of identifying fungal exposures in occupational environments, such as culture and microscopy-based approaches, have several limitations that have resulted in the exclusion of many species. Advances in the field over the last two decades have led occupational health researchers to turn to molecular-based approaches for identifying fungal hazards. These methods have resulted in the detection of many species within indoor and occupational environments that have not been detected using traditional methods. This protocol details an approach for determining fungal diversity within air samples through genomic DNA extraction, amplification, sequencing, and taxonomic identification of fungal internal transcribed spacer (ITS) regions. ITS sequencing results in the detection of many fungal species that are either not detected or difficult to identify to species level using culture or microscopy. While these methods do not provide quantitative measures of fungal burden, they offer a new approach to hazard identification and can be used to determine overall species richness and diversity within an occupational environment.

The species distribution within an environment can be assessed using relative abundance by determining the number of clones of each OTU identified in the air samples. Figure 6 is a Krona chart representing the taxonomically placed species within an indoor environment following 60 min of air sampling. It can be observed that the environment contains a variety of species within two major fungal phyla, Ascomycota and Basidiomycota, as well as species belonging to the phylum Zygomycota (Rhizopus microsporus). Traditional methods of assessment are biased toward Ascomycota species, as many Basidiomycota are not culturable or cannot be differentiated using microscopy-based methods. Sequencing of fungal ITS regions allows for the detection of numerous fungal species, specifically some Basidiomycota, that often go undetected. Analysis of this environment shows 68% of the fungal sequences identified belonged to the Basidiomycota with most of them placed in the order Polyporales.
The taxonomic data obtained using sequencing-based approaches can be used to determine the taxonomic diversity within an environment. Diversity indices, such as those presented in Table 1, determine how rich the environment is, which describes the number of species identified in each sample, as well as how diverse the environment is, which takes into account the number of species and the abundance of each. Table 1 shows the Chao 1 richness indices and Shannon diversity indices for two indoor environments. The indices indicate higher richness and diversity in air-conditioned environments compared to evaporative cooler environments. Also included are Bray-Curtis dissimilarity coefficients. These describe how dissimilar samples within the environment are regarding the species identified within each sample. The samples within both of the environments described in Table 1 are highly dissimilar at 98% and 97% for air conditioner and evaporative cooler environments, respectively.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56730/56730fig1.jpg" /> 
Figure 1: Schematic representation of a three-piece filter cassette assembly. The filter cassette is made up of three pieces: a top or cap piece (inlet), an extension cowl and a base piece (outlet). A support pad and filter are placed on the gridded surface of the base piece. The extension cowl is then sealed onto the base using a manual or pneumatic press. For use with the NIOSH bioaerosol cyclone sampler, the top piece is left off during sampling and is then replaced for storage and transport. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56730/56730fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56730/56730fig2.jpg" /> 
Figure 2: Schematic representation of the NIOSH bioaerosol cyclone sampler. The NIOSH cyclone aerosol sampler collects airborne particulate by drawing air into the sampler through the inlet, and producing a cyclone that deposits particles on the wall of the sample tubes. Air is first drawn into a 15 mL polypropylene tube, where large particles (&#62;4 &#181;m) collect on the walls of the tube. Air then flows out of the first tube and is drawn into a 1.5 mL microcentrifuge tube, where smaller particles (1 to 4 &#181;m) collect. The remaining particles (&#60;1 &#181;m) collect on a PTFE filter as the air is drawn out of the sampler. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56730/56730fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56730/56730fig3.jpg" /> 
Figure 3: Example of a static sampler set-up. The image demonstrates samplers set up for static area sampling. The NIOSH samplers are set up to collect 40 inches (102 cm) and 60 inches (152 cm) from the floor, representing heights of a sitting and standing adult, respectively. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56730/56730fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56730/56730fig4.jpg" /> 
Figure 4: Example of a personal sampler assembly. The image illustrates a NIOSH sampler worn on a backpack by an individual. The sampler and power switch are located on the straps of the pack while the sampling pump is located within the backpack itself. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56730/56730fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56730/56730fig5.jpg" /> 
Figure 5: Example of an agarose gel visualizing amplified ITS DNA from air samples. The image shows DNA bands between 750 and 1,000 base pairs. These bands represent amplified fungal ITS regions from extracted air samples. ITS regions vary in size depending on the species origin. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56730/56730fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56730/56730fig6.jpg" /> 
Figure 6: Example Krona chart presenting taxonomically placed fungal species identified within an indoor environment. The Krona chart depicts the relative abundance of fungal species found in 4 samples collected from homes following a 60 min air sampling period with the NIOSH bioaerosol cyclone sampler. 68% of the species identified belong to the phylum Basidiomycota with the remaining belonging to the Ascomycota (33%) and Zygomycota (1%). The most abundant ITS sequences in this environment belonged to the order Polyporales and were identified as Phanerodontia chrysosporium and Gelatoporia dichroa. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56730/56730fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="2"></td>
			<td>Chao-1 Richness</td>
			<td>Shannon Diversity</td>
			<td>Bray-Curtis Distance</td>
		</tr>
		<tr>
			<td>mean (min, max)</td>
			<td>mean (min, max)</td>
			<td>mean (min, max)</td>
		</tr>
		<tr>
			<td>Air Conditioner (n = 9)</td>
			<td>33.86 (15.0, 102.0)</td>
			<td>1.39 (0.68, 2.51)</td>
			<td>0.9778 (0.8313, 1.00)</td>
		</tr>
		<tr>
			<td>Evaporative Cooler (n = 10)</td>
			<td>25.46 (12.5, 49.5)</td>
			<td>1.10 (0.50, 1.72)</td>
			<td>0.9624 (0.3804, 1.00)</td>
		</tr>
	</tbody>
</table>
Table 1: Example data presenting diversity and species richness indices comparing indoor environments. The table shows Chao 1 richness indices (number of species per sample), Shannon diversity indices (number of species and abundance of each), and Bray-Curtis dissimilarity coefficients (how dissimilar the species distribution is between samples on a scale of 0 - 1). These data demonstrate higher richness and diversity in air-conditioned environments and high dissimilarity among samples in both environments. This table has been modified from Lemons et al.10 with permission from the Royal Society of Chemistry.

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Collection and Extraction of Occupational Air Samples for Analysis of Fungal DNA | Environment | Video

Collection and Extraction of Occupational Air Samples for Analysis of Fungal DNA

Determining the fungal diversity within an environment is a method utilized in occupational health studies to identify health hazards. This protocol describes DNA extraction from occupational air samples for amplification and sequencing of fungal ITS regions. This approach detects many fungal species that can be overlooked by traditional assessment methods.

Traditional methods of identifying fungal exposures in occupational environments, such as culture and microscopy-based approaches, have several ...

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12.3K Views.  Centers for Disease Control and Prevention. Het bepalen van de schimmel diversiteit binnen een omgeving is een methode gebruikt in studies van de gezondheid op het werk te identificeren van de gevaren voor de gezondheid. Dit protocol beschrijft DNA-extractie uit beroepsmatige lucht monsters voor de versterking en de sequencing van schimmel ITS-regio's. Deze aanpak detecteert vele schimmel soorten die kunnen worden overzien door traditionele beoordelingsmethoden.

Video: Collection and Extraction of Occupational Air Samples for Analysis of Fungal DNA

Air-sampled Filter Analysis for Endotoxins and DNA Content

Outdoor aerosol research commonly uses particulate matter sampled on filters. This procedure enables various characterizations of the collected particles to be performed in parallel. The purpose of the method presented here is to obtain a highly accurate and reliable analysis of the endotoxin and DNA content of bio-aerosols extracted from filters. The extraction of high molecular weight organic molecules, such as lipopolysaccharides, from sampled filters involves shaking the sample in a pyrogen-free water-based medium. The subsequent analysis is based on an enzymatic reaction that can be detected using a turbidimetric measurement. As a result of the high organic content on the sampled filters, the extraction of DNA from the samples is performed using a commercial DNA extraction kit that was originally designed for soils and modified to improve the DNA yield. The detection and quantification of specific microbial species using quantitative polymerase chain reaction (q-PCR) analysis are described and compared with other available methods.

Two complementary analyses of atmospheric biological particles from air sampled filters are described herein: the extraction and detection of endotoxin, and of DNA.

Outdoor aerosol research commonly uses particulate matter sampled on filters. This procedure enables various characterizations of the collected particles ...

Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered &#34;bee-safe,&#34; there is mounting evidence that fungicide residues in pollen are associated with bee declines (for both honey and bumble bee species). While the mechanisms remain relatively unknown, researchers have speculated that bee-microbe symbioses are involved. Microbes play a pivotal role in the preservation and/or processing of pollen, which serves as nutrition for larval bees. By altering the microbial community, it is likely that fungicides disrupt these microbe-mediated services, and thereby compromise bee health. This manuscript describes the protocols used to investigate the indirect mechanism(s) by which fungicides may be causing colony decline. Cage experiments exposing bees to fungicide-treated flowers have already provided the first evidence that fungicides cause profound colony losses in a native bumble bee (Bombus impatiens). Using field-relevant doses of fungicides, a series of experiments have been developed to provide a finer description of microbial community dynamics of fungicide-exposed pollen. Shifts in the structural composition of fungal and bacterial assemblages within the pollen microbiome are investigated by next-generation sequencing and metagenomic analysis. Experiments developed herein have been designed to provide a mechanistic understanding of how fungicides affect the microbiome of pollen-provisions. Ultimately, these findings should shed light on the indirect pathway through which fungicides may be causing colony declines.

Microbial consortia within bumble bee hives enrich and preserve pollen for bee larvae. Using next generation sequencing, along with laboratory and field-based experiments, this manuscript describes protocols used to test the hypothesis that fungicide residues alter the pollen microbiome, and colony demographics, ultimately leading to colony loss.

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered ...

Quantification of Fungal Colonization, Sporogenesis, and Production of Mycotoxins Using Kernel Bioassays

The rotting of grains by seed-infecting fungi poses one of the greatest economic challenges to cereal production worldwide, not to mention serious risks to human and animal health. Among cereal production, maize is arguably the most affected crop, due to pathogen-induced losses in grain integrity and mycotoxin seed contamination. The two most prevalent and problematic mycotoxins for maize growers and food and feed processors are aflatoxin and fumonisin, produced by Aspergillus flavus and Fusarium verticillioides, respectively.
Recent studies in molecular plant-pathogen interactions have demonstrated promise in understanding specific mechanisms associated with plant responses to fungal infection and mycotoxin contamination1,2,3,4,5,6. Because many labs are using kernel assays to study plant-pathogen interactions, there is a need for a standardized method for quantifying different biological parameters, so results from different laboratories can be cross-interpreted. For a robust and reproducible means for quantitative analyses on seeds, we have developed in-lab kernel assays and subsequent methods to quantify fungal growth, biomass, and mycotoxin contamination. Four sterilized maize kernels are inoculated in glass vials with a fungal suspension (106) and incubated for a predetermined period. Sample vials are then selected for enumeration of conidia by hemocytometer, ergosterol-based biomass analysis by high performance liquid chromatography (HPLC), aflatoxin quantification using an AflaTest fluorometer method, and fumonisin quantification by HPLC.

The devastation of cereal crops by seed-infecting fungi has prompted numerous research efforts to better understand plant-pathogen interactions. To study seed-fungal interactions in a laboratory setting, we developed a robust method for the quantification of fungal reproduction, biomass, and mycotoxin contamination using kernel bioassays.

The rotting of grains by seed-infecting fungi poses one of the greatest economic challenges to cereal production worldwide, not to mention serious risks ...

Immunology and Infection

Composition and Distribution Analysis of Bioaerosols Under Different Environmental Conditions

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. In this study, we described a protocol for multiple analyses of the biological compositions in environmental PM. Five experiments are presented: (1) PM number monitoring by using a laser particle counter; (2) PM collection by using a cyclonic aerosol sampler; (3) PM collection by using a high-volume air sampler with filters; (4) culturable microbes collection by the Andersen six-stage sampler; and (5) detection of biological composition of environmental PM by bacterial 16SrDNA and fungal ITS region sequencing. We selected hazy days and a livestock farm as two typical examples of the application in this protocol. In this study, these two sampling methods, cyclonic aerosol sampler and filter sampler, showed different sampling efficiency. The cyclonic aerosol sampler performed much better in terms of collecting bacteria, while these two methods showed the same efficiency in collecting fungi. Filter samplers can work under low temperature conditions while cyclonic aerosol samplers have a sampling limitation for temperature. A solid impacting sampler, such as an Andersen six-stage sampler, can be used to sample bioaerosols directly into the culture medium, which increases the survival rate of culturable microorganisms. However, this method mainly relies on culture while more than 99% of microbes cannot be cultured. DNA extracted from the culturable bacteria collected by the Andersen six-stage sampler and samples collected by cyclonic aerosol sampler and filter sampler were detected by bacterial 16S rDNA and fungal ITS region sequencing.All the methods above may have wide application in many fields of study, such as environmental monitoring and airborne pathogen detection. From these results, we can conclude that these methods can be used under different conditions and may help other researchers further explore the health impacts of environmental bioaerosols.

Understanding the biological composition of environmental particulate matter is important for the study of its significant impacts on human health and disease spread. Here, we used three types of bioaerosol sampling methods and a biological analysis of airborne microbes to better explore airborne microbial communities under different environmental conditions.

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. In this study, ...

Application of DNA Fingerprinting using the D1S80 Locus in Lab Classes

In biological sciences, DNA fingerprinting has been widely used for paternity testing, forensic applications and phylogenetic studies. Here, we describe a reliable and robust method for genotyping individuals by Variable Number of Tandem Repeat (VNTR) analysis in the context of undergraduate laboratory classes. The human D1S80 VNTR locus is used in this protocol as a highly polymorphic marker based on variation in the number of repetitive sequences.
This simple protocol conveys useful information for teachers and the implementation of DNA fingerprinting in practical laboratory classes. In the presented laboratory exercise, DNA extraction followed by PCR amplification is used to determine genetic variation at the D1S80 VNTR locus. Differences in the fragment size of PCR products are visualized by agarose gel electrophoresis. The fragment sizes and repeat numbers are calculated based on a linear regression of the size and migration distance of a DNA size standard.
Following this guide, students should be able to:
&#8226;&#160; Harvest and extract DNA from buccal mucosa epithelial cells 
&#8226;&#160; Perform a PCR experiment and understand the function of various reaction components 
&#8226;&#160; Analyze the amplicons by agarose gel electrophoresis and interpret the results 
&#8226;&#160; Understand the use of VNTRs in DNA fingerprinting and its application in biological sciences

Application of DNA Fingerprinting using the D1S80 Locus in Lab Classes

Here we describe a simple protocol to generate DNA fingerprinting profiles by amplifying the VNTR locus D1S80 from epithelial cell DNA.

In biological sciences, DNA fingerprinting has been widely used for paternity testing, forensic applications and phylogenetic studies. Here, we describe a ...

Genetics

Breath Collection from Children for Disease Biomarker Discovery

Breath collection and analysis can be used to discover volatile biomarkers in a number of infectious and non-infectious diseases, such as malaria, tuberculosis, lung cancer, and liver disease. This protocol describes a reproducible method for sampling breath in children and then stabilizing breath samples for further analysis with gas chromatography-mass spectrometry (GC-MS). The goal of this method is to establish a standardized protocol for the acquisition of breath samples for further chemical analysis, from children aged 4-15 years. First, breath is sampled using a cardboard mouthpiece attached to a 2-way valve, which is connected to a 3 L bag. Breath analytes are then transferred to a thermal desorption tube and stored at 4-5 &#176;C until analysis. This technique has been previously used to capture breath of children with malaria for successful breath biomarker identification. Subsequently, we have successfully applied this technique to additional pediatric cohorts. The advantage of this method is that it requires minimal cooperation on part of the patient (of particular value in pediatric populations), has a short collection period, does not require trained staff, and can be performed with portable equipment in resource-limited field settings.

This protocol describes a simple method for the acquisition of breath samples from children. Briefly, samples of mixed air are pre-concentrated in sorbent tubes prior to gas chromatography-mass spectrometry analysis. Breath biomarkers of infectious and non-infectious diseases can be identified using this breath collection method.

Breath collection and analysis can be used to discover volatile biomarkers in a number of infectious and non-infectious diseases, such as malaria, ...

Chemistry

Capturing Actively Produced Microbial Volatile Organic Compounds from Human-Associated Samples with Vacuum-Assisted Sorbent Extraction

Volatile organic compounds (VOCs) from biological samples have unknown origins. VOCs may originate from the host or different organisms from within the host&#39;s microbial community. To disentangle the origin of microbial VOCs, volatile headspace analysis of bacterial mono- and co-cultures of Staphylococcus aureus, Pseudomonas aeruginosa, and Acinetobacter baumannii, and stable isotope probing in biological samples of feces, saliva, sewage, and sputum were performed. Mono- and co-cultures were used to identify volatile production from individual bacterial species or in combination with stable isotope probing to identify the active metabolism of microbes from the biological samples.
Vacuum-assisted sorbent extraction (VASE) was employed to extract the VOCs. VASE is an easy-to-use, commercialized, solvent-free headspace extraction method for semi-volatile and volatile compounds. The lack of solvents and the near-vacuum conditions used during extraction make developing a method relatively easy and fast when compared to other extraction options such as tert-butylation and solid phase microextraction. The workflow described here was used to identify specific volatile signatures from mono- and co-cultures. Furthermore, analysis of the stable isotope probing of human associated biological samples identified VOCs that were either commonly or uniquely produced. This paper presents the general workflow and experimental considerations of VASE in conjunction with stable isotope probing of live microbial cultures.

This protocol describes the extraction of volatile organic compounds from a biological sample with the vacuum-assisted sorbent extraction method, gas chromatography coupled with mass spectrometry using the Entech Sample Preparation Rail, and data analysis. It also describes culture of biological samples and stable isotope probing.

Volatile organic compounds (VOCs) from biological samples have unknown origins. VOCs may originate from the host or different organisms from within the ...

Biochemistry

Histological Quantification to Determine Lung Fungal Burden in Experimental Aspergillosis

The quantification of lung fungal burden is critical for the determination of the relative levels of immune protection and fungal virulence in mouse models of pulmonary fungal infection. Although multiple methods are used to assess fungal burden, quantitative polymerase chain reaction (qPCR) of fungal DNA has emerged as a technique with several advantages over previous culture-based methods. Currently, a comprehensive assessment of lung pathology, leukocyte recruitment, fungal burden, and gene expression in mice with invasive aspergillosis (IA) necessitates the use of a significant number of experimental and control animals. Here the quantification of lung histological staining to determine fungal burden using a reduced number of animals was examined in detail. Lung sections were stained to identify fungal structures with Gomori&#39;s modified methanamine silver (GMS) staining. Images were taken from the GMS-stained sections from 4 discrete fields of each formalin-fixed paraffin-embedded lung. The GMS stained areas within each image were quantified using an image analysis program, and from this quantification, the mean percentage of stained area was determined for each sample. Using this strategy, eosinophil-deficient mice exhibited decreased fungal burden and disease with caspofungin therapy, while wild-type mice with IA did not improve with caspofungin. Similarly, fungal burden in mice lacking &#947;&#948; T cells were also improved by caspofungin, as measured by qPCR and GMS quantification. GMS quantification is therefore introduced as a method for the determination of relative lung fungal burden that may ultimately reduce the quantity of experimental animals required for comprehensive studies of invasive aspergillosis.

Here we describe a protocol to determine pulmonary fungal burden in mice with invasive aspergillosis by quantification of Gomori&#39;s modified methanamine silver staining in histological sections. Use of this method resulted in comparable results with less animals compared to assessment of fungal burden by quantitative PCR of lung fungal DNA.

The quantification of lung fungal burden is critical for the determination of the relative levels of immune protection and fungal virulence in mouse ...

Isolation of Culturable Yeasts and Molds from Soils to Investigate Fungal Population Structure

Soil is host to an incredible amount of microbial life, with each gram containing up to billions of bacterial, archaeal, and fungal cells. Multicellular fungi such as molds and unicellular fungi, broadly defined as yeasts, fulfill essential roles in soil ecosystems as decomposers of organic material and as food sources for other soil dwellers. Fungal species diversity in soil is dependent on a multitude of climatic factors such as rainfall and temperature, as well as soil properties including organic matter, pH, and moisture. Lack of adequate environmental sampling, especially in regions of Asia, Africa, South America, and Central America, hinders the characterization of soil fungal communities and the discovery of novel species. 
We characterized soil fungal communities in nine countries across six continents using ~4,000 soil samples and a protocol developed in the laboratory for the isolation of yeasts and molds. This protocol begins with separate selective enrichment for yeasts and the medically relevant mold Aspergillus fumigatus, in liquid media while inhibiting bacterial growth. Resulting colonies are then transferred to solid media and further processed to obtain pure cultures, followed by downstream genetic characterization. Yeast species identity is established via sequencing of their internal transcribed spacer (ITS) region of the nuclear ribosomal RNA gene cluster, while global population structure of A. fumigatus is explored via microsatellite marker analysis. 
The protocol was successfully applied to isolate and characterize soil yeast and A. fumigatus populations in Cameroon, Canada, China, Costa Rica, Iceland, Peru, New Zealand, and Saudi Arabia. These findings revealed much-needed insights on global patterns in soil yeast diversity, as well as global population structure and antifungal resistance profiles of A. fumigatus. This paper presents the method of isolating both yeasts and A. fumigatus from international soil samples.

This protocol is an effective, speedy method of culturing yeasts and the mold Aspergillus fumigatus from large sets of soil samples in as little as 7 days. The methods can be easily modified to accommodate a range of incubation media and temperatures as needed for experiments.

Soil is host to an incredible amount of microbial life, with each gram containing up to billions of bacterial, archaeal, and fungal cells. Multicellular ...

Isolation, Characterization, and Total DNA Extraction to Identify Endophytic Fungi in Mycoheterotrophic Plants

Mycoheterotrophic plants present one of the most extreme forms of mycorrhizal dependency, having totally lost their autotrophic capacity. As essential as any other vital resource, the fungi with which these plants intimately associate are essential for them. Hence, some of the most relevant techniques in studying mycoheterotrophic species are the ones that enable the investigation of associated fungi, especially those inhabiting roots and subterranean organs. In this context, techniques for identifying culture-dependent and culture-independent endophytic fungi are commonly applied. Isolating fungal endophytes provides a means for morphologically identifying them, analyzing their diversity, and maintaining inocula for applications in the symbiotic germination of orchid seeds. However, it is known that there is a large variety of non-culturable fungi inhabiting plant tissues. Thus, culture-independent molecular identification techniques offer a broader cover of species diversity and abundance. This article aims to provide the methodological support necessary for starting two investigation procedures: a culture-dependent and an independent one. Regarding the culture-dependent protocol, the processes of collecting and maintaining plant samples from collection sites to laboratory facilities are detailed, along with isolating filamentous fungi from subterranean and aerial organs of mycoheterotrophic plants, keeping a collection of isolates, morphologically characterizing hyphae by slide culture methodology, and molecular identification of fungi by total DNA extraction. Encompassing culture-independent methodologies, the detailed procedures include collecting plant samples for metagenomic analyses and total DNA extraction from achlorophyllous plant organs using a commercial kit. Finally, continuity protocols (e.g., polymerase chain reaction [PCR], sequencing) are also suggested for analyses, and techniques are presented here.

The current article aims to provide detailed and adequate protocols for the isolation of plant-associated endophytic fungi, long-term preservation of isolates, morphological characterization, and total DNA extraction for subsequent molecular identification and metagenomic analyses.

Mycoheterotrophic plants present one of the most extreme forms of mycorrhizal dependency, having totally lost their autotrophic capacity. As essential as ...

Collection and Extraction of Occupational Air Samples for Analysis of Fungal DNA

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