The authors acknowledge the Swiss National Science Foundation for Grant No. 31003A-132358/1, 31003A_152972 and No. 151948, and Fundation Pierre Mercier pour la Science.

1. Preparation of Chemicals and Equipment&#160;
<ol>
	<li>Make 500 ml of a 1% sodium hexametaphosphate (SHMP) (NaPO3)n solution and sterilize by autoclaving.</li>
	<li>Sterilize nitrocellulose (NC) filters (pore size 0.22 &#181;m, diameter 47 mm) by autoclaving in closed glass Petri dishes.</li>
	<li>Sterilize NC filters (pore size 0.22 &#181;m, diameter 25 mm) by autoclaving in closed glass Petri dishes.</li>
	<li>Weigh and note the empty weight of sterile 50 ml tubes (with cap on) (one tube per sample).</li>
	<li>Prepare Tris-EDTA-buffer (TE-buffer) 1x: make solution of 10 mM Tris (tris(hydroxymethyl)aminomethane) and 1 mM EDTA buffer. Adjust pH to 8 and sterilize by autoclaving.</li>
	<li>Prepare lysozyme solution (20 mg/ml) by dissolving 0.02 g of lysozyme in 1 ml TE-buffer. Ideally, make fresh every time. Store at 4 &#176;C for no more than 1 week.</li>
	<li>Prepare physiological solution by dissolving 8 g/L sodium chloride (NaCl) in distilled water. Sterilize by autoclaving.</li>
</ol>
2. Separation of Biomass from Sediment
<ol>
	<li>Add 3 g of sediment sample to pre-weighed sterile 50 ml tubes using ethanol-flamed metal scoops. Perform this in a clean, UV sterilized biosafety cabinet to avoid contamination.</li>
	<li>Add 15 ml of a 1% sterile (autoclaved) SHMP solution to the sample using a sterile graduated burette. The SHMP solution can also be filtered to avoid contamination.</li>
	<li>Homogenize the sediment and SHMP solution with a liquid disperser/homogenizer (e.g., Ultra-Turrax homogenizer). 70% ethanol-sterilize or autoclave the dispersion rotor prior to use. Run the homogenization for 1 min at 17,500 rpm. Let the sample rest for 2 min and repeat the homogenization for 1 min at the same rotor speed.</li>
	<li>Let the sample stand for 10 min. At this step, the heaviest particles (minerals) will settle. The cells and any organic components of the sample however will remain in solution. Afterwards, transfer the supernatant solution (containing cell biomass) into a clean 50 ml tube, while taking care not to disturb the sediment pellet.</li>
	<li>To the sediment pellet add again 15 ml of a 1% sterile (autoclaved) SHMP solution using a sterile graduated burette. Then repeat steps 2.3 and 2.4. This repetition ensures the separation of the maximum amount of cells and organic particles from the mineral component of the sediment. The supernatant of this second separation can be merged with the supernatant from the first separation. 
	Note: The following steps are all done on the supernatant (containing cell biomass). The mineral component (sediment pellet) can be discarded.</li>
	<li>Centrifuge the sample at 20 x g for 1 min. This step increases the g-force enough to settle small mineral particles while the biological cell material still remains in solution. After centrifugation, transfer the supernatant solution (containing cell biomass) into a clean 50 ml tube. Discard the mineral pellet.</li>
	<li>Determine the final volume of the solution containing biomass by weighing the sample. The weight determination avoids having to transfer the sample to a graduated cylinder and reduces risk of contamination.</li>
</ol>
3. Collection of Biomass on Filter Membrane
<ol>
	<li>Prepare filtration unit (for 47 mm diameter membranes) and vacuum pump. Sterilize the filtration unit either by autoclaving or (if Pyrex glass) by spraying it with 70% ethanol and flaming it with a Bunsen burner. Let it cool down before continuing with the protocol.</li>
	<li>Add sterile NC membrane to the filtration unit using ethanol-flamed sterilized forceps.</li>
	<li>Add half of the supernatant sample (from step 2.6) onto the membrane filtration unit and collect cells on the membrane using the vacuum pump.</li>
	<li>When the liquid has fully passed through the filter, stop the vacuum pump and carefully remove the membrane using ethanol-flame sterilized forceps. Place the membrane into a sterile Petri dish.
	<ol>
		<li>Cut the membrane in half using ethanol-flamed sterilized scissors. Add each half of the membrane to a separate 2 ml tube. One half of the membrane will be used for DNA extraction and analysis of the entire bacterial community. The other half of filter will be stored at -80 &#176;C and serves as a backup.</li>
	</ol>
	</li>
	<li>Place new NC membrane onto the filtration unit and collect biomass from the second half of sample volume (from step 2.6) using the vacuum pump.</li>
	<li>When the liquid has fully passed through the filter, stop the vacuum pump and carefully remove the membrane using ethanol-flamed sterilized forceps. Place the entire membrane into a separate 2 ml tube. This sample will be used for the treatment to separate endospores from vegetative cells. The sample can be stored at -20 &#176;C until use.</li>
</ol>
4. Lysis of Vegetative Cells
<ol>
	<li>Perform the treatment to separate endospores from vegetative cells on the biomass previously collected on a NC membrane (step 3.6).
	<ol>
		<li>If the membrane was frozen, leave it at RT for 10 min&#160;to thaw. Then place the membrane in a sterile Petri dish and cut it (approximately 4 times) into smaller pieces using ethanol-flamed sterilized scissors. Then place all membrane filter pieces into a sterile 2 ml tube.</li>
	</ol>
	</li>
	<li>Add 900 &#181;l of 1x TE (Tris-EDTA) buffer (see 1.5) to the tube containing the sample membrane and mix thoroughly by vortex. At this step, the biomass is removed from the membrane into the TE-buffer solution.</li>
	<li>Place the tube in an incubator at 65 &#176;C for 10 min and 80 rpm. Afterwards remove the tube from the incubator and let it cool down for 15 min.</li>
	<li>Add 100 &#181;l of freshly prepared lysozyme (see 1.5) to reach a final concentration of 2 mg/ml. Do not add the lysozyme before the sample has cooled down to 37 &#176;C, as this could degrade the enzyme.</li>
	<li>Incubate the sample at 37 &#176;C for 60 min and 80 rpm, the optimal conditions for the lysozyme to lyse vegetative cells.</li>
	<li>After lysis is complete, add 250 &#181;l of 3 N sodium hydroxide (NaOH) and 250 &#181;l of 6% sodium dodecyl sulfate (SDS) solution to the sample. By adding this, the sample volume reaches 1.5 ml and there is a final concentration of 0.5 N NaOH and final concentration of 1% SDS.</li>
	<li>Incubate this mix at RT for 60 min and 80 rpm. Adding the base and detergents will help in final cell lysis. The concentration of these detergents has been optimized as to not harm the endospores, while lysing vegetative cells.</li>
	<li>Prepare a sterile filtration unit that holds 25 mm diameter membranes by autoclaving, or (if Pyrex glass) spraying it with 70% ethanol and flaming it with a Bunsen burner. Let it cool down.</li>
	<li>Place a 0.2 &#181;m NC membrane (25 mm diameter) on the filtration unit using ethanol-flame sterilized forceps.</li>
	<li>Add the sample from step 4.7 onto the membrane and filter the liquid using the vacuum pump. When liquid has passed through, turn off vacuum pump. At this step, the lysed vegetative cell material is removed, as it is not retained on the membrane. Only endospores will remain on the membrane.</li>
	<li>Add 2 ml of sterile physiological solution to wash off residual detergents and filter the liquid using the vacuum pump.</li>
	<li>When liquid has fully filtered, turn off the vacuum pump. Leave the membrane on the filtration unit.</li>
</ol>
5. DNase Treatment
Note: Perform the DNase treatment directly on the filter membrane. It is important that the filtration unit does not leak and the vacuum pump is turned off.
<ol>
	<li>Add 450 &#181;l of sterile water, 50 &#181;l of DNase reaction buffer (1x) and 0.5 &#181;l DNase enzyme directly onto the filter membrane and let it stand for 15 min. If possible, do this digestion in a room that is slightly warmer than average RT, since the enzyme works better at temperatures of 25 &#176;C and above. 
	Note: Keeping the Bunsen burner aside the filtration unit also increases the temperature and has the added benefit of keeping the surroundings sterile, therefore reduced risk of contamination of the samples.</li>
	<li>When the DNase digestion is finished, turn on vacuum pump to remove the enzyme from the sample.</li>
	<li>Wash off residual enzyme by adding and filtering 1 ml of physiological solution.</li>
	<li>If liquid has fully passed, turn off vacuum pump and remove the filter membrane containing endospores using sterile forceps and place it into a sterile Petri dish. 
	Note: The sample of separated endospores is now ready for downstream analysis. Store at -20 &#176;C if used for DNA extraction or alternatively at 4&#176;C if used for germination and cultivation.</li>
</ol>

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The authors have nothing to disclose.

The resistance of endospores to external aggressive physicochemical factors (e.g., temperature or detergents) was used to devise a method to separate bacterial endospores from vegetative cells in environmental samples. This is the first comprehensive method to isolate endospores from environmental samples in a non-destructive manner. Previous methods to quantify, detect or analyze endospores in samples were based on the measurement of specific proxies for endospores such as dipicolinic acid or specific marker ge...

The goal of this work is to provide a protocol for the separation of bacterial endospores from vegetative bacterial cells in environmental samples. The formation of bacterial endospores is a survival strategy, usually triggered by starvation, found in a number of bacterial groups belonging to the phylum Firmicutes1. Endospore-forming bacteria are well studied, mainly because a number of strains are pathogens and hence of medical importance (e.g., Bacillus anthracis or Clostridium difficile). Environmental strains of endospore-forming bacteria have been isolated from virtually every environment (soil, water, sediment, air, ice, human gut, animals gut, and more)1-3. Therefore, Firmicutes are the second most abundant phylum in culture collections4.
Because of their hardy outer cortex and protective core proteins, endospores can survive extreme environmental conditions ranging from desiccation to high radiation, extreme temperatures and harmful chemicals5. This remarkable resistance makes it a challenge to extract DNA from endospores6-8. This likely explains why they have been overlooked in environmental sequencing studies9,10. Other methods, such as targeting of endospores in environmental samples by fluorescent antibodies11, quantification of dipicolinic acid (DPA) in soil12 and sediment13, flow cytometry14 or pasteurization and subsequent cultivation15,16 have been used to retrieve or quantify endospores in environmental samples. In recent years, optimized DNA extraction methods as well as specific molecular primers to target endospore-specific gene sequences have been developed10,17-20. This has helped to reveal more biodiversity among this group of bacteria21 and has also led to applications in industry and medicine for the detection of endospores, for example in milk powder19.
The protocol presented here is based on the difference in resistance to harmful physicochemical conditions (such as heat and detergents) of bacterial endospores relative to vegetative cells. To destroy vegetative cells in a sample, we consecutively apply heat, lysozyme and low concentrations of detergents. The time and strength of these treatments have been optimized so as not to destroy spores, but to lyse all vegetative cells. Some cells in an environmental cell pool are more resistant than others, so in order to increase the probability of destroying all vegetative cells, we apply three different treatments. The advantage and novelty of this method is that the endospores after the treatment are still intact and can be used for further downstream analyses. These include DNA extraction, quantitative PCR (qPCR) and amplicon or metagenomic sequencing (targeting specifically the group of endospores and thus reducing diversity, while increasing coverage). The endospores could also be used for downstream cultivation or quantification by fluorescence microscopy, flow cytometry, or detection of DPA. An important feature of this method is that by comparing an untreated sample with a treated sample, one can deduce the quantity and diversity of endospores in an environmental sample in addition to the component corresponding to vegetative cells.

<table><tbody><tr><td>Whatman nitrocellulose membrane filters, 0.2 &amp;#956;m pore size, 47 mm diameter</td><td>Sigma-Aldrich</td><td>WHA7182004 Aldrich&amp;nbsp;</td><td></td></tr><tr><td>Tris(hydroxymethyl)aminomethane</td><td>Sigma-Aldrich</td><td>252859 Sigma-Aldrich</td><td>CAS 77-86-1</td></tr><tr><td>EDTA</td><td>Sigma-Aldrich</td><td>E9884 Sigma-Aldrich</td><td>CAS&amp;nbsp; 60-00-4&amp;nbsp;</td></tr><tr><td>Lysozyme from chicken egg white</td><td>Sigma-Aldrich</td><td>62971 Fluka</td><td>powder, CAS&amp;nbsp; 12650-88-3&amp;nbsp;</td></tr><tr><td>Ultra-Turrax homogenizer T18 basic</td><td>IKA</td><td>3720000</td><td></td></tr><tr><td>Glass filter holders for 47&amp;nbsp;mm membranes, Pyrex glass</td><td>EMD Millipore</td><td>XX10 047 00</td><td></td></tr><tr><td>Chemical duty vacuum pump</td><td>Millipore</td><td>WP6122050</td><td>220 V/50 Hz</td></tr><tr><td>Manifold sampling filtration for 25&amp;nbsp;mm membranes</td><td>Millipore</td><td>1225 Sampling Manifold</td><td>polypropylene</td></tr><tr><td>Dnase</td><td>New England Biolabs</td><td>M0303S</td><td>Rnase free</td></tr><tr><td>NaOH</td><td>Sigma-Aldrich</td><td>S5881 Sigma-Aldrich</td><td>CAS&amp;nbsp; 1310-73-2&amp;nbsp;</td></tr><tr><td>Sodium dodecyl sulfphate (SDS)</td><td>Sigma-Aldrich</td><td>L3771 Sigma&amp;nbsp;</td><td></td></tr><tr><td>Whatman nitrocellulose membrane filters, 0.2 &amp;#956;m pore size, 25 mm diameter</td><td>Sigma-Aldrich</td><td>WHA7182002 Aldrich</td><td></td></tr></tbody></table>

physical isolation of endospores from environmental samples by targeted lysis of vegetative cells

Endospore formation is a survival strategy found among some bacteria from the phylum Firmicutes. During endospore formation, these bacteria enter a morpho-physiological resting state that enhances survival under adverse environmental conditions. Even though endospore-forming Firmicutes are one of the most frequently enriched and isolated bacterial groups in culturing studies, they are often absent from diversity studies based on molecular methods. The resistance of the spore core is considered one of the factors limiting the recovery of DNA from endospores. We developed a method that takes advantage of the higher resistance of endospores to separate them from other cells in a complex microbial community using physical, enzymatic and chemical lysis methods. The endospore-only preparation thus obtained can be used for re-culturing or to perform downstream analysis such as tailored DNA extraction optimized for endospores and subsequent DNA sequencing. This method, applied to sediment samples, has allowed the enrichment of endospores and after sequencing, has revealed a large diversity of endospore-formers in freshwater lake sediments. We expect that the application of this method to other samples will yield a similar outcome.

The results presented here have been published earlier10,21. Please refer to those articles for the environmental interpretation and discussion of the data.
The overall procedure is summarized in Figure 1 and corresponds to three main steps: first, the separation of biomass from sediment or any other environmental matrix; second, the destruction of vegetative cells; and third, the downstream analysis of the separated endospores. Downstream analysis could consist, fo...

Watch this Scientific Journal Video about Physical Isolation of Endospores from Environmental Samples by Targeted Lysis of Vegetative Cells at JoVE.com

Physical Isolation of Endospores from Environmental Samples by Targeted Lysis of Vegetative Cells

A method to single out bacterial endospores from complex microbial communities was developed to perform tailored culture or molecular studies of this group of bacteria.

Endospore formation is a survival strategy found among some bacteria from the phylum Firmicutes. During endospore formation, these bacteria enter a ...

physical-isolation-endospores-from-environmental-samples-targeted

Universidad de Cartagena UDEC

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JoVE Journal

Environment

14.1K Views.  Macquarie University. A method to single out bacterial endospores from complex microbial communities was developed to perform tailored culture or molecular studies of this group of bacteria.

Video: Physical Isolation of Endospores from Environmental Samples by Targeted Lysis of Vegetative Cells

Culturing and Maintaining Clostridium difficile in an Anaerobic Environment

Clostridium difficile is a Gram-positive, anaerobic, sporogenic bacterium that is primarily responsible for antibiotic associated diarrhea (AAD) and is a significant nosocomial pathogen. C. difficile is notoriously difficult to isolate and cultivate and is extremely sensitive to even low levels of oxygen in the environment. Here, methods for isolating C. difficile from fecal samples and subsequently culturing C. difficile for preparation of glycerol stocks for long-term storage are presented. Techniques for preparing and enumerating spore stocks in the laboratory for a variety of downstream applications including microscopy and animal studies are also described. These techniques necessitate an anaerobic chamber, which maintains a consistent anaerobic environment to ensure proper conditions for optimal C. difficile growth. We provide protocols for transferring materials in and out of the chamber without causing significant oxygen contamination along with suggestions for regular maintenance required to sustain the appropriate anaerobic environment for efficient and consistent C. difficile cultivation.

Culturing and Maintaining Clostridium difficile in an Anaerobic Environment

Clostridium difficile is a pathogenic bacterium that is a strict anaerobe and causes antibiotic associated diarrhea (AAD). Here, methods for isolating, culturing and maintaining C. difficile vegetative cells and spores are described. These techniques necessitate an anaerobic chamber, which requires regular maintenance to ensure proper conditions for optimal C. difficile cultivation.

Clostridium  difficile is a Gram-positive, anaerobic, sporogenic bacterium  that is primarily responsible for antibiotic associated diarrhea (AAD) and is ...

Immunology and Infection

Isolation, Propagation, and Identification of Bacterial Species with Hydrocarbon Metabolizing Properties from Aquatic Habitats

Hydrocarbon pollutants are recalcitrant to degradation and their accumulation in the environment is toxic to all life forms. Bacteria encode numerous catalytic enzymes and are naturally capable of metabolizing hydrocarbons. Scientists harness biodiversity in aquatic ecosystems to isolate bacteria with biodegradation and bioremediation potential. Such isolates from the environment provide a rich set of metabolic pathways and enzymes, which can be further utilized to scale up the degradation process at an industrial scale. In this article, we outline the general process of isolation, propagation, and identification of bacterial species from aquatic habitats and screen their ability to utilize hydrocarbons as the sole carbon source in vitro using simple techniques. The present protocol describes the isolation of various bacterial species and their subsequent identification using the 16S rRNA analysis. The protocol also presents steps for characterizing the hydrocarbon degrading potential of bacterial isolates. This protocol will be useful for researchers trying to isolate bacterial species from environmental habitats for their biotechnological applications.

We present the process of isolating, propagating, and characterizing hydrocarbon-degrading bacteria from aquatic habitats. The protocol outlines bacterial isolation, identification by the 16S rRNA method, and testing of their hydrocarbon-degrading potential. This article would help researchers in characterizing microbial biodiversity in environmental samples, and specifically screen for microbes with bioremediation potential.

Hydrocarbon pollutants are recalcitrant to degradation and their accumulation in the environment is toxic to all life forms. Bacteria encode numerous ...

Detecting Cortex Fragments During Bacterial Spore Germination

The process of endospore germination in Clostridium difficile, and other Clostridia, increasingly is being found to differ from the model spore-forming bacterium, Bacillus subtilis. Germination is triggered by small molecule germinants and occurs without the need for macromolecular synthesis. Though differences exist between the mechanisms of spore germination in species of Bacillus and Clostridium, a common requirement is the hydrolysis of the peptidoglycan-like cortex which allows the spore core to swell and rehydrate. After rehydration, metabolism can begin and this, eventually, leads to outgrowth of a vegetative cell. The detection of hydrolyzed cortex fragments during spore germination can be difficult and the modifications to the previously described assays can be confusing or difficult to reproduce. Thus, based on our recent report using this assay, we detail a step-by-step protocol for the colorimetric detection of cortex fragments during bacterial spore germination.

Herein, we describe a colorimetric assay to detect the presence of reducing sugars during bacterial spore germination.

The process of endospore germination in Clostridium difficile, and other Clostridia, increasingly is being found to differ from the model spore-forming ...

Visualization of Germinosomes and the Inner Membrane in Bacillus subtilis Spores

The small size of spores and the relatively low abundance of germination proteins, cause difficulties in their microscopic analyses using epifluorescence microscopy. Super-resolution three-dimensional Structured Illumination Microscopy (3D-SIM) is a promising tool to overcome this hurdle and reveal the molecular details of the process of germination of Bacillus subtilis (B. subtilis) spores. Here, we describe the use of a modified SIMcheck (ImageJ)-assistant 3D imaging process and fluorescent reporter proteins for SIM microscopy of B. subtilis spores&#8217; germinosomes, cluster(s) of germination proteins. We also present a (standard)3D-SIM imaging procedure for FM4-64 staining of B. subtilis spore membranes. By using these procedures, we obtained unsurpassed resolution for germinosome localization and show that &#62;80% of B. subtilis KGB80 dormant spores obtained after sporulation on defined minimal MOPS medium have one or two GerD-GFP and GerKB-mCherry foci. Bright foci were also observed in FM4-64 stained spores&#8217; 3D-SIM images suggesting that inner membrane lipid domains of different fluidity likely exist. Further studies that use double labeling procedures with membrane dyes and germinosome reporter proteins to assess co-localization and thus get an optimal overview of the organization of Bacillus germination proteins in the inner spore membrane are possible.

Visualization of Germinosomes and the Inner Membrane in Bacillus subtilis Spores

Germinant receptor proteins cluster in &#8216;germinosomes&#8217; in the inner membrane of Bacillus subtilis spores. We describe a protocol using super resolution microscopy and fluorescent reporter proteins to visualize germinosomes. The protocol also identifies spore inner membrane domains that are preferentially stained with the membrane dye FM4-64.

The small size of spores and the relatively low abundance of germination proteins, cause difficulties in their microscopic analyses using epifluorescence ...

Bioengineering

Investigating the Detrimental Effects of Low Pressure Plasma Sterilization on the Survival of Bacillus subtilis Spores Using Live Cell Microscopy

Plasma sterilization is a promising alternative to conventional sterilization methods for industrial, clinical, and spaceflight purposes. Low pressure plasma (LPP) discharges contain a broad spectrum of active species, which lead to rapid microbial inactivation. To study the efficiency and mechanisms of sterilization by LPP, we use spores of the test organism Bacillus subtilis because of their extraordinary resistance against conventional sterilization procedures. We describe the production of B. subtilis spore monolayers, the sterilization process by low pressure plasma in a double inductively coupled plasma reactor, the characterization of spore morphology using scanning electron microscopy (SEM), and the analysis of germination and outgrowth of spores by live cell microscopy. A major target of plasma species is genomic material (DNA) and repair of plasma-induced DNA lesions upon spore revival is crucial for survival of the organism. Here, we study the germination capacity of spores and the role of DNA repair during spore germination and outgrowth after treatment with LPP by tracking fluorescently-labelled DNA repair proteins (RecA) with time-resolved confocal fluorescence microscopy. Treated and untreated spore monolayers are activated for germination and visualized with an inverted confocal live cell microscope over time to follow the reaction of individual spores. Our observations reveal that the fraction of germinating and outgrowing spores is dependent on the duration of LPP-treatment reaching a minimum after 120 s. RecA-YFP (yellow fluorescence protein) fluorescence was detected only in few spores and developed in all outgrowing cells with a slight elevation in LPP-treated spores. Moreover, some of the vegetative bacteria derived from LPP-treated spores showed an increase in cytoplasm and tended to lyse. The described methods for analysis of individual spores could be exemplary for the study of other aspects of spore germination and outgrowth.

Investigating the Detrimental Effects of Low Pressure Plasma Sterilization on the Survival of Bacillus subtilis Spores Using Live Cell Microscopy

This protocol illustrates the important consecutive steps required to assess the relevance of monitoring vitality parameter and DNA repair processes in reviving Bacillus subtilis spores after treatment with low pressure plasma by tracking fluorescence-labelled DNA repair proteins via time-resolved confocal microscopy and scanning electron microscopy.

Plasma sterilization is a promising alternative to conventional sterilization methods for industrial, clinical, and spaceflight purposes. Low pressure ...

Biology

Investigating Teliospore Germination Using Microrespiration Analysis and Microdissection

Smut fungi are the etiological agents of several devastating agricultural diseases. They are characterized by the production of teliospores, which are thick-walled dispersal agents. Teliospores can remain dormant for decades. The dormancy is characterized by low metabolic rates, paused macromolecular biosynthesis and greatly reduced levels of respiration. Upon receiving required environmental signals, teliospores germinate to produce haploid cells, which can initiate new rounds of infection. Teliospore germination is characterized by the resumption of macromolecular biosynthesis, increased respiration and dramatic morphological changes. In order to precisely measure changes in cellular respiration during the early stages of germination, we have developed a simple protocol employing a Clark-type respirometer. The later stages of germination are distinguished by specific morphological changes, but germination is asynchronous. We developed a microdissection technique that enables us to collect teliospores at distinct germination stages.

Smut fungi cause many devastating agricultural diseases. They are dispersed as dormant teliospores that germinate in response to environmental cues. We outline two methods to investigate molecular changes during germination: measuring respiration increase to detect metabolic activation and assessing changing molecular events by isolating teliospores at distinct morphological stages.

Smut fungi are the etiological agents of several devastating agricultural diseases. They are characterized by the production of teliospores, which are ...

Isolation and Selection of Entomopathogenic Fungi from Soil Samples and Evaluation of Fungal Virulence against Insect Pests

Entomopathogenic fungi (EPF) are one of the microbial control agents for integrated pest management. To control local or invasive pests, it is important to isolate and select indigenous EPF. Therefore, the soil bait method combined with the insect bait (mealworm, Tenebrio molitor) system was used in this study with some modifications. The isolated EPF were then subjected to the virulence test against the agricultural pest Spodoptera litura. Furthermore, the potential EPF strains were subjected to morphological and molecular identifications. In addition, the conidia production and thermotolerance assay were performed for the promising EPF strains and compared; these data were further substituted into the formula of effective conidia number (ECN) for laboratory ranking. The soil bait-mealworm system and the ECN formula can be improved by replacing insect species and integrating more stress factors for the evaluation of commercialization and field application. This protocol provides a quick and efficient approach for EPF selection and will improve the research on biological control agents.

Here we present a protocol based on the&#160;mealworm (Tenebrio molitor)-bait system that was used for isolating and selecting entomopathogenic fungi (EPF) from soil samples. An effective conidia number (ECN) formula is used to select high stress tolerant EPF based on physiological characteristics for pest microbial control in the field.

Entomopathogenic fungi (EPF) are one of the microbial control agents for integrated pest management. To control local or invasive pests, it is important ...

Comparison of Methods for Isolating Entomopathogenic Fungi from Soil Samples

The goal of the present study is to compare the effectiveness of using insect baits versus artificial selective medium for isolating entomopathogenic fungi (EPF) from soil samples. The soil is a rich habitat for microorganisms, including EPF particularly belonging to the genera Metarhizium and Beauveria, which can regulate arthropod pests. Biological products based on fungi are available in the market mainly for agricultural arthropod pest control. Nevertheless, despite the high endemic biodiversity, only a few strains are used in commercial bioproducts worldwide. In the present study, 524 soil samples were cultured on potato dextrose agar enriched with yeast extract supplemented with chloramphenicol, thiabendazole, and cycloheximide (CTC medium). The growth of fungal colonies was observed for 3 weeks. All Metarhizium and Beauveria EPF were morphologically identified at the genus level. Additionally, some isolates were molecularly identified at the species level. Twenty-four out of these 524 soil samples were also surveyed for EPF occurrence using the insect bait method (Galleria mellonella and Tenebrio molitor). A total of 51 EPF strains were isolated (41 Metarhizium spp. and 10 Beauveria spp.) from the 524 soil samples. All fungal strains were isolated either from croplands or grasslands. Of the 24 samples selected for comparison, 91.7% were positive for EPF using Galleria bait, 62.5% using Tenebrio bait, and 41.7% using CTC. Our results suggested that using insect baits to isolate the EPF from the soil is more efficient than using the CTC medium. The comparison of isolation methods in addition to the identification and conservation of EPF has a positive impact on the knowledge about biodiversity. The improvement of EPF collection supports scientific development and technological innovation.

Entomopathogenic fungal colonies are isolated from tropical soil samples using Tenebrio bait, Galleria bait, as well as selective artificial medium, i.e., potato dextrose agar enriched with yeast extract supplemented with chloramphenicol, thiabendazole, and cycloheximide (CTC medium).

The goal of the present study is to compare the effectiveness of using insect baits versus artificial selective medium for isolating entomopathogenic ...

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 ...

Physical Isolation of Endospores from Environmental Samples by Targeted Lysis of Vegetative Cells

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Culturing and Maintaining Clostridium difficile in an Anaerobic Environment

Isolation, Propagation, and Identification of Bacterial Species with Hydrocarbon Metabolizing Properties from Aquatic Habitats

Detecting Cortex Fragments During Bacterial Spore Germination

Visualization of Germinosomes and the Inner Membrane in Bacillus subtilis Spores

Investigating the Detrimental Effects of Low Pressure Plasma Sterilization on the Survival of Bacillus subtilis Spores Using Live Cell Microscopy

Investigating Teliospore Germination Using Microrespiration Analysis and Microdissection

Isolation and Selection of Entomopathogenic Fungi from Soil Samples and Evaluation of Fungal Virulence against Insect Pests

Comparison of Methods for Isolating Entomopathogenic Fungi from Soil Samples

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

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