The authors would like to acknowledge Otterbein University for Undergraduate Student Research Fellowships and Student Research Fund Awards to GVK and SGK; and the Otterbein Professor Gilbert E. Mills Memorial Endowed Sabbatical Program Award and The Department of Biology and Earth Science Faculty Research and Scholarship Endowed Award to JAB. The Bert and Jane Horn Endowed Student Research Fund in the Sciences was awarded to GVK and SGK. The authors would also like to gratefully acknowledge Duquesne University funded Undergraduate Research Program Fellowships for GVK and SGK.&#160;Former Juniata College undergraduate research students, Ryan Johnson and Lindsey Draper contributed the microscopy images for Figures 2 and 3, respectively.

1. Prepare Instruments, Culture Media, Solutions, and Petri Dishes
<ol>
	<li>Autoclave toothpicks in a 50 mL beaker covered with aluminum foil to keep sterile.</li>
	<li>Sterilize all dry goods that will be used (tips, sticks, glassware, etc.) in an autoclave. 
	NOTE: Caution! Follow all safety directions for the autoclave model used. Autoclaves pose a risk for explosions, and burns from steam and contact with hot surfaces, materials, and media. For autoclave safety, view &#34;Proper Use of Autoclaves&#34;34.</li>
	<li>Prepare the MS agar (Mannitol Soya Flour (SFM))14 to grow cultures.
	<ol>
		<li>Add 5 g of soya flour and 5 g of agar to 1 L flask.</li>
		<li>Dissolve 5 g of mannitol in 250 mL of tap water and dispense into the flask containing the soy flour and agar.</li>
		<li>Add a foam plug and foil to the opening of the flask and autoclave at 121 &#176;C, 15 psi for at least 30 min. 
		CAUTION: This medium easily boils over. Use a flask that can hold at least four times the actual volume of MS agar being autoclaved. A standard pressure cooker can be used in place of an autoclave with similar safety considerations. Alternatively, a microwave oven may be used to sterilize media and materials if access to an autoclave is not possible35,36. High school teachers may also want to collaborate with local colleges or universities to access autoclaved supplies.</li>
		<li>Cool media until comfortable enough to handle. Be careful to not cool it for too long because agar solidifies at temperatures below 50 &#176;C.</li>
		<li>Pour MS agar from the flask into sterile Petri dishes until the bottom of each Petri dish is approximately half full (approximately 25 mL of media per Petri dish). Allow to solidify before using agar plates. Store agar plates in a plastic bag at room temperature until needed. 
		NOTE: Agar plates may also be stored in a refrigerator, especially if antibiotics are needed in the media.</li>
	</ol>
	</li>
</ol>
2. Streak Streptomyces onto Plates for Propagation
<ol>
	<li>Streak Streptomyces strains onto MS agar plates to single colonies using a standard method, such as the quadrant streak method as shown in Sanders, 201237. 
	NOTE: An inoculating loop or sterile toothpicks may be used. Optional: Other media are commonly used for Streptomyces species, such as R2YE and minimal media14.</li>
	<li>Incubate plates at 30 &#176;C.</li>
	<li>Restreak plates by picking a single colony every 4&#8211;6 days. As colonies age they become prone to random mutation.</li>
</ol>
3. Create Glycerol Mycelial Stocks for Bacterial Storage
NOTE: Mycelial stocks may be created for all Streptomyces strains, including whi and bld strains, and other developmental mutants that are unable to complete sporulation.
<ol>
	<li>Macerate 15&#8211;20 colonies in 200&#8211;500 &#181;L of 20% glycerol using a sterile wooden dowel applicator.</li>
	<li>Transfer the slurry of macerated colonies to a 1.5 mL freezer culture tube, using a micropipette or sterile transfer pipette. 
	NOTE: The basics of micropipette use are covered elsewhere38.</li>
	<li>Add enough 20% glycerol to make the final volume 1 mL and gently mix. Store samples at -80 &#176;C. 
	NOTE: Caution! Use cryogenic safety gloves and a lab coat to protect skin from contact with extreme cold. Alternatively, a -20 &#176;C freezer may be used with potential reduction in long term storage time. Avoid excessive freezing and thawing to extend viability.</li>
</ol>
4. Create Glycerol Spore Stocks for Strains Capable of Sporulation
NOTE: Spore stocks are preferable for long-term viability but are only feasible for strains that are capable of completing sporulation.
<ol>
	<li>Spread 100 &#181;L of macerated material (from step 3.1) on an agar plate to obtain a confluent lawn of growth37. 
	NOTE: Spread plating is shown in Sanders37 (Modification: A turntable is not required.). The plate can be turned with one hand while using the other hand to move the spreader in a gentle back and forth motion across the agar media.
	<ol>
		<li>Pipet 100 &#181;L of macerated mycelium to the center of an MS agar plate.</li>
		<li>Sterilize a glass or metal spreading tool by partially submerging in 70% ethanol and passing the spreader once slowly through a Bunsen burner flame to speed the evaporation of the ethanol. 
		CAUTION: Ethanol is highly flammable. Do not place the flaming spreader into ethanol or allow the flaming drop of ethanol to fall into the ethanol dish. Alternatively, use a disposable sterile cotton swab to spread the bacteria on the plate, which requires no ethanol or flame.</li>
		<li>Turn the MS agar plate with one hand, while using a back and forth motion to inoculate the plate with the sterile spreader.</li>
		<li>Incubate for 5&#8211;7 days at 30 &#176;C until the strain is well-sporulated. 
		NOTE: Incubation times will vary depending upon the species and strain.</li>
	</ol>
	</li>
	<li>Add 2 mL of sterile saline (0.8% NaCl) to the center of a confluent lawn of Streptomyces cells that has undergone sporulation.</li>
	<li>Harvest spores by gently rubbing the surface of the mycelium with a sterile inoculating loop (or a cotton bud), slowly moving towards the edge of the lawn.</li>
	<li>Pipet the saline containing spores into a 15 mL conical tube. Vortex vigorously to break spore chains into individual cells.</li>
	<li>Centrifuge at room temperature for 5 min at approximately 2,500 x g.</li>
	<li>Pipet the supernatant and discard. Disperse the spore pellet by vigorous vortex agitation in the small amount of remaining liquid.</li>
	<li>Resuspend the spores in 1 mL of 20% glycerol by drawing the solution up and down with a pipette.</li>
	<li>Transfer the glycerol spore stock to a 1.5 mL freezer tube. Store at -80 &#176;C. 
	CAUTION: Use cryogenic safety gloves and a lab coat to protect skin from contact with extreme cold. Alternatively, a -20 &#176;C freezer may be used with potential reduction of viability in long term storage time. Avoid excessive freezing and thawing to extend viability.</li>
</ol>
5. Perform Mutagenesis
<ol>
	<li>Perform mutagenesis based on the desired outcomes as referenced in the Introduction and recently reviewed by Baltz, 201639. 
	NOTE: Some phenotyping experiments will not require mutagenesis, such as the intent to identify new species from the environment. Methods of random mutagenesis include the use of a transposon8,9,10,11,12,13, UV radiation, or chemicals14. Site-directed mutagenesis15,16,17,18,19,20 techniques can be used to create specific point mutations, deletions, or other types of mutations.</li>
</ol>
6. Compare and Record the Visual Appearance
<ol>
	<li>Take note of colony morphology for each mutant as compared to the wild-type parent strain after streaking all strains as in step 2. Compare the new isolate to that of a well-studied species, such as S. coelicolor or S. venezuelae, when characterizing new Streptomyces species. Note characteristics (Figure 1) such as basic shape, surface of colony (fuzzy, bald, wrinkled, etc.), opacity, elevation, and pigmentation (distinguish between vegetative mycelium, aerial mycelium, and/or surrounding medium).</li>
	<li>Label an MS agar plate and streak wild-type and mutant strains in wedge patterns (Figure 1). Be careful that no strain touches another strain to avoid cross contamination. Note the date and time that strains are streaked onto the plate. Incubate the plates at 30 &#176;C.</li>
	<li>Place grown Petri dishes on colored or white paper to homogenize the background. Write on the paper the strain name, date, incubation temperature, and time from first plate streaking. Take digital picture(s) of the plate with the latter information written on the paper background so that confusion is minimized. 
	NOTE: The writing can be cropped from the photograph at a later time if it is to be used for figure preparation.</li>
</ol>
7. Perform Phase-Contrast Microscopy
<ol>
	<li>Sterilize tweezers by washing in 70% ethanol and then passing through a Bunsen burner flame to evaporate the ethanol. Allow to cool.</li>
	<li>Sterilize coverslips by washing in 70% ethanol and then allowing them to dry in a sterile Petri dish.</li>
	<li>Prepare a coverslip lift of the bacterial growth to be examined.
	<ol>
		<li>Pick up a sterile coverslip with sterile tweezers and place the coverslip on an MS agar plate where bacterial growth is dense. Press on the back of coverslip gently with tweezers to ensure sufficient transfer of bacterial spores and aerial mycelium.</li>
	</ol>
	</li>
	<li>Pick up the coverslip from the plate and place it so that the side with the cell material is facing toward the surface of a microscope slide, with a 15 &#181;L drop of 50% glycerol prepared in water for mounting. Reduce air bubbles by placing the coverslip at a 45&#176; angle to the slide and let it fall onto the 50% glycerol.</li>
	<li>(Optional) Seal with clear nail polish for preservation of the slide.</li>
	<li>Perform phase-contrast microscopy as described by Frohlich40 at various time intervals during the life cycles. 
	NOTE: Repeated imaging allows for a more complete analysis and the ability to detect delays in development. Independently repeat the observations to ensure accuracy and reproducibility.
	<ol>
		<li>Place the prepared slide on the microscope stage and add a drop of immersion oil to the center of the cover slip. Rotate the 100X phase objective in place and set the condenser turret to the proper &#34;matching&#34; phase setting. Focus the image using only the fine adjustment knob once the objective lens is in contact with the oil.</li>
		<li>Examine several fields of view to discern noticeable and consistent differences between mutant strains and the wild type, such as the ability to form spores, spore size and shape, and overall number of spores (see Figure 2). 
		NOTE: Alternatives to phase-contrast microscopy include differential interference contrast (DIC) microscopy40 and simple staining techniques for use with bright field microscopes41, such as crystal violet staining.</li>
	</ol>
	</li>
</ol>
8. Perform Fluorescence Microscopy
<ol>
	<li>Sterilize tweezers by washing in 70% ethanol and then running through a Bunsen burner flame.</li>
	<li>Pick up a sterile coverslip with sterile tweezers and place on an MS agar plate where bacterial growth is evident. Touch the back of the coverslip gently with the tweezers to ensure sufficient transfer of bacterial spores and aerial filaments.</li>
	<li>Pick up the coverslip from the plate and place it so that the side with the cell material is facing up, on cardstock for ease in manipulation of the coverslip.</li>
	<li>Flood the coverslip with ice-cold methanol (~300 &#181;L for a 20 x 20 mm2 coverslip) and let it completely air dry. 
	CAUTION: Methanol is a mutagen. Avoid direct contact with skin.</li>
	<li>Wash the coverslip three times in Phosphate Buffered Saline (PBS)42 by gently dispensing and removing the solution.</li>
	<li>Add 15 &#181;L of 100 &#956;g/ml Propidium Iodide and/or 10 &#956;g/mL Wheat-Germ-Agglutinin conjugated to fluorescein isothiocyanate (WGA-FITC) made in 50% glycerol to a labeled microscope slide.&#160; 
	NOTE: Propidium iodide stains DNA red to detect chromosomal DNA location while WGA-FITC stains the cell wall green.</li>
	<li>Invert coverslip face down onto the drop of fluorescent stain on the glass slide. (Optionally) seal with clear nail polish for preservation of slide.&#160; As the dyes are light-sensitive, incubate for 15 minutes in a darkened or dimly lit room.</li>
	<li>Observe the slides under a 100X objective lens using a phase-contrast or DIC microscope equipped with epifluorescence excitation cubes for propidium iodide (&#39;Texas Red&#39; Filter Set or 535/617 nm excitation/emission maxima) and FITC (FITC filter set or 490/525 nm excitation/emission maxima).</li>
	<li>Place the slide on the stage and focus using the coarse and fine adjustment knobs.</li>
	<li>Observe the fluorescence as shown elsewhere43.</li>
	<li>Use a free image analysis software package that can help in the identification of spores with lower fluorescence intensity44.</li>
</ol>

The authors have nothing to disclose.

Here we present protocols for beginning Streptomyces researchers to initiate studies by including the steps needed to propagate strains and prepare stocks for long-term storage. We then describe the protocols for visual and microscopic characterization of Streptomyces strains. Some typical initial steps in phenotyping developmental mutants are: 1) visual examination of the mutant colonies compared to wild type colonies on agar medium; 2) phase-contrast microscopy; and 3) fluorescence microscopy of sporogenic aerial hyphae. Based upon the phenotype displayed in these three steps, a variety of techniques may be employed to further discern the phenotype of a particular strain.
Initial phenotyping experiments are commonly used to characterize new species, identify mutants of interest, partially characterize mutants, and begin to discern the typical role of a particular gene based on the phenotype of an identified mutant. The methods described here have already been used in university teaching laboratories to identify and characterize a wide variety of Streptomyces mutants, including those with defects in cell division48,49,50,51,52,53,54, sporulation55,56, aerial mycelium formation57,58, antibiotic production59, second messenger signaling47, and chromosome segregation60. These techniques are the vital first steps to determining the phenotype of mutants in general and reveal a large amount of important information about the roles of genes of interest. The methods may easily be extended to other species of Streptomyces and have already been used to describe strains of S. griseus, S. venezuelae, S. scabies, and many other streptomycetes. The video protocols described here are expected to serve as an important resource for new researchers entering fields of Streptomyces research, such as in the area of drug discovery. This includes the new instructors who are working to combat the antibiotic resistance crisis and educating the countless numbers of new undergraduate student researchers joining the crowdsourcing efforts of the Small World Initiative.
The techniques described here can be easily adapted to college and high school classroom use in addition to research laboratories, using the modifications described in the video and text. Students in a first year microscopy module at a small liberal arts college were able to streak strains, take digital photographs of strains grown on agar media, and perform phase-contrast and fluorescence microscopy, which culminated in the submission of a portfolio of multi-paneled figures at the end of the 3 week module, representing 15 h of in-lab work. Approximately 160 first year students were responsible for the initial phenotyping of 320 novel transposon mutants. Undergraduate research students at three institutions participated in the initial phenotyping of additional mutants and the subsequent characterization of many of the strains. The comprehensive data obtained in a relatively short period of time, illustrate the value of the protocols described here. Hundreds of additional mutants have been stored as glycerol mycelial stocks for future characterization.
Following the initial experiments described here, a variety of methods may be employed to extend the quality of information pertaining to strains of interest. If the mutation is unknown, a genotyping method should be employed to determine the type and/or location of the mutation. For example, random transposon mutagenesis of the wild-type chromosome6,7,8,9,10,11,12,13 results in colonies that should undergo initial phenotypic screens such as those described above. Then the location of the transposon should be identified using a technique such as inverse polymerase chain reaction (iPCR)61. Determining the genotype of newly discovered mutants is an important step following initial characterization.
Some commonly used advanced methods for subsequent phenotyping analysis that may be mentioned in the classroom or explored through further research include green fluorescent protein (GFP) tagging to determine localization patterns for the protein of interest, gene expression analyses such as real time quantitative PCR (qPCR) and global gene expression patterns of wild-type versus mutant via RNA Sequencing (RNA-seq). Phenotyping skills are also required for genetic complementation analysis. In a complementation experiment, the wild-type copy of a gene is introduced into a mutated strain to determine whether the newly added allele can compensate for the loss-of-function of the mutated allele. Comparing the phenotype of the complemented strain to that of both the original mutant and the parental, wild-type strain is required.

Streptomycetes are Gram-positive, filamentous soil bacteria known for their ability to produce a variety of secondary metabolites, including over two-thirds of the commercially available antibiotics, as well as anti-tumor, anti-HIV, and anti-parasitic drugs1. S. coelicolor is the most genetically characterized member of the genus2,3 and is the species used in the methods described here. S. coelicolor has a complex life cycle that begins with germination of a single spore, progressing to an extensive, branching vegetative mycelium that grows into the agar medium. As the life cycle proceeds, aerial filaments are formed that break the surface tension of the substrate mycelium and are finally divided into long chains of cells that are ultimately converted into mature, grey-pigmented spores. Dispersion of these newly formed spores constitutes the beginning of the next life cycle4.
Because of its complex pattern of differentiation, S. coelicolor serves as an excellent model for the study of bacterial development. Historically, mutations result in blocks for two major stages of development and produce distinct visual phenotypes. The bld (bald) mutants are blocked for aerial mycelium formation and result in the lack of a fuzzy aerial mycelium imparting a &#34;bald&#34; colony appearance. Mutants inhibited in spore formation and maturation are referred to as whi (white) mutants because they typically fail to produce wild-type levels of the gray spore pigment and the aerial mycelium remains white. Other interesting mutants are inhibited in antibiotic production, cell division, chromosome segregation, or other important processes4,5.
Despite the discovery of many developmental genes in Streptomyces species, there are many more believed to exist based on the lack of saturation of mutant screens. Our laboratories continue to identify new developmental genes using a mini-transposon system that we have constructed. Novel mutant strains isolated in random mutagenesis experiments with our transposon undergo phenotypic screening to identify the possible role of each new gene discovered6,7. The methods for bacterial phenotyping described here are relevant to Streptomyces mutants isolated by transposon mutagenesis8,9,10,11,12,13 and other random methods, such as chemical and ultra-violet (UV) mutagenesis14, as well as the directed construction of mutations such as gene deletions using recombineering15,16 or CRISPR-Cas9 (Clustered Regularly Interspaced Short Palindromic Repeats) genome editing technology17,18,19 and point mutations20.
As antibiotic resistance among pathogens becomes increasingly prevalent, the need for new antibiotics becomes increasingly urgent21,22. The &#34;Small World Initiative&#34; (or SWI) is an effective Science, Technology, Engineering and Mathematics (STEM) learning23 and research strategy24 to combat antibiotic resistance through the crowdsourcing of college students, and more recently high school students. Supported courses work includes identifying new soil microbes that produce novel antibiotics (http://www.smallworldinitiative.org). It is believed that a major source of undiscovered antibiotics will continue to be Streptomyces species found in a wide range of soil and water habitats25,26,27,28,29,31. Recently, our laboratory and the work of others have discovered and characterized signaling genes in Streptomyces species that regulate morphology and development, including antibiotic production7,32,33. Changes in expression of these genes result in a change in the amount and timing of antibiotics produced. Skilled phenotyping of new species and new antibiotic-producing mutants will continue to be important. As new instructors and their students become major contributors to the field of drug discovery, training in bacterial phenotyping is necessary for the success of these novice individuals. Furthermore, these experiments are tractable for high school STEM or university microbiology education laboratories. They represent a demonstration of basic microbial genetic principles in a teaching lab setting.

<table><tbody><tr><td>50% Glycerol</td><td>Sigma-Aldrich</td><td>G5516</td><td></td></tr><tr><td>Immersion Oil (Type DF)</td><td>Cragille&amp;nbsp;</td><td>16482</td><td></td></tr><tr><td>Lens Paper</td><td>Fisherbrand</td><td>11-995</td><td></td></tr><tr><td>Sterile Water</td><td>GeneMate</td><td>G-3250-1L</td><td></td></tr><tr><td>100% Ethanol</td><td>Sigma-Aldrich</td><td>E7023</td><td></td></tr><tr><td>Propidium Iodide</td><td>Invitrogen</td><td>P1304MP</td><td></td></tr><tr><td>Toothpicks (flat)</td><td>Target</td><td>081-22-1957</td><td></td></tr><tr><td>Pipette Tips (10 mL)</td><td>GeneMate</td><td>P-1240-10</td><td></td></tr><tr><td>Pipette Tips (200 mL)</td><td>GeneMate</td><td>P-1240-200Y</td><td></td></tr><tr><td>Pipette Tips (1,250 mL)</td><td>GeneMate</td><td>P-1240-1250</td><td></td></tr><tr><td>Wooden Applicators 6&#039;&#039;</td><td>Solon Care</td><td>070809</td><td></td></tr><tr><td>Cotton Swabs</td><td>Fisherbrand</td><td>23-400-124</td><td></td></tr><tr><td>Soy Flour&amp;nbsp;</td><td>Bob&#039;s Red Mill&amp;nbsp;</td><td>1516C164</td><td></td></tr><tr><td>D-Mannitol</td><td>Sigma-Aldrich</td><td>M4125-1KG</td><td></td></tr><tr><td>Agar</td><td>Sigma-Aldrich</td><td>A1296-1KG</td><td></td></tr><tr><td>Glycerol 100%</td><td>VWR amresco life science&amp;nbsp;</td><td>0854-1L</td><td></td></tr><tr><td>0.8% NaCl (Saline)</td><td>Sigma-Aldrich</td><td>SLBB9000V</td><td></td></tr><tr><td>1.2 mL freezer tube</td><td>NEST</td><td>606101</td><td></td></tr><tr><td>Ultra low (-80 &amp;deg;C) freezer</td><td>SO-LOW</td><td>U85-18</td><td></td></tr><tr><td>Cryo Safety Gloves</td><td>Bel-Art</td><td>H13201</td><td></td></tr><tr><td>Petri dishes&amp;nbsp;</td><td>Sigma-Aldrich</td><td>P5856-500EA</td><td></td></tr><tr><td>Cover slips #1.5</td><td>Thomas Scientific</td><td>64-0721</td><td></td></tr><tr><td>Slides</td><td>Carolina</td><td>63-2010</td><td></td></tr><tr><td>Autoclave</td><td>Tuttnauer</td><td>2540E</td><td></td></tr><tr><td>Phase-Contrast Microscope&amp;nbsp;</td><td>Olympus</td><td>BX40</td><td></td></tr><tr><td>Forceps</td><td>Carolina Biological</td><td>624504</td><td></td></tr><tr><td>Bunsen Burner</td><td>Carolina Biological</td><td>706706</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>34860-1L-R</td><td></td></tr><tr><td>PBS (phosphate buffer saline)</td><td>Sigma-Aldrich</td><td>P4417-50TAB</td><td></td></tr><tr><td>WGA-FITC</td><td>Biotium</td><td>29022-1</td><td></td></tr><tr><td>Clear nail polish</td><td>OPI</td><td>22001009000</td><td></td></tr><tr><td>ImageJ</td><td>NIH</td><td>Free Software</td><td></td></tr><tr><td>100 mL beaker</td><td>Pyrex USA</td><td>1000-T</td><td></td></tr><tr><td>1 L beaker</td><td>Carolina Biological</td><td>721253</td><td></td></tr><tr><td>1 L flask</td><td>Fisherbrand</td><td>S63274</td><td></td></tr><tr><td>250 mL flask</td><td>Pyrex USA</td><td>4980</td><td></td></tr><tr><td>100 mL graduated cylinder</td><td>Carolina Biological</td><td>721788</td><td></td></tr><tr><td>500 mL graduated cylinder&amp;nbsp;</td><td>Carolina Biological</td><td>721792</td><td></td></tr><tr><td>Stir Bar</td><td>Fisher Scientific</td><td>22-271825</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5810R</td><td></td></tr><tr><td>Camera for Microscope</td><td>Olympus</td><td>DP72</td><td></td></tr><tr><td>Nitrile Examination Gloves (Med)</td><td>Bio Excell</td><td>71011002</td><td></td></tr><tr><td>Vortex Mixer</td><td>Carolina Biological</td><td>701077</td><td></td></tr></tbody></table>

visual and microscopic evaluation of streptomyces developmental mutants

Streptomycetes are filamentous soil bacteria belonging to the phylum Actinobacteria that are found throughout the world and produce a wide array of antibiotics and other secondary metabolites. Streptomyces coelicolor is a well-characterized, non-pathogenic species that is amenable to a variety of analyses in the lab. The phenotyping methods described here use S. coelicolor as a model streptomycete; however, the methods are applicable to all members of this large genus as well as some closely related actinomycetes. Phenotyping is necessary to characterize new species of Streptomyces identified in the environment, and it is also a vital first step in characterizing newly isolated mutant strains of Streptomyces. Proficiency in phenotyping is important for the many new researchers who are entering the field of Streptomyces research, which includes the study of bacterial development, cell division, chromosome segregation, and second messenger signaling. The recent crowdsourcing of antibiotic discovery through the isolation of new soil microbes has resulted in an increased need for training in phenotyping for instructors new to the field of Streptomyces research and their college or high school students. This manuscript describes methods for bacterial strain propagation, storage, and characterization through visual and microscopic examination. After reading this article, new researchers (microbiology education laboratories and citizen scientists) should be able to manipulate Streptomyces strains and begin visual characterization experiments.

Initial phenotyping experiments are necessary for characterizing new species and strains and can be used as a complimentary approach to the phylogenetics and DNA-DNA hybridization experiments that are used for characterizing new species. Streptomyces mutants resulting from random mutagenesis methods such as chemical, UV, or transposon mutagenesis are typically identified through direct, visual screens on agar plates. Colonies of Streptomyces are examined for changes in phenotype in comparison to the wild-type, parental strain. For example, a lighter colored aerial mycelium may indicate a lower level of gray pigment caused by a defect in sporulation, or the lack of a fuzzy appearance is indicative of a block in aerial mycelium formation (Figure 1). Many streptomycetes produce pigmented antibiotics in vegetative mycelium or surrounding agar. S. coelicolor produces two pigmented antibiotics as well as two non-pigmented ones. Actinorhodin is a blue-pigmented antibiotic and undecylprodigiosin is a red-pigmented antibiotic45. Strains that have undergone initial visual colony screens are then propagated by streaking for single colonies.
Following visual identification of potentially interesting mutants, strains are subjected to microscopic examination. Phase-contrast microscopy is especially suited to examining Streptomyces mutants for developmental defects, using the wild-type strain as a control (Figure 2). Wild type S. coelicolor colonies typically produce an aerial mycelium by about two days of growth at 30 &#176;C on MS agar, and long chains of spores by three days of growth. The life cycle will progress slightly slower or faster on other types of media. It is imperative that the mutants be analyzed under the same growth conditions as the wild type when drawing conclusions about the developmental defects and delays. Bald mutants may produce spores after prolonged growth on agar media. A class of mutants referred to as white mutants may either be delayed for spore formation, show a reduction in the abundance of spores produced, produce spores with shape and/or size defects, or simply produce lower levels of the mature, gray spore pigment46. Other mutants investigated thus far may be delayed or accelerated in the life cycle progression such as the mutants affected for the accumulation of signaling molecules (e.g., cyclic-di-GMP47).
A simple follow-up to the light microscopy technique described above is fluorescence microscopy using propidium iodide to stain chromosomal DNA nucleoids and fluorescently labeled wheat germ agglutinin to stain the cell wall of sporulation septa. Spores lacking a chromosome will be devoid of the red propidium iodide staining, while spores that contain less DNA than usual may be identified if they appear to have decreased staining (Figure 3). Wheat germ agglutinin may be used to stain the cell wall and differences in cell wall staining patterns (i.e., cell division defects) may be observed. In Figure 4 the wild-type strain of S. venezuelae, a species that is recently being used as another model streptomycete because of its faster life cycle and ability to sporulate in liquid45, has been stained with WGA-FITC to elucidate the ladder-like array of division septa that are typically seen early during sporulation.
The initial phenotyping strategies described here should result in the following: 1) Identification of mutant strains of interest for further study; 2) Acquired knowledge about the newly identified mutant and the potential role of the gene that has been mutated; 3) The formulation of a subsequent series of next experimental steps that can be used to further clarify the role of the gene in question. In the case of a newly identified streptomycete from the environment, the researcher will gain knowledge of the potential new species compared to already characterized species of Streptomyces.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57373/57373fig1.jpg" /> 
Figure 1: Photograph of a representative agar plate demonstrating macroscopic colony phenotypes of S. coelicolor. The agar plate shows the fuzzy, gray visual appearance of the aerial mycelium for wild-type S. coelicolor strain MT1110 (WT) in comparison to various random transposon insertion mutants with developmental phenotypes. Mutants are either lacking an aerial mycelium [bald (bld)] or an aerial mycelium with reduced spore pigmentation [white (whi)]. Transposon mutants were generated using mini-Tn5 for insertion mutagenesis6,7. The strains were grown on MS agar for 5 days at 30 &#176;C. Petri dish diameter, 100 mm.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57373/57373fig2.jpg" /> 
Figure 2: Phase-contrast micrographs showing a representative aerial filament for S. coelicolor white mutant strains containing random transposon insertion mutations. (A) Shown is a representative wild type aerial filament that has undergone synchronous, regularly-spaced cell divisions, producing evenly shaped and sized spores (MT1110). (B-F) The remaining panels show the vastly different microscopic phenotypes of various white mutants that were isolated via random transposon mutagenesis. Panel B shows an aerial filament that has not undergone sporulation, but instead has produced bulging areas within the filament. Long arrows in C, D, and F indicate abnormally large spores that are characteristic of mutants MIC42, MIC43, and TH49. The short arrow in panel D indicates an example of a lysed spore compartment. Panel E shows the partially constricted smaller compartments that are typical of the spore chains produced by mutant TH10. The strains were grown on MS agar for 5 days at 30 &#176;C. The mutants are unrelated to those shown in Figure 1. A wild type spore is approximately 1.1 &#181;m on the long axis. Scale bar =&#160;2 &#181;m. <a href="/files/ftp_upload/57373/57373fig2large.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/57373/57373fig3v2.jpg" /> 
Figure 3: Representative micrographs showing S. coelicolor mutant chromosome segregation phenotypes. (A) A diagram representation shows the typical wild-type, regularly-spaced staining appearance for a chain of spores (every spore contains chromosomal DNA) compared to the intermittent staining pattern for a mutant that displays a chromosome segregation defect. (B) Each pair of panels shows the same spore chain observed by Differential Interference Contrast (DIC) image on the left and a propidium iodide-stained fluorescence image is shown on the right. The wild-type phenotype (a, b) is compared to two random transposon insertion mutants (c, d and e, f). Arrows indicate spores devoid of DNA. The strains were grown on MS agar for 5 days at 30 &#176;C. The mutants shown are unrelated to those in Figure 1 and Figure 2. Scale bar =&#160;1 &#181;m. <a href="/files/ftp_upload/57373/57373fig3v2large.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/57373/57373fig4v3.jpg" /> 
Figure 4: Fluorescence micrograph of S. venezuelae wild-type strain stained with WGA-FITC. (A) A diagram of an aerial filament appears smooth with no indentations and no visible signs of sporulation using phase-contrast microscopy in comparison with the ladder-like array of cell wall staining that indicates an early stage of sporulation has begun in that same filament using WGA-FITC under fluorescence microscopy. (B) Micrographs of wild-type S. venezuelae. (a) Smooth aerial filaments are present among chains of spores. (b) Within the mycelium shown in panel a, one aerial filament at an early stage in development possesses a ladder-like array of cell wall deposition. Arrows indicate the regularly-spaced formation of cross-walls stained by WGA-FITC that develop synchronously within a single aerial filament as it undergoes developmentally-associated sporulation. The strain was grown on MYM(Maltose, Yeast extract, Malt extract) MYM agar for 38 h at 30 &#176;C. Scale bar =&#160;2 &#181;m. <a href="/files/ftp_upload/57373/57373fig4v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visual and Microscopic Evaluation of Streptomyces Developmental Mutants at JoVE.com

Visual and Microscopic Evaluation of Streptomyces Developmental Mutants

Here we present protocols for novice researchers to initiate phenotyping for the pharmacologically important bacterial genus Streptomyces.

Visual and Microscopic Evaluation of Streptomyces Developmental Mutants

Streptomycetes are filamentous soil bacteria belonging to the phylum Actinobacteria that are found throughout the world and produce a wide array of ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Immunology and Infection

17.6K Views.  Otterbein University. Here we present protocols for novice researchers to initiate phenotyping for the pharmacologically important bacterial genus Streptomyces.

Video: Visual and Microscopic Evaluation of Streptomyces Developmental Mutants

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the replacement of the entire gene with a selectable marker via homologous recombination. During homologous recombination, exchange of DNA takes place between sequences with high similarity. Therefore, linear genomic sequences flanking a target gene can be used to specifically direct a selectable marker to the desired integration site. Blunt ends of the deletion construct activate the cell&#39;s DNA repair systems and thereby promote integration of the construct either via homologous recombination or by non-homologous-end-joining. In organisms with efficient homologous recombination, the rate of successful gene deletion can reach more than 50% making this strategy a valuable gene disruption system. The smut fungus Ustilago maydis is a eukaryotic model microorganism showing such efficient homologous recombination. Out of its about 6,900 genes, many have been functionally characterized with the help of deletion mutants, and repeated failure of gene replacement attempts points at essential function of the gene. Subsequent characterization of the gene function by tagging with fluorescent markers or mutations of predicted domains also relies on DNA exchange via homologous recombination. Here, we present the U. maydis strain generation strategy in detail using the simplest example, the gene deletion.

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

We describe a robust gene replacement strategy to genetically manipulate the smut fungus Ustilago maydis. This protocol explains how to generate deletion mutants to investigate infection phenotypes. It can be extended to modify genes in any desired way, e.g., by adding a sequence encoding a fluorescent protein tag.

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the ...

Genetics

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not representative of the behavior, status or phenotype of single cells. Due to this new insight the number of single cell studies rises continuously (for recent reviews see 1,2,3). However, many of the single cell techniques applied do not allow monitoring the development and behavior of one specific single cell in time (e.g. flow cytometry or standard microscopy).
Here, we provide a detailed description of a microscopy method used in several recent studies 4, 5, 6, 7, which allows following and recording (fluorescence of) individual bacterial cells of Bacillus subtilis and Streptococcus pneumoniae through growth and division for many generations. The resulting movies can be used to construct phylogenetic lineage trees by tracing back the history of a single cell within a population that originated from one common ancestor. This time-lapse fluorescence microscopy method cannot only be used to investigate growth, division and differentiation of individual cells, but also to analyze the effect of cell history and ancestry on specific cellular behavior. Furthermore, time-lapse microscopy is ideally suited to examine gene expression dynamics and protein localization during the bacterial cell cycle. The method explains how to prepare the bacterial cells and construct the microscope slide to enable the outgrowth of single cells into a microcolony. In short, single cells are spotted on a semi-solid surface consisting of growth medium supplemented with agarose on which they grow and divide under a fluorescence microscope within a temperature controlled environmental chamber. Images are captured at specific intervals and are later analyzed using the open source software ImageJ.

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

This protocol provides a step-by-step procedure to monitor single cell behavior of different bacteria in time using automated fluorescence time-lapse microscopy. Furthermore, we provide guidelines how to analyze the microscopy images.

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not ...

Studying Oxidative Stress Caused by the Mitis Group Streptococci in Caenorhabditis elegans

Caenorhabditis elegans (C. elegans), a free-living nematode, has emerged as an attractive model to study host-pathogen interactions. The presented protocol uses this model to determine the pathogenicity caused by the mitis group streptococci via the production of H2O2. The mitis group streptococci are an emerging threat that cause many human diseases such as bacteremia, endocarditis, and orbital cellulitis. Described here is a protocol to determine the survival of these worms in response to H2O2 produced by this group of pathogens. Using the gene skn-1 encoding for an oxidative stress response transcription factor, it is shown that this model is important for identifying host genes that are essential against streptococcal infection. Furthermore, it is shown that activation of the oxidative stress response can be monitored in the presence of these pathogens using a transgenic reporter worm strain, in which SKN-1 is fused to green fluorescent protein (GFP). These assays provide the opportunity to study the oxidative stress response to H2O2 derived by a biological source as opposed to exogenously added reactive oxygen species (ROS) sources.

Studying Oxidative Stress Caused by the Mitis Group Streptococci in Caenorhabditis elegans

The nematode Caenorhabditis elegans is an excellent model to dissect host-pathogen interactions. Described here is a protocol to infect the worm with members of the mitis group streptococci and determine activation of the oxidative stress response against H2O2 produced by this group of organisms.

Caenorhabditis elegans (C. elegans), a free-living nematode, has emerged as an attractive model to study host-pathogen interactions. The presented ...

Developmental Biology

Analysis of the Development of a Morphological Phenotype as a Function of Protein Concentration in Budding Yeast

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of phenotype development as a function of protein concentration at population and single-cell levels in Saccharomyces cerevisiae. Although this protocol is based on the overexpression of a protein, it can easily be adapted for morphological phenotypes dependent on suppression of protein expression. Our lab is interested in studying the signaling properties of the endocytic adaptor protein epsin. To that purpose we used a dominant negative approach in which we over-expressed the conserved Epsin N-Terminal Homology (ENTH) domain in order to interfere with the functions of endogenous epsin-2 (Ent2 or YLR206W). We observed that overexpression of the ENTH domain of Ent2 (ENTH2) in wild type cells led to a cell division defect that is dependent on the mislocalization of a family of scaffolding proteins, septins.

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of phenotype development as a function of protein concentration at population and single-cell levels in Saccharomyces cerevisiae.

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of ...

Biology

Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

Live-cell imaging of biological processes at the single cell level has been instrumental to our current understanding of the subcellular organization of bacterial cells. However, the application of time-lapse microscopy to study the cell biological processes underpinning development in the sporulating filamentous bacteria Streptomyces has been hampered by technical difficulties. 
Here we present a protocol to overcome these limitations by growing the new model species, Streptomyces venezuelae, in a commercially available microfluidic device which is connected to an inverted fluorescence widefield microscope. Unlike the classical model species, Streptomyces coelicolor, S. venezuelae sporulates in liquid, allowing the application of microfluidic growth chambers to cultivate and microscopically monitor the cellular development and differentiation of S. venezuelae over long time periods. In addition to monitoring morphological changes, the spatio-temporal distribution of fluorescently labeled target proteins can also be visualized by time-lapse microscopy. Moreover, the microfluidic platform offers the experimental flexibility to exchange the culture medium, which is used in the detailed protocol to stimulate sporulation of S. venezuelae in the microfluidic chamber. Images of the entire S. venezuelae life cycle are acquired at specific intervals and processed in the open-source software Fiji to produce movies of the recorded time-series.

Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

Streptomyces are characterized by a complex life cycle that has been experimentally challenging to study by cell biological means. Here we present a protocol to perform fluorescence time-lapse microscopy of the complete life cycle by growing Streptomyces venezuelae in a microfluidic device.

Live-cell imaging of biological processes at the single cell level has been instrumental to our current understanding of the subcellular organization of ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

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

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

The intracellular bacterial pathogen Chlamydia trachomatis undergoes a developmental cycle consisting of two morphologically discrete developmental forms. The non-replicative elementary body (EB) initiates infection of the host. Once inside, the EB differentiates into the reticulate body (RB). The RB then undergoes multiple rounds of replication, before differentiating back to the infectious EB form. This cycle is essential for chlamydial survival as failure to switch between cell types prevents either host invasion or replication.
Limitations in genetic techniques due to the obligate intracellular nature of Chlamydia have hampered identification of the molecular mechanisms involved in the cell-type development. We designed a novel dual promoter-reporter plasmid system that, in conjunction with live-cell microscopy, allows for the visualization of cell type switching in real time. To identify genes involved in the regulation of cell-type development, the live-cell promoter-reporter system was leveraged for the development of a forward genetic approach by combining chemical mutagenesis of the dual reporter strain, imaging and tracking of Chlamydia with altered developmental kinetics, followed by clonal isolation of mutants. This forward genetic workflow is a flexible tool that can be modified for directed interrogation into a wide range of genetic pathways.

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

This protocol utilizes fluorescent promoter-reporters, live-cell microscopy, and individual inclusion extraction in a directed forward genetic approach to identify and isolate developmental mutants of Chlamydia trachomatis.

The intracellular bacterial pathogen Chlamydia trachomatis undergoes a developmental cycle consisting of two morphologically discrete developmental forms. ...

Use of In Vivo Imaging to Screen for Morphogenesis Phenotypes in Candida albicans Mutant Strains During Active Infection in a Mammalian Host

Candida albicans is an important human pathogen. Its ability to switch between morphologic forms is central to&#160;its pathogenesis; these morphologic changes are regulated by a complex signaling network controlled in response to environmental stimuli. These regulatory components have been highly studied, but almost all studies use a variety of in vitro stimuli to trigger filamentation. To determine how morphogenesis is regulated during the pathogenesis process, we developed an in vivo microscopy system to obtain high spatial resolution images of organisms undergoing hyphal formation within the mammalian host. The protocol presented here describes the use of this system to screen small collections of C. albicans mutant strains, allowing us to identify key regulators of morphogenesis as it occurs at the site of infection. Representative results are presented, demonstrating that some regulators of morphogenesis, such as the transcriptional regulator Efg1, have consistent phenotypes in vitro and in vivo, whereas other regulators, such as adenyl cyclase (Cyr1), have significantly different phenotypes in vivo compared to in vitro.

Use of In Vivo Imaging to Screen for Morphogenesis Phenotypes in Candida albicans Mutant Strains During Active Infection in a Mammalian Host

This manuscript describes a method for screening moderately sized Candida albicans mutant libraries for morphogenesis phenotypes during active infection in a mammalian host using non-invasive confocal microscopy.

Candida albicans is an important human pathogen. Its ability to switch between morphologic forms is central to&#160;its pathogenesis; these morphologic ...

Microscopy Techniques for Interpreting Fungal Colonization in Mycoheterotrophic Plants Tissues and Symbiotic Germination of Seeds

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching mycoheterotrophic plants (i.e., plants that obtain carbon from fungi), remarkable aspects of their structural adaptations, the patterns of tissue colonization by fungi, and the morphoanatomy of subterranean organs can enlighten their developmental strategies and their relationships with hyphae, the source of nutrients. Another important role of symbiotic fungi is related to the germination of orchid seeds; all Orchidaceae species are mycoheterotrophic during germination and seedling stage (initial mycoheterotrophy), even the ones that photosynthesize in adult stages. Due to the lack of nutritional reserves in orchid seeds, fungal symbionts are essential to provide substrates and enable germination. Analyzing germination stages by structural perspectives can also answer important questions regarding the fungi interaction with the seeds. Different imaging techniques can be applied to unveil fungi endophytes in plant tissues, as are proposed in this article. Freehand and thin sections of plant organs can be stained and then observed using light microscopy. A fluorochrome conjugated to wheat germ agglutinin can be applied to the fungi and co-incubated with Calcofluor White to highlight plant cell walls in confocal microscopy. In addition, the methodologies of scanning and transmission electron microscopy are detailed for mycoheterotrophic orchids, and the possibilities of applying such protocols in related plants is explored. Symbiotic germination of orchid seeds (i.e., in the presence of mycorrhizal fungi) is described in the protocol in detail, along with possibilities of preparing the structures obtained from different stages of germination for analyses with light, confocal, and electron microscopy.

This protocol aims to provide detailed procedures for collecting, fixing, and maintaining mycoheterotrophic plant samples, applying different microscopy techniques such as scanning and transmission electron microscopy, light, confocal, and fluorescence microscopy to study fungal colonization in plants tissues and seeds germinated with mycorrhizal fungi.

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching ...

Visual and Microscopic Evaluation of Streptomyces Developmental Mutants

Summary

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Chapters in this video

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

Studying Oxidative Stress Caused by the Mitis Group Streptococci in Caenorhabditis elegans

Analysis of the Development of a Morphological Phenotype as a Function of Protein Concentration in Budding Yeast

Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Visualization of Germinosomes and the Inner Membrane in Bacillus subtilis Spores

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

Use of In Vivo Imaging to Screen for Morphogenesis Phenotypes in Candida albicans Mutant Strains During Active Infection in a Mammalian Host

Microscopy Techniques for Interpreting Fungal Colonization in Mycoheterotrophic Plants Tissues and Symbiotic Germination of Seeds