The authors would like to thank Funda&#231;&#227;o de Amparo &#224; Pesquisa do Estado de S&#227;o Paulo&#160;(FAPESP)&#160;for providing financial support (Grant #2019/03746-0). MOB thanks for PhD scholarship received from FAPESP&#160;(process 2021/08013-0) and Coordena&#231;&#227;o de Aperfei&#231;oamento de Pessoal de N&#237;vel Superior - Brazil (CAPES) - Finance Code 001. AR also thanks Conselho Nacional de Desenvolvimento Cient&#237;fico e Tecnol&#243;gico (CNPq)&#160;for a research fellowship (#305269/2018). The authors would like to thank Marcia Regina de Moura Aouada and Antonio Teruyoshi Yabuki for helping with pilot tests for sample preparation, to Renato Barbosa Salaroli for technical assistance, and to Enzo Sorrentino for helping in photo shooting. This study was carried out under the access genetic heritage authorization # SISGen AA39A6D.

1. Sampling field colonies
NOTE: When collecting ant colonies, certify that all the permissions required by local legislation are obtained before collecting. In our case, the collecting permit #74585 was issued by Instituto Chico Mendes de Conserva&#231;&#227;o e Biodiversidade (ICMBio). When the samples come from a lab colony, go to section 2.
<ol>
	<li>Locate&#160;and mark the colony. Excavate a trench surrounding the nest area until the garden chamber is exposed (Figure 2A). 
	NOTE: Some attine species may build their colonies under the leaf litter or inside rotten logs. In such cases, carefully revolve the litter or carefully break the logs for collecting the samples. For detailed information on locating, collecting, and maintaining living colonies of diverse attine ant species, see Sosa-Calvo et al.46.</li>
	<li>Open the garden chamber laterally to prevent the soil from falling on the garden surface. Carefully collect garden samples using either entomological forceps, a spoon, or a kitchen skimmer, depending on the garden size. 
	NOTE: Be sure to sterilize the tools before collecting. When collecting leaf-cutting ants&#39; gardens, wear thick fabric gloves to prevent (or at least attenuate) worker bites. For other attine species, fabric gloves are optional.</li>
	<li>Transfer the garden samples to a clean plastic recipient containing a layer of plaster for balancing the garden&#39;s humidity. After transferring the garden and ant workers, hermetically close the recipient to avoid sample drying. Store garden samples at 23-25 &#176;C until processing.</li>
	<li>Close the trench with previously removed soil.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67334/67334fig02v3.jpg" /> 
Figure 2: Sample preparation protocol. (A) Sampling of field colonies. (B) Sample processing. (C)&#160;Brief fundamentals and workflow for sample preparation: 1. Fixation: for strengthening and preserving sample structure. 2. Dehydration: samples&#39; water content is exchanged for ethanol. 3. Critical point drying: liquid CO2&#160;replaces ethanol and is evaporated. 4. Mounting: sample displayed for analysis. 5. Sputter-coating with gold: prevent sample charging. 6. Imaging. Illustrations and photos by Mariana Barcoto. <a href="https://www.jove.com/files/ftp_upload/67334/67334fig02largev3.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Reagents
NOTE: Bear in mind that the following solutions should be prepared beforehand.
<ol>
	<li>Prepare 0.2 M sodium cacodylate buffer. For this, dissolve 42.8 g of sodium cacodylate in 800 mL of distilled water, stirring until dissolution and pH stabilization at 7.2 (if required, adjust pH using hydrochloric acid). Make up the volume of the solution to 1 L by adding distilled water. Store the solution at 4 &#176;C (for ~1 month). 
	CAUTION: Glutaraldehyde and paraformaldehyde are toxic and should be handled within a fume hood. Wear nitrile gloves and protective glasses all the time while handling such reagents.</li>
	<li>Prepare Karnovsky&#39;s fixative (modified from Karnovsky)47. For this, add 10 mL of 25% glutaraldehyde aqueous solution and 10 mL of 20% paraformaldehyde aqueous solution to 25 mL of 0.2 M sodium cacodylate buffer and mix. Add 1 mL of 0.1 M calcium chloride (CaCl2) and make up the volume to 100 mL by adding distilled water. Store the solution at 4 &#176;C (for up to 1 month). 
	CAUTION: Karnovsky&#39;s fixative is harmful when inhaled and may cause skin and eye irritation. Thus, avoid breathing vapors, using only outdoors or in a well-ventilated area. Wear nitrile gloves and protective glasses all the time when handling the solution.</li>
	<li>Using absolute ethanol (analytical grade), prepare 30%, 50%, 70%, and 90% ethanol solutions in distilled water; 100% ethanol is also required.</li>
</ol>
3. Sample fixation
NOTE: Fixatives harden and preserve samples, maintaining morphological features. Aldehydes (such as paraformaldehyde and glutaraldehyde) are non-coagulant fixatives of cross-linking type, inducing cross-links within and between proteins and nucleic acids48.
<ol>
	<li>Remove workers, eggs, pupae, and larvae from the garden samples using entomological forceps (Figure 2B). Set apart garden fragments no larger than 5 mm3. Add the fragments to a 2 mL tube (Figure 2C).</li>
	<li>To the tubes containing the samples, use a Pasteur glass pipette to add ~1 mL of Karnovsky fixative solution (ensure the sample is completely covered). Mix by gentle agitation to help the sample to soak and incubate at 4 &#176;C for at least 24 h before continuing sample processing (Figure 2C.1). 
	NOTE: When the following dehydration steps will not be carried out immediately after the fixation, the protocol may be paused at this step, and samples may be stored for ~1 year at 4 &#176;C. 
	We suggest using a Pasteur glass pipette, as it is composed of an inert material and is easier to clean for reuse afterward. Garden material is highly hydrophobic, tending to float over the surface of the fixative solution. It usually takes up to 5 min to be completely soaked in the fixative. Make sure the fixative covers the garden fragments since its volume tends to be reduced as it enters the sample&#39;s pores. We empirically verified that after the sample was soaked and became wet, its components (in particular, the delicate fungal mycelium) became susceptible to shattering when mixed further. Thus, we recommend avoiding shaking the samples as much as possible.</li>
</ol>
4. Sample dehydration
NOTE: The&#160;ethanol washing series gradually exchanges the water in samples for ethanol. It is important to start with a low-concentration ethanol solution (see below) to avoid excessively damaging or collapsing such delicate samples49.
<ol>
	<li>Completely remove Karnovsky&#39;s fixative solution using a glass pipette, being careful to not disrupt the sample (Figure 2C.1). 
	NOTE: Discard Karnovsky&#39;s fixative in a recipient properly labeled for toxic chemical residue management.</li>
	<li>Immediately after removing the fixative, add 1 mL of 30% ethanol, being careful to not disrupt the sample, and incubate for 10 min at room temperature (Figure 2C.2). 
	NOTE: Samples need to always be soaked in solution. Make sure to quickly replace solutions during the ethanol gradual washing series. As the garden loses its porous aspect and aggregates at the tube bottom, 1 mL of ethanol tends to be enough to cover the sample (when the sample is no larger than 5 mm3). However, if the samples are not completely covered, add ethanol until the sample is completely covered.</li>
	<li>Completely remove 30% ethanol with a glass pipette, being careful not to disrupt the sample. Discard 30% ethanol properly. 
	NOTE: For the entire ethanol gradual washing series, discard ethanol in a recipient properly labeled for toxic chemical residue management.</li>
	<li>Add 1 mL of 50% ethanol and incubate for 10 min at room temperature. Completely remove 50% ethanol with a glass pipette, being careful not to disrupt the sample. Discard 50% ethanol properly.</li>
	<li>Add 1 mL of 70% ethanol and incubate for 10 min at room temperature. Completely remove 70% ethanol with a glass pipette, being careful not to disrupt the sample. Discard 70% ethanol properly. 
	NOTE: At the 70% ethanol washing step, the user may pause the protocol if needed, as sample tubes may be stored overnight at 4 &#176;C, when the material will not be immediately processed.</li>
	<li>Add 1 mL of 90% ethanol and incubate for 10 min at room temperature. Completely remove 90% ethanol with a glass pipette, being careful not to disrupt the sample. Discard 90% ethanol properly.</li>
	<li>Add 1 mL of 100% ethanol and incubate for 10 min at room temperature. Completely remove 100% ethanol with a glass pipette, being careful to not disrupt the sample. Discard 100% ethanol properly.</li>
	<li>Using a forceps and/or a spatula, carefully transfer the samples to specimen containers for the critical point dryer&#160;(CPD), containing sample identification labels (made previously in paper and pencil).&#160;For avoiding samples drying, this transfer is carried out with the container placed in a Petri dish covered with 100% ethanol.</li>
	<li>Put the lids on the containers and immerse them in a graduated glass beaker containing 100% ethanol enough to submerge the containers. Cover the glass beaker and incubate&#160;for 10 min at room temperature; then properly discard 100% ethanol.</li>
	<li>Transfer the specimen containers to another graduated glass beaker containing enough 100% ethanol to submerge the containers. Cover the glass beaker and incubate for 10 min at room temperature; then, transfer specimen containers to the critical point dryer. 
	NOTE: After the washing series, glass pipettes and beakers should be abundantly rinsed with distilled water, and this residual water should be discarded in a recipient properly labeled for toxic chemical residue management. After rinsing, glass items may be washed with neutral detergent, rinsed with tap water, and air dried.</li>
</ol>
5. Critical point drying
NOTE: A Critical Point Dryer exchanges the ethanol in samples for liquid carbon dioxide (CO2), which evaporates from the sample at higher temperature and pressure. Please follow the manufacturer&#39;s instructions for such procedures.
<ol>
	<li>Switch on the equipment.</li>
	<li>Open the chamber, place the sample containers inside, and add 100% ethanol until it covers the containers. Close the chamber.</li>
	<li>Activate the Cool option, and wait until the temperature reaches 10 &#176;C.</li>
	<li>Open the CO2&#160;cylinder valve and activate the Stirrer option.</li>
	<li>Activate the CO2 in option, always checking the chamber to verify how much CO2&#160;has already filled it. When the chamber is almost filled, deactivate the CO2 in option and activate the Exchange option, keeping it activated until there is enough CO2&#160;only to cover the containers. Make sure that the containers are always covered with CO2&#160;(i.e., deactivate the Exchange option before all CO2 has left the chamber). Repeat step 5.5 6x.</li>
	<li>Activate CO2 in one last time and fill the chamber until the containers are covered.</li>
	<li>Activate the Heat option and deactivate the Stirrer option. Close the CO2&#160;cylinder valve.</li>
	<li>Wait until the temperature rises to 35 &#176;C; then, activate the Gas out option. 
	NOTE: At around 30 &#176;C, the chamber reaches pressures of 70-80 bar, achieving the critical point, where the liquid disappears.</li>
	<li>When the chamber pressure reaches 1 bar, all gas content has been removed. Open the chamber and remove the containers.</li>
	<li>Switch off the equipment.</li>
</ol>
6. Mounting
<ol>
	<li>Prepare SEM sample holders (i.e., aluminum stubs; Figure 2C.4).
	<ol>
		<li>Wrap the stubs with a piece of aluminum foil, covering only the top, to facilitate stub cleaning after the analysis.</li>
		<li>Identify the stubs by writing the sample code/number on the holder bottom, guaranteeing the identification of what is placed on top.</li>
		<li>Cover the upper part with double-sided carbon tape. Place the stubs into a specimen holder.</li>
	</ol>
	</li>
	<li>Open the lid of the specimen container, and carefully transfer the dried sample to a glass Petri dish using forceps and spatula. 
	NOTE: Critical point-dried gardens tend to agglomerate forming highly packed samples that should be carefully set apart as fragments no larger than 1 mm3in size.</li>
	<li>Carefully place fragments on the sticky surface of the tape-covered stub. Once the garden fragment touches the tape, it is very difficult to (re)move it, so be careful not to place it in unwanted places or positions. Add up to nine fragments per stub.</li>
	<li>Repeat steps 6.2 and 6.3 for each sample.</li>
</ol>
7. Sputter-coating with gold
NOTE: Coating the sample is required to prevent its charging. Follow the manufacturer&#39;s instructions for adjusting settings such as the operation gas pressure (0.5 &#215; 10-1 mm Hg of gas pressure in this protocol), the sputtering time (220 s), thickness of the gold layer (~120 &#197;), the current (50 mA), and the voltage supply. Sputtering tends to follow a common workflow though the equipment from different manufacturers may operate slightly differently.
<ol>
	<li>Open up the hinged target arm and place the stubs on the specimen table.</li>
	<li>Close down the hinged target arm and check whether the splinter shield of the glass vacuum chamber is properly embedded.</li>
	<li>Open the argon cylinder valve and switch on the main power switch.</li>
	<li>Follow the vacuum rising in the equipment display until it reaches the 0.5 &#215; 10-1 mm Hg mark in the display; then, activate Rinsing. Repeat the operation 5x.</li>
	<li>Switch on the water circulation system, activate the option HV On, and open the gold-film lid. Confirm that the plasma color is pinkish. To follow this protocol, set up 220 s of sputtering with 50 mA voltage, which will deposit a gold layer of ~120 &#197; (12 nm).</li>
	<li>HV On is automatically switched off. Turn off the water circulation system and close the gold-film lid.</li>
	<li>Turn off the main power switch, allowing air to enter the vacuum chamber. For garden and waste samples, repeat steps 7.1-7.7 3x. 
	NOTE: If the imaging will not occur immediately after sample preparation, store the stubs in a hermetic container filled with a silica layer to avoid rehydrating the samples.</li>
</ol>
8. Imaging
NOTE: Follow the manufacturer&#39;s instructions for adjusting SEM settings, such as objective aperture diameter, operating voltage, alignment of the electron beam system, axial alignment, and stigmators.
<ol>
	<li>Place the stubs in the sample holder, taking notes on each sample&#39;s position. 
	NOTE: Wear gloves when inserting or removing the sample holder, and keep the holder as clean as possible.</li>
	<li>Launch the operation software from the desktop.</li>
	<li>Select the instrument settings. Visualize garden samples with the objective aperture diameter&#160;of 30 &#181;m (i.e., in the second&#160;stage), operating in high vacuum, detecting signals from secondary electrons (SED), accelerating voltage of 20 kV, working distance of 15 - 20 mm, probe current of 40.0&#160;(in high current mode), and varying the magnification according to the sample.</li>
	<li>Following the navigation instructions, press the Vent icon and wait for the specimen chamber to be vented. A progress bar indicates the vacuum status.</li>
	<li>When atmospheric pressure has been reached, open the specimen chamber and carefully insert the sample holder.</li>
	<li>Gently close the chamber door and press the Evac icon to evacuate the instrument, following the vacuum status through the progress bar. The navigation system will show the moving stage position and provide a holder graph when the moving is completed. Press the Camera icon to record a photo of the holder to get a top view that will help to navigate between samples while imaging.</li>
	<li>Press the Observation icon to turn on the electron gun and wait for the image to form. Use the Manual User Interface (manual control) to manually move the Z-axis to the proper height (based on the sample height and the determined working distance). 
	NOTE: For prolonging the tungsten filament lifetime, pause the Observation function whenever not actively imaging.</li>
	<li>Use either the options shown on the screen or the manual control to change the raster rate and image position. Move the stage to get a thorough view of the sample, employing the RDC function to focus on specific areas. When observing a target (or an interesting) structure, adjust the magnification, focus, brightness, contrast, and stigmation accordingly. Correct stigmation by using the Manual User Interface to move the stage to X and Y directions.
	<ol>
		<li>Adjust the magnification between 100x and 700x to visualize general garden aspects (such as hyphal density, substrate, and colonization patterns).</li>
		<li>Adjust the magnification between 700x and 1,500x to visualize the microbiota spatial patterns.</li>
		<li>Adjust the magnification between 1,500x and 3,000x to observe the microbiota and biofilm physical interactions.</li>
		<li>Adjust the magnification between 3,000x and 4,000x to focus on specific microbial clusters.</li>
	</ol>
	</li>
	<li>To save an image, use the Freeze function, click on the Photo icon, and set up the file path. As a standard, analyze at least three garden fragments, imaging each one in all magnification ranges mentioned in step 8.8.&#160;This results in at least 12 images per sample, though more images may support more detailed descriptions. We suggest that 15-25 images per sample, ranging all the magnifications in step 8.8 tend to provide fine details for sample description. 
	NOTE: Rastering and saving an image takes a few seconds during which any vibration on the SEM floating table should be avoided.</li>
	<li>After finishing imaging, press the Vent option to vent the chamber, following the vacuum status through the progress bar. Carefully remove the holder, gently close the chamber door, and press Evac to evacuate the chamber.</li>
	<li>As soon as the evacuation is completed, exit the operation software.</li>
	<li>Adjust the brightness and contrast to improve visualization using an image editor.</li>
</ol>

 Microscale SEM Analysis of Microbial Communities in Fungus-Growing Ant Cultures

The authors have no conflicts of interest to disclose.

SEM uses an electron beam to scan the sample, generating an enlarged image of it such that one can visualize three-dimensional microstructures in high resolution. As SEM operates under high vacuum, the removal of up to/more than 99% of water from samples is required. Inside the SEM vacuum chamber, partially hydrated samples may dehydrate and collapse, besides scattering electrons. For high-resolution imaging in SEM, sample preparation should include procedures for removing water while keeping the changes in volume and mo...

Ecosystems are composed of organisms interconnected by processes in a specific geographical location (i.e., the environment). Organisms interact with their environment over time, from which complex and heterogeneous spatial patterns emerge. Spatial patterning determines ecological diversity and stability and, ultimately, ecosystem functioning1,2,3,4. In macroscale ecosystems, such as wetlands, savannas, coral reefs, and arid ecosystems, spatial patterns are correlated with resource flow and concentration. Permitting resource optimization, spatial heterogeneity&#160;and patterning result in more resilient ecosystems than homogeneous ones2. The spatial localization of organisms being at the basis of community ecology is also translated to the microbial world.
Microbial ecosystems, far from organisms randomly and homogeneously mixed throughout microhabitats, exhibit spatial patterns defining much of their functioning5,6,7. From Winogradsky columns to environmental- and host-associated microbiota, these ecosystems are heterogeneously organized in space, with spatial arrangements eliciting different phenotypic responses. Cohabiting species are more likely to interact and influence each other&#39;s physiology. Thus, the community spatial organization, more than its composition per se, delimits ecosystem properties and ecological niches5,7,8. Illustrating these concepts, changes in spatial patterning seem correlated to the pathological progression of dental plaques, caries, gum diseases9,10, in&#64258;ammatory bowel disease11, cystic fibrosis lung infections, chronic wound infections12,13, colorectal cancer, and adenomas14.
Under the scope of microbial biogeography (the study of biodiversity distribution and patterning across space and time on a microscale), the knowledge of microbial ecosystems is vastly benefitted by comprehending their spatial patterns6,13,15,16,17. We have looked into spatial patterns of an insect-built microbial ecosystem, found at the core of the charismatic fungus-growing attine ant (Hymenoptera: Formicidae: Myrmicinae: Attini: Attina) colonies. There resides a &#34;microbial garden,&#34; centered around a basidiomycete fungus in the tribe Leucocoprinae (Basidiomycota: Agaricaceae)&#160;or in the family Pterulaceae (Basidiomycota: Agaricales)18,19,20,21,22. The garden is a spongy structure emerging&#160;from an intertwined mesh of hyphae that grows by metabolizing the mostly plant-based substrate incorporated by the ants (Figure 1). These may include, according to the attine genera: dry plant parts, insect frass and carcasses, freshly cut leaves, seeds, and floral parts23,24. Analogous to an external herbivorous gut, the garden enzymatically and chemically converts recalcitrant polymers into labile nutritional resources, providing the ants with&#160;essential amino acids, lipids, and soluble sugars21,25,26,27,28.
Ultrastructural, enzymatic, and transcriptomic analyses carried out for gardens of the leaf-cutting genera Atta and Acromyrmex suggest these environments structure a continuum of substrate degradation&#160;and nutritional patches26,29,30,31,32. Young parts of the garden tend to be darker due to freshly incorporated substrate after being fragmented. These recently added substrates are often colonized from the edges, which were cut by ant workers and inoculated with mycelial clumps. Radiating from cut edges, fungal hyphae spread over the substrate29,32,33. Hyphal abundance increases as substrate degradation progresses, resulting in whitish and metabolically active regions30,31,32. Older regions, with more degraded substrate and an abundant microbiota29,32, tend to present brownish tones and higher humidity. Workers remove fragments of this region, separating them in waste piles, where they also take substrates that harm the fungal symbiont34,35,36. Waste piles, although physically detached from the garden, are a spot of continuous substrate degradation and nutrient cycling by the abundant inhabitant microbiota29,32,37,38,39.
A microbiota mainly composed of Enterobacter, Klebsiella, Pantoea, Pseudomonas, and Serratia, also inhabits the garden, apparently being shared by diverse attine fungiculture systems. Encoding metabolic pathways that could complement fungal metabolism, the microbiota potentially participates in the garden&#39;s physiological responses40,41,42,43,44. Not only did metagenomic data indicate that the microbiota was there41,42, but also Scanning Electron Microscopy (SEM) analysis of leaf-cutting ants&#39; fungiculture showed mostly rod-shaped bacteria over plant substrate32. Though bacteria (including cellulolytic strains) were isolated from the entire garden, they were visualized only in older parts of the garden and in waste piles, as well as in the initial pellet carried by foundress queens29,32. It was also uncertain whether the microbiota could form biofilms in vivo (i.e., in the garden and waste), as suggested by their metabolic capacity42&#160;and observed in vitro44.
Here, we employed SEM to further comprehend the microbiota spatial organization across the garden regions, detailing microbiota-substrate and microbiota-hyphae physical interactions. By providing images with larger focal depth, SEM permits observations of three-dimensional microscopic structures in high resolution, enabling a thorough analysis of the garden microbiota spatial patterns. We detail steps to fix, dehydrate, dry, sputter coat, and image such heterogeneous and delicate fungal-based samples. By removing the postfixation step using osmium tetroxide (OsO4) and reducing the dehydration time, we simplified the protocols32,33,45&#160;for preparing garden and waste samples for SEM analysis. This adapted protocol preserves hyphal structural patterns, as well as the microbiota and biofilm spatial organization, and could be applied to other delicate microbial ecosystems and biofilms.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67334/67334fig01v2.jpg" /> 
Figure 1: Attine microbial gardens. The garden is a sponge-like structure resulting from an intertwined mesh of hyphae that grows by metabolizing the mostly plant-based substrate incorporated by the ants. Also inhabiting the garden is the microbiota, which encodes metabolic pathways that could complement fungal metabolism. Metagenomic data and previous Scanning Electron Microscopy analysis indicated its presence, though we had scarce knowledge of its spatial organization and physical interactions with the substrate and fungal hyphae. We employed SEM to unveil the microbiota and biofilm spatial organization and patterning. Illustrations by Mariana Barcoto (garden and microbiota adapted from Barcoto and Rodrigues 94), and photos by Mariana Barcoto and Enzo Sorrentino. <a href="https://www.jove.com/files/ftp_upload/67334/67334fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2 mL tube</td>
			<td>Axygen</td>
			<td>MCT-200-C-BRA</td>
			<td>To fix and dehydrate samples</td>
		</tr>
		<tr>
			<td>Calcium chloride anhydrous</td>
			<td>Merck</td>
			<td>C4901</td>
			<td>CaCl2 anhydrous to prepare Karnovsky&#8217;s fixative</td>
		</tr>
		<tr>
			<td>Critical point dryer</td>
			<td>Leica</td>
			<td>EM CPD 300</td>
			<td>For critical point drying</td>
		</tr>
		<tr>
			<td>Double Sided Carbon Conductive Tape, 12 mm (W) X 5 M (L)</td>
			<td>Electron Microscopy Sciences</td>
			<td>77819-12</td>
			<td>For mounting samples</td>
		</tr>
		<tr>
			<td>Entomological forceps</td>
			<td>No specific supplier</td>
			<td></td>
			<td>To manipulate garden samples</td>
		</tr>
		<tr>
			<td>Ethyl alcohol (=ethanol), pure (&#8805;99.5%)</td>
			<td>Sigma-Aldrich</td>
			<td>459836</td>
			<td>For dehydration</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>No specific supplier</td>
			<td></td>
			<td>To manipulate garden samples</td>
		</tr>
		<tr>
			<td>Glass beaker</td>
			<td>No specific supplier</td>
			<td></td>
			<td>For dehydration</td>
		</tr>
		<tr>
			<td>Glass Petri dish</td>
			<td>No specific supplier</td>
			<td></td>
			<td>To manipulate garden samples</td>
		</tr>
		<tr>
			<td>Glass pipette</td>
			<td>No specific supplier</td>
			<td></td>
			<td>To fix and dehydrate samples</td>
		</tr>
		<tr>
			<td>Glutaraldehyde (Aqueous Glutaraldehyde EM Grade 25%)</td>
			<td>Electron Microscopy Sciences</td>
			<td>16220</td>
			<td>To prepare Karnovsky&#8217;s fixative</td>
		</tr>
		<tr>
			<td>Gold target&#160;</td>
			<td>Ted Pella, Inc.</td>
			<td>8071</td>
			<td>To sputter coat with gold</td>
		</tr>
		<tr>
			<td>Hydrochloric acid</td>
			<td>Sigma-Aldrich</td>
			<td>320331</td>
			<td>For adjusting solutions pH</td>
		</tr>
		<tr>
			<td>Image editor</td>
			<td>Photoshop</td>
			<td>any version</td>
			<td>To adjust images</td>
		</tr>
		<tr>
			<td>Paraformaldehyde (Paraformaldehyde 20% Aqueous Solution EM Grade)</td>
			<td>Electron Microscopy Sciences</td>
			<td>15713</td>
			<td>To prepare Karnovsky&#8217;s fixative</td>
		</tr>
		<tr>
			<td>Propilene recipient</td>
			<td>No specific supplier</td>
			<td></td>
			<td>For maintaining alive ant colonies</td>
		</tr>
		<tr>
			<td>Scanning Electron Microscope</td>
			<td>JEOL&#160;</td>
			<td>IT300 SEM</td>
			<td>For sample imaging&#160;</td>
		</tr>
		<tr>
			<td>Sodium cacodylate trihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>C0250</td>
			<td>For preparing sodium cacodylate buffer</td>
		</tr>
		<tr>
			<td>Spatula</td>
			<td>No specific supplier</td>
			<td></td>
			<td>To manipulate garden samples</td>
		</tr>
		<tr>
			<td>Specimen containers with 15 mm dia. x 10 mm high</td>
			<td>Ted Pella, Inc.</td>
			<td>4591</td>
			<td>For critical point drying</td>
		</tr>
		<tr>
			<td>Sputter coater</td>
			<td>Baltec&#160;</td>
			<td>SCD 050</td>
			<td>To coat with gold</td>
		</tr>
		<tr>
			<td>Stub (Aluminium mount, flat end pin) 12.7 mm x 8 mm&#160;</td>
			<td>Electron Microscopy Sciences</td>
			<td>75520</td>
			<td>For mounting samples</td>
		</tr>
	</tbody>
</table>

microbiota of attine ants' gardens: visualizing a microbial landscape by scanning electron microscopy

In macroscale ecosystems, such as rainforests or coral reefs, the spatial localization of organisms is the basis of our understanding of community ecology. In the microbial world, likewise, microscale ecosystems are far from a random and homogeneous mixture of organisms and habitats. Accessing the spatial distribution of microbes is fundamental for understanding the functioning and ecology of the microbiota, as cohabiting species are more likely to interact and influence each other&#39;s physiology.
An interkingdom microbial ecosystem is at the core of fungus-growing ant colonies, which cultivate basidiomycete fungi as a nutritional resource. Attine ants forage for diverse substrates (mostly plant-based), metabolized by the cultivated fungus while forming a spongy structure, a &#34;microbial garden&#34; that acts as an external gut. The garden is an intertwined mesh of fungal hyphae growing by metabolizing the substrate, opening niches for a characteristic and adapted microbiota to establish. The microbiota is thought to be a contributor to substrate degradation and fungal growth, though its spatial organization is yet to be determined.
Here, we describe how we employ Scanning Electron Microscopy (SEM) to investigate, with unprecedented detail, the microbiota and biofilm spatial organization across different fungiculture systems of fungus-growing ants. SEM imaging has provided a description of the microbiota spatial structure and organization. SEM revealed that microbiota commonly assemble in biofilms, a widespread structure of the microbial landscapes in fungiculture. We present the protocols employed to fix, dehydrate, dry, sputter coating, and image such a complex community. These protocols were optimized to deal with delicate and heterogeneous samples, comprising plant and fungal biomass, as well as the microbiota and the biofilm.

Here, we presented a simplified protocol to visualize the components of attine garden and waste samples, such as fungal hyphae, substrate, microbiota, and biofilms. SEM has enhanced our understanding of how the garden and waste scaffold the microbiota structural patterns (Figure 3). In attine gardens, fungal hyphae are branch-like structures covering portions of the substrate surface. Since fungal hyphae tend to be very sensitive to dehydration and rupture, the user may be guided by the hyph...

Microbiota of Attine Ants' Gardens: Visualizing a Microbial Landscape by Scanning Electron Microscopy

We propose an optimized Scanning Electron Microscopy protocol for visualizing highly heterogeneous and delicate samples containing plant and fungal biomass, together with microbiota and biofilm. This protocol allows describing the spatial dimensions of the microbiota organization.

In macroscale ecosystems, such as rainforests or coral reefs, the spatial localization of organisms is the basis of our understanding of community ...

microbiota-attine-ants-gardens-visualizing-microbial-landscape

Research

JoVE Journal

Biology

398 Views.  São Paulo State University (UNESP). We propose an optimized Scanning Electron Microscopy protocol for visualizing highly heterogeneous and delicate samples containing plant and fungal biomass, together with microbiota and biofilm. This protocol allows describing the spatial dimensions of the microbiota organization.

Video: Microbiota of Attine Ants' Gardens: Visualizing a Microbial Landscape by Scanning Electron Microscopy

Cryo-electron Microscopy Specimen Preparation By Means Of a Focused Ion Beam

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms or solutions. We make use of a custom built cryo-transfer station and a modified cryo-SEM preparation chamber2. The spores are taken from a culture, plunge-frozen in a liquid nitrogen slush and observed in the cryo-SEM to select a region of interest. A thin lamella is then extracted using the FIB, attached to a TEM grid and subsequently thinned to electron transparency. The grid is transferred to a cryo-TEM holder and into a TEM for high resolution studies. Thanks to the introduction of a cooled nanomanipulator tip and a cryo-transfer station, this protocol is a straightforward adaptation to cryogenic temperature of the routinely used FIB preparation of TEM samples. As such it has the advantages of requiring a small amount of modifications to existing instruments, setups and procedures; it is easy to implement; it has a broad range of applications, in principle the same as for cryo-TEM sample preparation. One limitation is that it requires skillful handling of the specimens at critical steps to avoid or minimize contaminations.

Cryo Electron Microscopes, either Scanning (SEM) or Transmission (TEM), are widely used for characterization of biological samples or other materials with a high water content1. A SEM/Focused Ion Beam (FIB) is used to identify features of interest in samples and extract a thin, electron-transparent lamella for transfer to a cryo-TEM.

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved substances from the blood into the liver parenchyma for metabolism and processing. Fenestrations are dynamic structures - both their size and/or number can be altered in response to various physiological states, drugs, and disease, making them an important target for modulation. An understanding of how LSEC morphology is influenced by various disease, toxic, and physiological states and how these changes impact on liver function requires accurate measurement of the size and number of fenestrations. In this paper, we describe scanning electron microscopy fixation and processing techniques used in our laboratory to ensure reproducible specimen preparation and accurate interpretation. The methods include perfusion fixation, secondary fixation and dehydration, preparation for the scanning electron microscope and analysis. Finally, we provide a step by step method for standardized image analysis which will benefit all researchers in the field.

The fenestrated liver sinusoidal endothelial cell is a biologically important filter system that is highly influenced by various diseases, toxins, and physiological states. These changes significantly impact on liver function. We describe methods for the standardisation of the measurement of the size and number of fenestrations in these cells.

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

Scanning Electron Microscopy (SEM) Protocols for Problematic Plant, Oomycete, and Fungal Samples

Common problems in the processing of biological samples for observations with the scanning electron microscope (SEM) include cell collapse, treatment of samples from wet microenvironments and cell destruction. Using young floral tissues, oomycete cysts, and fungi spores (Agaricales) as examples, specific protocols to process delicate samples are described here that overcome some of the main challenges in sample treatment for image capture under the SEM.
Floral meristems fixed with FAA (Formalin-Acetic-Alcohol) and processed with the Critical Point Dryer (CPD) did not display collapsed cellular walls or distorted organs. These results are crucial for the reconstruction of floral development. A similar CPD-based treatment of samples from wet microenvironments, such as the glutaraldehyde-fixed oomycete cysts, is optimal to test the differential growth of diagnostic characteristics (e.g., the cyst spines) on different types of substrates. Destruction of nurse cells attached to fungi spores was avoided after rehydration, dehydration, and the CPD treatment, an important step for further functional studies of these cells.
The protocols detailed here represent low-cost and rapid alternatives for the acquisition of good-quality images to reconstruct growth processes and to study diagnostic characteristics.

Scanning Electron Microscopy SEM Protocols for Problematic Plant, Oomycete, and Fungal Samples

Problems in the processing of biological samples for scanning electron microscopy observation include cell collapse, treatment of samples from wet microenvironments and cell destruction. Low-cost and relatively rapid protocols suited for preparing challenging samples such as floral meristems, oomycete cysts, and fungi (Agaricales) are compiled and detailed here.

Common problems in the processing of biological samples for observations with the scanning electron microscope (SEM) include cell collapse, treatment of ...

Preparation and Observation of Thick Biological Samples by Scanning Transmission Electron Tomography

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It also describes an imaging method for studying the 3D structure of thick biological specimens by scanning transmission electron tomography. The sample preparation protocol is based on conventional methods in which the sample is fixed using chemical agents, treated with a heavy atom salt contrasting agent, dehydrated in a series of ethanol baths, and embedded in resin. The specific imaging conditions for observing thick samples by scanning transmission electron microscopy are then described. Sections of the sample are observed using a through-focus method involving the collection of several images at various focal planes. This enables the recovery of in-focus information at various heights throughout the sample. This particular collection pattern is performed at each tilt angle during tomography data collection. A single image is then generated, merging the in-focus information from all the different focal planes. A classic tilt-series dataset is then generated. The advantage of the method is that the tilt-series alignment and reconstruction can be performed using standard tools. The collection of through-focal images allows the reconstruction of a 3D volume that contains all of the structural details of the sample in focus.

This report describes a sample preparation protocol and specific imaging conditions for performing scanning transmission electron tomography of thick biological specimens.

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It ...

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy (SEM)

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to cells, tissues, organelles, and even whole organisms. This protocol focuses on two chemical drying methods, hexamethyldisilazane (HMDS) and t-butyl alcohol (TBA), and their application to imaging of both prokaryotic and eukaryotic organisms using SEM. In this article, we describe how to fix, wash, dehydrate, dry, mount, sputter coat, and image three types of organisms: cyanobacteria (Toxifilum mysidocida, Golenkina sp., and an unknown sp.), two euglenoids from the genus Monomorphina (M. aenigmatica and M. pseudopyrum), and the fruit fly (Drosophila melanogaster). The purpose of this protocol is to describe a fast, inexpensive, and simple method to obtain detailed information about the structure, size, and surface characteristics of specimens that can be broadly applied to a large range of organisms for morphological assessment. Successful completion of this protocol will allow others to use SEM to visualize samples by applying these techniques to their system.

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy SEM

SEM analysis is an effective method to aid in species identification or phenotypic discrimination. This protocol describes the methods for examining specific morphological details of three representative types of organisms and would be broadly applicable to examining features of many organismal and tissue types.

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...