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><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>CaCl&lt;sub&gt;2&lt;/sub&gt; anhydrous to prepare Karnovsky&amp;rsquo;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 (&amp;ge;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&amp;rsquo;s fixative</td></tr><tr><td>Gold target&amp;nbsp;</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&amp;rsquo;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&amp;nbsp;</td><td>IT300 SEM</td><td>For sample imaging&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</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

532 Views.  São Paulo State University (UNESP). We onderzoeken de microbiota die in schimmeltuinen leven op schimmelgroeiende mieren. We willen weten wie ze zijn en hoe ze dit microbiële ecosysteem organiseren. In het bijzonder willen we weten hoe de microbiota fysiek interageert met de schimmel die de mieren kweken.En deze interacties worden meestal gemedieerd door biofilm. De microbiële tuinen die door de mieren worden gecultiveerd, zijn een complex en heterogeen monster dat is samengesteld uit ...

Video: Microbiota van attine mierentuinen: visualisatie van een microbieel landschap door scanning elektronenmicroscopie

Extracting DNA from the Gut Microbes of the Termite (Zootermopsis Angusticollis) and Visualizing Gut Microbes

Termites are among the few animals known to have the capacity to subsist solely by consuming wood.  The termite gut tract contains a dense and species-rich microbial population that assists in the degradation of lignocellulose predominantly into acetate, the key nutrient fueling termite metabolism (Odelson &amp; Breznak, 1983).  Within these microbial populations are bacteria, methanogenic archaea and, in some ("lower") termites, eukaryotic protozoa. Thus, termites are excellent research subjects for studying the interactions among microbial species and the numerous biochemical functions they perform to the benefit of their host.  The species composition of microbial populations in termite guts as well as key genes involved in various biochemical processes has been explored using molecular techniques (Kudo et al., 1998; Schmit-Wagner et al., 2003; Salmassi &amp; Leadbetter, 2003). These techniques depend on the extraction and purification of high-quality nucleic acids from the termite gut environment.  The extraction technique described in this video is a modified compilation of protocols developed for extraction and purification of nucleic acids from environmental samples (Mor  et al., 1994; Berthelet et al., 1996; Purdy et al., 1996; Salmassi &amp; Leadbetter, 2003; Ottesen et al. 2006) and it produces DNA from termite hindgut material suitable for use as template for polymerase chain reaction (PCR).

Extracting DNA from the Gut Microbes of the Termite Zootermopsis Angusticollis and Visualizing Gut Microbes

This video illustrates the technique for extracting DNA from the species of microbes resident in the termite hindgut. The preparation of a wet mount slide, which is useful for visualizing the gut microbial community is also illustrated, and a tour through the species-rich gut environment is given.

Termites are among the few animals known to have the capacity to subsist solely by consuming wood.  The termite gut tract contains a dense and ...

Layers of Symbiosis - Visualizing the Termite Hindgut Microbial Community

Jared Leadbetter takes us for a nature walk through the diversity of life resident in the termite hindgut - a microenvironment containing 250 different species found nowhere else on Earth.  Jared reveals that the symbiosis exhibited by this system is multi-layered and involves not only a relationship between the termite and its gut inhabitants, but also involves a complex web of symbiosis among the gut microbes themselves.

Jared Leadbetter takes us for a nature walk through the diversity of life resident in the termite hindgut - a microenvironment containing 250 different species found nowhere else on Earth. Jared reveals that the symbiosis exhibited by this system is multi-layered and involves not only a relationship between the termite and its gut inhabitants, but also involves a complex web of symbiosis among the gut microbes themselves.

Jared Leadbetter takes us for a nature walk through the diversity of life resident in the termite hindgut - a microenvironment containing 250 different ...

Visualization of Microbiota in Tick Guts by Whole-mount In Situ Hybridization

Infectious diseases transmitted by arthropod vectors continue to pose a significant threat to human health worldwide. The pathogens causing these diseases, do not exist in isolation when they colonize the vector; rather, they likely engage in interactions with resident microorganisms in the gut lumen. The vector microbiota has been demonstrated to play an important role in pathogen transmission for several vector-borne diseases. Whether resident bacteria in the gut of the Ixodes scapularis tick, the vector of several human pathogens including Borrelia burgdorferi, influence tick transmission of pathogens is not determined. We require methods for characterizing the composition of the bacteria associated with the tick gut to facilitate a better understanding of potential interspecies interactions in the tick gut. Using whole-mount in situ hybridization to visualize RNA transcripts associated with particular bacterial species allows for the collection of qualitative data regarding the abundance and distribution of the microbiota in intact tissue. This technique can be used to examine changes in the gut microbiota milieu over the course of tick feeding and can also be applied to analyze expression of tick genes. Staining of whole tick guts yield information about the gross spatial distribution of target RNA in the tissue without the need for three-dimensional reconstruction and is less affected by environmental contamination, which often confounds the sequencing-based methods frequently used to study complex microbial communities. Overall, this technique is a valuable tool that can be used to better understand vector-pathogen-microbiota interactions and their role in disease transmission.

Visualization of Microbiota in Tick Guts by Whole-mount In Situ Hybridization

Here, we present a protocol to spatially and temporally assess the presence of viable microbiota in tick guts using a modified whole-mount in situ hybridization approach.

Infectious diseases transmitted by arthropod vectors continue to pose a significant threat to human health worldwide. The pathogens causing these ...

Genetics

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Visualization of Gut Microbiota-host Interactions via Fluorescence In Situ Hybridization, Lectin Staining, and Imaging

Measuring the localization of microbes within their in vivo context is an essential step in revealing the functional relationships between the microbiota and the vertebrate gut. The spatial landscape of the gut microbiota is tightly controlled by physical features - intestinal mucus, crypts, and folds - and is affected by host-controlled properties such as pH, oxygen availability, and immune factors. These properties limit the ability of commensal microbes and pathogens alike to colonize the gut stably. At the micron-scale, microbial organization determines the close-range interactions between different microbes as well as the interactions between microbes and their host. These interactions then affect large-scale organ function and host health.
This protocol enables the visualization of the gut microbiota spatial organization from distances between cells to organ-wide scales. The method is based on fixing gut tissues while preserving intestinal structure and mucus properties. The fixed samples are then embedded, sectioned, and stained to highlight specific bacterial species through fluorescence in situ hybridization (FISH). Host features, such as mucus and host cell components, are labeled with fluorescently labeled lectins. Finally, the stained sections are imaged using a confocal microscope utilizing tile-scan imaging at high magnification to bridge the micron to centimeter length scales. This type of imaging can be applied to intestinal sections from animal models and biopsies from human tissues to determine the biogeography of the microbiota in the gut in health and disease.

Visualization of Gut Microbiota-host Interactions via Fluorescence In Situ Hybridization, Lectin Staining, and Imaging

This streamlined protocol details a workflow to detect and image bacteria in complex tissue samples, from fixing the tissue to staining microbes with fluorescent in situ hybridization.

Measuring the localization of microbes within their in vivo context is an essential step in revealing the functional relationships between the microbiota ...

Immunology and Infection

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 ...

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission Electron Microscopy (TEM) provided insight into cell ultrastructure, but in the recent decade, the development of modern Scanning Electron Microscopes (SEM) has changed the way of looking inside the cells. Even though the resolution of TEM is superior when protein-level structural details are needed, SEM-resolution is sufficient for the majority of the organelle-level cell biology-related questions. The advancement in technology enabled automatic volume acquisition solutions such as Serial block-face imaging (SBF-SEM) and Focused ion beam SEM (FIB-SEM). Nevertheless, to this day, these methods remain inefficient when the identification and navigation to areas of interest are crucial. Without the means for precise localization of target areas before imaging, operators need to acquire much more data than they need (in SBF-SEM), or, even worse, prepare many grids and image them all (in TEM). We propose the strategy of "lateral screening" using Array Tomography in SEM, which facilitates the localization of areas of interest, followed by automated imaging of the relevant fraction of the total sample volume. Array tomography samples are conserved during imaging, and they can be arranged into section libraries ready for repeated imaging. Several examples are shown in which lateral screening enables us to analyze structural details that are incredibly challenging to access with any other method.

We describe the preparation of ribbons of serial sections and their collection on large transfer support for use as Array Tomography samples, along with automated imaging procedures in a scanning electron microscope. The protocol allows screening, retrieval, and targeted imaging of local, rare events, and the acquisition of large data volumes.

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission ...

High-Precision Mosquito Organ Isolation for Microbiome Analyses

Dissection of Mosquito Ovaries, Midgut, and Salivary Glands for Microbiome Analyses at the Organ Level

The global burden of mosquito-transmitted diseases, including malaria, dengue, West Nile, Zika, Usutu, and yellow fever, continues to increase, posing a significant public health threat. With the rise of insecticide resistance and the absence of effective vaccines, new strategies are emerging that focus on the mosquito's microbiota. Nevertheless, the majority of symbionts remain resistant to cultivation. Characterizing the diversity and function of bacterial genomes in mosquito specimens, therefore, relies on metagenomics and subsequent assembly and binning strategies. The obtention and analysis of Metagenome-Assembled Genomes (MAGs) from separated organs can notably provide key information about the specific role of mosquito-associated microbes in the ovaries (the reproductive organs), the midgut (key for food digestion and immunity), or the salivary glands (essential for the transmission of vector-borne diseases as pathogens must colonize them to enter the saliva and reach the bloodstream during a blood meal). These newly reconstructed genomes can then pave the way for the development of novel vector biocontrol strategies. To this aim, it is required to isolate mosquito organs while avoiding cross-contamination between them or with microorganisms present in other mosquito organs. Here, we describe an optimized and contamination-free dissection protocol for studying mosquito microbiome at the organ level.

Author Spotlight: Optimizing Mosquito Organ Dissection for Studying Symbionts and Vector Microbiomes

Microbial communities in mosquitoes hold great promise for vector biocontrol strategies. Most symbionts are uncultivable, requiring metagenomic analyses. We describe a method to dissect female mosquitoes and separate ovaries, midgut, and salivary glands preventing cross-contamination, facilitating microbiome studies at the organ level, and enhancing understanding of microorganisms' roles in mosquito biology.

The global burden of mosquito-transmitted diseases, including malaria, dengue, West Nile, Zika, Usutu, and yellow fever, continues to increase, posing a ...

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

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