This work was supported by NSF CAREER AWARD 1553023 and NIH/NIAID award R01AI103369.

1. Growth of Pseudomonas aeruginosa Colony Biofilms
<ol>
	<li>Preparation of Medium-Bilayer Plates
	<ol>
		<li>Prepare a 10 g/L tryptone, 10 g/L agar (see Table of Materials) solution in deionized water.</li>
		<li>Autoclave for 20 min. Cool to 50-60 &#176;C in a water bath.</li>
		<li>Pour 45 mL of the agar-tryptone solution into a 100 mm x 100 mm square dish (see Table of Materials) using a 50-mL conical tube. Allow the agar to solidify (~20-30 min). Pour a second, 15-mL layer on top of first layer. Let solidify overnight, removing condensation from lids if necessary by wiping with a lint-free tissue.</li>
	</ol>
	</li>
	<li>Spotting Colony Biofilms
	<ol>
		<li>Streak the strains of interest from freezer stocks onto LB agar plates13 and incubate in the dark for 12-16 h at 37 &#176;C and 80-100% relative humidity.</li>
		<li>For each strain or replicate, use a single colony to inoculate 2 mL of LB and incubate in the dark for 12-16 h at 37 &#176;C with shaking at 250 rpm.</li>
		<li>Sub-culture by diluting 1:100 into fresh LB and incubating in the dark for 2.5-3 h at 37 &#176;C with shaking at 250 rpm.</li>
		<li>Measure optical density using a spectrophotometer and adjust with sterile LB to achieve a cell suspension with an OD500 nm of ~0.5.</li>
		<li>Depending on the desired colony size, pipet 2.5-10 &#181;L of cell suspension onto a medium-bilayer plate (prepared in step 1.1) and allow the spot to dry for ~20 min. Leaving the petri lid ajar near an open flame can facilitate drying.</li>
		<li>Incubate spot colonies at 25 &#176;C and 80-100% relative humidity, in the dark for up to 4 days.</li>
	</ol>
	</li>
</ol>
2. Preparation of the Fixative Solution
<ol>
	<li>Prepare the fixative solution on the day of sample harvesting. Dilute 37% formaldehyde (FA) in 1x PBS to a final concentration of 4% FA. 
	CAUTION: FA is volatile and toxic. Wear protective equipment and work in a well-ventilated fume-hood.</li>
	<li>Immediately before use, dissolve L-lysine hydrochloride in 4% FA (prepared in step 2.1) at room temperature to a final concentration of 50 mM L-lysine HCl.</li>
</ol>
3. Direct Application of Fixative to the Colony Biofilm [Optional]
NOTE: We have found that biofilm morphology is best preserved when the agar overlay is added before fixation. However, this step also constitutes a change in environmental conditions that could affect gene expression. Fluorescent reporter expression patterns should therefore be verified using a separate protocol in which the fixation step is carried out before the addition of the agar overlay, as described here.
<ol>
	<li>On the day of fixation, and working in a well-ventilated fume-hood, pipette the fixative prepared in step 2 directly to the agar surface around the colony&#39;s edge, allowing it to reach the colony&#39;s periphery without submerging it. Apply only as much fixative as is needed to fully surround the colony, approximately 500 &#181;L. Allow the fixative to diffuse into the colony and the surrounding agar.</li>
	<li>Once the fixative has been fully absorbed into the colony and surrounding media, about 20-30 min, and residual FA is no longer visible on the plate, repeat step 3.1. Allow this fixative to diffuse into the colony and the surrounding agar.</li>
	<li>Once the fixative has been fully absorbed into the colony and surrounding media repeat step 3.1, this time applying fixative to the surface of the colony directly. Allow this fixative to diffuse into the colony and the surrounding agar, proceeding to step 4 only after all fixative has been absorbed and is no longer visible on the plate.</li>
</ol>
4. Overlaying Colonies with Agar
<ol>
	<li>On the day of fixation, prepare a 10 g/L agar solution in deionized water and autoclave for 10-20 minutes to dissolve. Cool in a water bath to 50 &#730;C.</li>
	<li>Gently pour 15 mL of the agar solution over the growth medium and colony. Allow the agar to form a gel at room temperature for 5 min (Figure 1B).</li>
	<li>Use a sharp razor blade to cut a square, 3-layer chuck with the colony laminated between the top two layers. If preparing multiple samples, ensure that each chuck is cut to a comparable size.</li>
	<li>Gently remove excess agar from the colony-containing chuck.</li>
	<li>To lift the two upper layers of agar (containing the colony) away from the bottom layer of the chuck (Figure 1C), wet the flat head of a spatula in 1x PBS or water and gently insert between the top and bottom layers. The laminated colony should separate from the bottom layer. Be careful not to distort or bend the laminated colony when lifting it from its base.</li>
</ol>
5. Fixation
<ol>
	<li>Working in a well-ventilated fume-hood, immediately transfer the laminated colony (i.e., 2-layer chuck) into an embedding cassette (see Table of Materials), labeled with a chemical-resistant marking pen. Place the embedding cassette into a glass slide mailer (see table of materials) containing the fixative prepared in steps 2.1-2.2. Make sure there are no air bubbles trapped inside the embedding cassette.</li>
	<li>Incubate the sample in fixative overnight in the dark at room temperature.</li>
</ol>
6. Sample Processing: Buffer Wash, Dehydration, Clearing, and Infiltration
<ol>
	<li>After overnight fixation, wash the sample twice in 1x PBS for 1 h each. For best results, automate reagent homogenization using the spin function of an automatic tissue processor (see table of materials) set to a low setting. If no processor is available, processing can be run manually.</li>
	<li>Follow the buffer wash by dehydration through a graded series of ethanol (EtOH) dilutions in 1x PBS (25%, 50%, 70%, 95%) for 1 h each at room temperature.</li>
	<li>Dehydrate in three washes of 100% EtOH for 1 h each at room temperature.</li>
	<li>Clear the dehydrated sample in three washes of 100% orange oil-based clearing agent (see table of materials) for 1 h each at room temperature.</li>
	<li>Infiltrate the cleared sample twice with molten paraffin wax (see Table of Materials), heated to 55&#730;C, for 2 h each.</li>
</ol>
7. Sample Embedding
<ol>
	<li>Fill a wax mold with molten paraffin wax heated to 55 &#730;C. Use a heated, flat spatula to quickly transfer the infiltrated chuck from the embedding cassette into the wax mold, ensuring that the chuck rests parallel to the base of the mold. Avoid applying excessive pressure onto the chuck. Wax molds may be custom 3-D printed, or are commercially available (see Table of Materials).</li>
	<li>Allow wax to solidify overnight at 4 &#730;C. Samples molded in solid wax may be stored indefinitely at 4 &#730;C.</li>
	<li>Once the wax has solidified, excise the sample from the mold and trim excess wax from around the sample with a razor blade. Leave excess wax extending from one end of the sample, parallel to the plane of sectioning, that can be used to clamp it into the microtome (see Table of Materials). Also leave a sheath of wax, approximately 1-mm thick, around the sample and smooth its faces.</li>
</ol>
8. Sectioning
<ol>
	<li>Heat a water bath to 42 &#730;C. Clamp the sample into the microtome with the surface of the colony oriented perpendicularly to the edge of the blade (see Table of Materials).</li>
	<li>Trim the sample in 50-&#181;m intervals until the desired plane of the colony has been reached, from which sections will be collected.</li>
	<li>Cut a ribbon of the desired number of 10-&#181;m thick sections using a microtome set to a sectioning velocity of 75-80 rpm and a clearance angle of 6-10&#730;. Use a fine-tipped paintbrush to detach the ribbon from the blade.</li>
	<li>Using a drop of water at the tip of a Pasteur pipette (preferred) or forceps, gently transfer the ribbon to the water bath. Be careful to avoid trapping air bubbles under the section.</li>
	<li>Immediately insert a slide into the water bath and position it below the ribbon at a 45&#730; angle.</li>
	<li>Touch the narrow edge of the ribbon to the slide, just below the frosted label. The ribbon should adhere.</li>
	<li>Pull the slide out of the water bath, adjusting the angle to become perpendicular to the surface of the water, and allowing the ribbon to lay flat against the slide along its length. Avoid trapping excess water beneath the ribbon.</li>
	<li>Gently stand the slide onto an absorbent lint-free tissue, wicking excess water from the section.</li>
	<li>Lay the slide on a paper towel, and allow it to dry at room temperature overnight in the dark. Slides can be stored indefinitely at 4 &#730;C in the dark.</li>
</ol>
9. Heat-fixing, Rehydration, and Mounting
<ol>
	<li>Heat a leveled hot plate to 45 &#730;C. Heat the slide for 30-60 min. The wax will become semi-molten and flatten against the slide.</li>
	<li>Gently lift the slide from the hot plate and lay it flat onto a smooth, leveled, room temperature surface, using care to ensure that the molten wax does not pull to either side of the slide, until the wax solidifies (~1 min).</li>
	<li>Using a glass slide mailer, de-wax slides in four washes of clearing agent for 5 min each, using an B&#252;chner aspirator to remove solution between washes.</li>
	<li>Wash the slides three times in 100% EtOH for 1 min each.</li>
	<li>Rehydrate the slides in a graded series of ethanol in the reverse order of processing (95%, 70%, 50%, and 25%) for 1 min each followed by two 1-min washes in 1x PBS.</li>
	<li>Immediately mount the sections in a Tris-buffered mounting medium (see Table of Materials) and apply a coverslip, avoiding the introduction of air bubbles. Allow mounting medium to polymerize overnight at room temperature.</li>
	<li>Once polymerized, seal the coverslip to the slide using clear nail polish. Sealed slides may be stored indefinitely in the dark at 4 &#730;C.</li>
</ol>

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The authors have nothing to disclose.

Paraffin-embedding and thin-sectioning tissue samples is a classic histological technique that enables imaging of micro-morphological structures and is commonly used on eukaryotic tissues, and has been applied with some success to microbial samples8,9. While cryoembedding allows for strong retention of endogenous and immunofluorescent signal, paraffin embedding is generally preferable as it provides better preservation of morphology16. In adapting this method for application to microbial biofilms, we focused on characteristics that are unique to our system. The cells and matrix of tissues are often held together relatively robustly, while biofilms tend to be more delicate; we therefore optimized routine fixation and wax embedding protocols to maximize sample retention. Maintaining the sample on its growth substrate and overlaying the colony with agar immediately prior to fixation ensures that the sample is not lost from the top or the base. However, overlaying colonies with agar that is too hot can damage the colonies, so extra care should be taken during this step.
Fixation is a critical step for retaining structure through the harsh dehydration, clearing, and infiltration steps required for paraffin embedding. We found that lysine-formaldehyde fixation best preserves the integrity of the sample. Soluble lysine, a diamine, can react with aldehydes to form cross-linked polymers and has been proposed to reinforce exopolymeric substance (EPS) against mechanical degradation17,18.
We have observed that background fluorescence decreases with successive washes in the orange oil-based clearing agent, or through prolonged exposure, during sample rehydration (step 9.3). Over-treatment with the clearing agent can, however, result in brittle samples that are difficult to section19. It may be necessary to adjust the volume or number of washes to reflect the amount of wax present during each rehydration. Increased background fluorescence can likewise arise from contamination of the tissue processing reagents, where the wax, which is slightly autofluorescent, is either fully or partially miscible in the clearing solvent or ethanol used in these steps (Steps 6.1-6.4). Additionally, we have found that ethanol diminishes reporter signal, and it is critical to re-constitute the fluorophore in two washes of PBS prior to mounting (step 9.5-9.6).
This method also allows for the localization of constitutive and promoter-driven fluorescence at the resolution of cells. However, we advise caution when using this method to interpret gene expression. The overlay step introduced in this protocol (Step 4.2), applied prior to fixation at 50 &#730;C, presents a considerable environmental change relative to the initial growth conditions that may influence gene expression patterns. It may be useful to verify expression patterns by applying fixative to the colony directly, immediately prior to pouring the overlay, but at a cost to morphology (Optional Step 3.1-3.3).
Another factor influencing the interpretation of fluorescence is the topography of the section itself. Different regions of the biofilm may exhibit varying degrees of structural integrity and be more or less amenable to preservation via fixation, depending upon which, if any, functional groups those components are able to contribute to fixative cross-linking 20. As a result, sections may lose biomass disproportionately from the section surface when transitioning from one morphological or metabolic zone to another along the colony&#39;s z-axis. This may be addressed by imaging an intact focal slice of the section with a confocal microscope.
In addition to applying this technique to strains engineered to express fluorescent proteins, we have examined micromorphological features by transmission electron microscopy (TEM) and in biofilm samples stained with specific fluorescent dyes (Figure 3A). Jennings et al. recently described a dye specific for Pel, the major polysaccharide produced by P. aeruginosa PA14 biofilms 21. As described in the protocol and representative results above, application of this dye to sections allows for visualization of Pel distribution in PA14 colony biofilms at high resolution (Figure 3C) 3. Conversely, fluorescent dyes such as Congo red may be added to the growth medium and imaged post-sectioning (Figure 3B). The application of an overlay also preserves colony biofilm morphology and cell structure throughout processing for TEM (Figure 3D).
The optimized method described here enables the preparation of colony biofilms for sectioning via paraffin embedding, wherein endogenous signal and features or complexed dyes may be localized in vivo with superior resolution while minimizing sample loss and preserving native colony macro- and micro-morphologies. We acknowledge that substantial effort, time and reagent use are required for this method. However, we feel these drawbacks are countered by the degree of morphological preservation, the resolution of in situ fluorescence, and the amenability to pre- and post-staining procedures or alternative processing strategies (such as preparation for TEM) afforded by this technique.

Most microbes have the capacity to form biofilms, communities of cells held together by self-produced matrices. Biofilms can be grown in many types of physical setups, with various regimes of nutrient and substrate provision. Specific assays for biofilm formation tend to yield reproducible multicellular structures, and common architectures are observed for phylogenetically diverse species at the community or macroscopic level. When microbes are grown as colonies on solid medium under an atmosphere, macroscopic morphology conveys information about the capacity for matrix production and often correlates with other traits 1,2,3. The internal architecture of microbial colonies can also provide clues regarding biofilm-specific chemistry and physiology, but has been difficult to characterize. Recent applications of cryoembedding and cryosectioning techniques to bacterial colonies have enabled imaging and visualization of specific features at unprecedented resolution 4,5,6. However, studies with animal tissue have shown that paraffin embedding provides superior preservation of morphology when compared to cryoembedding 7 and has been used to visualize bacteria in tissues 8,9. We have therefore developed a protocol for fixation, paraffin embedding, and thin sectioning of microbial colony biofilms. Here, we will describe the preparation of Pseudomonas aeruginosa PA14 colony-biofilm thin sections 10,11, but we have also successfully applied this technique to biofilms formed by the bacteria Pseudomonas synxantha, Bacillus subtilis, and Vibrio cholerae12.
The process of paraffin-embedding and thin-sectioning biofilms follows a simple logic. First, the biofilms are encased in a layer of agar to preserve morphology during processing. Second, the encased-biofilms are submerged in a fixative to crosslink macromolecules and preserve micromorphology. These are then dehydrated with alcohol, cleared with a more non-polar solvent, and then infiltrated with liquid paraffin wax. Once infiltrated, the samples are embedded into wax blocks for sectioning. Sections are cut, mounted on slides, and then rehydrated in order to return them to a more native state. From this point, they can be stained or covered in mounting medium for microscopic analysis.
This protocol produces thin sections of microbial biofilms suitable for histological analysis. Colony biofilm substructures are visible when thin sections prepared using this method are imaged by light microscopy. Biofilms can also be grown on media containing fluorescent stains specific for individual features or stained at the rehydration step, immediately prior to mounting (steps 9.5-9.6). Finally, microbes can be engineered to produce fluorescent proteins in a constitutive or regulated fashion allowing in situ reporting of cell distribution or gene expression within these communities. We have used these methods to determine colony biofilm depth, cell distribution, matrix distribution, growth patterns, and spatiotemporal gene expression.

<table><tbody><tr><td>5 3/4&quot; Pasteur pipette</td><td>Fisher Scientific</td><td>13-678-6A</td><td>Purchased from univeristy biostores&amp;nbsp;</td></tr><tr><td>Agar&amp;nbsp;</td><td>Teknova&amp;nbsp;</td><td>A7777</td><td></td></tr><tr><td>Buchner Aspirator (Vacuum) Flask&amp;nbsp;</td><td>Pyrex</td><td>5340</td><td>Purchased from univeristy biostores&amp;nbsp;</td></tr><tr><td>Chemically-resistant Marking Pen</td><td>VWR</td><td>103051-182</td><td>Manufacturer: Leica</td></tr><tr><td>Clear Fingernail Polish&amp;nbsp;</td><td>********</td><td>********</td><td>Store bought</td></tr><tr><td>Congo Red Indicator Grade</td><td>VWR</td><td>AAAB24310-14</td><td>Manufacturer: Alfa Aesar</td></tr><tr><td>Coomassie Blue&amp;nbsp;</td><td>VWR</td><td>EM-3340</td><td>Manufacturer: EMD Millipore</td></tr><tr><td>TRIS-buffered Mounting Medium (w/ DAPI)&amp;nbsp;</td><td>Fisher Scientific</td><td>50 247 04</td><td>Manufacturer: Electron Microscopy Sciences</td></tr><tr><td>Embedding Mold&amp;nbsp;</td><td>********</td><td>********</td><td>3D printed in-house</td></tr><tr><td>Embedding Mold (commercial)&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>70182</td><td></td></tr><tr><td>Ethanol 200P</td><td>Decon Labs, Inc.&amp;nbsp;</td><td>2701</td><td>Purchased from univeristy biostores&amp;nbsp;</td></tr><tr><td>Fine-tipped Brush</td><td>********</td><td>********</td><td>Store bought, paint brush</td></tr><tr><td>Glass Coverslips 60x22mm</td><td>Fisher Scientific</td><td>12-519-21C</td><td></td></tr><tr><td>Glass Rehydration Mailer&amp;nbsp;</td><td>Ted Pella</td><td>21043</td><td>20 slide mailer&amp;nbsp;</td></tr><tr><td>Histoclear-II, orange oil-based clearing agent&amp;nbsp;</td><td>Fisher Scientific</td><td>50 899 90150</td><td>Manufacturer: National Diagnostics&amp;nbsp;</td></tr><tr><td>Histosette, Embedding Casette</td><td>Fisher Scientific</td><td>15 182 701A</td><td></td></tr><tr><td>L-lysine hydrochloride&amp;nbsp;</td><td>Fisher Scientific</td><td>BP386 100</td><td></td></tr><tr><td>Low Profile Microtome Blades</td><td>Fisher Scientific</td><td>22 210 048</td><td>Manufacturer: Sturkey&amp;nbsp;</td></tr><tr><td>Micropipette&amp;nbsp;</td><td>VWR</td><td>89080-004</td><td>Promo-pack</td></tr><tr><td>Micropipette Tips&amp;nbsp;</td><td>See comments section</td><td>See comments section</td><td>p10 (Fisher Scientific, 02 707 469), p200 (VWR, 89079-474), p1250 (VWR, 89079-486)</td></tr><tr><td>Microtome&amp;nbsp;</td><td>Fisher Scientific</td><td>905200U/00016050</td><td>Model: HM355S, Manufacturer: Microm, NON-CATALOG, Vendor Catalog # 905200U/00016050</td></tr><tr><td>Formaldehyde, 37% Aqueous (Formalin)</td><td>Ricca Chemical</td><td>RSOF0010-500A</td><td></td></tr><tr><td>Paraplast Xtra (paraffin wax)</td><td>VWR</td><td>15159-486</td><td>Manufacturer: McCormick Scientific&amp;nbsp;</td></tr><tr><td>Petri Dishes Square 100x100x15mm</td><td>Laboratory Disposable Products&amp;nbsp;</td><td>D210-16</td><td></td></tr><tr><td>Potassium chloride&amp;nbsp;</td><td>EMD Chemicals&amp;nbsp;</td><td>PX1405-1</td><td>Component of phosphate buffered saline, prepared in-house&amp;nbsp;</td></tr><tr><td>Potassium phosphate&amp;nbsp;</td><td>Fisher Scientific</td><td>P380-500</td><td>Component of phosphate buffered saline, prepared in-house&amp;nbsp;</td></tr><tr><td>Razor Blades&amp;nbsp;</td><td>VWR</td><td>55411-050</td><td>Purchased from univeristy biostores&amp;nbsp;</td></tr><tr><td>Slide Warmer&amp;nbsp;</td><td>Fisher Scientific</td><td>NC0865259</td><td>NON-CATALOG, Vendor Catalog # 12857D</td></tr><tr><td>Sodium chloride&amp;nbsp;</td><td>VWR</td><td>0241-1KG</td><td>Component of phosphate buffered saline, prepared in-house&amp;nbsp;</td></tr><tr><td>Sodium phosphate&amp;nbsp;</td><td>VWR</td><td>BDH9296.500</td><td>,Component of phosphate buffered saline, prepared in-house&amp;nbsp;</td></tr><tr><td>Suprafrost Histology Slides&amp;nbsp;</td><td>Fisher Scientific</td><td>12-544-2</td><td></td></tr><tr><td>Tissue Flotation Water Bath&amp;nbsp;</td><td>Fisher Scientific</td><td>NC0815797</td><td>Manufacturer: Ted Pella, Vendor Catalog # 28156-B</td></tr><tr><td>Automatic Tissue Processor&amp;nbsp;</td><td>Fisher Scientific</td><td>813160U/Q#00009061</td><td>Model: STP120 Tissue Processor</td></tr><tr><td>Tryptone&amp;nbsp;</td><td>Teknova&amp;nbsp;</td><td>T9012</td><td></td></tr><tr><td>Yeast extract</td><td>Teknova&amp;nbsp;</td><td>Y9010</td><td></td></tr></tbody></table>

paraffin embedding and thin sectioning of microbial colony biofilms for microscopic analysis

Sectioning via paraffin embedding is a broadly established technique in eukaryotic systems. Here we provide a method for the fixation, embedding, and sectioning of intact microbial colony biofilms using perfused paraffin wax. To adapt this method for use on colony biofilms, we developed techniques for maintaining each sample on its growth substrate and laminating it with an agar overlayer, and added lysine to the fixative solution. These optimizations improve sample retention and preservation of micromorphological features. Samples prepared in this manner are amenable to thin sectioning and imaging by light, fluorescence, and transmission electron microscopy. We have applied this technique to colony biofilms of Pseudomonas aeruginosa, Pseudomonas synxantha, Bacillus subtilis, and Vibrio cholerae. The high level of detail visible in samples generated by this method, combined with reporter strain engineering or the use of specific dyes, can provide exciting insights into the physiology and development of microbial communities.

This method generates biofilm thin-sections wherein distinct morphological features and zones of gene expression can be imaged by DIC, fluorescence microscopy, and TEM. While DIC imaging using a 40X oil immersion objective can be sufficient to show some morphological features (Figure 2E), we have found that fluorescence microscopy of strains engineered to constitutively express fluorescent protein provides enhanced visualization of cell distribution within the sample (Figure 2D). Images of individual sections can be stitched together to generate a cross-section of the entire colony (Figure 2B) and to provide context for the localization of structural features within the overall morphology at the macroscopic level (compare Figure 2A, B, and C). Morphological features and fluorescent signals can be measured using imaging software14. Structures within the biofilm or transitions between zones of gene expression can then be correlated with distributions of metabolites or chemical gradients11 .
This protocol can be amended and adapted in several ways. Use of a strain engineered to express fluorescent protein under the control of a specific promoter enables visualization of the distribution of gene expression (Figure 3A). Colonies can be grown on medium containing dyes, or dyes can be added post-sectioning, to stain specific polysaccharides (Figures 3B and C). Finally, samples prepared in a modified version of this protocol can be examined by TEM (Figure 3D) to allow for visualization at higher resolution than that enabled by DIC or fluorescence microscopy.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57196/57196fig1.jpg" /> 
Figure 1: Schematic depicting setup for colony growth prior to fixation. 
(A) Colonies are grown on top of two layers of agar-solidified growth medium (top and bottom) and (B) are then overlaid with an additional layer (overlay), which preserves colony morphology. (C) The colony sandwiched between the overlay and top layer is then separated from the bottom layer for further processing. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57196/57196fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57196/57196fig2.jpg" /> 
Figure 2: Micromorphological features and YFP fluorescence in cross-sections of paraffin-embedded colonies. 
Representative data for a Pseudomonas aeruginosa PA14 &#916;phz colony15 constitutively expressing YFP and grown for 3 days on 1% tryptone, 1% agar containing Congo red and Coomassie blue. (A) Top-view, fluorescence image showing half of a colony prior to fixation. (B) Cross section of the entire colony after paraffin-embedding. The boxed area was imaged with a 10X objective lens (C). The boxed area in (C) was then imaged with a 40X oil immersion objective lens using fluorescence (D) and DIC (E). Scale bars represent 2 mm (A), 500 &#181;m (B), and 100 &#181;m (C-E). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57196/57196fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57196/57196fig3.jpg" /> 
Figure 3: Reporter expression, pre- and post-staining of biofilm features, and TEM imaging of paraffin-embedded colony biofilm samples. 
(A) mexGPr-GFP reporter fluorescence in a P. aeruginosa PA14 colony grown for 3 days on 1% tryptone, 1% agar. (B) Fluorescence of bound Congo red in a P. synxantha colony grown for 5 days on a medium containing the dye. (C) Fluorescence of the Wisteria floribunda lectin stain for Pel polysaccharide in a P. aeruginosa PA14 &#916;phz colony (grown for 2 days on 1% tryptone, 1% agar containing Congo red and Coomassie blue), applied after sectioning. (D) TEM image of a P. aeruginosa PA14 &#916;phz colony (grown for 3 days on 1% tryptone, 1% agar containing Congo red and Coomassie blue), and processed for sectioning via paraffin embedding*. Scale bar represents 40 &#181;m (A-B), 20 &#181;m (C), and 5 &#181;m (D). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57196/57196fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
*This sample was processed for sectioning via paraffin-embedding through Step 4 of the above protocol, omitting Steps 4.1-4.3, and then processed separately for TEM analysis.

Watch this Scientific Journal Video about Paraffin Embedding and Thin Sectioning of Microbial Colony Biofilms for Microscopic Analysis at JoVE.com

Paraffin Embedding and Thin Sectioning of Microbial Colony Biofilms for Microscopic Analysis

We describe fixation, paraffin embedding, and thin sectioning techniques for microbial colony biofilms. In prepared samples, biofilm substructure and reporter expression patterns can be visualized by microscopy.

Sectioning via paraffin embedding is a broadly established technique in eukaryotic systems. Here we provide a method for the fixation, embedding, and ...

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Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

14.3K Views.  Columbia University. We beschrijven fixatie, paraffine insluiten en dunne vectorafbeeldingsbestanden technieken voor microbiële kolonie biofilms. In bereid monsters, kunnen biofilm substructuur en verslaggever expressiepatronen worden gevisualiseerd door microscopie.

Video: Paraffin Embedding and Thin Sectioning of Microbial Colony Biofilms for Microscopic Analysis

A Method for Obtaining Serial Ultrathin Sections of Microorganisms in Transmission Electron Microscopy

Observing cells and cell components in three dimensions at high magnification in transmission electron microscopy requires preparing serial ultrathin sections of the specimen. Although preparing serial ultrathin sections is considered to be very difficult, it is rather easy if the proper method is used. In this paper, we show a step-by-step procedure for safely obtaining serial ultrathin sections of microorganisms. The key points of this method are: 1) to use the large part of the specimen and adjust the specimen surface and knife edge so that they are parallel to each other; 2) to cut serial sections in groups and avoid difficulty in separating sections using a pair of hair strands when retrieving a group of serial sections onto the slit grids; 3) to use a 'Section-holding loop' and avoid mixing up the order of the section groups; 4) to use a 'Water-surface-raising loop' and make sure the sections are positioned on the apex of the water and that they touch the grid first, in order to place them in the desired position on the grids; 5) to use the support film on an aluminum rack and make it easier to recover the sections on the grids and to avoid wrinkling of the support film; and 6) to use a staining tube and avoid accidentally breaking the support films with tweezers. This new method enables obtaining serial ultrathin sections without difficulty. The method makes it possible to analyze cell structures of microorganisms at high resolution in 3D, which cannot be achieved by using the automatic tape-collecting ultramicrotome method and serial block-face or focused ion beam scanning electron microscopy.

This study presents reliable and easy procedures for obtaining serial ultrathin sections of a microorganism without expensive equipment in transmission electron microscopy.

Observing cells and cell components in three dimensions at high magnification in transmission electron microscopy requires preparing serial ultrathin ...

Engineering

Methodologies for Studying B. subtilis Biofilms as a Model for Characterizing Small Molecule Biofilm Inhibitors

This work assesses different methodologies to study the impact of small molecule biofilm inhibitors, such as D-amino acids, on the development and resilience of Bacillus subtilis biofilms. First, methods are presented that select for small molecule inhibitors with biofilm-specific targets in order to separate the effect of the small molecule inhibitors on planktonic growth from their effect on biofilm formation. Next, we focus on how inoculation conditions affect the sensitivity of multicellular, floating B. subtilis cultures to small molecule inhibitors. The results suggest that discrepancies in the reported effects of such inhibitors such as D-amino acids are due to inconsistent pre-culture conditions. Furthermore, a recently developed protocol is described for evaluating the contribution of small molecule treatments towards biofilm resistance to antibacterial substances. Lastly, scanning electron microscopy (SEM) techniques are presented to analyze the three-dimensional spatial arrangement of cells and their surrounding extracellular matrix in a B. subtilis biofilm. SEM facilitates insight into the three-dimensional biofilm architecture and the matrix texture. A combination of the methods described here can greatly assist the study of biofilm development in the presence and absence of biofilm inhibitors, and shed light on the mechanism of action of these inhibitors.

Methodologies for Studying B. subtilis Biofilms as a Model for Characterizing Small Molecule Biofilm Inhibitors

This study presents the development of reproducible methodologies to study biofilm inhibitors and their effects on Bacillus subtilis multicellularity.

This work assesses different methodologies to study the impact of small molecule biofilm inhibitors, such as D-amino acids, on the development and ...

A New Method for Qualitative Multi-scale Analysis of Bacterial Biofilms on Filamentous Fungal Colonies Using Confocal and Electron Microscopy

Bacterial biofilms frequently form on fungal surfaces and can be involved in numerous bacterial-fungal interaction processes, such as metabolic cooperation, competition, or predation. The study of biofilms is important in many biological fields, including environmental science, food production, and medicine. However, few studies have focused on such bacterial biofilms, partially due to the difficulty of investigating them. Most of the methods for qualitative and quantitative biofilm analyses described in the literature are only suitable for biofilms forming on abiotic surfaces or on homogeneous and thin biotic surfaces, such as a monolayer of epithelial cells.
While laser scanning confocal microscopy (LSCM) is often used to analyze in situ and in vivo biofilms, this technology becomes very challenging when applied to bacterial biofilms on fungal hyphae, due to the thickness and the three dimensions of the hyphal networks. To overcome this shortcoming, we developed a protocol combining microscopy with a method to limit the accumulation of hyphal layers in fungal colonies. Using this method, we were able to investigate the development of bacterial biofilms on fungal hyphae at multiple scales using both LSCM and scanning electron microscopy (SEM). This report describes the protocol, including microorganism cultures, bacterial biofilm formation conditions, biofilm staining, and LSCM and SEM visualizations.

This protocol describes a new method to grow and qualitatively analyze bacterial biofilms on fungal hyphae by confocal and electron microscopy.

Bacterial biofilms frequently form on fungal surfaces and can be involved in numerous bacterial-fungal interaction processes, such as metabolic ...

Yeast Colony Embedding Method

Patterning of different cell types in embryos is a key mechanism in metazoan development. Communities of microorganisms, such as colonies and biofilms also display patterns of cell types. For example, in the yeast S. cerevisiae, sporulated cells and pseudohyphal cells are not uniformly distributed in colonies. The functional importance of patterning and the molecular mechanisms that underlie these patterns are still poorly understood.
One challenge with respect to investigating patterns of cell types in fungal colonies is that unlike metazoan tissue, cells in colonies are relatively weakly attached to one another. In particular, fungal colonies do not contain the same extensive level of extracellular matrix found in most tissues . Here we report on a method for embedding and sectioning yeast colonies that reveals the interior patterns of cell types in these colonies. The method can be used to prepare thick sections (0.5 &mu;) useful for light microscopy and thin sections (0.1 &mu;) suitable for transmission electron microscopy. Asci and pseudohyphal cells can easily be distinguished from ovoid yeast cells by light microscopy , while the interior structure of these cells can be visualized by EM.
The method is based on surrounding colonies with agar, infiltrating them with Spurr's medium, and then sectioning. Colonies with a diameter in the range of 1-2 mm are suitable for this protocol. In addition to visualizing the interior of colonies, the method allows visualization of the region of the colony that invades the underlying agar.

A method for embedding yeast colonies allowing sectioning for light and electron microscopy. This protocol allows determination of the distribution of sporulated cells and pseudohyphal cells within colonies providing a new tool toward understanding the organization of cell types within a fungal community.

Patterning of  different cell types in embryos is a key mechanism in metazoan development. Communities of microorganisms, such as colonies and biofilms ...

Biology

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in every tissue to be identified and characterized. Our laboratory is actively working to make technological advances in X-ray micro-computed tomography (micro-CT) that will bring the diagnostic power of histology to the study of full tissue volumes at cellular resolution (i.e., an X-ray Histo-tomography modality). Toward this end, we have made targeted improvements to the sample preparation pipeline. One key optimization, and the focus of the present work, is a straightforward method for rigid embedding of fixed and stained millimeter-scale samples. Many of the published methods for sample immobilization and correlative micro-CT imaging rely on placing the samples in paraffin wax, agarose, or liquids such as alcohol. Our approach extends this work with custom procedures and the design of a 3-dimensional printable apparatus to embed the samples in an acrylic resin directly into polyimide tubing, which is relatively transparent to X-rays. Herein, sample preparation procedures are described for the samples from 0.5 to 10 mm in diameter, which would be suitable for whole zebrafish larvae and juveniles, or other animals and tissue samples of similar dimensions. As proof of concept, we have embedded the specimens from Danio, Drosophila, Daphnia, and a mouse embryo; representative images from 3-dimensional scans for three of these samples are shown. Importantly, our methodology leads to multiple benefits including rigid immobilization, long-term preservation of laboriously-created resources, and the ability to re-interrogate samples.

We developed protocols and designed a custom apparatus to enable embedding of millimeter-scale specimens. We present sample preparation procedures with an emphasis on embedding in acrylic resin and polyimide tubing to achieve rigid immobilization and long-term storage of specimens for the interrogation of tissue architecture and cell morphology by micro-CT.

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in ...

Bioengineering

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

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

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

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

Histological Sample Preparation for Light Microscopy

Histology is the study of cells and tissues, which is typically aided by the use of a light microscope. The preparation of histological samples can vary greatly based on the inherent properties of the samples such as size and hardness as well as expected post-processing which includes planned staining techniques or other down-stream applications. As described in this video, specimen preparation typically begins with a fixation procedure to prevent degradation of the sample by naturally occurring enzymes that are released by the cells upon death. Once fixed, samples are placed into an embedding medium that is able to sufficiently support the sample. Most commonly this is paraffin wax, but other materials such as a glycerin based freezing medium and agars are also used to surround the sample during sectioning. Sectioning then takes place on a microtome or other cutting device that allows the user to shave the sample into thin slices ranging from a few microns to a few millimeters in thickness. Once cut, sections are mounted on a glass slide and stained to bring out specific features of the sample before being imaged on a microscope.

Histological Sample Prep for a Light Microscope

Histology is the study of cells and tissues, which is typically aided by the use of a light microscope.  The preparation of histological samples can vary ...

Education

JoVE Science Education

Basic Biology

General Laboratory Techniques

Formalin-Fixed Paraffin Embedded (FFPE) Tissue Preservation: A Method for Studying Tissue Morphology

Source: Nagaraj A.S., et al., <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/58569/establishment-analysis-tumor-slice-explants-as-prerequisite-for">Establishment and Analysis of Tumor Slice Explants As a Prerequisite for Diagnostic Testing.</a> J. Vis. Exp. (2018)
The following video describes how to preserve morphology of a tissue with formalin fixation and paraffin embedding.

Source: Nagaraj A.S., et al., Establishment and Analysis of Tumor Slice Explants As a Prerequisite for Diagnostic Testing. J. Vis. Exp. (2018)
The ...

JoVE Encyclopedia of Experiments

Cancer Research

Lung Cancer

Ex vivo study

A Method for Sample Preparation to Visualize Biofilm Forming Bacteria on Fungal Hyphae

Source: Guennoc, Cora Miquel, et al, <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/54771">A New Method for Qualitative Multi-scale Analysis of Bacterial Biofilms on Filamentous Fungal Colonies Using Confocal and Electron Microscopy.</a> J. Vis. Exp. (2017).
This video describes a method to grow and qualitatively analyze bacterial biofilms on fungal hyphae using microscopy. Bacterial biofilms frequently form on fungal surfaces and involve numerous bacterial-fungal interaction processes, such as metabolic cooperation, competition, or predation.

1. Preparation of an In Vitro Biofilm of Bacteria on the Fungal Colony

	Centrifuge the bacterial culture at 5,000 x g for 3 min and suspend the pellet in ...

Immunology

Immunodiagnostics

In Vitro and In Vivo Models

Fixation and Sectioning

Two basic types of preparation are used to visualize specimens with a light microscope: wet mounts and fixed specimens.
The simplest type of preparation is the wet mount, in which the specimen is placed in a drop of liquid on the slide. A liquid specimen can be directly deposited on the slide using a dropper. Solid specimens, such as skin scraping, can be placed on the slide before adding a drop of liquid to prepare the wet mount. Sometimes the liquid is simply water, but stains are often added to enhance contrast. Once the liquid has been added to the slide, a coverslip is placed on top, and the specimen is ready for examination under the microscope.
The second method of preparing specimens for light microscopy is fixation. The &#8220;fixing&#8221; of a sample refers to the process of attaching cells to a slide. Fixation is often achieved by heating (heat fixing) or chemically treating the specimen. In addition to attaching the specimen to the slide, fixation also kills microorganisms in the specimen, stopping their movement and metabolism while preserving the integrity of their cellular components for observation.
To heat-fix a sample, a thin layer of the specimen is spread on the slide (called a smear), and the slide is then briefly heated over a heat source. Chemical fixatives are often preferable to heat for tissue specimens. Chemical agents such as acetic acid, ethanol, methanol, formaldehyde (formalin), and glutaraldehyde can denature proteins, stop biochemical reactions, and stabilize cell structures in tissue samples.
In addition to fixation, staining is almost always applied to color certain features of a specimen before examining it under a light microscope. Stains, or dyes, contain salts made up of a positive ion and a negative ion. Depending on the type of dye, the positive or the negative ion may be the chromophore (the colored ion); and the other uncolored ion is called the counterion. If the chromophore is the positively charged ion, the stain is classified as a basic dye; if the negative ion is the chromophore, the stain is considered an acidic dye.
Dyes are selected for staining based on the chemical properties of the dye and the specimen being observed, which determine how the dye will interact with the specimen. In most cases, it is preferable to use a positive stain, a dye that will be absorbed by the cells or organisms being observed, adding color to objects of interest to make them stand out against the background. However, there are scenarios where it is advantageous to use a negative stain absorbed by the background but not by the cells or organisms in the specimen. Negative staining produces an outline or silhouette of the organisms against a colorful background.
Because cells typically have negatively charged cell walls, the positive chromophores in basic dyes tend to stick to the cell walls, making them positive stains. Thus, basic dyes such as basic fuchsin, crystal violet, malachite green, methylene blue, and safranin typically serve as positive stains. On the other hand, the negatively charged chromophores in acidic dyes are repelled by negatively charged cell walls, making them negative stains. Commonly used acidic dyes include acid fuchsin, eosin, and rose bengal.
Some staining techniques involve the application of only one dye to the sample; others require more than one dye. In simple staining, a single dye is used to emphasize particular structures in the specimen. A simple stain will generally make all of the organisms in a sample appear the same color, even if the sample contains more than one type of organism. In contrast, differential staining distinguishes organisms based on their interactions with multiple stains. In other words, two organisms in a differentially stained sample may appear different colors. Differential staining techniques commonly used in clinical settings include Gram staining, acid-fast staining, endospore staining, flagella staining, and capsule staining.
This text is adapted from <a href="https://openstax.org/books/microbiology/pages/2-4-staining-microscopic-specimens">Openstax, Microbiology 2e, Section 2.4: </a><a href="https://openstax.org/books/microbiology/pages/2-4-staining-microscopic-specimens">Staining microscopic specimens</a>

Two basic types of preparation are used to visualize specimens with a light microscope: wet mounts and fixed specimens.
The simplest type of preparation ...

Paraffin Embedding and Thin Sectioning of Microbial Colony Biofilms for Microscopic Analysis

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