This work was supported by the French National Research Agency through the Laboratory of Excellence ARBRE (ANR-11-LABX-0002-01), the Plant-Microbe Interfaces Scientific Focus Area in the Genomic Science Program, and the Office of Biological and Environmental Research in the DOE Office of Science. Oak Ridge National Laboratory is managed by UT-Battelle, LLC, for the United States Department of Energy under contract DE-AC05-00OR22725.

1. Preparation of Bacterial and Fungal Cultures
<ol>
	<li>Preparation of fungal cultures
	<ol>
		<li>Prepare cellophane membranes with the same diameter as the Petri dishes and boil them in EDTA (1 g/L in deionized water) for 30 min. Rinse the sheets with deionized water and autoclave them twice.</li>
		<li>Prepare a fungal pre-culture by inoculating fungal agar plugs on the appropriate agar medium covered with an EDTA pre-treated cellophane membrane. Incubate it at the optimal growth temperature to obtain colonies approximately 1 cm in diameter.
		<ol>
			<li>For L. bicolor S238N, incubate at 25 &#176;C for 5 days on Pachlewski P5 agar medium, made of 0.5 g of diNH4 tartrate, 1 g of KH2PO4, 0.5 g of MgSO4.7H2O, 5 g of Maltose D+, 20 g of Glucose D+, 1 ml of 10% Thiamine-HCl, 1 ml of 1/10 diluted micronutrient solution (6 g/L ferrous sulfate; 0.27 g/L molybdenum trioxyde; 8.45 g/L boric acid; 5 g/L manganese sulfate; 5 g/L cupric sulfate; 2.27 g/L zinc sulfate) made up to 1 L with deionized water; and 20 g of agar; pH 5.5.</li>
		</ol>
		</li>
		<li>From the fungal pre-culture, inoculate a new agar plate covered with the EDTA pre-treated cellophane membrane containing a low-carbon agar medium to promote the radial expansion of the colonies .
		<ol>
			<li>To inoculate the Petri dish, gently scratch the external area (where cells have the same physiological state) of a fungal colony from the pre-culture with a scalpel and transfer the hyphae to the new agar plate.</li>
			<li>Incubate at the optimal growth temperature until the new colonies are approximately 1 cm in diameter. For L. bicolor S238N, incubate them at 25 &#176;C for 5 days on P20 agar medium (0.25 g of diNH4 tartrate; 0.5 g of KH2PO4; 0.25 g of MgSO4.7H2O; 1 g of Glucose D+; 0.5 ml of 10% Thiamine-HCl; 0.5 ml of 1/10 diluted micronutrient solution (cf 1.1.2.1), made up to 1 L with deionized water; and 20 g of agar; pH 5.5). 
			Note: Due to the pre-culture on cellophane, the second culture will not contain agar media in the central plugs. The agar plugs should be removed for further analysis of the biofilms, since these plugs introduce agar and nutrients that can create biases in the analysis. This step concerns only fungal species that are kept and propagated on agar plates, and it is not necessary for fungi that are propagated from spores or frozen stocks.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparation of bacterial cultures
	<ol>
		<li>Use a sterile loop to collect 2 to 3 individual bacterial colonies from a culture on the appropriate agar medium and inoculate 25 ml of liquid Luria-Bertani (LB) medium (or any other appropriate medium, depending on the strain used). For P. fluorescens BBc6, incubate an overnight culture at 28 &#176;C with gentle shaking (~150 rpm). 
		Note: Using a strain tagged with fluorescent protein, such as GFP, is preferable, because this avoids performing a double-staining (fungus and bacteria) prior to microscopic observations. The timing, the stirring speed, and the temperature of the incubation depend on the strain used, the goal being to obtain a culture in late exponential growth.</li>
	</ol>
	</li>
</ol>
2. Preparation of N In Vitro Biofilm of Bacteria on the Fungal Colony
<ol>
	<li>Centrifuge the bacterial culture at 5,000 x g for 3 min and suspend the pellet in 25 ml of sterile 0.1 M potassium phosphate buffer (25 g/L KH2PO4 and 2.78 g/L K2HPO4 , pH 5.8). Repeat this step once and adjust the final bacterial concentration to 109 cells/ml with the same buffer.</li>
	<li>Fill a 6-well microplate with 5 ml of the bacterial suspension (or sterile 0.1 M potassium phosphate buffer for the negative control).</li>
	<li>Cut the cellophane membrane of the fungal culture with a sterile razor blade to obtain squares of cellophane with a single fungal colony on each membrane. Carefully remove the cellophane squares containing hyphae from the solid medium using forceps and transfer the square of cellophane containing hyphae to a well of the microplate containing the bacterial suspension.</li>
	<li>Gently shake the microplate while the fungal colonies are still attached to the cellophane; then, remove the cellophane sheets, leaving the fungal colonies in the plate.</li>
	<li>Incubate the microplate with gentle agitation (~60 rpm) at a temperature adapted to the studied microorganisms. 
	Note: The time of incubation depends on the speed of establishment of the biofilm and the stage to be analyzed. For P. fluorescens BBc6/L. bicolor, incubate at 20 &#176;C for 30 min to get early-stage biofilms and up to 16 hr to get mature biofilms.</li>
</ol>
3. Laser Scanning Confocal Microscopy Analysis of the Biofilm Formation
<ol>
	<li>Sample preparation
	<ol>
		<li>To remove planktonic bacteria and bacteria electrostatically-attached to the hyphae, rinse the fungus by transferring it to a new 6-well microplate filled with 5 ml of a strong salt solution (NaCl, 17 g/L) 17 ; gently shake for 1 min.</li>
		<li>Transfer the fungus to a new 6-well microplate containing 5 ml of sterile 0.1 M potassium phosphate buffer, gently shake for 2 min, and transfer the fungus to fresh potassium phosphate buffer.</li>
		<li>Cut the part of the fungal colony to be imaged with a scalpel while keeping it in the potassium phosphate buffer, such that the obtained section contains about half of the fungal colony.</li>
		<li>To stain the sample, transfer it to a Petri dish filled with sterile water containing an appropriate fluorescent dye (depending on the aim of the analysis), and incubate it in the dark.
		<ol>
			<li>To visualize the fungal network, use, for example, Congo Red (0.1% final concentration, 5 min of incubation), wheat germ agglutin (WGA) lectin conjugated to Alexa Fluor 633 (10 &#181;g/ml final concentration, 10 min of incubation) or FUN1 (10 &#181;M final concentration, 10 min of incubation).</li>
			<li>If using a non-fluorescent-tagged bacterium, counterstain the bacterial cells with a DNA-specific fluorescent probe among the numerous cell-permeant DNA dyes commercially available. For example, use DAPI (0.25 &#181;g/ml final concentration, 10 min of incubation).</li>
			<li>To visualize the matrix proteins, use a protein stain21.</li>
		</ol>
		</li>
		<li>After staining, rinse the sample by transferring it to a Petri dish lid containing 10 ml of sterile 0.1 M potassium phosphate and gently shake for 1 min.</li>
		<li>Half submerge a slide in the Petri dish lid and delicately bring the cut section to float above the slide. Then, slowly remove the slide from the buffer solution, allowing the sample to gently settle on the slide. 
		Note: For this step, it is very important to proceed gently to avoid biofilm disturbance.</li>
		<li>Finally, add 10 &#956;l of anti-fading mounting medium to the sample and cover it with a glass coverslip. 
		Note: Once the slide is prepared, the microscopic analysis must be performed as soon as possible (within at most 30 min) to avoid modification in the biofilm structure.</li>
	</ol>
	</li>
	<li>Confocal microscopic analysis
	<ol>
		<li>Examine the slides under a confocal laser microscope with a 10X or 40X objective lens coupled to imaging software. 
		Note: Here, a multi-beam scanning confocal microscope equipped with 405, 488, and 561 nm excitation lasers, a GaAsp PMT detector, and 10X 0.3 NA and 40X 1.2 NA objectives was used.
		<ol>
			<li>Perform imaging using parameters adapted to the type of data requested and to the time constrains. Use laser excitation/emission wavelengths according to the stain used and to the manufacturer&#39;s recommendations. Use a combination of tile scan and the Z-stack functions to obtain global 3D views of the fungal colony and the biofilms. 
			Note: Images on Figure 3 were acquired using the following parameters: 8-bit/pixel, averaging = 2, pixel dwell = 1.58 &#181;sec, tiles scan of 5x5 images with online stitching (10% coverage, threshold = 0.7); Z-step = 2 &#181;m.</li>
		</ol>
		</li>
		<li>For 3D data visualization, use one of the many free or commercial softwares (these offer many options for 3D rendering).
		<ol>
			<li>For example, to display Z-stack data as 2D maximum intensity projections, subtract the bright-field channel, adjust the brightness and contrast, and merge channels to create a composite image. If they are present, eliminate slices without signal at the Z-stack extremities. Then, perform a 2D maximum-intensity projection and convert the result into RGB color. Use the &#34;Z project&#34; function in the ImageJ software22 to obtain the images presented here by 2D maximum intensity projections.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Electron Microscopy Analysis
<ol>
	<li>Prepare the samples as described for LSCM, except in step 3.1.2, transfer the biofilms to sterile water instead of potassium phosphate buffer after the rinse steps. This avoids the formation of salt crystals during the dehydration step, which would perturb SEM imaging.</li>
	<li>Transfer the biofilms to a sample holder and remove excess water; only allow a small pellicle of water to cover the sample.</li>
	<li>Transfer the sample to the chamber of a variable-pressure scanning electron microscope (VP-SEM); freeze it to -50 &#176;C on a Peltier cooling stage; and allow it to slowly freeze-dry, directly in the SEM chamber, with 100 Pa of variable pressure set for 15 hr.</li>
	<li>After completion of the freeze-drying, retrieve the sample and transfer it into a high-vacuum film deposition system. Coat the sample with 2 nm of platinum (quartz measurement) under argon plasma (2.5 x 10-2 mbar, 35 mA).</li>
	<li>Observe the coated sample with an SEM equipped with a field emission gun (FEG) using the high resolution &#34;in lens&#34; detector and an electronic high tension of 1 kV.</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microplate+spectrofluorometric+assay+for+bacterial+biofilms.">A microplate spectrofluorometric assay for bacterial biofilms.</a> J. Ind. Microbiol. Biotechnol. 34 (1), 1-4 (2007).</li><li>Berne, C., Ducret, A., Hardy, G. G., Brun, Y. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adhesins+Involved+in+Attachment+to+Abiotic+Surfaces+by+Gram-Negative+Bacteria.">Adhesins Involved in Attachment to Abiotic Surfaces by Gram-Negative Bacteria.</a> Microbiol. Spectr. 3 (4), 1-45 (2015).</li><li>Schlafer, S., Garcia, J. E., Greve, M., Raarup, M. K., Nyvad, B., Dige, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ratiometric+imaging+of+extracellular+pH+in+bacterial+biofilms+with+C-SNARF-4.">Ratiometric imaging of extracellular pH in bacterial biofilms with C-SNARF-4.</a> Appl. Environ. Microbiol. 81 (4), 1267-1273 (2015).</li><li>Daddi Oubekka, S., Briandet, R., Fontaine-Aupart, M. P., Steenkeste, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+time-resolved+fluorescence+microscopy+to+assess+antibiotic+diffusion-reaction+in+biofilms.">Correlative time-resolved fluorescence microscopy to assess antibiotic diffusion-reaction in biofilms.</a> Antimicrob. Agents Chemother. 56 (6), 3349-3358 (2012).</li><li>Almeida, C., Azevedo, N. F., Santos, S., Keevil, C. W., Vieira, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Discriminating+multi-species+populations+in+biofilms+with+peptide+nucleic+acid+fluorescence+in+situ+hybridization+(PNA+FISH).">Discriminating multi-species populations in biofilms with peptide nucleic acid fluorescence in situ hybridization (PNA FISH).</a> PLoS One. 6 (3), (2011).</li><li>Malic, S., Hill, K. E., Hayes, A., Percival, S. L., Thomas, D. W., Williams, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+and+identification+of+specific+bacteria+in+wound+biofilms+using+peptide+nucleic+acid+fluorescent+in+situ+hybridization+(PNA+FISH).">Detection and identification of specific bacteria in wound biofilms using peptide nucleic acid fluorescent in situ hybridization (PNA FISH).</a> Microbiology. 155 (8), 2603-2611 (2009).</li></ol>

The authors have nothing to disclose.

Bacterial biofilms are retrieved in many environments and have been studied since the 1950s, leading to the development of a number of methods to analyze them24. Classical methods to quantify and monitor biofilms include micro-titer assays and, the most widely-used method, crystal violet (CV) staining. These methods are fast, low cost, and easy to handle25 and are particularly useful to quantify total biofilm biomass or to perform viability and matrix quantification assays. On an other hand, &#34;omics&#34; methods are also useful in biofilm studies, allowing for quantitative and functional analyses of biofilms26,27. Despite the advantages of micro-titer plate and &#34;omics&#34; methods, several essential features of biofilms cannot be captured with these techniques, hindering a complete understanding of this process. Such features include matrix structures, bacterial colony architectures, cell/cell interactions, and colonization patterns, which are key data for understanding both the functioning of biofilms and the dynamics of their formation. Despite the capacity of microscopy to capture these features, microscopy analysis of bacterial biofilms on filamentous fungi are still scarce. This is mainly due to the growth of filamentous fungi, which often forms colonies of thick, complex, tridimensional networks. Formation of bacterial biofilms on fungi is common in diverse environments and is significantly involved in various fields4 (e.g., medicine, agriculture, and environment); hence, it is critical to develop new methods to facilitate their investigation. To this end, we combined a method to generate very thin fungal colonies with microscopic imaging of the bacterial biofilms. In addition, we proposed a set of microscopy tools to qualitatively analyze those biofilms. The success of the method relies upon the ability to produce very thin hyphal colonies and to apply the appropriate dyes. These points are discussed below.
Due to the complex structures of the biofilms, understanding their function requires a multi-scale approach28,29. Distribution patterns of the biofilms, bacterial colony architecture, and matrix structure and composition are analyzed at different scales (i.e., meso-scale and micro-scale). Moreover, nanoscale resolution allows access to the cell/cell physical interactions and the nano-structure of the matrix. Thus, the developed method easily enables a multi-scale analysis of the bacterial biofilms formed on the fungal colony.
In most studies, LSCM analyses of biofilms are limited to the micro-scale, the meso-scale usually being performed by optical coherence tomography30,31,32. The method presented here enables both micro- and meso-scale analyses by LSCM. It demonstrates the utility of combining both analyses in the same region of the sample and even on the same image using new-generation confocal microscopes with high resolution (Figure 3). Thus, issues linked to compiled data gathered at different scales with different methods are here avoided.
This combination of analyses gave access concomitantly to the biofilm repartition on the fungal colony, the bacterial colony architecture along the developing biofilm, and the matrix structure. The meso-scale analysis showed a heterogenic distribution of the bacterial biofilms on the fungal colonies (Figures 2 and 3). This observation would not have been possible with protocols that only permit imaging of a small portion of the fungal colony, which is not necessarily representative of the entire colony. Thus, while often neglected, the meso-scale analysis can give precious information about biofilm distribution patterns.
Finally, the developed method can be used to analyze samples with different microscopy techniques, including scanning electron microscopy. Here, SEM was used to reach the nano-scale and to obtain the bacterial spatial organization within the biofilm. It performed very well with the thin fungal colonies, while SEM only permitted surface imaging. In contrast to LSCM, SEM, however, required sample dehydration and, most often, coating with a conductive metal. This dehydration process might alter biological structures when it is not properly executed and may require optimization. Here, sample dehydration using slow lyophilization was used33. Nevertheless, applying both LSCM and SEM to the samples will allow the performance of correlative microscopy at the same location of the sample.
Despite the advantages described above, some limitations exist. Firstly, it may not be applicable to all kind of fungi. Indeed, this culturing method is developed for fungi spreading radially on the surface of solid media. This method may not be suitable for fungi forming mainly aerial hyphae (e.g., Fusarium sp.) or for micro-aerobic fungi spreading mainly inside agar. Moreover, fungi degrading cellophane may be problematic as well (e.g., Trichoderma sp.). Secondly, it is important to note that the staining strategy is a critical point and the choice of the stain must be made carefully, as the stain must not disturb the biofilm. For example, we noticed that Calcofluor White caused partial biofilm disruption (data not shown), likely due to the high pH of this stain. Also, some dyes produced heterogeneous staining (e.g., Congo Red), while others produced homogeneous staining (e.g., cell wall staining with WGA lectin), giving a heterogeneous image quality. Moreover, it is important to be aware that some dyes might not be fully specific. For example, WGA stains not only fungal cell walls but also N-acetylneuraminic acid in gram-positive bacterial cell walls and adhesins produced by gram-positive and -negative bacteria during biofilm formation34,35. Therefore, using fluorescent protein-tagged bacteria and/or fungi is recommended to avoid multiple staining. If multiple dyes are used, they must not chemically interfere, and their emission spectra should not overlap.
Meso-scale analyses require a large scanned area, and therefore, LSCM may be time-consuming (40 min to 1 hr, depending on the sample thickness) and bottleneck the analysis of a large number of samples. Nonetheless, adjustments can be made depending on the type of data required. It is possible to decrease acquisition time and image size by altering the image quality. For example, high resolution is not necessary to analyze the biofilm general repartition.
Finally, some limitations need to be considered when choosing to display Z- stack data as 2D or 3D projections. Two-dimensional projections are a good way to summarize data, but depth information is lost, and overlapped structures become hidden. On the other hand, 3D projections allow the visualization from different points of view, but they often render poorly in case of spatial complexity.
In conclusion, we have reported a method for the characterization of bacterial biofilms on hyphae at the structural level. The methodology can be extended to other applications. Indeed, this method allows the performance of functional or chemical characterization of bacterial biofilms forming on fungal hyphae. Due to the great variety of existing fluorescent reporter systems, LSCM analysis can be used for multiple purposes29. For example, the fluorescence microscopy could be used to monitor pH gradient36 or molecule diffusion in biofilms37. Additionally, the method allows for community analysis in multispecies biofilms. For example, fluorescence in situ hybridization targeting specific bacterial groups is particularly useful to study specific bacterial repartition in multispecies biofilms38,39. Last, numerous fluorescent dyes can be used to characterize the matrix composition of the biofilms21. Here, proteins were targeted using Sypro, which stains a large range of proteins, among them matrix proteins (Figure 4), but other dyes allow for the visualization of other important matrix constituents, such as exopolysaccharides or extracellular DNA. Interestingly, all these analyses could be performed at the meso-scale using the described method. Since LSCM can be performed on living samples, it is also possible to achieve time-lapse imaging using, for example, coverwell chambers, particularly suitable for thin fungal colonies. This option is particularly interesting, as biofilm formation is a complex, dynamic process. Finally, for a quantitative purpose, the reported method may improve the accuracy of automatic quantitative analysis by making this quantification possible on meso-scale images. This may overcome biofilm heterogeneity and statistical issues29.

Fungi and bacteria have many opportunities to interact with each other because they cohabit in most terrestrial environments. Due to their diversity and their ubiquity, these interactions are important in many biological fields, including biotechnology, agriculture, food processing, and medicine1,2. Molecular interactions require a certain degree of proximity to allow exchanges between the partners, and in some cases, a physical association of the partners is necessary for a functional interaction3. A common physical association between bacteria and fungi is the formation of bacterial biofilms on fungal surfaces4. This direct contact between bacterial cells and fungal hyphae permits intimate interactions that are involved in various biological processes. For example, in medicine, the study of biofilm formation of Pseudomonas aeruginosa on the opportunistic fungal pathogen Candida albicans could provide insights into the link between biofilm formation and virulence5. In agriculture, studies suggest that plant-growth-promoting rhizobacteria and biocontrol bacteria have an increased efficiency when associated with a fungus in a mixed biofilm. For example, Bradyrhizobium elkanii have enhanced N2-fixing activity when associated with Pleurotus ostreatus in a mixed biofilm6. Finally, in bioremediation, bacterial-fungal mixed biofilms have been used for the remediation of polluted sites7,8.
LSCM is particularly suitable to study biofilms since it allows for a three-dimensional observation of living hydrated biofilms with minimum pretreatments, thereby maintaining biofilm structure and organization. Thus, biofilm analysis by LSCM is very informative, especially to determine the time course of the biofilm formation and the detection of characteristic stages9,10, from the adhesion step to the development of a mature biofilm. It is also particularly adapted to visualize the biofilm structure and matrix11,12 or to quantify the biofilm size13,14. Although this method is suitable to study biofilms on abiotic or thin biotic surfaces, studying bacterial biofilm on a fungal filamentous colony is still very challenging. Indeed, most filamentous fungi build thick, complex, tridimensional networks in culture. Even if thick objects can be imaged by confocal microscopy, the attenuation of the laser penetration and the fluorescence emission often decrease the quality of the final images over a depth of 50 &#181;m15. Moreover, because fungal colonies are not rigid, it is difficult to handle the microorganisms without disturbing the biofilms. Due to the thickness of the samples, the few microscopic analyses of bacterial biofilms on fungal hyphae are usually only performed on a small part of the fungal colony, therefore containing only few hyphae16,17,18. All this limits our ability to describe biofilm distribution on the fungal colony and thus can bring biases into the analysis in case of the heterogenic distribution of the biofilm within the fungal colony.
To overcome such difficulties, we report a method for the growth and the analysis of bacterial biofilm on fungal hyphae. This method was applied to study the biofilm formation in Pseudomonas fluorescens BBc6 on the hyphae of the ectomycorrhizal basidiomycete Laccaria bicolor S238N. These two forest-soil microorganisms were previously described to form mixed biofilm-like structures19,20. This method can easily be further adapted to other filamentous fungal/bacterial systems. The method presented here is based on the combination of a fungal culture method, allowing for the growth of very thin fungal colonies, with LSCM and SEM imaging. This permitted us to obtain micro- (&#181;m range) and meso- (mm range) scale views of the interaction between the two microorganisms, allowing for the qualitative characterization of the biofilm. We also showed that the samples can be observed with SEM, permitting structural analysis of the biofilm at the nano-scale level (nm range).

<table><tbody><tr><td>6 well Falcon Tissue Culture Plates</td><td>Fisher Scientific</td><td>08-772-33</td><td>Used in 2.2 &amp;amp; 3.1</td></tr><tr><td>Congo Red</td><td>Fisher Scientific</td><td>C580-25</td><td>Used in 3.1.4.1</td></tr><tr><td>FUN 1 Cell Stain</td><td>Thermo Fisher Scientific</td><td>F7030</td><td>Used in 3.1.4.1</td></tr><tr><td>Wheat Germ Agglutinin, Alexa Fluor 633 Conjugate</td><td>Thermo Fisher Scientific</td><td>W21404</td><td>Used in 3.1.4.1</td></tr><tr><td>DAPI solution</td><td>Thermo Fisher Scientific</td><td>62248</td><td>Used in 3.1.4.2</td></tr><tr><td>Propidium iodide</td><td>Thermo Fisher Scientific</td><td>P3566</td><td>Used in 3.1.4.3</td></tr><tr><td>FilmTracer SYPRO Ruby Biofilm Matrix protein Stain</td><td>Thermo Fisher Scientific</td><td>F10318</td><td>Used in 3.1.4.4</td></tr><tr><td>Fluoromount-G Slide Mounting Medium</td><td>Fisher Scientific</td><td>OB100-01</td><td>Used in 3.1.7</td></tr><tr><td>LSM780 Axio Observer Z1</td><td>Zeiss</td><td></td><td>Used in 3.2.1</td></tr><tr><td>ZEN 2.1 lite black software&amp;nbsp;</td><td>Zeiss</td><td></td><td>Used in 3.2.1</td></tr><tr><td>High Vacuum Coater Leica EM ACE600</td><td>Leica</td><td></td><td>Used in 4</td></tr><tr><td>GeminiSEM-FEG</td><td>Zeiss</td><td></td><td>Used in 4</td></tr></tbody></table>

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.

The overall schematic procedure of fungal culture and biofilm preparation are given in Figure 1. The culture method allowed us to obtain fungal colonies 20&#160; to 50 &#181;m thick containing a few layers of hyphae, allowing micro- and meso-scale analyses of hydrated living biofilm using LSCM. The application of the method permitted the acquisition of high-quality images of P. fluorescens BBc6 biofilms on L. bicolor hyphae along the formation of the biofilm (Figures 2-4).
The meso-scale analysis of the biofilms demonstrated the heterogenic distribution of the biofilm of P. fluorescens BBc6 on the hyphae of L. bicolor S238N (Figures 2 and 3). Meso-scale analysis also allowed for the tracking of the biofilm formation over time (Figure 3) from the early steps, in which only some bacteria were attached to hyphae (Figure 3a), to the formation of a thick, mature biofilm (Figure 3b). The high resolution of the meso-scale images allowed us to perform micro-scale analysis on the same images in order to obtain the colony architectures (Figure 3c-d).
The micro-scale analysis combined with the specific labeling enabled us to go a step further in biofilm characterization. Here, SYPRO Ruby23, which labels most classes of proteins, was applied to the samples (Figure 4) and shows the presence of proteins in the matrix.
Finally, further details of the biofilm structure were obtained by SEM imaging of the same sample after dehydration and coating (Figure 5). SEM imaging provided access to the nanoscale level, thus giving access to the matrix structure.
<img alt="Figure 1" src="/files/ftp_upload/54771/54771fig1.jpg" /> 
Figure 1: Overall schematic procedure of fungal culture and biofilm preparation. This figure describes the main steps of the method, from microorganism cultures to sample analyses. More details are given in the protocol. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54771/54771fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54771/54771fig2.jpg" /> 
Figure 2: BBc6 biofilm repartition on the fungal colony. The sample was imaged after 18 hr of interaction at 10X magnification. The image was obtained via 2D maximum intensity projection of 3D confocal microscopy images. The grey grid on the figure depicts the mosaic positions. L. bicolor hyphae are stained with Congo Red (red) and bacteria are GFP-tagged (green). <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54771/54771fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54771/54771fig3.jpg" /> 
Figure 3: Early and late stages of BBc6 biofilm formation on L. bicolor hyphae. This image was obtained via 2D maximum intensity projection of 3D confocal microscopy images. Imaging was performed at 40X magnification. (a) Biofilms repartition after 2 hr of interaction. (b) Biofilm repartition after 14 hr of interaction. (c) Enlargement of the white rectangle in (a). (d) Enlargement of the white rectangle in (b). Fungal hyphae are stained with Congo Red (red) and bacteria are GFP-tagged (green). <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54771/54771fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54771/54771fig4.jpg" /> 
Figure 4: Matrix staining of BBc6 biofilm on L. bicolor hyphae. (a) Biofilms after 16 hr of interaction. (b) Enlargement of the white rectangle in (a). Imaging was performed at 40X magnification, and these images were obtained via 2D maximum intensity projection of 3D confocal microscopy images. Bacteria are GFP-tagged (green), fungus is stained with Fun1 (dark green), and proteins are stained with S. Ruby (red). <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54771/54771fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54771/54771fig5.jpg" /> 
Figure 5: SEM imaging of BBc6 biofilms on L. bicolor hyphae. SEM Imaging was performed at 2360X, EHT: 1kV. Before imaging, the sample was slowly freeze-dried in the SEM chamber and coated with platinum. Green arrows point to bacterial biofilms and yellow arrows point to fungal hyphae. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54771/54771fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A New Method for Qualitative Multi-scale Analysis of Bacterial Biofilms on Filamentous Fungal Colonies Using Confocal and Electron Microscopy at JoVE.com

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

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 ...

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Immunology and Infection

19.4K Views.  INRA Université de Lorraine. This protocol describes a new method to grow and qualitatively analyze bacterial biofilms on fungal hyphae by confocal and electron microscopy.

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

Oral Biofilm Analysis of Palatal Expanders by Fluorescence In-Situ Hybridization and Confocal Laser Scanning Microscopy

Confocal laser scanning microscopy (CLSM) of natural heterogeneous biofilm is today facilitated by a comprehensive range of staining techniques, one of them being fluorescence in situ hybridization (FISH).1,2 We performed a pilot study in which oral biofilm samples collected from fixed orthodontic appliances (palatal expanders) were stained by FISH, the objective being to assess the three-dimensional organization of natural biofilm and plaque accumulation.3,4 FISH creates an opportunity to stain cells in their native biofilm environment by the use of fluorescently labeled 16S rRNA-targeting probes.4-7,19 Compared to alternative techniques like immunofluorescent labeling, this is an inexpensive, precise and straightforward labeling technique to investigate different bacterial groups in mixed biofilm consortia.18,20 General probes were used that bind to Eubacteria (EUB338 + EUB338II + EUB338III; hereafter EUBmix),8-10 Firmicutes (LGC354 A-C; hereafter LGCmix),9,10 and Bacteroidetes (Bac303).11 In addition, specific probes binding to Streptococcus mutans (MUT590)12,13 and Porphyromonas gingivalis (POGI)13,14 were used. The extreme hardness of the surface materials involved (stainless steel and acrylic resin) compelled us to find new ways of preparing the biofilm. As these surface materials could not be readily cut with a cryotome, various sampling methods were explored to obtain intact oral biofilm. The most workable of these approaches is presented in this communication. Small flakes of the biofilm-carrying acrylic resin were scraped off with a sterile scalpel, taking care not to damage the biofilm structure. Forceps were used to collect biofilm from the steel surfaces. Once collected, the samples were fixed and placed directly on polysine coated glass slides. FISH was performed directly on these slides with the probes mentioned above. Various FISH protocols were combined and modified to create a new protocol that was easy to handle.5,10,14,15 Subsequently the samples were analyzed by confocal laser scanning microscopy. Well-known configurations3,4,16,17 could be visualized, including mushroom-style formations and clusters of coccoid bacteria pervaded by channels. In addition, the bacterial composition of these typical biofilm structures were analyzed and 2D and 3D images created.

We present a protocol for structural and compositional analysis of natural oral biofilm from orthodontic appliances with in situ hybridization (FISH) and confocal laser scanning microscopy (CLSM). Oral biofilm samples were collected from palatal expanders, scraping acrylic-resin flakes off their surface and referring them for molecular processing.

Confocal laser scanning microscopy (CLSM) of natural heterogeneous biofilm is today facilitated by a comprehensive range of staining techniques, one of ...

Medicine

Characterization of Aquatic Biofilms with Flow Cytometry

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond quickly to environmental changes and can be thus used as indicators of water quality. Currently, biofilm assessment is mostly based on integrative and functional endpoints, such as photosynthetic or respiratory activity, which do not provide information on the biofilm community structure. Flow cytometry and computational visualization offer an alternative, sensitive, and easy-to-use method for assessment of the community composition, particularly of the photoautotrophic part of freshwater biofilms. It requires only basic sample preparation, after which the entire sample is run through the flow cytometer. The single-cell optical and fluorescent information is used for computational visualization and biological interpretation. Its main advantages over other methods are the speed of analysis and the high-information-content nature. Flow cytometry provides information on several cellular and biofilm traits in a single measurement: particle size, density, pigment content, abiotic content in the biofilm, and coarse taxonomic information. However, it does not provide information on biofilm composition on the species level. We see high potential in the use of the method for environmental monitoring of aquatic ecosystems and as an initial biofilm evaluation step that informs downstream detailed investigations by complementary and more detailed methods.

Flow cytometry in combination with visual clustering offers an easy-to-use and fast method for studying aquatic biofilms. It can be used for biofilm characterization, detection of changes in biofilm community structure, and detection of abiotic particles embedded in the biofilm.

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond ...

Environment

Concurrent Quantification of Cellular and Extracellular Components of Biofilms

Confocal laser scanning microscopy (CLSM) is a powerful tool for investigation of biofilms. Very few investigations have successfully quantified concurrent distribution of more than two components within biofilms because: 1)&#160;selection of fluorescent dyes having minimal spectral overlap is complicated, and 2)&#160;quantification of multiple fluorochromes poses a multifactorial problem. Objectives: Report a methodology to quantify and compare concurrent 3-dimensional distributions of three cellular/extracellular components of biofilms grown on relevant substrates. Methods: The method consists of distinct, interconnected steps involving biofilm growth, staining, CLSM imaging, biofilm structural analysis and visualization, and statistical analysis of structural parameters. Biofilms of Streptococcus mutans (strain UA159) were grown for 48 hr on sterile specimens of Point 4 and TPH3 resin composites. Specimens were subsequently immersed for 60 sec in either Biot&#232;ne PBF (BIO) or Listerine Total Care (LTO) mouthwashes, or water (control group; n=5/group). Biofilms were stained with fluorochromes for extracellular polymeric substances, proteins and nucleic acids before imaging with CLSM. Biofilm structural parameters calculated using ISA3D image analysis software were biovolume and mean biofilm thickness. Mixed models statistical analyses compared structural parameters between mouthwash and control groups (SAS software; &#945;=0.05). Volocity software permitted visualization of 3D distributions of overlaid biofilm components (fluorochromes). Results: Mouthwash BIO produced biofilm structures that differed significantly from the control (p&#60;0.05) on both resin composites, whereas LTO did not produce differences (p&#62;0.05) on either product. Conclusions: This methodology efficiently and successfully quantified and compared concurrent 3D distributions of three major components within S. mutans biofilms on relevant substrates, thus overcoming two challenges to simultaneous assessment of biofilm components. This method can also be used to determine the efficacy of antibacterial/antifouling agents against multiple biofilm components, as shown using mouthwashes. Furthermore, this method has broad application because it facilitates comparison of 3D structures/architecture of biofilms in a variety of disciplines.

A protocol for the concurrent quantification and comparison of three cellular and extracellular components within biofilms is presented. The methodology involves the use of confocal laser scanning microscopy, biofilm structural analysis and visualization software, and statistical analysis software.

Confocal laser scanning microscopy (CLSM) is a powerful tool for investigation of biofilms. Very few investigations have successfully quantified ...

Methods for Characterizing the Co-development of Biofilm and Habitat Heterogeneity

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is strongly regulated by the surrounding flow and nutritional environment. Biofilm growth also increases the heterogeneity of the local microenvironment by generating complex flow fields and solute transport patterns. To investigate the development of heterogeneity in biofilms and interactions between biofilms and their local micro-habitat, we grew mono-species biofilms of Pseudomonas aeruginosa and dual-species biofilms of P. aeruginosa and Escherichia coli under nutritional gradients in a microfluidic flow cell. We provide detailed protocols for creating nutrient gradients within the flow cell and for growing and visualizing biofilm development under these conditions. We also present protocols for a series of optical methods to quantify spatial patterns in biofilm structure, flow distributions over biofilms, and mass transport around and within biofilm colonies. These methods support comprehensive investigations of the co-development of biofilm and habitat heterogeneity.

Biofilms have complex interactions with their surrounding environment. To comprehensively investigate biofilm-environment interactions, we present here a series of methods to create heterogeneous chemical environment for biofilm development, to quantify local flow velocity, and to analyze mass transport in and around biofilm colonies.

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is ...

Bioengineering

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 ...

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.

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 ...

A Semi-quantitative Approach to Assess Biofilm Formation Using Wrinkled Colony Development

Biofilms, or surface-attached communities of cells encapsulated in an extracellular matrix, represent a common lifestyle for many bacteria. Within a biofilm, bacterial cells often exhibit altered physiology, including enhanced resistance to antibiotics and other environmental stresses 1. Additionally, biofilms can play important roles in host-microbe interactions. Biofilms develop when bacteria transition from individual, planktonic cells to form complex, multi-cellular communities 2. In the laboratory, biofilms are studied by assessing the development of specific biofilm phenotypes. A common biofilm phenotype involves the formation of wrinkled or rugose bacterial colonies on solid agar media 3. Wrinkled colony formation provides a particularly simple and useful means to identify and characterize bacterial strains exhibiting altered biofilm phenotypes, and to investigate environmental conditions that impact biofilm formation. Wrinkled colony formation serves as an indicator of biofilm formation in a variety of bacteria, including both Gram-positive bacteria, such as Bacillus subtilis 4, and Gram-negative bacteria, such as Vibrio cholerae 5, Vibrio parahaemolyticus 6, Pseudomonas aeruginosa 7, and Vibrio fischeri 8.
The marine bacterium V. fischeri has become a model for biofilm formation due to the critical role of biofilms during host colonization: biofilms produced by V. fischeri promote its colonization of the Hawaiian bobtail squid Euprymna scolopes 8-10. Importantly, biofilm phenotypes observed in vitro correlate with the ability of V. fischeri cells to effectively colonize host animals: strains impaired for biofilm formation in vitro possess a colonization defect 9,11, while strains exhibiting increased biofilm phenotypes are enhanced for colonization 8,12. V. fischeri therefore provides a simple model system to assess the mechanisms by which bacteria regulate biofilm formation and how biofilms impact host colonization.
In this report, we describe a semi-quantitative method to assess biofilm formation using V. fischeri as a model system. This method involves the careful spotting of bacterial cultures at defined concentrations and volumes onto solid agar media; a spotted culture is synonymous to a single bacterial colony. This 'spotted culture' technique can be utilized to compare gross biofilm phenotypes at single, specified time-points (end-point assays), or to identify and characterize subtle biofilm phenotypes through time-course assays of biofilm development and measurements of the colony diameter, which is influenced by biofilm formation. Thus, this technique provides a semi-quantitative analysis of biofilm formation, permitting evaluation of the timing and patterning of wrinkled colony development and the relative size of the developing structure, characteristics that extend beyond the simple overall morphology.

We provide a simple, semi-quantitative method to investigate biofilm formation in vitro. This method takes advantage of the Zeiss stemi 2000-C Dissecting Microscope (with camera attachment) to monitor both the timing and pattern of biofilm formation, as assessed by the development of wrinkled colonies.

Biofilms, or surface-attached communities of cells encapsulated in an extracellular matrix, represent a common lifestyle for many bacteria. Within a ...

Biology

Monitoring Extracellular pH in Cross-Kingdom Biofilms using Confocal Microscopy

Cross-kingdom biofilms consisting of both fungal and bacterial cells are involved in a variety of oral diseases, such as endodontic infections, periodontitis, mucosal infections and, most notably, early childhood caries. In all of these conditions, the pH in the biofilm matrix impacts microbe-host interactions and thus the disease progression. The present protocol describes a confocal microscopy-based method to monitor pH dynamics inside cross-kingdom biofilms comprising Candida albicans and Streptococcus mutans. The pH-dependent dual-emission spectrum and the staining properties of the ratiometric probe C-SNARF-4 are exploited to determine drops in pH in extracellular areas of the biofilms. Use of pH ratiometry with the probe requires a meticulous choice of imaging parameters, a thorough calibration of the dye, and careful, threshold-based post-processing of the image data. When used correctly, the technique allows for the rapid assessment of extracellular pH in different areas of a biofilm and thus the monitoring of both horizontal and vertical pH gradients over time. While the use of confocal microscopy limits Z-profiling to thin biofilms of 75 &#181;m or less, the use of pH ratiometry is ideally suited for the noninvasive study of an important virulence factor in cross-kingdom biofilms.

The protocol describes the cultivation of cross-kingdom biofilms consisting of Candida albicans and Streptococcus mutans and presents a confocal microscopy-based method for the monitoring of extracellular pH inside these biofilms.

Cross-kingdom biofilms consisting of both fungal and bacterial cells are involved in a variety of oral diseases, such as endodontic infections, ...

Identifying Drugs that Affect Polymerization of Bacterial Cytoskeletal Proteins

Fission Yeast as a Platform for Antibacterial Drug Screens Targeting Bacterial Cytoskeleton Proteins

Bacterial cytoskeletal proteins such as FtsZ and MreB perform essential functions such as cell division and cell shape maintenance. Further, FtsZ and MreB have emerged as important targets for novel antimicrobial discovery. Several assays have been developed to identify compounds targeting nucleotide binding and polymerization of these cytoskeletal proteins, primarily focused on FtsZ. Moreover, many of the assays are either laborious or cost-intensive, and ascertaining whether these proteins are the cellular target of the drug often requires multiple methods. Finally, the toxicity of the drugs to eukaryotic cells also poses a problem. Here, we describe a single-step cell-based assay to discover novel molecules targeting bacterial cytoskeleton and minimize hits that might be potentially toxic to eukaryotic cells. Fission yeast is amenable to high-throughput screens based on microscopy, and a visual screen can easily identify any molecule that alters the polymerization of FtsZ or MreB. Our assay utilizes the standard 96-well plate and relies on the ability of the bacterial cytoskeletal proteins to polymerize in a eukaryotic cell such as the fission yeast. While the protocols described here are for fission yeast and utilize FtsZ from Staphylococcus aureus and MreB from Escherichia coli, they are easily adaptable to other bacterial cytoskeletal proteins that readily assemble into polymers in any eukaryotic expression hosts. The method described here should help facilitate further discovery of novel antimicrobials targeting bacterial cytoskeletal proteins.

Author Spotlight: Exploring Cytoskeletal Dynamics to Unveil Novel Antibiotics Through Innovative Cell-Based Assays

Fission yeast is used here as a heterologous host to express bacterial cytoskeletal proteins such as FtsZ and MreB as translational fusion proteins with GFP to visualize their polymerization. Also, compounds that affect polymerization are identified by imaging using a fluorescence microscope.

Bacterial cytoskeletal proteins such as FtsZ and MreB perform essential functions such as cell division and cell shape maintenance. Further, FtsZ and MreB ...

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.bdigitaluss.remotexs.co/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 ...

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

Summary

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Chapters in this video

Oral Biofilm Analysis of Palatal Expanders by Fluorescence In-Situ Hybridization and Confocal Laser Scanning Microscopy

Characterization of Aquatic Biofilms with Flow Cytometry

Concurrent Quantification of Cellular and Extracellular Components of Biofilms

Methods for Characterizing the Co-development of Biofilm and Habitat Heterogeneity

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

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

A Semi-quantitative Approach to Assess Biofilm Formation Using Wrinkled Colony Development

Monitoring Extracellular pH in Cross-Kingdom Biofilms using Confocal Microscopy

Fission Yeast as a Platform for Antibacterial Drug Screens Targeting Bacterial Cytoskeleton Proteins

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