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 border="0" cellpadding="0" cellspacing="0" width="1069">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">6 well Falcon Tissue Culture Plates</td>
			<td style="width:381px;">Fisher Scientific</td>
			<td style="width:136px;">08-772-33</td>
			<td style="width:141px;">Used in 2.2 &#38; 3.1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">Congo Red</td>
			<td style="width:381px;">Fisher Scientific</td>
			<td style="width:136px;">C580-25</td>
			<td style="width:141px;">Used in 3.1.4.1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">FUN 1 Cell Stain</td>
			<td style="width:381px;">Thermo Fisher Scientific</td>
			<td style="width:136px;">F7030</td>
			<td style="width:141px;">Used in 3.1.4.1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">Wheat Germ Agglutinin, Alexa Fluor 633 Conjugate</td>
			<td style="width:381px;">Thermo Fisher Scientific</td>
			<td style="width:136px;">W21404</td>
			<td style="width:141px;">Used in 3.1.4.1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">DAPI solution</td>
			<td style="width:381px;">Thermo Fisher Scientific</td>
			<td style="width:136px;">62248</td>
			<td style="width:141px;">Used in 3.1.4.2</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">Propidium iodide</td>
			<td style="width:381px;">Thermo Fisher Scientific</td>
			<td style="width:136px;">P3566</td>
			<td style="width:141px;">Used in 3.1.4.3</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">FilmTracer SYPRO Ruby Biofilm Matrix protein Stain</td>
			<td style="width:381px;">Thermo Fisher Scientific</td>
			<td style="width:136px;">F10318</td>
			<td style="width:141px;">Used in 3.1.4.4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">Fluoromount-G Slide Mounting Medium</td>
			<td style="width:381px;">Fisher Scientific</td>
			<td style="width:136px;">OB100-01</td>
			<td style="width:141px;">Used in 3.1.7</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">LSM780 Axio Observer Z1</td>
			<td style="width:381px;">Zeiss</td>
			<td style="width:136px;"></td>
			<td style="width:141px;">Used in 3.2.1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ZEN 2.1 lite black software&#160;</td>
			<td style="width:381px;">Zeiss</td>
			<td style="width:136px;"></td>
			<td style="width:141px;">Used in 3.2.1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">High Vacuum Coater Leica EM ACE600</td>
			<td style="width:381px;">Leica</td>
			<td style="width:136px;"></td>
			<td style="width:141px;">Used in 4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:411px;">GeminiSEM-FEG</td>
			<td style="width:381px;">Zeiss</td>
			<td style="width:136px;"></td>
			<td style="width:141px;">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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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

Microwave Assisted Rapid Diagnosis of Plant Virus Diseases by Transmission Electron Microscopy

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample preparation for transmission electron microscopy (TEM) such investigations can take several days1,2 and are therefore not suited for a rapid diagnosis of plant virus diseases. Microwave fixation can be used to drastically reduce sample preparation time for TEM investigations with similar ultrastructural results as observed after conventionally sample preparation3-5. Many different custom made microwave devices are currently available which can be used for the successful fixation and embedding of biological samples for TEM investigations5-8. In this study we demonstrate on Tobacco Mosaic Virus (TMV) infected Nicotiana tabacum plants that it is possible to diagnose ultrastructural alterations in leaves in about half a day by using microwave assisted sample preparation for TEM. We have chosen to perform this study with a commercially available microwave device as it performs sample preparation almost fully automatically5 in contrast to the other available devices where many steps still have to be performed manually6-8 and are therefore more time and labor consuming. As sample preparation is performed fully automatically negative staining of viral particles in the sap of the remaining TMV-infected leaves and the following examination of ultrastructure and size can be performed during fixation and embedding.

This study describes a method that allows the rapid and clear diagnosis of plant virus diseases in about half a day by using a combination of microwave assisted plant sample preparation for transmission electron microscopy and negative staining methods.

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample ...

Intravital Microscopy of the Spleen: Quantitative Analysis of Parasite Mobility and Blood Flow

The advent of intravital microscopy in experimental rodent malaria models has allowed major advances to the knowledge of parasite-host interactions 1,2. Thus, in vivo imaging of malaria parasites during pre-erythrocytic stages have revealed the active entrance of parasites into skin lymph nodes 3, the complete development of the parasite in the skin 4, and the formation of a hepatocyte-derived merosome to assure migration and release of merozoites into the blood stream 5. Moreover, the development of individual parasites in erythrocytes has been recently documented using 4D imaging and challenged our current view on protein export in malaria 6. Thus, intravital imaging has radically changed our view on key events in Plasmodium development. Unfortunately, studies of the dynamic passage of malaria parasites through the spleen, a major lymphoid organ exquisitely adapted to clear infected red blood cells are lacking due to technical constraints.
Using the murine model of malaria Plasmodium yoelii in Balb/c mice, we have implemented intravital imaging of the spleen and reported a differential remodeling of it and adherence of parasitized red blood cells (pRBCs) to barrier cells of fibroblastic origin in the red pulp during infection with the non-lethal parasite line P.yoelii 17X as opposed to infections with the P.yoelii 17XL lethal parasite line 7. To reach these conclusions, a specific methodology using ImageJ free software was developed to enable characterization of the fast three-dimensional movement of single-pRBCs. Results obtained with this protocol allow determining velocity, directionality and residence time of parasites in the spleen, all parameters addressing adherence in vivo. In addition, we report the methodology for blood flow quantification using intravital microscopy and the use of different colouring agents to gain insight into the complex microcirculatory structure of the spleen. 
Ethics statement
All the animal studies were performed at the animal facilities of University of Barcelona in accordance with guidelines and protocols approved by the Ethics Committee for Animal Experimentation of the University of Barcelona CEEA-UB (Protocol No DMAH: 5429). Female Balb/c mice of 6-8 weeks of age were obtained from Charles River Laboratories.

We show the method for performing intravital microscopy of the spleen using GFP transgenic malaria parasites and the quantification of parasite mobility and blood flow within this organ.

The advent of intravital microscopy in experimental rodent malaria models has allowed major advances to the knowledge of parasite-host interactions 1,2. ...

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly constituted by proteins and exopolysaccharides, among other factors 7-10. The microbial community encased within the biofilm often shows the differentiation of distinct subpopulation of specialized cells 11-17. These subpopulations coexist and often show spatial and temporal organization within the biofilm 18-21.
Biofilm formation in the model organism Bacillus subtilis requires the differentiation of distinct subpopulations of specialized cells. Among them, the subpopulation of matrix producers, responsible to produce and secrete the extracellular matrix of the biofilm is essential for biofilm formation 11,19. Hence, differentiation of matrix producers is a hallmark of biofilm formation in B. subtilis.
We have used fluorescent reporters to visualize and quantify the subpopulation of matrix producers in biofilms of B. subtilis 15,19,22-24. Concretely, we have observed that the subpopulation of matrix producers differentiates in response to the presence of self-produced extracellular signal surfactin 25. Interestingly, surfactin is produced by a subpopulation of specialized cells different from the subpopulation of matrix producers 15.
We have detailed in this report the technical approach necessary to visualize and quantify the subpopulation of matrix producers and surfactin producers within the biofilms of B. subtilis. To do this, fluorescent reporters of genes required for matrix production and surfactin production are inserted into the chromosome of B. subtilis. Reporters are expressed only in a subpopulation of specialized cells. Then, the subpopulations can be monitored using fluorescence microscopy and flow cytometry (See Fig 1).
The fact that different subpopulations of specialized cells coexist within multicellular communities of bacteria gives us a different perspective about the regulation of gene expression in prokaryotes. This protocol addresses this phenomenon experimentally and it can be easily adapted to any other working model, to elucidate the molecular mechanisms underlying phenotypic heterogeneity within a microbial community.

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Microbial biofilms are generally constituted by distinct subpopulations of specialized cells. Single-cell analysis of these subpopulations requires the use of fluorescent reporters. Here we describe a protocol to visualize and monitor several subpopulationswithin B. subtilis biofilms using fluorescence microscopy and flow cytometry.

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly ...

Visualization of Bacterial Toxin Induced Responses Using Live Cell Fluorescence Microscopy

Bacterial toxins bind to cholesterol in membranes, forming pores that allow for leakage of cellular contents and influx of materials from the external environment. The cell can either recover from this insult, which requires active membrane repair processes, or else die depending on the amount of toxin exposure and cell type1. In addition, these toxins induce strong inflammatory responses in infected hosts through activation of immune cells, including macrophages, which produce an array of pro-inflammatory cytokines2. Many Gram positive bacteria produce cholesterol binding toxins which have been shown to contribute to their virulence through largely uncharacterized mechanisms.
Morphologic changes in the plasma membrane of cells exposed to these toxins include their sequestration into cholesterol-enriched surface protrusions, which can be shed into the extracellular space, suggesting an intrinsic cellular defense mechanism3,4. This process occurs on all cells in the absence of metabolic activity, and can be visualized using EM after chemical fixation4. In immune cells such as macrophages that mediate inflammation in response to toxin exposure, induced membrane vesicles are suggested to contain cytokines of the IL-1 family and may be responsible both for shedding toxin and disseminating these pro-inflammatory cytokines5,6,7. A link between IL-1&beta; release and a specific type of cell death, termed pyroptosis has been suggested, as both are caspase-1 dependent processes8. To sort out the complexities of this macrophage response, which includes toxin binding, shedding of membrane vesicles, cytokine release, and potentially cell death, we have developed labeling techniques and fluorescence microscopy methods that allow for real time visualization of toxin-cell interactions, including measurements of dysfunction and death (Figure 1). Use of live cell imaging is necessary due to limitations in other techniques. Biochemical approaches cannot resolve effects occurring in individual cells, while flow cytometry does not offer high resolution, real-time visualization of individual cells. The methods described here can be applied to kinetic analysis of responses induced by other stimuli involving complex phenotypic changes in cells.

Methods for purifying the cholesterol binding toxin streptolysin O from recombinant E. coli and visualization of toxin binding to live eukaryotic cells are described. Localized delivery of toxin induces rapid and complex changes in targeted cells revealing novel aspects of toxin biology.

Bacterial toxins bind to cholesterol in membranes, forming pores that allow for leakage of cellular contents and influx of materials from the external ...

Live-cell Video Microscopy of Fungal Pathogen Phagocytosis

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of phagocytes to the site where pathogens are located; (ii) recognition of pathogen-associated molecular patterns (PAMPs) through pattern recognition receptors (PRRs); (iii) engulfment of microorganisms bound to the phagocyte cell membrane, and (iv) processing of engulfed cells within maturing phagosomes and digestion of the ingested particle. Studies that assess phagocytosis in its entirety are informative1, 2, 3, 4, 5 but are limited in that they do not normally break the process down into migration, engulfment and phagosome maturation, which may be affected differentially. Furthermore, such studies assess uptake as a single event, rather than as a continuous dynamic process. We have recently developed advanced live-cell imaging technologies, and have combined these with genetic functional analysis of both pathogen and host cells to create a cross-disciplinary platform for the analysis of innate immune cell function and fungal pathogenesis. These studies have revealed novel aspects of phagocytosis that could only be observed using systematic temporal analysis of the molecular and cellular interactions between human phagocytes and fungal pathogens and infectious microorganisms more generally. For example, we have begun to define the following: (a) the components of the cell surface required for each stage of the process of recognition, engulfment and killing of fungal cells1, 6, 7, 8; (b) how surface geometry influences the efficiency of macrophage uptake and killing of yeast and hyphal cells7; and (c) how engulfment leads to alteration of the cell cycle and behavior of macrophages 9, 10.
In contrast to single time point snapshots, live-cell video microscopy enables a wide variety of host cells and pathogens to be studied as continuous sequences over lengthy time periods, providing spatial and temporal information on a broad range of dynamic processes, including cell migration, replication and vesicular trafficking. Here we describe in detail how to prepare host and fungal cells, and to conduct the video microscopy experiments. These methods can provide a user-guide for future studies with other phagocytes and microorganisms.

We describe methods for live-cell video microscopy of Candida albicans phagocytosis by macrophages. These methods enable stage-specific analysis of macrophage migration, recognition, engulfment and phagosome maturation and reveal novel aspects of phagocytosis.

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of ...

Assessing Anti-fungal Activity of Isolated Alveolar Macrophages by Confocal Microscopy

The lung is an interface where host cells are routinely exposed to microbes and microbial products. Alveolar macrophages are the first-line phagocytic cells that encounter inhaled fungi and other microbes. Macrophages and other immune cells recognize Aspergillus motifs by pathogen recognition receptors and initiate downstream inflammatory responses. The phagocyte NADPH oxidase generates reactive oxygen intermediates (ROIs) and is critical for host defense. Although NADPH oxidase is critical for neutrophil-mediated host defense1-3, the importance of NADPH oxidase in macrophages is not well defined. The goal of this study was to delineate the specific role of NADPH oxidase in macrophages in mediating host defense against A. fumigatus. We found that NADPH oxidase in alveolar macrophages controls the growth of phagocytosed A. fumigatus spores4. Here, we describe a method for assessing the ability of mouse alveolar macrophages (AMs) to control the growth of phagocytosed Aspergillus spores (conidia). Alveolar macrophages are stained in vivo and ten days later isolated from mice by bronchoalveolar lavage (BAL). Macrophages are plated onto glass coverslips, then seeded with green fluorescent protein (GFP)-expressing A. fumigatus spores. At specified times, cells are fixed and the number of intact macrophages with phagocytosed spores is assessed by confocal microscopy.

A method to evaluate the ability of isolated mouse alveolar macrophages to control the growth of phagocytosed Aspergillus spores by confocal microscopy.

The lung is an interface where host cells are routinely exposed to microbes and microbial products. Alveolar macrophages are the first-line phagocytic ...

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 ...

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research investigating immunological responses against A. fumigatus has been limited by the lack of consistent and reliable assays for measuring the antifungal activity of specific immune cells in vitro. A new method is described to assess the antifungal activity of primary monocytes and neutrophils from human donors against A. fumigatus using FLuorescent Aspergillus REporter (FLARE) conidia. These conidia contain a genetically encoded dsRed reporter, which is constitutively expressed by live FLARE conidia, and are externally labeled with Alexa Fluor 633, which is resistant to degradation within the phagolysosome, thus allowing a distinction between live and dead A. fumigatus conidia. Video microscopy and flow cytometry are subsequently used to visualize the interaction between conidia and innate immune cells, assessing fungicidal activity whilst also providing a wealth of information on phagocyte migration, phagocytosis and the inhibition of fungal growth. This novel technique has already provided exciting new insights into the host-pathogen interaction of primary immune cells against A. fumigatus. It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.
It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Here, we describe a protocol to assess antifungal activity of primary human immune cells in real-time using fluorescent Aspergillus reporter conidia in conjunction with live-cell video microscopy and flow cytometry. Generated data provide insight into host cell-Aspergillus interactions such as fungicidal activity, phagocytosis, cell migration and inhibition of fungal growth.

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research ...

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal microscopy methods to visualize macrophage extracellular traps (METs). These extracellular traps are defined by the presence of extracellular chromatin with co-expression of other components such as granule proteases, citrullinated histones, and peptidyl arginase deiminase (PAD). The expression of METs is generally measured after exposure to a stimulus and compared to un-stimulated samples. Samples are also included for background and isotype control. Cells are analyzed using well-defined image analysis software. Confocal microscopy may be used to define the presence of METs both in vitro and in vivo in lung tissue.

Macrophage extracellular traps are a newly described entity. This article will concentrate on confocal microscopy methods and how they are visualized in vitro and in vivo from lung samples.

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal ...

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.

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

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Microwave Assisted Rapid Diagnosis of Plant Virus Diseases by Transmission Electron Microscopy

Intravital Microscopy of the Spleen: Quantitative Analysis of Parasite Mobility and Blood Flow

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Visualization of Bacterial Toxin Induced Responses Using Live Cell Fluorescence Microscopy

Live-cell Video Microscopy of Fungal Pathogen Phagocytosis

Assessing Anti-fungal Activity of Isolated Alveolar Macrophages by Confocal Microscopy

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

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

Monitoring Extracellular pH in Cross-Kingdom Biofilms using Confocal Microscopy