This project was supported by Midwestern University, IL Office of Research and Sponsored Programs (ORSP) and Midwestern University College of Dental Medicine-Illinois (CDMI).

1. HSV Strains and Handling
Note: Recombinant non-spreading HSV-1(KOS) gL86 and HSV-2 (KOS) 333gJ- with beta-galactosidase reporter activity used were provided by V. Twiari36,37.
<ol>
	<li>Use virus from a single lot and store at -80 &#176;C at a 1:1 ratio of Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) with 20% fetal bovine serum (FBS) and skim milk until use. Before viral lot storage, determine virus concentration by o-nitrophenyl-&#946;-D-galactopyranoside (ONPG) and 5-bromo-4-chloro-3-indolyl-&#946;-D-galactopyranoside (X-Gal) assay.</li>
	<li>Determine virus viability and multiplicity of infection (MOI) by X-Gal staining for each experimental assay run using reporter virus entry assay, as previously described (Figure 1)14.</li>
	<li>Dilute virus (Opt-MEM) to desired MOI. Fix monolayers (paraformaldehyde; 0.5 ml/well) before staining. Place virus viability controls in a separate microtiter plate in parallel with polymicrobial assay plates.</li>
</ol>
2. HeLa 299 Cell Handling
<ol>
	<li>Grow at 37 &#176;C, 5% CO2 in DMEM with 4.5 g/L glucose, 10% heat-inactivated fetal bovine serum (FBS), gentamicin (50 &#181;g/ml) and L-glutamine. Passage cells at 80% confluence with Trypsin solution (ethylenediaminetetraacetic acid, 0.53 M EDTA; 0.05% Trypsin; 5 ml/flask).</li>
</ol>
3. C. albicans Handling
Note: C. albicans obtained from a clinical laboratory source is stored at -80 &#176;C in Remmel skim milk 2x medium.
<ol>
	<li>Culture frozen stock onto Sabouraud Dextrose Medium (37 &#176;C). After 24 hr subculture the C. albicans onto Fungisel medium (37 &#176;C; 48 hr) for use.</li>
	<li>Generate germ tube (GT) forms (pick representative colonies; 3 ml FBS; 3 hr; 37 &#176;C; Abs600, 0.3). After incubation, wash GT (HBSS; 2x; 4,000 x g). Add washed GT to warmed HBSS (37 &#176;C; 0.32 Abs600). GT forms should be 99% of cells observed as determined by hemocytometer count.</li>
	<li>Make yeast form (YF) stock suspensions by picking representative Fungisel colonies (HBSS, 3 ml; 0.32 Abs600). Count the number of YF forms/ml microscopically using a hemocytometer. YF forms should be 99% of cells observed as determined by hemocytometer count.</li>
	<li>Make working fungal stock (250 &#181;l of GT or YF stock in 25 ml HBSS; 37 &#176;C; 105 CFU/ml)</li>
</ol>
4. S. aureus Handling
<ol>
	<li>Store S. aureus ATCC 25923 (-80 &#176;C; Remmel skim milk 2x). Culture onto sheep blood agar (5%; 37 &#176;C; 24 hr). Pick representative colonies and transfer to mannitol salts medium within 2 days for stock (37 &#176;C; 18 hr).</li>
	<li>Make S. aureus stock suspension (3 ml HBSS; 1.32 Abs600 ;108 CFU/ml)
	<ol>
		<li>Make working S. aureus stock (100 &#181;l of the stock in 25 ml HBSS; 105 CFU/ml).</li>
	</ol>
	</li>
</ol>
5. Candida and S. aureus Suspensions
<ol>
	<li>Make mixed C. albicans and S. aureus suspension (250 &#181;l YF or GT stock and 100 &#181;l S. aureus stock in 25 ml HBSS).</li>
</ol>
6. Polymicrobial Biofilm Assay
<ol>
	<li>Seed 96 well plates with 200 &#181;l of 2 x 105 HeLa cells/ml (85% confluence level). Rock plates (30 - 45 min; 37 &#176;C) before incubation (37 &#176;C; 5% CO2 incubator; 18 hr). Wash monolayers (1x; Opt-MEM ) then seed with HSV (HSV-1 (KOS) gL86 or HSV-2 (KOS) 33 gJ- in 100 &#181;l Opt-MEM ; MOI 50 and 10). Incubate plates (3 hr; 37 &#176;C; 5% CO2). Use only one viral strain per day.</li>
	<li>Wash infected monolayers (1x; phosphate buffered saline (PBS) with Mg+2 and Ca+2; 100 &#181;l). Replace PBS with warm HBSS leaving 25 &#181;l in each well.
	<ol>
		<li>Add YF, GT and/or S. aureus working suspensions (100 &#181;l; target to cell ratio =5:1; n=16) as indicated in Table 1. Incubate plates (static; 30 min; 37 &#176;C; 5% CO2).</li>
	</ol>
	</li>
	<li>After incubation, aspirate one column at a time immediately refilling with 300 &#181;l PBS with Mg+2 and Ca+2. Repeat this step twice then add radio-immunoprecipitation assay lysis buffer (RIPA; filter sterilized; 200 &#181;l of a 1:50 dilution).</li>
	<li>Rapidly triturate the HeLa cell lysate then place 50 &#181;l onto mannitol salts (MS) and/or Fungisel (F) media (Figure 2). Spread the lysate using a glass rod bent at a 90&#176; angle. Incubate the plates (18 hr at 37 &#176;C). Manually count the number of colonies per plate. Controls consist of S. aureus and/or C. albicans adherence to HSV-uninfected HeLa cells.</li>
</ol>
7. Imaging Studies
<ol>
	<li>Wash each round glass coverslip (12 mm; 50 ml acetone in 100 ml beaker). Dry and sterilize coverslips (Kimwipes; glass petri dishes). Place dry sterile coverslips into the wells of sterile 24 well plates with alcohol flame sterilized forceps.</li>
	<li>Add HeLa cells (1 ml; 5x volume used for 96 well plates) to the wells of 24 well plates containing the washed sterile round glass coverslips. Add the virus, bacteria and fungi according to the template (Table 2) at 5x the volume used for the 96 well plates, then incubate and process as described in steps 6.2.1 to 6.4 above.</li>
	<li>After the final wash, fix the cells for microscopy by flooding the slide with methanol and allowing it to evaporate. Store the plates at RT until staining.</li>
	<li>For bright field microscopy (1,000x final original magnification) fill the wells containing methanol-fixed coverslips with deionized water. Immediately aspirate water. Cover each coverslip with Grams crystal violet. Wash coverslips free of non-bound stain (deionized water). Dry coverslips in situ then adhere them with hard set mounting medium to a labeled slide (Figure 3).</li>
	<li>For fluorescent microscopy (100x objective; 1,000x final original magnification) wash coverslips free of methanol essentially as described in step 7.4. Dry the coverslips in the wells.
	<ol>
		<li>After drying, remove the cover slips and place them on labeled slides. Then add a sufficient amount of 1:20 dilution of fluorescein isothiocyanate (FITC)-conjugated Herpes Simplex Virus Type 1 + 2 gD antibody to cover the coverslip (1 - 5 &#181;l).</li>
		<li>Incubate the slides in a moisture chamber at 37 &#176;C for 30 min. After incubation, wash the coverslips in 4 changes of PBS.</li>
		<li>After the final wash, return the coverslip to the labeled slide. Stain with 4&#8242;,6-diamidino-2-phenylindole (DAPI; moisture chamber; 37 &#176;C for 30 min). After incubation wash the coverslips in 4 changes of PBS, then affix to the labeled slide with hard set mounting medium.</li>
		<li>Allow the mounting medium to cure for 24 hr at RT in the dark. Examine the coverslips under oil objective on either a bright field microscope or fluorescent microscope with FITC and DAPI cutoff filters. Examine pictures of at least 50 fields (100 cells per organism minimum) for co-localization (Figure 4).</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Currently no information is available on complex interactions between permanent to semi-permanent members of the host microbiome that cross multiple taxonomic domains, i.e., prokaryotic, eukaryotic and viral. Therefore we developed a novel in vitro model system to study biofilm initiation by S. aureus and C. albicans on HSV-1 or HSV-2 infected HeLa 229 (HeLa) cells 38. The HeLa cell model system presents a unique advantage. This is due to their lack of surface fibronectin ex...

The human microbiome includes diverse organisms from multiple taxonomic kingdoms that share geographic regions in the body. Adherence to cell surfaces is an essential first step in biofilm formation, which is part of the microbiome colonization process. Included in the microbiome can be viruses that cause chronic and persistent infections. The chronic cell infection by these viruses can cause an alteration in putative receptor availability.1,2 In addition, cell entry by intracellular pathogens could also affect host membrane fluidity/hydrophobicity which in turn may alter attachment of other microbiome members, including bacteria and fungi. In order to understand the interactions that can occur between these multiple pathogens that co-localize in the same geographic regions of the human host, we must be able to study the interaction of pathogens that represent the spectrum of taxonomic kingdoms present at the mucosal surface.
The Herpesviridae are a family of microbes present in 100% of humans as permanent members of the microbiome3,4. In addition they can also be persistently shed both in the presence and absence of symptoms. Specifically, herpes simplex virus-1 and herpes simplex virus-2 (HSV-1 and HSV-2, respectively) are permanent members of the microbiome in the oronasopharynx and genital tract. In immune-competent individuals, both HSV-1 and HSV-2 cause gingivostomatitis, as well as genital herpes5-8. At these sites, HSV causes a latent infection characterized by chronic persistent asymptomatic viral shedding9. Entry of HSV into cells results in alterations in surface expression of nectins, heparan sulfate, lipid rafts and herpesvirus entry mediator/tumor necrosis factor receptor (HVEM/TNFr)10-25. These potentially represent shared receptors for some bacteria and fungi, e.g. S. aureus and C. albicans,which while opportunistic pathogens, can also reside as members of the mucosal microbiome of the oronasopharynx 26,27. Within the oronasopharynx S. aureus and C. albicans occupy two distinct sites of colonization. In hosts with natural teeth, the oral mucosa is shared by HSV-1 and C. albicans, while the anterior nasal nares are occupied by S. aureus28. However, despite in vitro findings that S. aureusadheres to mouth epithelial cells, 29,30 S. aureus is infrequently isolated from oral specimens when normal tissue is present29,30. Little is known concerning genital tract co-colonization niches beyond the clinical findings that S. aureus is associated with aerobic vaginitis, characterized by genital inflammation, discharge and dyspareunia, while C. albicans produces mucosal lesions similar to that observed in the oral cavity31-35. Thus, although these members of the oral and genital microbiome cross taxonomic kingdoms little is known concerning their interaction as it impacts their ability to initiate biofilm formation through adherence to the host cell surface5. This protocol has been effectively applied to determine the functional consequences of co-colonization/infection.

<table><tbody><tr><td>&lt;strong&gt;&lt;em&gt;C.albicans&lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>BBL Sabouraud Dextrose</td><td>BD</td><td>211584</td><td></td></tr><tr><td>Fungisel Agar</td><td>Dot Scientific</td><td>7205A</td><td></td></tr><tr><td>&lt;strong&gt;&lt;em&gt;S.aureus&lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Mannitol Salt Agar</td><td>Troy Biologicals</td><td>7143B</td><td></td></tr><tr><td>Sheep blood agar</td><td>Troy Biologicals</td><td>221239</td><td></td></tr><tr><td>&lt;strong&gt;Hela cells&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1x DMEM (Dubelcco&#039;s Modified Eagle Medium, with 4.5 g/L glucose and L-glutamine, without sodium pyruvate</td><td>Corning</td><td>10-017-CM</td><td></td></tr><tr><td>Gentamicin 50 mg/ml</td><td>Sigma</td><td>1397</td><td>50 &amp;micro;g/ml final concentration in the complete DMEM</td></tr><tr><td>Trypsin EDTA (0.05% Trypsin, 0.53 M EDTA)Solution 1x</td><td>Corning</td><td>25-052-CI</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Atlanta Biologicals</td><td>S11150</td><td>10% final concentration in the complete DMEM</td></tr><tr><td>&lt;strong&gt;Other medium and reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>ONPG</td><td>Thermo Scientific</td><td>34055</td><td></td></tr><tr><td>Ultra-Pure X gal</td><td>Invitrogen</td><td>15520-018</td><td></td></tr><tr><td>1x HBSS (Hanks&#039; Balanced Salt Solution)</td><td>Corning</td><td>20-021-CV</td><td></td></tr><tr><td>1x PBS</td><td>Dot Scientific</td><td>30042-500</td><td></td></tr><tr><td>RIPA Lysis</td><td>Life Technologies</td><td>89901</td><td></td></tr><tr><td>&lt;strong&gt;Staining&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>A433P-4</td><td></td></tr><tr><td>HSV 1&amp;amp;2, specific for gD</td><td>ViroStat</td><td>196</td><td></td></tr><tr><td>DAPI</td><td>SIGMA</td><td>D8417-5MG</td><td></td></tr><tr><td>Gram Crystal Violet</td><td>Troy Biologicals</td><td>212527</td><td></td></tr><tr><td>&lt;strong&gt;Supplies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Petri dish 100 x 15</td><td>Dot Scientific</td><td>229693&amp;nbsp;</td><td></td></tr><tr><td>Petri dish 150 x 15</td><td>Kord Valmark</td><td>2902</td><td></td></tr><tr><td>96-Well plates</td><td>Evergreen Scientific</td><td>222-8030-01F</td><td></td></tr><tr><td>24-well plates</td><td>Evergreen Scientific</td><td>222-8044-01F</td><td></td></tr><tr><td>Culture tubes 100 x 13</td><td>Thomas Scientific</td><td>9187L61</td><td></td></tr><tr><td>Cover slip circles, 12 mm</td><td>Deckglaser</td><td>CB00120RA1</td><td></td></tr></tbody></table>

determination of biofilm initiation on virus-infected cells by bacteria and fungi

The study of polymicrobial interactions across the taxonomic kingdoms that include fungi, bacteria and virus have not been previously examined with respect to how viral members of the microbiome affect subsequent microbe interactions with these virus-infected host cells. The co-habitation of virus with bacteria and fungi is principally present on the mucosal surfaces of the oral cavity and genital tract. Mucosal cells, particularly those with persistent chronic or persistent latent viral infections, could have a significant impact on members of the microbiome through virus alteration in number and type of receptors expressed. Modification in host cell membrane architecture would result in altered ability of subsequent members of the normal flora and opportunistic pathogens to initiate the first step in biofilm formation, i.e., adherence. This study describes a method for quantitation and visual examination of HSV&#39;s effect on the initiation of biofilm formation (adherence) of S. aureus and C. albicans.

The level of robustness of data obtainable from system described in this report is shown in Figure 2 a-f 38. Through the use of this system the modulation of staphylococcal and fungal interaction with virally infected cells and their effect on each other&#39;s adherence can be delineated. These types of studies require microscopic examination of the interaction as shown in Figures 3 and 4 38 in order to determine whe...

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Determination of Biofilm Initiation on Virus-infected Cells by Bacteria and Fungi

A method is described herein for the determination of inter-Kingdom association and competition (bacterial and fungal) for adherence to virus-infected HeLa cell monolayers. This protocol can be extended to multiple combinations of prokaryotes, eukaryotes, and viruses.

The study of polymicrobial interactions across the taxonomic kingdoms that include fungi, bacteria and virus have not been previously examined with ...

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Immunology and Infection

9.6K Views.  Midwestern University. A method is described herein for the determination of inter-Kingdom association and competition (bacterial and fungal) for adherence to virus-infected HeLa cell monolayers. This protocol can be extended to multiple combinations of prokaryotes, eukaryotes, and viruses.

Video: Determination of Biofilm Initiation on Virus-infected Cells by Bacteria and Fungi

Microtiter Dish Biofilm Formation Assay

Biofilms are communities of microbes attached to surfaces, which can be found in medical, industrial and natural settings. In fact, life in a biofilm probably represents the predominate mode of growth for microbes in most environments. Mature biofilms have a few distinct characteristics. Biofilm microbes are typically surrounded by an extracellular matrix that provides structure and protection to the community. Microbes growing in a biofilm also have a characteristic architecture generally comprised of macrocolonies (containing thousands of cells) surrounded by fluid-filled channels. Biofilm-grown microbes are also notorious for their resistance to a range of antimicrobial agents including clinically relevant antibiotics.
The microtiter dish assay is an important tool for the study of the early stages in biofilm formation, and has been applied primarily for the study of bacterial biofilms, although this assay has also been used to study fungal biofilm formation. Because this assay uses static, batch-growth conditions, it does not allow for the formation of the mature biofilms typically associated with flow cell systems. However, the assay has been effective at identifying many factors required for initiation of biofilm formation (i.e, flagella, pili, adhesins, enzymes involved in cyclic-di-GMP binding and metabolism) and well as genes involved in extracellular polysaccharide production. Furthermore, published work indicates that biofilms grown in microtiter dishes do develop some properties of mature biofilms, such a antibiotic tolerance and resistance to immune system effectors.
This simple microtiter dish assay allows for the formation of a biofilm on the wall and/or bottom of a microtiter dish. The high throughput nature of the assay makes it useful for genetic screens, as well as testing biofilm formation by multiple strains under various growth conditions. Variants of this assay have been used to assess early biofilm formation for a wide variety of microbes, including but not limited to, pseudomonads, Vibrio cholerae, Escherichia coli, staphylocci, enterococci, mycobacteria and fungi.
In the protocol described here, we will focus on the use of this assay to study biofilm formation by the model organism Pseudomonas aeruginosa. In this assay, the extent of biofilm formation is measured using the dye crystal violet (CV). However, a number of other colorimetric and metabolic stains have been reported for the quantification of biofilm formation using the microtiter plate assay. The ease, low cost and flexibility of the microtiter plate assay has made it a critical tool for the study of biofilms.

The assay describes a rapid means to measure early biofilm formation in bacteria and fungi. This method uses a microtiter plate as the substratum for microbial biofilm formation, and the biofilm is visualized using crystal violet strain. The assay provides either a qualitative or quantitative assay for early biofilm formation.

Biofilms are communities of microbes attached to surfaces, which can be found in medical, industrial and natural settings.  In fact, life in a biofilm ...

In vitro Biofilm Formation in an 8-well Chamber Slide

The chronic nature of many diseases is attributed to the formation of bacterial biofilms which are recalcitrant to traditional antibiotic therapy. Biofilms are community-associated bacteria attached to a surface and encased in a matrix. The role of the extracellular matrix is multifaceted, including facilitating nutrient acquisition, and offers significant protection against environmental stresses (e.g. host immune responses). In an effort to acquire a better understanding as to how the bacteria within a biofilm respond to environmental stresses we have used a protocol wherein we visualize bacterial biofilms which have formed in an 8-well chamber slide. The biofilms were stained with the BacLight Live/Dead stain and examined using a confocal microscope to characterize the relative biofilm size, and structure under varying incubation conditions. Z-stack images were collected via confocal microscopy and analyzed by COMSTAT. This protocol can be used to help elucidate the mechanism and kinetics by which biofilms form, as well as identify components that are important to biofilm structure and stability.

In vitro Biofilm Formation in an 8-well Chamber Slide

This article describes the procedure for the formation and visualization of a bacterial biofilm grown within an 8-well chamber slide

The chronic nature of many diseases is attributed to the formation of bacterial biofilms which are recalcitrant to traditional antibiotic therapy. ...

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

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

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

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

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

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

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

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

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

Visualization of Biofilm Formation in Candida albicans Using an Automated Microfluidic Device

Candida albicans is the most common fungal pathogen of humans, causing about 15% of hospital-acquired sepsis cases. A major virulence attribute of C. albicans is its ability to form biofilms, structured communities of cells attached to biotic and abiotic surfaces. C. albicans biofilms can form on host tissues, such as mucosal layers, and on medical devices, such as catheters, pacemakers, dentures, and joint prostheses. Biofilms pose significant clinical challenges because they are highly resistant to physical and chemical perturbations, and can act as reservoirs to seed disseminated infections. Various in vitro assays have been utilized to study C. albicans biofilm formation, such as microtiter plate assays, dry weight measurements, cell viability assays, and confocal scanning laser microscopy. All of these assays are single end-point assays, where biofilm formation is assessed at a specific time point. Here, we describe a protocol to study biofilm formation in real-time using an automated microfluidic device under laminar flow conditions. This method allows for the observation of biofilm formation as the biofilm develops over time, using customizable conditions that mimic those of the host, such as those encountered in vascular catheters. This protocol can be used to assess the biofilm defects of genetic mutants as well as the inhibitory effects of antimicrobial agents on biofilm development in real-time.

Visualization of Biofilm Formation in Candida albicans Using an Automated Microfluidic Device

This protocol describes the use of a customizable automated microfluidic device to visualize biofilm formation in Candida albicans under host physiological conditions.

Candida albicans is the most common fungal pathogen of humans, causing about 15% of hospital-acquired sepsis cases. A major virulence attribute of C. ...

Bile Salt-induced Biofilm Formation in Enteric Pathogens: Techniques for Identification and Quantification

Biofilm formation is a dynamic, multistage process that occurs in bacteria under harsh environmental conditions or times of stress. For enteric pathogens, a significant stress response is induced during gastrointestinal transit and upon bile exposure, a normal component of human digestion. To overcome the bactericidal effects of bile, many enteric pathogens form a biofilm hypothesized to permit survival when transiting through the small intestine. Here we present methodologies to define biofilm formation through solid-phase adherence assays as well as extracellular polymeric substance (EPS) matrix detection and visualization. Furthermore, biofilm dispersion assessment is presented to mimic the analysis of events triggering release of bacteria during the infection process. Crystal violet staining is used to detect adherent bacteria in a high-throughput 96-well plate adherence assay. EPS production assessment is determined by two assays, namely microscopy staining of the EPS matrix and semi-quantitative analysis with a fluorescently-conjugated polysaccharide binding lectin. Finally, biofilm dispersion is measured through colony counts and plating. Positive data from multiple assays support the characterization of biofilms and can be utilized to identify bile salt-induced biofilm formation in other bacterial strains.

This protocol enables the reader to analyze bile salt-induced biofilm formation in enteric pathogens using a multifaceted approach to capture the dynamic nature of bacterial biofilms by assessing adherence, extracellular polymeric substance matrix formation, and dispersion.

Biofilm formation is a dynamic, multistage process that occurs in bacteria under harsh environmental conditions or times of stress. For enteric pathogens, ...

Measuring the Migration and Biofilm Formation of Various Bacteria

As microbes that thrive in the host body primarily have adaptive abilities that facilitate their survival, methods for classifying and identifying their nature would be beneficial in facilitating their characterization. Currently, most studies focus only on one specific characterization method; however, the isolation and identification of microorganisms from the host is a continuous process and usually requires several combinatorial characterization methods. Herein, we describe methods of identifying the microbial biofilm-forming ability, the microbial respiration state, and their chemotaxis behavior. The methods are used to identify five microbes, three of which were isolated from the bone tissue of Sprague-Dawley (SD) rats (Corynebacterium stationis, Staphylococcus cohnii subsp. urealyticus, and Enterococcus faecalis) and two from the American Type Culture Collection (ATCC)-Staphylococcus aureus ATCC 25923&#160;and Enterococcus faecalis V583. The microbes isolated from the SD rat bone tissue include the gram-positive microbes. These microbes have adapted to thrive under stressful and nutrient-limiting environments within the bone matrix. This article aims to provide the readers with the specific know-how of determining the nature and behavior of the isolated microbes within a laboratory setting.

Here, we present a practical method for the isolation and identification of microorganisms within the host. In this way, the physicochemical properties of microorganisms and possible ways of living in the host are clearly described.

As microbes that thrive in the host body primarily have adaptive abilities that facilitate their survival, methods for classifying and identifying their ...

Biology

Assessment of Acinetobacter Biofilm Formation and Characterization of Biofilms

Quantification, Viability Assessment, and Visualization Strategies for Acinetobacter Biofilms

Acinetobacter causes nosocomial infections and its biofilm formation can contribute to the survival on dry surfaces such as hospital environments. Thus, biofilm quantification and visualization are important methods to assess the potential of Acinetobacter strains to cause nosocomial infections. The biofilms forming on the surface of the microplate can be quantified in terms of volume and cell numbers. Biofilm volumes can be quantified by staining using crystal violet, washing, destaining using ethanol, then measuring the solubilized dye using a microplate reader. To quantify the number of cells embedded in the biofilms, the biofilms are scrapped off using cell scrapers, harvested in the saline, vigorously agitated in the presence of glass beads, and spread on Acinetobacter agar. Then, the plates are incubated at 30 &#176;C for 24-42 h. After incubation, the red colonies are enumerated to estimate the number of cells in biofilms. This viable count method can also be useful for counting Acinetobacter cells in mixed-species biofilms. Acinetobacter biofilms can be visualized using fluorescent dyes. A commercially available microplate designed for microscopic analysis is employed to form biofilms. Then, the bottom-surface attached biofilms are stained with SYTO9 and propidium iodide dyes, washed, then visualized with confocal laser scanning microscopy.

Author Spotlight: A Comprehensive Protocol for Acinetobacter Biofilm Quantification, Assessment, and Visualization 

This protocol describes the preparation of the inoculum, the biofilm quantification on microtiter plates using crystal violet dye, the viable count in biofilms, and the visualization of biofilms of Acinetobacter.

Acinetobacter causes nosocomial infections and its biofilm formation can contribute to the survival on dry surfaces such as hospital environments. Thus, ...

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/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 ...

JoVE Encyclopedia of Experiments

Immunology

Immunodiagnostics

In Vitro and In Vivo Models

An Assay to Determine Polymicrobial Biofilm Initiation on Virus-Infected Cells

Source: Plotkin, B. J. et al., <a href="https://app.jove.com/v/54162">Determination of Biofilm Initiation on Virus-infected Cells by Bacteria and Fungi.</a> J. Vis. Exp. (2016)
This video illustrates the initiation of polymicrobial biofilm formation on virus-infected cells by bacteria and fungi. Herpes simplex virus infection impedes bacterial attachment to the cells, preventing bacterial-fungal co-localization. In contrast, uninfected cells display bacterial-fungal co-localization and initiation of biofilm formation.

1. HSV Strains and Handling
NOTE: Recombinant non-spreading HSV-1(KOS) gL86 and HSV-2 (KOS) 333gJ-&#160;with beta-galactosidase reporter activity is used ...

Determination of Biofilm Initiation on Virus-infected Cells by Bacteria and Fungi

Summary

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Chapters in this video

Microtiter Dish Biofilm Formation Assay

In vitro Biofilm Formation in an 8-well Chamber Slide

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

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

Visualization of Biofilm Formation in Candida albicans Using an Automated Microfluidic Device

Bile Salt-induced Biofilm Formation in Enteric Pathogens: Techniques for Identification and Quantification

Measuring the Migration and Biofilm Formation of Various Bacteria

Quantification, Viability Assessment, and Visualization Strategies for Acinetobacter Biofilms

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

An Assay to Determine Polymicrobial Biofilm Initiation on Virus-Infected Cells