We thank Rachael B. Chanin&#160;and Alejandro Llanos-Chea for technical assistance. We thank Anthony T. Maurelli, Bryan P. Hurley, Alessio Fasano, Brett E. Swierczewski, and Bobby Cherayil for the strains used in this study. This work was supported by the National Institute of Allergy and Infectious Diseases Grant K22AI104755 (C.S.F.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

1. Preparation of Reagents
<ol>
	<li>Bile salts medium: To prepare tryptic soy broth (TSB) containing 0.4% bile salts (weight/volume), resuspend 200 mg of bile salts in 50 mL autoclaved TSB. Filter sterilize using a 0.22 &#181;m filter. Make fresh medium weekly. 
	Notes: The bile salts routinely used is a 1:1 mixture of sodium cholate and sodium deoxycholate isolated from ovine and bovine gallbladders. As demonstrated previously4, the presence of glucose was required for bile salt-induced biofilm formation. TSB has added glucose relative to Luria-Bertani (LB) broth; and therefore, was sufficient to induce biofilm formation in Shigella and the other enteric pathogens analyzed. Depending on the bacteria to be analyzed, different glucose concentrations or different sugar requirements might be needed.</li>
	<li>0.5% w/v crystal violet in water: Dissolve 2.5 g of crystal violet in 500 mL distilled water. Filter sterilize using a 0.22 &#181;m filter.</li>
	<li>Concanavalin A (ConA) conjugated to fluorescein isothiocyanate (FITC): Reconstitute the stock in 1x PBS. Dilute the 10 mg concentrated stock with 400 &#956;L of 1x PBS to a final concentration of 25 &#181;g/mL, and protect from light.</li>
	<li>PBS + Glucose: Dissolve 0.2 g glucose in 10 mL 1x PBS (2% w/v glucose final). Filter sterilize using a 0.22 &#181;m filter. Make fresh on the day of use.</li>
	<li>PBS + Bile Salts: Dissolve 40 mg in 10 mL 1x PBS (0.4% w/v bile salts final). Filter sterilize using a 0.22 &#181;m filter. Make fresh on the day of use.</li>
	<li>PBS + glucose and bile salts: Dissolve 40 mg bile salts and 0.2 g glucose in 10 mL 1x PBS (0.4% w/v bile salts and 2% w/v glucose final). Filter sterilize using a 0.22 &#181;m filter. Make fresh on the day of use.</li>
	<li>Prepare LB agar plates.</li>
	<li>Formaldehyde/glutaraldehyde fix: Add 810 &#181;L formaldehyde (37% stock solution, 3% final concentration) and 125 &#181;L glutaraldehyde (25% stock solution, 0.25% final concentration) to 14 mL 1x PBS. Mix thoroughly and store at 4 &#176;C. The fix should be cold for proper use. 
	Caution: The fix is toxic and requires hazardous waste disposal.</li>
	<li>Antifade mountant solution: Use antifade mountant solution containing 4,6-diamidino-2-phenylindole (DAPI) stain to inhibit photobleaching of immunofluorescent microscopy samples while fluorescently staining the DNA of the bacteria.</li>
</ol>
2. Preparation of Bacteria
<ol>
	<li>Grow overnight cultures of the bacterial strains to be tested by inoculating 3 mL of TSB with a single, well-isolated colony in a sterile culture tube. Incubate at 37 &#176;C with shaking at 225 rpm for overnight incubation (16 - 24 h). 
	NOTE: Strains should be restreaked from freezer stocks every 2 to 4 weeks, and maintained on plates no more than 2 weeks old.</li>
</ol>
3. Solid-phase Adherence Assay
NOTE: This assay quantifies adherent bacteria using a 96-well plate method. Bacteria are grown statically in flat bottom plates. Washing is performed to remove non-adherent bacteria and adherent bacteria are stained with crystal violet. The crystal violet stain binds peptidoglycan in the bacterial cell wall and can be solubilized using ethanol. The number of adherent bacteria is determined based on crystal violet retention.
<ol>
	<li>Set up two 1.5 mL tubes. Label with TSB or TSB + bile salts (BS).</li>
	<li>Add 1 mL of TSB or TSB + BS to the respective tubes.</li>
	<li>Inoculate tubes with 20 &#181;L of overnight culture (at a 1:50 dilution).</li>
	<li>In a sterile, clear, flat-bottomed, tissue culture-treated 96-well plate, add 130 &#181;L/well of uninoculated control media to three wells to serve as the blank control. Set up three control wells for each media type (TSB and TSB + BS) to be tested.</li>
	<li>Add 130 &#181;L/well of inoculated culture into three wells, and repeat until all experimental conditions are plated in triplicate.</li>
	<li>Incubate for 4 - 24 h at 37 &#176;C statically.</li>
	<li>Using a plate reader, record the OD600. Set the control wells as &#39;blank.&#39;&#160;Confirm the control medium is clear with no evidence of turbidity. If any turbidity is detected, discard the experiment. The OD600 values can be used to normalize the data if there are significant differences in growth rate between bacterial strains.</li>
	<li>Remove the culture medium using a vacuum line by gently tilting the plate and slowly aspirating the medium at the lower edge of the well. Be sure to collect all the culture medium without disrupting the adherent bacterial population located on the plastic surface. If EPS matrix was produced during the incubation time, the matrix will be visualized as a white precipitate. Do not disturb the EPS matrix.</li>
	<li>Gently wash the wells once with 200 &#181;L sterile PBS. Remove the PBS wash using the vacuum line.</li>
	<li>Invert the plate to dry. Allow a minimum of 20 min to dry. 
	NOTE: Since the biofilm must be thoroughly dried before staining, the protocol can be paused at this step for a few hours or even overnight. The added drying time will not alter the staining procedure while incomplete drying will affect quantification of the results and reproducibility.</li>
	<li>Add 150 &#181;L of 0.5% crystal violet to each experimental and control well.</li>
	<li>Incubate for 5 min at room temperature (RT).</li>
	<li>Wash the wells once with 400 &#181;L of distilled water. The added volume helps to remove residual crystal violet stain from the sides of the wells. Remove the wash with the vacuum line.</li>
	<li>Wash the wells five times with 200 &#181;L of distilled water. Remove the wash with the vacuum line. 
	NOTE: Thorough washing is important for quantification. When the blank wells are clear from the distilled water and do not contain any residual crystal violet stain, proceed to the next step.</li>
	<li>Invert the plate to dry, protected from light. Ensure complete drying of the plate as noted above.</li>
	<li>Destain the wells with 200 &#181;L of 95% ethanol. Incubate the plate on the shaker for 30 min. To avoid evaporation, particularly at higher temperatures, perform this step at 4 &#176;C.</li>
	<li>Using a plate reader, record the OD540. 
	NOTE: The wavelength for maximum absorbance of crystal violet is near 590 nm. The literature reports a range from OD540&#160;- OD5954,13,14,15,16 for crystal violet absorbance; thus choose a wavelength available based on the plate reader.</li>
</ol>
4. EPS Matrix Detection
NOTE: These complimentary assays quantify and visualize the EPS. In both, EPS is detected using a lectin to bind polysaccharides. The fluorescently-conjugated protein allows quantification (step 4.1) or visualization (step 4.2).
<ol>
	<li>Semi-quantitative detection of EPS
	<ol>
		<li>Set up two 1.5 mL tubes. Label with TSB or TSB + BS.</li>
		<li>Add 1 mL of TSB or TSB + BS to the respective tubes.</li>
		<li>Inoculate tubes with 20 &#181;L of overnight culture (a 1:50 dilution).</li>
		<li>In a sterile, black, flat-bottomed, tissue culture-treated 96-well plate, add 130 &#181;L/well of uninoculated control media to three wells to serve as the blank control. Set up three control wells for each medium type to be tested. Add 130 &#181;L/well of inoculated culture to the wells, and repeat until all experimental conditions are plated in triplicate.</li>
		<li>Incubate for 4 - 24 h at 37 &#176;C statically.</li>
		<li>Transfer the culture medium to a clear 96-well plate with a pipette, ideally a multichannel pipette. Be cautious to collect all the culture medium without disrupting the adherent population and/or the EPS located on the plastic surface. Set aside.</li>
		<li>Fix the black plate for 15 min at RT using 200 &#181;L/well of the formaldehyde/glutaraldehyde in 1x PBS.</li>
		<li>While the adherent population is fixing, evaluate the supernatant fraction from step 4.1.6. Using a plate reader, record the OD600. Set the control wells as &#39;blank.&#39;&#160;Confirm the control medium is clear with no evidence of turbidity. If any turbidity is detected, discard the experiment.
		<ol>
			<li>Alternatively, the entire assay can be performed in a black clear bottom plate. In those circumstances, the OD600 values can be recorded and the culture medium can be subsequently discarded prior to proceeding to step 4.1.7. The OD600 values can be used to normalize the data in the case there are significant differences in growth rate between bacterial strains.</li>
		</ol>
		</li>
		<li>Remove the fix and dispose in the hazardous waste.</li>
		<li>Gently wash the wells twice with 200 &#181;L/well of sterile PBS. Remove the PBS wash using the vacuum line by gently tilting the plate and slowly aspirating the wash at the lower edge of the well.</li>
		<li>Add 150 &#181;L/well of 25 &#181;g/mL ConA-FITC, and incubate for 15 min at RT.</li>
		<li>Gently wash twice with 200 &#181;L of PBS.</li>
		<li>Add 150 &#181;L of PBS to each well. Record the fluorescence at 488 nm.</li>
	</ol>
	</li>
	<li>Confocal microscopy visualization of EPS production and calculation of biofilm thickness
	<ol>
		<li>Set up two 1.5 mL tubes. Label with TSB or TSB + BS.</li>
		<li>Add 1 mL of TSB or TSB + BS to the respective tubes.</li>
		<li>Inoculate tubes with 20 &#181;L of overnight culture (at a 1:50 dilution).</li>
		<li>Set up a sterile 24-well plate with 12 mm sterile round glass coverslips. Add 400 &#181;L/well of uninoculated control media to three wells to serve as the blank control. Set up three control wells for each medium type to be tested.</li>
		<li>Add 400 &#181;L of inoculated culture to wells in duplicate, and repeat until all experimental conditions are plated.</li>
		<li>Incubate for 4 - 24 h at 37 &#176;C statically.</li>
		<li>Visually confirm growth in the TSB condition, growth and EPS precipitate (white) in the TSB + BS condition, and no growth and/or white precipitate in sterile control wells. Remove the supernatants.</li>
		<li>Fix for 15 min at RT using 200 &#181;L/well of formaldehyde/glutaraldehyde solution in 1x PBS.</li>
		<li>Remove the fix and dispose in hazardous waste.</li>
		<li>Gently wash the wells twice with 200 &#181;L/well of sterile PBS. Remove the PBS wash using the vacuum line.</li>
		<li>Add 150 &#181;L/well of 25 &#181;g/mL ConA-FITC and incubate for 15 min at RT.</li>
		<li>Gently wash twice with 200 &#181;L/well of PBS.</li>
		<li>Mount with antifade mountant solution with DAPI. 
		NOTE: The solution is a ready-made, glycerol-based mount solution containing DAPI for preserving the fluorescent label while simultaneously staining the DNA in bacterial cells for visualization as a counterstain. Follow the kit&#39;s directions and recommendations for incubation times prior to imaging samples. The DAPI staining procedure can be performed prior to applying antifade mountant solution that does not contain DAPI.</li>
		<li>Evaluate by confocal microscopy. On a confocal microscope, set the laser to 495 nm/519 nm excitation/emission for FITC visualization. Set a second laser to 360 nm/460 nm excitation/emission to visualize DAPI. Locate the biofilm by focusing on the DAPI channel. Use both the DAPI and FITC channels to determine the full thickness and set the upper and lower imaging borders. Record full thickness Z-stack images by capturing images every 0.25 &#181;m after setting perimeters to the top and bottom of the z-stack. For more information on confocal microscopy, please refer to publications by Paddock et al.17,18,19,20</li>
		<li>Reconstruct the 3D images in ImageJ (https://imagej.nih.gov/ij/) to visualize the full biofilm formation and calculate the biofilm thickness. The average thickness obtained for S. flexneri bile-salt induced biofilms was 14 &#181;m4.</li>
	</ol>
	</li>
</ol>
5. Dispersion Assay
NOTE: In this assay, the disassembly of biofilm through bacterial dispersion is detected. Here, mature biofilms are established and subsequently (usually the next day), the media are replaced with PBS or supplemented PBS. The supernatant component is then assessed to quantitate the number of bacteria that have dissociated from the biofilm.
<ol>
	<li>Set up two 1.5 mL tubes. Label with TSB or TSB + BS.</li>
	<li>Add 1 mL of TSB or TSB + BS to the respective tubes.</li>
	<li>Inoculate tubes with 20 &#181;L of overnight culture (at a 1:50 dilution).</li>
	<li>In a sterile, clear, flat-bottomed, tissue culture-treated 96-well plate, add 130 &#181;L/well of uninoculated control media to three wells to serve as the blank control. Set up three control wells for each media type to be tested.</li>
	<li>Add 130 &#181;L/well of inoculated culture to three wells, and repeat until all experimental conditions are plated in triplicate.</li>
	<li>Incubate the plate for 4 - 24 h at 37 &#176;C statically.</li>
	<li>Before continuing, warm the PBS, PBS + glucose, PBS + bile salts, and PBS + bile salts + glucose to 37 &#176;C. Prepare these reagents fresh daily.</li>
	<li>Using a plate reader, record the OD600. Set the control wells as &#39;blank.&#39;&#160;Confirm the medium is clear with no evidence of turbidity. If any turbidity is detected, discard the experiment. The OD600 values can be used to normalize the data if there are significant differences in growth rate between bacterial strains.</li>
	<li>Remove the culture medium using a vacuum line. Be sure to collect all the culture medium without disrupting the adherent population located on the plastic surface.</li>
	<li>Gently wash the wells twice with 200 &#181;L/well of sterile PBS. Remove the PBS wash using the vacuum line.</li>
	<li>Replace the wash with 130 &#181;L/well of the following into three wells each: PBS, PBS + glucose, PBS + bile salts, and PBS + bile salts + glucose. These reagents must be pre-warmed to 37 &#176;C.</li>
	<li>Incubate the plate for 30 min at 37 &#176;C.</li>
	<li>Carefully remove the plate from the 37 &#176;C incubator. Transfer the supernatants to a fresh, sterile 96-well plate.</li>
	<li>Using a 96-well plate or dilution block, prepare 10-fold (1:10) serial dilutions of the supernatant into sterile PBS. 
	NOTE: The dilution series should range from undiluted to 10-6 depending on the bacterial strain to be tested. Pilot experiments can be performed to determine the optimal dilution range.</li>
	<li>Spot plate 5 &#181;L of each dilution onto LB agar using a multichannel pipette. Incubate at 37 &#176;C overnight. Count colonies on the next day and account for the dilution factor when determining the recovery colony forming units (CFU). To calculate the percent dispersion relative to the 1x PBS control (set at 100%), divide the recovery CFU from each treatment condition by the recovery CFU from the 1x PBS control sample. 
	NOTE: Routine media plates for the bacterial strain of interest can also be used. For example, Shigella can be plated on Congo red plates. Ensure the plates are dry for appropriate spot-plating techniques.</li>
</ol>

The authors have no disclosures.

Analysis of biofilm formation is challenging due to the dynamic nature of biofilms and the variability between strains, materials, laboratories, and assays. Here, several strategies are presented to determine biofilm formation in enteric pathogens following bile salts exposure with experimental insight provided to promote reproducibility. There are additional considerations to ensure reproducibility. First and foremost, we recommend performing at least three independent experiments each with technical triplicates to confirm observations and statistical significance due to the variation that can occur. Second, the use of positive and negative controls is vital to ensuring reproducibility of the assay. It is highly recommended to use the wild type strain as a positive control when evaluating mutants for biofilm formation. Furthermore, as genes that are important for biofilm formation are identified (for example, see Vibrio22, Salmonella7, and E. coli23,24,25), the use of biofilm mutants as negative controls will validate the reproducibility of the assay. Third, the above methods are based on focused research with Shigella and S. flexneri, in particular. Adjustments to the protocols may be required depending on the growth requirements of each enteric pathogen evaluated. Fourth, the assays measure a bacterial stress response; and therefore, it is critical to work with appropriately maintained bacteria from frozen stocks (-80 &#176;C) or storage plates kept under refrigeration (maximum 4 &#176;C) for no more than two weeks. It is important to note that some bacteria including the enteric pathogens in the Yersinia genus are psychrotrophs that can grow at temperatures close to 0 &#176;C26. Since the storage at refrigerated temperatures can alter the bacterial physiology and possibly result in genotypic or phenotypic shifts27,28,29, it is highly recommended to prepare aliquots of the stock culture at -80 &#176;C for routine inoculation into starter cultures. Furthermore, working with clinical isolates requires more frequent restreaks from -80 &#176;C stocks as genetic shifts are rapidly observed when transitioning a clinically-derived bacterial strain into the laboratory setting30,31. Especially working with Shigella isolates, strains rapidly adapt and phenotype shifts occur often32,33. It is recommended to restreak bacteria from freezer stocks once a month, and restreak the stock plate every two weeks.
Another important factor to ensure reproducibility is fresh preparation of the bile salts media to limit variability of the assay. Bile salts media older than 1 week negatively impacted the robust phenotype observed with fresh media preparation. Moreover, depending on the pathogen and the type of exposure to be analyzed (small intestine versus colon), different sources and mixtures of bile salts may be required. For example, individual bile salts, conjugated and de-conjugated sources, or crude bile extracts that include additional bile components such as cholesterol and bilirubin may be utilized5,6. Shigella analyses have focused on bacterial adaptation to the host during transit of the small intestine prior to infection in the colon. Therefore, the use of a cholate and deoxycholate mixture mimics conditions of the small intestine5,6,34. Also with regards to the dispersion assay (section 5), it is specified that the PBS-supplemented reagents are to be made fresh daily and pre-warmed to 37 &#176;C prior to assay. When reagents older than a few days were used, reproducibility of the assay was severely compromised. It was also critical to pre-warm these reagents to 37 &#176;C as dispersion did not occur readily with room-temperature reagents. Finally, dispersion methods for assessment of complete titer enumeration of the biofilm can be used, which include sonication35,36 or enzymatic digestion37. The approach to the dispersion assay was directed toward assessing the effect of bile salt reabsorption as Shigella transits from the terminal ileum and into the colon. The analysis was based on the hypotheses that the loss of bile salts would enable dispersion of the transient biofilm and that Shigella must disperse the biofilm prior to causing infection in the colonic epithelium4,5. Conditions such as sonication and enzymatic digestion represent total release of microbial mass from the biofilm while the strategy presented here emulates the ileum to colon transition to capture a physiological phenotype for Shigella. Depending on the desired experimental conditions, additional methods for dispersion analysis or biofilm enumeration may be required.
The assays are designed to be moderately high-throughput with the use of 96-well plates to enable testing of multiple bacterial strains and/or conditions. To further ensure reproducibility of biofilm formation, flat-bottomed, 96-well plates that were tissue culture treated were used. For technical reasons, plate readers cannot accurately measure absorbance or fluorescence in round bottom plates. Additionally, uncoated plates offered variable adherence profiles in Shigella, which is an avenue of current investigation. The assays can be scaled to larger well formats proportionally, but will require the alterations to the protocol that can reduce experimental efficiency. For example, the culture media will need to be transferred to cuvettes to record the OD600 values with a spectrophotometer. Additionally, the crystal violet-stained biofilm can be scraped following a 30 - 60 min incubation in the 95% ethanol destain step and the contents can be transferred to a cuvette for spectrophotometer readings at OD540. Care must be taken when scrapping the contents of the biofilm in each well to avoid splashing, and we have found that scrapping is easier following a 60 min incubation.
Analysis with Shigella, Salmonella, and pathogenic E. coli4 demonstrated a repeated observation that biofilm formation occurs in an accelerated time scale (within 24 h) for enteric pathogens exposed to bile salts. Contrary to other bacteria that require 48 to 72 h to observe biofilm formation under normal conditions, bile salt-induced enteric biofilm begins as early as 4 h with the initial adherence phase. Subsequently, EPS matrix formation is evident by 6 h, and maturation occurs by 24 h. As such, the adherence phenotype at 24 h may be too robust to decipher the role of different adherence factors in biofilm formation. Performing early analyses, for example as early as 4 h, allows for identification of factors important in early biofilm formation that would otherwise be missed at a later time point. Finally, and as mentioned above in the protocol steps, bile salt-induced biofilm formation for Shigella was not detected in the absence of glucose4. Since the EPS matrix is comprised primarily of polysaccharides and is a critical step in biofilm formation for most bacteria38, the presence of glucose or other sugar in the media is most likely important to observe biofilm formation. Depending on the bacterial pathogen analyzed, different base media formulations may be required.
Biofilm formation is a multilayered process that often occurs in an open system where nutrient flux and fluid motion readily occur1,2,3. The methods are limited by the closed culture and static growth requirements; however, the methods enable assessment of biofilm formation in the laboratory setting. Upon identification of the biofilm phenotype, further experiments using fluid cells or continuous culture strategies39,40,41 may provide additional mechanistic insight into physiological or clinical significance of biofilm formation. For identification of the EPS matrix, the protocols use the polysaccharide-binding lectin ConA. ConA binds &#945;-D-glucosyl or sterically-closely related residues42. Different lectins may be required depending on type of EPS produced for different genera or species. An alternative strategy to identify EPS production is mass spectrometry, which will provide both verification that EPS is produced and determine the composition of the EPS matrix43,44. Additionally, the use of the DAPI counterstain can result in significant brightness given the number of bacteria in the biofilm or the occurrence of extracellular DNA that is often present in the EPS matrix45,46.
There are alternatives to the above methods described in the literature. For example, the solid-phase binding assay has been reported for use in other pathogens (for example, references4,25,47,48,49). There are modifications to the protocol between publications; and therefore, the strategies presented here are an easy-to-replicate format. It is important to note that the protocols specify air-drying biofilms prior to staining (step 3.10) while some groups prefer to fix biofilms. The two different approaches may represent varying pathogen-specific strategies. Additionally, by preparing the crystal violet stain in water, a consistent biofilm formation phenotype was detected. Other groups use ethanol to solubilize crystal violet; which for Shigella, led to destabilization of biofilms and compromised reproducibility. We are currently investigating the factors that may be ethanol soluble in the Shigella biofilm. Another example is the quantification of the EPS production by ConA staining. A 96-well plate fluorescent detection method (step 4.1) is presented to determine semi-quantitative values reflecting the amount of FITC-conjugated ConA binding to the biofilm. To our knowledge, the method is the first time this strategy has been reported. EPS detection via microscopic examination is accepted in the literature and allows for a different, but complimentary, strategy to visualize EPS. The semi-quantitative fluorescence detection combined with the confocal microscopic detection of EPS certainly strengthened the data, especially given the limitations of the DAPI counterstain procedure in the microscopy analysis described above. In all, a significant strength in the set of assays described here is that each technique complements another approach to give rise to a dynamic assessment of bile salt-induced biofilm formation in enteric pathogens.
As bile salts and virulent biofilms are incorporated into the paradigm of enteric bacterial pathogenesis, it is anticipated that these methods will be used to validate models, identify roles of specific genes, identify genes of unknown function, and provide general characterization of clinical strains. This multifaceted approach allows the identification of genes important in one aspect of biofilm formation that may be lost once the mature biofilm is established, which therefore enables identification of specialized functions of each bacterial component of the biofilm. Furthermore, as evident by the fact that pathogen adaptation within the human intestine is a rapidly occurring phenomenon, the role of bile salt-induced biofilm formation is an example of a newly appreciated, conserved, and critical phenomenon that occurs in Shigella infection as well as additional enteric pathogens.

Biofilm formation is an important bacterial survival strategy induced during harsh environmental conditions. Exposure to bactericidal compounds like antibiotics or changes in nutrient or oxygen availability induces a stressed state in bacteria that can be alleviated through biofilm formation. A biofilm is characterized by bacterial attachment to a surface or other bacteria and is accompanied by the secretion of an EPS matrix primarily composed of polysaccharides1,2,3. Biofilm formation is a dynamic process in which a cascade of events culminates in formation of a mature adherent bacterial community1,2,3. Bacteria produce adhesins to facilitate early attachment while shifting adhesin gene expression profiles to strengthen attachment during biofilm maturation. Simultaneously, EPS production occurs to coat the bacterial community in a matrix to protect the cells from the initial stressor. Bacteria contained within the biofilm are slow growing; and as such, renders most antibiotics ineffective. Furthermore, the slow growth conserves energy until conditions change to favor bacterial growth1,2,3. After the harsh conditions have passed, bacteria disperse the biofilm and resume a planktonic lifestyle1,2,3. Traditionally, biofilms are observed on surfaces and represent a persistent clinical challenge due to infection reservoirs present on catheters and in-dwelling devices1,2,3.
Biofilm formation was recently described for several enteric pathogens; bacteria that infect the small intestine or colon4. For Shigella species, infection occurs in the human colon after a transit through the majority of the gastrointestinal tract. During passage through the small intestine, Shigella is exposed to bile; a lipid-degrading detergent secreted into the intestine to facilitate digestion of lipids while simultaneously killing most bacteria5. Enteric pathogens have a unique ability to resist the bactericidal effects of bile6. Our recent analysis utilized in vivo-like combinations of glucose and bile salts to demonstrate robust biofilm formation in S. flexneri as well as other species of Shigella, pathogenic Escherichia coli, and Salmonella4. Previously, Salmonella enterica serovar Typhi was shown to form a bile-induced biofilm due to unique colonization of the gallbladder during chronic infection7,8,9,10. Additionally, prior research with Vibrio11&#160;and Campylobacter12 demonstrated biofilm formation in response to bile. Therefore, the analyses extended the bile-induced biofilm formation observations to other pathogens and help to establish demonstration of a conserved enteric pathogen response to bile. Unlike chronic biofilms in which bacterial gene transcription is limited and cell senescence can occur1,2,3, we propose that the enteric bile-induced biofilm is more transient in nature. This transient, virulent biofilm is hallmarked by a rapid disassembly (as seen in the dispersion assay) and enhanced virulence gene expression observed in the biofilm population4,6.
As biofilm formation is a multifaceted, dynamic process and the use of bile salts as an initiating factor has only been recently described for most enteric pathogens, the tools and techniques used are unique and creative applications of traditional methods. Thus, presented here are three complimentary strategies to quantify several important characteristics of bile salt-induced biofilm formation, including bacterial adherence, production of the EPS matrix, and dispersion of viable bacteria from the biofilm. These techniques have been utilized primarily for research with Shigella; and therefore, evaluation of other enteric pathogens may require optimization. Nevertheless, positive data from all three assays support identification of biofilms and establish reproducible protocols for bile salt-induced biofilm formation.

<table><tbody><tr><td>Tryptic Soy Broth</td><td>Sigma-Aldrich</td><td>22092-500G</td><td></td></tr><tr><td>Crystal Violet</td><td>Sigma</td><td>C6158-50</td><td></td></tr><tr><td>Concanavalin-A FITC</td><td>Sigma</td><td>C7642-10mg</td><td></td></tr><tr><td>Glucose</td><td>Sigma</td><td>G7021-1KG</td><td></td></tr><tr><td>Bile Salts</td><td>Sigma</td><td>B8756-100G</td><td></td></tr><tr><td>LB Agar</td><td>Sigma</td><td>L7533-1KG</td><td></td></tr><tr><td>14 mL culture tubes, 17 x 100 mm, plastic, sterile</td><td>Fisher</td><td>14-959-11B</td><td></td></tr><tr><td>Vectashield hard-set antifade with DAPI</td><td>Vector Laboratories</td><td>H-1500</td><td></td></tr><tr><td>Formaldehyde</td><td>Sigma-Aldrich</td><td>F1635-500</td><td></td></tr><tr><td>Gluteraldehyde</td><td>Sigma-Aldrich</td><td>G6257</td><td></td></tr><tr><td>Flat-bottomed 96-well plates (clear)</td><td>TPP</td><td>92696</td><td></td></tr><tr><td>Flat-bottomed 96-well plates (black)</td><td>Greiner Bio-One</td><td>655076</td><td></td></tr><tr><td>Flat-bottomed 24-well plates (clear)</td><td>TPP</td><td>92424</td><td></td></tr><tr><td>Glass coverslips 12mm, round</td><td>Fisher</td><td>08-774-383</td><td></td></tr><tr><td>96-well plate reader</td><td>Spectramax</td><td></td><td></td></tr><tr><td>Flourescent plate reader</td><td>Biotek Synergy 2</td><td></td><td></td></tr><tr><td>Confocal or Fluorescent Microscope</td><td>Nikon A1 confocal microscope</td><td></td><td></td></tr><tr><td>37&amp;deg;C Shaking Incubator</td><td>New Brunswick Scientific Excella E25</td><td></td><td></td></tr><tr><td>37&amp;deg;C Plate Incubator</td><td>Thermolyne Series 5000</td><td></td><td></td></tr></tbody></table>

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.

In Figure 1, biofilm formation is induced in most of the six enteric pathogens tested following growth in media containing bile salts. A significant increase in adherent bacteria after bile salts exposure is observed in nearly all strains tested. The exception is enteroaggregative E. coli (EAEC); however, note the induced observation of the &#916;aaf mutant4. The results indicate that additional adherence mechanisms are induced by bile salts exposure in EAEC in the absence of aggregative adherence fimbriae I (AAF/I)21. To draw conclusions from this data set, plot the OD540 values for each strain and media type tested. Comparing the values from each strain in the media control (-BS) relative to media containing bile salts will determine if bile salts significantly induce biofilm formation. Comparisons across bacterial strains and/or mutants can also be performed for further characterization analyses.
The ConA-FITC assessment of EPS is presented in Figure 2. Figure 2A represents the semi-quantitative assessment of EPS production in which ConA-FITC is used to stain the biofilms. The ConA binds to the polysaccharides in the EPS matrix and the amount retained is detected by a fluorescent plate reader. To present the data, plot the average fluorescence units for each condition and compare the biofilm-inducing conditions to the negative control. Additionally, comparisons across bacterial strains or mutants can be performed. It is critical to analyze a media-only, negative control to determine the amount of background ConA-FITC fluorescence that occurs for the plate. The TSB condition was not significantly different from the media control, while the fluorescence readings significantly increased following bile salts exposure. The data confirm that EPS production is indicative of biofilm formation and does not occur in the absence of bile salts for S. flexneri. The images of bile salt-induced biofilms in S. flexneri is represented in Figure 2B. ConA-FITC is used to stain polysaccharides in the EPS matrix while a DAPI counterstain is used to visualize the bacteria by staining the DNA. For bacteria not exposed to bile salts (negative control), only DAPI is detected. Notably, the density of bacteria is much lower than in the bile salts condition. A clear background on the FITC channel is obtained to indicate the lack of EPS production. As expected, the data demonstrate that EPS production is correlated with biofilm formation1,2,3 following bile salts exposure. The -ConA staining control in the bile salt-treated samples is provided to demonstrate the specificity of the FITC fluorescence. For these images, 1x PBS was used in place of ConA-FITC and the DAPI counterstain was maintained. Imaging can be performed with a confocal microscope to assess the thickness of biofilms with the EPS-positive samples4.
Bacterial dispersion from biofilms in Figure 3 is detected by incubating established biofilms in PBS or supplemented PBS. Physiologically, bile is reabsorbed into circulation in the small intestine and only 5% of bile enters the colon5. As Shigella invades the colonic epithelium, we hypothesize that reabsorption of bile is an important signal for biofilm dispersion. In this figure, dispersion occurs in biofilms exposed to PBS or PBS + glucose; however, dispersion is inhibited in PBS + bile salts regardless of the presence of glucose. The data demonstrate that the removal of bile salts is necessary and sufficient to induce dispersion of S. flexneri from bile salt-induced biofilms4. To plot the data, either the recovered CFU of the bacteria or the relative percent to the control (as shown here) can be plotted.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57322/57322fig1.jpg" /> 
Figure 1: Bile salts induce biofilm formation in enteric pathogens. 18 h biofilms were grown on tissue-culture treated 96-well plates in either TSB or TSB + 0.4% bile salts media and stained with 0.5% crystal violet. Quantitative values from ethanol solubilization of the crystal violet stain were determined by spectrophotometry at OD540. Bile salts exposure significantly increased biofilm formation in Shigella flexneri, S. dysenteriae, enteropathogenic Escherichia coli, Salmonella enterica serovar Typhi, and S. enterica serovar Typhimurium. Statistical significance was determined by the student&#39;s T-test for each strain comparing the bile salts treatment to the media control. The standard error of the mean (SEM) is represented by the error bars. All p-values are &#60;0.0001. Data shown are from one (n = 3) representative experiment, with a full set of data provided in a previous publication4. <a href="/files/ftp_upload/57322/57322fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57322/57322fig2.jpg" /> 
Figure 2: Bile salt-induced biofilm formation is characterized by production of a EPS matrix that is detected with FITC-labeled Concanavalin A. (A) Sterile black, 96-well plate were seeded with S. flexneri at a 1:50 dilution into TSB or TSB + bile salts and grown for 18 h. Biofilms were fixed, gently washed, and stained with ConA-FITC. The amount of retained Concanavalin-A FITC was determined by a fluorescent plate reader. The values plotted demonstrate EPS quantification following growth of S. flexneri grown in TSB or TSB-containing bile salts. Significance was determined by one-way ANOVA and p-values are &#60;0.01. The SEM is represented by the error bars. Data shown are from one (n = 3) representative experiment, with a full set of data provided in a previous publication4. (B) Microscopy images from a bile salt-induced Shigella biofilm. Sterile coverslips were seeded with S. flexneri at a 1:50 dilution in TSB or TSB + bile salts and grown for 18 h. The coverslips were subsequently fixed and stained with ConA-FITC and/or counterstained with DAPI. ConA-FITC was only detected in the bile salts condition due to the presence of the EPS matrix. A coverslip stained only with DAPI in the TSB + bile salts treatment was used to demonstrate minimal background fluorescence for the FITC setting on the confocal microscope. Images for each wavelength channel and a composite overlay are provided for all treatment conditions. Scale bars indicate 10 &#181;m. Data shown are from one (n = 3) representative experiment, with a full set of data provided in a previous publication4. <a href="/files/ftp_upload/57322/57322fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57322/57322fig3.jpg" /> 
Figure 3: Analysis of bacterial dispersion from bile salt-induced biofilms. 18 h bile salt-induced Shigella flexneri biofilms were exposed to PBS or PBS supplemented with glucose, bile salts, or a combination of glucose and bile salts. The supernatant was subsequently serially diluted and plated on agar plates to enumerate the colony forming units (CFUs) that dispersed the biofilm. The number of bacteria recovered was plotted relative to the PBS control. Bacteria dispersed from the biofilm in the PBS or PBS + glucose conditions, but the presence of bile salts was sufficient to inhibit bacterial dispersion from the biofilm. Data shown are from one (n = 3) representative experiment, with a full set of data provided in a previous publication4. For reference, 100-fold more bacteria were routinely recovered from the PBS control treatment (approximately 7 x 107 CFU) compared to the PBS + bile salts treatment (approximately 7 x 105 CFU). Significance was determined by one-way ANOVA and p-values are &#60;0.0001. The SEM is represented by the error bars.

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

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, ...

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Nanyang Technological University

Research

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Immunology and Infection

12.4K Views.  Massachusetts General Hospital. Dit protocol laat de lezer om te analyseren bile salt-geïnduceerde biofilm vorming in de darmen ziekteverwekkers met behulp van een veelzijdige aanpak om vast te leggen van het dynamische karakter van het bacteriële biofilms door beoordeling van de naleving, extracellulaire stof van polymere matrix vorming, en dispersie.

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

An Analytical Tool-box for Comprehensive Biochemical, Structural and Transcriptome Evaluation of Oral Biofilms Mediated by Mutans Streptococci

Biofilms are highly dynamic, organized and structured communities of microbial cells enmeshed in an extracellular matrix of variable density and composition 1, 2. In general, biofilms develop from initial microbial attachment on a surface followed by formation of cell clusters (or microcolonies) and further development and stabilization of the microcolonies, which occur in a complex extracellular matrix. The majority of biofilm matrices harbor exopolysaccharides (EPS), and dental biofilms are no exception; especially those associated with caries disease, which are mostly mediated by mutans streptococci 3. The EPS are synthesized by microorganisms (S. mutans, a key contributor) by means of extracellular enzymes, such as glucosyltransferases using sucrose primarily as substrate 3.
Studies of biofilms formed on tooth surfaces are particularly challenging owing to their constant exposure to environmental challenges associated with complex diet-host-microbial interactions occurring in the oral cavity. Better understanding of the dynamic changes of the structural organization and composition of the matrix, physiology and transcriptome/proteome profile of biofilm-cells in response to these complex interactions would further advance the current knowledge of how oral biofilms modulate pathogenicity. Therefore, we have developed an analytical tool-box to facilitate biofilm analysis at structural, biochemical and molecular levels by combining commonly available and novel techniques with custom-made software for data analysis. Standard analytical (colorimetric assays, RT-qPCR and microarrays) and novel fluorescence techniques (for simultaneous labeling of bacteria and EPS) were integrated with specific software for data analysis to address the complex nature of oral biofilm research.
The tool-box is comprised of 4 distinct but interconnected steps (Figure 1): 1) Bioassays, 2) Raw Data Input, 3) Data Processing, and 4) Data Analysis. We used our in vitro biofilm model and specific experimental conditions to demonstrate the usefulness and flexibility of the tool-box. The biofilm model is simple, reproducible and multiple replicates of a single experiment can be done simultaneously 4, 5. Moreover, it allows temporal evaluation, inclusion of various microbial species 5 and assessment of the effects of distinct experimental conditions (e.g. treatments 6; comparison of knockout mutants vs. parental strain 5; carbohydrates availability 7). Here, we describe two specific components of the tool-box, including (i) new software for microarray data mining/organization (MDV) and fluorescence imaging analysis (DUOSTAT), and (ii) in situ EPS-labeling. We also provide an experimental case showing how the tool-box can assist with biofilms analysis, data organization, integration and interpretation.

Biofilms formed on tooth surfaces are highly complex and exposed to constant innate and exogenous environmental challenges, which modulate their architecture, physiology and transcriptome. We developed a toolbox to examine the composition, structural organization and gene expression of oral biofilms, which can be adapted to other areas of biofilm research.

Biofilms are highly dynamic, organized and structured communities of microbial cells enmeshed in an extracellular matrix of variable density and ...

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 ...

Concurrent Quantification of Cellular and Extracellular Components of Biofilms

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

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

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

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 ...

Protocols for Analyzing Shigella Infection in Colonic Epithelial Cells

Epithelial Cell Infection Analyses with Shigella

The human-adapted enteric bacterial pathogen Shigella causes millions of infections each year, creates long-term growth effects among pediatric patients, and is a leading cause of diarrheal deaths worldwide. Infection induces watery or bloody diarrhea as a result of the pathogen transiting the gastrointestinal tract and infecting the epithelial cells lining the colon. With staggering increases in antibiotic resistance and the current lack of approved vaccines, standardized research protocols are critical to studying this formidable pathogen. Here, methodologies are presented to examine the molecular pathogenesis of Shigella using in vitro analyses of bacterial adherence, invasion, and intracellular replication in colonic epithelial cells. Prior to infection analyses, the virulence phenotype of Shigella colonies was verified by the uptake of the Congo red dye on agar plates. Supplemented laboratory media can also be considered during bacterial culturing to mimic in vivo conditions. Bacterial cells are then used in a standardized protocol to infect colonic epithelial cells in tissue culture plates at an established multiplicity of infection with adaptations to analyze each stage of infection. For adherence assays, Shigella cells are incubated with reduced media levels to promote bacterial contact with epithelial cells. For both invasion and intracellular replication assays, gentamicin is applied for various time intervals to eliminate extracellular bacteria and enable assessment of invasion and/or the quantification of intracellular replication rates. All infection protocols enumerate adherent, invaded, and/or intracellular bacteria by serially diluting infected epithelial cell lysates and plating bacterial colony forming units relative to infecting titers on Congo red agar plates. Together, these protocols enable independent characterization and comparisons for each stage of Shigella infection of epithelial cells to study this pathogen successfully.

Author Spotlight: Advancements in Understanding and Combatting Shigella Infections

The present protocol describes infection assays to interrogate Shigella adherence, invasion, and intracellular replication using in vitro epithelial cell lines.

The human-adapted enteric bacterial pathogen Shigella causes millions of infections each year, creates long-term growth effects among pediatric patients, ...

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 Solid Phase Adherence Assay to Assess Biofilm Formation

Source: Nickerson, K. P. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/57322">Bile Salt-induced Biofilm Formation in Enteric Pathogens: Techniques for Identification and Quantification.</a>&#160;J. Vis. Exp.&#160;(2018)
The video demonstrates a solid-phase adherence assay for assessing biofilm formation. In a multi-well plate, bacteria in bile salt-supplemented media adhere to solid surfaces, forming biofilms, which is confirmed by colorimetry through crystal violet staining and ethanol treatment.

1. Preparation of Reagents

	Bile salts medium: To prepare tryptic soy broth (TSB) containing 0.4% bile salts (weight/volume), resuspend 200 mg of bile ...

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Immunology

Immunodiagnostics

Specialized Assays

An Assay to Study Bile-Salt Induced Biofilm Formation through EPS Matrix Detection

Source: Nickerson, K. P., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/57322">Bile Salt-induced Biofilm Formation in Enteric Pathogens: Techniques for Identification and Quantification.</a> J. Vis. Exp. (2018).
This video demonstrates a fluorescence-based method for quantifying bile salt-induced biofilm formation. Upon the addition of bile salts in the culture of a pathogenic enteric bacteria, biofilm production is detected using a fluorescent lectin that binds to the polysaccharides in the extracellular polymeric substance (EPS) matrix of the biofilm.

1. Preparation of Reagents

	Bile salts (BS) medium: To prepare tryptic soy broth (TSB) containing 0.4% bile salts (weight/volume), resuspend 200 mg of ...

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

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