Name: a platform of anti-biofilm assays suited to the exploration of natural compound libraries
Uploaded: 2016-12-27T12:00:00.000+00:00
Duration: 9 min 39 s
Description: Watch this Scientific Journal Video about A Platform of Anti-biofilm Assays Suited to the Exploration of Natural Compound Libraries at JoVE.com

The authors thank professor Paul Cos and LMPH, University of Antwerp, Belgium for his support during the filming process in his laboratory. This work was funded by Academy of Finland projects (projects 272266 and 282981) and Svenska Tekniska Vetenskapsakademien i Finland. The technical contributions of MSc Janni Kujala are acknowledged.

1. Growing the Bacteria
<ol>
	<li>Pre-culture the bacteria overnight in a tryptic soy broth (TSB) at 37 &#176;C with 220 rpm shaking (16-18 hr).</li>
	<li>Dilute the pre-culture 100-1,000 times (depending on the growth rate of the bacteria; here, 1,000 times is used for S. aureus) in fresh TSB and let it grow at 37 &#176;C and 200 rpm to reach exponential growth (optical density at 595 nm (OD595) between 0.2 and 0.6). 
	NOTE: This step requires strain-specific optimization.</li>
</ol>
2. Biofilm Formation: Pre- and Post-exposure
<ol>
	<li>Dilute the exponentially grown culture 100 times (this equals approximately 106 colony-forming units per milliliter (CFU/ml)).</li>
	<li>Pre-exposure protocol
	<ol>
		<li>For untreated control samples, add 200 &#181;l of the diluted bacterial culture per well of a sterile 96-microwell plate.</li>
		<li>Add 4 &#181;l of a test compound or a control antibiotic (50x stock solution) and 196 &#181;l of the diluted bacterial culture per well.</li>
		<li>Incubate it at 37 &#176;C on a plate shaker at 200 rpm for 18 hr. 
		Note: Here we use a shaker with a 2 mm orbit.</li>
	</ol>
	</li>
	<li>Post-exposure protocol
	<ol>
		<li>For all samples, add 200 &#181;l of the diluted bacterial culture per well of a sterile 96-microwell plate.</li>
		<li>Incubate it at 37 &#176;C on a plate shaker at 200 rpm for 18 hr. 
		Note: Here we use a shaker with a 2 mm orbit.</li>
		<li>Remove the entire planktonic solution carefully, without touching the biofilm, using a multichannel pipette.</li>
		<li>Add 4 &#181;l of a test compound or a control antibiotic (50x stock solution) and 196 &#181;l of TSB per well.</li>
		<li>Incubate it at 37 &#176;C on a plate shaker at 200 rpm for an additional 24 hr. 
		Note: Here we use a shaker with a 2 mm orbit.</li>
	</ol>
	</li>
</ol>
3. Resazurin Staining Protocol for the Viability Assessment of Biofilms
<ol>
	<li>Prepare a stock solution of 0.1 mg/ml resazurin (0.4 mM) in sterile PBS. Keep this stock sterile, protected from light exposure, and at 4 &#176;C.</li>
	<li>Dilute the resazurin stock 1:50 in sterile phosphate-buffered saline (PBS) to achieve a final concentration of 20 &#181;M.</li>
	<li>Transfer the entire planktonic solution (leaving the biofilms in the wells) carefully, without touching the biofilms or creating air bubbles, to a separate, clean 96-well plate using a multichannel pipette.</li>
	<li>Wash the biofilms once with sterile PBS by adding 200 &#181;l per well, and remove it carefully using a multichannel pipette.</li>
	<li>Add 200 &#181;l of the diluted resazurin per well of the biofilm plate using a multichannel pipette.</li>
	<li>Incubate it in darkness, at room temperature (RT), and 200 rpm, shaking for approximately 20 min until the untreated biofilm controls are evenly pink.</li>
	<li>Measure the fluorescence at &#955;exc = 560 nm and &#955;em = 590 nm with a plate reader (top reading).</li>
</ol>
4. Crystal Violet Staining Protocol for Biomass Quantification of Biofilms Using the Same Plate as in Step 3
<ol>
	<li>Remove the resazurin stain carefully from the wells using a multichannel pipette.</li>
	<li>Fix the biofilms with 200 &#181;l of methanol for 15 min.</li>
	<li>Let the plate air dry for 10 min.</li>
	<li>Add 190 &#181;l of 0.02 % (vol/vol, diluted in deionized water) crystal violet solution, carefully using a multichannel pipette. Avoid touching the sides of the wells with the stain while pipetting and prevent the formation of air bubbles by not pressing the pipette to complete blow-out.</li>
	<li>Incubate it for 5 min at RT.</li>
	<li>Remove the stain carefully, using a multichannel pipette.</li>
	<li>Wash it twice with deionized water (200 &#181;l each time).</li>
	<li>Let the wells dry for 5 min at RT and dissolve the remaining stain in 96% ethanol or 33% acetic acid.</li>
	<li>Incubate it for 1 hr at RT and read the absorbance at 595 nm.</li>
</ol>
5. Evaluating the Bactericidal Effect on Planktonic Bacteria Using the Same Sample Plate as for the Biofilms
<ol>
	<li>Measure the turbidity at 595 nm of the plate with the planktonic solution from 3.3.</li>
	<li>Stain the planktonic bacteria with resazurin by adding 10 &#181;l of the resazurin stock per well. Mix well by pipetting.</li>
	<li>Incubate it in darkness at RT for approximately 5 min, until the untreated controls are evenly pink.</li>
	<li>Measure the fluorescence at &#955;exc = 560 nm and &#955;em = 590 nm.</li>
</ol>
6. Wheat Germ Agglutinin Staining for Matrix Quantification and Fluorescence Microscopy Imaging of Biofilms
<ol>
	<li>Matrix quantification
	<ol>
		<li>Prepare a 5 &#181;g/ml solution of the WGA probe in sterile PBS.</li>
		<li>Use a parallel sample plate for this staining. Remove the planktonic solution from the wells and wash once with sterile PBS (200 &#181;l per well), carefully using a multichannel pipette without touching the biofilms.</li>
		<li>Add 200 &#181;l of WGA solution per well to be stained.</li>
		<li>Incubate it in darkness at 4 &#176;C for 2 hr.</li>
		<li>Remove the unbound stain by washing the wells with 200 &#181;l of PBS three times.</li>
		<li>Let the plate dry for 15 min at RT.</li>
		<li>Dissolve the bound stain in 33% acetic acid, using 200 &#181;l per well.</li>
		<li>Seal the wells with strip caps and sonicate them using a water bath sonicator for 30 sec at RT and 40 kHz.</li>
		<li>Incubate the plate for 1 hr at RT.</li>
		<li>Repeat the sonication step. The wells can remain sealed between the sonication steps.</li>
		<li>Measure the fluorescence at &#955;ex = 495 nm and &#955;em = 520 nm with a plate reader (top reading).</li>
	</ol>
	</li>
	<li>Visualizing the matrix with fluorescence microscopy
	<ol>
		<li>Use the same protocol as above until step 6.1.6.</li>
		<li>After the drying step, visualize the samples with a fluorescence microscope, using a FITC filter (or another suitable green excitation filter).</li>
	</ol>
	</li>
</ol>
7. Staining Viable and Dead Cells within the Biofilms for Imaging with Fluorescence Microscopy and Quantification of the Signal for the Green-to-red-ratio (G/R)
<ol>
	<li>Imaging with fluorescence microscopy
	<ol>
		<li>Prepare a solution of each probe (one staining viable cells and the other one staining dead cells), according to the manufacturer&#39;s guidelines.</li>
		<li>Remove the planktonic solution from the wells and wash once with sterile PBS (200 &#181;l per well) using a multichannel pipette.</li>
		<li>Add 6 &#181;l of the staining solution per well.</li>
		<li>Incubate the plate in darkness for 15 min.</li>
		<li>Before the microscopy, remove the excess liquid manually using a multichannel pipette.</li>
		<li>Capture the images using a fluorescence microscope, using for instance a FITC filter (for green fluorescence, viable cells) or a TRITC filter (red fluorescence, dead cells).</li>
	</ol>
	</li>
	<li>Quantification of the signal as a green-to-red-fluorescence ratio (G/R)
	<ol>
		<li>Follow the same protocol as in steps 7.1.1 and 7.1.2.</li>
		<li>Add 200 &#181;l of the staining solution per well and incubate them for 15 min in darkness.</li>
		<li>Measure the fluorescence with a plate reader at excitation/emission wavelengths 485/535 nm and 485/635 for green and red fluorescence, respectively (top reading).</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to disclose.

There is no single method that can simultaneously measure the effect of a compound on the viability, biomass, and biofilm matrix. Therefore, there is a need for combining assays in order to detect an effect on the three endpoints, preferably at a primary screening stage.
Resazurin is a very simple staining protocol consisting only of the addition of the redox probe. However, establishing the optimal incubation time of the biofilms with the resazurin is crucial to the success of this assay. In some bacterial strains, the reduction of the resazurin probe to the pink, fluorescent resorufin occurs very quickly, while in others it can last several hours19. Moreover, within the same bacterial strain, there may also be significant differences between planktonic bacteria and biofilms. The planktonic bacteria are typically more easily reached, mainly because the bacterial density is lower in planktonic populations than in biofilm populations, which promotes faster turnover of the reaction. In a published comparison of biofilm viability staining probes, resazurin was concluded to be one of the most accurate, simple to use, and least expensive assays19. Moreover, it has been shown to be applicable to a range of bacterial and fungal organisms19. On the other hand, the crystal violet assay is the most widely used for biofilm mass quantification19,32,33. The most critical steps of this protocol are the addition and removal of the crystal violet stain solution. These steps have to be conducted very carefully to avoid unspecific staining (e.g., due to drops of stain on the walls of the wells). One of the main benefits of using resazurin as the initial staining assay is the fact that it is non-toxic to the cells, which enables the use of the same plate for crystal violet staining. The combination of both assays significantly simplifies the workflow, lowers the costs, saves consumables, and minimizes the use of the test compounds, which is particularly valuable in a screening environment.
As indicated earlier, the self-produced extracellular polysaccharide matrix is an essential component of biofilms. To measure the matrix content (particularly essential polysaccharides), a third assay was included here, based on fluorescence-labeled WGA. Originally the WGA quantification was described as an enzyme-linked lectin-sorbent assay (ELLA)24. A similar assay based on staining glycosaminoglycans (GAGs) in the matrix of S. aureus biofilms with dimethylmethylene blue (DMMB) has also been used31,34. GAGs are similar to the polysaccharide intercellular adhesins (PIAs) found in the matrix of Staphylococcus spp. biofilms35. The WGA-assay similarly targets PIAs, but WGA more specifically binds to poly-N-acetyl-glucosamine residues, which play an essential role in the biofilm matrix36. Targeting the matrix is of high importance due to the fact that biofilms can easily regrow if the matrix is left after a chemical or antibiotic treatment. Bacteria can also more easily attach to a surface that is matrix pre-coated35. It has been shown that some antibiotics can effectively lower the viability and biofilm biomass, but without any effect on the matrix. In our experiments, this is exemplified by the case of ciprofloxacin31. Some antibiotics can even increase matrix production, as demonstrated here with penicillin G26. Based on these changes, antibiotics can be classified into the categories listed in Table 2. All the hits in our screening studies (Figure 2) can be classified, together with ciprofloxacin, as compounds that kill the bacteria in the biofilm and reduce the biomass but do not disassemble or disrupt the biofilm matrix.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Category</td>
			<td>Effect on bacteria</td>
			<td>Effect on matrix</td>
			<td>Example antibiotic (target biofilm)</td>
			<td>Reference</td>
		</tr>
		<tr>
			<td>1</td>
			<td>none</td>
			<td>none</td>
			<td>ampicillin (SA)</td>
			<td>(Tot&#233; et al. 2009)</td>
		</tr>
		<tr>
			<td>2</td>
			<td>none</td>
			<td>&#8210;</td>
			<td>cefalotin (SA)</td>
			<td>(Tot&#233; et al. 2009)</td>
		</tr>
		<tr>
			<td>3</td>
			<td>&#8210;</td>
			<td>none</td>
			<td>polymyxin (SA)</td>
			<td>(Tot&#233; et al. 2009)</td>
		</tr>
		<tr>
			<td rowspan="2">4</td>
			<td rowspan="2">&#8210;</td>
			<td rowspan="2">&#8210;</td>
			<td>kanamycin (PA)</td>
			<td>(Tot&#233; et al. 2009)</td>
		</tr>
		<tr>
			<td>ciprofloxacin (SA)</td>
			<td>(Skogman et al. 2012)</td>
		</tr>
		<tr>
			<td>5</td>
			<td>&#8210; (+)</td>
			<td>&#8210;</td>
			<td>doxycycline (PA)</td>
			<td>(Tot&#233; et al. 2009)</td>
		</tr>
		<tr>
			<td>6</td>
			<td>&#8210;</td>
			<td>+</td>
			<td>penicillin G (SA, EC)</td>
			<td>(Skogman et al. 2012)</td>
		</tr>
	</tbody>
</table>
Table 2: Classification of antibiotics. The antibiotics are divided into categories based on their effect on the viability of the biofilm bacteria (using resazurin staining) and the matrix (using WGA or dimethylmethylene blue (DMMB) staining). SA: Staphylococcus aureus; PA: Pseudomonas aeruginosa; EC: Escherichia coli. Modified from Tot&#233; et al.31
Furthermore, this platform is ideal for performing primary screening campaigns. The most cost- and time-effective strategy is to apply resazurin and crystal violet-based assays as a first-tier strategy and to move on to the WGA-matrix assay in second-tier (follow-up) studies. This is due to the fact that the WGA probe is rather expensive and that this assay consists of several steps, which makes it more laborious and time-consuming.
A relevant feature of this platform is the possibility to evaluate the long-term effects of the identified antimicrobials. Long-term chemotherapeutic effects can only be achieved if the biofilm matrix is disassembled. It has been shown that the risk of resuscitation of the biofilm infection is much higher if the matrix is left behind. With this platform, it is possible to first assess if the overall biofilm biomass is affected using crystal violet staining. A more detailed assessment of the biofilm matrix polysaccharides, using the WGA assay, can follow. Of note, the likelihood of identifying a single molecule that can both disassemble the matrix and exert antibacterial (biocidal) effects may be regarded as low. Indeed, thus far, the only compounds of this type that we have identified are antimicrobial peptides37. To address this, follow-ups are performed in this platform with either resazurin- or crystal violet-active hits, since this strategy permits the identification of compounds with separate biocidal or matrix-degrading activity. Such leads can be potentially interesting for multi-component strategies. A combination of matrix-degrading molecules with biocidal or antibiotic-type compounds is expected to provide more efficient biofilm eradication.
In summary, the platform presented here provides a good basis for anti-biofilm screening using viability and biomass measurements together with matrix quantification and visualization. Both the resazurin and crystal violet staining assays can also be used for other kinds of biofilm- forming microorganisms. However, these assays require separate optimization for both growing and staining conditions. The applicability of the WGA staining assay is limited to those bacteria in which the poly-N-acetyl-glucosamine is a main component of the biofilm matrix. Other matrix quantification assays (such as the one based on dimethyl-methylene blue staining) can be applied in other cases. Altogether, the assays in this platform are fairly easy to perform, and all reagents are easily available as well as generally affordable. This platform is suitable for low- to medium-throughput anti-biofilm screening without the need for expensive equipment investments.

Bacteria can switch between two very different lifestyles, planktonic and sessile, of which a biofilm is the most common example. In biofilms, bacteria form structured communities embedded in a self-produced matrix1. This self-produced matrix is a barrier between the bacteria and their external environment, and it protects the microbial cells, keeping them in close proximity. The composition of the biofilm matrix varies between and even within species, but it mostly consists of a tight network of lipopolysaccharides, extracellular DNA, and proteins. The matrix serves as a physical barrier inhibiting the entrance of harmful agents, but it also protects the biofilm from dehydration and prevents nutrients from escaping the cell2.
Biofilms are regarded as one of the most challenging topics of modern biomedicine, and they are purportedly responsible for over 80% of antibiotic-tolerant infections3. They display an inherently high tolerance against external threats: humidity, osmotic pressure, mechanical stress4, heat, UV radiation5, disinfectants, antimicrobial agents, and the host immune system1. For example, the required antimicrobial agent concentration required to kill a biofilm has been shown to be up to 1,000 times higher in comparison to that required to kill planktonic bacteria. The explanation for this higher tolerance seems to be multifactorial. The matrix provides a physical barrier that decreases the penetration of antibiotics into the biofilm. Also, cells within the biofilms are phenotypically diverse; they transition between different metabolic states due to an existing gradient of oxygen, nutrients, and metabolites between the inner and outer parts of the biofilm6. Hence, in some biofilm regions, such as the core, bacteria are deprived of oxygen and nutrients and live in a metabolically less active or a completely dormant state7. The completely dormant cells are referred to as persister cells, and they are not susceptible to conventional antimicrobial treatments8. Thus, it is likely that biofilm resilience arises from a combination of the presently suggested and other, yet unknown, mechanisms. Staphylococcus spp. are still among the most problematic gram-positive bacteria, causing severe, often biofilm-related, infections4. It is suggested that up to 99% of all bacteria are associated within biofilms, making it the predominant bacterial lifestyle3. However, all of the currently existing antibiotics have been developed against single-cell (planktonic) bacteria. So far, a very limited repertoire of molecules exists that can selectively act on mature biofilms. This situation has driven a progressive paradigm shift in drug discovery, in which searching for anti-biofilms has been urged to occupy a more prominent place.
From a methodological perspective, additional challenges exist, as only a limited number of biofilm methods have been developed by standard-setting organizations, especially those applicable to the high-throughput screening of chemical libraries. All standardized assays (with only one exception) are based upon biofilm reactors, and these methods require large working volumes and large amounts of compounds to be tested, which are usually unavailable during the early investigational stage9-12. The only existing standardized screening-applicable assay is the so-called Calgary Biofilm Device, from which the commercially available minimum biofilm eliminating concentration (MBEC) system was developed13-15. However, the limitation of this assay is that the biofilms are grown on pegs, and not all bacterial species or even strains within the same species are able to form biofilms on this device. Moreover, methods that can be particularly applied to the exploration of natural compounds are needed. Natural products have been the major source for innovation in antimicrobial drug discovery over the past century16. They can provide novel anti-biofilm compounds with unique mechanisms of action that can also be effective against persister cells. Thus, the exploration of natural and naturally inspired libraries has high chances of producing promising and unique anti-biofilm leads.
Here, we present the experimental details of a platform of assays that was developed for the chemical screening of anti-biofilm compounds using three assays to measure the effects on the viability, total biomass, and matrix of Staphylococcus aureus biofilms. The first assay measures biofilm viability, and it is based on resazurin staining. Resazurin is a redox stain that is blue and non-fluorescent in its oxidized state and turns into pink, highly fluorescent resorufin when reduced by the metabolic activity of the bacteria. It is a very simple and fast method suitable for primary screenings17-20. The second assay, based on crystal violet staining, measures total biofilm mass. Crystal violet is a widely used stain for studying bacteria and bacteria in biofilms19,21-23. The assay is based on inexpensive reagents and has a simple absorbance endpoint reading. Finally, the third assay targets the extracellular polymeric substance (EPS)-matrix of the biofilm via wheat germ agglutinin (WGA), which binds specifically to poly-N-acetyl-glucosamine residues (PNAG) present in the matrix of staphylococcal biofilms24. The WGA is conjugated with a fluorophore that can be detected using fluorescence intensity readers25. We present here the rationale and details of the platform we developed, including examples of applications.

<table><tbody><tr><td>Resazurin</td><td>Sigma Aldrich</td><td>R7017</td><td></td></tr><tr><td>Crystal violet</td><td>Sigma Aldrich</td><td>HT90132&amp;nbsp;</td><td></td></tr><tr><td>Wheat Germ Agglutinin, Alexa Fluor 488 Conjugate</td><td>Thermo Fisher Scientific</td><td>W11261</td><td></td></tr><tr><td>LIVE/DEAD &lt;em&gt;BacLight&lt;/em&gt;&amp;nbsp;</td><td>Molecular Probes</td><td>L7012</td><td>SYTO 9 for staining viable cells green and propidium iodide for staining dead cells red</td></tr><tr><td>Phosphate Buffered Saline</td><td></td><td></td><td></td></tr><tr><td>Tryptone soy agar</td><td>Lab M, Neogen</td><td>LAB011</td><td></td></tr><tr><td>Tryptine soy broth</td><td>Lab M, Neogen</td><td>LAB004</td><td></td></tr><tr><td>F96 Well Plate Polystyrene Sterile Clear Flat bottom</td><td>Thermo Fisher Scientific</td><td>161093</td><td></td></tr><tr><td>BRAND caps, strips of 8</td><td>Sigma Aldrich</td><td>BR781413-300EA</td><td></td></tr><tr><td>Branson CPX series ultrasonic bath</td><td>Sigma Aldrich</td><td>Z769428-1EA</td><td></td></tr><tr><td>Multipipette</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Multidrop dispenser</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Biomek 3000</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Varioskan Flash Multiplate reader</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>&lt;em&gt;Staphylococcus aureus&lt;/em&gt;</td><td>ATCC&amp;nbsp;</td><td>25923</td><td></td></tr></tbody></table>

a platform of anti-biofilm assays suited to the exploration of natural compound libraries

Biofilms are regarded as one of the most challenging topics of modern biomedicine, and they are potentially responsible for over 80% of antibiotic-tolerant infections. Biofilms have displayed an exceptionally high tolerance for chemotherapy, which is thought to be multifactorial. For instance, the matrix provides a physical barrier that decreases the penetration of antibiotics into the biofilm. Also, cells within the biofilms are phenotypically diverse. Likely, biofilm resilience arises from a combination of these and other, yet unknown, mechanisms. All of the currently existing antibiotics have been developed against single-cells (planktonic) bacteria. Therefore, so far, a very limited repertoire of molecules exists that can selectively act on mature biofilms. This situation has driven a progressive paradigm shift in drug discovery, in which searching for anti-biofilms has been urged to occupy a more prominent place. An additional challenge is that there are a very limited number of standardized methods for biofilm research, especially those that can be used for large-throughput screening of chemical libraries. Here, an experimental anti-biofilm platform for chemical screening is presented. It uses three assays to measure biofilm viability (with resazurin staining), total biomass (with crystal violet staining), and biofilm matrix (using a wheat germ agglutinin, WGA-fluorescence-based staining of the poly-N-acetyl-glucosamine, PNAG, fraction). All the assays were developed using Staphylococcus aureus as the model bacteria. Examples of how the platform can be used for primary screening as well as for functional characterization of identified anti-biofilm hits are presented. This experimental sequence further allows for the classification of the hits based upon the measured end-points. It also provides information on their mode of action, especially on long-term versus short-term chemotherapeutic effects. Thus, it is very advantageous for the quick identification of high-quality hit compounds that can serve as starting points for various biomedical applications.

In the proposed platform, the effects on the viability, biomass, and biofilm matrix are quantified. In the working sequence (Figure 1), one sample plate is stained with resazurin and subsequently with crystal violet to simultaneously evaluate the effects on bacterial biofilm viability and on total biofilm biomass. Both assays can be performed consecutively in the same plate because it was demonstrated earlier that a first staining with resazurin had no statistically significant impact on the crystal violet staining result (p = 0.4149 for comparison of maximal signal absorbance units between crystal violet plates stained separately or after resazurin-staining, n = 20)26. The effect on planktonic bacteria can be simultaneously evaluated.
When hits are identified from either the viability or the biomass-based assay, a second plate is stained with the WGA probe to quantify the effect of the compounds on the polysaccharide component (PNAG) of the biofilm matrix. This workflow can be applied to the screening of chemical libraries, as well as to functional follow-up studies for potency measurements of hit compounds, as exemplified below.
<img alt="Figure 1" src="/files/ftp_upload/54829/54829fig1.jpg" /> 
Figure 1: Workflow of the three-assay platform. Schematic representation of the workflow combining the three assays on biofilm samples. The platform includes staining for an effect on viability, biomass, and EPS layer; imaging using fluorescence microscopy; and evaluating the effect on the planktonic phase. &#34;A&#34; and &#34;NA&#34; stand for &#34;Active&#34; and &#34;Not Active,&#34; respectively. Modified from Skogman et al.26 <a href="http://cloudflare.jove.com/files/ftp_upload/54829/54829fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Performance of the Three-assay Platform for Screening Purposes
Here, two screening runs are shown as representative results of the performance of the platform. In the first campaign (Figure 2A), a small library of cinchona alkaloid derivatives was screened for anti-biofilm activity against S. aureus biofilms27, while in the second example (Figure 2B), a natural and naturally inspired library of abietane-type diterpenoids and derivatives was explored28,29. The active hits were determined using the calculated hit limits (thresholds; Equation 6, Table 1). In each screening study, the results of the first two assays correlated very well; the hits were all able to reduce the viability and the biofilm biomass, while the inactive compounds showed no effect on either assay. All the identified hits were then tested on the third (WGA) assay, but they had no matrix-disassembly (or matrix-degrading) effects (results not shown).
<img alt="Figure 2" src="/files/ftp_upload/54829/54829fig2.jpg" /> 
Figure 2: Representative results of screening campaigns using the platform with S. aureus biofilms. Two examples of proof-of-concept validation screens performed using the assay platform. The results are presented as a percent of the untreated biofilm control, the hit compounds are indicated in red, and the lines represent the calculated hit limits. (A) The screening of a cinchona-alkaloid derivative library identified one active compound. (B) The screening of a library of abietane-type diterpenoids and derivatives identified five active compounds. <a href="http://cloudflare.jove.com/files/ftp_upload/54829/54829fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Performance of the Three-assay Platform for Follow-up Studies
Representative results can be seen in Figure 3, where two known antibiotics were tested at a very broad range of concentrations (0.5 nM-5 mM) against S. aureus biofilms. Minimum inhibitory concentration (MIC) values of reference compounds against planktonic bacteria (either from the literature or performed in the lab) serve as guidelines for choosing the concentrations to test against biofilms of the same species. In addition, knowledge of the concentration values in which no cytotoxicity is detected in mammalian cells can also help in the selection of the concentration to test for anti-biofilm follow-ups. Ideally, if both antimicrobial and cytotoxicity data for a certain compound are available, a parameter known as the &#34;Biocompatibility Index&#34; (BI) can be calculated, as defined by M&#252;ller and Kramer30. Here, MIC values against planktonic S. aureus for penicillin G and ciprofloxacin were determined to be 0.04 &#181;M and 6 &#181;M, respectively (results not shown). Thus, the concentration interval ranged from approximately 105 x MIC to 10-2 x MIC and 103 x MIC to 10-4 x MIC for penicillin and ciprofloxacin, respectively. Both antibiotics could significantly decrease the viability, biomass, and biofilm matrix PNAG content when they were exposed to single-cell bacteria prior to the initiation of the biofilm formation process (Figure 3a). However, despite the very high concentration of antibiotics tested on preformed (18 hr) biofilms, the viability and the biomass were only reduced to approximately 50% of those of untreated biofilms (Figure 3b). The most prominent result here was the increase of the biofilm matrix (to over 200%) when preformed biofilms were treated with high concentrations of penicillin G. No changes in the content of the biofilm matrix were detected when preformed biofilms were treated with ciprofloxacin.
<img alt="Figure 3" src="/files/ftp_upload/54829/54829fig3.jpg" /> 
Figure 3: Example of a follow-up study of anti-biofilm effects using two model antibiotics against S. aureus biofilms. The effects of penicillin G and ciprofloxacin are presented here: (A) prior to biofilm formation (pre-exposure) and (B) post-biofilm formation (post-exposure). For figure clarity, only the lowest and the highest concentrations tested are shown. The standard deviations do not exceed 20%. *** equals p &#60; 0.01. Modified from Skogman et al.26 <a href="http://cloudflare.jove.com/files/ftp_upload/54829/54829fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Fluorescence Microscopy-based Imaging
The result that penicillin G at 400 &#181;M (as well as higher concentrations, as shown in Figure 3b) kills about 50% of the preformed S. aureus biofilm bacterial population was confirmed with another viability staining assay. The average of the green-to-red (G/R) fluorescence ratios for the untreated biofilm wells was 2.75, indicating a predominance of live (green-stained) cells, over dead (red-stained) cells. However, in penicillin-treated cells the G/R ratio decreased to 1.54, corresponding to 56% of the control wells. The cells remaining alive after penicillin treatment produced a significantly higher amount of EPS, as judged from the detected increase in green (WGA) fluorescence when compared with untreated biofilms (Figure 4b). We recommend, whenever possible, to perform these imaging-based experiments to further confirm the results of the platform, especially during the stage of characterization of hit candidates.
<img alt="Figure 4" src="/files/ftp_upload/54829/54829fig4.jpg" /> 
Figure 4: Fluorescence microscopy. The effect of penicillin addition (400 &#181;M) on viability and EPS production are shown here. The top images (A) correspond to untreated biofilms and penicillin-treated biofilms, where viable cells are stained green and dead cells are stained red. The inserted graph illustrates the calculations of G/R fluorescence ratios. The bottom images (B) correspond to untreated biofilms and penicillin-treated cells stained with the WGA probe. The inserted graph presents the quantification of the WGA signals for treated and untreated biofilm samples, and error bars represent the standard deviations. Modified from Skogman et al.26 <a href="http://cloudflare.jove.com/files/ftp_upload/54829/54829fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Data Processing and Statistical Analysis
Statistical parameters are calculated to characterize the quality of the assays and to follow their performance during screening runs. The equations of the most important parameters used, as well as the obtained and target values, are listed in Table 1. In all equations, SDmin, &#181;min, SDmax, and &#181;max represent the standard deviations and means of the minimal (min) and maximal (max) signals, respectively. In the results shown here, paired comparisons of the original values were done with an unpaired t-test with Welch&#39;s correction, where p &#60; 0.05 was considered to be statistically significant.
<img alt="Figure 1" src="/files/ftp_upload/54829/54829table1.jpg" /> 
Table 1: Statistical parameters used to evaluate the performance of the assays. In all equations, SDmin, &#181;min, SDmax, and &#181;max represent the standard deviations and means of the minimal (min) and maximal (max) signals, respectively. The staining methods, resazurin, crystal violet, and wheat germ agglutinin, are abbreviated RES, CrV, and WGA, respectively. RFU: relative fluorescence units; RAU: relative absorbance units. <a href="http://cloudflare.jove.com/files/ftp_upload/54829/Table1.xlsx">Please click here to download this file.</a>

Watch this Scientific Journal Video about A Platform of Anti-biofilm Assays Suited to the Exploration of Natural Compound Libraries at JoVE.com

A Platform of Anti-biofilm Assays Suited to the Exploration of Natural Compound Libraries

Biofilm infections show high tolerance towards chemotherapy. No single assay captures the complexity of biofilms. Instead, complementary assays are needed. We present a screening platform (developed for S. aureus) that combines assays for viability, biomass, and biofilm matrix. It allows anti-biofilm drug discovery, including the assessment of long-term chemotherapeutic effects.

Biofilms are regarded as one of the most challenging topics of modern biomedicine, and they are potentially responsible for over 80% of ...

a-platform-anti-biofilm-assays-suited-to-exploration-natural-compound

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

17.5K Views.  University of Helsinki. Biofilm infections show high tolerance towards chemotherapy. No single assay captures the complexity of biofilms. Instead, complementary assays are needed. We present a screening platform (developed for S. aureus) that combines assays for viability, biomass, and biofilm matrix. It allows anti-biofilm drug discovery, including the assessment of long-term chemotherapeutic effects. 

Video: A Platform of Anti-biofilm Assays Suited to the Exploration of Natural Compound Libraries

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 ...

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 ...

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 ...

Targeting Biofilm Associated Staphylococcus aureus Using Resazurin Based Drug-susceptibility Assay

Most pathogenic bacteria are able to form biofilms during infection, but due to the difficulty of manipulating and assessing biofilms, the vast majority of laboratory work is conducted with planktonic cells. Here, we describe a peg plate biofilm assay as performed with Staphylococcus aureus. Bacterial biofilms are grown on pegs attached to a 96-well microtiter plate lid, washed through gentle submersion in buffer, and placed in a drug challenge plate. After subsequent incubation they are again washed and moved to a final recovery plate, in which the fluorescent dye resazurin serves as a viability indicator. This assay offers greatly increased ease-of-use, reliability, and reproducibility, as well as a wealth of data when conducted as a kinetic read. Moreover, this assay can be adapted to a medium-throughput drug screening approach by which an endpoint fluorescent readout is taken instead, offering a path for drug discovery efforts.

Targeting Biofilm Associated Staphylococcus aureus Using Resazurin Based Drug-susceptibility Assay

Most bacterial infections produce a biofilm. By virtue of their environment, biofilm associated bacteria are often phenotypically drug resistant. Novel antibacterial molecules that kill bacteria in biofilms are thus a high priority. We establish an assay to quickly screen for antimicrobial compounds that are effective at eradicating biofilms.

Most pathogenic bacteria are able to form biofilms during infection, but due to the difficulty of manipulating and assessing biofilms, the vast majority ...

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 ...

Adaptable CF Model for Antimicrobial Testing and Microbial Dynamics Analysis

Development of a Polymicrobial Colony Biofilm Model to Test Antimicrobials in Cystic Fibrosis

A range of bacteria biofilm models exist for the testing of antibiotics. However, many of these are limited to a single experimental output, such as colony-forming units or metabolic activity. Furthermore, many biofilm models do not reflect the biological and physiochemical properties of the human host environment. This is an important issue in many conditions, but most noticeably in cystic fibrosis (CF). A large proportion of people with CF suffer from both chronic and intermittent infections, and in vitro, antibiotic susceptibility testing poorly correlates with patient treatment outcomes. Some biofilm models incorporate CF lung-relevant media, including synthetic sputum mimics, but do not consider the polymicrobial nature of the environment, which alters biofilm architecture, physiology, and the way microbes respond to treatment. The solid-air interface colony biofilm model described here is highly adaptable and incorporates both CF-relevant media and a polymicrobial context. This model can also be used for mid-throughput screening of antimicrobials and to study their effect on polymicrobial dynamics. Output measurements from the model can be colony-forming units, metabolic activity, and confocal microscopy analysis. The model can easily be adapted to different microorganisms, media, temperatures, and variable oxygen conditions and can be used to test a wide range of chemical, biological, and physical treatments.

This methodology allows for the establishment of a polymicrobial biofilm model in cystic fibrosis for antimicrobial sensitivity testing in research and clinical laboratories. This model provides accurate and reliable results over a range of outputs.

A range of bacteria biofilm models exist for the testing of antibiotics. However, many of these are limited to a single experimental output, such as ...

Systematic Approach to Identify Novel Antimicrobial and Antibiofilm Molecules from Plants' Extracts and Fractions to Prevent Dental Caries

Natural products provide structurally different substances, with a myriad of biological activities. However, the identification and isolation of active compounds from plants are challenging because of the complex plant matrix and time-consuming isolation and identification procedures. Therefore, a stepwise approach for screening natural compounds from plants, including the isolation and identification of potentially active molecules, is presented. It includes the collection of the plant material; preparation and fractionation of crude extracts; chromatography and spectrometry (UHPLC-DAD-HRMS and NMR) approaches for analysis and compounds identification; bioassays (antimicrobial and antibiofilm activities; bacterial &#34;adhesion strength&#34; to the salivary pellicle and initial glucan matrix treated with selected treatments); and data analysis. The model is simple, reproducible, and allows high-throughput screening of multiple compounds, concentrations, and treatment steps can be consistently controlled. The data obtained provide the foundation for future studies, including formulations with the most active extracts and/or fractions, isolation of molecules, modeling molecules to specific targets in microbial cells and biofilms. For example, one target to control cariogenic biofilm is to inhibit the activity of Streptococcus mutans glucosyltransferases that synthesize the extracellular matrix&#8217; glucans. The inhibition of those enzymes prevents the biofilm build-up, decreasing its virulence.

Natural products represent promising starting points for the development of new drugs and therapeutic agents. However, due to the high chemical diversity, finding new therapeutic compounds from plants is a challenging and time-consuming task. We describe a simplified approach to identify antimicrobial and antibiofilm molecules from plant extracts and fractions.

Natural products provide structurally different substances, with a myriad of biological activities. However, the identification and isolation of active ...

Biology

A Peg Plate Biofilm Assay for Screening Antibacterial Agents

Source: Dalecki, A. G., et al. <a href="https://app.jove.com/v/53925">Targeting Biofilm Associated Staphylococcus aureus Using Resazurin Based Drug-susceptibility Assay.</a> J. Vis. Exp. (2016)
This video demonstrates an assay for screening antimicrobial compounds that efficiently eliminate biofilms using a peg-plate. The methodology monitors the biofilm&#39;s metabolic state after treatment with antimicrobial compounds, facilitating the determination of the minimum biofilm eradication concentration.

1. Biofilm Initiation

	Grow a culture of biofilm-producing organisms in a nutrient-rich medium. Inoculate&#160;Staphylococcus aureus&#160;strain Newman ...

JoVE Encyclopedia of Experiments

Immunology

Immunodiagnostics

Specialized Assays

A Solid Phase Adherence Assay to Assess Biofilm Formation

Source: Nickerson, K. P. et al., <a href="https://app.jove.com/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 ...

Detection of Biofilm Disassembly Through a Dispersion Assay

Source: Nickerson, K. P. et. al., <a href="https://app.jove.com/v/57322">Bile Salt-induced Biofilm Formation in Enteric Pathogens: Techniques for Identification and Quantification.</a>&#160;J. Vis. Exp.&#160;(2018)
The video showcases a biofilm dispersion assay using a multi-well plate. Buffer and glucose treatment disassemble the biofilm, releasing bacteria, while bile salts induce stress, retaining bacteria in the protective extracellular polymeric substance. Higher dispersion with buffer and glucose confirms successful biofilm disassembly, estimated by colony forming units measurement.

A Platform of Anti-biofilm Assays Suited to the Exploration of Natural Compound Libraries

Summary

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Chapters in this video

Microtiter Dish Biofilm Formation Assay

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

Concurrent Quantification of Cellular and Extracellular Components of Biofilms

Targeting Biofilm Associated Staphylococcus aureus Using Resazurin Based Drug-susceptibility Assay

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

Development of a Polymicrobial Colony Biofilm Model to Test Antimicrobials in Cystic Fibrosis

Systematic Approach to Identify Novel Antimicrobial and Antibiofilm Molecules from Plants' Extracts and Fractions to Prevent Dental Caries

A Peg Plate Biofilm Assay for Screening Antibacterial Agents

A Solid Phase Adherence Assay to Assess Biofilm Formation

Detection of Biofilm Disassembly Through a Dispersion Assay