We wish to acknowledge Danielle Orr Goveia, Blaine Fritz, Jennifer Summers and Fei San Lee for their contributions to this paper.

1. Vortexing and sonication
<ol>
	<li>Grow a mature P. aeruginosa ATCC 15442 biofilm grown according to ASTM Standard E25622.</li>
	<li>At the end of the 48 h growth period, prepare to treat the biofilm and sample coupons according to ASTM Standard E28718</li>
	<li>Aseptically insert autoclaved splash guards into sterile 50 mL conical tubes using flame-sterilized forceps. Repeat for all tubes that will receive treatment. Tubes for control coupons do not need a splash guard.</li>
	<li>Aseptically remove a randomly selected rod from the CDC Biofilm Reactor. Rinse coupons to remove loosely attached cells by gently dipping the rod into 30 mL of sterile buffered water.</li>
	<li>Hold the rod parallel to the bench top, over an empty, sterile 50 mL conical tube and using a flame-sterilized Allen wrench, loosen set screw to drop a biofilm coated coupon into tube. Repeat for the desired number of coupons. Remove splash guards and place in a separate container for sterilization.</li>
	<li>Using a 5 mL serological pipette, slowly pipette 4 mL of the treatment or control into the tubes so that the liquid flows down the inside of the wall of the tube.</li>
	<li>Gently tap the bottom of the tube so that any air bubbles under the coupon are displaced. Allow 30 - 60 s between each addition.</li>
	<li>At the end of the specified contact time, pipette 36 mL of neutralizer into the tubes in the same order that the treatment (or control) was applied. 
	NOTE: The final volume of combined treatment and neutralizer is important for accurately determining biofilm log density.</li>
	<li>Vortex each tube on the highest setting for 30 &#177; 5 s. Ensure that a complete vortex is achieved. 
	NOTE: Caution should be exercised when vortexing heavy coupons such as stainless steel in glass vials where breakage could occur.</li>
	<li>Determine the optimal number of tubes per bath and placement within the test tube rack prior to processing actual samples. If processing multiple samples, confirm that the water temperature in the sonicating bath is 21 &#177; 2 oC.</li>
	<li>Place tubes in tube rack suspended in degassed sonicator such that water level in bath is equal to liquid level in tubes. Sonicate at 45 kHz, 100% power and Normal function for 30 &#177; 5 s. Repeat vortex and sonication cycles and then end with a final vortex (5 cycles total). 
	NOTE: These tubes with the harvested and disaggregated biofilm are the100&#160;or 0 dilution.</li>
	<li>Serially dilute sample in buffered water. Plate on R2A agar using appropriate plating method. Incubate at 36 &#177; 2 oC for 24 h. Count colonies as appropriate to plating method used and record data.</li>
</ol>
2. Scraping and homogenization
<ol>
	<li>Grow a mature P. aeruginosa ATCC 15442 biofilm according to the ASTM Standard E264713.</li>
	<li>Set up the sampling station to include sampling board, 95% ethanol in a beaker, alcohol burner, hemostats, coupon removal tool, beakers with sterile dilution water and dilution tubes for rinsing the coupons.</li>
	<li>Turn off the pump. Remove a channel cover and use a sterile coupon removal tool and hemostats to remove the coupon, being careful not to disturb the biofilm.</li>
	<li>Rinse the coupon by gently immersing, with a fluid motion, in 45 mL of sterile dilution water (contained in a 50 mL centrifuge tube). Immediately reverse the motion to remove the coupon.</li>
	<li>Place the coupon into a beaker containing 45 mL of sterile dilution water. Scrape the biofilm-covered coupon surface in a downward direction for approximately 15 s, using a sterile spatula or scraper. Rinse the spatula or scraper by stirring it in the beaker. Repeat the scraping and rinsing process 3-4 times, ensuring full coverage of the coupon surface.</li>
	<li>Rinse the coupon by holding it at a 60&#176; angle over the sterile beaker and pipetting 1 mL of sterile dilution water over the surface of the coupon. Repeat for a total of 5 rinses. The final volume in the beaker is 50 mL. 
	NOTE: The final volume of combined treatment and neutralizer is important for accurately determining biofilm log density.</li>
	<li>Replace each channel cover as the coupons are removed.</li>
	<li>Working in the biosafety cabinet, homogenize the scraped biofilm sample. Attach a sterile homogenizer probe to the homogenizer, place the probe tip in the liquid, turn the homogenizer on and ramp up to 20,500 rpm.</li>
	<li>Homogenize the sample for 30 s. Turn down the RPMs and switch the homogenizer off.</li>
	<li>Sanitize the probe between biofilm samples by homogenizing a 9 mL of sterile dilution blank at 20,500 rpm for 30 s as described above. Homogenize a 9 mL tube of 70% ethanol for 30 s, detach the probe and let stand in the ethanol tube for 1 min. Homogenize two additional dilution blanks. 
	NOTE: A disposable homogenizer probe may be used for each sample.</li>
	<li>Serially dilute the samples in buffered water. Plate on R2A agar using the appropriate plating method. Incubate plates at 36 &#177; 2oC for 24 h, count colonies as appropriate to plating method used and record data.</li>
</ol>
3. Scraping, vortexing and sonication
<ol>
	<li>Grow a mature Escherichia coli ATCC 53498 biofilm in silicone catheter tubing10.</li>
	<li>Prepare sampling materials: rinse tubes, sterile centrifuge tube, empty sterile Petri dish, flame sterilized stainless steel hemostat and scissors, timer, and ruler.</li>
	<li>With the pump paused, use 70% ethanol to clean the outside of the tubing. Measure 2 cm from the end, avoiding the area attached to the connector, and mark the tubing to determine cutting locations.</li>
	<li>With flame sterilized scissors, cut the tubing on the 2 cm mark and place the segment in empty sterile Petri dish. Wipe the tubing with 70% ethanol and reconnect the distal end to waste tubing.</li>
	<li>Rinse tubing segment to remove planktonic cells. With flame sterilized forceps, gently immerse tubing segment into 20 mL of sterile dilution water and then immediately remove. Place the segment into 10 mL of neutralizer.</li>
	<li>With flame sterilized forceps, hold the tubing segment and scrape with sterile wooden applicator stick until all inner areas of the tubing have been scraped. Occasionally rinse the stick in the 10 mL of neutralizer and place the segment back into the sample tube. The scraped tubing segment is the 100 or 0 dilution. 
	NOTE: The final volume of combined treatment and neutralizer is important for accurately determining biofilm log density.</li>
	<li>Vortex each tube on the highest setting for 30 &#177; 5 s. Place the tube in tube rack suspended in sonicator such that water level in bath is equal to liquid level in tubes. Sonicate at 45 kHz, 100% power and Normal function for 30 &#177; 5 seconds. Repeat vortex and sonication cycles then end with a final vortex. 
	NOTE: This tube with the harvested and disaggregated biofilm is the 100 dilution.</li>
	<li>Serially dilute the samples in buffered water. Plate on Tryptic Soy Agar using the appropriate plating method.</li>
	<li>Incubate plates at 36 &#177; 2 oC for 24 h. Count colonies as appropriate to plating method used, record data and calculate the arithmetic mean.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biofilms:+microbial+life+on+surfaces.">Biofilms: microbial life on surfaces.</a> Emerging Infectious Diseases. 8 (9), 881-890 (2002).</li><li>ASTM International. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ASTM+Standard+E2562,+2017,+Standard+Test+Method+for+Quantification+of+Pseudomonas+aeruginosa+Biofilm+Grown+with+High+Shear+and+Continuous+Flow+using+CDC+Biofilm+Reactor.">ASTM Standard E2562, 2017, Standard Test Method for Quantification of Pseudomonas aeruginosa Biofilm Grown with High Shear and Continuous Flow using CDC Biofilm Reactor.</a> ASTM International. , (2017).</li><li>Azeredo, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Critical+review+on+biofilm+methods.">Critical review on biofilm methods.</a> Critical Reviews In Microbiology. 43 (3), 313-351 (2017).</li><li>Gomes, I. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Standardized+reactors+for+the+study+of+medical+biofilms:+a+review+of+the+principles+and+latest+modifications.">Standardized reactors for the study of medical biofilms: a review of the principles and latest modifications.</a> Critical Reviews in Biotechnology. 38 (5), 657-670 (2018).</li><li>Goeres, D. M., et al., Simoes, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+and+fabrication+of+biofilm+reactors.">Design and fabrication of biofilm reactors.</a> Recent Trends in Biofilm Sciences and Technology. , 71-88 (2020).</li><li>Goeres, D. 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M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Validation+of+a+Biofilm+Efficacy+Test:+The+Single+Tube+Method.">Validation of a Biofilm Efficacy Test: The Single Tube Method.</a> Journal of Microbiological Methods. , (2019).</li><li>The development and validation of a standard in vitro method to evaluate the efficacy of surface modified urinary catheters. Theses and Dissertations at Montana State University Available from: <a target="_blank" href="https://scholarworks.montana.edu/xmlui/handle/1/15149">https://scholarworks.montana.edu/xmlui/handle/1/15149</a> (2019)</li><li>Hamilton, M. A., Buckingham-Meyer, K., Goeres, D. 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W., Tuohy, M. J., Fujishiro, T., Procop, G. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brief+Ultrasonication+Improves+Detection+of+Biofilm-formative+Bacteria+Around+a+Metal+Implant.">Brief Ultrasonication Improves Detection of Biofilm-formative Bacteria Around a Metal Implant.</a> Clinical Orthopaedics and Related Research. 457, 210-213 (2007).</li><li>Lourenco, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Minimum+information+about+a+biofilm+experiment+(MIABiE),+standards+for+reporting+experiments+and+data+on+sessile+microbial+communities+living+at+interfaces.">Minimum information about a biofilm experiment (MIABiE), standards for reporting experiments and data on sessile microbial communities living at interfaces.</a> Pathogens and Disease. , 1-7 (2014).</li><li>Nascentes, C. C., Korn, M., Sousa, C. S., Arruda, M. A. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+Ultrasonic+Baths+for+Analytical+Applications:+A+New+Approach+for+Optimisation+Conditions.">Use of Ultrasonic Baths for Analytical Applications: A New Approach for Optimisation Conditions.</a> Journal of Brazilian Chemical Society. 12 (1), 57-63 (2001).</li><li>Elma GmbH &amp; Co. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=KG+TI-H+Ultrasonic+Cleaning+Units:+Operating+Instructions.">KG TI-H Ultrasonic Cleaning Units: Operating Instructions.</a> Elma GmbH &amp; Co. , (2009).</li><li>Stamper, D. M., Holm, E. R., Brizzolara, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exposure+times+and+energy+densities+for+ultrasonic+disinfection+of+Escherichia+coli,+Pseudomonas+aeruginosa,+Enterococcus+avium+and+sewage.">Exposure times and energy densities for ultrasonic disinfection of Escherichia coli, Pseudomonas aeruginosa, Enterococcus avium and sewage.</a> Journal of Environmental Engineering and Science. 7 (2), 139-146 (2008).</li><li>Suslick, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sonochemistry.">Sonochemistry.</a> Science. 247 (4949), (1990).</li></ol>

The authors have no disclosures.

Minimum Information for Harvesting and Disaggregation Methods 
To create reproducible biofilm data across the scientific community, it is imperative that authors include as much detail as possible regarding each of the growth, treatment, sampling and analysis steps of a biofilm method. The standardization of biofilm methods has aided in this endeavor as it allows the researcher to reference a specific method and any relevant modifications. However, many papers include only a sentence or two to describe biofilm harvesting and disaggregation. For better reproducibility, we recommend that minimum information for biofilm harvesting be included in publications. This builds on the Minimum Information About a Biofilm Experiment (MIABiE) initiative presented by Lourenco et al.17. In the case of using sonication for biofilm harvesting and disaggregation parameters the information should include: position of tubes within the bath (manufacturers&#39; recommendations to avoid damaging the transducers), number of tubes sonicated at the same time, tube material, volume and type of liquid in the tube, presence of surfactant, position of liquid in tubes relative to liquid level of ultrasonic bath, device used to hold the tubes in the bath (glass beakers vs. test tube rack), degassing of water bath prior to sonication of samples (if degassing is not a manufacturer option, simple operation of the bath before inserting the samples will help remove some dissolved gasses from the bath liquid), temperature of water bath (temperatures can rise after long periods of sonication), frequency (25 kHz or 45 kHz, for example), bath function (sweep or normal, for example) and power range delivered to transducers (10 - 100%, for example). These settings should be optimized for the biofilm being studied, the system used to grow the biofilm and the specific make/model of ultrasonic bath18.
Sonicator settings may be changed to optimize desired harvesting effects. Degassing removes dissolved air within the bath liquid thereby enhancing cleaning power. Frequency can be adjusted to a low or high setting. A low setting such as 25 kHz, for example, would aid in harvesting tenacious samples while a higher setting of 45 kHz would be more appropriate for sensitive samples. A bath function of sweep or normal allows for distribution of cavitation. The sweep function creates a continuous shifting of the sound pressure maxima. The normal function allows the transducers to operate in double half wave mode which may result in dead zones, thereby making the sonication less efficacious. Power ranges can be altered from 10 - 100% of the power delivered to the transducers19. It is known that sonication can detrimentally effect bacterial viability. A 2008 study by Stamper et al.20 exposed bacterial cultures to increasing ultrasonice energy over time to create bacterial kill curves. We recommend that users confirm that a particular combination of sonication settings does not cause a decrease in viable bacteria20.
There is no one perfect method to harvest and disaggregate biofilm, but particular methods do work better for some surfaces/microbe combinations than others. We advocate for the reader to determine which parameters are important for their particular biofilm scenario. Key parameters of importance for harvesting and disaggregation are included in Supplementary File 1.
For the sonication studies done to assess harvesting and disaggregation bias, we found that sonication is convenient, efficient and able to be standardized for harvesting and disaggregating biofilm from surfaces. Placement of coupons in vials minimizes technician to technician variability that would be encountered in methods where coupons are physically scraped by laboratory personnel, for example. Although it seems simple enough to place vials in a sonicating water bath, there are many parameters that need to be considered to achieve optimal harvesting of biofilm.
Two types of sonication equipment are available, ultrasonic baths and ultrasonic probes. This paper focuses primarily on ultrasonic baths where ultrasonic energy is generated in the range of high (20 - 45 kHz) to normal (40 - 60 Hz) frequency.
Three main processes are at play when using an ultrasonic device to clean a surface. Electrical energy is converted to acoustic energy when a high frequency current is sent to a piezoelectric or magnetostrictive transducer that oscillates in response to the current. The oscillation generates compression (rarefaction) waves in the liquid. Cavitation bubbles form due to negative pressure during rarefaction. The bubbles grow until they reach an unstable size and collapse, creating a water jet that cleans surfaces21.
Three harvesting and disaggregation approaches are demonstrated in this article: sonication and vortexing methods for harvesting and disaggregating biofilm when grown on polycarbonate coupons in the CDC Biofilm Reactor according to the Single Tube Method. Scraping and homogenization methods for harvesting and disaggregating biofilm are shown when grown on glass coupons using the Drip Flow Biofilm Reactor. Scraping, sonication and vortexing methods for harvesting and disaggregating biofilm are shown when grown in silicone tubing.
There is no perfect method to harvest and disaggregate the biofilm, but some approaches are better for different surfaces and/or applications. What is important is to take the time to validate the method used. In this paper, we discussed the use of crystal violet and microscopy, but other choices exist depending on the sensitivity required. If research includes efficacy testing, then it is critical to confirm the validity of the approach in the presence of the antimicrobial6. All equipment is slightly different, so even if the harvesting and disaggregation method have been standardized, it is still prudent to confirm the process for the equipment used. Harvesting and disaggregation methods are specific for surface associated and biofilm bacteria. Research has demonstrated that improper choices may lead to biased test results. Nonetheless, harvesting and disaggregation are the least studied of the four steps in biofilm methods. It is generally also the unvalidated step (taken for granted as working) with the least information present in published paper making it challenging to reproduce the process in a different lab. This paper and accompanying video show three common approaches for two surface types and suggests how to validate a method for an individual lab. This information will help researchers make more informed decisions on which method to use and provides guidance on what to report to improve reproducibility.

The definition of biofilm has evolved over the last few decades and encompasses microbial association with a variety of biological and/or non-biological surfaces, inclusion of noncellular components1 that display differing growth and genetic expression2 within a matrix. Biofilm provides protection from environmental stresses such as drying and may render the action of chemical disinfectants less effective resulting in the survival of microbes. The survivors within a biofilm can potentially provide a source of pathogenic microorganisms that are a public health concern3.
Biofilm methods are comprised of four steps, growth, treatment, sampling (harvesting and disaggregation), and analysis. Growth, the first step, where the user determines the organism growth conditions, temperature, media, etc., is the most considered and reported upon in the biofilm literature4,5,6,7. The treatment step evaluates antimicrobials (e.g., disinfectants) to determine their efficacy either against a mature biofilm3,8,9 or the antimicrobial may be incorporated into the surface to determine the ability of the product to prevent or reduce biofilm growth10. The third step, sampling, includes steps to harvest the biofilm from the surface on which it was growing and to disaggregate the removed clumps3,8,11. The fourth step, analysis, may include viable cell counts, microscopy, fluorescence measurements, molecular outcomes, and/or a matrix component assessment8,9. Assessment of data provides information about the outcome of an experiment. Of the four, sampling is often the most overlooked step because it presumes that the chosen biofilm harvesting and/or disaggregation technique is 100% effective, often without verification11.
Planktonic suspensions of bacteria, often considered to be homogenous, require simple vortexing prior to analysis. Biofilms, however, are complex communities composed of microorganisms (prokaryotic and/or eukaryotic), exopolysaccharides, proteins, lipids, extracellular DNA and host cells12. Steps beyond traditional planktonic microbiological culture methods are needed in order to adequately harvest biofilm from a surface and then disaggregate it into a homogenous single cell suspension. An extensive literature review (information not included in this publication) demonstrated that the choice of the removal and disaggregation technique is dependent on a number of factors, including species present in the biofilm, surface that the biofilm is attached to (non-porous or porous), accessibility to growth surfaces (easily removable coupon or physical destruction of apparatus in which the biofilm is growing), surface geometry (area and shape), density of biofilm on growth surfaces, and available laboratory equipment.
When biofilm is harvested from a surface, the resulting cell suspension is heterogenous. If this nonuniform suspension is to be accurately enumerated, it must be disaggregated into individual cells. Viable plate counts assume that a colony forming unit originates from one bacterium. If aggregates of biofilm are placed on the growth medium, it is impossible to distinguish individual cells which could lead to inaccurate estimates. For example, during disinfectant efficacy testing, if a treatment removes biofilm very effectively from a surface compared to the control, the log reduction could appear artificially large compared to the control. On the other hand, a chemical disinfectant that fixes biofilm onto a surface compared to the control will appear to have a lower log reduction11. This type of scenario could lead to biased interpretation of experimental data.
In preparation for the&#160;publication, a review of the literature determined that common approaches to harvesting and disaggregating biofilm include scraping, swabbing, sonication, vortexing or a combination of these. Scraping is defined as physical removal of biofilm from surfaces with a sterile stick, spatula or other tool. Swabbing refers to removal of biofilm from surfaces with a cotton tipped stick or other fixed absorbent material. Sonication refers to disruption of biofilm from surfaces via ultrasonic waves distributed through water. Vortexing refers to the use of a mixer to achieve a liquid vortex of the sample inside a tube. Homogenization uses rotating blades to shear harvested biofilm clumps into a single cell suspension. In this paper, we present three harvesting and disaggregation methods for two different surface types, hard/non-porous and porous.
A list of recommended minimum information that researchers should include in the methods sections of publications is provided. We hope that inclusion of this information enables other researchers to reproduce their work. There is no perfect harvesting and disaggregation method, therefore, recommendations for how to check the technique are also provided.
Three common methods to harvest and disaggregate biofilm from common growth surfaces are demonstrated in this article. This information will enable researchers to better understand the overall precision and bias of a biofilm test method. Methods described are as follows: (1) A Pseudomonas aeruginosa biofilm grown on polycarbonate coupons (hard non-porous surface) under high fluid shear in the CDC Biofilm Reactor is harvested and disaggregated following a five step combination of vortexing and sonication to achieve biofilm harvest and disaggregation (2) A P. aeruginosa biofilm grown on borosilicate glass coupons (hard non-porous surface) in the drip flow reactor under low fluid shear is harvested and disaggregated using scraping and homogenization (3) An Escherichia coli biofilm grown in silicone tubing (porous surface) is harvested and disaggregated using scraping, followed by sonication and vortexing.

<table><tbody><tr><td>50 mL conical vials</td><td>Thermo Scientific</td><td>339652</td><td></td></tr><tr><td>100 mL glass beakers</td><td>Fisher Scientific</td><td>FB102100</td><td></td></tr><tr><td>5 mL serological pipettes</td><td>Fisher Scientific</td><td>13-678-12D</td><td>For adding treatment to vials containing coupons.</td></tr><tr><td>50 mL serological pipettes</td><td>Fisher Scientific</td><td>13-678-14C</td><td>For adding neutralizer to vials at the end of treatment contact time.</td></tr><tr><td>Applicator sticks</td><td>Puritan</td><td>807</td><td></td></tr><tr><td>Hemostats</td><td>Fisher Scientific</td><td>16-100-115</td><td></td></tr><tr><td>Metal spatula</td><td>Fisher Scientific</td><td>14-373</td><td></td></tr><tr><td>PTFE policemen</td><td>Saint-Gobain</td><td>06369-04</td><td></td></tr><tr><td>S 10 N - 10 G - ST Dispersing tool</td><td>IKA</td><td>4446700</td><td>For homogenization of biofilm samples.</td></tr><tr><td>Scissors</td><td>Fisher Scientific</td><td>08-951-20</td><td></td></tr><tr><td>Silicone Foley catheter, size 16 French</td><td>Medline Industries</td><td>DYND11502</td><td></td></tr><tr><td>Silicone tubing, size 16</td><td>Cole-Parmer</td><td>EW96400-16</td><td></td></tr><tr><td>Splash Guards</td><td>BioSurface Technologies, Inc.</td><td>CBR 2232</td><td></td></tr><tr><td>T 10 basic ULTRA-TURRAX Disperser</td><td>IKA</td><td>3737001</td><td>For homogenization of biofilm samples.</td></tr><tr><td>Tubing connectors</td><td>Cole-Parmer</td><td>EW02023-86</td><td></td></tr><tr><td>Ultrasonic Cleaner</td><td>Elma</td><td>TI-H15</td><td></td></tr><tr><td>Vortex-Genie 2</td><td>Scientific Industries</td><td>SI-0236</td><td></td></tr><tr><td>Vortex-Genie 2 Vertical 50 mL Tube Holder</td><td>Scientific Industries</td><td>SI-V506</td><td></td></tr></tbody></table>

harvesting and disaggregation: an overlooked step in biofilm methods research

Biofilm methods consist of four distinct steps: growing the biofilm in a relevant model, treating the mature biofilm, harvesting the biofilm from the surface and disaggregating the clumps, and analyzing the sample. Of the four steps, harvesting and disaggregation are the least studied but nonetheless critical when considering the potential for test bias. This article demonstrates commonly used harvesting and disaggregation techniques for biofilm grown on three different surfaces. The three biofilm harvesting and disaggregation techniques, gleaned from an extensive literature review, include vortexing and sonication, scraping and homogenization, and scraping, vortexing and sonication. Two surface types are considered: hard non-porous (polycarbonate and borosilicate glass) and porous (silicone). Additionally, we provide recommendations for the minimum information that should be included when reporting the harvesting technique followed and an accompanying method to check for bias.

Validation/Confirmation of a Harvesting Method 
Several studies that were conducted in our laboratory examined the ability of vortexing and sonication to effectively harvest biofilm grown in the biofilm reactor (ASTM E2562)2 using the Single Tube Method (ASTM E2871)8.
A P. aeruginosa ATCC 15442 biofilm was grown according to ASTM E25622 on borosilicate glass coupons. After 48 hours, four coupons were placed into vials, &#34;treated&#34; with 4 mL sterile buffered water and neutralized with 36 mL of 2x D/E Neutralizing Broth. The initial sonication setting of 45 kHz, 10% power, Sweep setting, 30 &#177; 5 s was used to harvest and disaggregate the biofilm from three of the four coupons. Upon completion of the vortex and sonication cycle, each coupon was stained with crystal violet and photographed. Figure 1 demonstrates the amount of biofilm remaining on the three coupons after vortexing and sonication as compared to the control.
To test this further, a P. aeruginosa ATCC 15442 biofilm was grown as described previously and two sonication settings were compared: 1) 45 kHz, 10% power, Sweep setting, 30 &#177; 5 seconds and 2) 45 kHz, 100% power, Normal setting, 30 &#177; 5 seconds. One coupon from each set of the three was stained with BacLight Live/Dead stain and imaged using confocal microscopy (CM). The remaining two coupons from each set were diluted, plated and enumerated for viable cells. The viable plate count results were 9.230 Log10 CFU/coupon &#177; 0.007 (SD_R) for setting 1 and 9.272 Log10 CFU/coupon &#177; 0.066 (SD_R) for sonication setting 2. This data corroborated a 2015 EPA Single Tube Method Collaborative Study where 9.03 Log10 CFU/coupon &#177; 0.272 (SD_R) was achieved9. According to the viable plate counts, it appears that all three means are similar enough to not warrant further investigation into differences between the two harvesting methods. However, the microscopic images shown in Figure 2 may suggest that more biofilm remained after use of setting 1 than setting 2. While we observe that biofilm remaining on the coupons appears dead (red in color), interpretation of viability when using BacLight Live/Dead stain is difficult14, 15. Rather than focusing on viability implications of the stained biofilm, we used this stain to visualize biofilm remaining on the surfaces. Additionally, while we acknowledge that sonication could be deleterious to bacterial viability, a 2007 publication by Kobayashi et al.16 demonstrated that increased sonication time beyond 5 minues resulted in decreased viable plate counts. Since our study used a total of 1 minute sonication, we are confident that few cells were killed via sonication as shown by the &#62;9.2 LOG10 CFU/coupons for the two sonication parameters. It is interesting that a complete harvest of the biofilm from the surface was not achieved by either method. This finding demonstrates that viable plate counts alone are not adequate to determine harvesting and disaggregation bias and therefore must be paired with an additional method, microscopy, for example.
In addition to the sonicator settings, we investigated other important factors that affect sonication. These included the volume of liquid in the vials (10 or 40 mL), the type of liquid in the vial (buffered water or 2X D/E Neutralizing Broth) and the number of vials placed in the bath at the same time (3 or 12 vials)9.
P. aeruginosa biofilms on CDC Biofilm Reactor coupons were sonicated using sonicator setting 2 (45 kHz, 100% power, Normal setting, 30 &#177; 5 seconds). All samples were vortexed either 3 at a time or 6 at a time using a vortexer fitted with a 6-place tube attachment then sonicated as described in Figure 3. One coupon from each of the following categories was imaged using CM (Figure 3).
In the second study where sonication parameters were investigated, the microscopic images (Figure 3) suggest that minimizing the volume in the tubes, reducing the number of tubes processed at once and use of D/E Neutralizing Broth (which contains surfactant) all contribute to enhanced biofilm harvesting from the coupons.
Biocides may positively or negatively enhance harvesting and disaggregation. Similar to confirming that a neutralizer effectively stops the active while not increasing kill prior to performing an efficacy test, it is important to confirm that a biocide does not differentially impact harvesting and disaggregation. For efficacy testing, bias results if and only if there is differential removal for the control vs treated biofilm samples11.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62390/62390fig01.jpg" /> 
Figure 1: Photos of coupons stained with crystal violet demonstrating residual biofilm. A) Coupon removed from the reactor and stained with crystal violet. B, C, D) Three replicate coupons were separately &#34;treated&#34; with sterile buffered water then neutralized. To harvest and disaggregate, the coupons were vortexed (30 &#177; 5 seconds) and sonicated (45 kHz, 10% power, Sweep setting, 30 &#177; 5 seconds) twice then received a final vortex. Images courtesy of Danielle Orr and Blaine Fritz. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62390/62390fig01large.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/62390/62390fig02.jpg" /> 
Figure 2: Confocal microscopy images of coupons comparing two different sonication settings. Coupons (12.5X magnification) processed via sonication setting 1 (45 kHz, 10% power, Sweep setting) on the left or sonication setting 2 (45 kHz, 100% power, Normal setting) on the right. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62390/62390fig02large.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/62390/62390fig03.jpg" /> 
Figure 3. Confocal microscopy images of coupons comparing volumes, sonication liquid and number of tubes. Coupons (12.5X magnification) processed in 10 or 40 mL volumes, in dilution water (DW) or D/E Neutralizing Broth (D/E), 3 or 12 tubes at a time with optimized sonication setting (45 kHz, 100% power, Normal setting). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62390/62390fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1: Key parameters of importance for harvesting and disaggagregation <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62390/Supp file 1 _FINAL.docx" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Harvesting and Disaggregation: An Overlooked Step in Biofilm Methods Research at JoVE.com

Harvesting and Disaggregation: An Overlooked Step in Biofilm Methods Research

This paper details methods that demonstrate three common biofilm harvesting and disaggregation techniques on two surface types, ruggedness testing of a harvesting method and minimum information to consider when choosing and optimizing harvesting and disaggregation techniques to increase reproducibility.

Biofilm methods consist of four distinct steps: growing the biofilm in a relevant model, treating the mature biofilm, harvesting the biofilm from the ...

harvesting-disaggregation-an-overlooked-step-biofilm-methods

Universidad San Sebastian

Research

JoVE Journal

Biology

3.8K Views.  Montana State University. This paper details methods that demonstrate three common biofilm harvesting and disaggregation techniques on two surface types, ruggedness testing of a harvesting method and minimum information to consider when choosing and optimizing harvesting and disaggregation techniques to increase reproducibility.

Video: Harvesting and Disaggregation: An Overlooked Step in Biofilm Methods Research

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 ...

Immunology and Infection

Characterization of Aquatic Biofilms with Flow Cytometry

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

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

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

Environment

Discovering CsgD Regulatory Targets in Salmonella Biofilm Using Chromatin Immunoprecipitation and High-Throughput Sequencing (ChIP-seq)

Chromatin immunoprecipitation followed by sequencing (ChIP-seq) is a technique that can be used to discover the regulatory targets of transcription factors, histone modifications, and other DNA-associated proteins. ChIP-seq data can also be used to find differential binding of transcription factors in different environmental conditions or cell types. Initially, ChIP was performed through hybridization on a microarray (ChIP-chip); however, ChIP-seq has become the preferred method through technological advancements, decreasing financial barriers to sequencing, and massive amounts of high-quality data output. Techniques of performing ChIP-seq with bacterial biofilms, a major source of persistent and chronic infections, are described in this protocol. ChIP-seq is performed on Salmonella enterica serovar Typhimurium biofilm and planktonic cells, targeting the master biofilm regulator, CsgD, to determine differential binding in the two cell types. Here, we demonstrate the appropriate amount of biofilm to harvest, normalizing to a planktonic control sample, homogenizing biofilm for cross-linker access, and performing routine ChIP-seq steps to obtain high quality sequencing results.

Discovering CsgD Regulatory Targets in Salmonella Biofilm Using Chromatin Immunoprecipitation and High-Throughput Sequencing ChIP-seq

Chromatin immunoprecipitation coupled with next-generation sequencing (ChIP-seq) is a method used to establish interactions between transcription factors and the genomic sequences they control. This protocol outlines techniques for performing ChIP-seq with bacterial biofilms, using Salmonella enterica serovar Typhimurium bacterial biofilm as an example.

Chromatin immunoprecipitation followed by sequencing (ChIP-seq) is a technique that can be used to discover the regulatory targets of transcription ...

Genetics

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 ...

Use of a High-throughput In Vitro Microfluidic System to Develop Oral Multi-species Biofilms

There are few high-throughput in vitro systems which facilitate the development of multi-species biofilms that contain numerous species commonly detected within in vivo oral biofilms. Furthermore, a system that uses natural human saliva as the nutrient source, instead of artificial media, is particularly desirable in order to support the expression of cellular and biofilm-specific properties that mimic the in vivo communities. We describe a method for the development of multi-species oral biofilms that are comparable, with respect to species composition, to supragingival dental plaque, under conditions similar to the human oral cavity. Specifically, this methods article will describe how a commercially available microfluidic system can be adapted to facilitate the development of multi-species oral biofilms derived from and grown within pooled saliva. Furthermore, a description of how the system can be used in conjunction with a confocal laser scanning microscope to generate 3-D biofilm reconstructions for architectural and viability analyses will be presented. Given the broad diversity of microorganisms that grow within biofilms in the microfluidic system (including Streptococcus, Neisseria, Veillonella, Gemella, and Porphyromonas), a protocol will also be presented describing how to harvest the biofilm cells for further subculture&#160;or DNA extraction and analysis. The limits of both the microfluidic biofilm system and the current state-of-the-art data analyses will be addressed. Ultimately, it is envisioned that this article will provide a baseline technique that will improve the study of oral biofilms and aid in the development of additional technologies that can be integrated with the microfluidic platform.

Use of a High-throughput In Vitro Microfluidic System to Develop Oral Multi-species Biofilms

The goal of this methods paper is to describe the use of a microfluidic system for the development of multi-species biofilms that contain species typically identified in human supragingival dental plaque.&#160;Methods to describe biofilm architecture, biofilm viability, and an approach to harvest biofilm for culture-dependent or culture-independent analyses are highlighted.

There are few high-throughput in vitro systems which facilitate the development of multi-species biofilms that contain numerous species commonly detected ...

Bioengineering

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

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

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

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

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 ...

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.

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 ...

Quantifying the Effects of Antimicrobials on In vitro Biofilm Architecture using COMSTAT Software

Biofilms are aggregates of microorganisms that rely on a self-produced matrix of extracellular polymeric substance for protection and structural integrity. The nosocomial pathogen, Pseudomonas aeruginosa, is known to adopt a biofilm mode of growth, causing chronic pulmonary infection in patients with cystic fibrosis (CF). The computer program, COMSTAT, is a useful tool for quantifying antimicrobial-induced changes in P. aeruginosa biofilm architecture by extracting data from three-dimensional confocal images. However, standardized operation of the software is less commonly addressed, which is important for optimal reporting of biofilm behavior and cross-center comparison. Thus, the aim of this protocol is to provide a simple and reproducible framework for quantifying in vitro biofilm structures under varying antimicrobial conditions via COMSTAT. The technique is modeled using a CF P. aeruginosa isolate, grown in the form of biofilm replicates, and exposed to tobramycin and the anti-Psl monoclonal antibody, Psl0096. The step-by-step approach aims to reduce user ambiguity and minimize the chance of overlooking crucial image-processing steps. Specifically, the protocol emphasizes the elimination of subjective variations associated with the manual operation of COMSTAT, including image segmentation and the selection of appropriate quantitative analysis functions. Although this method requires users to spend additional time processing confocal images prior to running COMSTAT, it helps minimize misrepresented biofilm heterogenicity in automated outputs.

Quantifying the Effects of Antimicrobials on In vitro Biofilm Architecture using COMSTAT Software

Antimicrobial-induced changes to Pseudomonas aeruginosa biofilm architecture differ among clinical isolates cultured from patients with cystic fibrosis and chronic pulmonary infection. Following confocal microscopy, COMSTAT software can be utilized to quantify variations in biofilm architecture (e.g., surface area, thickness, biomass) for individual isolates to assess the efficacy of anti-infective agents.

Harvesting and Disaggregation: An Overlooked Step in Biofilm Methods Research

Summary

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Chapters in this video

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

Characterization of Aquatic Biofilms with Flow Cytometry

Discovering CsgD Regulatory Targets in Salmonella Biofilm Using Chromatin Immunoprecipitation and High-Throughput Sequencing (ChIP-seq)

Concurrent Quantification of Cellular and Extracellular Components of Biofilms

Use of a High-throughput In Vitro Microfluidic System to Develop Oral Multi-species Biofilms

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

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

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

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

Quantifying the Effects of Antimicrobials on In vitro Biofilm Architecture using COMSTAT Software