Funding for this work was provided by operating grants to DCB from the Natural Science and Engineering Research Council of Canada Discovery Grant (RGPIN- 2016-05891) and to MRM and GRG from the Government of Canadian Genomics Research and Development Initiative Grant (GRDI) program (GRDI7 2254557).

1. Aseptic culture preparation for biofilm growth (Days 0-1)
<ol>
	<li>On Day 0, streak the desired bacterial strain(s) for testing from a cryo-preserved stock (in glycerol or dimethylsulfoxide (DMSO)) directly onto a nutrient agar plate with antimicrobial selection as the strain requires. Incubate agar plates at the optimal temperature (37&#160;&#176;C) and time (18-22 h) for strain growth.</li>
	<li>The following day (Day 1), inoculate a colony from the agar plate in 5 mL of growth media at optimal growth conditions. These cultures will be used as Day 2 starting inoculates for biofilm devices (see step 2.2; Figure 1). 
	NOTE: For this study, E. coli K-12 BW21153 and P. aeruginosa PAO1 were grown in Luria-Bertani (LB) medium at 37 &#176;C for 18 h with shaking at 160 revolutions per minute (rpm).
	<ol>
		<li>Prepare at least 3 independently cultured strains to generate biological replicates for biofilm measurement due to the statistical variability of biofilm biomass formation on the pegged lid devices.</li>
	</ol>
	</li>
</ol>
2. Preparation and inoculation of plates (Day 2)
<ol>
	<li>Fill the outer wells of an autoclaved 96-well, 1.1 mL/well polypropylene plate with 750 &#181;L/well sterile water (Figure 2A). Fill the negative control wells (uninoculated media wells; column B) with 750 &#181;L growth media and the remaining wells with 675 &#181;L growth media. 
	NOTE: Step 2.1 above and steps below list&#160;appropriate volumes for the deep well biofilm device. If using the standard biofilm device, the volumes are to be changed as follows: 75 &#181;L of inoculum culture to 20 &#181;L; 675 &#181;L of growth media to 180 &#181;L; 750 &#181;L of sterile water/ growth media to 200 &#181;L; 800 &#181;L of CV stain/acetic acid/ rinse solutions to 210 &#181;L. Additionally, for step 3.6 below, the Absorbance at 500nm (A550 nm)&#160;can be read directly in the polystyrene microplate after mixing when using the standard biofilm device microplate.</li>
	<li>Using Day 1 overnight cultures, standardize the cultures to an optical density at 600 nm (OD600nm) =1.0 or to a McFarland standard of 0.5-1 units.</li>
	<li>Create a 10-fold serial dilution of each standardized culture in test tubes or a 96-well deep well microtiter plate until a 10-3 dilution is reached.</li>
	<li>Inoculate the media containing wells as shown in Figure 2B with 75 &#181;L of the 1 x 10-3 diluted culture, for a final in-well bacterial dilution of 10-4.</li>
	<li>Carefully insert an autoclaved PCR plate (pegged lid) into the inoculated deep well plate. Incubate plates at optimal strain conditions (37 &#176;C) with shaking (maximum 160 rpm) in an incubator with 50-60% humidity for 24 h.</li>
</ol>
3. Biofilm biomass determination from devices using CV staining (Day 3)
<ol>
	<li>Aseptically remove the biofilm pegged lid from each device and rinse by inserting into an autoclaved deep well plate prepared with 800 &#181;L of sterile phosphate buffered saline (PBS)/well for 30 s to remove planktonic cells.</li>
	<li>For biomass CV staining, transfer the pegged PCR plate lid into a new plate prepared with 800 &#181;L/well 0.1% (w/v) CV in dH2O and stain for 5 minutes.</li>
	<li>Remove excess stain by rinsing the pegged lid in a deep well plate prepared with 800 &#181;L/well sterile PBS. Allow the plates to dry for 10 minutes in a biosafety cabinet with pegs facing upward.</li>
	<li>Transfer the dried PCR plate lid to a plate containing 800 &#181;L/well 30% (v/v) acetic acid and allow the lid to de-stain for 5 minutes.</li>
	<li>Remove the destained PCR peg lid and mix the solution thoroughly by pipetting. Transfer 200 &#181;L from the deep well plates to a standard 96 well microplate and measure the absorbance of the de-staining solution at a wavelength of 550 nm (A550nm) in an ultraviolet (UV)/ visible (Vis) range microplate reader.</li>
	<li>Using the measured A550nm values, subtract the average blank (negative control) A550nm value from each biofilm sample value. Average each blank-subtracted A550nm biofilm sample value representing sample biological replicates.</li>
</ol>
4. Challenging biofilms with antimicrobials to calculate their 24 h antimicrobial MBEC value (Day 3)
<ol>
	<li>Prepare an antimicrobial stock solution in water that is two times (2x) the concentration of the desired highest antimicrobial concentration to be tested.</li>
	<li>Create a two-fold dilution series of the antimicrobial stock solution in 2x concentrated growth media so that there are 8 antimicrobial concentrations.</li>
	<li>As shown in Figure 2B, fill the outer wells of an autoclaved deep well plate with 750 &#181;L/well sterile water. Fill columns B (negative control; no bacteria) and C (positive control; no antimicrobial exposure) of the plate with 750 &#181;L/well growth media. Fill the remaining wells with 750 &#181;L/well of the antimicrobial dilution series as shown in Figure 2B.</li>
	<li>Remove and rinse the pegged lid as described in step 3.1 and transfer them into the antimicrobial challenge plate.</li>
	<li>Incubate plates with shaking (max 160 rpm) in appropriate strain conditions and for the desired exposure timeframe.</li>
</ol>
5. Recovery of biofilm biomass from pegged lids (Day 4)
<ol>
	<li>Remove antimicrobial exposed pegged lids and rinse as described in step 3.1. Transfer the pegged lid to a deep well plate with 750 &#181;L/well recovery media in the inner wells and 750 &#181;L/well sterile dH2O in the outer wells.
	<ol>
		<li>To prepare 70 mL recovery media, combine 35 mL of 2x concentrated LB, 0.7 mL of 100% tween-20, 3.5 mL of 20x universal neutralizing solution, and 30.8 mL of sterile dH2O in a sterile bottle.</li>
		<li>To prepare 20x concentrated universal neutralizing solution, add 1.0 g of L-histidine, 1.0 g of L-cysteine, and 2.0 g of reduced glutathione to a final volume of 20 mL of dH2O. Filter sterilize the solution through a 0.2 &#181;m filter and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Put the device from step 5.1 into a secondary bin fitted inside a sonicating water bath. Sonicate the devices for 30 minutes to dislodge the biofilm from pegs into the recovery media.</li>
	<li>After sonication, remove the pegged lid and place a sterile flat plate cover lid on the deep well recovery media plate. The pegged lid can be discarded.</li>
	<li>Incubate plates with recovery media from step 5.3 at the optimal strain growth conditions overnight (16-18 h).</li>
</ol>
6. Determination of device MBEC values (Day 5)
<ol>
	<li>The following day, transfer 200 &#181;L from each well of the incubated deep well plate from step 5.4 to a new 96-well microtiter plate. Read the OD600nm of each well using a UV/Vis range microplate reader.</li>
	<li>Subtract negative control (uninoculated blank) OD600nm values from each biofilm containing sample well.</li>
	<li>Calculate the MBEC value for each sample using OD600nm values, where the MBEC value is the lowest antimicrobial concentration that resulted in the lowest OD600nm value (indistinguishable from the uninoculated control) for a specific antimicrobial treated species/strain sample.</li>
</ol>

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The authors have no disclosures to make.

This study describes methods for using a larger volume 96-well high throughput static biofilm growth device involving a polypropylene deep well microplate fitted with a semi-skirted PCR-plate lid for biofilm formation (deep well biofilm device; Table of Materials). We compared biofilms generated with this device to a commercially available polystyrene standard biofilm device (Table of Materials). The deep well device method uses the same methodological steps and solutions as the standard device17 at adjusted volumes modified for the deep well devices, making this device ideal for large-scale biofilm formation and experimental analysis. The growth of E. coli BW25113 and P. aeruginosa PAO1, two Gram-negative species that are known to form biofilms, were examined for their biomass formation and disinfectant (BZK/bleach) MBEC values using on both devices. Comparison of CV-stained biomass formed on pegs from each device showed that both E. coli and P. aeruginosa formed biofilms with higher biomass on the deep well biofilm device when compared to the standard device (Figure 3A-B). Increased biofilm biomass reflected the larger surface area of deep well pegs when compared&#160;to the standard device peg surface area. When we accounted for differences in peg surface areas (mm2) with both devices, differences in biomass formation were noted, where E. coli formed significantly greater CV biomass per mm2 on polystyrene standard device pegs than with the deep well device&#39;s PCR-tube pegs (Table 1). P. aeruginosa formed greater CV-stained biomass/ peg mm2 as compared to standard devices (Table 1). These findings may highlight species-specific differences in biofilm biomass formation on the different devices.
It should be noted that bleach eradication of E. coli and P. aeruginosa biofilms on deep well devices occurred at 2-4-fold lower concentrations than standard device (Figure 4A-B, Table 1). The differences in bleach MBEC values identified from each device are likely impacted by differences in device peg shapes (deep well &#34;pegs&#34; are tapered tubes and standard pegs are cylindrical), plastic composition differences (polypropylene versus polystyrene), and volume differences (750 &#181;L versus 200 &#181;L). For example, P. aeruginosa had greater CV M biomass/ mm2 on deep well device pegs when compared to standard devices, but E. coli had less biomass on deep well pegs (Figure 3). This suggests that disinfectant concentrations required for biofilm eradication may be impacted by the biomass formed by each species as well as the available surface area. Additionally, differences in device peg shape may influence various growth conditions for particular species. In our study, P. aeruginosa may form greater biomass on deep well pegs due to their greater surface area and aeration, as this species is an obligate aerobe in contrast to E. coli, which is a facultative anaerobe. To date, we have not identified any published studies that have directly compared E. coli and P. aeruginosa biofilm formation together on both polypropylene27,28,29 and polystyrene30,31 materials. However, reports of robust E. coli and P. aeruginosa biofilm formation have been noted in independent studies examining either polypropylene or polystyrene materials. With respect to Pseudomonads, many Pseudomonas spp. can use plastics such as polypropylene as potential carbon sources32. Hence, the availability of this polypropylene deep well biofilm device is a useful advance in static biofilm studies. Polypropylene is chemically more durable than polystyrene and is a clinically relevant material, as it is frequently used in medical implants, sutures, and meshes for hernia or pelvic surgeries33,34.
Although biofilm biomass was formed by both devices, the deep well device had slightly higher variability in biomass based on the CV staining method and OD600nm biofilm eradication MBEC values for bleach and BZK. This may be explained by 3 main factors: 1) Deep well devices have greater peg surface area than standard devices that were angled as compared to standard device pegs. 2) Both species tested may have differing abilities to adhere to polypropylene and polystyrene materials of each device. 3) The volumes of growth media used in each device (750 &#181;L deep well, 200 &#181;L standard) and spacing between the inserted peg to the well side walls differs. These issues are not a problem if only one type of device is used for all biofilm experiments, however, if both devices are selected then the comparisons&#160;we conducted herein should be performed to identify differences36,37. Due to the differences in plastic material used in each device, the CV-stained biofilm biomass and MBEC values should not be directly compared between different devices. However, if methods and experiments are conducted on the same device (deep well or standard), the results obtained for species and antimicrobials tested are comparable.
This protocol shows that self-assembled deep well PCR-plate devices are larger volume biofilm device for measuring biofilm formation and eradication that is also cost-effective. From a cost perspective, standard biofilm devices with a 96 well pegged lids range retail at $29-36 US dollars (USD) per device (Table of Materials). Polystyrene standard biofilm devices are not autoclavable and are less tolerant to solvents/acids due to its plastic chemical properties. The self-assembled polypropylene deep well plates described herein, fitted with a separate semi-skirted 96 well PCR-plate (Table of Materials) cost a total of $14 USD per assembled device, which is half the standard biofilm device cost. It should be noted that prices may vary depending on region, distributor, and availability, and our costs after institutional discounts worked out to $9 USD/deep well device. These self-assembled deep well polypropylene PCR-plate devices have the added advantage of being autoclavable for sterilization and offer 2-4 times more biofilm biomass than standard devices.
In conclusion, this protocol and the representative findings from biofilm growth comparisons of the deep well and standard biofilm devices show that both devices are capable of cultivating bacterial biofilms, but deep well devices form 2-4 times more biofilm. The deep well biofilm device offers a viable&#160;and affordable alternative for larger volume high-throughput biofilm formation experiments such as drug susceptibility screening studies. This technique may generate biofilms useful for downstream &#39;-omics&#39; extractions (proteomic, metabolomic, transcriptomic) or experimental assays (enzymatic, fluorescent) that may require larger quantities of biofilm materials for analyses. The deep well biofilm device is recommended for labs that would like to study biofilms in a high-throughput 96-well assay using lower-cost, larger volume, chemically durable plastic materials that are clinically relevant.

Pseudomonas aeruginosa and Escherichia coli are Gram-negative proteobacteria that are commonly studied for their ability to form sessile, surface-attached cell communities known as biofilms1. Both species, when growing as biofilms, can secrete a matrix of extracellular polymeric substances (EPS) composed primarily of different polysaccharides and proteins, that can also include extracellular DNA and/or lipids, providing added cellular protection and enhanced survival in harsh, nutrient-limited environments2,3. Biofilm physiology and formation by both species is of clinical relevance, as they represent the top five most commonly isolated antimicrobial resistant pathogens from hospital patient blood, urinary tract and lung infections in Canada4,5. It is also important to note that biofilms are estimated to make up nearly 80% of all chronic and recurrent infections caused by these bacterial species6. Due to the secreted EPS substances and slower metabolic activity7,8, established biofilms become more difficult to eradicate when they form on surfaces such as implanted medical devices, scar tissues, and in the lungs of cystic fibrosis patients9,10, adding to their antimicrobial resistance.
The recalcitrant growth properties of bacterial biofilms often make them more resistant to antimicrobial inhibition and/or eradication2,9. Thus, establishing in vitro methods to study bacterial biofilm antimicrobial eradication is paramount for selecting effective compounds to eradicate infections when they form on medical plastics (e.g., in-dwelling catheters) and medical implants. Most rapid in vitro biofilm antimicrobial eradication culturing methods examine biofilm growth as a &#34;static&#34; biofilm culture rather than &#34;continuous&#34; cultures, where static bacterial growth is monitored in early to late stages of biofilm formation over a short (24-96 h) timeframe in the same growth medium. Continuous biofilm methods are cumbersome for rapid analysis as they require biofilm growth assessed over longer timeframes grown in chambers that allow the continuous flow and replacement of growth media with fewer replicates11. Due to the time and effort required to maintain and establish continuous biofilms, static in vitro biofilms are the most popular, as they are easily adapted for high-throughput antimicrobial susceptibility testing assays in plastic 96-well microtiter plates rather than elaborate flow chamber systems that limit the number&#160;of cultures simultaneously tested 12,13. The simplest static &#34;in-well&#34; biofilm microplate assays use standard polystyrene or vinyl (300 &#181;L capacity) microtiter plates to measure biofilm formation on the sides and bottoms of each well and often as rings at the air to liquid surface interface. Bacterial in-well biofilm formation is measured by staining the deposited biomass adhered to the wells after the growth medium liquid is removed and the biofilms are washed12,13. These assays are economically popular but often have reproducibility issues due to their inherent design, as deposited biofilms are prone to damage or loss during rinsing for cell recovery and biomass staining procedures11,14.
To reduce biofilm losses, standard commercial biofilm devices have improved upon in-well biofilm microplate designs by adding an insertable 96-well pegged polystyrene lid to the standard 96 in-well plate design, referred to as the &#34;standard biofilm device&#34; herein. The addition of a pegged lid expands the available surface area in each microplate well, allowing for enhanced surface adherence and biofilm biomass formation15,16. Standard biofilm pegged lid devices allow for greater biofilm recovery, removal, and rinsing for subsequent antimicrobial biofilm susceptibility and eradication testing when pegged lids are inserted into new microtiter plates containing drug or growth condition challenges. Similar to in-well biofilm microplate techniques, the recovered materials from the removed and washed pegged lid devices allows for cell survival testing and biofilm biomass staining, typically involving crystal violet (CV) dye formulations 17,18,19. Standard biofilm devices are also optimal for screening biofilm antimicrobial susceptibilities. These assays monitor biofilm growth inhibition in two ways: 1) When antimicrobials are added to cells at the start of growth, it can determine the minimum biofilm inhibition concentration (MBIC) value. 2) When established biofilms are formed on the pegs after 24 hours and then exposed to antimicrobials, it can determine the minimum biofilm eradication concentration (MBEC) value 17,20. Similar to in-well biofilm microplate devices, standard biofilm devices have some notable limitations, such as their high cost per device, they are non-autoclavable, and less durable to chemical solvents due to the polystyrene plate material used. Standard biofilm devices also have a low surface area to peg ratio, which limits the maximum working volumes in each well to 200 &#181;L. These factors can make standard biofilm devices more challenging to use for studies that demand greater quantities of biofilm in a high-throughput format.
Here, we describe a static biofilm pegged-lid 96-well method developed in our lab using commercially available polypropylene semi-skirted 0.5 mL 96-well PCR plates fitted to 96-well microtiter plates having wells deeper than the standard plate (Table of Materials). These assembled devices have a maximum working volume of 750 &#181;L when used for growing biofilm (herein known as &#34;deep well biofilm device&#34;). The advantages of using these deep well biofilm devices are their lower cost when compared to standard biofilm devices, they can be sterilized by autoclaving, and they increase the surface area for biofilm formation on the larger external PCR-plate &#34;tube/pegs&#34;. With this method we show the applications of these devices for growing and characterizing biofilm biomass formed by Escherichia coli BW25113 and P. aeruginosa PAO1. Methods to determine MBEC values using biofilm eradication assays are described using the&#160;quaternary ammonium compound disinfectant benzalkonium chloride (BZK) and sodium hypochlorite (bleach) antimicrobials. The antiseptic/disinfectant BZK was selected&#160;as this compound is frequently used to inhibit biofilms from contaminated surfaces, but it is reportedly less effective at eradicating well-established biofilms21. Bleach&#160;is a highly effective chemical for the eradication of established biofilms and is a mainstay in chemical disinfection 22. Both disinfectants provide a useful comparison of MBEC values for each biofilm device21. A protocol for this biofilm device assessment using CV staining and biofilm eradication assays for MBEC determination is summarized in this article. A protocol flowchart has been included for a simplified overview of the workflow for these methods (Figure 1).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10 mL serological pipets, individ wrap paper/plastic (200/CS)</td>
			<td>Fisher Scientific</td>
			<td>13-678-11F</td>
			<td>disposable serological pipettes for aseptic media/ culture transfer</td>
		</tr>
		<tr>
			<td>2 mL serological pipets, individ wrap paper/plastic, 500/CS</td>
			<td>Fisher Scientific</td>
			<td>13-678-11C</td>
			<td>disposable serological pipettes for aseptic media/ culture transfer</td>
		</tr>
		<tr>
			<td>Axygen Aerosol Filter Tips, Sterile, 5 racks/ PK, 1000 tips/rack</td>
			<td>Fisher Scientific</td>
			<td>14-222-766</td>
			<td>sterile pipettor tips for media aliquoting</td>
		</tr>
		<tr>
			<td>Axygen deep well storage microplates, round wells, 1.1 mL cap, 5/PK, P-DW-11-C</td>
			<td>Fisher Scientific</td>
			<td>P-DW-11-C</td>
			<td>microplate for 96 well deepwell device biofilm cultivation</td>
		</tr>
		<tr>
			<td>Axygen Filter tips, 350 &#181;L tips racked, 96/rack, 10 racks, low retention barrier tips</td>
			<td>Fisher Scientific</td>
			<td>TF-350-L-R-S</td>
			<td>sterile pipettor tips for media aliquoting</td>
		</tr>
		<tr>
			<td>Basin/reservoir natural PS 50 mL, Sterile, 5/bag, 40 bags, CS200</td>
			<td>Avantor/ VWR</td>
			<td>89094-676</td>
			<td>sterile basins/ reservoirs for microplate preparation</td>
		</tr>
		<tr>
			<td>BD Difco Dehydrated Culture Media: Granulated Agar, 500 g</td>
			<td>Fisher Scientific</td>
			<td>DF0145-17-0</td>
			<td>materials for LB agar preparation</td>
		</tr>
		<tr>
			<td>Biotek Synergy Neo2 multimode plate reader</td>
			<td>Biotek</td>
			<td>NEO2MB</td>
			<td>microplate UV/Vis region plate reader</td>
		</tr>
		<tr>
			<td>Branson M3800 Ultrasonic Bath, 117 V</td>
			<td>Avantor/ VWR</td>
			<td>CPX-952-316R</td>
			<td>sonicating water bath</td>
		</tr>
		<tr>
			<td>crystal violet (CV), ACS grade, 100 g</td>
			<td>Fisher Scientific</td>
			<td>C581-100</td>
			<td>biofilm biomass stain</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO), 1 L, ACS grade 99.9%, poly bottle, BDH</td>
			<td>Avantor/ VWR</td>
			<td>CABDH1115-1LP</td>
			<td>media components for cryopreservation</td>
		</tr>
		<tr>
			<td>Easypet 3, pipet controller</td>
			<td>Avantor/ VWR</td>
			<td>CA11027-980</td>
			<td>serological pipettor for aseptic media/ culture transfer</td>
		</tr>
		<tr>
			<td>Eppendorf Research Plus 8 multi-channel pipettor , 10-100 &#181;L</td>
			<td>Avantor/ VWR</td>
			<td>CA89125-338</td>
			<td>multichannel pipettes for aseptic media/ culture transfer</td>
		</tr>
		<tr>
			<td>Eppendorf Research Plus 8 multi-channel pipettor , 30-300 &#181;L</td>
			<td>Avantor/ VWR</td>
			<td>CA89125-340</td>
			<td>multichannel pipettes for aseptic media/ culture transfer</td>
		</tr>
		<tr>
			<td>Glacial acetic acid, CAS 64-19-7, 2.5L, ACS grade</td>
			<td>Fisher Scientific</td>
			<td>A38-212</td>
			<td>CV destain</td>
		</tr>
		<tr>
			<td>Glass test tubes, 150 mm x 18 mm, 72/Pack, PYREX</td>
			<td>Fisher Scientific</td>
			<td>14957H/ 9820-18</td>
			<td>materials for cell culturing</td>
		</tr>
		<tr>
			<td>Glycerol, 4 L glass bottle ACS</td>
			<td>Fisher Scientific</td>
			<td>BP229-4</td>
			<td>media components for cryopreservation</td>
		</tr>
		<tr>
			<td>L-Cysteine, 98%, 250 g</td>
			<td>Avantor/ VWR</td>
			<td>97061-204</td>
			<td>universal neutralizing solution</td>
		</tr>
		<tr>
			<td>L-glutathione reduced, 98%, 25 g</td>
			<td>Fisher Scientific</td>
			<td>AAJ6216614</td>
			<td>universal neutralizing solution</td>
		</tr>
		<tr>
			<td>L-Hisitidine, 98%, 100 g</td>
			<td>Avantor/ VWR</td>
			<td>CA97062-598</td>
			<td>universal neutralizing solution</td>
		</tr>
		<tr>
			<td>MBEC Assay Inoculator with 96 well tray, 100/CS</td>
			<td>Innovotech</td>
			<td>19112</td>
			<td>material for 96 well MBEC device biofilm cultivation</td>
		</tr>
		<tr>
			<td>McFarland Standard, 0.5 EA</td>
			<td>Fisher Scientific</td>
			<td>R20410</td>
			<td>cell culture standardization</td>
		</tr>
		<tr>
			<td>McFarland Standard, 1.0 EA</td>
			<td>Fisher Scientific</td>
			<td>R20411</td>
			<td>cell culture standardization</td>
		</tr>
		<tr>
			<td>NUNC 96-well microtiter plates, w/lid, 50/CS</td>
			<td>Fisher Scientific</td>
			<td>167008</td>
			<td>microplate for 96 well MBEC device biofilm cultivation and OD measurements</td>
		</tr>
		<tr>
			<td>PCR plate, semi-skirted 96 well, fast PCR, polypropylene, 25ea/PK</td>
			<td>Sarstedt</td>
			<td>72.1981.202</td>
			<td>pegged lid for 96 well deepwell device biofilm cultivation</td>
		</tr>
		<tr>
			<td>Petri dish, 100 mm x 15 mm, semi-TK CS/500</td>
			<td>Fisher Scientific</td>
			<td>FB0875712</td>
			<td>materials for LB agar preparation</td>
		</tr>
		<tr>
			<td>Potassium phosphate dibasic, ACS 500 g</td>
			<td>Fisher Scientific</td>
			<td>P288-500</td>
			<td>PBS component/ buffer</td>
		</tr>
		<tr>
			<td>Sodium chloride, ACS grade, 3 kg</td>
			<td>Fisher Scientific</td>
			<td>S2713</td>
			<td>media components for LB broth and PBS</td>
		</tr>
		<tr>
			<td>sodium phosphate monobasic, 1 kg</td>
			<td>Fisher Scientific</td>
			<td>S369-1</td>
			<td>PBS component/ buffer</td>
		</tr>
		<tr>
			<td>Syringe filters, Sterile, PES 0.45 um, 25 mm, PK50</td>
			<td>Avantor/ VWR</td>
			<td>28145-505</td>
			<td>non-autoclavable solution sterilization</td>
		</tr>
		<tr>
			<td>Tin foil, heavy duty, 50 feet</td>
			<td>Grocery store</td>
			<td>---</td>
			<td>materials for deepwell device sterilization</td>
		</tr>
		<tr>
			<td>Tryptone (peptone from casein), 2.2 kg/EA</td>
			<td>Fisher Scientific</td>
			<td>B11922</td>
			<td>media components for LB broth</td>
		</tr>
		<tr>
			<td>Tween-20, 100 mL</td>
			<td>Fisher Scientific</td>
			<td>BP337-100</td>
			<td>recovery media solution</td>
		</tr>
		<tr>
			<td>Ultra-deepwell, 2.5 mL deep well plates (square well), with lid, polypropylene, 10/CS</td>
			<td>Avantor/ VWR</td>
			<td>37001-520</td>
			<td>materials for biofilm dilution preparation</td>
		</tr>
		<tr>
			<td>Yeast Extract, Fisher Bioreagents, 500 g</td>
			<td>Fisher Scientific</td>
			<td>BP1422-500</td>
			<td>media components for LB broth</td>
		</tr>
	</tbody>
</table>

generation of greater bacterial biofilm biomass using pcr-plate deep well microplate devices

Bacterial biofilms are difficult to eradicate from surfaces using conventional antimicrobial interventions. High-throughput 96-well microplate methods are frequently used to cultivate bacterial biofilms for rapid antimicrobial susceptibility testing to calculate minimal biofilm eradication concentration (MBEC) values. Standard biofilm devices consist of polystyrene pegged-lids fitted to 96-well microplates and are ideal for measuring biofilm biomass and MBEC values, but these devices are limited by available peg surface area for biomass accumulation and cost. Here, we outline a protocol to use self-assembled polypropylene 96-well deep well PCR-plate pegged-lid device to grow Escherichia coli BW25113 and Pseudomonas aeruginosa PAO1 biofilms. A comparison of 24-hour biofilms formed on standard and deep well devices by each species using crystal violet biomass staining and MBEC determination assays are described. The larger surface area of deep well devices expectedly increased overall biofilm formation by both species 2-4-fold. P. aeruginosa formed significantly greater biomass/mm2 on deep well pegs as compared to the standard device. E. coli had greater biomass/mm2 on standard polystyrene devices as compared the deep well device. Biofilm eradication assays with disinfectants such as sodium hypochlorite (bleach) or benzalkonium chloride (BZK) showed that both compounds could eliminate E. coli and P. aeruginosa biofilms from both devices but at different MBEC values. BZK biofilm eradication resulted in variable E. coli MBEC values between devices, however, bleach demonstrated reproducible MBEC values for both species and devices. This study provides a high throughput deep well method for growing larger quantities of biofilms on polypropylene devices for downstream studies requiring higher amounts of static biofilm.

The aim of this study was to provide a method for conducting larger volume, high-throughput (96-well), static biofilm device measurements using deep well biofilm devices. Here, we compared a self-assembled semi-skirted PCR-plate inserted into a deep well microplate, known as the deep well device, to a commonly used standard biofilm pegged lid device to examine their capabilities to form static biofilms. CV-stained biomass and biofilm eradication assays (MBEC) were used to assess biofilm formation by both devices.
To compare biofilm formation by different species on each device, we assessed biofilm biomasses formed by E. coli BW25113 and P. aeruginosa PAO1 using the biofilm CV staining protocol17. Although&#160;CV staining is generally reported as A550nm values, due to the differences in growth volumes of each device and their available surface areas&#160;we converted CV stain A550nm values to molar CV concentrations in solution. Molar CV concentrations accounted for volume differences of each device and allowed a comparison of CV concentrations representing biomasses recovered from each device. The results showed that both species produced significantly more biomass on deep well biofilm devices (2.1 fold&#160;E. coli;&#160;4.1 fold P. aeruginosa) compared to the standard biofilm device (Figure 3A-B). This outcome was expected given the larger surface area of the deep well PCR peg (320.4 mm2) as compared to the standard device peg surface area (108.9 mm2). This finding is also consistent with the increased the increased volumes (750 &#181;L) used in the deep well biofilm device as compared to the standard device wells (200 &#181;L). Hence, deep well devices increased biofilm biomass accumulation on PCR tube pegs as compared to smaller pegs with standard biofilm devices.
Both biofilm devices formed reproducible biofilms when we compared biologically replicated biofilms formed by E. coli and P. aeruginosa (Figure 3A-B). Despite observing greater variability in technical replicate CV-stained M A550nm values for either strain grown on the deep well device, there were no statistically significant differences when we compared the median CV M values for each species&#39; biological replicates on the deep well or standard devices using pairwise two-way Analysis of variance (ANOVA) or Student&#39;s T-tests (both p &#62; 0.05). This finding shows that biofilm formation by deep well and standard devices form reproducible biofilms. However, the CV-staining results also showed that when we accounted peg surface area differences on each device, the biofilm biomass formation by E. coli and P. aeruginosa showed statistically significant differences in biomass accumulation (Table 1). Calculation of mean CV-stained biomass (M) per peg surface area in mm2 (CV M/mm2) for E. coli showed that biofilm formation was 1.5-fold lower on polypropylene deep well devices when compared to the standard biofilm device (Table 1). However, the opposite result was obtained for P. aeruginosa, which demonstrated 1.4-fold higher CV M/mm2 on polypropylene deep well devices when compared to standard biofilm devices (Table 1). Despite the observed species-specific differences in biofilm biomass accumulation with each device, the deep well device still demonstrated greater overall (2-4-fold increases) biofilm biomass formation by each species (Figure 3).
To determine the antimicrobial susceptibility testing applications of the deep well biofilm device, we compared two commonly used disinfectants, BZK and bleach, for their biofilm eradicating potential. Both chemicals are commonly used to prevent (BZK) and/or eradicate (bleach) bacterial biofilms in clinical and industrial applications21,22,23. Each compound was added in increasing 2-fold concentrations to biofilms formed by E. coli and P. aeruginosa on the standard and deep well biofilm devices22,23(Figures 1, 2B). The lowest concentration of BZK or bleach that resulted in OD600nm values that were indistinguishable from negative control wells were defined as the MBEC value. Treatment of biofilms formed on standard biofilm devices with bleach resulted in the same bleach MBEC values of 0.625% for E. coli and P. aeruginosa (Table 1, Figure 4A-B). Bleach MBEC values determined for E. coli and P. aeruginosa biofilm formed on deep well devices showed 2-4 fold lower MBEC values by both species (E. coli; 0.156%; P. aeruginosa 0.313%) when compared to standard devices (Table 1). A 2-fold difference in bleach MBEC values between species was noted for the deep well device, where P. aeruginosa required a 2-fold higher concentration of bleach to eradicate biofilms as compared to E. coli (Figure 4A-B, Table 1). The 2-fold bleach MBEC difference between P. aeruginosa and E. coli in deep well device appears to inversely correlate with 1.5-fold increased CV-stained biomass formation by P. aeruginosa as compared to E. coli (Figure 3A-B). The larger amount of biomass formed by P. aeruginosa on deep well device pegs may also explain why a higher bleach concentration was required to eradicate P. aeruginosa biofilms when compared to E. coli on deep wells (Figure 3A-B, Table 1). Hence, deep well device biofilm eradication assays show that both species were susceptible to bleach but at lower (2-4-fold) bleach concentrations as compared to standard devices. This inversely correlates with the 3-fold greater peg surface area and volumes of the deep well device PCR plate pegs. This suggests&#160;that for bleach exposure, greater biomass surface area may lower the bleach concentrations necessary for biofilm eradication.
BZK biofilm eradication testing showed greater variability in recovered growth (OD600nm values) and MBEC values by each species when compared to bleach results (Figure 4C-D). This variability is not unexpected based on previous studies showing that BZK was more effective at preventing biofilm formation than eradicating well-established biofilms24,25,26. Using the standard biofilm device, only P. aeruginosa biofilms treated with BZK showed a consistent MBEC value (2560 &#181;g/mL) that was 8-fold lower than the MBEC value (20480 &#181;g/mL) obtained for this strain in the deep well device (Table 1, Figure 4C). These results may reflect differences in the amounts of P. aeruginosa biomass formed on deep well pegged surfaces and plastic compositions when compared to the standard device pegs. BZK eradication of biofilms formed by E. coli on both the deep well and standard devices were equally poor, resulting in a broad range of BZK MBEC values from 2560-40960 &#181;g/mL for the deep well device and 320-10240 &#181;g/mL on the standard device (Figure 4D). For E. coli, this variability was explained by the occasional instance where 1-2 replicate wells showed low but statistically significant growth in recovery media, that increased error and reduced BZK MBEC determination&#160;accuracy with either device (Figure 4D, Table 1). This variability highlights the ineffectiveness of using BZK in eradicating E. coli biofilms as previously noted in studies24,25,26. In contrast P. aeruginosa biofilms could be reliably eradicated by BZK at a defined concentration with both devices, but with an&#160;8-fold BZK concentration difference (Figure 4C). In summary, BZK eradication MBEC values of biofilms formed by P. aeruginosa show distinct MBEC values with each device, however, both devices are equally poor in distinguishing E. coli precise BZK eradication phenotypes.
Hence, the deep well biofilm device method described herein is similarly effective for forming reproducible biofilms when compared to a standard biofilm device..
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63069/63069fig01.jpg" /> 
Figure 1. Schematic overview of experiment. Day 0 isolation of single colonies from cryopreserved stock; Day 1 inoculation of growth media with single colonies; Day 2 inoculation of biofilm plate; Day 3 CV staining or&#160;antimicrobial challenge; Day 4 biofilm sonication into recovery media; and Day 5 reading OD600nm&#160;of recovery plate to determine MBEC values. Some images in figure provided by Servier Medical Art (smart.servier.com). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63069/63069fig01large.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/63069/63069fig02.jpg" /> 
Figure 2. Example plate layouts shown for various steps of the protocol. A) The final plate setup for the initial 24 h biofilm growth using Day 1&#160;bacterial cultures from two bacterial strains (E. coli and P. aeruginosa), each with three biological replicates. Negative control wells (grey) are used as sterility controls (no bacteria added). B) Plate setup for antimicrobial challenge of biofilms. Darkening colours indicate increasing antimicrobial concentration. Negative control are wells with no bacteria added (Grey) and positive controls are biofilms with no antimicrobial exposure (orange). Biofilm peg lids taken from panel A are transferred to this deep well plate for antimicrobial exposure. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63069/63069fig02large.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/63069/63069fig3v2.jpg" /> 
Figure 3. The molar concentration (M) of CV-stained E. coli BW25113. (A) and P. aeruginosa PAO1 (B) biofilm biomass recovered from standard (circles) and deep well (triangles) devices. CV staining was measured by reading the absorbance of CV at 550 nm (A550nm), and deep well A550nm readings were adjusted by a factor of 3.809 (800 &#181;L/ 210 &#181;L) to account for the volume differences between the devices. CV molar concentration (M) was determined by the Beer-Lambert law (CV Concentration =A550nm/&#949;l); where the pathlength (l) of 210 &#181;L in the 96 well flat bottom microtitre plate was determined to be 0.56 cm and the extinction coefficient (&#949;) of CV in water at A550nm of 251500 cm-1M-1&#160;39. Results represent 9 technical replicates from three biological replicates of each strain grown as a biofilm and each biological replicate&#39;s median value is shown as a horizontal bar in each plot. Statistically significant differences in biomass between the devices was determined between biological replicate median values using a two-tailed paired t-test (E. coli p= 0.008-0.018 (*); P. aeruginosa p= 0.002-0.009 (**)) shown as bars with asterisks. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63069/63069fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63069/63069fig4v2.jpg" /> 
Figure 4. Determination of antimicrobial MBEC concentration. Determination of antimicrobial MBEC concentration for bleach (A,B) and BZK (C,D) for P. aeruginosa PAO1 (A, C) and E. coli BW25113 (B,D) when grown on standard (white bars) and deep well (grey bars) biofilm pegged devices. Results were determined by reading of the OD600nm&#160;of biofilm recovered after sonication into recovery media and overnight incubation. Deep well OD600nm&#160;was adjusted by a factor of 3.75 (750 &#181;L /210 &#181;L) to account for volume differences between the devices. Results are representative of three biological replicates and error bars show the standard deviation between replicates at each antimicrobial concentration39. Asterisks (*) above each bar plot indicate statistically significant differences in OD600nm values from blank controls with p-values &#60; 0.05 using two-tailed Student&#39;s T-test calculations. Circle symbols (o) above each bar plot indicate measurements where only 1 or 2 replicates had statistically significant OD600nm values from blank controls. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63069/63069fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td colspan="2">BZK MBEC (&#181;g/mL)</td>
			<td colspan="2">Bleach MBEC (% v/v)</td>
		</tr>
		<tr>
			<td>Strain tested</td>
			<td>Deep well Device</td>
			<td>Standard Device</td>
			<td>Deep well Device</td>
			<td>Standard Device</td>
		</tr>
		<tr>
			<td>E. coli BW25113</td>
			<td>2560-40960</td>
			<td>320-10240</td>
			<td>0.156</td>
			<td>0.625</td>
		</tr>
		<tr>
			<td>P. aeruginosa PAO1</td>
			<td>20480</td>
			<td>2560</td>
			<td>0.313</td>
			<td>0.625</td>
		</tr>
		<tr>
			<td></td>
			<td>Deep well Device</td>
			<td>Standard Device</td>
			<td rowspan="2">p-value**</td>
			<td rowspan="2">Fold change (Deep well/ Standard)</td>
		</tr>
		<tr>
			<td>Strain tested</td>
			<td>Mean CV M*/ mm2 &#177; SD</td>
			<td>Mean CV M*/ mm2 &#177; SD</td>
		</tr>
		<tr>
			<td>E. coli BW25113</td>
			<td>1.99 x 10-8 &#177; 3.25 x10-9</td>
			<td>3.03 x 10-8 &#177; 3.31 x 10-9</td>
			<td>0.0176</td>
			<td>-1.52</td>
		</tr>
		<tr>
			<td>P. aeruginosa PAO1</td>
			<td>8.01 x 10-8 &#177; 9.42 x 10-9</td>
			<td>5.72 x 10-8 &#177; 9.42 x 10-9</td>
			<td>0.034</td>
			<td>1.4</td>
		</tr>
	</tbody>
</table>
Table 1. A summary of E. coli BW25113 and P. aeruginosa PAO1 mean CV-stained biomass M/ mm2 values and biofilm eradication MBEC values for BZK and bleach on various devices. 
*Mean CV M/mm2 values calculated using median biological replicate CV M values (Figure 3). 
**p-values determined using Two-tailed Student&#39;s T-test.

Watch this Scientific Journal Video about Generation of Greater Bacterial Biofilm Biomass using PCR-Plate Deep Well Microplate Devices at JoVE.com

Generation of Greater Bacterial Biofilm Biomass using PCR-Plate Deep Well Microplate Devices

This protocol presents methodology to perform biofilm growth and biomass measurements using self-assembled deep well PCR-plate devices for high-throughput 96-well pegged lid static biofilm screening.

Bacterial biofilms are difficult to eradicate from surfaces using conventional antimicrobial interventions. High-throughput 96-well microplate methods are ...

generation-greater-bacterial-biofilm-biomass-using-pcr-plate-deep

Research

JoVE Journal

Immunology and Infection

7.2K Views.  University of Manitoba. This protocol presents methodology to perform biofilm growth and biomass measurements using self-assembled deep well PCR-plate devices for high-throughput 96-well pegged lid static biofilm screening.

Video: Generation of Greater Bacterial Biofilm Biomass using PCR-Plate Deep Well Microplate Devices

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 ...

Protocol for Biofilm Streamer Formation in a Microfluidic Device with Micro-pillars

Several bacterial species possess the ability to attach to surfaces and colonize them in the form of thin films called biofilms. Biofilms that grow in porous media are relevant to several industrial and environmental processes such as wastewater treatment and CO2 sequestration. We used Pseudomonas fluorescens, a Gram-negative aerobic bacterium, to investigate biofilm formation in a microfluidic device that mimics porous media. The microfluidic device consists of an array of micro-posts, which were fabricated using soft-lithography. Subsequently, biofilm formation in these devices with flow was investigated and we demonstrate the formation of filamentous biofilms known as streamers in our device. The detailed protocols for fabrication and assembly of microfluidic device are provided here along with the bacterial culture protocols. Detailed procedures for experimentation with the microfluidic device are also presented along with representative results.

Protocols for the study of biofilm formation in a microfluidic device that mimics porous media are discussed. The microfluidic device consists of an array of micro-pillars and biofilm formation by Pseudomonas fluorescens in this device is investigated.

Several bacterial species possess the ability to attach to surfaces and colonize them in the form of thin films called biofilms. Biofilms that grow in ...

Bioengineering

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 ...

Enhanced Reproducibility and Precision of High-Throughput Quantification of Bacterial Growth Data Using a Microplate Reader

This study aimed to develop a repeatable, reliable, high-throughput protocol to monitor bacterial growth in 96-well plates and analyze the maximum growth rate. The growth curves and maximum growth rates of two bacterial species were determined. Issues including (i) lid condensation, (ii) pathlength correction, (iii) inoculation size, (iv) sampling time interval, and (v) spatial bias were investigated. The repeatability of the protocol was assessed with three independent technical replications, with a standard deviation of 0.03 between the runs. The maximum growth rates of Bacillus mycoides and Paenibacillus tundrae were determined to be&#160;(mean&#160;&#177; SD) 0.99 h&#8722;1 &#177;&#160; 0.03 h&#8722;1 and 0.85 h&#8722;1 &#177; 0.025 h&#8722;1, respectively. These bacteria are more challenging to monitor optically due to their affinity to clump together. This study demonstrates the critical importance of inoculation size, path length correction, lid warming, sampling time intervals, and well-plate spatial bias to obtain reliable, accurate, and reproducible data on microplate readers. The developed protocol and its verification steps can be expanded to other methods using microplate readers and high-throughput protocols, reducing the researchers&#39; innate errors and material costs.

Here, a high-throughput protocol is presented to measure growth data, including growth curves, growth rate, and maximum growth rate. The protocol was verified and validated using two biofilm-producing bacteria. The results and approach applied in this study can be expanded to other high-throughput protocols using microplate readers.

This study aimed to develop a repeatable, reliable, high-throughput protocol to monitor bacterial growth in 96-well plates and analyze the maximum growth ...

Environment

Quantifying Bacterial Surface Swarming Motility on Inducer Gradient Plates

Bacterial swarming motility is a common microbiological phenotype that bacterial communities use to migrate over semisolid surfaces. In investigations of induced swarming motility, specific concentration&#160;of an inducer may not be able to report events occurring within the optimal concentration range to elicit the desired responses from a species. Semisolid plates containing multiple concentrations are commonly used to investigate the response within an inducer concentration range. However, separate semisolid plates increase variations in medium viscosity and moisture content within each plate due to nonuniform solidification time.
This paper describes a one-step method to simultaneously test surface swarming motility on a single gradient plate, where the isometrically arranged test wells allow the simultaneous acquisition of multiconcentration responses. In the present work, the surface swarming of Escherichia coli K12 and Pseudomonas aeruginosa PAO1 were evaluated in response to a concentration gradient of inducers such as resveratrol and arabinose. Periodically, the swarm morphologies were imaged using an imaging system to capture the entire surface swarming process.
The quantitative measurement of the swarm morphologies was acquired using ImageJ software, providing analyzable information of the swarm area. This paper presents a simple gradient swarm plate method that provides qualitative and quantitative information about the inducers&#39; effects on surface swarming, which can be extended to study the effects of other inducers on a broader range of motile bacterial species.

Here, we describe the use of inducer gradient plates to evaluate bacterial swarming motility while simultaneously obtaining multiple concentration responses.

Bacterial swarming motility is a common microbiological phenotype that bacterial communities use to migrate over semisolid surfaces. In investigations of ...

Biochemistry

Assembly and Tracking of Microbial Community Development within a Microwell Array Platform

The development of microbial communities depends on a combination of complex deterministic and stochastic factors that can dramatically alter the spatial distribution and activities of community members. We have developed a microwell array platform that can be used to rapidly assemble and track thousands of bacterial communities in parallel. This protocol highlights the utility of the platform and describes its use for optically monitoring the development of simple, two-member communities within an ensemble of arrays within the platform. This demonstration uses two mutants of Pseudomonas aeruginosa, part of a series of mutants developed to study Type VI secretion pathogenicity. Chromosomal inserts of either mCherry or GFP genes facilitate the constitutive expression of fluorescent proteins with distinct emission wavelengths that can be used to monitor community member abundance and location within each microwell. This protocol describes a detailed method for assembling mixtures of bacteria into the wells of the array and using time-lapse fluorescence imaging and quantitative image analysis to measure the relative growth of each member population over time. The seeding and assembly of the microwell platform, the imaging procedures necessary for the quantitative analysis of microbial communities within the array, and the methods that can be used to reveal interactions between microbial species area all discussed.

The development of microbial communities depends on a combination of factors, including environmental architecture, member abundance, traits, and interactions. This protocol describes a synthetic, microfabricated environment for the simultaneous tracking of thousands of communities contained in femtoliter wells, where key factors such as niche size and confinement can be approximated.

The development of microbial communities depends on a combination of complex deterministic and stochastic factors that can dramatically alter the spatial ...

Precise, High-throughput Analysis of Bacterial Growth

Bacterial growth is a central concept in the development of modern microbial physiology, as well as in the investigation of cellular dynamics at the systems level. Recent studies have reported correlations between bacterial growth and genome-wide events, such as genome reduction and transcriptome reorganization. Correctly analyzing bacterial growth is crucial for understanding the growth-dependent coordination of gene functions and cellular components. Accordingly, the precise quantitative evaluation of bacterial growth in a high-throughput manner is required. Emerging technological developments offer new experimental tools that allow updates of the methods used for studying bacterial growth. The protocol introduced here employs a microplate reader with a highly optimized experimental procedure for the reproducible and precise evaluation of bacterial growth. This protocol was used to evaluate the growth of several previously described Escherichia coli strains. The main steps of the protocol are as follows: the preparation of a large number of cell stocks in small vials for repeated tests with reproducible results, the use of 96-well plates for high-throughput growth evaluation, and the manual calculation of two major parameters (i.e., maximal growth rate and population density) representing the growth dynamics. In comparison to the traditional colony-forming unit (CFU) assay, which counts the cells that are cultured in glass tubes over time on agar plates, the present method is more efficient and provides more detailed temporal records of growth changes, but has a stricter detection limit at low population densities. In summary, the described method is advantageous for the precise and reproducible high-throughput analysis of bacterial growth, which can be used to draw conceptual conclusions or to make theoretical observations.

Quantitative evaluation of bacterial growth is essential to understanding microbial physiology as a systems-level phenomenon. A protocol for experimental manipulation and an analytical approach are introduced, allowing for precise, high-throughput analysis of bacterial growth, which is a key subject of interest in systems biology.

Bacterial growth is a central concept in the development of modern microbial physiology, as well as in the investigation of cellular dynamics at the ...

Biology

Measuring the Migration and Biofilm Formation of Various Bacteria

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

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

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

In Vitro Polymicrobial Interactions in Cystic Fibrosis Lung Infections

Growing a Cystic Fibrosis-Relevant Polymicrobial Biofilm to Probe Community Phenotypes

Most in vitro models lack the capacity to fully probe bacterial phenotypes emerging from the complex interactions observed in real-life environments. This is particularly true in the context of hard-to-treat, chronic, and polymicrobial biofilm-based infections detected in the airways of individuals living with cystic fibrosis (CF), a multiorgan genetic disease. While multiple microbiome studies have defined the microbial compositions detected in the airway of people with CF (pwCF), no in vitro models thus far have fully integrated critical CF-relevant lung features. Therefore, a significant knowledge gap exists in the capacity to investigate the mechanisms driving the pathogenesis of mixed species CF lung infections. Here, we describe a recently developed four-species microbial community model, including Pseudomonas aeruginosa, Staphylococcus aureus, Streptococcus sanguinis, and Prevotella melaninogenica grown in CF-like conditions. Through the utilization of this system, clinically relevant phenotypes such as antimicrobial&#160;recalcitrance of several pathogens were observed and explored at the molecular level. The usefulness of this in vitro model resides in its standardized workflow that can facilitate the study of interspecies interactions in the context of chronic CF lung infections.

This protocol describes a cystic fibrosis (CF) lung-relevant four-species polymicrobial biofilm model that can be used to explore the impact of bacterial interspecies interactions.

Most in vitro models lack the capacity to fully probe bacterial phenotypes emerging from the complex interactions observed in real-life environments. This ...

Developing Synergistic Multispecies Biofilms from Rhizosphere Soil

An Approach to Constructing Multispecies Biofilm Communities from Rhizosphere Soil

The multispecies biofilm is a naturally occurring and dominant lifestyle of bacteria in nature, including in rhizosphere soil, although the current understanding of it is limited. Here, we provide an approach to rapidly establish synergistic multispecies biofilm communities. The first step is to extract cells from rhizosphere soil using the differential centrifugation method. Afterward, these soil cells are inoculated into the culture medium to form pellicle biofilm. After 36 h of incubation, the bacterial composition of the biofilm and the solution underneath are determined using the 16S rRNA gene amplicon sequencing method. Meanwhile, high-throughput bacterial isolation from pellicle biofilm is conducted using the limiting dilution method. Then, the top 5 bacterial taxa are selected with the highest abundance in the 16S rRNA gene amplicon sequencing data (pellicle biofilm samples) for further use in constructing multispecies biofilm communities. All combinations of the 5 bacterial taxa were quickly established using a 24-well plate, selected for the strongest biofilm formation ability by the crystal violet staining assay, and quantified by qPCR. Finally, the most robust synthetic bacterial multispecies biofilm communities were obtained through the methods above. This methodology provides informative guidance for conducting research on rhizosphere multispecies biofilm and identifying representative communities for studying the principles governing interactions among these species.

Author Spotlight: Developing Synthetic Microbial Communities for Generating Second-Generation Biofertilizers

A rapid and standardized procedure for establishing synergistic multispecies biofilm communities from various rhizosphere soils is presented here. It is a unique protocol designed to probe and simulate the complex rhizosphere soil microbiota.

The multispecies biofilm is a naturally occurring and dominant lifestyle of bacteria in nature, including in rhizosphere soil, although the current ...

Generation of Greater Bacterial Biofilm Biomass using PCR-Plate Deep Well Microplate Devices

Summary

Explore More Videos

Chapters in this video

Microtiter Dish Biofilm Formation Assay

Protocol for Biofilm Streamer Formation in a Microfluidic Device with Micro-pillars

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

Enhanced Reproducibility and Precision of High-Throughput Quantification of Bacterial Growth Data Using a Microplate Reader

Quantifying Bacterial Surface Swarming Motility on Inducer Gradient Plates

Assembly and Tracking of Microbial Community Development within a Microwell Array Platform

Precise, High-throughput Analysis of Bacterial Growth

Measuring the Migration and Biofilm Formation of Various Bacteria

Growing a Cystic Fibrosis-Relevant Polymicrobial Biofilm to Probe Community Phenotypes

An Approach to Constructing Multispecies Biofilm Communities from Rhizosphere Soil