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/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/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/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/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 ...

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Immunology and Infection

7.1K 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 ...

In vitro Biofilm Formation in an 8-well Chamber Slide

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

In vitro Biofilm Formation in an 8-well Chamber Slide

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

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

Immuno-fluorescence Assay of Leptospiral Surface-exposed Proteins

Bacterial surface proteins are involved in direct contact with host cells and in uptake of nutrients from the environment 1. For this reason, cellular localization can provide insights into the functional role of bacterial proteins. Surface localization of bacterial proteins is a key step towards identification of virulence factors involved in mechanisms of pathogenicity. 
Methods for fractionating leptospiral membranes 2-5 may be selective for a certain class of outer-membrane proteins (OMPs), such as lipoproteins vs. transmembrane OMPs, and therefore lead to misclassification. This likely is due to structural differences and how they are associated to the outer membrane. Lipoproteins are associated with membranes via a hydrophobic interaction between the N-terminal lipid moiety (three fatty acids) and the lipid bilayer phospholipids 6, 7. In contrast, transmembrane OMPs are typically integrated into the lipid bilayer by amphipathic &beta;-sheets arranged in a barrel-like structure 8, 9. In addition, presence of a protein in the outer-membrane does not necessarily guarantee that the protein or its domains are exposed on the surface. Spirochetal outer membranes are known to be fragile and therefore necessitate methods involving gentle manipulation of cells and inclusion of sub-surface protein controls to assess the integrity of the outer membrane.
Here, we present an immunofluorescence assay (IFA) method to directly assess surface exposure of proteins on intact leptospires. This method is based on recognition of leptospiral surface proteins by antigen-specific antibodies. Herein, antibodies specific for OmpL5410 are detetcted aftero binding to native, surface exposed epitopes. Comparison of antibody reactivity to intact versus permeabilized cells enables evaluation of cellular distribution and whether or not a protein is selectively present on leptospiral surface. The integrity of outer membrane should be assessed using antibody to one or more subsurface proteins, preferably located in the periplasm.
The surface IFA method can be used to analyze surface exposure of any leptospiral protein to which specific antibodies are available. Both the usefulness and limitation of the method depends on whether the antibodies employed are able to bind to native epitopes. Since antibodies often are raised against recombinant proteins, epitopes of native, surface-exposed proteins may not be recognized. Nevertheless, the surface IFA method is a valuable tool for studying components of intact bacterial surfaces. This method can be applied not only for leptospires but also other spirochetes and gram-negative bacteria. For stronger conclusions regarding surface-exposure of OMPs, a comprehensive approach involving several cell localization methods is recommended 10.

An efficient method to assess surface-exposure of leptospiral proteins is described. The method is specifically designed to avoid disruption of the fragile outer membrane of leptospiral cells. This technique requires employment of several negative controls to assess the integrity of the outer membrane and specificity of antibody reaction.

Bacterial surface proteins are involved in direct contact with host cells and in uptake of nutrients from the environment 1. For this reason, cellular ...

Measurement of Factor V Activity in Human Plasma Using a Microplate Coagulation Assay

In response to injury, blood coagulation is activated and results in generation of the clotting protease, thrombin. Thrombin cleaves fibrinogen to fibrin which forms an insoluble clot that stops hemorrhage. Factor V (FV) in its activated form, FVa, is a critical cofactor for the protease FXa and accelerator of thrombin generation during fibrin clot formation as part of prothrombinase 1, 2. Manual FV assays have been described 3, 4, but they are time consuming and subjective. Automated FV assays have been reported 5-7, but the analyzer and reagents are expensive and generally provide only the clot time, not the rate and extent of fibrin formation. The microplate platform is preferred for measuring enzyme-catalyzed events because of convenience, time, cost, small volume, continuous monitoring, and high-throughput 8, 9. Microplate assays have been reported for clot lysis 10, platelet aggregation 11, and coagulation Factors 12, but not for FV activity in human plasma. The goal of the method was to develop a microplate assay that measures FV activity during fibrin formation in human plasma.
This novel microplate method outlines a simple, inexpensive, and rapid assay of FV activity in human plasma. The assay utilizes a kinetic microplate reader to monitor the absorbance change at 405nm during fibrin formation in human plasma (Figure 1) 13. The assay accurately measures the time, initial rate, and extent of fibrin clot formation. It requires only &mu;l quantities of plasma, is complete in 6 min, has high-throughput, is sensitive to 24-80pM FV, and measures the amount of unintentionally activated (1-stage activity) and thrombin-activated FV (2-stage activity) to obtain a complete assessment of its total functional activity (2-stage activity - 1-stage activity).
Disseminated intravascular coagulation (DIC) is an acquired coagulopathy that most often develops from pre-existing infections 14. DIC is associated with a poor prognosis and increases mortality above the pre-existing pathology 15. The assay was used to show that in 9 patients with DIC, the FV 1-stage, 2-stage, and total activities were decreased, on average, by 54%, 44%, and 42%, respectively, compared with normal pooled human reference plasma (NHP).
The FV microplate assay is easily adaptable to measure the activity of any coagulation factor. This assay will increase our understanding of FV biochemistry through a more accurate and complete measurement of its activity in research and clinical settings. This information will positively impact healthcare environments through earlier diagnosis and development of more effective treatments for coagulation disorders, such as DIC.

This study describes a novel microplate assay that measures FV coagulation activity during fibrin clot formation in human plasma which has not been reported previously. The method uses a kinetic microplate reader to continuously measure the change in absorbance at 405nm during fibrin clot formation in human plasma.

 In response to injury, blood coagulation is activated and results in generation of the clotting protease, thrombin. Thrombin cleaves fibrinogen to fibrin ...

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne relapsing fever (Borrelia spp.), Rocky Mountain Spotted fever (Rickettsia rickettsii), ehrlichiosis (Ehrlichia chaffeensis and E. equi), anaplasmosis (Anaplasma phagocytophilum), encephalitis (tick-borne encephalitis virus), babesiosis (Babesia spp.), Colorado tick fever (Coltivirus), and tularemia (Francisella tularensis) 1-8. To be properly transmitted into the host these infectious agents differentially regulate gene expression, interact with tick proteins, and migrate through the tick 3,9-13. For example, the Lyme disease agent, Borrelia burgdorferi, adapts through differential gene expression to the feast and famine stages of the tick's enzootic cycle 14,15. Furthermore, as an Ixodes tick consumes a bloodmeal Borrelia replicate and migrate from the midgut into the hemocoel, where they travel to the salivary glands and are transmitted into the host with the expelled saliva 9,16-19.
As a tick feeds the host typically responds with a strong hemostatic and innate immune response 11,13,20-22. Despite these host responses, I. scapularis can feed for several days because tick saliva contains proteins that are immunomodulatory, lytic agents, anticoagulants, and fibrinolysins to aid the tick feeding 3,11,20,21,23. The immunomodulatory activities possessed by tick saliva or salivary gland extract (SGE) facilitate transmission, proliferation, and dissemination of numerous tick-borne pathogens 3,20,24-27. To further understand how tick-borne infectious agents cause disease it is essential to dissect actively feeding ticks and collect tick saliva. This video protocol demonstrates dissection techniques for the collection of hemolymph and the removal of salivary glands from actively feeding I. scapularis nymphs after 48 and 72 hours post mouse placement. We also demonstrate saliva collection from an adult female I. scapularis tick.

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

The collection of infected tick hemolymph, salivary glands, and saliva is important to study how tick-borne pathogens cause disease. In this protocol we demonstrate how to collect hemolymph and salivary glands from feeding Ixodes scapularis nymphs. We also demonstrate saliva collection from female I. scapularis adults.

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne ...

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence factors, which can subvert and manipulate key host functions. The study of host/pathogen interactions is therefore extremely important to understand bacterial infections and develop alternative strategies to counter infectious diseases. This approach however, requires the development of new high-throughput assays for the unbiased, automated identification and characterization of bacterial virulence determinants. Here, we describe a method for the generation of a GFP-tagged mutant library by transposon mutagenesis and the development of high-content screening approaches for the simultaneous identification of multiple transposon-associated phenotypes. Our working model is the intracellular bacterial pathogen Coxiellaburnetii, the etiological agent of the zoonosis Q fever, which is associated with severe outbreaks with a consequent health and economic burden. The obligate intracellular nature of this pathogen has, until recently, severely hampered the identification of bacterial factors involved in host pathogen interactions, making of Coxiella the ideal model for the implementation of high-throughput/high-content approaches.

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Coxiella burnetii is an obligate intracellular Gram-negative bacterium responsible for the zoonotic disease Q fever. Here we describe methods for the generation of Coxiella fluorescent transposon mutants as well as the automated identification and analysis of the resulting internalization, replication and cytotoxic phenotypes.

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence ...

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

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

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

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

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

Visualizing the Effects of Sputum on Biofilm Development Using a Chambered Coverglass Model

Biofilms consist of groups of bacteria encased in a self-secreted matrix. They play an important role in industrial contamination as well as in the development and persistence of many health related infections. One of the most well described and studied biofilms in human disease occurs in chronic pulmonary infection of cystic fibrosis patients. When studying biofilms in the context of the host, many factors can impact biofilm formation and development. In order to identify how host factors may affect biofilm formation and development, we used a static chambered coverglass method to grow biofilms in the presence of host-derived factors in the form of sputum supernatants. Bacteria are seeded into chambers and exposed to sputum filtrates. Following 48 hr of growth, biofilms are stained with a commercial biofilm viability kit prior to confocal microscopy and analysis. Following image acquisition, biofilm properties can be assessed using different software platforms. This method allows us to visualize key properties of biofilm growth in presence of different substances including antibiotics.

This protocol describes the visualization of biofilm development following exposure to host-factors using a slide chamber model. This model allows for direct visualization of biofilm development as well as analysis of biofilm parameters using computer software programs.

Biofilms consist of groups of bacteria encased in a self-secreted matrix. They play an important role in industrial contamination as well as in the ...

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

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

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

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

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

A Multi-well Format Polyacrylamide-based Assay for Studying the Effect of Extracellular Matrix Stiffness on the Bacterial Infection of Adherent Cells

Extracellular matrix stiffness comprises one of the multiple environmental mechanical stimuli that are well known to influence cellular behavior, function, and fate in general. Although increasingly more adherent cell types&#39; responses to matrix stiffness have been characterized, how adherent cells&#39; susceptibility to bacterial infection depends on matrix stiffness is largely unknown, as is the effect of bacterial infection on the biomechanics of host cells. We hypothesize that the susceptibility of host endothelial cells to a bacterial infection depends on the stiffness of the matrix on which these cells reside, and that the infection of the host cells with bacteria will change their biomechanics. To test these two hypotheses, endothelial cells were used as model hosts and Listeria monocytogenes as a model pathogen. By developing a novel multi-well format assay, we show that the effect of matrix stiffness on infection of endothelial cells by L. monocytogenes can be quantitatively assessed through flow cytometry and immunostaining followed by microscopy. In addition, using traction force microscopy, the effect of L. monocytogenes infection on host endothelial cell biomechanics can be studied. The proposed method allows for the analysis of the effect of tissue-relevant mechanics on bacterial infection of adherent cells, which is a critical step towards understanding the biomechanical interactions between cells, their extracellular matrix, and pathogenic bacteria. This method is also applicable to a wide variety of other types of studies on cell biomechanics and response to substrate stiffness where it is important to be able to perform many replicates in parallel in each experiment.

We have developed a multi-well format polyacrylamide-based assay for probing the effect of extracellular matrix stiffness on bacterial infection of adherent cells. This assay is compatible with flow cytometry, immunostaining, and traction force microscopy, allowing for quantitative measurements of the biomechanical interactions between cells, their extracellular matrix, and pathogenic bacteria.