This work was funded by The MITRE Corporation through the MITRE Innovation Program (Project 25MSR621-CA). The authors would like to thank Mark Maybury, Ph.D., Richard Games, Ph.D., James Patton, Ellen Mac Garrigle, Ph.D., Carl Picconatto, Ph.D., and Caroline Gary for their support and review of this work.

1. Preparation of Bacterial and Peptidoglycan Substrates
Note: Whole cell bacterial substrates and purified peptidoglycans are used as enzyme substrates in both the microslide diffusion assay and the dye-release assay. These substrates require preparation prior to conducting the enzyme reactions. The following protocol describes their preparation.
<ol>
	<li>Whole Bacterial Cell Substrate
	<ol>
		<li>Produce bacterial cell substrate by inoculating 500 ml of nutrient broth with a 2 ml overnight culture of Bacillus subtilis 168 (American Type Culture Collection; ATCC 23857). Incubate at 30 &#176;C with shaking (125 rpm) until the culture reaches an exponential phase of growth, defined as a rapid growth phase resulting in the doubling of the bacterial culture. For the cultivation of Salmonella enterica subsp. enterica (ATCC 10708), use nutrient broth as a growth medium at 37 &#176;C with shaking (125 rpm).</li>
		<li>Heat-kill each culture by autoclaving for 10 min at 121 &#176;C under 3 atm of pressure.</li>
		<li>Harvest the heat-killed bacterial substrate by centrifugation for 20 min at 5,000 x g. Wash the pellet three times with Type 1 water and re-suspend in a minimal amount of water. In this study, suspend the substrates in 1,200 &#181;l.</li>
		<li>Aliquot 300 &#181;l of the bacterial cell substrates to 1.5 ml microfuge tubes and store at 20 &#176;C.</li>
	</ol>
	</li>
	<li>Purified Peptidoglycan Substrate
	<ol>
		<li>Purify peptidoglycan from the target substrate bacterium 7-10 or acquire from a vendor (see Materials and Equipment Table). Purify crude peptidoglycan preparations from accessory cell wall polymers.</li>
	</ol>
	</li>
</ol>
2. Qualitative Microslide Diffusion Assay [Modified from Lachica, et al. 11 ]
Note: The microslide diffusion assay is a qualitative method for detecting the presence of antimicrobial enzyme in a sample. As the enzyme diffuses through an agarose matrix containing substrate, a zone of clearing develops as the enzyme hydrolyzes the substrate. The following protocol describes the preparation and performance of the microslide diffusion assay for qualitatively detecting the presence of protein antimicrobials.
<ol>
	<li>Determine the concentration of the antimicrobial enzyme to be evaluated using a Bicinchoninic Acid (BCA) Protein Assay Kit according to manufacturer&#39;s protocol.</li>
	<li>Utilize phosphate-buffered saline (PBS) as an enzyme buffer; however, empirically determine the buffer appropriate for the given enzyme.</li>
	<li>Perform a serial dilution of the antimicrobial enzyme using phosphate-buffered saline (PBS) as a reaction buffer. The serial dilution used in these microslide diffusion assays produced final assay protein masses of 0 pg, 10 pg, 100 pg, 1 ng, 10 ng, 100 ng, 1 &#181;g, and 10 &#181;g per reaction volume.</li>
	<li>Dispense protein masses into individual microfuge tubes and adjust protein volumes to 20 &#181;l using PBS in preparation for addition to microslide reaction wells.</li>
	<li>Make a 0.5% agarose solution by dissolving 0.25 g of agarose in 50 ml of PBS. Heat the solution to a boil until the agarose is completely dissolved. Determine water lost during the melting process by weight and add back to the solution.</li>
	<li>Add 50 &#181;l of 10% sodium azide in water to the agarose solution and adjust the temperature of the solution to 50 &#176;C in a water bath. The sodium azide is added to inhibit contaminating bacterial growth during incubation of the microslide diffusion assay. If viable cells are used as substrate, omit the sodium azide from the agarose solution.</li>
	<li>Thaw the heat-killed bacterial substrate on ice.</li>
	<li>Re-suspend the bacterial substrate in 12 ml of the agarose solution to approximately match the turbidity of a 2.0 McFarland equivalence standard (see Materials Table). Compare the turbidities of the solutions by visualizing a black line on a white background through the solutions, adding bacterial substrate to the agarose until the approximate turbidity is achieved.</li>
	<li>Immediately pipet 3 ml of the agarose-substrate solution to each microslide (25 x 75 x 1 mm3).</li>
	<li>After slides solidify, punch three wells in the agarose-substrate layer of each microslide using a cork borer (4.8 mm diameter). Add 20 &#181;l of each adjusted protein serial dilution to their respective wells.</li>
	<li>Use PBS (20 &#181;l) or bovine serum albumin (5 &#181;g in 20 &#181;l PBS) in a negative control well. Use known bacteriolytic enzymes appropriate for the substrate, such as lysozyme, as a positive control.</li>
	<li>Incubate the slide in a humidity chamber at 37 &#176;C or the optimal activity temperature of the enzyme. The length of incubation depends upon the concentration and specific activity of the antimicrobial enzyme. A typical reaction time is overnight (approximately 16 hr).</li>
	<li>After incubation, observe the slides under indirect light and capture images of the developing assay using a digital single-lens reflex camera with a 60 mm lens at a focal distance of approximately 30 cm. 
	Note: Enzymatic activity of the protein antimicrobial for the given substrate is correlated with the development of a clear zone of hydrolysis, which forms around the well as the enzyme diffuses into the agarose.</li>
	<li>To qualify the enzymatic activity, observe the sizes of zones that form around each well. High enzymatic activity qualitatively relates to a larger zone, while low enzymatic activity qualitatively relates to a smaller zone.</li>
</ol>
3. Substrate Labeling - Remazol Brilliant Blue R Dye Labeling [Modified from Zhou 12 ]
Note: In the dye-release assay, the substrate is covalently linked to Remazol brilliant blue R dye. The following protocol describes the preparation of dyed enzyme substrates.
<ol>
	<li>Make a 250 mM sodium hydroxide (NaOH) solution by dissolving 1 g NaOH in 99 ml of Type I water to be used for making a 200 mM Remazol brilliant blue R dye (RBB) solution.</li>
	<li>Make a 200 mM RBB solution by dissolving 1.25 g RBB in 98.75 ml &#177; 1 ml of a fresh 250 mM NaOH solution (step 3.1).</li>
	<li>Re-suspend heat-killed bacterial cells at a concentration of 0.5 g wet weight in 30 ml of RBB solution. For purified peptidoglycan, re-suspend the peptidoglycan at a concentration of 0.3 g wet weight in 30 ml of RBB solution.</li>
	<li>Incubate the reaction mixture in an Erlenmeyer flask on a rotating platform for 6 hr at 37 &#176;C with gentle mixing.</li>
	<li>Transfer the reaction mixture to a 4 &#176;C incubator, and incubate for an additional 12 hr with gentle mixing.</li>
	<li>After incubation, harvest the dyed substrate by centrifugation at 3,000 x g for 30 min. Decant the dye solution from the substrate pellet.</li>
	<li>Remove non-covalently linked soluble dye from the substrates by washing the dye-labeled cells or peptidoglycan repeatedly (approximately 3-5 washes) with Type I water followed by centrifugation. With each water addition, re-suspend the pellet thoroughly. 
	Note: When unbound, soluble dye is no longer visible in the water wash after centrifugation. The substrate should be given one additional wash. Note that the substrate will remain blue, while the supernatant of the last wash will be clear.</li>
	<li>Store the dyed substrates suspended in a minimal amount of water at -20 &#176;C for later use.</li>
</ol>
4. Quantitative Dye-release Assay
Note: During the dye-release assay, the hydrolysis of RBB-dyed substrate in the enzymatic reaction releases dyed products into the reaction supernatant. Colorimetric measurement of the amount of dye released indicates the amount of enzymatic activity present within the sample for a given enzyme. The quantitative method allows the comparison of different enzymes across substrates and allows for variation of environmental conditions influencing the enzymatic reaction (e.g., temperature, salinity, and pH). The following protocol describes the preparation and performance of the dye-release assay for quantitatively detecting the enzymatic activity of protein antimicrobials.
<ol>
	<li>Preparation for the Dye-release Assay
	<ol>
		<li>Allow the frozen dyed substrate to return to room temperature and wash twice with assay buffer (PBS), which is empirically determined for the given enzyme.</li>
		<li>Calculate the volume of substrate suspension that is needed by multiplying the 200 &#181;l reaction volume by the number of dye-release assays to be performed.</li>
		<li>Prepare the substrate suspension by adding dyed substrate to the volume of assay buffer, determined in 4.1.2, until an optical density (OD) of 2.0 is achieved at 595 nm using a spectrophotometer. To remain within the functional limitations of the spectrophotometer, measure the 2.0 OD595 as 1.0 for a 1:1 dilution of the concentrated solution. Use assay buffer as a blank. 
		Note: The substrate turbidity for the reaction can be raised beyond 2.0 to match the activity levels of highly efficient enzymes.</li>
		<li>Suspend the protein antimicrobial to be evaluated in assay buffer at an estimated concentration of 1 mg/ml.</li>
		<li>Using the microplate assay of a BCA Protein Assay Kit according to manufacturer&#39;s protocol, determine the actual concentration of the protein sample to be assayed. Adjust the concentration of the stock suspension to 1 mg/ml using assay buffer.</li>
	</ol>
	</li>
	<li>Using the Dye-release Assay to Determine the Optimal Incubation Conditions for a Protein Antimicrobial
	<ol>
		<li>Dilute the stock protein antimicrobial suspension to 100 ng/&#181;l. This concentration gives a final reaction mass of 1 &#181;g of protein per volume (10 &#181;l) added to the assay reaction.</li>
		<li>Determine the thermal range to be assessed for the bacteriolytic protein. The thermal range in these assays included 5 &#176;C, 15 &#176;C, 25 &#176;C, 35 &#176;C, 45 &#176;C, 55 &#176;C, and 65 &#176;C.</li>
		<li>Perform the reaction assays in 0.5 ml microfuge tubes. For each thermal condition, add 10 &#181;l of the stock protein to 200 &#181;l of prepared substrate suspension.</li>
		<li>Incubate in a thermal cycler for 8 hr with mixing by inversion once per hour.</li>
		<li>Following incubation, arrest the reactions by adding 25 &#181;l of ethanol.</li>
		<li>Remove undigested, insoluble substrate by centrifugation at 3,000 x g for 2 min, and transfer 150 &#181;l of the reaction supernatant for each reaction mixture to a 96-well flat-bottom microplate, taking care not to disrupt the pellet of undigested substrate.</li>
		<li>Measure enzymatic activity of the protein antimicrobial for the given substrate by determining the amount of soluble RBB dye dissociated from the substrate after enzymatic hydrolysis. To quantify the enzymatic activity, measure the absorbance of the supernatant at 595 nm using a microplate spectrophotometer. Increased absorbance by the soluble dye released into the supernatant from the labeled substrate is a quantitative measurement of enzymatic activity. 
		Note: The change in absorbance is represented by activity units (AU), where 1 AU results in a 0.01 increase in the optical density of the dye-release reaction supernatant at 595 nm. A blank reaction, incubated with all reaction components except enzyme, is used to subtract any dye that releases from the RBB-labeled substrate due to the incubation. The optimal enzymatic temperature provides the peak activity unit measurement.</li>
		<li>Repeat steps 4.2.1 through 4.2.7 using various incubation buffers, to determine the optimal buffer conditions for the antimicrobial enzyme. In these assays, use PBS.</li>
	</ol>
	</li>
	<li>Using the Dye-release Assay to Determine the Minimal Active Concentration at the Optimal Incubation Temperature for a Protein Antimicrobial
	<ol>
		<li>Perform a serial dilution of the stock antimicrobial protein suspension using assay buffer. The dilution series range will vary with the activity of the antimicrobial enzyme. The serial dilution used in these assays produced final assay protein amounts of 0 pg, 10 pg, 100 pg, 1 ng, 10 ng, 100 ng, 1 &#181;g, and 10 &#181;g.</li>
		<li>Perform each reaction assay in a 48-well flat bottom microplate. For each assay condition, add 10 &#181;l of the respective protein suspension to 200 &#181;l of prepared substrate suspension. Adjust any variation in volume of the protein solution added to the reaction to 10 &#181;l using reaction buffer.</li>
		<li>Seal the microplate with plate sealing film to avoid volume variation due to evaporation. Incubate in a shaker incubator at the determined optimal incubation temperature for the given protein antimicrobial for 16 hr with shaking.</li>
		<li>Following incubation, arrest the reaction by adding 25 &#181;l of ethanol to each well and remove undigested substrate by centrifugation at 3,000 x g for 2 min.</li>
		<li>Transfer 150 &#181;l of the reaction supernatant for each reaction mixture to a 96-well flat-bottom microplate, taking care not to disrupt the pellet of the undigested substrate.</li>
		<li>To quantify the enzymatic activity, measure the absorbance of the supernatant at 595 nm using a microplate spectrophotometer. A blank reaction, incubated with all reaction components except enzyme, is used to subtract any dye that releases from the RBB-labeled substrate due to the incubation. Increased absorbance by the soluble dye released into the supernatant from the labeled substrate provides a quantitative measure to enzymatic activity. 
		Note: The change in absorbance is represented by activity units (AU), where 1 AU results in a 0.01 increase in the optical density of the dye-release reaction supernatant at 595 nm.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.&#160;The MITRE Corporation&#160;is a not-for-profit company that operates multiple federally funded research and development centers (FFRDCs).

The microslide diffusion assay and the dye-release assay provide advantages for detecting and characterizing previously unidentified protein antimicrobials. These rapid and sensitive methods allow high-throughput screening of organism source libraries for the identification of enzymes of interest prior to investing greater research resources. As high profile target enzymes are identified, variations of the detection assays can be used to rapidly detect the enzyme or to determine biochemical characteristics of the enzyme. For example, during a multistep protein purification scheme, the microslide diffusion assay can rapidly detect fractions containing the enzyme of interest. In addition, reaction optima for the enzyme can be determined by altering the reaction buffers and incubation temperatures in the dye-release assay.
The range of enzyme mass required for detection in the microslide diffusion assay is a function of enzyme characteristics. Detection limitations of the assay generally require the use of higher amounts of enzyme than the dye-release assay. In this study, comparison of the detection limits of the microslide diffusion assay (Figure 1) to the dye-release assay (Figure 4) illustrated that the microslide diffusion assay is a less sensitive technique than the dye-release assay. When enzyme concentration is not a concern, the microslide assay allows for rapid initial screening of libraries for the production of protein antimicrobials against target substrates with low labor and equipment demands. Although requiring more effort and equipment, the dye-release assay is more sensitive and reproducible, yielding quantitative results that allow comparison between dye-release assays of the same or different enzymes for a given substrate. The two methods are easily performed in most microbiological laboratories with no need for highly specialized instrumentation. In the representative assays, the control enzyme, &#945;-chymotrypsin, was detected at amounts as low as 100 ng (Figure 5) using the dye-release assay, while the minimal amount detected in the microslide diffusion assay was 1 &#181;g (data not shown).
Providing utility as a qualitative detection assay for antimicrobial enzymes, a number of variations of the diffusion assay have been reported 11,13. In reviewing these assays, the microslide diffusion assay was developed for its relatively high sensitivity as an initial screen and for its ease of reaction observance. Modified from the work of Lachica et al. 11, in which agarose overlays were used to detect the production of nucleases by strains of Staphylococcus aureus, the microslide diffusion assay operates similarly to the radial immunodiffusion method 14 as the protein diffuses from the well into the agarose containing the substrate. The radial immunodiffusion method halts the expansion of the zone of immunoprecipitation when the system reaches a complex formation equilibrium between the diffusing antigen and antibody within the agarose. In contrast, the substrate of the microslide diffusion assay is initially digested by the diffusing protein antimicrobial in a continually expanding zone of lysis surrounding the well. The increasing diameter of the zone continues until the enzyme loses activity or the substrate is exhausted. The rate of the production of the zone of lysis is quickened as the purity, and thus the specific activity, of the enzyme or the concentration of the enzyme added to the well increases.
For quantitatively assessing the enzymatic activity of protein antimicrobials against heat-killed B. subtilis, the dye-release assay was chosen. Colorimetric assays using Remazol brilliant blue R dye (RBB)-labeled substrates allow versatile and sensitive evaluation of protein antimicrobial activity. Enzymatic hydrolysis of RBB-labeled substrate results in the release of soluble blue products that can be readily measured by a spectrophotometer at 595 nm. Several variations of the RBB dye-release assay, developed to characterize other protein antimicrobial enzymes, show the flexibility of this assay for application with other substrates. For example, in determining the effects of lysostaphin bacteriolytic activity, Zhou, et al., reported a dye-release assay using RBB-dyed staphylococcal cells and staphylococcal peptidoglycan as substrates 12. In a modification of the application of this RBB dye-release assay, RBB-labeled Micrococcus luteus substrate was proposed for use as a rapid and sensitive means to diagnose and screen for diseases such as bacterial infection and the presence of a malignant carcinoma, which can be correlated to human lysozyme expression levels in blood serum 15. In addition, RBB-labeled bacterial cell substrates have also been used in a zymogram method for the detection of lysozyme 16.
We demonstrate the lowered detection limit, convenience, and reproducibility of the dye-release assay, allowing for rapid library screening for enzyme production. Modifications of the assay allow for downstream quantitative biochemical characterization assays, including effects of temperature, pH, and salinity as well as the effects of other proteolytic enzymes on the activity of antimicrobial proteins 6. In addition, the quantitative dye-release assay can be used to compare the activities of multiple enzymes for a given substrate. The RBB dye-release assay has reported utility for enzymes with activity against polysaccharides in addition to the characterization and detection of protein antimicrobial agents. Pettersson and Eriksson reported an assay for detecting endoglycanase activity using amorphous, RBB-dyed polysaccharide beads of cellulose, xylan, mannan, and chitin 17. In an expansion of these findings, a sensitive method for detection of chitinase enzymes produced by Bacillus thuringiensis Bt-107 was developed using colloidal chitin labeled with RBB 18. From these and other studies, commercially available sources of RBB-dyed substrates have emerged for use in detection of glycolytic activity, including those against keratin, amylopectin, amylose, glycogen, laminarin, d-xylan, Azo-barley glucan, and starch.
In this study, we demonstrate the utility of the dye-release assay in a microplate format, reducing the volume of RBB-labeled substrate and enzyme required to produce the colorimetric result. As illustrated in Figure 4 and Figure 5, the microplate dye-release assay provides a lower detection limit than the microslide diffusion assay for enzymatic activity detection. With regard to rapid use, conservation of labor, and ease of interpretation, the microslide diffusion assay provides utility in initial high-throughput screens for the presence of antimicrobial activity as well as the indication of antimicrobial protein presence in downstream protein isolation and purification steps. The combination of these two assays complement each other in the initial screening and ultimate characterization of novel protein antimicrobials

Antimicrobial agents play an essential role in targeting a wide range of microorganisms such as viruses, fungi, and bacteria. The antimicrobial activities produced by various bacteria range in target specificity from broad-spectrum to species-specific, including clinically relevant bacterial pathogens 1. In nature, protein antimicrobial compounds like bacteriocins and lysins represent a microbial arsenal of tools for self-defense, replication, and nutrient acquisition 2,3. Serving as mediators of population dynamics within microbial communities, protein antimicrobials provide a competitive advantage to the producing organism over nonproducing, sensitive species 4. As recombinant enzymes and probiotics, these proteins also have an economic and societal importance as the need for new sources of pathogen control increases with each new expansion of antimicrobial resistance within the bacterial population 5.
The first evaluations of newly discovered protein antimicrobials include determining the basic physical characteristics that are inherent to the enzyme. Methods for rapid screening of potential antimicrobials allow selection of candidates with hydrolytic activities against the desired target substrates. The microslide diffusion assay and the quantitative dye-release assay allow for the use of a variety of substrates in the detection of enzymatic hydrolysis. For protein antimicrobial enzymes with activity against the bacterial cell wall, the versatility of these assays allow rapid and effective, qualitative and quantitative screening against viable bacterial cells (microslide assay only), heat-killed whole bacterial cell substrates, or purified peptidoglycan from the target bacterium. These assays have utility in antimicrobial enzyme discovery for use in fields such as alternative therapeutics, food supplements, and inhibitors of biofilm production.
The microslide diffusion assay and the dye-release assay are demonstrated methods for detection and characterization of novel protein antimicrobials. The microslide diffusion assay provides a relative (qualitative) estimate of antimicrobial activity and allows for minimal inference of enzyme concentration and activity. Using this method, activity is readily visualized as the development of a zone of lysis with the absence of activity resulting in a lack of zone formation. The rate of zone development and the zone diameter are indicative of the amount and purity of the enzyme present in the sample; however, this rate and the quality of the substrate clearing within the zone can also be affected by the presence of contaminants, the completeness of the hydrolysis reaction, and the type of substrate. Interfering contaminants can affect the rate of enzyme diffusion from the well, while incomplete hydrolysis or variation of the substrate type can result in a turbid zone of lysis 6.
The dye-release assay quantitatively assesses the amount of covalently-linked Remazol brilliant blue R dye products released into the reaction supernatant from the enzymatically hydrolyzed substrate. The method has only slight variability associated with a difference in substrate, thus allowing greater sensitivity for detection of hydrolysis as compared to the microslide diffusion assay. These properties allow the method to have utility in the characterization of biochemical properties of the enzyme and in the standardization of the assay for comparison between different antimicrobial enzymes.
In this protocol, we provide methods to perform the basic microslide diffusion assay and dye-release assay, while demonstrating the scale of detection limits and variability for each. The assays are used to perform basic evaluations of an unknown protein antimicrobial purified from the culture supernatant of an uncharacterized soil bacterial isolate. Observed activities against bacterial cell substrates within the assays allow comparison of reaction activities between a previously uncharacterized protein antimicrobial and &#945;-chymotrypsin, a control proteolytic enzymes. These protocols provide a foundation for the application of the two techniques that can be expanded and modified to accommodate varied applications.

<table><tbody><tr><td>48-well flat-bottom microplate with low evaporation lid</td><td>Becton Dickinson</td><td>3078</td><td></td></tr><tr><td>96-well flat-bottom microplate (Costar 3595)</td><td>Corning, Inc.</td><td>3595</td><td></td></tr><tr><td>agarose</td><td>Fisher Scientific</td><td>BP162-100</td><td></td></tr><tr><td>&amp;alpha;-chymotrypsin</td><td>MP Biomedicals</td><td>215227205</td><td></td></tr><tr><td>Bacillus subtilis 168</td><td>American Type Culture Collection</td><td>23857</td><td></td></tr><tr><td>C1000 Touch Thermal Cycler</td><td>Bio-Rad</td><td></td><td></td></tr><tr><td>cork borer</td><td>Humboldt</td><td>H-9661</td><td></td></tr><tr><td>lysozyme</td><td>Sigma-Aldrich</td><td>L6876-1G</td><td></td></tr><tr><td>McFarland equivalence standard (2.0)</td><td>Fisher Scientific</td><td>R20412</td><td></td></tr><tr><td>microslides</td><td>Fisher Scientific</td><td>22-38-103</td><td></td></tr><tr><td>nutrient broth</td><td>Fisher Scientific</td><td>S25959B</td><td></td></tr><tr><td>peptidoglycan of &lt;em&gt;Bacillus subtilis&lt;/em&gt; 168</td><td>Sigma-Aldrich</td><td>69554-10MG-F</td><td></td></tr><tr><td>phosphate-buffered saline (1x)</td><td>Teknova</td><td>P0200</td><td></td></tr><tr><td>Pierce BCA Protein Assay Kit</td><td>Life Technologies</td><td>23227</td><td></td></tr><tr><td>Synergy HT microplate spectrophotometer</td><td>BioTek Instruments, Inc.</td><td></td><td></td></tr><tr><td>Remazol brilliant blue R dye (RBB)</td><td>Sigma-Aldrich</td><td>R8001</td><td></td></tr><tr><td>Salmonella enterica subsp. enterica</td><td>American Type Culture Collection</td><td>10708</td><td></td></tr><tr><td>sodium azide</td><td>Fisher Scientific</td><td>S227-100</td><td></td></tr><tr><td>sodium hydroxide (NaOH)</td><td>Fisher Scientific</td><td>S318-500</td><td></td></tr><tr><td>Titer Tops pre-cut adhesive polyethylene film</td><td>Diversified Biotechnology</td><td>T-TOPS-50</td><td></td></tr><tr><td>UltraPure distilled water</td><td>Invitrogen</td><td>10977-015</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>agarose solution&lt;br/&gt; 50 ml PBS&lt;br/&gt; 0.25 g agarose</td><td></td><td></td><td>Add 10% sodium azide to the molten PBS/agarose solution</td></tr><tr><td>nutrient broth medium&lt;br/&gt; 4 g nutrient broth&lt;br/&gt; 500 ml Type I water</td><td></td><td></td><td></td></tr><tr><td>250 mM sodium hydroxide (NaOH) solution&lt;br/&gt; 1 g NaOH&lt;br/&gt; 99 ml Type I water</td><td></td><td></td><td></td></tr><tr><td>200 mM Remazol brilliant blue R dye (RBB)&lt;br/&gt; 1.25 g RBB&lt;br/&gt; 98.75 ml 250 mM NaOH solution</td><td></td><td></td><td></td></tr></tbody></table>

qualitative and quantitative assays for detection and characterization of protein antimicrobials

Initial evaluations of large microbial libraries for potential producers of novel antimicrobial proteins require both qualitative and quantitative methods to screen for target enzymes prior to investing greater research effort and resources. The goal of this protocol is to demonstrate two complementary assays for conducting these initial evaluations. The microslide diffusion assay provides an initial or simple detection screen to enable the qualitative and rapid assessment of proteolytic activity against an array of both viable and heat-killed bacterial target substrates. As a counterpart, the increased sensitivity and reproducibility of the dye-release assay provides a quantitative platform for evaluating and comparing environmental influences affecting the hydrolytic activity of protein antimicrobials. The ability to label specific heat-killed cell culture substrates with Remazol brilliant blue R dye expands this capability to tailor the dye-release assay to characterize enzymatic activity of interest.

The microslide diffusion assay and the quantitative dye-release assay are effective methods for screening and measuring initial investigations of new protein antimicrobials. Each assay has advantages and limitations; however, when performed in conjunction, they allow rapid initial screening and basic characterization of an antimicrobial.
The microslide diffusion assay efficiently allows for the rapid screening of microbial libraries, producing protein antimicrobials. When the enzyme concentration levels are of concern, sensitivity of detection constraints may limit the assay, requiring a greater amount of enzyme to be added to the reaction well than the dye-release assay. As illustrated in Figure 1, the qualitative properties of the assay allow the observer to compare relative enzyme amounts present within each well.
In the developing zone of lysis, illustrated in Figure 1A (25 &#181;g of enzyme), the leading edge of the zone displays turbid or incomplete lysis of the substrate, while the zone closest to the well displays a more complete substrate lysis. This phenomenon, a product of the diminishing concentration of the diffusing enzyme at the leading edge, complicates the accurate measurement of the zone diameter. As the wells B (15 &#181;g), C (10 &#181;g), D (5 &#181;g), E (1.0 &#181;g), and F (0.1 &#181;g) are observed in Figure 1, the diameter of the zone of complete lysis dissipates to extinction correlating to the reduced amount of enzyme. For condition F (0.1 &#181;g), the enzyme concentration within the agarose is below the limit that will allow the diffusing enzyme to be visualized as it moves through the agarose, hydrolyzing the substrate. The microslide diffusion assay was also run using purified Bacillus subtilis peptidoglycan as substrate (Figure 2). While the zone is less defined than those observed with whole cell Salmonella enterica due to the reluctance of the peptidoglycan to evenly suspend in the agarose, the hydrolysis of the peptidoglycan by the unknown antimicrobial enzyme is apparent.
The dye-release assay is a more sensitive and versatile assay than the microslide diffusion assay, allowing a lower detection limit and variation of environmental factors affecting the enzyme reaction. In the representative assays, temperature was varied to determine the optimal temperature for the antimicrobial enzyme, determined to be 35 &#176;C in PBS (Figure 3). This optimum is seen clearly in the reaction supernatants as increased amounts of blue color (Figure 3B) as well as represented in activity units derived from absorbance measurements at 595 nm (Figure 3A). The versatility of the dye-release assay allows the researcher to vary not only the temperatures but also the reaction buffer and buffer components to rapidly determine optimum incubation conditions for a given enzyme.
The activity level of the unknown antimicrobial enzyme (Figure 4) and the &#945;-chymotrypsin control enzyme (Figure 5) were measured at the determined optimum incubation temperature of 35 &#176;C in PBS against RBB-labeled Bacillus subtilis heat-killed substrate. Comparison of results from Figure 4 and Figure 5 indicates that the unknown antimicrobial enzyme has almost twice the affinity for the B. subtilis substrate. In addition, the &#945;-chymotrypsin control did not completely digest the heat-killed B. subtilis substrate within the well (data not shown). The activity of the &#945;-chymotrypsin control begins to plateau around 0.3 &#181;g as compared to the continued rise in activity units across all enzyme amounts for the unknown antimicrobial enzyme (Figure 4 and Figure 5). This may indicate that the unknown enzyme has a greater sustained activity or that there are a greater number of cleavage sites available to the enzyme within the B. subtilis substrate.
<img alt="Figure 1" src="/files/ftp_upload/53819/53819fig1.jpg" /> 
Figure 1: Enzyme Activity against Salmonella enterica Whole Cell Substrate. The microslide diffusion assay was used to qualitatively evaluate the activity of an unknown protein antimicrobial against heat-killed Salmonella enterica subsp. enterica (ATCC 10708). The protein masses of the unknown antimicrobial suspended in phosphate-buffered saline (PBS) that were added to the respective wells of the slides included 25 &#181;g (well A), 15 &#181;g (well B), 10 &#181;g (well C), 5 &#181;g (well D), 1 &#181;g (well E), and 0.1 &#181;g (well F). PBS alone was used for the negative controls of the assays. Zones of lysis were imaged after a 6 hr incubation. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53819/53819fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53819/53819fig2.jpg" /> 
Figure 2: Enzyme Activity Against Bacillus subtilis Peptidoglycan Cell Wall. The microslide diffusion assay was used to qualitatively evaluate the activity of an unknown protein antimicrobial against peptidoglycan of Bacillus subtilis 168. Suspended in 20 &#181;l of PBS, 10 &#181;g of the unknown antimicrobial was added to well A of the microslides. PBS alone was used as a negative control for the assay (well B). The zone of lysis was imaged after a 6-hour incubation at 37 &#176;C. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53819/53819fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53819/53819fig3.jpg" /> 
Figure 3: Optimal Reaction Temperature for a Protein Antimicrobial. The versatility of the dye-release assay allows the variation of physical and chemical parameters, such as incubation temperatures and buffers, to determine their influences on the activity of the enzyme of interest 6. As illustrated in this figure, the activity of the unknown protein antimicrobial (1 &#181;g) was evaluated in PBS against RBB-labeled heat-killed Bacillus subtilis under a range of incubation temperatures. Displayed as activity units (AU) in (A), the absorbance of RBB-bound products released into the supernatant after hydrolysis was measured at 595 nm using a microplate spectrophotometer. At each temperature condition, reactions with no enzyme added were used to subtract the effects of any released dye that resulted from incubation at the various temperatures. The reaction supernatants corresponding to each incubation temperature were imaged prior to absorbance measurement (B). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53819/53819fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53819/53819fig4.jpg" /> 
Figure 4: Activity Range for a Protein Antimicrobial. The activity range for the unknown protein antimicrobial was measured as activity units after overnight incubations for 0, 100, 200, 300, 400, 500, 600, 700, 800, 900, and 1,000 ng, as measured by the BCA protein assay (A). For these reactions, the RBB-labeled B. subtilis substrate level was increased to give an optical density of 5.0 at 595 nm to ensure that the reaction with the highest amount of enzyme (1,000 ng) did not completely hydrolyze all available substrate within the reaction incubation period. Control reactions with no enzyme added were used to subtract the effects of any released dye caused by the incubation alone. The reaction supernatants corresponding to each enzyme amount were imaged prior to absorbance measurements (B). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53819/53819fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/53819/53819fig5.jpg" /> 
Figure 5: Activity Range for &#945;-Chymotrypsin. The activity range for &#945;-chymotrypsin was measured as activity units after overnight incubations for 0, 100, 200, 300, 400, 500, 600, 700, 800, 900, and 1,000 ng of enzyme (A). For these reactions, the RBB-labeled B. subtilis substrate level was increased to give an optical density of 5.0 at 595 nm to ensure that the reaction with the highest amount of enzyme (1,000 ng) did not completely hydrolyze all available substrate within the reaction incubation period. Control reactions with no enzyme added were used to subtract the effects of any released dye caused by the incubation alone. The reaction supernatants corresponding to each enzyme amount were imaged prior to absorbance measurements (B). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53819/53819fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Qualitative and Quantitative Assays for Detection and Characterization of Protein Antimicrobials at JoVE.com

Qualitative and Quantitative Assays for Detection and Characterization of Protein Antimicrobials

Here, we present protocols to rapidly screen and characterize enzymes for antimicrobial activity. The microslide diffusion assay and the dye-release assay utilize target bacterial substrates for qualitative and quantitative enzymatic activity evaluation.

Initial evaluations of large microbial libraries for potential producers of novel antimicrobial proteins require both qualitative and quantitative methods ...

qualitative-quantitative-assays-for-detection-characterization

Universidad de Cartagena UDEC

Research

JoVE Journal

Immunology and Infection

18.4K Views.  The MITRE Corporation. Here, we present protocols to rapidly screen and characterize enzymes for antimicrobial activity. The microslide diffusion assay and the dye-release assay utilize target bacterial substrates for qualitative and quantitative enzymatic activity evaluation.

Video: Qualitative and Quantitative Assays for Detection and Characterization of Protein Antimicrobials

An HS-MRM Assay for the Quantification of Host-cell Proteins in Protein Biopharmaceuticals by Liquid Chromatography Ion Mobility QTOF Mass Spectrometry

The analysis of low-level (1-100 ppm) protein impurities (e.g., host-cell proteins (HCPs)) in protein biotherapeutics is a challenging assay requiring high sensitivity and a wide dynamic range. Mass spectrometry-based quantification assays for proteins typically involve protein digestion followed by the selective reaction monitoring/multiple reaction monitoring (SRM/MRM) quantification of peptides using a low-resolution (Rs ~1,000) tandem quadrupole mass spectrometer. One of the limitations of this approach is the interference phenomenon observed when the peptide of interest has the "same" precursor and fragment mass (in terms of m/z values) as other co-eluting peptides present in the sample (within a 1-Da window). To avoid this phenomenon, we propose an alternative mass spectrometric approach, a high selectivity (HS) MRM assay that combines the ion mobility separation of peptide precursors with the high-resolution (Rs ~30,000) MS detection of peptide fragments. We explored the capabilities of this approach to quantify low-abundance peptide standards spiked in a monoclonal antibody (mAb) digest and demonstrated that it has the sensitivity and dynamic range (at least 3 orders of magnitude) typically achieved in HCP analysis. All six peptide standards were detected at concentrations as low as 0.1 nM (1 femtomole loaded on a 2.1-mm ID chromatographic column) in the presence of a high-abundance peptide background (2 &#181;g of a mAb digest loaded on-column). When considering the MW of rabbit phosphorylase (97.2 kDa), from which the spiked peptides were derived, the LOQ of this assay is lower than 50 ppm. Relative standard deviations (RSD) of peak areas (n = 4 replicates) were less than 15% across the entire concentration range investigated (0.1-100 nM or 1-1,000 ppm) in this study.

Here, we describe a chromatographic assay coupled with the ion mobility separation of peptide precursors followed by the high-resolution (~30,000) MS-detection of peptide fragments for the quantification of spiked peptide standards in a monoclonal antibody digest.

The analysis of low-level (1-100 ppm) protein impurities (e.g., host-cell proteins (HCPs)) in protein biotherapeutics is a challenging assay requiring ...

Chemistry

Nanomechanics of Drug-target Interactions and Antibacterial Resistance Detection

The cantilever sensor, which acts as a transducer of reactions between model bacterial cell wall matrix immobilized on its surface and antibiotic drugs in solution, has shown considerable potential in biochemical sensing applications with unprecedented sensitivity and specificity1-5. The drug-target interactions generate surface stress, causing the cantilever to bend, and the signal can be analyzed optically when it is illuminated by a laser. The change in surface stress measured with nano-scale precision allows disruptions of the biomechanics of model bacterial cell wall targets to be tracked in real time. Despite offering considerable advantages, multiple cantilever sensor arrays have never been applied in quantifying drug-target binding interactions.
Here, we report on the use of silicon multiple cantilever arrays coated with alkanethiol self-assembled monolayers mimicking bacterial cell wall matrix to quantitatively study antibiotic binding interactions. To understand the impact of vancomycin on the mechanics of bacterial cell wall structures1,6,7.&#160;We developed a new model1 which proposes that cantilever bending can be described by two independent factors; i) namely a chemical factor, which is given by a classical Langmuir adsorption isotherm, from which we calculate the thermodynamic equilibrium dissociation constant (Kd) and ii) a geometrical factor, essentially a measure of how bacterial peptide receptors are distributed on the cantilever surface. The surface distribution of peptide receptors (p) is used to investigate the dependence of geometry and ligand loading. It is shown that a threshold value of p ~10% is critical to sensing applications. Below which there is no detectable bending signal while above this value, the bending signal increases almost linearly,&#160;revealing that stress is a product of a local chemical binding factor and a geometrical factor combined by the mechanical connectivity of reacted regions and provides a new paradigm for design of powerful agents to combat superbug infections.

Acquired resistance to antibiotics is a major public healthcare problem and is presently ranked by the WHO as one of the greatest threats to human life. Here we describe the use of cantilever technology to quantify antibacterial resistance, critical to the discovery of novel and powerful agents against multidrug resistant bacteria.

The cantilever sensor, which acts as a transducer of reactions between model bacterial cell wall matrix immobilized on its surface and antibiotic drugs in ...

Isolation and Identification of Waterborne Antibiotic-Resistant Bacteria and Molecular Characterization of their Antibiotic Resistance Genes

The development and spread of antibiotic resistance (AR) through microbiota associated with freshwater bodies is a major global health concern. In the present study, freshwater samples were collected and analyzed with respect to the total bacterial diversity and AR genes (ARGs) using both conventional culture-based techniques and a high-throughput culture-independent metagenomic approach. This paper presents a systematic protocol for the enumeration of the total and antibiotic-resistant culturable bacteria from freshwater samples and the determination of phenotypic and genotypic resistance in the culturable isolates. Further, we report the use of whole metagenomic analysis of the total metagenomic DNA extracted from the freshwater sample for the identification of the overall bacterial diversity, including non-culturable bacteria, and the identification of the total pool of different ARGs (resistome) in the water body. Following these detailed protocols, we observed a high antibiotic-resistant bacteria load in the range of 9.6 &#215; 105-1.2 &#215; 109 CFU/mL. Most isolates were resistant to the multiple tested antibiotics, including cefotaxime, ampicillin, levofloxacin, chloramphenicol, ceftriaxone, gentamicin, neomycin, trimethoprim, and ciprofloxacin, with multiple antibiotic resistance (MAR) indexes of &#8805;0.2, indicating high levels of resistance in the isolates. The 16S rRNA sequencing identified potential human pathogens, such as Klebsiella pneumoniae,&#160;and opportunistic bacteria, such as Comamonas spp., Micrococcus spp., Arthrobacter spp., and Aeromonas spp. The molecular characterization of the isolates showed the presence of various ARGs, such as blaTEM, blaCTX-M (&#946;-lactams), aadA, aac (6&#39;)-Ib (aminoglycosides), and dfr1 (trimethoprims), which was also confirmed by the whole metagenomic DNA analysis. A high prevalence of other ARGs encoding for antibiotic efflux pumps-mtrA, macB, mdtA, acrD, &#946;-lactamases-SMB-1, VIM-20, ccrA, ampC, blaZ, the&#160;chloramphenicol acetyltransferase gene catB10, and&#160;the rifampicin resistance gene rphB-was also detected in the metagenomic DNA. With the help of the protocols discussed in this study, we confirmed the presence of waterborne MAR bacteria with diverse AR phenotypic and genotypic traits. Thus, whole metagenomic DNA analysis can be used as a complementary technique to conventional culture-based techniques to determine the overall AR status of a water body.

Here, we present a detailed protocol for the isolation and identification of antibiotic-resistant bacteria from water and the molecular characterization of their antibiotic resistance genes (ARGs). The use of culture-based and non-culture-based (metagenomic analysis) techniques provides complete information about the total bacterial diversity and the total pool of different ARGs present in freshwaters from Mumbai, India.

The development and spread of antibiotic resistance (AR) through microbiota associated with freshwater bodies is a major global health concern. In the ...

Environment

High Throughput, Real-time, Dual-readout Testing of Intracellular Antimicrobial Activity and Eukaryotic Cell Cytotoxicity

Traditional measures of intracellular antimicrobial activity and eukaryotic cell cytotoxicity rely on endpoint assays. Such endpoint assays require several additional experimental steps prior to readout, such as cell lysis, colony forming unit determination, or reagent addition. When performing thousands of assays, for example, during high-throughput screening, the downstream effort required for these types of assays is considerable. Therefore, to facilitate high-throughput antimicrobial discovery, we developed a real-time assay to simultaneously identify inhibitors of intracellular bacterial growth and assess eukaryotic cell cytotoxicity. Specifically, real-time intracellular bacterial growth detection was enabled by marking bacterial screening strains with either a bacterial lux operon (1st generation assay) or fluorescent protein reporters (2nd generation, orthogonal assay). A non-toxic, cell membrane-impermeant, nucleic acid-binding dye was also added during initial infection of macrophages. These dyes are excluded from viable cells. However, non-viable host cells lose membrane integrity permitting entry and fluorescent labeling of nuclear DNA (deoxyribonucleic acid). Notably, DNA binding is associated with a large increase in fluorescent quantum yield that provides a solution-based readout of host cell death. We have used this combined assay to perform a high-throughput screen in microplate format, and to assess intracellular growth and cytotoxicity by microscopy. Notably, antimicrobials may demonstrate synergy in which the combined effect of two or more antimicrobials when applied together is greater than when applied separately. Testing for in vitro synergy against intracellular pathogens is normally a prodigious task as combinatorial permutations of antibiotics at different concentrations must be assessed. However, we found that our real-time assay combined with automated, digital dispensing technology permitted facile synergy testing. Using these approaches, we were able to systematically survey action of a large number of antimicrobials alone and in combination against the intracellular pathogen, Legionella pneumophila.

A high throughput, real-time assay was developed to simultaneously identify (1) eukaryotic cell-penetrant antimicrobials targeting an intracellular bacterial pathogen, and (2) assess eukaryotic cell cytotoxicity. A variation on the same technology was thereafter combined with digital dispensing technology to enable facile, high-resolution, dose-response, and two- and three-dimensional synergy studies.

Traditional measures of intracellular antimicrobial activity and eukaryotic cell cytotoxicity rely on endpoint assays. Such endpoint assays require ...

Use of the Soft-agar Overlay Technique to Screen for Bacterially Produced Inhibitory Compounds

The soft-agar overlay technique was originally developed over 70 years ago and has been widely used in several areas of microbiological research, including work with bacteriophages and bacteriocins, proteinaceous antibacterial agents. This approach is relatively inexpensive, with minimal resource requirements. This technique consists of spotting supernatant from a donor strain (potentially harboring a toxic compound(s)) onto a solidified soft agar overlay that is seeded with a bacterial test strain (potentially sensitive to the toxic compound(s)). We utilized this technique to screen a library of Pseudomonas syringae strains for intraspecific killing. By combining this approach with a precipitation step and targeted gene deletions, multiple toxic compounds produced by the same strain can be differentiated. The two antagonistic agents commonly recovered using this technique are bacteriophages and bacteriocins. These two agents can be differentiated using two simple additional tests. Performing a serial dilution on a supernatant containing bacteriophage will result in individual plaques becoming less in number with greater dilution, whereas serial dilution of a supernatant containing bacteriocin will result a clearing zone that becomes uniformly more turbid with greater dilution. Additionally, a bacteriophage will produce a clearing zone when spotted onto a fresh soft agar overlay seeded with the same strain, whereas a bacteriocin will not produce a clearing zone when transferred to a fresh soft agar lawn, owing to the dilution of the bacteriocin.

We describe a simple method for screening bacterial cultures for the production of compounds inhibitory towards other bacteria.

The soft-agar overlay technique was originally developed over 70 years ago and has been widely used in several areas of microbiological research, ...

A Protein Microarray Assay for Serological Determination of Antigen-specific Antibody Responses Following Clostridium difficile Infection

We provide a detailed overview of a novel high-throughput protein microarray assay for the determination of anti-Clostridium difficile antibody levels in human sera and in separate preparations of polyclonal intravenous immunoglobulin (IVIg). The protocol describes the methodological steps involved in sample preparation, printing of arrays, assay procedure, and data analysis. In addition, this protocol could be further developed to incorporate diverse clinical samples including plasma and cell culture supernatants. We show how protein microarray can be used to determine a combination of isotype (IgG, IgA, IgM), subclass (IgG1, IgG2, IgG3, IgG4, IgA1, IgA2), and strain-specific antibodies to highly purified whole C. difficile toxins A and B (toxinotype 0, strain VPI 10463, ribotype 087), toxin B from a C. difficile toxin-B-only expressing strain (CCUG 20309), a precursor form of a B fragment of binary toxin, pCDTb, ribotype-specific whole surface layer proteins (SLPs; 001, 002, 027), and control proteins (tetanus toxoid and Candida albicans). During the experiment, microarrays are probed with sera from individuals with C. difficile infection (CDI), individuals with cystic fibrosis (CF) without diarrhea, healthy controls (HC), and from individuals pre- and post-IVIg therapy for the treatment of CDI, combined immunodeficiency disorder, and chronic inflammatory demyelinating polyradiculopathy. We encounter significant differences in toxin neutralization efficacies and multi-isotype specific antibody levels between patient groups, commercial preparations of IVIg, and sera before and following IVIg administration. Also, there is a significant correlation between microarray and enzyme-linked immunosorbent assay (ELISA) for antitoxin IgG levels in serum samples. These results suggest that microarray could become a promising tool for profiling antibody responses to C.difficile antigens in vaccinated or infected humans. With further refinement of antigen panels and a reduction in production costs, we anticipate that microarray technology may help optimize and select the most clinically useful immunotherapies for C. difficile infection in a patient-specific manner.

A Protein Microarray Assay for Serological Determination of Antigen-specific Antibody Responses Following Clostridium difficile Infection

This article describes a simple protein microarray method for profiling humoral immune responses to a 7-plex panel of highly purified Clostridium difficile antigens in human sera. The protocol can be extended for the determination of specific antibody responses in preparations of polyclonal intravenous immunoglobulin.

We provide a detailed overview of a novel high-throughput protein microarray assay for the determination of anti-Clostridium difficile antibody levels in ...

Antimicrobial Synergy Testing by the Inkjet Printer-assisted Automated Checkerboard Array and the Manual Time-kill Method

As rates of multidrug-resistant (MDR) pathogens continue to rise, outpacing the development of new antimicrobials, novel approaches to treatment of MDR bacteria are increasingly becoming a necessity. One such approach is combination therapy, in which two or more antibiotics are used together to treat an infection against which one or both of the drugs may be ineffective alone. When two drugs, in combination, exert a greater than additive effect, they are considered synergistic. In vitro investigation of synergistic activity is an important first step in evaluating the possible efficacy of drug combinations. Two main in vitro synergy testing methods have been developed: the checkerboard array and the time-kill study. In this paper, we present an automated checkerboard array method that makes use of inkjet printing technology to increase the efficiency and accuracy of this technique, as well as a standard manual time-kill synergy method. The automated checkerboard array can serve as a high-throughput screening assay, while the manual time-kill study provides additional, complementary data on synergistic activity and killing.
The checkerboard array is a modification of standard minimum inhibitory concentration (MIC) testing, in which bacteria are incubated with antibiotics at different concentration combinations and evaluated for growth inhibition after overnight incubation. Manual performance of the checkerboard array requires a laborious and error-prone series of calculations and dilutions. In the automated method presented here, the calculation and dispensing of required antibiotic stock solution volumes are automated through the use of inkjet printer technology. In the time-kill synergy assay, bacteria are incubated with the antibiotics of interest, both together and individually, and sampled at intervals over the course of 24 h for quantitative culture. The results can determine whether a combination is synergistic and whether it is bactericidal, and provide data on inhibition and killing of bacteria over time.

Antimicrobial synergy testing is used to evaluate the effect of two or more antibiotics used in combination and is typically performed by one of two methods: the checkerboard array or the time-kill assay. Here, we present an automated, inkjet printer-assisted checkerboard array synergy technique and a classic time-kill synergy study.

As rates of multidrug-resistant (MDR) pathogens continue to rise, outpacing the development of new antimicrobials, novel approaches to treatment of MDR ...

High-Throughput Screening Assay for Mycobacterium abscessus Using Dual Reporters

Assay Development for High-Throughput Drug Screening Against Mycobacteria

Mycobacterium abscessus (Mab) infections are challenging to treat due to high intrinsic drug resistance, comparable to multidrug-resistant tuberculosis. Treatments are extremely ineffective and based on a multi-drug regimen, resulting in low patient compliance. Consequently, the scientific community is urged to identify new and effective drugs to treat these infections. One of the strategies employed to this end is drug repurposing - the process of identifying new therapeutic opportunities for existing drugs in the market, circumventing the time required to establish pharmacokinetic and safety profiles of new drugs. With most studies on drug development against Mab relying on traditional and time-consuming methods, an assay for high-throughput drug screening was developed against mycobacteria using an in house developed double-reporter strain of Mab. Using liquid-handling robotics, automated microscopy, and analysis, alongside in house developed double reporter strains, bacterial viability can be rapidly measured using two different readouts, luminescence and fluorescence, without adding reagents or performing any extra steps. This reduces time and variability between assays, a major advantage for high-throughput screenings. The described protocol was validated by screening a library of 1280 compounds. The obtained results were corroborated by the literature, with efficient detection of active compounds. Thus, this work fulfilled the aim of supplying the field with a new tool to help fight this extremely drug-resistant bacteria.

Author Spotlight: Scalable Drug Screening Protocol for Efficient Discovery of M. abscessus Treatments

This study developed an assay for high-throughput screening against Mycobacterium abscessus with a&#160;recently created double reporter strain, using fluorescence and luminescence to assess bacterial viability quickly. This protocol will be relevant for researchers aiming to screen new drugs against this drug-resistant bacteria.

Mycobacterium abscessus (Mab) infections are challenging to treat due to high intrinsic drug resistance, comparable to multidrug-resistant tuberculosis. ...

Antibiotic Susceptibility Testing: Epsilometer Tests to Determine MIC Values of Two Antibiotics and Evaluate Antibiotic Synergy

Source: Anna Bl&#228;ckberg1, Rolf Lood1 
1 Department of Clinical Sciences Lund, Division of Infection Medicine, Biomedical Center, Lund University, 221 00 Lund Sweden
Knowledge of the interactions between antibiotics and bacteria is important in understanding how microbes evolve antibiotic resistance. In 1928, Alexander Fleming discovered penicillin, an antibiotic that exerts its antibacterial function by interfering with cell wall regeneration (1). Other antibiotics with diverse mechanisms of action have subsequently been discovered, including drugs that inhibit DNA replication and protein translation in bacteria; however, no new antibiotics have been developed in recent years. Resistance to current antibiotics has been increasing, resulting in severe infectious diseases that cannot be effectively treated (2). Here, we describe several methods to assess antibiotic resistance in bacterial populations. Each of these methods works, regardless of the mechanism of action of the antibiotics used, because bacterial death is the measured outcome. Antibiotic resistance is not only rapidly disseminated specifically through hospital settings, but also throughout society. In order to investigate such means of resistance, different methods have been developed including the Epsilometer test (E-test) and the broth dilution test (3).
The E-test is a well-established method and is a cost-effective tool that quantifies Minimal Inhibitory Concentration (MIC) data, the lowest concentration of an antimicrobial that inhibits visible growth of a microorganism. Depending on the bacterial strain and antibiotics used, the MIC value can vary between sub &#956;g/mL to &#62;1000 &#956;g/mL (4). The E-test is performed using a plastic strip containing a predefined antibiotic gradient, which is imprinted with the MIC reading scale in &#956;g/mL. This strip is directly transferred on the agar matrix when applied to the inoculated agar plate. After incubation, a symmetrical elliptic inhibition zone is visible along the strip as bacterial growth is prevented. MIC is defined by the area of inhibition, which is the endpoint where the ellipse intersects the strip. Another common method to determine MIC is the microbroth dilution method. Microbroth dilution incorporates different concentrations of the antimicrobial agent added to a broth medium containing inoculated bacteria. After incubation, MIC is defined as the lowest concentration of antibiotic that prevents visible growth (5). It is also a quantitative method and can be applied to several bacteria. Disadvantages of this method include the possibility of errors when preparing the concentrations of the reagents and the large number of reagents required for the experiment. Measuring antibiotic resistance is imperative from both a clinical and research perspective, and these in vitro methods of investigating resistance are discussed and showcased below.
The profile of resistance for a specific bacterium can be applied in order to optimize antibiotic treatment to determine whether a patient would benefit from combination treatment versus single therapy. For the usage of more than one antibiotic at a single time, it is imperative to know their interactions with each other and if they have an additive, synergistic, or antagonistic effect. An additive effect can be seen when the joint effect of the antibiotics equals the potency of the individual antibiotics given at an equal dose. Synergy between antibiotics, on the other hand, is present when the joint effect of the antibiotics is more potent than if the drug would be given alone (6). Applying combinations of antimicrobial treatment is used to avoid the occurrence of antimicrobial resistance thus to enhance the effect of the individual antibiotic treatment (7). Knowledge of antagonism is also as important to prevent unnecessary use of antimicrobial combinations. E-test methodology offers simple and several ways to determine possible synergy and antagonism between different antimicrobial agents. In order to face the proliferation of antibiotic resistant pathogens, knowledge of possible synergistic and antagonistic mechanisms of certain antibiotics is important resulting in clinical efficacy and battling multidrug-resistance.
Determination of synergy using E-tests can be divided into two broad approaches: cross and non-cross testing. While both synergy tests rely on the previous knowledge of individual MIC values, the two approaches are slightly different in methodology and conceptual approach. In a non-cross synergy test, the first antibiotic in the pair to be tested is placed on an agar plate inoculated with bacteria. After allowing the antibiotics from the first strip infuse the plate (e.g. after 1 hour), the strip is removed and a new strip containing the second antibiotic is placed on the exact same spot as the first, making sure to place the two individual MIC values on top of each other. The resulting inhibition zone can then be analyzed as described above, and synergy calculated based on the Equation 1.
Equation 1 - Fractional Inhibitory Concentrations (FIC)
<img src="/files/ftp_upload/10512/10512eq1.jpg" />
Values &#62;0.5 demonstrates synergy.
While rewarding the examiner with easy-to-analyze plates, the method is somewhat laborious and time-consuming due to the change of strips, as well as the need of using two plates per experiment. Instead, a cross test is often employed. Instead of adding the two different E-test strips subsequently on top of each other (after removal of the first), both are placed simultaneously but in the form of a cross (<img align="center" src="/files/ftp_upload/10512/10512eq3.jpg" /> 90&#176; angle), with the two previously determined MIC values forming the 90&#176; angle. By this approach only one plate is needed per synergy test, as well as less work, making it a preferred choice despite being slightly more difficult to analyze. The new MIC values in the combined antibiotics approach can be visualized as the modified inhibition zones, after which synergy can be determined by Equation 1.
Instead of using an agar plate approach, a microbroth approach can often be preferential due to its higher flexibility (e.g. ability to choose specific concentrations of antibiotics outside the limits of an E-test strip). Further, microbroth tests are suggested to be more sensitive due to their even distribution of antibiotics in a liquid solution, not depending on the dissociation within a solid phase (agar plate). Wells in a 96-well microplate will be inoculated with a set number of bacteria (106 cfu/mL: bacterial concentration can be estimated by OD600 nm measurements, turbidity standards, or by spread plating samples from 10x bacterial serial dilutions), and antibiotics in different dilutions will be added to the wells. Similarly, to the E-test strips MIC is determined as the intersection (well/spot) with the lowest concentration of antibiotics inhibiting visible growth of bacteria.
Experimental Aim
<ul>
	<li>The project below describes strategies to determine MIC-values of penicillin G and gentamicin of Streptococcus group G by two different methods, E-test and microbroth dilution. For E-test, Mueller-Hinton agar plates inoculated with Streptococcus group G were used in combination with gradient strips of penicillin G and/or gentamicin; while MH-broth with 50% lysed horse blood and 20 mg/mL &#946;-NAD were used with soluble antibiotics together with Streptococcus group G in a microbroth approach.</li>
</ul>
Materials
<ul>
	<li>Bacterial colonies on a blood agar plate, stored &#60;7 days in 4&#176;C</li>
	<li>Blood agar plates</li>
	<li>0.5 McFarland standard
	<ul>
		<li>1% BaCl2</li>
		<li>1% H2SO4</li>
	</ul>
	</li>
	<li>Saline tube (2 mL)</li>
	<li>Cotton tipped applicator</li>
	<li>Mueller-Hinton agar plates (MHA plates)</li>
	<li>MH broth with 50% lysed horse blood and 20 mg/mL &#946;-NAD (MH-F)</li>
	<li>E-test penicillin/gentamicin (or antibiotics of interest) (BioMerieux, Marcy l&#39;Etoile, France, Sweden)</li>
	<li>Antibiotics penicillin/gentamicin (or antibiotics of interest (powder/solution))</li>
</ul>
Note: The specific media used for bacterial growth may vary for different species.

Antibiotic Susceptibility Testing, MIC and Antibiotic Synergy

Source: Anna Bl&#228;ckberg1, Rolf Lood1
1 Department of Clinical Sciences Lund, Division of Infection Medicine, Biomedical Center, Lund University, 221 ...

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Advanced Biology

Microbiology

Photometric Protein Determination

Measuring the concentration is a fundamental step of many biochemical assays. Photometric protein determination takes advantage of the fact that the more a sample contains light-absorbing substances, the less the light will transmit through it. Since the relationship between concentration and absorption is linear, this phenomenon can be used to measure the concentration in samples where it is unknown.
This video describes the basics of photometric protein determination and introduces the Bradford Assay and the Lowry Method. The procedure in the video will cover a typical Bradford assay. Applications covered include direct measurement of very small volumes of nucleic acids to characterize concentration and purity, determination of coupling efficiency of a biomimetic material, and another variation of photometric protein determination using Remazol dye.
<div class="chapter_text" id="script_0">
	Determining the concentration of a protein in samples is a fundamental step in many biochemical assays. Photometric determination can be done with small sample sizes. The more a sample contains light-absorbing substances, the less the light will transmit through it. This provides a quantitative measurement of the absorbing substances. These concepts are so fundamental to science that the articles that introduced two of the techniques are in the three most cited papers of all time. This video will show the concepts behind some of the most common photometric protein determination techniques, how they are performed, and how the gathered data is analyzed.
</div>
<div class="chapter_text" id="script_1">
	Photometric protein determination is based on the relationship between concentration and light absorbency. This is known as the Beer-Lambert Law, which states that the concentration of a light-absorbing species is proportional to its absorbance.
	This principle underlies all photometric protein determination methods. 
	For direct absorption analysis, the absorbance values of unaltered protein samples are measured. Because of their aromatic side chains, tryptophan and tyrosine residues give the highest absorbance readings at a wavelength of 280 nm.
	However, these amino acids-which are two of the least frequently found in proteins-are present in different amounts in every protein, so each determination is unique. To overcome this limitation, more complex assays-that are not dependent on these amino acids-were developed.
	One example is the Bradford Assay, where colored dye is added to the sample. The dye, known as Coomassie Blue, responds proportionally-the more protein present, the more binding events with the dye.
	Then, protein concentration is determined by measuring the absorbance of the bound Coomassie Blue dye, which absorbs light at 594 nm. However, the Bradford assay is linear over a short range of concentrations, so dilutions are often required before analysis.
	The Lowry Method combines the Biuret reagent, an alkaline solution of copper ions that react with peptide bonds, and the Folin-Cioc&#226;lteu reagent, which oxidizes aromatic protein residues. The resulting color change of the sample is proportional to the protein concentration.
	The absorbance of the reduced Folin reagent can be determined at 750 nm. Like direct absorption, each protein has a unique response, and must be calibrated for the protein of interest. Now that we've reviewed the basic principles behind some of the most common assays, let's look at how direct absorption and the Bradford assay are performed.
</div>
<div class="chapter_text" id="script_2">
	To begin a direct absorption analysis, the spectrophotometer is calibrated with a blank to determine zero absorbance. Standard solutions are prepared for use in creating the calibration curve. Then, an aliquot of the first standard is added to a cuvette, and placed into the spectrophotometer.
	The absorbance value at 280 nm is then recorded. This process is repeated for each standard, using a clean cuvette for each run. Once complete, a calibration curve is created by plotting the absorbance versus concentration. The slope of this line is the molar attenuation coefficient, which relates absorbance to concentration.
	Next, the unknown sample is added to a cuvette, and the absorbance value is recorded. As the data analysis for the different photometric determination methods is similar, we will cover that after we look at the Bradford assay.
	Here, the Bradford protein assay is performed with a BSA standard on a 96-well plate. To begin, BSA stock solutions are prepared.
	The unknown solutions are diluted with deionized water to ensure that the concentrations are within the assay's range. Depending on the kit, the Coomassie dye may also require dilution. Then, the calibration curve is set up by adding the BSA standards to the 96-well plate.
	Deionized water is added to reach the needed concentration to generate a standard curve. The unknown sample should be added to the plate in triplicates to ensure an accurate measurement is taken. Coomassie dye is next added to each well, mixing with the pipette.
	Deionized water is added to an empty well as a blank, to measure the absorbance. After waiting 5 min for the dye to bind, the absorbance is measured in a plate-reader at 590 nm.
</div>
<div class="chapter_text" id="script_3">
	Now that we've performed a few assays, let's look at how to analyze the data. Each photometric protein determination method is based on the Beer-Lambert Law.
	The measured absorbance of the standards is used to create a calibration curve, which is then used to determine the concentration of unknown samples. This curve can be manually plotted, though newer spectrophotometric tools will create the calibration curve once all standards have been measured. These systems will also calculate protein concentration as unknown samples are analyzed.
</div>
<div class="chapter_text" id="script_4">
	Now that we've reviewed how to analyze photometric protein determination data, let's look at some of the ways these procedures are utilized.
	The principles of photometric protein determination can also be used to directly measure nucleic acid concentration. The nanodrop spectrophotometer accepts samples of very small volume onto an optically active pedestal. The absorbance is then measured, and the system automatically determines the nucleic acid concentration. Because proteins and other sources can interfere with measurements, sample purity is determined by analyzing the 260 to 280 nm and 260 to 230 nm absorbance ratios. Pure nucleic acids typically yield ratios of approximately 1.8 and approximately 2.0 for DNA and RNA, respectively.
	Photometric protein determination can also be used in the production of biomimetic materials, which are inspired from nature to elicit specific cellular responses. Recombinant adhesins are bound to polystyrene beads to simulate bacterial attachment to host cells. The Bradford assay is used to determine the coupling efficiency of the recombinant adhesion to the beads in the production of the biomimetic material.
	Alternative photometric protein assays can be used in the detection and characterization of protein antimicrobials. Remazol brilliant blue R dye is covalently bonded to heat-killed bacteria. The protein antimicrobial is incubated in the dyed solution. Then, the sample is centrifuged, and the absorbance of the supernatant at 595 nm is measured using a microplate spectrophotometer. Increased absorbance, by the soluble dye released into the supernatant from the labeled bacteria, is a quantitative measurement of enzymatic activity.
</div>
<div class="chapter_text" id="script_5">
	You've just watched JoVE's video on photometric protein determination. This video described the underlying principles of photometric determination, went over general procedures for some common assays, and covered some new advances in techniques. Thanks for watching!
</div>

Photometric Protein Determination: Bradford Assay

Measuring the concentration is a fundamental step of many biochemical assays. Photometric protein determination takes advantage of the fact that the more ...

Qualitative and Quantitative Assays for Detection and Characterization of Protein Antimicrobials

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