The funding for this research project was provided by the Natural Sciences and Engineering Research Council of Canada (NSERC) via a Discovery Grant to YL.

1. Preparation of Reagents and Buffers
<ol>
	<li>0.5 M Ethylenediaminetetraacetic Acid (EDTA)
	<ol>
		<li>In a 2 L beaker, add 186.1 g EDTA to 800 ml of deionized-distilled water (ddH2O). Adjust the pH of the solution to 8.0 using NaOH pellets. Add ddH2O to a final volume of 1.0 L and transfer the solution to an autoclavable glass bottle for autoclaving and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>10&#215; Tris-borate EDTA (10x TBE)
	<ol>
		<li>In a 4 L plastic beaker, add 432 g Tris-base, 200 g boric acid, 80 ml of 0.5 M EDTA (pH 8.0) and ddH2O to a final volume of 4 L. Mix using a stir bar until all the components are dissolved. Transfer the solution to 1 L glass bottles for autoclaving and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>10% Denaturing Polyacrylamide Stock
	<ol>
		<li>In a 4 L plastic beaker, add 1681.7 g urea, 400 ml 10x TBE, 1 L of 40% acrylamide/bisacrylamide (29:1) solution and ddH2O until the final volume is 4 L. Mix using a stir bar until the urea is dissolved. Transfer the solution to 1 L amber glass bottles and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>2x Gel Loading Buffer (2x GLB)
	<ol>
		<li>In a 200 ml glass beaker, add 44 g urea, 8 g sucrose, 10 mg bromophenol blue, 10 mg xylene cyanol FF, 400 &#181;l of 10% sodium dodecyl sulfate, and 4 ml of 10&#215; TBE. Add ddH2O to a final volume of 40 ml and dissolve the components with mild heating at 50 &#176;C while mixing with a magnetic stir bar. Transfer 1 ml aliquot to 1.5 ml microfuge tubes and store at 4 &#176;C. 
		NOTE: Prior use requires brief heating at 90 &#176;C to redissolve any solids.</li>
	</ol>
	</li>
	<li>1 M Tris-hydrochloride (Tris-HCl) (pH 7.5)
	<ol>
		<li>In a glass bottle, add 12.1 g of Tris-base and 70 ml of ddH2O and mix until the solid is dissolved. Adjust the pH to 7.5 using 1 M hydrochloric acid (HCl). Add ddH2O to a final volume of 100 ml, autoclave and store at 4 &#176;C</li>
	</ol>
	</li>
	<li>5 M Sodium Chloride (NaCl)
	<ol>
		<li>In a glass bottle, dissolve 58.4 g of NaCl in 150 ml of ddH2O. Adjust the volume to 200 ml with ddH2O. Store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>DNA Elution Buffer
	<ol>
		<li>In a glass bottle, mix 2.0 ml of 1 M Tris-HCl (pH 7.5), 8.0 ml of 5 M NaCl and 0.4 ml of 0.5 M EDTA (pH 8.0). Adjust the volume to 200 ml with ddH2O, autoclave and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>1 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (pH 7.4)
	<ol>
		<li>In a glass bottle, dissolve 2.38 g of HEPES in 80 ml of ddH2O. Adjust the pH to 7.4 using 5 N NaOH and add ddH2O to a final volume of 100 ml.</li>
	</ol>
	</li>
	<li>1 M Magnesium Chloride (MgCl2)
	<ol>
		<li>In a glass bottle, add 2.03 g of MgCl2-6H2O and bring the volume to 100 ml with ddH2O.</li>
	</ol>
	</li>
	<li>Reaction Buffer (RB)
	<ol>
		<li>In a 50 ml conical tube, add 50 &#181;l of 1 M HEPES (pH 7.4), 1.5 ml of 5 M NaCl, 0.75 ml of 1 M MgCl2, and 5 &#181;l of Tween-20. Add ddH2O to a final volume of 50 ml. Mix the solution and filter into another conical tube using a syringe-driven filter (0.22 &#181;m) and store at 4 &#176;C until use.</li>
	</ol>
	</li>
	<li>Binding Buffer (BB)
	<ol>
		<li>In a 50 ml conical tube, add 500 &#181;l of 1 M Tris-HCl (pH 7.5), 8.8 g of NaCl, 50 &#181;l of 1 M of MgCl2, and 5 &#181;l of Tween 20. Add ddH2O to a final volume of 50 ml. Mix the solution and filter into another conical tube using a syringe-driven filter (0.22 &#181;m) and store at 4 &#176;C until use.</li>
	</ol>
	</li>
	<li>Substrate Solution (SS)
	<ol>
		<li>In a 50 ml conical tube, add 5.8 g of NaCl, 3 ml of 1 M MgCl2, 1.5 g urea, and 40 ml of ddH2O. Adjust the pH of the solution to 5.0 using 10 mM HCl. Because the solution is not buffered, adjust pH using HCl carefully through the addition of small HCl aliquots. Add ddH2O to a final volume of 50 ml.</li>
		<li>Mix the solution and filter into another conical tube using a syringe-driven filter (0.22 &#181;m) and store at 4 &#176;C until use.</li>
	</ol>
	</li>
	<li>Luria Bertani (LB) Broth
	<ol>
		<li>In a beaker, dissolve 20.0 g LB powder in 1 L of ddH2O. Transfer to a glass bottle, autoclave, and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>1.5% LB agar
	<ol>
		<li>In a 250 ml flask, add 1.5 g agar and 100 ml of LB broth. Autoclave and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Agar Plates
	<ol>
		<li>Redissolve the LB agar in a microwave and cool the solution to ~50 &#176;C. Pour the solution into petri dishes, making ~5 plates and allow them to solidify.</li>
	</ol>
	</li>
</ol>
2. Synthesis and Purification of E. coli-responsive DNAzyme EC1
<ol>
	<li>Synthesis of EC1 by Template Mediated Enzymatic Ligation
	<ol>
		<li>Purify commercially synthesized oligonucleotides BS1, DE1, and T1 (sequences provided in Table 1) by 10% denaturing polyacrylamide gel electrophoresis (dPAGE) according to standard protocols.</li>
		<li>Prepare a 100 &#181;M stock of BS1, DE1, and T1. Store at -20 &#176;C until use.</li>
		<li>In a 1.5 ml microfuge tube, add 38.5 &#181;l of ddH2O, 5 &#181;l of DE1, and 5 &#181;l of 10x T4 polynucleotide kinase reaction buffer provided by enzyme supplier (500 mM Tris-HCl (pH 7.6), 100 mM MgCl2, 50 mM dithiothreitol (DTT), 1.0 mM spermidine).</li>
		<li>Add 1 &#181;l of adenosine triphosphate (ATP) (100 mM). Add 5 units of T4 polynucleotide kinase (10 U/&#181;l). Mix by carefully pipetting the reaction mixture.</li>
		<li>Incubate the reaction at 37 &#176;C for 30 min.</li>
		<li>Quench the reaction by heating to 90 &#176;C for 5 min.</li>
		<li>Add 118 &#181;l of ddH2O, 5 &#181;l of BS1, and 5 &#181;l of T1. Heat the reaction to 90 &#176;C for 2 min and then cool to room temperature over 10 min.</li>
		<li>Add 20 &#181;l of 10&#215; T4 DNA ligase reaction buffer provided by the enzyme supplier (400 mM Tris-HCl (pH 7.8), 100 mM MgCl2, 100 mM DTT, 5 mM ATP). Add 10 units of T4 DNA ligase (5 U/&#181;l) and carefully mix by pipetting.</li>
		<li>Incubate at room temperature for 2 hr.</li>
		<li>Add 20 &#181;l of 3 M sodium acetate (NaOAc) (pH 5.2), 500 &#181;l of cold 100% ethanol. Mix the solution by vortexing and place the tube in a -20 &#176;C freezer for 30 min.</li>
		<li>Centrifuge the microfuge at 20,000 x g for 20 min at 4 &#176;C. Carefully remove the supernatant by pipetting.</li>
		<li>Wash the pellet with cold 70% ethanol and centrifuge again at 20,000 x g for 10 min at 4 &#176;C. Once again remove the supernatant and dry the pellet under vacuum for 10 min.</li>
		<li>Resuspend the DNA with 15 &#181;l of ddH2O and then add 15 &#181;l of 2x GLB.</li>
		<li>Vortex vigorously and heat to 90 &#176;C for 2 min. Purify the full-length DNA by 10% dPAGE as described below.</li>
	</ol>
	</li>
	<li>Setting Up 10% dPAGE
	<ol>
		<li>Clean two glass plates (one full plate and one notched), two 0.75 mm thick spacers, and a 12-well comb. Lay one glass plate on a flat surface with two spacers on each side and the notched plate on top. Clip the two plates together using the four clips provided by the supplier.</li>
		<li>In a 150 ml plastic beaker, pour 40 ml of 10% dPAGE, 40 &#181;l of tetramethylethylenediamine (TEMED), and 400 &#181;l of 10% ammonium persulfate (APS). Mix the components and pour the solution between the plates slowly.</li>
		<li>Insert the comb and then allow the gel to polymerize over 10 min. Once the gel is polymerized, slowly remove the comb and flush the wells with ddH2O.</li>
		<li>Mount the plates onto the gel electrophoresis apparatus. Use a metal plate to dissipate heat generated to prevent overheating.</li>
		<li>Pour 1x TBE onto the top and bottom chambers and ensure that the wells are well submerged in the buffer. Flush the wells with 1x TBE using a pipette or syringe.</li>
		<li>Set the apparatus to run at 35 mA and pre-run for 15 min before loading samples.</li>
	</ol>
	</li>
	<li>Elution of Ligated EC1 from 10% dPAGE
	<ol>
		<li>Following step 2.2.6, run the gel at 35 mA until the bottom dye (bromophenol blue) reaches the bottom of the glass plate. This should take approximately 1.5 hr. Turn off the power and remove the plates from the apparatus.</li>
		<li>Lay the plates on a few sheets of paper towels and carefully remove the spacers from the glass plates. Carefully remove the top glass plate and wrap the gel with plastic wrap.</li>
		<li>Flip the gel over to remove the second glass plate and cover with plastic wrap again. Take care to avoid wrinkles of the plastic wrap.</li>
		<li>Visualize the ligated product by using UV shadowing (260 nm), which will produce 4 distinct DNA bands (Fully ligated EC1, DE1, BS1, and T1).</li>
		<li>Excise the top band (EC1) with a sterile razor blade and transfer to a 1.5 ml microfuge tube. Crush the gel with a sterile pipette tip (200 &#181;l tip size) until it turns into a fine paste. Add 550 &#181;l of DNA elution buffer, cover the tube with aluminum foil to protect the embedded fluorophore, and shake for 15 min.</li>
		<li>Centrifuge the gel solution at 20,000 x g for 5 min and carefully transfer 400 &#181;l of the supernatant to another 1.5 ml microfuge tube. Avoid withdrawing gel pieces during pipetting.</li>
		<li>To the microfuge tube with the transferred supernatant add 40 &#181;l of 3 M NaOAc (pH 5.2), and 1.0 ml of cold 100% ethanol. Mix the solution by vortexing and place the tube in a -20 &#176;C freezer for 30 min.</li>
		<li>Centrifuge the microfuge at 20,000 x g for 20 min at 4 &#176;C. Carefully remove the supernatant by pipetting.</li>
		<li>Wash the pellet with cold 70% ethanol and centrifuge again at 20,000 x g for 10 min at 4 &#176;C. Once again, remove the supernatant and dry the pellet under vacuum for 10 min.</li>
		<li>Determine the concentration of EC1 by measuring UV absorbance at 260 nm. Make a 10 &#181;M stock and store the sample at -20 &#176;C until use.</li>
	</ol>
	</li>
</ol>
3. Conjugation of Urease to DNA
<ol>
	<li>Preparation of Succinimidyl 4-(N-maleimidomethyl) cyclohexane-1-carboxylate (SMCC) Stock
	<ol>
		<li>Dissolve 1 mg SMCC in 676 &#181;l of DMSO. Vortex and place on ice until use.</li>
	</ol>
	</li>
	<li>Preparation of Urease Stock
	<ol>
		<li>Dissolve 1 mg urease in 1 ml of 1&#215; PBS (no Mg2+ or Ca2+). Place on ice until use. 
		NOTE: Crystallized urease is slow to dissolve and gentle mixing is needed to avoid denaturation.</li>
	</ol>
	</li>
	<li>Synthesis of Urease-DNA (UrDNA)
	<ol>
		<li>Add 10 &#181;l of 100 &#181;M LD1 to a 2.5 ml microfuge tube. Add 140 &#181;l of ddH2O, 40 &#181;l of 10x PBS, and vortex.</li>
		<li>Add 80.5 &#181;l of SMCC stock, 159.5 &#181;l of DMSO, vortex, and briefly centrifuge using a benchtop centrifuge.</li>
		<li>Incubate at 37 &#176;C for 60 min. Avoid condensation under the microfuge cap.</li>
		<li>Add 200 &#181;l of 1x PBS, 60 &#181;l of 3 M NaOAc (pH 5.2), and 1.5 ml of cold 100% ethanol. Mix the solution by vortexing and incubate at -20 &#176;C for 30 min.</li>
		<li>Centrifuge the solution at 20,000 x g for 20 min at 4 &#176;C. Remove the supernatant and dry the pellet under vacuum. Avoid over-drying.</li>
		<li>To the dried pellet add 400 &#181;l of urease stock and incubate at room temperature for 5 hr.</li>
		<li>Transfer 200 &#181;l of crude conjugate to a pre-washed 100k MWCO centrifugal filter column. Centrifuge the column at 14,000 x g for 5 min.</li>
		<li>Transfer the remaining 200 &#181;l of crude conjugate to the column and centrifuge at 14,000 x g for 5 min. Remove the column and place it upside down in a new 2.0 ml microfuge tube (&#34;collection tube&#34;).</li>
		<li>Centrifuge the collection tube (with the inverted column) at 1,000 x g for 2 min. Remove the collection tube and add 30 &#181;l of 1x PBS to the centrifugal column to wash the membrane for additional recovery of conjugates.</li>
		<li>Once again invert the column and place back into the collection tube. Centrifuge the collection tube (with the inverted column) at 1,000 x g for 2 min. Remove and dispose the column.</li>
		<li>Store the UrDNA at 4 &#176;C until use.</li>
	</ol>
	</li>
</ol>
4. Assembly of EC1 and UrDNA onto Magnetic Beads
<ol>
	<li>Mix the magnetic bead (MB) stock well and transfer 100 &#181;l of MB suspension to a 1.5 ml microfuge tube. Place the microfuge tube on a magnetic rack holder for isolating the MB.</li>
	<li>Remove the supernatant by pipetting and add 150 &#181;l of binding buffer (BB) to the tube. Remove the tube from the holder and carefully tap the tube to resuspend the MB to a homogeneous solution. Avoid splashing the suspension to the top of the tube or cap. If this happens, use a benchtop centrifuge to briefly spin the residue back to the suspension.</li>
	<li>Repeat step 4.2 two more times.</li>
	<li>To this suspension add 10 &#181;l of 10 &#181;M EC1. Carefully mix by tapping on the tube.</li>
	<li>Incubate the solution with mild shaking for 30 min. Tap the tube every 2-3 min to avoid the precipitation of the MB.</li>
	<li>Place the microfuge tube back on the magnetic rack to isolate the MB and remove the supernatant by pipetting. Wash the MB three times with 150 &#181;l of BB (as described in step 4.2).</li>
	<li>Once the washing is complete, suspend the MB in a total of 150 &#181;l of BB. To this solution, add 15 &#181;l of UrDNA and heat the solution to 45 &#176;C for 2 min. Cool the solution to room temperature and incubate for 2 hr.</li>
	<li>Place the microfuge tube back on the magnetic rack to isolate the MB and remove the supernatant by pipetting.</li>
	<li>Add 100 &#181;l of reaction buffer (RB). Remove the microfuge tube from the magnetic rack and carefully resuspend the MB.</li>
	<li>Wash the MB three more times by repeating step 4.9. 
	NOTE: The washed supernatant can be quickly tested to determine if unhybridized UrDNA is still present, which may result in a false positive signal. The test can be done by adding 10 &#181;l of 50 mM urea and 10 &#181;l of 0.04% phenol red to the wash solution. Continue to wash the MB until the wash solution does not cause a color change. The 100 &#181;l suspension is stored at 4 &#176;C until use.</li>
</ol>
5. Preparation of Bacterial Cells20
<ol>
	<li>Culturing E. coli from Stocks
	<ol>
		<li>Plate E. coli K12 (MG1655) onto LB agar plates from a glycerol stock under a flame or in a biological safety cabinet.</li>
		<li>Using a sterile pipette tip, gently touch the glycerol stock and lightly streak the surface of an agar plate to avoid puncturing the LB agar.</li>
		<li>Invert the streaked plate and incubate at 37 &#176;C for 14 hr.</li>
		<li>Seal the plate with Parafilm and store at 4 &#176;C for a maximum of 3 weeks.</li>
	</ol>
	</li>
	<li>Culturing E. coli for Cell Counting
	<ol>
		<li>In a sterile 14 ml culture tube, dispense 2 ml of LB broth.</li>
		<li>Using a sterile pipette tip, pick a single colony from the streaked agar plate prepared in step 5.1 and transfer it to the culture tube.</li>
		<li>Incubate the culture at 37 &#176;C and shake at 230 rpm for 14 hr.</li>
		<li>Serially dilute the bacterial culture in 10-fold intervals.</li>
		<li>For each diluted sample, evenly plate five 100 &#181;l aliquots on separate LB agar plates. Invert the plates and incubate at 37 &#176;C for 14 hr.</li>
		<li>Count the cells of each sample to obtain the average cell concentration of each dilution. 
		NOTE: 107 cells are often used to set up a reference litmus test for E. coli as this level of E. coli can trigger a quick color change. However, a properly executed litmus test can detect as low as 500 cells, as discussed in the Results section.</li>
	</ol>
	</li>
	<li>Preparing E. coli Cells for Testing
	<ol>
		<li>For a desired cell suspension, transfer 1 ml of cultured stock to a 1.5 ml microfuge tube.</li>
		<li>Centrifuge the cells at 6,000 x g for 10 min at 4 &#176;C. Carefully remove the supernatant without disturbing the cell pellet.</li>
		<li>Add 10 &#181;l of reaction buffer to the cell pellet and resuspend the cells. Sonicate the cell suspension for 5 min. Transfer the cell suspension to an ice box for 5 min.</li>
		<li>Sonicate the cell suspension for another 5 min.</li>
		<li>Centrifuge the cell suspension at 13,000 x g for 10 min at 4 &#176;C. Use the supernatant for testing (10 &#181;l).</li>
	</ol>
	</li>
</ol>
6. Litmus Test
<ol>
	<li>In a 1.5 ml microfuge tube, prewash the tube by adding and vortexing 100 &#181;l of reaction buffer (RB) in the microfuge tube and discarding the buffer.</li>
	<li>Transfer 15 &#181;l of assembled EC1 (protocol 4) to the washed microfuge tube.</li>
	<li>Wash the magnetic beads by placing the microfuge tube on a magnetic rack. Remove the supernatant by pipetting. Remove the microfuge tube from the rack, add 100 &#181;l of RB, and carefully resuspend the magnetic beads.</li>
	<li>Wash the MB two more times by repeating step 6.3.</li>
	<li>Place the microfuge tube back on the magnetic rack, remove the supernatant and add the 10 &#181;l E. coli sample prepared from step 5.3.</li>
	<li>Mix the sample and magnetic beads carefully by gently tapping on the microfuge tube.</li>
	<li>Incubate the reaction at room temperature for 1 hr.</li>
	<li>To the reaction, add 90 &#181;l of ddH2O and place the microfuge tube onto a magnetic rack.</li>
	<li>After approximately 3 min of magnetic separation, carefully transfer 85 &#181;l of the supernatant to a 0.5 ml microfuge tube. Withdraw the supernatant slowly to avoid collecting any magnetic beads.</li>
	<li>To the above microfuge tube add 15 &#181;l of 0.04% phenol red, and 100 &#181;l of substrate solution.</li>
	<li>Take a photograph at specific time intervals to record color change. 
	NOTE: The change in pH can also be monitored using a pH meter with a microelectrode. The starting pH should be approximately 5.2-5.5 (the solution is yellow). If not, the solution can be adjusted by addition of 1 mM Acetate Buffer pH 5.0).</li>
</ol>

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<li>Chiuman, W., Li, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Simple+fluorescent+sensors+engineered+with+catalytic+DNA+%27MgZ%27+based+on+a+non-classic+allosteric+design%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">Simple fluorescent sensors engineered with catalytic DNA 'MgZ' based on a non-classic allosteric design.</a> PLoS One. 2, e1224(2007).</li>
<li>Xiang, Y., Lu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Using+personal+glucose+meters+and+functional+DNA+sensors+to+quantify+a+variety+of+analytical+targets%5BTitle%5D)+AND+%22Nat.+Chem%22%5BJournal%5D)">Using personal glucose meters and functional DNA sensors to quantify a variety of analytical targets.</a> Nat. Chem. 3, 697-703 (2011).</li>
<li>Aguirre, S. D., Ali, M. M., Salena, B. J., Li, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+sensitive+DNA+enzyme-based+fluorescent+assay+for+bacterial+detection%5BTitle%5D)+AND+%22Biomolecules%22%5BJournal%5D)">A sensitive DNA enzyme-based fluorescent assay for bacterial detection.</a> Biomolecules. 3, 563-577 (2013).</li>
<li>Aguirre, S. D., Ali, M. M., Kanda, P., Li, Y. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Detection+of+Bacteria+Using+Fluorogenic+DNAzymes%5BTitle%5D)+AND+%22J.+Vis.+Exp%22%5BJournal%5D)">Detection of Bacteria Using Fluorogenic DNAzymes.</a> J. Vis. Exp. (63), e3961(2012).</li>
<li>Shen, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+catalytic+DNA+activated+by+a+specific+strain+of+bacterial+pathogen%5BTitle%5D)+AND+%22Angew.+Chem.+Int.+Ed%22%5BJournal%5D)">A catalytic DNA activated by a specific strain of bacterial pathogen.</a> Angew. Chem. Int. Ed. 54, (2015).</li>
<li>He, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Highly+specific+recognition+of+breast+tumors+by+an+RNA-cleaving+fluorogenic+DNAzyme+probe%5BTitle%5D)+AND+%22Anal.+Chem%22%5BJournal%5D)">Highly specific recognition of breast tumors by an RNA-cleaving fluorogenic DNAzyme probe.</a> Anal. Chem. 87, 569-577 (2015).</li>
<li>Sumner, J. B., Hand, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Isoelectric+point+of+crystalline+urease%5BTitle%5D)+AND+%22J.+Am.+Chem.+Soc%22%5BJournal%5D)">Isoelectric point of crystalline urease.</a> J. Am. Chem. Soc. 51, 1255-1260 (1929).</li>
<li>Karplus, P. A., Pearson, M., Hausinger, R. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((70+Years+of+crystalline+urease%3A+What+have+we+learned%5BTitle%5D)+AND+%22Acc.+Chem.+Res%22%5BJournal%5D)">70 Years of crystalline urease: What have we learned.</a> Acc. Chem. Res. 30, 330-337 (1997).</li>
<li>Omiccioli, E., Amagliani, G., Brandi, G., Magnani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+new+platform+for+Real-Time+PCR+detection+of+Salmonella+spp.%2C+Listeria+monocytogenes+and+Escherichia+coli+O157+in+milk%5BTitle%5D)+AND+%22Food+Microbiol%22%5BJournal%5D)">A new platform for Real-Time PCR detection of Salmonella spp., Listeria monocytogenes and Escherichia coli O157 in milk.</a> Food Microbiol. 26, 615-622 (2009).</li>
<li>Cui, S., Schroeder, C. M., Zhang, D. Y., Meng, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rapid+sample+preparation+method+for+PCR-based+detection+of+Escherichia+coli+O157%3AH7+in+ground+beef%5BTitle%5D)+AND+%22J.+Appl.+Microbiol%22%5BJournal%5D)">Rapid sample preparation method for PCR-based detection of Escherichia coli O157:H7 in ground beef.</a> J. Appl. Microbiol. 95, 129-134 (2003).</li>
<li>Ibekwe, A. M., Watt, P. M., Grieve, C. M., Sharma, V. K., Lyons, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Multiplex+fluorogenic+real-time+PCR+for+detection+and+quantification+of+Escherichia+coli+O157%3AH7+in+dairy+wastewater+wetlands%5BTitle%5D)+AND+%22Appl.+Environ.+Microbiol%22%5BJournal%5D)">Multiplex fluorogenic real-time PCR for detection and quantification of Escherichia coli O157:H7 in dairy wastewater wetlands.</a> Appl. Environ. Microbiol. 68, 4853-4862 (2002).</li>
<li>Strachan, N. J., Ogden, I. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+sensitive+microsphere+coagulation+ELISA+for+Escherichia+coli+O157%3AH7+using+Russell%27s+viper+venom%5BTitle%5D)+AND+%22FEMS+Microbiol+Lett%22%5BJournal%5D)">A sensitive microsphere coagulation ELISA for Escherichia coli O157:H7 using Russell's viper venom.</a> FEMS Microbiol Lett. 186, 79-84 (2000).</li>
<li>de Boer, E., Beumer, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Methodology+for+detection+and+typing+of+foodborne+microorganisms%5BTitle%5D)+AND+%22Int.+J.+Food+Microbiol%22%5BJournal%5D)">Methodology for detection and typing of foodborne microorganisms.</a> Int. J. Food Microbiol. 50, 119-130 (1999).</li>
<li>Gracias, K. S., McKillip, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+review+of+conventional+detection+and+enumeration+methods+for+pathogenic+bacteria+in+food%5BTitle%5D)+AND+%22Can.+J.+Microbiol%22%5BJournal%5D)">A review of conventional detection and enumeration methods for pathogenic bacteria in food.</a> Can. J. Microbiol. 50, 883-890 (2004).</li>
</ol>

The authors have nothing to disclose.

The translation of the action of the RNA cleavage activity of a bacterium-responsive DNAzyme to a litmus test is made possible through the use of urease and magnetic separation, as illustrated by Figure 1. Although the demonstration of the modified litmus test for bacterial detection is done with an E. coli-dependent RNA-cleaving DNAzyme,5,19,20 the design can be generally extended for any RNA-cleaving DNAzyme. Given the great availability of RNA-cleaving DNAzymes for different analyt...

Bacterial pathogens are one of the major causes of global morbidity and mortality. Outbreaks from hospital-acquired infections, food-borne pathogens, and bacterial contaminants in the environment pose serious and on-going threats to public health and safety. To prevent these outbreaks, effective tools are needed that permit pathogen detection in a timely fashion under a variety of settings. Simple but still effective tests that are portable and cost-effective are greatly coveted, especially in regions that are susceptible to outbreaks but cannot afford expensive testing facilities.1-3 Although there exists a multitude of methods to detect bacteria, many of them are not suitable as screening or on-site testing tools because they require long test times, expensive instruments and complicated testing procedures. 
Colorimetric tests are particularly attractive for point-of-care or field applications as color changes can be easily detected by the naked eye. The litmus test for pH is simple, quick, and effective. Although it is a very old technology, it is still widely used today because of its simplicity and effectiveness. Surprisingly, this simple test had never been modified to achieve the detection of other analytes before we recently developed an approach of modifying this test for E. coli testing.4
The expanded litmus test for E. coli employs three additional components: an E. coli activated RNA-cleaving DNAzyme (EC1), 5 urease, and magnetic beads. DNAzymes refer to synthetic single-stranded DNA molecules with catalytic activity.6 They can be isolated from random-sequence DNA pools using in vitro selection.7,8 They are highly stable and can be produced cost-effectively using high-efficiency automated DNA synthesis.9 For these reasons, DNAzymes, particularly RNA-cleaving DNAzymes, have been widely examined for biosensing applications.6,10,11 RNA-cleaving DNAzyme sensors have been developed to detect metal ions,12-16 small molecules,17,18 bacterial pathogens5,19-21 and cancer cells.22 Given the great availability of target-induced RNA-cleaving DNAzymes, any assay that utilizes a DNAzyme can be potentially expanded to detect a diverse range of analytes.
Urease is chosen for its ability to hydrolyze urea into ammonia,23,24 resulting in a pH increase. Urease is also highly efficient, stable and amenable for conjugation to other biomolecules. Therefore, we postulated that a conjugate of an RNA-cleavage DNAzyme with urease would allow the use of litmus test for the detection of other targets.5
The action of the RNA-cleaving DNAzyme is relayed to urease-mediated increase of pH through the use of magnetic beads that are immobilized with the DNAzyme-urease conjugate. Because the activity of the DNAzyme under investigation is strictly dependent on E. coli, the presence of this bacterium in the test solution will result in the release of urease from the magnetic beads to the solution, which is then taken and used to hydrolyze urea in a reporter solution that contains a pH-sensitive dye. The final outcome of this procedure is a color change that can be conveniently reported by the dye or pH paper.

<table><tbody><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>VWR AMRESCO</td><td>0105</td><td></td></tr><tr><td>Sodium Hydroxide (NaOH) pellets</td><td>BIO BASIC CANADA INC.</td><td>SB6789</td><td></td></tr><tr><td>Tris-base</td><td>VWR AMRESCO</td><td>0497</td><td></td></tr><tr><td>Boric acid</td><td>AMRESCO</td><td>0588</td><td></td></tr><tr><td>Urea</td><td>VWR AMRESCO</td><td>M123</td><td></td></tr><tr><td>40% acrylamide/bisacrylamide (29:1) solution</td><td>BIO BASIC CANADA INC.</td><td>A0007</td><td></td></tr><tr><td>Sucrose</td><td>Bioshop Canada inc.</td><td>SUC507</td><td></td></tr><tr><td>Bromophenol blue</td><td>Bioshop Canada inc.</td><td>BRO777</td><td></td></tr><tr><td>Xylenecyanol FF</td><td>SIGMA-ALDRICH</td><td>X-4126</td><td></td></tr><tr><td>10% sodium dodecyl sulfate</td><td>Bioshop Canada inc.</td><td>SDS001</td><td></td></tr><tr><td>Hydrochloric Acid (HCl)</td><td>CALEDON LABORATORIES LTD</td><td>6026</td><td></td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Bioshop Canada inc.</td><td>SOD001</td><td></td></tr><tr><td>4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)</td><td>Bioshop Canada inc.</td><td>HEP001</td><td></td></tr><tr><td>Magnesium Chloride (II) hexahydrate</td><td>VWR AMRESCO</td><td>0288</td><td></td></tr><tr><td>Tween 20</td><td>Bioshop Canada inc.</td><td>TW508</td><td></td></tr><tr><td>Adenosine Triphospahte (ATP)</td><td>AMRESCO</td><td>0220</td><td></td></tr><tr><td>Sodium Acetate trihydrate (NaOAc)</td><td>SIGMA-ALDRICH</td><td>S8625</td><td></td></tr><tr><td>Ethanol</td><td>Commercial Alcohols</td><td>P016EAAN</td><td></td></tr><tr><td>Tetramethyleneethylenediamine (TEMED)</td><td>AMRESCO</td><td>0761</td><td></td></tr><tr><td>10% Ammonium persulfate (APS)</td><td>BIO BASIC CANADA INC.</td><td>AB0072</td><td></td></tr><tr><td>Succinimidyl 4-(&lt;em&gt;N&lt;/em&gt;-maleimidomethyl) cyclohexane-1-carboxylate (SMCC)</td><td>ThermoFisher SCIENTIFIC</td><td>22360</td><td></td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>CALEDON LABORATORIES</td><td>803540</td><td></td></tr><tr><td>Urease</td><td>SIGMA-ALDRICH</td><td>U0251</td><td></td></tr><tr><td>1x Phosphate Buffered Saline (PBS)</td><td>ThermoFisher SCIENTIFIC</td><td>70011-069</td><td></td></tr><tr><td>0.04% Phenol red</td><td>SIGMA-ALDRICH</td><td>P3532</td><td></td></tr><tr><td>10x T4 polynucleotide kinase reaction buffer</td><td>Lucigen</td><td>30061-1</td><td></td></tr><tr><td>10x T4 DNA ligase reaction buffer</td><td>Bio Basics Canada</td><td>B1122-B</td><td></td></tr><tr><td>T4 DNA ligase (5 U/&amp;#956;l)</td><td>Thermo Fischer Scientific</td><td>B1122</td><td></td></tr><tr><td>Luria Bertani (LB) Broth</td><td>AMRESCO</td><td>J106</td><td></td></tr><tr><td>Agar</td><td>AMRESCO</td><td>J637</td><td></td></tr><tr><td>T4 polynucleotide kinase (10 U/&amp;#956;l)</td><td>Lucigen</td><td>30061-1</td><td></td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; K12 (MG1655)</td><td>ATCC</td><td>ATCC700926</td><td></td></tr><tr><td>Centrifuge</td><td>Beckman Coulter, Inc.</td><td>392187</td><td></td></tr><tr><td>Glass plates</td><td>CBS scientific</td><td>ngp-250nr</td><td></td></tr><tr><td>0.75 mm thick spacers</td><td>CBS scientific</td><td>VGS-0725r</td><td></td></tr><tr><td>12-well comb</td><td>CBS scientific</td><td>VGC-7512</td><td></td></tr><tr><td>UV Lamp</td><td>UVP</td><td>95-0017-09</td><td></td></tr><tr><td>Spectrophotometer (NanoVue)</td><td>GE Healthcare</td><td>N/A</td><td></td></tr><tr><td>Metal plate</td><td>CBS scientific</td><td>CPA165-250</td><td></td></tr><tr><td>Vortex</td><td>VWR International</td><td>58816-123</td><td></td></tr><tr><td>Gel electrophoresis apparatus</td><td>CBS scientific</td><td>ASG-250</td><td></td></tr><tr><td>Petri dishes</td><td>VWR International</td><td>25384-342</td><td></td></tr><tr><td>100 kDa MWCO centrifugal filters</td><td>EMD Millipore</td><td>UFC510024</td><td></td></tr><tr><td>Magnetic Bead (BioMag)</td><td>Bangs Laboratories Inc</td><td>BM568</td><td></td></tr><tr><td>Magnetic Seperation Rack</td><td>New England BioLabs</td><td>S1506S</td><td></td></tr><tr><td>Microfuge tubes</td><td>Sarstedt</td><td>72.69</td><td></td></tr><tr><td>Syringe filter (0.22 &amp;#956;m)</td><td>VWR International</td><td>28145-501</td><td></td></tr><tr><td>14 ml culture tube</td><td>VWR International</td><td>60818-725</td><td></td></tr><tr><td>Cell culture incubator</td><td>Eppendorf Scientific</td><td>M13520000</td><td></td></tr><tr><td>Branson Ultrasonic cleaner</td><td>Branson</td><td>N/A</td><td></td></tr><tr><td>Camera (Canon Powershot G11)</td><td>Canon</td><td>N/A</td><td></td></tr><tr><td>50 ml conical tube</td><td>VWR International</td><td>89004-364</td><td></td></tr></tbody></table>

colorimetric detection of bacteria using litmus test

There are increasing demands for simple but still effective methods that can be used to detect specific pathogens for point-of-care or field applications. Such methods need to be user-friendly and produce reliable results that can be easily interpreted by both specialists and non-professionals. The litmus test for pH is simple, quick, and effective as it reports the pH of a test sample via a simple color change. We have developed an approach to take advantage of the litmus test for bacterial detection. The method exploits a bacterium-specific RNA-cleaving DNAzyme to achieve two functions: recognizing a bacterium of interest and providing a mechanism to control the activity of urease. Through the use of magnetic beads immobilized with a DNAzyme-urease conjugate, the presence of bacteria in a test sample is relayed to the release of urease from beads to solution. The released urease is transferred to a test solution to hydrolyze urea into ammonia, resulting in an increase of pH that can be visualized using the classic litmus test.

The principle of the bacterial litmus test is explained in Figure 1. The test uses three key materials: an RNA-cleaving DNAzyme that is activated by a specific bacterium, urease and magnetic beads. The DNAzyme is used as the molecular recognition element to achieve highly specific detection of a bacterium of interest. Urease and magnetic beads are used to achieve signal transduction of the RNA-cleavage activity of the DNAzyme. This involves the creation of magnetic beads ...

Watch this Scientific Journal Video about Colorimetric Detection of Bacteria Using Litmus Test at JoVE.com

Colorimetric Detection of Bacteria Using Litmus Test

We describe a protocol for colorimetric detection of E. coli using a modified litmus test that takes advantage of an RNA-cleaving DNAzyme, urease, and magnetic beads.

There are increasing demands for simple but still effective methods that can be used to detect specific pathogens for point-of-care or field applications. ...

colorimetric-detection-of-bacteria-using-litmus-test

Research

JoVE Journal

Biology

9.2K Views.  McMaster University. We describe a protocol for colorimetric detection of E. coli using a modified litmus test that takes advantage of an RNA-cleaving DNAzyme, urease, and magnetic beads. 

Video: Colorimetric Detection of Bacteria Using Litmus Test

Colorimetric Paper-based Detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from Large Volumes of Agricultural Water

This protocol describes rapid colorimetric detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from large volumes (10 L) of agricultural waters. Here, water is filtered through sterile Modified Moore Swabs (MMS), which consist&#160;of a simple gauze filter enclosed in a plastic cartridge, to concentrate bacteria. Following filtration, non-selective or selective enrichments for the target bacteria are performed in the MMS. For colorimetric detection of the target bacteria, the enrichments are then assayed using paper-based analytical devices (&#181;PADs) embedded with bacteria-indicative substrates. Each substrate reacts with target-indicative bacterial enzymes, generating colored products that can be detected visually (qualitative detection) on the &#181;PAD. Alternatively, digital images of the reacted &#181;PADs can be generated with common scanning or photographic devices and analyzed using ImageJ software, allowing for more objective and standardized interpretation of results. Although the biochemical screening procedures are designed to identify the aforementioned bacterial pathogens, in some cases enzymes produced by background microbiota or the degradation of the colorimetric substrates may produce a false positive. Therefore, confirmation using a more discriminatory diagnostic is needed. Nonetheless, this bacterial concentration and detection platform is inexpensive, sensitive (0.1 CFU/ml detection limit), easy to perform, and rapid (concentration, enrichment, and detection are performed within approximately 24&#160;hr), justifying its use as an initial screening method for the microbiological quality of agricultural water.

Colorimetric Paper-based Detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from Large Volumes of Agricultural Water

A protocol involving integrated concentration, enrichment, and end-point colorimetric detection of foodborne pathogens in large volumes of agricultural water is presented here. Water is filtered through Modified Moore Swabs (MMS), enriched with selective or non-selective media, and detection is performed using paper-based analytical devices (&#181;PAD) imbedded with bacterial-indicative colorimetric substrates.

This protocol describes rapid colorimetric detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from large volumes (10 L) of ...

Environment

'Bioluminescent' Reporter Phage for the Detection of Category A Bacterial Pathogens

Yersinia pestis and Bacillus anthracis are Category A bacterial pathogens that are the causative agents of the plague and anthrax, respectively 1. Although the natural occurrence of both diseases&#39; is now relatively rare, the possibility of terrorist groups using these pathogens as a bioweapon is real. Because of the disease&#39;s inherent communicability, rapid clinical course, and high mortality rate, it is critical that an outbreak be detected quickly. Therefore methodologies that provide rapid detection and diagnosis are essential to ensure immediate implementation of public health measures and activation of crisis management.
Recombinant reporter phage may provide a rapid and specific approach for the detection of Y. pestis and B. anthracis. The Centers for Disease Control and Prevention currently use the classical phage lysis assays for the confirmed identification of these bacterial pathogens 2-4. These assays take advantage of naturally occurring phage which are specific and lytic for their bacterial hosts. After overnight growth of the cultivated bacterium in the presence of the specific phage, the formation of plaques (bacterial lysis) provides a positive identification of the bacterial target. Although these assays are robust, they suffer from three shortcomings: 1) they are laboratory based; 2) they require bacterial isolation and cultivation from the suspected sample, and 3) they take 24-36 h to complete. To address these issues, recombinant &#34;light-tagged&#34; reporter phage were genetically engineered by integrating the Vibrio harveyi luxAB genes into the genome of Y. pestis and B. anthracis specific phage 5-8. The resulting luxAB reporter phage were able to detect their specific target by rapidly (within minutes) and sensitively conferring a bioluminescent phenotype to recipient cells. Importantly, detection was obtained either with cultivated recipient cells or with mock-infected clinical specimens 7.
For demonstration purposes, here we describe the method for the phage-mediated detection of a known Y. pestis isolate using a luxAB reporter phage constructed from the CDC plague diagnostic phage &#934;A1122 6,7 (Figure 1). A similar method, with minor modifications (e.g. change in growth temperature and media), may be used for the detection of B. anthracis isolates using the B. anthracis reporter phage W&#946;::luxAB 8. The method describes the phage-mediated transduction of a biolumescent phenotype to cultivated Y. pestis cells which are subsequently measured using a microplate luminometer. The major advantages of this method over the traditional phage lysis assays is the ease of use, the rapid results, and the ability to test multiple samples simultaneously in a 96-well microtiter plate format.
<img alt="Figure 1" src="/files/ftp_upload/2740/2740fig1.jpg" /> 
Figure 1. Detection schematic. The phage are mixed with the sample, the phage infects the cell, luxAB are expressed, and the cell bioluminesces. Sample processing is not necessary; the phage and cells are mixed and subsequently measured for light.

A simple method for the identification of priority bacterial pathogens is to use genetically engineered reporter phage. These reporter phage, which are specific to their particular host species, are capable of rapidly transducing a bioluminescent signal response to host cells. Herein, we describe the use of reporter phage for the detection of Yersinia pestis.

Yersinia pestis and Bacillus anthracis are Category A bacterial pathogens that are the causative agents of the plague and anthrax, respectively 1. ...

Immunology and Infection

Bacterial Detection &amp; Identification Using Electrochemical Sensors

Electrochemical sensors are widely used for rapid and accurate measurement of blood glucose and can be adapted for detection of a wide variety of analytes. Electrochemical sensors operate by transducing a biological recognition event into a useful electrical signal. Signal transduction occurs by coupling the activity of a redox enzyme to an amperometric electrode. Sensor specificity is either an inherent characteristic of the enzyme, glucose oxidase in the case of a glucose sensor, or a product of linkage between the enzyme and an antibody or probe.
Here, we describe an electrochemical sensor assay method to directly detect and identify bacteria. In every case, the probes described here are DNA oligonucleotides. This method is based on sandwich hybridization of capture and detector probes with target ribosomal RNA (rRNA). The capture probe is anchored to the sensor surface, while the detector probe is linked to horseradish peroxidase (HRP). When a substrate such as 3,3',5,5'-tetramethylbenzidine (TMB) is added to an electrode with capture-target-detector complexes bound to its surface, the substrate is oxidized by HRP and reduced by the working electrode. This redox cycle results in shuttling of electrons by the substrate from the electrode to HRP, producing current flow in the electrode.

Bacterial Detection & Identification Using Electrochemical Sensors

We describe an electrochemical sensor assay method for rapid bacterial detection and identification. The assay involves a sensor array functionalized with DNA oligonucleotide capture probes for ribosomal RNA (rRNA) species-specific sequences. Sandwich hybridization of target rRNA with the capture probe and a horseradish peroxidase-linked DNA oligonucleotide detector probe produces a measurable amperometric current.

Electrochemical  sensors are widely used for rapid and accurate measurement of blood glucose and  can be adapted for detection of a wide variety of ...

Bioengineering

Detection of Bacteria Using Fluorogenic DNAzymes

Outbreaks linked to food-borne and hospital-acquired pathogens account for millions of deaths and hospitalizations as well as colossal economic losses each and every year. Prevention of such outbreaks and minimization of the impact of an ongoing epidemic place an ever-increasing demand for analytical methods that can accurately identify culprit pathogens at the earliest stage. Although there is a large array of effective methods for pathogen detection, none of them can satisfy all the following five premier requirements embodied for an ideal detection method: high specificity (detecting only the bacterium of interest), high sensitivity (capable of detecting as low as a single live bacterial cell), short time-to-results (minutes to hours), great operational simplicity (no need for lengthy sampling procedures and the use of specialized equipment), and cost effectiveness. For example, classical microbiological methods are highly specific but require a long time (days to weeks) to acquire a definitive result.1 PCR- and antibody-based techniques offer shorter waiting times (hours to days), but they require the use of expensive reagents and/or sophisticated equipment.2-4 Consequently, there is still a great demand for scientific research towards developing innovative bacterial detection methods that offer improved characteristics in one or more of the aforementioned requirements. Our laboratory is interested in examining the potential of DNAzymes as a novel class of molecular probes for biosensing applications including bacterial detection.5
DNAzymes (also known as deoxyribozymes or DNA enzymes) are man-made single-stranded DNA molecules with the capability of catalyzing chemical reactions.6-8 These molecules can be isolated from a vast random-sequence DNA pool (which contains as many as 1016 individual sequences) by a process known as "in vitro selection" or "SELEX" (systematic evolution of ligands by exponential enrichment).9-16 These special DNA molecules have been widely examined in recent years as molecular tools for biosensing applications.6-8
Our laboratory has established in vitro selection procedures for isolating RNA-cleaving fluorescent DNAzymes (RFDs; Fig. 1) and investigated the use of RFDs as analytical tools.17-29 RFDs catalyze the cleavage of a DNA-RNA chimeric substrate at a single ribonucleotide junction (R) that is flanked by a fluorophore (F) and a quencher (Q). The close proximity of F and Q renders the uncleaved substrate minimal fluorescence. However, the cleavage event leads to the separation of F and Q, which is accompanied by significant increase of fluorescence intensity.
More recently, we developed a method of isolating RFDs for bacterial detection.5 These special RFDs were isolated to "light up" in the presence of the crude extracellular mixture (CEM) left behind by a specific type of bacteria in their environment or in the media they are cultured (Fig. 1). The use of crude mixture circumvents the tedious process of purifying and identifying a suitable target from the microbe of interest for biosensor development (which could take months or years to complete). The use of extracellular targets means the assaying procedure is simple because there is no need for steps to obtain intracellular targets.
Using the above approach, we derived an RFD that cleaves its substrate (FS1; Fig. 2A) only in the presence of the CEM produced by E. coli (CEM-EC).5 This E. coli-sensing RFD, named RFD-EC1 (Fig. 2A), was found to be strictly responsive to CEM-EC but nonresponsive to CEMs from a host of other bacteria (Fig. 3).
Here we present the key experimental procedures for setting up E. coli detection assays using RFD-EC1 and representative results.

We have recently reported a novel approach for generating fluorogenic DNAzyme probes that can be applied to set up a simple, "mix-and-read" fluorescent assay for bacterial detection. These special DNA probes catalyze the cleavage of a chromophore-modified DNA-RNA chimeric substrate in the presence of crude extracellular mixture (CEM) produced by a specific bacterium, thereby translating bacterial detection into fluorescence signal generation. In this report we will describe key experimental procedures where a specific DNAzyme probe denoted "RFD-EC1" is employed for the detection of the model bacterium, Escherichia coli (E. coli).

Outbreaks linked to food-borne and hospital-acquired pathogens account for millions of deaths and hospitalizations as well as colossal economic losses ...

Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols to address these questions include colony count assays, gentamicin protection assays, and electron microscopy. Colony count and gentamicin protection assays only assess the viability of the entire bacterial population and are unable to determine individual bacterial viability. Electron microscopy can be used to determine the viability of individual bacteria and provide information regarding their localization in host cells. However, bacteria often display a range of electron densities, making assessment of viability difficult. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells. These assays were developed originally to assess survival of Neisseria gonorrhoeae in primary human neutrophils, but should be applicable to any bacterium-host cell interaction. These protocols combine membrane-permeable fluorescent dyes (SYTO9 and 4&#39;,6-diamidino-2-phenylindole [DAPI]), which stain all bacteria, with membrane-impermeable fluorescent dyes (propidium iodide and SYTOX Green), which are only accessible to nonviable bacteria. Prior to eukaryotic cell permeabilization, an antibody or fluorescent reagent is added to identify extracellular bacteria. Thus these assays discriminate the viability of bacteria adherent to and inside eukaryotic cells. A protocol is also provided for using the viability dyes in combination with fluorescent antibodies to eukaryotic cell markers, in order to determine the subcellular localization of individual bacteria. The bacterial viability dyes discussed in this article are a sensitive complement and/or alternative to traditional microbiology techniques to evaluate the viability of individual bacteria and provide information regarding where bacteria survive in host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols ...

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact cellular functions, including energetics and signal transductions. While both fluorescent and electrophysiological methods, including electrode usage and patch-clamping, have been well developed for measuring these events in eukaryotic cells, methodology for measuring similar events in microorganisms have proven more challenging to develop given their small size in combination with the more complex outer surface of bacteria shielding the membrane. During our studies of death-initiation in Streptococcus pneumoniae (pneumococcus), we wanted to elucidate the role of membrane events, including changes in polarity, integrity, and intracellular ion concentrations. Searching the literature, we found that very few studies exist. Other investigators had monitored radioisotope uptake or equilibrium to measure ion fluxes and membrane potential and a limited number of studies, mostly in Gram-negative organisms, had seen some success using carbocyanine or oxonol fluorescent dyes to measure membrane potential, or loading bacteria with cell-permeant acetoxymethyl (AM) ester versions of ion-sensitive fluorescent indicator dyes. We therefore established and optimized protocols for measuring membrane potential, rupture, and ion-transport in the Gram-positive organism S. pneumoniae. We developed protocols using the bis-oxonol dye DiBAC4(3) and the cell-impermeant dye propidium iodide to measure membrane depolarization and rupture, respectively, as well as methods to optimally load the pneumococci with the AM esters of the ratiometric dyes Fura-2, PBFI, and BCECF to detect changes in intracellular concentrations of Ca2+, K+, and H+, respectively, using a fluorescence-detection plate reader. These protocols are the first of their kind for the pneumococcus and the majority of these dyes have not been used in any other bacterial species. Though our protocols have been optimized for S. pneumoniae, we believe these approaches should form an excellent starting-point for similar studies in other bacterial species.

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Unlike that seen for eukaryotes, there is a paucity of studies that detail membrane depolarization and ion concentration changes in bacteria, primarily as their small size makes conventional methods of measurement difficult. Here, we detail protocols for monitoring such events in the significant Gram-positive pathogen Streptococcus pneumoniae utilizing fluorescence techniques.

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact ...

Visualizing Bacterial Motility Based on a Color Reaction

Bacterial motility is crucial for bacterial pathogenicity, biofilm formation, and drug resistance. Bacterial motility is crucial for the invasion and/or dissemination of many pathogenic species. Therefore, it is important to detect bacterial motility. Bacterial growth conditions, such as oxygen, pH, and temperature, can affect bacterial growth and the expression of bacterial flagella. This can lead to reduced motility or even loss of motility, resulting in the inaccurate evaluation of bacterial motility. Based on the color reaction of 2,3,5-triphenyl tetrazolium chloride (TTC) by intracellular dehydrogenases of living bacteria, TTC was added to traditional semisolid agar for bacterial motility detection. The results showed that this TTC semisolid agar method for the detection of bacterial motility is simple, easy to operate, and does not involve large and expensive instruments. The results also showed that the highest motility was observed in semisolid medium prepared with 0.3% agar. Compared with the traditional semisolid medium, the results are easier to evaluate and more accurate.

Here, we present a protocol to detect bacterial motility based on a color reaction. Key advantages of this method are that it is easy to evaluate and more accurate, and does not require specialized equipment.

Bacterial motility is crucial for bacterial pathogenicity, biofilm formation, and drug resistance. Bacterial motility is crucial for the invasion and/or ...

Water Quality Analysis via Indicator Organisms

Source: Laboratories of Dr. Ian Pepper and Dr. Charles Gerba -The University of Arizona 
Demonstrating Author: Luisa Ikner
Water quality analysis monitors anthropogenic influences such as pollutants, nutrients, pathogens, and any other constituent that can impact the water&#8217;s integrity as a resource. Fecal contamination contributes microbial pathogens that threaten plant, animal, and human health with disease or illness. Increasing water demands and strict quality standards require that water being supplied for human or environmental resources be monitored for low pathogen levels. However, monitoring each pathogen associated with fecal pollution is not feasible, as laboratory techniques involve extensive labor, time, and costs. Therefore, detection for indicator organisms provides a simple, rapid, and cost effective technique to monitor pathogens associated with unsanitary conditions.

Microbial Indicators and Most Probable Number Analysis for Water Quality

Source: Laboratories of Dr. Ian Pepper and Dr. Charles Gerba -The University of Arizona
Demonstrating Author: Luisa Ikner
Water quality analysis monitors ...

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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.
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<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!
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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 ...

Chemistry

Biochemistry

Colorimetric Detection of a Bacterial Biomarker Using a Modified Litmus Test

After preparing E. coli cells according to the text protocol, pre-wash a 1.5-milliliter microfuge tube by adding and vortexing 100 microliters of RB. Then, discard the buffer.
Add 15 microliters of assembled EC1 to the washed tube, then, place the tube on the magnetic rack, and aspirate the supernatant. Remove the tube from the rack, add 100 microliters of RB, and carefully resuspend the magnetic beads.
After using RB to wash the beads two more times and removing the RB, add 10 microliters of the E. coli sample to the tube, and gently mix the contents by tapping on the tube. Then, incubate the reaction at room temperature for one hour.
After the incubation, add 90 microliters of doubly-distilled water, and place the tube on the magnetic rack. Following approximately three minutes of magnetic separation, carefully transfer 85 microliters of the supernatant to a 0.5-milliliter microfuge tube.
It is very important to pipette the solution carefully without taking up any of the magnetic beads. These beads are loaded with urease and will generate a false positive signal.
Add 15 microliters of 0.04% phenol red and 100 microliters of substrate solution. Then, take photographs at specific time intervals to record a color change.

Colorimetric Detection of Bacteria Using Litmus Test

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Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Visualizing Bacterial Motility Based on a Color Reaction

Water Quality Analysis via Indicator Organisms

Photometric Protein Determination

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