This project was supported by University of Alabama at Birmingham Department of Microbiology startup funds and NIH grant R35GM124590 (to MJG), and NIH grant R01AI121364 (to FW).

1. Purifying Yeast Exopolyphosphatase (ScPPX)
<ol>
	<li>Transform the E. coli protein overexpression strain BL21(DE3)31 with plasmid pScPPX26 by electroporation32 or chemical transformation33.</li>
	<li>Inoculate 1 L of lysogeny broth (LB) containing 100 &#181;g mL-1 ampicillin in a 2 L unbaffled flask with a single colony of BL21(DE3) containing pScPPX2 and incubate overnight at 37 &#176;C without shaking, to an absorbance at 600 nm (A600) of approximately 0.3, as measured in a spectrophotometer.</li>
	<li>Start the culture shaking (180 rpm) and incubate for 30 min at 37 &#176;C, during which time the cells will grow to an A600 of 0.4 - 0.5.</li>
	<li>Add isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM and an additional 100 &#181;g mL-1 ampicillin, then incubate for 4 h at 37 &#176;C with shaking at 180 rpm. 
	NOTE: This overexpression protocol generates a large amount of soluble ScPPX. However, ScPPX overexpression is very forgiving, and a variety of other common protein overexpression protocols have been used to successfully purify ScPPX.</li>
	<li>Transfer the cells to a 1 L centrifuge bottle and harvest them by centrifuging for 20 min at 5000 x g at 4 &#176;C. Remove the supernatant and transfer the cell pellet to a 50 mL conical tube. 
	NOTE: Cell pellets can be stored indefinitely at -80 &#176;C.</li>
	<li>Thaw the pellet on ice, then resuspend it in 9 mL of 50 mM HEPES, 0.5 M NaCl, and 5 mM imidazole (pH 8) for a total volume of about 10 mL.</li>
	<li>Add (final concentrations) 1 mg mL-1 lysozyme, 2 mM MgCl2, and 50 units mL-1 of an RNA- and DNA-degrading endonuclease (see Table of Materials) and incubate for 30 min on ice. 
	NOTE: ScPPX binds nucleic acids (data not shown), so nuclease treatment is essential during purification.</li>
	<li>Using a sonicator with a microtip (see Table of Materials), lyse the cells by 2 cycles of sonication on ice at 50% power, pulsing 5 s on and 5 s off, with a 2 min rest between cycles. 
	NOTE: Use appropriate hearing protection during sonication. Similarly to the case with overexpression, a variety of cell lysis methods are acceptable for ScPPX purification.</li>
	<li>Remove insoluble debris by centrifuging the lysate for 20 min at 20,000 x g at 4 &#176;C. Filter the resulting supernatant (approximately 10 mL) through a 0.8 &#181;m pore size cellulose acetate syringe filter.</li>
	<li>Load the lysate onto a nickel-charged 5 mL chelating column (see Table of Materials) using either a peristaltic pump or a large syringe.</li>
	<li>Rinse the column with 50 mL of 50 mM HEPES, 0.5 M NaCl, 5 mM imidazole (pH 8).</li>
	<li>Rinse the column with 50 mL of 50 mM HEPES, 0.5 M NaCl, 20 mM imidazole (pH 8).</li>
	<li>Elute ScPPX with 50 mM HEPES, 0.5 M NaCl, and 0.5 M imidazole (pH 8), collecting 15 1-mL fractions. Test these elution fractions for protein content with the Bradford protein assay34 (see Table of Materials) and run an SDS-PAGE gel35 to confirm which fractions contain purified ScPPX (molecular weight = 45 kDa).</li>
	<li>Pool the fractions containing pure ScPPX and adjust the concentration of the pooled protein to about 2 mg mL-1 with 50 mM HEPES and 0.5 M NaCl. Higher protein concentrations may precipitate in the next step.</li>
	<li>Exchange the purified ScPPX into storage buffer (20 mM Tris-HCl [pH 7.5], 50 mM KCl, 10% [v/v] glycerol) by dialysis. Place pooled ScPPX fractions in a sealed length of 12,000 - 14,000 Da molecular weight cutoff dialysis membrane tubing (see Table of Materials) and suspend in 1 L of storage buffer with continuous stirring at 4 &#176;C for at least 4 h. Repeat this step with fresh storage buffer for a total of 3 buffer changes.</li>
	<li>Transfer the purified, dialyzed protein to a centrifuge tube or tubes and centrifuge for 20 min at 20,000 x g at 4 &#176;C to remove any insoluble aggregates. Carefully transfer the supernatant to a clean 15 mL conical tube.</li>
	<li>Adjust the concentration of the purified ScPPX to 1 mg mL-1 with storage buffer, add 0.1% (w/v) protease-free bovine serum albumin (BSA; final concentration), and store for up to 6 months at 4 &#176;C. 
	NOTE: ScPPX loses more than 90% of its activity when it is frozen13.</li>
</ol>
2. Harvesting Samples for Polyphosphate Extraction
<ol>
	<li>Grow bacteria under the conditions of interest for determining polyP content. For this protocol, grow Lactobacillus reuteri36 overnight at 37 &#176;C without shaking in malic enzyme induction (MEI) medium37 without cysteine (MEI-C).</li>
	<li>Harvest enough cells by centrifugation in a 1.5 mL microcentrifuge tube to total 50 &#8211; 100 &#181;g of cellular protein (see step 3 below). For E. coli, this is 1 mL of a log phase culture at an A600 of 0.2 - 0.4. For L. reuteri, this is 1 mL of an overnight culture. Adjust as necessary for other species of interest.</li>
	<li>Completely remove the supernatant from the cell pellets.</li>
	<li>Resuspend the cell pellets in 250 &#181;L of GITC lysis buffer (4 M guanidine isothiocyanate, 50 mM Tris-HCl, pH 7) and lyse by incubation at 95 &#176;C for 10 min. Store lysates at -80 &#176;C. 
	NOTE: Be consistent with lysis time, since extended incubation at high temperature may result in degradation of polyP by hydrolysis38. Lysates can be stored indefinitely at -80 &#176;C. 
	CAUTION: Guanidine isothiocyanate is a chaotropic salt and should be handled with gloves and disposed of as hazardous waste.</li>
</ol>
3. Measuring the Protein Content of Cell Lysates
<ol>
	<li>Prepare BSA standards containing 0, 0.1, 0.2, and 0.4 mg mL-1 of BSA in GITC lysis buffer. 
	NOTE: It is important to make the BSA standards in GITC, since GITC influences the color development of the Bradford assay. BSA standards can be stored indefinitely at -20 &#176;C.</li>
	<li>Aliquot 5 &#181;L of thawed, well-mixed cell lysates and of BSA standards to separate wells in a clear 96-well plate.</li>
	<li>Add 195 &#181;L of Bradford reagent34 to each well and measure absorbance at 595 nm (A595) in a plate reader (see Table of Materials).</li>
	<li>Calculate the amount of protein in each well by comparison to the BSA standard curve, using the formula y = mx + b, where y is A595, x is &#181;g of BSA, m is the slope of the standard curve, and b is the y-intercept of the standard curve. Multiply the resulting value by 0.05 to determine the total mg of protein in each sample.</li>
</ol>
4. Extracting Polyphosphate
<ol>
	<li>Add 250 &#181;L of 95% ethanol to each GITC-lysed sample and vortex to mix.</li>
	<li>Apply that mixture to a silica membrane spin column and centrifuge for 30 s at 16,100 x g.</li>
	<li>Discard the flow-through, then add 750 &#181;L of 5 mM Tris-HCl (pH 7.5), 50 mM NaCl, 5 mM EDTA, 50% ethanol and centrifuge for 30 s at 16,100 x g.</li>
	<li>Discard the flow-through and centrifuge for 2 min at 16,100 x g.</li>
	<li>Place the column in a clean 1.5 mL microfuge tube and add 150 &#181;L of 50 mM Tris-HCl (pH 8).</li>
	<li>Incubate at room temperature for 5 min, then elute polyP by centrifuging for 2 min at 8,000 x g. 
	NOTE: If desired, the eluates can be stored at -20 &#176;C for at least 1 week.</li>
</ol>
5. Digesting Polyphosphate
<ol>
	<li>Prepare standards containing 0, 5, 50, or 200 &#181;M potassium phosphate in 50 mM Tris-HCl (pH 8). 
	NOTE: Potassium phosphate standards can be stored indefinitely at room temperature.</li>
	<li>Aliquot 100 &#181;L of each phosphate standard and of extracted polyP samples into separate wells of a clear 96-well plate.</li>
	<li>Prepare a master mix containing (per sample): 30 &#181;L of 5x ScPPX reaction buffer (100 mM Tris-HCl, 25 mM MgCl2, 250 mM ammonium acetate, pH 7.5)13, 19 &#181;L of H2O, and 1 &#181;L of purified ScPPX (1 mg mL-1).</li>
	<li>Add 50 &#181;L of the master mix to each well of the 96-well plate. Incubate for 15 min at 37 &#176;C. 
	NOTE: If desired, the digested polyP samples can be stored at -20 &#176;C indefinitely.</li>
</ol>
6. Detecting Free Phosphate24
<ol>
	<li>Prepare detection solution base by dissolving 0.185 g of antimony potassium tartrate in 200 mL of water, adding 150 mL of 4 N H2SO4, then adding 1.49 g of ammonium heptamolybdate. Stir to dissolve and then bring to a final volume of 456 mL. Filter the solution to remove particulates and store protected from light at 4 &#176;C for up to 1 month.</li>
	<li>Prepare a stock solution of 1 M ascorbic acid. Store protected from light at 4 &#176;C for up to 1 month.</li>
	<li>Prepare a fresh working stock of detection solution each day by mixing 9.12 mL of detection solution base with 0.88 mL of 1 M ascorbic acid. Allow the detection solution to come to room temperature before use.</li>
	<li>Add 50 &#181;L of detection solution to each sample and standard in the 96-well plate and incubate at room temperature for about 2 min to allow color development.</li>
	<li>Measure absorbance at 882 nm with a plate reader and calculate the phosphate concentration of each sample by comparison to the potassium phosphate standard curve. 
	CAUTION: The detection solution contains toxic salts and strong acids. Wear gloves and treat excess solution as toxic waste.</li>
</ol>
7. Calculating Cellular Polyphosphate Content
<ol>
	<li>Convert the phosphate concentrations determined in step 6.5 to nanomoles of polyP-derived phosphate in each entire cell lysate according to the following formula: 
	&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;nmol polyP = 1.5 x (&#181;M phosphate / 10) 
	NOTE: The standard curve-based method in step 6 determines the concentration of phosphate (in &#181;M) in each 100 &#181;L polyP sample aliquoted in step 5.2. To convert this concentration to a number of nmol, 100 &#181;L is divided by 106 to give a volume in L, multiplied by the concentration (&#181;mol L-1), then multiplied by 1,000 (the number of nmol in a &#181;mol). This reduces to dividing the concentration by 10. The total extract volume prepared in step 4 is 150 &#181;L, so it is necessary to multiply the resulting value by 1.5 to calculate the nmol of phosphate present in the entire extract.</li>
	<li>Normalize cellular polyP content to total cellular protein by dividing nmol polyP by the mg total protein in each sample determined in step 3.4. PolyP levels are expressed in terms of the concentration of individual polyP-derived free phosphate. 
	NOTE: In some cases, the amount of polyP-derived phosphate present in a sample may fall outside the linear range of the phosphate standard curve (step 6.5). If the levels of polyP are very high, the excess polyP-containing eluate from step 4.6 can be diluted 1:10 or 1:100, as necessary, and then measured again as described in steps 5 through 7.</li>
</ol>

The authors have nothing to disclose.

The protocol described here simplifies and accelerates quantification of polyP levels in diverse bacteria, with a typical set of 24 samples taking about 1.5 h to fully process. This permits rapid screening of samples and analysis of mutant libraries, and simplifies kinetic experiments measuring the accumulation of polyP over time. We have demonstrated that the protocol works effectively on representatives of three different phyla: proteobacteria, firmicutes, and actinobacteria, which are notorious for their resilient, di...

Inorganic polyphosphate (polyP) is a linear biopolymer of phosphoanhydride-linked phosphate units that is found in all domains of life1,2,3. In diverse bacteria, polyP is essential for stress response, motility, biofilm formation, cell cycle control, antibiotic resistance, and virulence4,5,6,7,8,9,10,11. Studies of polyP metabolism in bacteria therefore have the potential to yield fundamental insights into the ability of bacteria to cause disease and thrive in diverse environments. In many cases, however, the methods available for quantifying polyP in bacterial cells are a limiting factor in these studies.
There are several methods currently used to measure polyP levels in biological materials. These methods typically involve two distinct steps: extracting polyP and quantifying the polyP present in those extracts. The current gold standard method, developed for the yeast Saccharomyces cerevisiae by Bru and colleagues12, extracts polyP along with DNA and RNA using phenol and chloroform, followed by ethanol precipitation, treatment with deoxyribonuclease (DNase) and ribonuclease (RNase), and digestion of the resulting purified polyP with the S. cerevisiae polyP-degrading enzyme exopolyphosphatase (ScPPX)13 to yield free phosphate, which is then quantified using a malachite green-based colorimetric assay. This procedure is highly quantitative but labor-intensive, limiting the number of samples that can be processed in a single experiment, and is not optimized for bacterial samples. Others have reported extracting polyP from a variety of cells and tissues using silica beads (&#34;glassmilk&#34;) or silica membrane columns6,14,15,16,17,18. These methods do not efficiently extract short chain polyP (less than 60 phosphate units)12,14,15, although this is of less concern for bacteria, which are generally thought to synthesize primarily long-chain polyP3. Older methods of polyP extraction using strong acids19,20 are no longer widely used, since polyP is unstable under acidic conditions12.
There are also a variety of reported methods for quantifying polyP. Among the most common is 4&#8242;,6-diamidino-2-phenylindole (DAPI), a fluorescent dye more typically used to stain DNA. DAPI-polyP complexes have different fluorescence excitation and emission maxima than DAPI-DNA complexes21,22, but there is considerable interference from other cellular components, including RNA, nucleotides, and inositol phosphates12,15,16,23, reducing the specificity and sensitivity of polyP measurements made using this method. Alternatively, polyP and adenosine diphosphate (ADP) can be converted into adenosine triphosphate (ATP) using purified Escherichia coli polyP kinase (PPK) and the resulting ATP quantified using luciferase14,17,18. This allows the detection of very small amounts of polyP, but requires two enzymatic reaction steps and both luciferin and very pure ADP, which are expensive reagents. ScPPX specifically digests polyP into free phosphate6,12,13,24, which can be detected using simpler methods, but ScPPX is inhibited by DNA and RNA12, necessitating DNase and RNase treatment of polyP-containing extracts. Neither PPK nor ScPPX are commercially available, and PPK purification is relatively complex25,26.
PolyP in cell lysates or extracts can also be visualized on polyacrylamide gels by DAPI negative staining27,28,29,30, a method that does allow assessment of chain length, but is low-throughput and poorly quantitative.
We now report a fast, inexpensive, medium-throughput polyP assay that allows rapid quantification of polyP levels in diverse bacterial species. This method begins by lysing bacterial cells at 95 &#176;C in 4 M guanidine isothiocyanate (GITC)14 to inactivate cellular phosphatases, followed by a silica membrane column extraction optimized for rapid processing of multiple samples. The resulting polyP-containing extract is then digested with a large excess of ScPPX, eliminating the need for DNase and RNase treatment. We include a protocol for straightforward nickel affinity purification of mg quantities of ScPPX. Finally, polyP-derived free phosphate is quantified with a simple, sensitive, ascorbic acid-based colorimetric assay24 and normalized to total cellular protein. This method streamlines the measurement of polyP in bacterial cells, and we demonstrate its use with representative species of Gram-negative bacteria, Gram-positive bacteria, and mycobacteria.

<table>
	<tbody>
		<tr>
			<td>E. coli BL21(DE3)</td>
			<td>Millipore Sigma</td>
			<td>69450</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid pScPPX2</td>
			<td>Addgene</td>
			<td>112877</td>
			<td>available to academic and other non-profit institutions</td>
		</tr>
		<tr>
			<td>LB broth</td>
			<td>Fisher Scientific</td>
			<td>BP1427-2</td>
			<td>E. coli growth medium</td>
		</tr>
		<tr>
			<td>ampicillin</td>
			<td>Fisher Scientific</td>
			<td>BP176025</td>
			<td></td>
		</tr>
		<tr>
			<td>isopropyl &#946;-D-1-thiogalactopyranoside (IPTG)</td>
			<td>Gold Biotechnology</td>
			<td>I2481C</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES buffer</td>
			<td>Gold Biotechnology</td>
			<td>H-400-1</td>
			<td></td>
		</tr>
		<tr>
			<td>potassium hydroxide (KOH)</td>
			<td>Fisher Scientific</td>
			<td>P250500</td>
			<td>for adjusting the pH of HEPES-buffered solutions</td>
		</tr>
		<tr>
			<td>sodium chloride (NaCl)</td>
			<td>Fisher Scientific</td>
			<td>S27110</td>
			<td></td>
		</tr>
		<tr>
			<td>imidazole</td>
			<td>Fisher Scientific</td>
			<td>O3196500</td>
			<td></td>
		</tr>
		<tr>
			<td>lysozyme</td>
			<td>Fisher Scientific</td>
			<td>AAJ6070106</td>
			<td></td>
		</tr>
		<tr>
			<td>magnesium chloride (MgCl2)</td>
			<td>Fisher Scientific</td>
			<td>BP214-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Pierce Universal Nuclease</td>
			<td>Fisher Scientific</td>
			<td>PI88700</td>
			<td>Benzonase (Sigma-Aldrich cat. # E1014) is an acceptable substitute</td>
		</tr>
		<tr>
			<td>Model 120 Sonic Dismembrator</td>
			<td>Fisher Scientific</td>
			<td>FB-120</td>
			<td>other cell lysis methods (e.g. French Press) can also be effective</td>
		</tr>
		<tr>
			<td>5 mL HiTrap chelating HP column</td>
			<td>GE Life Sciences</td>
			<td>17040901</td>
			<td>any nickel-affinity chromatography column or resin could be substituted</td>
		</tr>
		<tr>
			<td>nickel(II) sulfate hexahydrate</td>
			<td>Fisher Scientific</td>
			<td>AC415611000</td>
			<td>for charging HiTrap column</td>
		</tr>
		<tr>
			<td>0.8 &#181;m pore size cellulose acetate syringe filters</td>
			<td>Fisher Scientific</td>
			<td>09-302-168</td>
			<td></td>
		</tr>
		<tr>
			<td>Bradford reagent</td>
			<td>Bio-Rad</td>
			<td>5000205</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris buffer</td>
			<td>Fisher Scientific</td>
			<td>BP1525</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectrum&#160;Spectra/Por 4 RC Dialysis Membrane Tubing 12,000 to 14,000 Dalton MWCO</td>
			<td>Fisher Scientific</td>
			<td>08-667B</td>
			<td>other dialysis membranes with MWCO &#60; 30,000 Da should also work</td>
		</tr>
		<tr>
			<td>hydrochloric acid (HCl)</td>
			<td>Fisher Scientific</td>
			<td>A144-212</td>
			<td>for adjusting the pH of Tris-buffered solutions</td>
		</tr>
		<tr>
			<td>potassium chloride (KCl)</td>
			<td>Fisher Scientific</td>
			<td>P217500</td>
			<td></td>
		</tr>
		<tr>
			<td>glycerol</td>
			<td>Fisher Scientific</td>
			<td>BP2294</td>
			<td></td>
		</tr>
		<tr>
			<td>10x MOPS medium mixture</td>
			<td>Teknova</td>
			<td>M2101</td>
			<td>E. coli growth medium</td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>Fisher Scientific</td>
			<td>D161</td>
			<td></td>
		</tr>
		<tr>
			<td>monobasic potassium phosphate (KH2PO4)</td>
			<td>Fisher Scientific</td>
			<td>BP362-500</td>
			<td></td>
		</tr>
		<tr>
			<td>dibasic potassium phosphate (K2HPO4)</td>
			<td>Fisher Scientific</td>
			<td>BP363-500</td>
			<td></td>
		</tr>
		<tr>
			<td>dehydrated yeast extract</td>
			<td>Fisher Scientific</td>
			<td>DF0886-17-0</td>
			<td></td>
		</tr>
		<tr>
			<td>tryptone</td>
			<td>Fisher Scientific</td>
			<td>BP1421-500</td>
			<td></td>
		</tr>
		<tr>
			<td>magnesium sulfate heptahydrate</td>
			<td>Fisher Scientific</td>
			<td>M63-50</td>
			<td></td>
		</tr>
		<tr>
			<td>manganese sulfate monohydrate</td>
			<td>Fisher Scientific</td>
			<td>M113-500</td>
			<td></td>
		</tr>
		<tr>
			<td>guanidine isothiocyanate</td>
			<td>Fisher Scientific</td>
			<td>BP221-250</td>
			<td></td>
		</tr>
		<tr>
			<td>bovine serum albumin (protease-free)</td>
			<td>Fisher Scientific</td>
			<td>BP9703100</td>
			<td></td>
		</tr>
		<tr>
			<td>clear flat bottom 96-well plates</td>
			<td>Sigma-Aldrich</td>
			<td>M0812-100EA</td>
			<td>any clear 96-well plate will work</td>
		</tr>
		<tr>
			<td>Tecan M1000 Infinite plate reader</td>
			<td>Tecan, Inc.</td>
			<td>not applicable</td>
			<td>any plate reader capable of measuring absorbance at 595 and 882 nm will work</td>
		</tr>
		<tr>
			<td>ethanol</td>
			<td>Fisher Scientific</td>
			<td>04-355-451</td>
			<td></td>
		</tr>
		<tr>
			<td>silica membrane spin columns</td>
			<td>Epoch Life Science</td>
			<td>1910-050/250</td>
			<td></td>
		</tr>
		<tr>
			<td>ethylenediaminetetraacetic acid (EDTA)</td>
			<td>Fisher Scientific</td>
			<td>BP120500</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL microfuge tubes</td>
			<td>Fisher Scientific</td>
			<td>NC9580154</td>
			<td></td>
		</tr>
		<tr>
			<td>ammonium acetate</td>
			<td>Fisher Scientific</td>
			<td>A637-500</td>
			<td></td>
		</tr>
		<tr>
			<td>antimony potassium tartrate</td>
			<td>Fisher Scientific</td>
			<td>AAA1088922</td>
			<td></td>
		</tr>
		<tr>
			<td>4 N sulfuric acid (H2SO4)</td>
			<td>Fisher Scientific</td>
			<td>SA818-500</td>
			<td></td>
		</tr>
		<tr>
			<td>ammonium heptamolybdate</td>
			<td>Fisher Scientific</td>
			<td>AAA1376630</td>
			<td></td>
		</tr>
		<tr>
			<td>ascorbic acid</td>
			<td>Fisher Scientific</td>
			<td>AC401471000</td>
			<td></td>
		</tr>
	</tbody>
</table>

assaying for inorganic polyphosphate in bacteria

Inorganic polyphosphate (polyP) is a biological polymer found in cells from all domains of life, and is required for virulence and stress response in many bacteria. There are a variety of methods for quantifying polyP in biological materials, many of which are either labor-intensive or insensitive, limiting their usefulness. We present here a streamlined method for polyP quantification in bacteria, using a silica membrane column extraction optimized for rapid processing of multiple samples, digestion of polyP with the polyP-specific exopolyphosphatase ScPPX, and detection of the resulting free phosphate with a sensitive ascorbic acid-based colorimetric assay. This procedure is straightforward, inexpensive, and allows reliable polyP quantification in diverse bacterial species. We present representative polyP quantification from the Gram-negative bacterium (Escherichia coli), the Gram-positive lactic acid bacterium (Lactobacillus reuteri), and the mycobacterial species (Mycobacterium smegmatis). We also include a simple protocol for nickel affinity purification of mg quantities of ScPPX, which is not currently commercially available.

The key steps of the protocol are diagrammed in simplified form in Figure 1.
To demonstrate the use of this protocol with Gram-negative bacteria, wild-type E. coli MG165539 was grown to mid-log phase in LB rich medium at 37 &#176;C with shaking (200 rpm), then rinsed and incubated for an additional 2 h in morpholinopropanesulfonate-buffered (MOPS) minimal medium<sup ...

Watch this Scientific Journal Video about Assaying for Inorganic Polyphosphate in Bacteria at JoVE.com

Assaying for Inorganic Polyphosphate in Bacteria

We describe a simple method for rapid quantification of inorganic polyphosphate in diverse bacteria, including Gram-negative, Gram-positive, and mycobacterial species.

Inorganic polyphosphate (polyP) is a biological polymer found in cells from all domains of life, and is required for virulence and stress response in many ...

assaying-for-inorganic-polyphosphate-in-bacteria

Research

JoVE Journal

Immunology and Infection

8.5K Views.  University of Alabama at Birmingham. We describe a simple method for rapid quantification of inorganic polyphosphate in diverse bacteria, including Gram-negative, Gram-positive, and mycobacterial species.

Video: Assaying for Inorganic Polyphosphate in Bacteria

Genotypic Inference of HIV-1 Tropism Using Population-based Sequencing of V3

Background: Prior to receiving a drug from CCR5-antagonist class in HIV therapy, a patient must undergo an HIV tropism test to confirm that his or her viral population uses the CCR5 coreceptor for cellular entry, and not an alternative coreceptor. One approach to tropism testing is to examine the sequence of the V3 region of the HIV envelope, which interacts with the coreceptor.
Methods: Viral RNA is extracted from blood plasma. The V3 region is amplified in triplicate with nested reverse transcriptase-PCR. The amplifications are then sequenced and analyzed using the software, RE_Call. Sequences are then submitted to a bioinformatic algorithm such as geno2pheno to infer viral tropism from the V3 region. Sequences are inferred to be non-R5 if their geno2pheno false positive rate falls below 5.75%. If any one of the three sequences from a sample is inferred to be non-R5, the patient is unlikely to respond to a CCR5-antagonist.

HIV tropism can be inferred from the V3 region of the viral envelope. V3 is PCR amplified in triplicate using nested RT-PCR, sequenced, and interpreted using bioinformatic software. Samples with with 1 or more sequence(s) with low g2P scores are classified as non-R5 virus.

Background: Prior to receiving a drug from CCR5-antagonist class in HIV therapy, a patient must undergo an HIV tropism test to confirm that his or her ...

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 ...

Tractable Mammalian Cell Infections with Protozoan-primed Bacteria

Many intracellular bacterial pathogens use freshwater protozoans as a natural reservoir for proliferation in the environment. Legionella pneumophila, the causative agent of Legionnaires' pneumonia, gains a pathogenic advantage over in vitro cultured bacteria when first harvested from protozoan cells prior to infection of mammalian macrophages. This suggests that important virulence factors may not be properly expressed in vitro. We have developed a tractable system for priming L. pneumophila through its natural protozoan host Acanthamoeba castellanii prior to mammalian cell infection. The contribution of any virulence factor can be examined by comparing intracellular growth of a mutant strain to wild-type bacteria after protozoan priming. GFP-expressing wild-type and mutant L. pneumophila strains are used to infect protozoan monolayers in a priming step and allowed to reach late stages of intracellular growth. Fluorescent bacteria are then harvested from these infected cells and normalized by spectrophotometry to generate comparable numbers of bacteria for a subsequent infection into mammalian macrophages. For quantification, live bacteria are monitored after infection using fluorescence microscopy, flow cytometry, and by colony plating. This technique highlights and relies on the contribution of host cell-dependent gene expression by mimicking the environment that would be encountered in a natural acquisition route. This approach can be modified to accommodate any bacterium that uses an intermediary host as a means for gaining a pathogenic advantage.

This technique provides a method to harvest, normalize and quantify intracellular growth of bacterial pathogens that are pre-cultivated in natural protozoan host cells prior to infections of mammalian cells. This method can be modified to accommodate a wide variety of host cells for the priming stage as well as target cell types.

Many intracellular bacterial pathogens use freshwater protozoans as a natural reservoir for proliferation in the environment. Legionella pneumophila, the ...

Measuring Growth and Gene Expression Dynamics of Tumor-Targeted S. Typhimurium Bacteria

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium bacterial colonies growing inside tumors. 
We generated model xenograft tumors in mice by subcutaneous injection of a human ovarian cancer cell line, OVCAR-8 (NCI DCTD Tumor Repository, Frederick, MD). 
We transformed attenuated strains of S. typhimurium bacteria (ELH430:SL1344 phoPQ- 1) with a constitutively expressed luciferase (luxCDABE) plasmid for visualization2. These strains specifically colonize tumors while remaining essentially non-virulent to the mouse1. 
Once measurable tumors were established, bacteria were injected intravenously via the tail vein with varying dosage. Tumor-localized, bacterial gene expression was monitored in real time over the course of 60 hours using an in vivo imaging system (IVIS). At each time point, tumors were excised, homogenized, and plated to quantitate bacterial colonies for correlation with gene expression data. 
Together, this data yields a quantitative measure of the in vivo growth and gene expression dynamics of bacteria growing inside tumors.

Measuring Growth and Gene Expression Dynamics of Tumor-Targeted S. Typhimurium Bacteria

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium bacterial colonies growing inside tumors. This video covers tumor cell preparation and implantation, bacteria preparation and injection, whole-animal luminescence imaging, tumor excision, and bacterial colony counting.

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium ...

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 ...

Fluorescence in situ Hybridizations (FISH) for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells and can be further modified to visualize other components in the cell such as infection with viruses and bacteria. Spatial localization and visualization of viruses and bacteria during the infection process is an essential step that complements expression profiling experiments such as microarrays and RNAseq in response to different stimuli. Understanding the spatiotemporal infections with these agents complements biological experiments aimed at understanding their interaction with cellular components. Several techniques for visualizing viruses and bacteria such as reporter gene systems or immunohistochemical methods are time-consuming, and some are limited to work with model organisms&#160;and involve complex methodologies. FISH that targets RNA or DNA species in the cell is a relatively easy and fast method for studying spatiotemporal localization of genes and for diagnostic purposes. This method can be robust and relatively easy to implement when the protocols employ short hybridizing, commercially-purchased probes, which are not expensive. This is particularly robust when sample preparation, fixation, hybridization, and microscopic visualization do not involve complex steps. Here we describe a protocol for localization of bacteria and viruses in insect and plant tissues. The method is based on simple preparation, fixation, and hybridization of insect whole mounts and dissected organs or hand-made plant sections, with 20 base pairs short DNA probes conjugated to fluorescent dyes on their 5&#39; or 3&#39; ends. This protocol has been successfully applied to a number of insect and plant tissues, and can be used to analyze expression of mRNAs or other RNA or DNA species in the cell.

Fluorescence in situ Hybridizations FISH for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

We describe here a simple fluorescence in situ hybridization (FISH) method for the localization of viruses and bacteria in insect and plant tissues. This protocol can be extended for the visualization of mRNA in whole mount and microscopic sections.

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells ...

Following in Real Time the Impact of Pneumococcal Virulence Factors in an Acute Mouse Pneumonia Model Using Bioluminescent Bacteria

Pneumonia is one of the major health care problems in developing and industrialized countries and is associated with considerable morbidity and mortality. Despite advances in knowledge of this illness, the availability of intensive care units (ICU), and the use of potent antimicrobial agents and effective vaccines, the mortality rates remain high1. Streptococcus pneumoniae is the leading pathogen of community-acquired pneumonia (CAP) and one of the most common causes of bacteremia in humans. This pathogen is equipped with an armamentarium of surface-exposed adhesins and virulence factors contributing to pneumonia and invasive pneumococcal disease (IPD). The assessment of the in vivo role of bacterial fitness or virulence factors is of utmost importance to unravel S. pneumoniae pathogenicity mechanisms. Murine models of pneumonia, bacteremia, and meningitis are being used to determine the impact of pneumococcal factors at different stages of the infection. Here we describe a protocol to monitor in real-time pneumococcal dissemination in mice after intranasal or intraperitoneal infections with bioluminescent bacteria. The results show the multiplication and dissemination of pneumococci in the lower respiratory tract and blood, which can be visualized and evaluated using an imaging system and the accompanying analysis software.

Streptococcus pneumoniae is the leading pathogen causing severe community-acquired pneumonia and responsible for over 2&#160;million deaths worldwide. The impact of bacterial factors implicated in fitness or virulence can be monitored in real-time in an acute mouse pneumonia or bacteremia model using bioluminescent bacteria.

Pneumonia is one of the major health care problems in developing and industrialized countries and is associated with considerable morbidity and mortality. ...

Rapid Identification of Gram Negative Bacteria from Blood Culture Broth Using MALDI-TOF Mass Spectrometry

An important role of the clinical microbiology laboratory is to provide rapid identification of bacteria causing bloodstream infection. Traditional identification requires the sub-culture of signaled blood culture broth with identification available only after colonies on solid agar have matured. MALDI-TOF MS is a reliable, rapid method for identification of the majority of clinically relevant bacteria when applied to colonies on solid media. The application of MALDI-TOF MS directly to blood culture broth is an attractive approach as it has potential to accelerate species identification of bacteria and improve clinical management. However, an important problem to overcome is the pre-analysis removal of interfering resins, proteins and hemoglobin&#160;contained in blood culture specimens which, if not removed, interfere with the MS spectra and can result in insufficient or low discrimination identification scores. In addition it is necessary to concentrate bacteria to develop spectra of sufficient quality. The presented method describes the concentration, purification, and extraction of Gram negative bacteria allowing for the early identification of bacteria from a signaled blood culture broth.

The application of matrix-assisted laser desorption/ionization time of flight (MALDI-TOF) mass spectrometry (MS) directly to blood culture broth expedites the identification of bacteria. The presented method is a rapid and reliable method for identification of Gram negative bacteria directly from blood culture broth.

An important role of the clinical microbiology laboratory is to provide rapid identification of bacteria causing bloodstream infection. Traditional ...

A Reference Broth Microdilution Method for Dalbavancin In Vitro Susceptibility Testing of Bacteria that Grow Aerobically

Antimicrobial susceptibility testing (AST) is performed to assess the in vitro activity of antimicrobial agents against various bacteria. The AST results, which are expressed as minimum inhibitory concentrations (MICs) are used in research for antimicrobial development and monitoring of resistance development and in the clinical setting for antimicrobial therapy guidance. Dalbavancin is a semi-synthetic lipoglycopeptide antimicrobial agent that was approved in May 2014 by the Food and Drug Administration (FDA) for the treatment of acute bacterial skin and skin structure infections caused by Gram-positive organisms. The advantage of dalbavancin over current anti-staphylococcal therapies is its long half-life, which allows for once-weekly dosing. Dalbavancin has activity against Staphylococcus aureus (including both methicillin-susceptible S. aureus [MSSA] and methicillin-resistant S. aureus [MRSA]), coagulase-negative staphylococci, Streptococcus pneumoniae, Streptococcus anginosus group, &#946;-hemolytic streptococci and vancomycin susceptible enterococci. Similar to other recent lipoglycopeptide agents, optimization of CLSI and ISO broth susceptibility test methods includes the use of dimethyl sulfoxide (DMSO) as a solvent when preparing stock solutions and polysorbate 80 (P80) to alleviate adherence of the agent to plastic. Prior to the clinical studies and during the initial development of dalbavancin, susceptibility studies were not performed with the use of P-80 and MIC results tended to be 2-4 fold higher and similarly higher MIC results were obtained with the agar dilution susceptibility method. Dalbavancin was first included in CLSI broth microdilution methodology tables in 2005 and amended in 2006 to clarify use of DMSO and P-80. The broth microdilution (BMD) procedure shown here is specific to dalbavancin and is in accordance with the CLSI and ISO methods, with step-by-step detail and focus on the critical steps added for clarity.

A Reference Broth Microdilution Method for Dalbavancin In Vitro Susceptibility Testing of Bacteria that Grow Aerobically

Here, we present a protocol for determination of dalbavancin susceptibility of clinically relevant Gram-positive bacteria using a broth microdilution reference methodology.

Antimicrobial susceptibility testing (AST) is performed to assess the in vitro activity of antimicrobial agents against various bacteria. The AST results, ...

Biomimetic Materials to Characterize Bacteria-host Interactions

Bacterial attachment to host cells is one of the earliest events during bacterial colonization of host tissues and thus a key step during infection. The biochemical and functional characterization of adhesins mediating these initial bacteria-host interactions is often compromised by the presence of other bacterial factors, such as cell wall components or secreted molecules, which interfere with the analysis. This protocol describes the production and use of biomimetic materials, consisting of pure recombinant adhesins chemically coupled to commercially available, functionalized polystyrene beads, which have been used successfully to dissect the biochemical and functional interactions between individual bacterial adhesins and host cell receptors. Protocols for different coupling chemistries, allowing directional immobilization of recombinant adhesins on polymer scaffolds, and for assessment of the coupling efficiency of the resulting &#8220;bacteriomimetic&#8221; materials are also discussed. We further describe how these materials can be used as a tool to inhibit pathogen mediated cytotoxicity and discuss scope, limitations and further applications of this approach in studying bacterial - host interactions.

Bacterial attachment to host cells is a key step during host colonization and infection. This protocol describes the generation of polymer-coupled recombinant adhesins as biomimetic materials which allow analysis of the contribution of individual adhesins to these processes, independent of other bacterial factors.

Bacterial attachment to host cells is one of the earliest events during bacterial colonization of host tissues and thus a key step during infection. The ...