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, difficult&#160;to lyse cell walls49.
Detection of polyP by digestion with ScPPX is more specific and more sensitive than fluorescence detection with DAPI, and is cheaper and&#160;more technically straightforward than conversion of polyP to ATP using PPK. Although ScPPX is inhibited by DNA and RNA12, we have overcome this by using a very large excess of enzyme (1 &#181;g per sample) to achieve complete digestion even in bacteria containing more than 6,000 nmol polyP mg-1 protein29. Other published methods use 10 to 15 ng of ScPPX per reaction12,24. Our protocol for ScPPX purification typically yields more than 5 mg of pure ScPPX per liter of overexpression culture, which is enough for more than 5,000 assays. We have found that several different protocols for nickel-affinity purification of His-tagged proteins work well to purify ScPPX overexpressed from pScPPX2 (data not shown), and there are a wide variety of such protocols available to suit labs with varying technical capacities.
Previous protocols using ScPPX digestion for polyP detection have often relied on malachite green-based colorimetric assays to quantify free phosphate6,12. These assays are sensitive, but typically require carefully timed sequential addition of two or three separate reagents to make accurate measurements24,50,51. The ascorbic acid-based detection system used here24 has a wider linear range of detection than malachite green with no loss of sensitivity, only involves addition of a single reagent, and does not require precise timing.
There are several steps that are critical to the success of this protocol. First, bacterial samples should be lysed in GITC immediately after harvesting, to ensure that polyP levels do not change after the time of sampling and to rapidly inactivate cellular phosphatases. Second, do not incubate GITC lysates at 95 &#176;C for more than 10 min, and store them immediately afterwards at -80 &#176;C to minimize possible polyP hydrolysis. Third, be careful when pipetting samples into 96-well plates for either protein or phosphate measurements not to splash or drip anything from one sample into another. The colorimetric assays, particularly for phosphate, are very sensitive, and small amounts of contamination can strongly skew the results. Finally, some reagents used in this protocol (i.e. ScPPX, detection solution) have limited shelf lives. Prepare fresh ScPPX every 6 months and fresh detection solution every month.
One limitation of our method is that silica columns do not bind polyP less than 60 phosphate units long very well12,14,15. Although bacteria typically synthesize mostly long-chain polyP3, we recommend preliminary experiments using polyacrylamide gel electrophoresis27,30 to assess chain length in a given bacterial species before using our procedure. For species where short-chain polyP makes up a substantial fraction of the polyP pool, our protocol is not appropriate and we recommend extracting polyP by a different method (e.g., phenol-chloroform extraction12), and would also recommend supplementing ScPPX digestion with commercially available S. cereviseae pyrophosphatase (PPA1)24, since ScPPX does not digest pyrophosphate13 and this has a proportionally higher impact on measurements of shorter-chain polyP. As an alternative to ScPPX, acid hydrolysis38 could be used to hydrolyze polyP to free phosphate, but this would require additional DNase, RNase, and purification steps to ensure that only polyP-derived phosphate was being measured. In some cases, it may be useful to use an alternative extraction method to test whether the efficiency of polyP extraction with GITC varies between strains or conditions, particularly with bacteria known to have thick cell walls, such as mycobacteria. If levels of polyP below the limit of detection of this assay are important for studies of a particular species, it may be necessary to use a different detection scheme, such as using PPK to convert polyP to ATP, which can be detected extremely sensitively using commercially available luciferase-based assays14,17,18.

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>&lt;em&gt;E. coli &lt;/em&gt;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>&lt;em&gt;E. coli&lt;/em&gt; growth medium</td></tr><tr><td>ampicillin</td><td>Fisher Scientific</td><td>BP176025</td><td></td></tr><tr><td>isopropyl &amp;beta;-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 (MgCl&lt;sub&gt;2&lt;/sub&gt;)</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 &amp;micro;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&amp;nbsp;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 &amp;lt; 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>&lt;em&gt;E. coli&lt;/em&gt; growth medium</td></tr><tr><td>glucose</td><td>Fisher Scientific</td><td>D161</td><td></td></tr><tr><td>monobasic potassium phosphate (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Fisher Scientific</td><td>BP362-500</td><td></td></tr><tr><td>dibasic potassium phosphate (K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</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 (H&lt;sub&gt;2&lt;/sub&gt;SO4)</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 medium40 containing 4 g L-1 glucose and 0.1 mM K2HPO4, a condition known to induce production of polyP14,29. As shown in Figure 2A, we detected no polyP in wild-type E. coli grown in LB and 192 &#177; 14 (mean &#177; standard deviation [SD]) nmol polyP mg-1 total protein in MOPS, consistent with previous reports14,29. As expected, a &#8710;ppk mutant6, which lacks polyP kinase41, produced no polyP in either medium, a &#8710;ppx mutant6, which lacks exopolyphosphatase42, produced approximately the same amount of polyP as the wild-type, and a &#8710;phoB mutant29, which is defective in phosphate transport, produced significantly less polyP than the wild-type14,29,43.
To demonstrate the use of this protocol with Gram-positive bacteria, wild-type L. reuteri ATCC PTA 647536 and an isogenic ppk1 null mutant lacking polyP kinase, constructed using oligo-directed recombineering44, were grown overnight in MEI-C at 37 &#176;C without shaking. As shown in Figure 2B, the wild-type accumulated 51 &#177; 6 nmol polyP mg-1 total protein under these conditions, while the ppk1 mutant contained less than half this amount. L. reuteri contains a second polyP kinase, encoded by the ppk2 gene45, which presumably accounts for the polyP present in the ppk1 null mutant.
To demonstrate the use of this protocol with mycobacteria, Mycobacterium smegmatis strain SMR546 was grown to mid-log phase in Hartmans-de Bont (HdB) medium47, then rinsed and diluted five-fold in HdB or HdB with 2% ethanol. These cultures were then incubated overnight at 37 &#176;C with shaking (180 rpm). As shown in Figure 2C, in the absence of ethanol, M. smegmatis accumulated 141 &#177; 52 nmol polyP mg-1 total protein, while ethanol treatment resulted in a three-fold increase to 437 &#177; 102 nmol polyP mg-1 total protein. This result was expected, since ethanol has previously been reported to elevate polyP levels in M. smegmatis48.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58818/58818fig1.jpg" /> 
Figure 1: Diagram of polyphosphate extraction and measurement procedure. The essential steps of the polyP quantification protocol are illustrated. Bacterial cell pellets are lysed at 95 &#176;C in GITC (guanidine isothiocyanate). Small aliquots of the lysates are assayed for protein content using the Bradford assay. PolyP is extracted using silica membrane columns and then digested with ScPPX. The resulting free phosphate is quantified using the ascorbic acid assay. Total polyP-derived phosphate is normalized to total protein for each sample. A595 = absorbance at 595 nm; A882 = absorbance at 882 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58818/58818fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58818/58818fig2.jpg" /> 
Figure 2: Representative results with diverse bacteria. (A) E. coli MG1655 wild-type and isogenic &#8710;ppk, &#8710;ppx, and &#8710;phoB mutants were grown at 37 &#176;C with shaking (200 rpm) to A600 = 0.2 - 0.4 in rich LB medium (black circles) and then shifted to minimal medium (MOPS containing 4 g L-1 glucose and 0.1 mM K2HPO4) for 2 h (white circles; n = 3, &#177; SD). (B) L. reuteri ATCC PTA 6475 (circles) and an isogenic ppk1 null mutant (triangles) were grown overnight at 37 &#176;C in MEI-C medium without shaking (n = 3, &#177; SD). (C)&#160;M. smegmatis SMR5 was grown at 37 &#176;C with shaking (180 rpm) to A600 = 1 in HdB medium with (closed squares) or without (open squares) 2% ethanol (n = 3, &#177; SD). PolyP levels are expressed in terms of the concentration of individual polyP-derived free phosphate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58818/58818fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

8.6K 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

Optimized Procedure for Determining the Adsorption of Phosphonates onto Granular Ferric Hydroxide using a Miniaturized Phosphorus Determination Method

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric hydroxide (GFH), with little effort and high reliability. The phosphonate, e.g., nitrilotrimethylphosphonic acid (NTMP), is brought into contact with the GFH in a rotator in a solution buffered by an organic acid (e.g., acetic acid) or Good buffer (e.g., 2-(N-morpholino)ethanesulfonic acid) [MES] and N-cyclohexyl-2-hydroxyl-3-aminopropanesulfonic acid [CAPSO]) in a concentration of 10 mM for a specific time in 50 mL centrifuge tubes. Subsequently, after membrane filtration (0.45 &#181;m pore size), the total phosphorus (total P) concentration is measured using a specifically developed determination method (ISOmini). This method is a modification and simplification of the ISO 6878 method: a 4 mL sample is mixed with H2SO4 and K2S2O8 in a screw cap vial, heated to 148-150 &#176;C for 1 h and then mixed with NaOH, ascorbic acid and acidified molybdate with antimony(III) (final volume of 10 mL) to produce a blue complex. The color intensity, which is linearly proportional to the phosphorus concentration, is measured spectrophotometrically (880 nm). It is demonstrated that the buffer concentration used has no significant effect on the adsorption of phosphonate between pH 4 and 12. The buffers, therefore, do not compete with the phosphonate for adsorption sites. Furthermore, the relatively high concentration of the buffer requires a higher dosage concentration of oxidizing agent (K2S2O8) for digestion than that specified in ISO 6878, which, together with the NaOH dosage, is matched to each buffer. Despite the simplification, the ISOmini method does not lose any of its accuracy compared to the standardized method.

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric hydroxide, with little effort and high reliability. In a buffered solution, the phosphonate is brought into contact with the adsorbent using a rotator and then analyzed via a miniaturized phosphorus determination method.

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric ...

Environment

Screening for Thermotoga maritima Membrane-Bound Pyrophosphatase Inhibitors

Membrane-bound pyrophosphatases (mPPases) are dimeric enzymes that occur in bacteria, archaea, plants, and protist parasites. These proteins cleave pyrophosphate into two orthophosphate molecules, which is coupled with proton and/or sodium ion pumping across the membrane. Since no homologous proteins occur in animals and humans, mPPases are good candidates in the design of potential drug targets. Here we present a detailed protocol to screen for mPPase inhibitors utilizing the molybdenum blue reaction in a 96 well plate system. We use mPPase from the thermophilic bacterium Thermotoga maritima (TmPPase) as a model enzyme. This protocol is simple and inexpensive, producing a consistent and robust result. It takes only about one hour to complete the activity assay protocol from the start of the assay until the absorbance measurement. Since the blue color produced in this assay is stable for a long period of time, subsequent assay(s) can be performed immediately after the previous batch, and the absorbance can be measured later for all batches at once. The drawback of this protocol is that it is done manually and thus can be exhausting as well as require good skills of pipetting and time keeping. Furthermore, the arsenite-citrate solution used in this assay contains sodium arsenite, which is toxic and should be handled with necessary precautions.

Screening for Thermotoga maritima Membrane-Bound Pyrophosphatase Inhibitors

Here we present a screening method for membrane-bound pyrophosphatase (from Thermotoga maritima) inhibitors based on the molybdenum blue reaction in a 96 well plate format.

Membrane-bound pyrophosphatases (mPPases) are dimeric enzymes that occur in bacteria, archaea, plants, and protist parasites. These proteins cleave ...

Biochemistry

Nonradioactive Assay to Measure Polynucleotide Phosphorylation of Small Nucleotide Substrates

Polynucleotide kinases (PNKs) are enzymes that catalyze the phosphorylation of the 5&#39; hydroxyl end of DNA and RNA oligonucleotides. The activity of PNKs can be quantified using direct or indirect approaches. Presented here is a direct, in vitro approach to measure PNK activity that relies on a fluorescently-labeled oligonucleotide substrate and polyacrylamide gel electrophoresis. This approach provides resolution of the phosphorylated products while avoiding the use of radiolabeled substrates. The protocol details how to set up the phosphorylation reaction, prepare and run large polyacrylamide gels, and quantify the reaction products. The most technically challenging part of this assay is pouring and running the large polyacrylamide gels; thus, important details to overcome common difficulties are provided. This protocol was optimized for Grc3, a PNK that assembles into an obligate pre-ribosomal RNA processing complex with its binding partner, the Las1 nuclease. However, this protocol can be adapted to measure the activity of other PNK enzymes. Moreover, this assay can also be modified to determine the effects of different components of the reaction, such as the nucleoside triphosphate, metal ions, and oligonucleotides.

This protocol describes a nonradioactive assay to measure kinase activity of polynucleotide kinases (PNKs) on small DNA and RNA substrates.

Polynucleotide kinases (PNKs) are enzymes that catalyze the phosphorylation of the 5&#39; hydroxyl end of DNA and RNA oligonucleotides. The activity of ...

Fluorescence-Based Measurements of Phosphatidylserine/Phosphatidylinositol 4-Phosphate Exchange Between Membranes

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found to represent a novel lipid transfer protein (LTP) group in yeast and human cells. They transfer phosphatidylserine (PS) from the endoplasmic reticulum (ER) to the plasma membrane (PM) via PS/phosphatidylinositol 4-phosphate (PI(4)P) exchange cycles. This finding allows a better understanding of how PS, which is critical for signaling processes, is distributed throughout the cell and the investigation of the link between this process and phosphoinositide (PIP) metabolism. The development of new fluorescence-based protocols has been instrumental in the discovery and characterization of this new cellular mechanism in vitro at the molecular level. This paper describes the production and the use of two fluorescently labelled lipid sensors, NBD-C2Lact and NBD-PHFAPP, to measure the ability of a protein to extract PS or PI(4)P and to transfer these lipids between artificial membranes. First, the protocol describes how to produce, label, and obtain high-purity samples of these two constructs. Secondly, this paper explains how to use these sensors with a fluorescence microplate reader to determine whether a protein can extract PS or PI(4)P from liposomes, using Osh6p as a case study. Finally, this protocol shows how to accurately measure the kinetics of PS/PI(4)P exchange between liposomes of defined lipid composition and to determine lipid transfer rates by fluorescence resonance energy transfer (FRET) using a standard fluorometer.

Here, we describe protocols using fluorescent lipid sensors and liposomes to determine whether a protein extracts and transports phosphatidylserine or phosphatidylinositol 4-phosphate in vitro.

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found ...

Absolute Quantitation of Inositol Pyrophosphates by Capillary Electrophoresis Electrospray Ionization Mass Spectrometry

Inositol pyrophosphates (PP-InsPs) are an important group of intracellular signaling molecules. Derived from inositol phosphates (InsPs), these molecules feature the presence of at least one energetic pyrophosphate moiety on the myo-inositol ring. They exist ubiquitously in eukaryotes and operate as metabolic messengers surveying phosphate homeostasis, insulin sensitivity, and cellular energy charge. Owing to the absence of a chromophore in these metabolites, a very high charge density, and low abundance, their analysis requires radioactive tracer, and thus it is convoluted and expensive. Here, the study presents a detailed protocol to perform absolute and high throughput quantitation of inositol pyrophosphates from mammalian cells by capillary electrophoresis electrospray ionization mass spectrometry (CE-ESI-MS). This method enables the sensitive profiling of all biologically relevant PP-InsPs species in mammalian cells, enabling baseline separation of regioisomers. Absolute cellular concentrations of PP-InsPs, including minor isomers, and monitoring of their temporal changes in HCT116 cells under several experimental conditions are presented.

A procedure for capillary electrophoresis electrospray ionization mass spectrometry for the absolute quantitation of inositol pyrophosphates from mammalian cell extracts is described.

Inositol pyrophosphates (PP-InsPs) are an important group of intracellular signaling molecules. Derived from inositol phosphates (InsPs), these molecules ...

A Mass Spectrometry-Based Approach to Identify Phosphoprotein Phosphatases and their Interactors

Most cellular processes are regulated by dynamic protein phosphorylation. More than three-quarters of proteins are phosphorylated, and phosphoprotein phosphatases (PPPs) coordinate over 90% of all cellular serine/threonine dephosphorylation. Deregulation of protein phosphorylation has been implicated in the pathophysiology of various diseases, including cancer and neurodegeneration. Despite their widespread activity, the molecular mechanisms controlling PPPs and those controlled by PPPs are poorly characterized. Here, a proteomic approach termed phosphatase inhibitor beads and mass spectrometry (PIB-MS) is described to identify and quantify PPPs, their posttranslational modifications, and their interactors in as little as 12 h using any cell line or tissue. PIB-MS utilizes a non-selective PPP inhibitor, microcystin-LR (MCLR), immobilized on sepharose beads to capture and enrich endogenous PPPs and their associated proteins (termed the PPPome). This method does not require the exogenous expression of tagged versions of PPPs or the use of specific antibodies. PIB-MS offers an innovative way to study the evolutionarily conserved PPPs and expand our current understanding of dephosphorylation signaling.

Here, we present a protocol for the enrichment of endogenous phosphoprotein phosphatases and their interacting proteins from cells and tissues and their identification and quantification by mass spectrometry-based proteomics.

Most cellular processes are regulated by dynamic protein phosphorylation. More than three-quarters of proteins are phosphorylated, and phosphoprotein ...

High-Throughput Measurement and Classification of Organic P in Environmental Samples

Many types of organic phosphorus (P) molecules exist in environmental samples1. Traditional P measurements do not detect these organic P compounds since they do not react with colorimetric reagents2,3. Enzymatic hydrolysis (EH) is an emerging method for accurately characterizing organic P forms in environmental samples4,5. This method is only trumped in accuracy by Phosphorus-31 Nuclear Magnetic Resonance Spectroscopy (31P-NMR) -a method that is expensive and requires specialized technical training6. We have adapted an enzymatic hydrolysis method capable of measuring three classes of phosphorus (monoester P, diester P and inorganic P) to a microplate reader system7. This method provides researchers with a fast, accurate, affordable and user-friendly means to measure P species in soils, sediments, manures and, if concentrated, aquatic samples. This is the only high-throughput method for measuring the forms and enzyme-lability of organic P that can be performed in a standard laboratory. The resulting data provides insight to scientists studying system nutrient content and eutrophication potential.

This protocol describes a high-throughput method of enzymatic hydrolysis that utilizes a microplate reader to measure and classify soil phosphorus as P monoesters, P diesters and inorganic P. Up to 96 samples can be measured at one time in a standard laboratory.

Many types of organic phosphorus (P) molecules exist in environmental samples1.  Traditional P measurements do not detect these organic P compounds since ...

Biology

Colorimetric Analysis of Alkaline Phosphatase Activity in S. aureus Biofilm

Alkaline phosphatase (ALP) is a common enzyme expressed in both prokaryotic and eukaryotic cells. It catalyzes the hydrolysis of phosphate monoesters from many molecules at basic pH and plays an indispensable role in phosphate metabolism. In humans, eukaryotic ALP is one of the most frequently used enzymatic signals in diagnosing various diseases, such as cholestasis and rickets. In S. aureus, ALP is detected exclusively on the cell membrane; it is also expressed as a secretory form as well. Yet, little is known about its function in biofilm formation.
The purpose of this manuscript is to develop a quick and reliable assay to measure ALP activity in S. aureus biofilm that does not require protein isolation. Using p-nitrophenyl phosphate (pNPP) as a substrate, we measured ALP activity in S. aureus biofilm formed in 96-well tissue culture plates. Activity was based on the formation of the soluble reaction product measured by 405 nm absorbance. The high throughput nature of the 96 well tissue culture plate method provides a sensitive and reproducible method for ALP activity assays. The same experimental set up can also be extended to measure other extracellular molecular markers related to biofilm formation.

Colorimetric Analysis of Alkaline Phosphatase Activity in S. aureus Biofilm

In this manuscript, we established a high throughput method to quantify alkaline phosphatase activity in S. aureus biofilm culture from 96-well tissue culture plate

Alkaline phosphatase (ALP) is a common enzyme expressed in both prokaryotic and eukaryotic cells. It catalyzes the hydrolysis of phosphate monoesters from ...

Bioengineering

Preparation of Quality Inositol Pyrophosphates

Myo-inositol is present in nature either unmodified or in more complex phosphorylated derivates. Of the latest, the two most abundant in eukaryotic cells are inositol pentakisphosphate (IP5) and inositol hexakisphosphate (phytic acid or IP6). IP5 and IP6 are the precursors of inositol pyrophosphate molecules that contain one or more pyrophosphate bonds1. Phosphorylation of IP6 generates diphoshoinositolpentakisphosphate (IP7 or PP-IP5) and bisdiphoshoinositoltetrakisphosphate (IP8 or (PP)2-IP4). Inositol pyrophosphates have been isolated from all eukaryotic organisms so far studied. In addition, the two distinct classes of enzymes responsible for inositol pyrophosphate synthesis are highly conserved throughout evolution2-4.
The IP6 kinases (IP6Ks) posses an enormous catalytic flexibility, converting IP5 and IP6 to PP-IP4 and IP7 respectively and subsequently, by using these products as substrates, promote the generation of more complex molecules5,6. Recently, a second class of pyrophosphate generating enzymes was identified in the form of the yeast protein VIP1 (also referred as PP-IP5K), which is able to convert IP6 to IP7 and IP87,8.
Inositol pyrophosphates regulate many disparate cellular processes such as insulin secretion9, telomere length10,11, chemotaxis12, vesicular trafficking13, phosphate homeostasis14 and HIV-1 gag release15. Two mechanisms of actions have been proposed for this class of molecules. They can affect cellular function by allosterically interacting with specific proteins like AKT16. Alternatively, the pyrophosphate group can donate a phosphate to pre-phosphorylated proteins17. The enormous potential of this research field is hampered by the absence of a commercial source of inositol pyrophosphates, which is preventing many scientists from studying these molecules and this new post-translational modification. The methods currently available to isolate inositol pyrophosphates require sophisticated chromatographic apparatus18,19. These procedures use acidic conditions that might lead to inositol pyrophosphate degradation20 and thus to poor recovery. Furthermore, the cumbersome post-column desalting procedures restrict their use to specialized laboratories.
In this study we describe an undemanding method for the generation, isolation and purification of the products of the IP6-kinase and PP-IP5-kinases reactions. This method was possible by the ability of polyacrylamide gel electrophoresis (PAGE) to resolve highly phosphorylated inositol polyphosphates20. Following IP6K1 and PP-IP5K enzymatic reactions using IP6 as the substrate, PAGE was used to separate the generated inositol pyrophosphates that were subsequently eluted in water.

Inositol pyrophosphates play an important role in human pathologies such cancer, diabetes and obesity; however, the exact mechanism of action is a matter of dispute. The lack of commercially available inositol pyrophosphates renders detailed studies problematic. Here we describe a simple protocol to produce and isolate milligrams of inositol pyrophosphates.

Myo-inositol is present in nature either unmodified or in more complex phosphorylated derivates. Of the latest, the two most abundant in eukaryotic cells ...

A Purification and In Vitro Activity Assay for a (p)ppGpp Synthetase from Clostridium difficile

Kinase and pyrophosphokinase enzymes transfer the gamma phosphate or the beta-gamma pyrophosphate moiety from nucleotide triphosphate precursors to substrates to create phosphorylated products. The use of &#947;-32-P labeled NTP precursors allows simultaneous monitoring of substrate utilization and product formation by radiography. Thin layer chromatography (TLC) on cellulose plates allows rapid separation and sensitive quantification of substrate and product. We present a method for utilizing the thin-layer chromatography to assay the pyrophosphokinase activity of a purified (p)ppGpp synthetase. This method has previously been used to characterize the activity of cyclic nucleotide and dinucleotide synthetases and is broadly suitable for characterizing the activity of any enzyme that hydrolyzes a nucleotide triphosphate bond or transfers a terminal phosphate from a phosphate donor to another molecule.

A Purification and In Vitro Activity Assay for a pppGpp Synthetase from Clostridium difficile

Here, we describe a method for purifying histidine-tagged pyrophosphokinase enzymes and utilizing thin layer chromatography of radiolabelled substrates and products to assay for the enzymatic activity in vitro. The enzyme activity assay is broadly applicable to any kinase, nucleotide cyclase, or phosphor-transfer reaction whose mechanism includes nucleotide triphosphate hydrolysis.

Kinase and pyrophosphokinase enzymes transfer the gamma phosphate or the beta-gamma pyrophosphate moiety from nucleotide triphosphate precursors to ...

Assaying for Inorganic Polyphosphate in Bacteria

Summary

Explore More Videos

Optimized Procedure for Determining the Adsorption of Phosphonates onto Granular Ferric Hydroxide using a Miniaturized Phosphorus Determination Method

Screening for Thermotoga maritima Membrane-Bound Pyrophosphatase Inhibitors

Nonradioactive Assay to Measure Polynucleotide Phosphorylation of Small Nucleotide Substrates

Fluorescence-Based Measurements of Phosphatidylserine/Phosphatidylinositol 4-Phosphate Exchange Between Membranes

Absolute Quantitation of Inositol Pyrophosphates by Capillary Electrophoresis Electrospray Ionization Mass Spectrometry

A Mass Spectrometry-Based Approach to Identify Phosphoprotein Phosphatases and their Interactors

High-Throughput Measurement and Classification of Organic P in Environmental Samples

Colorimetric Analysis of Alkaline Phosphatase Activity in S. aureus Biofilm

Preparation of Quality Inositol Pyrophosphates

A Purification and In Vitro Activity Assay for a (p)ppGpp Synthetase from Clostridium difficile