This work was supported by Grants AI064184 and AI76322 from the National Institutes of Health (NIH) and by Grant 61789-MA-MUR from the Army Research Office to M.S.T. Research was also supported by Welch Foundation Grant F1155 and NIH grant R01GM103655 to J.S. B.

All solutions should be prepared with ultrapure water and HPLC grade methanol and chloroform. Prepared solutions that contain organic solvents such as methanol, chloroform, or pyridine and concentrated acids or bases should be prepared and used under a chemical fume hood. All solutions can be stored at RT. Solvents should be measured in a graduated glass cylinder and stored in glass solvent bottles with PTFE lined caps. For long-term storage chloroform-containing solvents should be stored in tinted amber glass bottles to avoid the production of phosgene, a highly reactive acid chloride. PTFE centrifuge tubes and rotary evaporator flasks should be rinsed with methanol and chloroform before use. Follow necessary federal, state and/or institutional waste disposal regulations when disposing of solvents and/or radioactive waste.
1. Large Scale Lipid A Extraction (50 ml to 1.5 L)
<ol>
	<li>Prepare a single-phase Bligh-Dyer mixture: chloroform-methanol-1X phosphate buffered saline (PBS), pH 7.4; 1:2:0.8 v/v). Combine 200 ml of chloroform, 400 ml of methanol and 160 ml of PBS in 1 L solvent bottle. Cap bottle and mix by inversion (&gt;30x). Loosen the cap periodically during mixing to vent and store sealed.</li>
	<li>Prepare a pre-equilibrated two-phase Bligh-Dyer mixture- chloroform:methanol:water (2:2:1.8 v/v). Combine 400 ml of chloroform, 400 ml of methanol and 180 ml of water in 1 L solvent bottle. Cap bottle and mix by inversion, making sure to loosen cap periodically during mixing to vent. Let equilibrate O/N and store sealed.</li>
	<li>Prepare the mild acid hydrolysis buffer (50 mM sodium acetate, pH 4.5; 1% sodium dodecyl sulfate (SDS)). For 500 ml of mild acid hydrolysis buffer, weigh 2.05 g of sodium acetate and transfer to a 500 ml beaker. Add water to a volume of ~350 ml and stir, then add 50 ml of 10% SDS. Mix and adjust pH to 4.5, transfer to a graduated cylinder, and add water to 500 ml.</li>
	<li>Prepare chloroform:methanol (4:1, v/v): Measure 100 ml of chloroform and transfer to solvent bottle. Measure 25 ml of methanol and mix with chloroform. Store in amber glass bottle.</li>
	<li>Inoculate 5 ml of media (Luria broth or other) from a single bacterial colony. Grow O/N at 37 &deg;C, or at required &deg;C for growth. A diagram of the extraction procedure is shown in Figure 2.</li>
	<li>The next day, measure the OD600 and use the 5 ml O/N culture to inoculate 200 ml of culture to a starting OD600 of 0.05. Grow cells until an OD600 of 0.8-1.0 is reached.</li>
	<li>Harvest cells via centrifugation at 10,000 x g for 10 min. Longer spins may be required for strains that pellet poorly. Pour off media supernatant.</li>
	<li>Wash cell pellet with 50 ml of 1x phosphate buffered saline (PBS). Repeat centrifugation to pellet cells. Pour off supernatant and store cell pellet at -20 &deg;C, or proceed to step 1.9.</li>
	<li>Resuspend cells in 40 ml of 1x PBS and divide between two 250 ml PTFE centrifuge tubes (yielding 20 ml of cell suspension/tube). Add 25 ml of chloroform and 50 ml of methanol to each tube, for a single phase Bligh-Dyer (chloroform: methanol:water; 1:2:0.8 v/v) (Figure 2). Mix by inversion and incubate at RT for &gt;20 min to ensure complete cell lysis.</li>
	<li>Centrifuge the mixture at 2,000 x g for 20 min. LPS will pellet along with proteins and nucleic acids (Figure 2); however, phospholipids, isoprenyl lipids, and small, hydrophobic peptides will remain in the supernatant. Discard the supernatant.</li>
	<li>Wash the LPS pellet with ~100 ml single phase Bligh-Dyer mixture. Centrifuge at 2,000 x g for 20 min. Discard supernatant. When isolating lipid A from E. coli or Salmonella, only one wash is required; however, additional wash steps may be needed to reduce phospholipid contamination when isolating lipid A from some organisms.</li>
	<li>Add 27 ml of mild acid hydrolysis buffer (50 mM sodium acetate, pH 4.5; 1% SDS; see step 1.3) to LPS pellet, and mix by pipetting up and down until only small particles remain. Sonicate to homogeneously resuspend LPS pellet in solution with probe tip sonicator at a constant duty cycle for 20 sec at 50% output. Repeat sonication of sample 2x (20 sec/burst, ~5 sec between bursts).</li>
	<li>Boil samples in a water bath for 30 min. Caution: make sure the caps are tight, but not fully sealed. Some organisms, like Vibrio cholerae (V. cholerae) require longer incubation times (1 hr) to increase overall yield of lipid A. Remove bottles from water bath and allow sample to cool to RT before proceeding.</li>
	<li>To extract lipids after hydrolysis, convert the SDS solution into a two-phase Bligh-Dyer (Figure 2) mixture by adding 30 ml of chloroform and 30 ml of methanol, for a chloroform:methanol:water (2:2:1.8, v/v) mixture. Mix by inversion and centrifuge the sample for 10 min at 2,000 x g. Extract the lower phase into a clean PTFE centrifuge tube using a glass pipette.</li>
	<li>Perform a second extraction by adding 30 ml of lower phase from a pre-equilibrated two-phase Bligh-Dyer mixture (step 1.2), to the upper phase from step 1.14. Mix, then centrifuge at 2,000 x g for 10 min. Extract the lower phase, and pool with the lower phase extracted in step 1.14.</li>
	<li>Wash the pooled lower phases (60 ml total) by adding 114 ml of pre-equilibrated two phase Bligh-Dyer upper phase (step 1.2) to create a two-phase Bligh-Dyer mixture (chloroform:methanol:water; 2:2:1.8, v/v). Mix. Centrifuge at 2,000 x g for 10 min.</li>
	<li>Remove the lower phase to a clean glass rotary evaporator flask and dry sample using rotary evaporation (Figure 2).</li>
	<li>Add 5 ml of chloroform:methanol (4:1, v/v) to the rotary flask, and bath-sonicate (&gt;30 sec) to aid in suspension of lipid from sides of flask. Use a glass transfer pipette to transfer lipid to a clean glass tube (13 x 100 mm or larger) capped with PTFE lined phenolic screw caps. Dry sample under a stream of nitrogen using a nitrogen dryer.</li>
	<li>Resuspend dried lipid in 1 ml of chloroform:methanol (4:1, v/v). Transfer to small glass sample vial (12 x 32 mm) with a tapered base for more quantitative resuspension using small volumes (see Table of Equipment/Reagents). Dry using a nitrogen dryer.</li>
	<li>Dried sample can be stored at -20 &deg;C until subsequent TLC (Protocol 2) or MS analysis (Protocol 3, 4, or 5). (Note: Amount of material isolated and used in subsequent protocols is suggested based on what's sufficient for most organisms. The same lipid sample can be used in Protocols 2-5, taking note of % material removed. See Discussion for more details.).</li>
</ol>
2. Visualization of Lipid A Species via Thin Layer Chromatography
<ol>
	<li>Prepare the TLC mobile phase solvent system (chloroform:pyridine:88% formic acid:water; 50:50:16:5 v/v). Combine 200 ml of chloroform, 200 ml of pyridine, 64 ml of 88% formic acid, and 20 of ml water in a 1 L solvent bottle. Cap bottle, mix by inversion multiple times, and vent.</li>
	<li>Use a TLC tank that will accommodate 20 x 20 cm plates. Line TLC tank with ~40 cm chromatography paper.</li>
	<li>To TLC tank add 200 ml of chloroform:pyridine:88% formic acid:water (50:50:16:5, v/v) mixture. Allow tank to pre-equilibrate for &gt;3 hr, often O/N is preferred and more convenient.</li>
	<li>Remove dried lipid samples from freezer (step 1.20) and let warm to RT.</li>
	<li>With a razor remove the silica from the top edge of a silica gel 60 TLC plate. Using a dull pencil, draw a line parallel to the bottom of the plate, 2 cm from the bottom. This line is the origin for spotting samples. Mark increments along this reference line to spot samples 1 cm apart.</li>
	<li>Dissolve dried lipid from step 2.4 in 200 &mu;l of chloroform:methanol (4:1, v/v), vortex and sonicate (3x, ~15 sec each), yields ~100-500 ng/&mu;l lipid A if extracted from E. coli.</li>
	<li>Using a microcapillary glass pipet, spot one-tenth of the volume (20 &mu;l) onto the TLC plate as marked in step 2.5. Allow samples to air-dry on TLC plate for ~15 min. (Note: Samples in small glass vials can be dried using a nitrogen dryer and stored at -20 &deg;C for subsequent use in protocols 3-5. Take note of % material removed).</li>
	<li>Place TLC plate containing spotted samples into the pre-equilibrated tank. Once solvent front reaches the top of the TLC plate (~2.5-3 hr), remove the plate and air dry (&gt;60 min). A cold-air gun can be used to ensure complete dryness.</li>
	<li>While plate is drying, turn on hot plate at 250 &deg;C for charring.</li>
	<li>In a ventilated chemical fume hood, prepare 10% sulfuric acid-ethanol mixture for charring lipids resolved on the silica TLC plate (concentrated sulfuric acid:100% ethanol; 1:9 v/v). Measure 10 ml of sulfuric acid and add slowly to 90 ml of 100% ethanol. Carefully mix and transfer to a glass chromatographic reagent atomizer.</li>
	<li>In the fume hood, use a glass chromatographic reagent atomizer to spray the dried TLC plate with 10% sulfuric acid-ethanol mixture. Spray mixture evenly across the plate.</li>
	<li>Place the TLC plate on the 250 &deg;C hot plate until charred lipid samples appear as black/brown spots (&lt;1 min). Do not overexpose the plate, as this will cause the entire plate to turn brown and make visualization of lipid A species difficult.</li>
</ol>
3. Structural Characterization of Lipid A via MALDI-TOF Mass Spectrometry
<ol>
	<li>From step 1.20, (or step 2.7) remove lipid A sample from freezer and let warm to RT. Resuspend dried lipid A sample in ~20 &mu;l chloroform:methanol (4:1, v/v) and vortex to obtain a ~1-5 &mu;g/&mu;l lipid A solution if extracted from E. coli. (Note: 10% less concentrated if material used from step 2.7).</li>
	<li>Prepare ATT matrix components. Saturated 6-aza-2-thiothymine in 50% acetonitrile: add 500 &mu;l of water and 500 &mu;l of acetonitrile to a 1.5 ml microcentrifuge tube, and then add &gt;10 mg 6-aza-2-thiothymine so that the 50% acetonitrile is super-saturated. Saturated tribasic ammonium citrate solution: Add &gt;1 mg to 500 &mu;l water, precipitate should be readily apparent. Vortex and centrifuge solutions before use; only saturated supernatant is used.</li>
	<li>Prepare ATT matrix mixture: saturated 6-aza-2-thiothymine in 50% acetonitrile, saturated tribasic ammonium citrate (20:1, v/v). Mix matrix components together by adding 20 &mu;l of ATT solution to 1 &mu;l saturated tribasic ammonium citrate solution in 500 &mu;l microcentrifuge tube, vortex to mix, and centrifuge before applying to MALDI plate.</li>
	<li>Prepare MALDI plate by adding 0.5 &mu;l calibrant mixture to the MALDI plate on a spot near where the samples will be deposited, to provide the most accurate mass/charge ratio determination.</li>
	<li>Deposit 0.5 &mu;l of ATT matrix onto MALDI plate on each spot where lipid A sample will be deposited.</li>
	<li>Deposit 0.5 &mu;l of sample (2.5% of total sample) onto the spot of ATT matrix, to mix on plate, and acquire spectra by scanning sample for optimal ion signals (Figure 3). Note: Amounts for the most optimal MS signal can vary with organism and need to be determined empirically. To add more material one can spot 0.5 &mu;l multiple times allowing spotted mixture to dry in between addition of more sample. Sample can also be concentrated (dry and resuspend in a smaller volume) or diluted as necessary. More starting material (volume of starting cell culture) can also be used (See Discussion).</li>
</ol>
4. Electrospray Ionization Mass Spectrometry and Collision Induced Dissociation of Lipid A
<ol>
	<li>Prepare a chloroform:methanol solvent mixture (1:1, v/v) by mixing a 200 &mu;l stock of HPLC grade methanol with 200 ml HPLC grade chloroform in a glass solvent bottle.</li>
	<li>Transfer 200 &mu;l of the chloroform:methanol solvent mixture to the vial with lipid A (e.g. from step 1.20, 2.7, or dried after Protocol 3) and sonicate the lipid a solution for 5 min or until all material is dissolved.</li>
	<li>Set up the mass spectrometer for negative mode electrospray ionization.</li>
	<li>Using a 250 &mu;l syringe and a syringe pump, directly infuse the diluted lipid A sample at a flow rate of 2.0-3.5 &mu;l/min.</li>
	<li>Optimize and enhance the lipid A ion signal by tuning the ion optics. A full mass spectrum can now be collected of the lipid A species (Figure 4).</li>
	<li>Isolate and activate the target lipid A by selecting CID as the MS/MS method.</li>
	<li>Increase the CID voltage (or normalized collision energy, NCE) until the precursor lipid A species is about 10% relative abundance compared to the highest product ion.</li>
	<li>Acquire and average spectra until sufficient signal-to-noise is achieved for the product ions. The number of scans needed is dependent on the signal intensity of the original precursor and can range from 3-300 scans (Figure 5A).</li>
</ol>
5. MS/MS on Lipid A by Ultraviolet Photodissociation
<ol>
	<li>Prepare sample and mass spectrometer as in steps 4.1-4.5.</li>
	<li>Turn on the laser interfaced to the mass spectrometer. The mass spectrometer was outfitted with a 193 nm excimer laser and modified to allow UV activation in the HCD cell of the instrument. Photodissociation was implemented in a manner similar to that described previously35. We use a modified vacuum manifold with a CaF2 optical window to transmit photons into the higher-energy C-trap dissociation (HCD) cell. The laser is triggered during MS/MS by a transistor-transistor logic (TTL) signal from mass spectrometer to a pulse/delay generator.</li>
	<li>Set up the instrument software so that the laser will trigger the excimer laser when the ions enter the HCD cell. This requires a modest modification of the software.</li>
	<li>Turn on the pulse generator such that the laser is pulsed every 2 msec (500 Hz).</li>
	<li>Isolate the lipid A precursor ion by selecting HCD as MS/MS method and adjust the collisional energy to 1% NCE. This allows the isolation of precursor ions for ultraviolet dissociation within the HCD cell. Although the HCD cell is used, the software is modified to perform UVPD during this interval.</li>
	<li>Activate the isolated lipid A ion by increasing the laser energy and adjusting the number of laser pulses. A typical UVPD experiment will be performed using ten 6 ml pulses.</li>
	<li>As in step 4.8, acquire and average spectra until sufficient signal-to-noise is achieved for the UVPD product ions. The number of scans needed is dependent on the signal intensity of the original precursor and can range from 3-300 scans (Figure 5B).</li>
</ol>
6. 32P-Labeling of Lipid A and Subsequent Isolation
<ol>
	<li>Inoculate 5 ml of media (Luria broth or other media) from a single colony. Grow O/N at 37 &deg;C or at required temperature.</li>
	<li>The next day, measure the OD600 and use the O/N culture to inoculate 7 ml of culture to a starting OD600 of ~0.05, grown in a standard 20 x 150 mm disposable glass culture tube. Add 2.5 &mu;Ci/ml of inorganic 32P. Grow cells until an OD600 of 0.8-1.0 is reached. Please follow federal, state, and/or institutional guidelines for the safe and proper handling of radioactive materials.</li>
	<li>Harvest cells in 16 x 125 mm glass centrifuge tubes with PTFE lined cap using a fixed angle clinical centrifuge at 1,500 x g for 10 min. Remove supernatant into appropriate radioactive waste container. All radioactive waste (e.g. liquid, glass, biohazard) generated in subsequent steps should be disposed of in accordance with federal, state, and/or institutional guidelines.</li>
	<li>Wash cell pellet with 5 ml of 1x phosphate buffered saline (PBS). Centrifuge for 10 min at 1,500 x g. Discard supernatant.</li>
	<li>Resuspend cells in 5 ml of single phase Bligh-Dyer mixture (Figure 2) consisting of chloroform:methanol:water (1:2:0.8, v/v). Vortex to mix, and incubate at RT for &gt;20 min to ensure complete cell lysis.</li>
	<li>Centrifuge in clinical centrifuge at 1,500 x g for 20 min. Gently pour off supernatant, which contains phospholipids and isoprenyl lipids.</li>
	<li>Resuspend the LPS pellet in 1.8 ml of 50 mM sodium acetate, pH 4.5; 1% SDS buffer by vortexing. Sonicate sample using a bath sonicator until pellet is evenly dispersed (~30 sec).</li>
	<li>Incubate sample for 30 min in boiling water bath. Make sure caps are tight but not sealed.</li>
	<li>Remove from water bath and allow sample to cool at RT for 5-10 min.</li>
	<li>Convert the solution into a two-phase Bligh-Dyer mixture by adding 2 ml of chloroform and 2 ml of methanol, yielding a chloroform:methanol:aqueous (2:2:1.8 v/v) mixture. Vortex to mix, and centrifuge for 10 min in clinical centrifuge at 1,500 x g to separate phases.</li>
	<li>Extract the lower phase into a clean glass centrifuge tube using a glass pipette (first extraction).</li>
	<li>Perform a second extraction on the sample by adding 2 ml of pre-equilibrated two-phase Bligh-Dyer lower phase (prepared as in step 1.2) to the remaining upper phase from step 11. Vortex and centrifuge 10 min. Remove lower phase and combine with the lower phase from the first extraction.</li>
	<li>To the pooled lower phase (~4 ml total volume), add 7.6 ml of pre-equilibrated upper phase (step 1.2), yielding a two-phase Bligh-Dyer solution (chloroform:methanol:water; 2:2:1.8, v/v). Vortex and centrifuge for 10 min.</li>
	<li>Remove the lower phase to a clean glass tube and dry using a nitrogen dryer. Dried samples can be stored at -20 &deg;C until further use.</li>
</ol>
7. Visualization of 32P-labeled Lipid A Species via Thin Layer Chromatography
<ol>
	<li>Prepare 32P-labeled lipid A sample, TLC tank, and TLC plates as described in steps 2.1-2.8.</li>
	<li>Dissolve 32P-labeled sample in 500 &mu;l of 4:1 chloroform-methanol (v/v). Vortex and bath sonicate to completely dissolve lipid material. Add 5 &mu;l of sample to scintillation vial containing 5 ml of scintillation cocktail. Count in scintillation counter and calculate total counts/min of sample.</li>
	<li>When visualizing radiolabeled lipid species, 10 x 20 cm or 20 x 20 cm TLC plates can be used. For 10x20 cm plates, samples should be spotted along the origin marked in step 2.5 so that the plate's longest dimension is vertical (i.e. samples spotted at origin marked 2 cm above the 10 cm edge).</li>
	<li>Using a microcapillary pipette, spot 10,000-20,000 cpm/sample on plate and allow spots to dry (&gt;15 min). Samples might need to be concentrated by drying under nitrogen and resuspended in an appropriate volume, to achieve 10,000-20,000 counts/min/spot (2 &mu;l or 2 &mu;l at a time for &lt;10 &mu;l total).</li>
	<li>Place TLC plate containing radiolabeled samples into the pre-equilibrated tank. Once solvent front reaches the top of the TLC plate (~3 hr), remove the plate and air dry (&gt;60 min). Caution: TLC plates run in the presence of pyridine must be dried thoroughly in a chemical fume hood, as trace amounts of pyridine can damage phosphorimaging screens. A cold-air gun can be used to ensure complete dryness.</li>
	<li>Wrap the plate in plastic wrap and expose to phosphorImager screen in an autoradiography cassette O/N. The next morning, scan the screen to obtain the image (Figure 6). Using image densitometry analysis software, this method allows for accurate, relative quantification of lipid A species.</li>
</ol>

No conflicts of interest declared.

In this protocol we have detailed the isolation of lipid A species from whole cells of bacteria, and described TLC or MS based analytical methods to chemically characterize this isolated material. Tandem mass spectrometry is a powerful strategy for de novo structural characterization of biological compounds, and is invaluable for the chemical characterization of the panoply of lipid A molecules observed in nature. CID and UVPD are two complementary activation methods that create different types of product ions t...

Lipopolysaccharide (LPS) is the major outer surface molecule of nearly all gram-negative organisms and consists of three molecular domains: a distal O-antigen polysaccharide, a core oligosaccharide, and the membrane-associated lipid A domain deposited on the outer leaflet of the outer membrane bilayer1,2. The lipid A domain consists of 3-deoxy-D-manno-oct-2-ulosonic (Kdo) residues and a lipid A molecular species, where lipid A can be defined as the chloroform soluble portion of LPS upon mild-acid hydrolysis1,2. The standard lipid A molecule can be chemically defined as a diglucosamine backbone that is hexa-acylated and bis-phosphorylated; consistent with the major lipid A species observed in the model organism Escherichia coli (E. coli)1,2. Nine constitutively expressed genes, conserved throughout gram-negative bacteria, are responsible for the production of the lipid A domain (Figure 1)1,2. Most bacteria have an additional set of genes, which vary in degree of phylogenetic conservation, that participate in further chemical modification of lipid A3. Dephosphorylation, removal of acyl chains, and the addition of chemical moieties such as amino sugars (e.g. aminoarabinose) and/or phosphoethanolamine are the most commonly observed activities (Figure 1). Many of the enzymes responsible for lipid A modification are directly activated by environmental signals, such as divalent cations, or their expression is regulated by two component response-regulator systems3.
Recognition of lipid A species by the host innate immune system is mediated by the Toll-like receptor 4/myeloid differentiation factor 2 (TLR4/MD2) co-receptor4. Hydrophobic forces between MD2 and the lipid A acyl chains, as well as between TLR4 and the 1 and 4 'phosphate groups of lipid A promote the strong association of lipid A with TLR4/MD24,5. Modifications that alter acylation state or the negative charge of lipid A impact TLR4/MD2 based lipid A recognition and downstream stimulation of the innate immune response activators NF-&kappa;B and mediators of inflammation such as TNF&alpha; and IL1-&beta;6,7. Modifications that mask the negative charge of lipid A also prevent bactericidal cationic antimicrobial peptides from binding to gram-negative cell surfaces3,8. Many lipid A modifications are hypothesized to increase bacterial fitness under specific environmental conditions, such as inside the human host or in an ecological niche. For this reason many modification enzymes are attractive targets in the rational development of antimicrobial compounds. The chemical diversity of lipid A structures, with respect to organism and/or environment, and the biological implications of these diverse structures make the structural characterization of lipid A an important endeavor in the study of gram-negative bacteria.
Isolation of lipid A molecules from whole bacteria involves the extraction of LPS from the bacterial cell surface, a hydrolytic step to liberate lipid A, followed by a final purification procedure9-11. The most frequently cited LPS extraction procedure is the hot-phenol water extraction procedure, first introduced by Westphal and Jann10. After extraction whole LPS is subjected to mild-acid hydrolysis, which chemically separates Kdo from the 6'-hydroxyl of the distal glucosamine sugar of lipid A (Figure 1). Numerous pitfalls exist for the hot-phenol water procedure including the use of a high hazard reagent, the need to degrade co-extracted nucleic acids and proteins, and several days are required to complete the protocol10.
Our lab has further developed the extraction and isolation of lipid A as first developed by Caroff and Raetz12,13. Compared to hot-phenol water procedures, the method presented here is more rapid and efficient and accommodates a wide range of culture volumes from 5 ml to multiple liters. Moreover, unlike hot-phenol water extractions, our method does not select for rough- or smooth-types of LPS, providing optimal recovery of lipid A species. In our protocol, chemical lysis of whole bacterial cells is performed using a mixture of chloroform, methanol and water, where LPS can be pelleted by centrifugation. A combination of mild-acid hydrolysis and solvent extractions (Bligh-Dyer) are used to liberate lipid A from covalently attached polysaccharide. The method of Bligh and Dyer was first applied to the extraction of lipid species from a variety of animal and plant tissues14, modified here to separate hydrolyzed polysaccharide from lipid A. In this final separation step, chloroform soluble lipids selectively partition into the lower organic phase. To further purify lipid A, reverse-phase or anionic exchange column chromatography can be used12.
After isolation of lipid A species from whole cells, a number of analytical methods can be used to characterize the chemical structure of the isolated material such as NMR, TLC, and MS-based analysis. NMR allows for non-destructive structural elucidation, and provides structural detail related to glycosidic linkages, unequivocal assignment of acyl chain positions, and assignment of attachment sites for lipid A modifications like aminoarabinose or phosphoethanolamine15-17. NMR analysis of lipid A is not discussed within our protocol, but has been described adequately elsewhere15,16. For rapid analysis TLC based methods are frequently used, but fail to provide direct information regarding fine chemical structure. MS based protocols are the most frequently employed method to characterize lipid A structures18,19. Matrix associated laser desorption ionization (MALDI)-MS is often used to initially survey intact lipid A species. Singly charged ions are generated from analyte prepared according to our extraction procedures. As more fine structural analysis is required, MS/MS based methods prove more informative than MALDI-MS. Coupled to electrospray ionization (ESI) singly or multiply charged lipid A precursor ions are further fragmented by collision induced dissociation (CID) or ultraviolet photodissociation (UVPD), to generate structurally informative product ions18,20,21. Neutral loss products from lipid A precursor ions are also frequently generated during ESI-MS providing an additional layer of structural information.
Tandem mass spectrometry (MS/MS) has proven to be an indispensable and versatile method for the elucidation of lipid A structures. During MS/MS, ions are activated to yield a diagnostic fragmentation pattern that can be used to elucidate the structure of the precursor ion. The most widely available MS/MS method is CID. This method produces fragment ions via collisions of the selected precursor ion with an inert target gas, resulting in energy deposition that leads to dissociation. CID has proven a critical tool in the assignment of lipid A structure for a wide range of bacterial species22-33.
Although CID is the most universally implemented MS/MS method, it generates a limited array of product ions. 193 nm UVPD is an alternative and complementary MS/MS method. This method uses a laser to irradiate ions, and the absorption of photons results in energization of the ions and subsequent dissociation. This higher energy MS/MS technique produces a more diverse array of product ions than CID and thus provides more informative fragmentation patterns. In particular, UVPD affords information about subtle changes in lipid A species based on cleavages at glycosidic, amine, acyl and C-C linked bonds18,21,34.

<table border="1">
		<tbody>
		 <tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		 </tr>
		 <tr>
			<td>Chloroform</td>
			<td>Thermo Fisher Scientific</td>
			<td>C607</td>
			<td>HPLC Grade</td>
		 </tr>
		 <tr>
			<td>Methanol</td>
			<td>Thermo Fisher Scientific</td>
			<td>A452</td>
			<td>HPLC Grade</td>
		 </tr>
		 <tr>
			<td>Teflon FEP Centrifuge Bottles</td>
			<td>Thermo Fisher Scientific</td>
			<td>05-562-21</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Silica Gel 60 TLC Plates</td>
			<td>EMD Biosciences</td>
			<td>5626-6</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Grade No. 3MM Chromatography Paper</td>
			<td>Whatman</td>
			<td>3030700</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Orbitrap Elite</td>
			<td>Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Mass Spectrometer</td>
 <td></td>
 <td></td>
 <td></td>
		 </tr>
		 <tr>
			<td>ExciStar XS Excimer Lasrer</td>
			<td>Coherent Inc.</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>PicoTip Nanospray ESI emitters</td>
			<td>New Obectives</td>
			<td></td>
			<td>&ge; 30 &mu;m to reduce clogging</td>
		 </tr>
		 <tr>
			<td>Model 505 Pulse/Delay Generator</td>
			<td>Berkeley Nucleonics Corporation</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Hot Plate Thermoylne 2200</td>
			<td>Barnstead/Thermolyne</td>
			<td>HPA2235MQ</td>
			<td></td>
		 </tr>
		 <tr>
			<td>16x125 mm GPI 15-415 Threaded Disposable Borosilicate Culture Tubes</td>
			<td>Corning Pyrex</td>
			<td>99449-16X</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Reusable Threaded PTFE screw caps GPI 45-415</td>
			<td>Corning </td>
			<td>9999-152</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Personal Molecular Imager System (phosphorimager)</td>
			<td>BioRad</td>
			<td>170-9400</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Autoradiography Cassette</td>
			<td>Thermo Fisher Scientific</td>
			<td>FBCS810</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Phosphorscreen SO230</td>
			<td>Kodak</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Peptide Mass Standards Kit</td>
			<td>Sequazyme</td>
			<td>P2-3143-00</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Sonifier S250-A</td>
			<td>Branson</td>
			<td>101063196</td>
			<td></td>
		 </tr>
		 <tr>
			<td>1.5 ml 12x32 mm Tapered Base Screw Thread Vial</td>
			<td>Thermo Fisher Scientific</td>
			<td>C4000-V1</td>
			<td></td>
		 </tr>
		</tbody>
 </table>

isolation and chemical characterization of lipid a from gram-negative bacteria

Lipopolysaccharide (LPS) is the major cell surface molecule of gram-negative bacteria, deposited on the outer leaflet of the outer membrane bilayer. LPS can be subdivided into three domains: the distal O-polysaccharide, a core oligosaccharide, and the lipid A domain consisting of a lipid A molecular species and 3-deoxy-D-manno-oct-2-ulosonic acid residues (Kdo). The lipid A domain is the only component essential for bacterial cell survival. Following its synthesis, lipid A is chemically modified in response to environmental stresses such as pH or temperature, to promote resistance to antibiotic compounds, and to evade recognition by mediators of the host innate immune response. The following protocol details the small- and large-scale isolation of lipid A from gram-negative bacteria. Isolated material is then chemically characterized by thin layer chromatography (TLC) or mass-spectrometry (MS). In addition to matrix-assisted laser desorption/ionization-time of flight (MALDI-TOF) MS, we also describe tandem MS protocols for analyzing lipid A molecular species using electrospray ionization (ESI) coupled to collision induced dissociation (CID) and newly employed ultraviolet photodissociation (UVPD) methods. Our MS protocols allow for unequivocal determination of chemical structure, paramount to characterization of lipid A molecules that contain unique or novel chemical modifications. We also describe the radioisotopic labeling, and subsequent isolation, of lipid A from bacterial cells for analysis by TLC. Relative to MS-based protocols, TLC provides a more economical and rapid characterization method, but cannot be used to unambiguously assign lipid A chemical structures without the use of standards of known chemical structure. Over the last two decades isolation and characterization of lipid A has led to numerous exciting discoveries that have improved our understanding of the physiology of gram-negative bacteria, mechanisms of antibiotic resistance, the human innate immune response, and have provided many new targets in the development of antibacterial compounds.

Canonical lipid A of E. coli and Salmonella enterica serovar Typhimurium is a hexa-acylated disaccharide of glucosamine with phosphate groups at the 1- and 4 '-positions. During growth in rich media (e.g. Luria Broth) a portion of the lipid A contains a pyrophosphate group at the 1-position yielding a tris-phosphorylated species36 (Figure 1). Kdo (3-deoxy-D-manno-octulosonic acid), is attached at the 6'-hydroxyl and serves as a bridge to link lipid ...

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Isolation and Chemical Characterization of Lipid A from Gram-negative Bacteria

Isolation and characterization of the lipid A domain of lipopolysaccharide (LPS) from gram-negative bacteria provides insight into cell surface based mechanisms of antibiotic resistance, bacterial survival and fitness, and how chemically diverse lipid A molecular species differentially modulate host innate immune responses.

Lipopolysaccharide (LPS) is the major cell surface molecule of gram-negative bacteria, deposited on the outer leaflet of the outer membrane bilayer. LPS ...

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31.5K Views.  The University of Texas at Austin. Isolation and characterization of the lipid A domain of lipopolysaccharide (LPS) from gram-negative bacteria provides insight into cell surface based mechanisms of antibiotic resistance, bacterial survival and fitness, and how chemically diverse lipid A molecular species differentially modulate host innate immune responses.

Video: Isolation and Chemical Characterization of Lipid A from Gram-negative Bacteria

Free Radicals in Chemical Biology: from Chemical Behavior to Biomarker Development

The involvement of free radicals in life sciences has constantly increased with time and has been connected to several physiological and pathological processes. This subject embraces diverse scientific areas, spanning from physical, biological and bioorganic chemistry to biology and medicine, with applications to the amelioration of quality of life, health and aging. Multidisciplinary skills are required for the full investigation of the many facets of radical processes in the biological environment and chemical knowledge plays a crucial role in unveiling basic processes and mechanisms. We developed a chemical biology approach able to connect free radical chemical reactivity with biological processes, providing information on the mechanistic pathways and products. The core of this approach is the design of biomimetic models to study biomolecule behavior (lipids, nucleic acids and proteins) in aqueous systems, obtaining insights of the reaction pathways as well as building up molecular libraries of the free radical reaction products. This context can be successfully used for biomarker discovery and examples are provided with two classes of compounds: mono-trans isomers of cholesteryl esters, which are synthesized and used as references for detection in human plasma, and purine 5',8-cyclo-2'-deoxyribonucleosides, prepared and used as reference in the protocol for detection of such lesions in DNA samples, after ionizing radiations or obtained from different health conditions.

Radical-based biomimetic chemistry has been applied to building-up libraries necessary for biomarker development.

The involvement of free radicals in life sciences has constantly increased with time and has been connected to several physiological and pathological ...

Folding and Characterization of a Bio-responsive Robot from DNA Origami

The DNA nanorobot is a hollow hexagonal nanometric device, designed to open in response to specific stimuli and present cargo sequestered inside. Both stimuli and cargo can be tailored according to specific needs. Here we describe the DNA nanorobot fabrication protocol, with the use of the DNA origami technique. The procedure initiates by mixing short single-strand DNA staples into a stock mixture which is then added to a long, circular, single-strand DNA scaffold in presence of a folding buffer. A standard thermo cycler is programmed to gradually lower the mixing reaction temperature to facilitate the staples-to-scaffold annealing, which is the guiding force behind the folding of the nanorobot. Once the 60 hr folding reaction is complete, excess staples are discarded using a centrifugal filter, followed by visualization via agarose-gel electrophoresis (AGE). Finally, successful fabrication of the nanorobot is verified by transmission electron microscopy (TEM), with the use of uranyl-formate as negative stain.

DNA origami is a powerful method for fabricating precise nanoscale objects by programming the self-assembly of DNA molecules. Here we describe a protocol for the folding of a bio-responsive robot from DNA origami, its purification and negative staining for transmission electron microscopic imaging (TEM).

The DNA nanorobot is a hollow hexagonal nanometric device, designed to open in response to specific stimuli and present cargo sequestered inside. Both ...

Nucleoside Triphosphates - From Synthesis to Biochemical Characterization

The traditional strategy for the introduction of chemical functionalities is the use of solid-phase synthesis by appending suitably modified phosphoramidite precursors to the nascent chain. However, the conditions used during the synthesis and the restriction to rather short sequences hamper the applicability of this methodology. On the other hand, modified nucleoside triphosphates are activated building blocks that have been employed for the mild introduction of numerous functional groups into nucleic acids, a strategy that paves the way for the use of modified nucleic acids in a wide-ranging palette of practical applications such as functional tagging and generation of ribozymes and DNAzymes. One of the major challenges resides in the intricacy of the methodology leading to the isolation and characterization of these nucleoside analogues.
	
	In this video article, we present a detailed protocol for the synthesis of these modified analogues using phosphorous(III)-based reagents. In addition, the procedure for their biochemical characterization is divulged, with a special emphasis on primer extension reactions and TdT tailing polymerization. This detailed protocol will be of use for the crafting of modified dNTPs and their further use in chemical biology.

The protocol described herein aims to explain and abridge the numerous obstacles in the way of the intricate route leading to modified nucleoside triphosphates. Consequently, this protocol facilitates both the synthesis of these activated building-blocks and their availability for practical applications.

The  traditional strategy for the introduction of chemical functionalities is the  use of solid-phase synthesis by appending suitably modified ...

An Inverse Analysis Approach to the Characterization of Chemical Transport in Paints

The ability to directly characterize chemical transport and interactions that occur within a material (i.e., subsurface dynamics) is a vital component in understanding contaminant mass transport and the ability to decontaminate materials. If a material is contaminated, over time, the transport of highly toxic chemicals (such as chemical warfare agent species) out of the material can result in vapor exposure or transfer to the skin, which can result in percutaneous exposure to personnel who interact with the material. Due to the high toxicity of chemical warfare agents, the release of trace chemical quantities is of significant concern. Mapping subsurface concentration distribution and transport characteristics of absorbed agents enables exposure hazards to be assessed in untested conditions. Furthermore, these tools can be used to characterize subsurface reaction dynamics to ultimately design improved decontaminants or decontamination procedures. To achieve this goal, an inverse analysis mass transport modeling approach was developed that utilizes time-resolved mass spectroscopy measurements of vapor emission from contaminated paint coatings as the input parameter for calculation of subsurface concentration profiles. Details are provided on sample preparation, including contaminant and material handling, the application of mass spectrometry for the measurement of emitted contaminant vapor, and the implementation of inverse analysis using a physics-based diffusion model to determine transport properties of live chemical warfare agents including distilled mustard (HD) and the nerve agent VX.

In this paper, a procedure for quantifying the mass transport parameters of chemicals in various materials is presented. This process involves employing an inverse-analysis based diffusion model to vapor emission profiles recorded by real-time, mass spectrometry in high vacuum.

The ability to directly characterize chemical transport and interactions that occur within a material (i.e., subsurface dynamics) is a vital component in ...

Synthesis and Characterization of Functionalized Metal-organic Frameworks

Metal-organic frameworks have attracted extraordinary amounts of research attention, as they are attractive candidates for numerous industrial and technological applications. Their signature property is their ultrahigh porosity, which however imparts a series of challenges when it comes to both constructing them and working with them. Securing desired MOF chemical and physical functionality by linker/node assembly into a highly porous framework of choice can pose difficulties, as less porous and more thermodynamically stable congeners (e.g., other crystalline polymorphs, catenated analogues) are often preferentially obtained by conventional synthesis methods. Once the desired product is obtained, its characterization often requires specialized techniques that address complications potentially arising from, for example, guest-molecule loss or preferential orientation of microcrystallites. Finally, accessing the large voids inside the MOFs for use in applications that involve gases can be problematic, as frameworks may be subject to collapse during removal of solvent molecules (remnants of solvothermal synthesis). In this paper, we describe synthesis and characterization methods routinely utilized in our lab either to solve or circumvent these issues. The methods include solvent-assisted linker exchange, powder X-ray diffraction in capillaries, and materials activation (cavity evacuation) by supercritical CO2 drying. Finally, we provide a protocol for determining a suitable pressure region for applying the Brunauer-Emmett-Teller analysis to nitrogen isotherms, so as to estimate surface area of MOFs with good accuracy.

Synthesis, activation, and characterization of intentionally designed metal-organic framework materials is challenging, especially when building blocks are incompatible or unwanted polymorphs are thermodynamically favored over desired forms. We describe how applications of solvent-assisted linker exchange, powder X-ray diffraction in capillaries and activation via supercritical CO2 drying, can address some of these challenges.

Metal-organic frameworks have attracted extraordinary amounts of research attention, as they are attractive candidates for numerous industrial and ...

Preparing Adherent Cells for X-ray Fluorescence Imaging by Chemical Fixation

X-ray fluorescence imaging allows us to non-destructively measure the spatial distribution and concentration of multiple elements simultaneously over large or small sample areas. It has been applied in many areas of science, including materials science, geoscience, studying works of cultural heritage, and in chemical biology. In the case of chemical biology, for example, visualizing the metal distributions within cells allows us to study both naturally-occurring metal ions in the cells, as well as exogenously-introduced metals such as drugs and nanoparticles. Due to the fully hydrated nature of nearly all biological samples, cryo-fixation followed by imaging under cryogenic temperature represents the ideal imaging modality currently available. However, under the circumstances that such a combination is not easily accessible or practical, aldehyde based chemical fixation remains useful and sometimes inevitable. This article describes in as much detail as possible in the preparation of adherent mammalian cells by chemical fixation for X-ray fluorescent imaging.

Here, we present a protocol on how to determine the quantity and distribution of metals in a sample using synchrotron X-ray fluorescence. We focus on adherent cells, and describe the chemical fixation method to prepare this sample. We then describe how to mount and image the sample using synchrotron X-rays.

X-ray fluorescence imaging allows us to non-destructively measure the spatial distribution and concentration of multiple elements simultaneously over ...

Synthesis and Characterization of Supramolecular Colloids

Control over colloidal assembly is of utmost importance for the development of functional colloidal materials with tailored structural and mechanical properties for applications in photonics, drug delivery and coating technology. Here we present a new family of colloidal building blocks, coined supramolecular colloids, whose self-assembly is controlled through surface-functionalization with a benzene-1,3,5-tricarboxamide (BTA) derived supramolecular moiety. Such BTAs interact via directional, strong, yet reversible hydrogen-bonds with other identical BTAs. Herein, a protocol is presented that describes how to couple these BTAs to colloids and how to quantify the number of coupling sites, which determines the multivalency of the supramolecular colloids. Light scattering measurements show that the refractive index of the colloids is almost matched with that of the solvent, which strongly reduces the van der Waals forces between the colloids. Before photo-activation, the colloids remain well dispersed, as the BTAs are equipped with a photo-labile group that blocks the formation of hydrogen-bonds. Controlled deprotection with UV-light activates the short-range hydrogen-bonds between the BTAs, which triggers the colloidal self-assembly. The evolution from the dispersed state to the clustered state is monitored by confocal microscopy. These results are further quantified by image analysis with simple routines using ImageJ and Matlab. This merger of supramolecular chemistry and colloidal science offers a direct route towards light- and thermo-responsive colloidal assembly encoded in the surface-grafted monolayer.

A protocol for the synthesis and characterization of colloids coated with supramolecular moieties is described. These supramolecular colloids undergo self-assembly upon the activation of the hydrogen-bonds between the surface-anchored molecules by UV-light.

Control over colloidal assembly is of utmost importance for the development of functional colloidal materials with tailored structural and mechanical ...

Microfluidic Dry-spinning and Characterization of Regenerated Silk Fibroin Fibers

The protocol demonstrates a method for mimicking the spinning process of silkworm. In the native spinning process, the contracting spinning duct enables the silk proteins to be compact and ordered by shearing and elongation forces. Here, a biomimetic microfluidic channel was designed to mimic the specific geometry of the spinning duct of the silkworm. Regenerated silk fibroin (RSF) spinning doped with high concentration, was extruded through the microchannel to dry-spin fibers at ambient temperature and pressure. In the post-treated process, the as-spun fibers were drawn and stored in ethanol aqueous solution. Synchrotron radiation wide-angle X-ray diffraction (SR-WAXD) technology was used to investigate the microstructure of single RSF fibers, which were fixed to a sample holder with the RSF fiber axis normal to the microbeam of the X-ray. The crystallinity, crystallite size, and crystalline orientation of the fiber were calculated from the WAXD data. The diffraction arcs near the equator of the two-dimensional WAXD pattern indicate that the post-treated RSF fiber has a high orientation degree.

A protocol for the microfluidic spinning and microstructure characterization of regenerated silk fibroin monofilament is presented.

The protocol demonstrates a method for mimicking the spinning process of silkworm. In the native spinning process, the contracting spinning duct enables ...

Synthesis and Evaluation of a Ruthenium-based Mitochondrial Calcium Uptake Inhibitor

We detail the synthesis and purification of a mitochondrial calcium uptake inhibitor, [(OH2)(NH3)4Ru(&#181;-O)Ru(NH3)4(OH2)]5+. The optimized synthesis of this compound commences from [Ru(NH3)5Cl]Cl2 in 1 M NH4OH in a closed container, yielding a green solution. Purification is accomplished with cation-exchange chromatography. This compound is characterized and verified to be pure by UV-vis and IR spectroscopy. The mitochondrial calcium uptake inhibitory properties are assessed in permeabilized HeLa cells by fluorescence spectroscopy.

A protocol for the synthesis, purification, and characterization of a ruthenium-based inhibitor of mitochondrial calcium uptake is presented. A procedure to evaluate its efficacy in permeabilized mammalian cells is demonstrated.

We detail the synthesis and purification of a mitochondrial calcium uptake inhibitor, [(OH2)(NH3)4Ru(&#181;-O)Ru(NH3)4(OH2)]5+. The optimized synthesis of ...

An Engineered Split-TET2 Enzyme for Chemical-inducible DNA Hydroxymethylation and Epigenetic Remodeling

DNA methylation is a stable and heritable epigenetic modification in the mammalian genome and is involved in regulating gene expression to control cellular functions. The reversal of DNA methylation, or DNA demethylation, is mediated by the ten-eleven translocation (TET) protein family of dioxygenases. Although it has been widely reported that aberrant DNA methylation and demethylation are associated with developmental defects and cancer, how these epigenetic changes directly contribute to the subsequent alteration in gene expression or disease progression remains unclear, largely owing to the lack of reliable tools to accurately add or remove DNA modifications in the genome at defined temporal and spatial resolution. To overcome this hurdle, we designed a split-TET2 enzyme to enable temporal control of 5-methylcytosine (5mC) oxidation and subsequent remodeling of epigenetic states in mammalian cells by simply adding chemicals. Here, we describe methods for introducing a chemical-inducible epigenome remodeling tool (CiDER), based on an engineered split-TET2 enzyme, into mammalian cells and quantifying the chemical inducible production of 5-hydroxymethylcytosine (5hmC) with immunostaining, flow cytometry or a dot-blot assay. This chemical-inducible epigenome remodeling tool will find broad use in interrogating cellular systems without altering the genetic code, as well as in probing the epigenotype&#8722;phenotype relations in various biological systems.

Detailed protocols for introducing an engineered split-TET2 enzyme (CiDER) into mammalian cells for chemical inducible DNA hydroxymethylation and epigenetic remodeling are presented.