Name: enrichment of bacterial lipoproteins and preparation of n-terminal lipopeptides for structural determination by mass spectrometry
Uploaded: 2018-05-21T12:30:00.000+00:00
Duration: 10 min 59 s
Description: Watch this Scientific Journal Video about Enrichment of Bacterial Lipoproteins and Preparation of N-terminal Lipopeptides for Structural Determination by Mass Spectrometry at JoVE.com

Research in the Meredith lab was supported by startup funds provided by the Eberly College of Science (Pennsylvania State University). We thank Dr. Tatiana Laremore for expert technical advice and access to equipment at the Penn State Proteomics and Mass Spectrometry Core Facility, University Park, PA, where mass spectrometric analyses were performed.

1. Cell Growth and Lysis
<ol>
	<li>Grow bacteria in 15 mL tryptic soy broth (TSB) or similar rich media to late exponential phase (OD600 of 1.0 - 1.5). Harvest cells by centrifugation, wash once with Tris-buffered saline/EDTA (TBSE) and continue with protocol or freeze until use. 
	NOTE: TBSE: 20 mM Tris-hydrochloride (HCl), pH 8.0, 130 mM sodium chloride (NaCl), and 5 mM ethylenediaminetetraacetic acid (EDTA)). Lipoprotein expression and modification can be influenced by growth conditions (e.g. acidity, salinity, growth media) and growth phase15. Cell growth can be scaled as desired, but 15-mL of cells is recommended for a single preparation of lipoproteins as excess biomass can decrease lipoprotein yield. One 15-mL preparation generally yields enough sample for optimal loading of ~2-4 lanes on a standard SDS-PAGE mini gel.</li>
	<li>Resuspend cells in 800 &#956;L TBSE with 1 mM phenylmethyl sulfonyl fluoride (PMSF) and 0.5 mg/mL lysozyme. Transfer solution to a 2.0-mL threaded microcentrifuge tube with screw cap and O-ring and incubate for 20 min at 37 &#176;C. 
	NOTE: Certain species are not susceptible to lysis by lysozyme. An appropriate lytic enzyme should be substituted to aid in lysis.</li>
	<li>Add ~800 &#956;L 0.1 mm zirconia/silica beads to the tube and disrupt cells by shaking at maximum speed (7,000 rpm) on a homogenizer for 5 cycles of 30 s each, with 2 min rest on ice between each cycle.</li>
	<li>Centrifuge sample at 3000 x g for 5 min at 4 &#176;C (to pellet beads and unbroken cells). Transfer the supernatant to a new 2.0-mL microcentrifuge tube and keep on ice.</li>
	<li>Add 200 &#956;L TBSE to the remaining pellet and return to the homogenizer for an additional cycle. Centrifuge (as above) and combine supernatant with the previous supernatant (should be ~800-1000 &#956;L total volume).</li>
</ol>
2. Enrichment of Lipoproteins by TX-114 Phase Partitioning
<ol>
	<li>Supplement the supernatant with TX-114 surfactant to a final concentration of 2% (vol/vol)by adding an equal volume of 4% (vol/vol) the surfactant in ice-cold TBSE and incubate on ice for 1 h, mixing by inversion every ~15 min. 
	NOTE: When chilled, the supernatant and surfactant will be miscible.</li>
	<li>Transfer the tube to a 37 &#176;C water bath and incubate for 10 min to induce phase separation. Centrifuge the sample at 10,000 x g for 10 min at room temperature to maintain bi-phasic separation.</li>
	<li>Gently pipette off the upper aqueous phase and discard. Add ice-cold TBSE to the lower surfactant phase to refill the tube to its original volume and invert to mix. Incubate on ice for 10 min.</li>
	<li>Transfer the tube to a 37 &#176;C water bath and incubate for 10 min to induce phase separation, then centrifuge at 10,000 x g for 10 min at room temperature.</li>
	<li>Repeat steps 2.3-2.4 once more for a total of 3 separations. Remove the upper aqueous phase and discard.</li>
	<li>Remove pellet of precipitated proteins that formed during the course of extractions (visible at the bottom of the tube), by adding 1 volume of ice-cold TBSE to the surfactant phase. Centrifuge at 4 &#176;C at 16,000 x g for 2 min to pellet insoluble protein. 
	NOTE: Sample should remain a single phase. If phase separation occurs, re-chill sample and centrifuge again. The pellet is generally composed of non-lipoprotein contaminants, but can be retained for further analysis.</li>
	<li>Immediately transfer the supernatant to a fresh 2.0-mL microcentrifuge tube containing 1250 &#956;L of 100% acetone. Pipette up and down thoroughly to wash the sample from the tip as it will be viscous. Mix by inversion and incubate overnight at -20 &#176;C to precipitate protein.</li>
	<li>Centrifuge the sample at 16,000 x g for 20 min at room temperature to pellet lipoproteins with attention to the orientation of the tube. Lipoproteins will form a thin, white film along the outside wall of the tube.</li>
	<li>Wash pellet twice with 100% acetone. Decant acetone and allow sample to air dry.</li>
	<li>Add 20-40 &#956;L of 10 mM Tris-HCl, pH 8.0 or a standard Laemmli SDS-PAGE sample buffer16 and thoroughly resuspend by pipetting up and down against the wall with the precipitated lipoproteins. Scrape the side of the tube with the pipette tip to dislodge lipoproteins. 
	NOTE: Lipoproteins will not dissolve in Tris buffer, but rather form a suspension.
	<ol>
		<li>Store lipoproteins at -20 &#176;C until use.</li>
	</ol>
	</li>
</ol>
3. SDS-PAGE, Electroblotting, and Staining with Ponceau S
<ol>
	<li>Separate lipoproteins by SDS-PAGE using standard methods16. 
	NOTE: The appropriate gel acrylamide percentage will vary depending on its intended use and size of lipoproteins. Here, E. faecalis lipoproteins were separated over a 10% Tris-glycine gel, while a 16.5% Tris-tricine gel was used for the smaller E. coli lipoprotein Lpp17.</li>
	<li>Transfer the lipoproteins to a nitrocellulose transfer membrane using a standard electroblotting procedure. 
	NOTE: A semi-dry transfer using Bjerrum Schafer-Nielsen buffer plus 0.1% SDS was performed at 25 V, 1.3 mA for 15 min18. Alternately, polyvinylidene difluoride (PVDF) membrane could be used, however as it is more hydrophobic than nitrocellulose, it may reduce efficient elution of hydrophobic lipopeptides from the membrane at later steps.</li>
	<li>Transfer the nitrocellulose membrane to a container and cover with Ponceau S solution (0.2% (w/v) Ponceau S in 5% acetic acid). Rock gently for 5 min or until red-pink bands are visible.</li>
	<li>Pour off Ponceau S solution and carefully rinse the nitrocellulose membrane with dH2O to remove excess stain. 
	NOTE: Ponceau-stained bands will destain rapidly. If bands disappear, repeat staining process.</li>
	<li>With a clean razor blade, excise the desired band and transfer to a microcentrifuge tube. Wash three times with 1 mL of dH2O to completely destain the band. 
	NOTE: The protocol can be paused here. Store the excised section covered with water at -20 &#176;C until use.</li>
	<li>Transfer the section to a clean surface and with a clean razor blade, dice the nitrocellulose strip into small pieces of approximately 1 mm x 1 mm. Collect the pieces into a 0.5-mL low protein binding microcentrifuge tube.</li>
	<li>Wash the pieces twice with 0.5&#160;mL of freshly-prepared 50 mM ammonium bicarbonate (NH4HCO3), pH 7.8 in high-performance liquid chromatography (HPLC) grade water.</li>
</ol>
4. Tryptic Digestion, Lipopeptide Extraction from Nitrocellulose Membrane, Deposition onto MALDI Target, and Data Acquisition
<ol>
	<li>Resuspend nitrocellulose pieces in 20 &#956;L of a 20 &#956;g/mL solution of trypsin in 50 mM NH4HCO3, pH 7.8 in HPLC grade water. Vortex to mix, then spin briefly to ensure that all pieces are fully covered by the trypsin solution. Cover the tube lid using paraffin film to prevent evaporation and incubate digest overnight at 37 &#176;C.</li>
	<li>Spin the sample at 16,000 x g for 30 s and remove the liquid by pipetting. 
	NOTE: This removes trypsinized hydrophilic peptides that can be examined later by MALDI-TOF MS for protein identification.</li>
	<li>Add 50 &#956;L of 0.5% trifluoroacetic acid (TFA) in HPLC grade water. Vortex to mix, then spin the sample briefly to ensure all pieces are covered by the solution. Incubate for 10 min at room temperature. Remove the liquid by pipetting. 
	CAUTION: TFA is harmful if inhaled. Handle with caution and use in the chemical fume hood. Avoid contact with skin.</li>
	<li>Repeat step 4.3 with 50 &#956;L of 10% acetonitrile in HPLC grade water. 
	CAUTION: Acetonitrile is harmful if inhaled. Handle with caution and use in the chemical fume hood. Avoid contact with skin.</li>
	<li>Repeat step 4.3 with 50 &#956;L of 20% acetonitrile in HPLC grade water. 
	NOTE: This removes any moderately hydrophobic peptides loosely bound to the nitrocellulose.</li>
	<li>To elute the tightly-bound lipopeptides, add 15 &#956;L of freshly-made 10 mg/mL &#945;-cyano-4-hydroxycinnamic acid (CHCA) matrix dissolved in chloroform-methanol (2:1, v/v) and incubate for 10 min at room temperature with intermittent vortexing.
	<ol>
		<li>Alternately, add 15 &#956;L chloroform-methanol to elute lipopeptides and mix with matrix at a later time. Other matrices, such as 2,5-dihydroxybenzoic acid (DHB), should be tested as alternatives to CHCA if unsatisfactory results are obtained for a particular lipopeptide composition. 
		CAUTION: Both chloroform and methanol are harmful if inhaled. Handle with caution and use in the chemical fume hood. Avoid contact with skin.</li>
		<li>Optional: To promote sodium adduct formation, supplement the solution of CHCA in chloroform-methanol with aqueous sodium bicarbonate (NaHCO3) to a final concentration of 1 mM.</li>
	</ol>
	</li>
	<li>Spin the sample briefly. With a pipette, carefully transfer the liquid to a new low protein binding microcentrifuge tube. This solution will contain the bulk of the N-terminal lipopeptides. 
	NOTE: The nitrocellulose may partially or fully dissolve in the chloroform-methanol solution. This will not negatively affect MS results, and may be used as an alternate method to increase lipopeptide return. PVDF membrane will not dissolve in the same manner.</li>
	<li>Deposit 1 &#956;L of the eluted lipopeptides with CHCA onto a polished steel MALDI target. 
	NOTE: The chloroform-methanol will evaporate quickly, leaving behind crystallized CHCA and lipopeptides. An optional second 1 &#956;L aliquot can be deposited onto the same spot to increase sample concentration. Chloroform-methanol may spread significantly after deposition onto the target and as such, care should be taken to avoid sample mixing on the target. It is recommended to deposit and shoot multiple spots of each sample.</li>
	<li>Immediately proceed to mass spectrometry. Scan all areas of the spot for lipopeptide signal, especially if the spot contains two layers of sample, as the second layer may push the lipopeptides to the spot&#39;s outer rim. 
	NOTE: Recommended starting laser intensity is 25%, though it may be necessary to increase intensity to acquire signal and sum multiple spectra (20 or more scans) to achieve adequate signal-to-noise ratio. Here, spectra were acquired on a MALDI-TOF-TOF instrument (see the Table of Materials) using a factory-configured instrument method for the reflector positive-ion detection over the 700-3500 m/z range. The instrument was calibrated with a bovine serum albumin (BSA) tryptic peptide mixture.</li>
</ol>

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The authors have nothing to disclose.

The protocol herein describes two distinct stages of lipoprotein characterization: enrichment by TX-114 phase partitioning and structural determination by MALDI-TOF MS. During TX-114 extraction, additional centrifugation removes contaminating proteins that precipitate during this process, followed by acetone precipitation to yield highly enriched lipoproteins. By limiting the scale of each preparation to 15-mL worth of cells, several samples can be easily processed in parallel, and if desired, pooled at the end of the pr...

Bacterial lipoproteins are characterized by a conserved N-terminal lipid-modified cysteine that anchors the globular protein domain to the cell membrane surface. They are universally distributed in bacteria, constituting 2 to 5% of all cellular genes within a typical genome1. Lipoproteins play critical roles in a wide variety of cellular processes, including nutrient uptake, signal transduction, assembly of protein complexes, and in maintaining cell envelope structural integrity2. In pathogenic bacteria, lipoproteins serve as virulence factors3,4. During an infection, recognition of N-terminal lipopeptides by Toll-like receptor (TLR) 2 incites an innate immune response to remove invading pathogens. Depending on the N-terminal acylation state, lipoproteins are generally recognized by alternate TLR2 heterodimeric complexes. TLR2-TLR1 recognizes N-acylated lipopeptides, while TLR2-TLR6 binds free lipopeptide &#945;-amino termini. Once bound, the signaling pathways converge to induce secretion of proinflammatory cytokines3,4.
It was previously thought that lipoproteins from Gram-positive bacteria were diacylated and those from Gram-negative bacteria were triacylated, differing in the absence or presence of an amide-linked fatty acid on the conserved N-terminal cysteine residue. This assumption was supported by the lack of sequence orthologs in Gram-positive genomes to Lnt, the Gram-negative N-acyl transferase that forms triacylated lipoproteins5. However, recent studies have revealed lipoprotein triacylation in Gram-positive Firmicutes that lack lnt, as well as three novel N-terminal lipoprotein structures, termed the peptidyl, lyso, and N-acetyl forms6,7,8. These findings raise questions about possible yet-to-be-discovered lipoprotein forms, alongside fundamental questions about how these novel lipoproteins are made and what physiological purpose or advantage the various forms impart. Furthermore, they clearly demonstrate the current inability of genomics to predict lipoprotein structure. Indeed, we recently identified a novel class of lipoprotein N-acyl transferases, called Lit, from Enterococcus faecalis and Bacillus cereus that makes lyso-form lipoproteins9. This indicates the need to experimentally verify lipoprotein structure, which can be challenging due to their extremely hydrophobic nature and limited methods available to characterize their molecular structure.
To facilitate studies on lipoprotein induction of the host immune response, as well as N-terminal structural determination, we have adapted several previously-described protocols in order to purify bacterial lipoproteins and prepare the N-terminal tryptic lipopeptides for analysis by MALDI-TOF MS6,10,11,12. Lipoproteins are enriched using an established Triton X-114 (henceforth referred to as surfactant or TX-114) phase partitioning method, with optimization to remove contaminating non-lipoproteins and increase lipoprotein yield. These lipoproteins are suitable for direct use in TLR assays or for further purification by SDS-PAGE. For MALDI-TOF MS, transfer of the lipoproteins to nitrocellulose membrane provides a scaffold for efficient in situ trypsin digestion, washing, and subsequent elution from the membrane surface, resulting in highly purified N-terminal lipopeptides. Nitrocellulose has been shown to facilitate sample handling and improve sequence coverage for highly hydrophobic peptides from integral membrane proteins13,14, as well as lipoproteins9,10. The method has the additional advantage of fractionating peptides based on polarity, so that intermediate wash solutions can be analyzed for high confidence protein identification simultaneously with N-terminal structural determination in a single experiment. This protocol uniquely features intentional sodium adduct formation to promote parent ion fragmentation towards dehydroalanyl ions during MS/MS, aiding in structural assignment of the N-acylation state. The N-terminus is both the most variable and key feature related to TLR recognition of lipoproteins. Taken together, this protocol has allowed intensive and reproducible studies on lipoproteins, with the individual stages of purification and structural determination by MALDI-TOF MS easily adapted depending on the overall goal of the experiment.

<table><tbody><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.01 mm Zirconia/silica beads</td><td>BioSpec Products</td><td>110791012</td><td></td></tr><tr><td>Acetic acid</td><td>EMD</td><td>AX0073-9</td><td></td></tr><tr><td>Acetone</td><td>EMD</td><td>AX0116-6</td><td></td></tr><tr><td>Acetonitrile</td><td>EMD</td><td>AX0142-6</td><td></td></tr><tr><td>Ammonium bicarbonate</td><td>Fluka Analytical</td><td>09830</td><td></td></tr><tr><td>BioTrace NT Nitrocellulose</td><td>PALL Life Sciences</td><td>66485</td><td></td></tr><tr><td>Bovine serum albumin (BSA) digest standard</td><td>Protea</td><td>PS-204-1</td><td></td></tr><tr><td>Chloroform</td><td>Acros Organics</td><td>423550025</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Fisher Scientific</td><td>BP118</td><td></td></tr><tr><td>HPLC Grade water</td><td>EMD</td><td>WX0008-1</td><td></td></tr><tr><td>Lysozyme</td><td>Fisher Scientific</td><td>BP535-1</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>34860</td><td></td></tr><tr><td>Phenylmethyl sulfonyl fluoride (PMSF)</td><td>Amresco</td><td>0754</td><td></td></tr><tr><td>Pierce trypsin protease</td><td>Thermo Scientific</td><td>90057</td><td></td></tr><tr><td>Ponceau S</td><td>Acros Organics</td><td>161470100</td><td></td></tr><tr><td>Protein LoBind Tube 0.5mL</td><td>Eppendorf</td><td>022431064</td><td></td></tr><tr><td>Sodium bicarbonate</td><td>Sigma-Aldrich</td><td>792519</td><td></td></tr><tr><td>Sodium chloride</td><td>Macron Fine Chemicals</td><td>7581-06</td><td></td></tr><tr><td>Trifluoroacetic acid (TFA)</td><td>Sigma-Aldrich</td><td>299537</td><td></td></tr><tr><td>Tris-hydrochloride (HCl)</td><td>Fisher Scientific</td><td>BP152</td><td></td></tr><tr><td>Triton X-114</td><td>Sigma-Aldrich</td><td>93422</td><td></td></tr><tr><td>Tryptic soy broth (TSB)</td><td>BD</td><td>211822</td><td></td></tr><tr><td>&amp;alpha;-cyano-4-hydroxycinnamic acid (MS Grade)</td><td>Sigma-Aldrich</td><td>C8982</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>MagNALyser</td><td>Roche</td><td></td><td></td></tr><tr><td>Trans-Blot Turbo Transfer System</td><td>Bio-Rad</td><td></td><td></td></tr><tr><td>MALDI-TOF target</td><td>Bruker Daltonics</td><td></td><td></td></tr><tr><td>Ultraflextreme MALDI-TOF-TOF</td><td>Bruker Daltonics</td><td></td><td>Equipped with a 355 nm frequency-tripled Nd:YAG smartbeam-II laser</td></tr></tbody></table>

enrichment of bacterial lipoproteins and preparation of n-terminal lipopeptides for structural determination by mass spectrometry

Lipoproteins are important constituents of the bacterial cell envelope and potent activators of the mammalian innate immune response. Despite their significance to both cell physiology and immunology, much remains to be discovered about novel lipoprotein forms, how they are synthesized, and the effect of the various forms on host immunity. To enable thorough studies on lipoproteins, this protocol describes a method for bacterial lipoprotein enrichment and preparation of N-terminal tryptic lipopeptides for structural determination by matrix-assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF MS). Expanding on an established Triton X-114 phase partitioning method for lipoprotein extraction and enrichment from the bacterial cell membrane, the protocol includes additional steps to remove non-lipoprotein contaminants, increasing lipoprotein yield and purity. Since lipoproteins are commonly used in Toll-like receptor (TLR) assays, it is critical to first characterize the N-terminal structure by MALDI-TOF MS. Herein, a method is presented to isolate concentrated hydrophobic peptides enriched in N-terminal lipopeptides suitable for direct analysis by MALDI-TOF MS/MS. Lipoproteins that have been separated by Sodium Dodecyl Sulfate Poly-Acrylamide Gel Electrophoresis (SDS-PAGE) are transferred to a nitrocellulose membrane, digested in situ with trypsin, sequentially washed to remove polar tryptic peptides, and finally eluted with chloroform-methanol. When coupled with MS of the more polar trypsinized peptides from wash solutions, this method provides the ability to both identify the lipoprotein and characterize its N-terminus in a single experiment. Intentional sodium adduct formation can also be employed as a tool to promote more structurally informative fragmentation spectra. Ultimately, enrichment of lipoproteins and determination of their N-terminal structures will permit more extensive studies on this ubiquitous class of bacterial proteins.

A schematic of the protocol is provided in Figure 1. The lipoprotein-enriched fraction extracted from Enterococcus faecalis ATCC 19433 by TX-114 is shown in Figure 2. For comparison, the banding pattern of the precipitated protein fraction is also shown. Proteins from this fraction were confirmed by MALDI-MS to be highly abundant contaminating proteins other than lipoproteins (Table 1). The mass spectra ...

Watch this Scientific Journal Video about Enrichment of Bacterial Lipoproteins and Preparation of N-terminal Lipopeptides for Structural Determination by Mass Spectrometry at JoVE.com

Enrichment of Bacterial Lipoproteins and Preparation of N-terminal Lipopeptides for Structural Determination by Mass Spectrometry

The enrichment of bacterial lipoproteins using a non-ionic surfactant phase partitioning method is described for direct use in TLR assays or other applications. Further steps are detailed to prepare N-terminal tryptic lipopeptides for structural characterization by mass spectrometry.

Lipoproteins are important constituents of the bacterial cell envelope and potent activators of the mammalian innate immune response. Despite their ...

enrichment-bacterial-lipoproteins-preparation-n-terminal-lipopeptides

Research

JoVE Journal

Immunology and Infection

8.6K Views.  Pennsylvania State University. The enrichment of bacterial lipoproteins using a non-ionic surfactant phase partitioning method is described for direct use in TLR assays or other applications. Further steps are detailed to prepare N-terminal tryptic lipopeptides for structural characterization by mass spectrometry.

Video: Enrichment of Bacterial Lipoproteins and Preparation of N-terminal Lipopeptides for Structural Determination by Mass Spectrometry

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.

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 ...

Chemistry

Defining Substrate Specificities for Lipase and Phospholipase Candidates

Microorganisms produce a wide spectrum of (phospho)lipases that are secreted in order to make external substrates available for the organism. Alternatively, other (phospho)lipases may be physically associated with the producing organism causing a turnover of intrinsic lipids and frequently giving rise to a remodeling of the cellular membranes. Although potential (phospho)lipases can be predicted with a number of algorithms when the gene/protein sequence is available, experimental proof of the enzyme activities, substrate specificities, and potential physiological functions has frequently not been obtained. This manuscript describes the optimization of assay conditions for prospective (phospho)lipases with unknown substrate specificities and how to employ these optimized conditions in the search for the natural substrate of a respective (phospho)lipase. Using artificial chromogenic substrates, such as p-nitrophenyl derivatives, may help to detect a minor enzymatic activity for a predicted (phospho)lipase under standard conditions. Having encountered such a minor enzymatic activity, the distinct parameters of an enzyme assay can be varied in order to obtain a more efficient hydrolysis of the artificial substrate. After having determined the conditions under which an enzyme works well, a variety of potential natural substrates should be assayed for their degradation, a process that can be followed employing distinct chromatographic methods. The definition of substrate specificities for new enzymes, often provides hypotheses for a potential physiological role of these enzymes, which then can be tested experimentally. Following these guidelines, we were able to identify a phospholipase C (SMc00171) that degrades phosphatidylcholine to phosphocholine and diacylglycerol, in a crucial step for the remodeling of membranes in the bacterium Sinorhizobium meliloti upon phosphorus-limiting conditions of growth. For two predicted patatin-like phospholipases (SMc00930 and SMc01003) of the same organism, we could redefine their substrate specificities and clarify that SMc01003 is a diacylglycerol lipase.

Many predicted (phospho)lipases are poorly characterized with regard to their substrate specificities and physiological functions. Here we provide a protocol to optimize enzyme activities, search for natural substrates, and propose physiological functions for these enzymes.

Microorganisms produce a wide spectrum of (phospho)lipases that are secreted in order to make external substrates available for the organism. ...

Biochemistry

Isolation and Preparation of Bacterial Cell Walls for Compositional Analysis by Ultra Performance Liquid Chromatography

The bacterial cell wall is critical for the determination of cell shape during growth and division, and maintains the mechanical integrity of cells in the face of turgor pressures several atmospheres in magnitude. Across the diverse shapes and sizes of the bacterial kingdom, the cell wall is composed of peptidoglycan, a macromolecular network of sugar strands crosslinked by short peptides. Peptidoglycan&#8217;s central importance to bacterial physiology underlies its use as an antibiotic target and has motivated genetic, structural, and cell biological studies of how it is robustly assembled during growth and division. Nonetheless, extensive investigations are still required to fully characterize the key enzymatic activities in peptidoglycan synthesis and the chemical composition of bacterial cell walls. High Performance Liquid Chromatography (HPLC) is a powerful analytical method for quantifying differences in the chemical composition of the walls of bacteria grown under a variety of environmental and genetic conditions, but its throughput is often limited. Here, we present a straightforward procedure for the isolation and preparation of bacterial cell walls for biological analyses of peptidoglycan via HPLC and Ultra Performance Liquid Chromatography (UPLC), an extension of HPLC that utilizes pumps to deliver ultra-high pressures of up to 15,000 psi, compared with 6,000 psi for HPLC. In combination with the preparation of bacterial cell walls presented here, the low-volume sample injectors, detectors with high sampling rates, smaller sample volumes, and shorter run times of UPLC will enable high resolution and throughput for novel discoveries of peptidoglycan composition and fundamental bacterial cell biology in most biological laboratories with access to an ultracentrifuge and UPLC.

The bacterial cell wall is composed of peptidoglycan, a macromolecular network of sugar strands crosslinked by peptides. Ultra Performance Liquid Chromatography provides high resolution and throughput for novel discoveries of peptidoglycan composition. We present a procedure for the isolation of cell walls (sacculi) and their subsequent preparation for analysis via UPLC.

The bacterial cell wall is critical for the determination of cell shape during growth and division, and maintains the mechanical integrity of cells in the ...

Lipid Droplet Isolation for Quantitative Mass Spectrometry Analysis

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of virion morphogenesis. Quantitative lipid droplet proteome analysis can be used to identify proteins that localize to or are displaced from lipid droplets under conditions such as virus infections. Here, we describe a protocol that has been successfully used to characterize the changes in the lipid droplet proteome following infection with HCV. We use Stable Isotope Labeling with Amino Acids in Cell Culture (SILAC) and thus label the complete proteome of one population of cells with &#34;heavy&#34; amino acids to quantitate the proteins by mass spectrometry. For lipid droplet isolation, the two cell populations (i.e.&#160;HCV-infected/&#34;light&#34; amino acids and uninfected control/&#34;heavy&#34; amino acids) are mixed 1:1 and lysed mechanically in hypotonic buffer. After removing the nuclei and cell debris by low speed centrifugation, lipid droplet-associated proteins are enriched by two subsequent ultracentrifugation steps followed by three washing steps in isotonic buffer. The purity of the lipid droplet fractions is analyzed by western blotting with antibodies recognizing different subcellular compartments. Lipid droplet-associated proteins are then separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coomassie staining. After tryptic digest, the peptides are quantified by liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS). Using this method, we identified proteins recruited to lipid droplets upon HCV infection that might represent pro- or antiviral host factors. Our method can be applied to a variety of different cells and culture conditions, such as infection with pathogens, environmental stress, or drug treatment.

Lipid droplets are important organelles for the replication of several pathogens, including the Hepatitis C Virus (HCV). We describe a method to isolate lipid droplets for quantitative mass spectrometry of associated proteins; it can be used under a variety of conditions, such as virus infection, environmental stress, or drug treatment.

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of ...

Isolation of Lipoprotein Particles from Chicken Egg Yolk for the Study of Bacterial Pathogen Fatty Acid Incorporation into Membrane Phospholipids

Staphylococcus aureus and other Gram-positive pathogens incorporate fatty acids from the environment into membrane phospholipids. During infection, the majority of exogenous fatty acids are present within host lipoprotein particles. Uncertainty remains as to the reservoirs of host fatty acids and the mechanisms by which bacteria extract fatty acids from the lipoprotein particles. In this work, we describe protocols for enrichment of low-density lipoprotein (LDL) particles from chicken egg yolk and determining whether LDLs serve as fatty acid reservoirs for S. aureus. This method exploits unbiased lipidomic analysis and chicken LDLs, an effective and economical model for the exploration of interactions between LDLs and bacteria. The analysis of S. aureus integration of exogenous fatty acids from LDLs is performed using high-resolution/accurate mass spectrometry and tandem mass spectrometry, enabling the characterization of the fatty acid composition of the bacterial membrane and unbiased identification of novel combinations of fatty acids that arise in bacterial membrane lipids upon exposure to LDLs. These advanced mass spectrometry techniques offer an unparalleled perspective of fatty acid incorporation by revealing the specific exogenous fatty acids incorporated into the phospholipids. The methods outlined here are adaptable to the study of other bacterial pathogens and alternative sources of complex fatty acids.

This method provides a framework for studying incorporation of exogenous fatty acids from complex host sources into bacterial membranes, particularly Staphylococcus aureus. To achieve this, protocols for the enrichment of lipoprotein particles from chicken egg yolk and subsequent fatty acid profiling of bacterial phospholipids utilizing mass spectrometry are described.

Staphylococcus aureus and other Gram-positive pathogens incorporate fatty acids from the environment into membrane phospholipids. During infection, the ...

Click-Chemistry Based Fluorometric Assay for Apolipoprotein N-acyltransferase from Enzyme Characterization to High-Throughput Screening

Lipoproteins from proteobacteria are posttranslationally modified by fatty acids derived from membrane phospholipids by the action of three integral membrane enzymes, resulting in triacylated proteins. The first step in the lipoprotein modification pathway involves the transfer of a diacylglyceryl group from phosphatidylglycerol onto the prolipoprotein, resulting in diacylglyceryl prolipoprotein. In the second step, the signal peptide of prolipoprotein is cleaved, forming an apolipoprotein, which in turn is modified by a third fatty acid derived from a phospholipid. This last step is catalyzed by apolipoprotein N-acyltransferase (Lnt). The lipoprotein modification pathway is essential in most &#947;-proteobacteria, making it a potential target for the development of novel antibacterial agents. Described here is a sensitive assay for Lnt that is compatible with high-throughput screening of small inhibitory molecules. The enzyme and substrates are membrane-embedded molecules; therefore, the development of an in vitro test is not straightforward. This includes the purification of the active enzyme in the presence of detergent, the availability of alkyne-phospholipids and diacylglyceryl peptide substrates, and the reaction conditions in mixed micelles. Furthermore, in order to use the activity test in a high-throughput screening (HTS) setup, direct readout of the reaction product is preferred over coupled enzymatic reactions. In this fluorometric enzyme assay, the alkyne-triacylated peptide product is rendered fluorescent through a click-chemistry reaction and detected in a multiwell plate format. This method is applicable to other acyltransferases that use fatty acid-containing substrates, including phospholipids and acyl-CoA.

Presented here is a sensitive fluorescence assay to monitor apolipoprotein N-acyltransferase activity using diacylglyceryl peptide and alkyne-phospholipids as substrates with click-chemistry.

Lipoproteins from proteobacteria are posttranslationally modified by fatty acids derived from membrane phospholipids by the action of three integral ...

Semi-Quantitative Analysis of Peptidoglycan by Liquid Chromatography Mass Spectrometry and Bioinformatics

Peptidoglycan is an important component of bacterial cell walls and a common cellular target for antimicrobials. Although aspects of peptidoglycan structure are fairly conserved across all bacteria, there is also considerable variation between Gram-positives/negatives and between species. In addition, there are numerous known variations, modifications, or adaptations to the peptidoglycan that can occur within a bacterial species in response to growth phase and/or environmental stimuli. These variations produce a highly dynamic structure that is known to participate in many cellular functions, including growth/division, antibiotic resistance, and host defense avoidance. To understand the variation within peptidoglycan, the overall structure must be broken down into its constitutive parts (known as muropeptides) and assessed for overall cellular composition. Peptidoglycomics uses advanced mass spectrometry combined with high-powered bioinformatic data analysis to examine peptidoglycan composition in fine detail. The following protocol describes the purification of peptidoglycan from bacterial cultures, the acquisition of muropeptide intensity data through a liquid chromatograph&#8212;mass spectrometer, and the differential analysis of peptidoglycan composition using bioinformatics.

This protocol covers a detailed analysis of peptidoglycan composition using liquid chromatography mass spectrometry coupled with advanced feature extraction and bioinformatic analysis software.

Peptidoglycan is an important component of bacterial cell walls and a common cellular target for antimicrobials. Although aspects of peptidoglycan ...

Cell-Free Production of Proteoliposomes for Functional Analysis and Antibody Development Targeting Membrane Proteins

Membrane proteins play essential roles in a variety of cellular processes and perform vital functions. Membrane proteins are medically important in drug discovery because they are the targets of more than half of all drugs. An obstacle to conducting biochemical, biophysical, and structural studies of membrane proteins as well as antibody development has been the difficulty in producing large amounts of high-quality membrane protein with correct conformation and activity. Here we describe a &#8220;bilayer-dialysis method&#8221; using a wheat germ cell-free system, liposomes, and dialysis cups to efficiently synthesize membrane proteins and prepare purified proteoliposomes in a short time with a high success rate. Membrane proteins can be produced as much as in several milligrams, such as GPCRs, ion channels, transporters, and tetraspanins. This cell-free method contributes to reducing the time, cost and effort for preparing high-quality proteoliposomes, and provides suitable means for functional analysis of membrane proteins, drug targets screening, and antibody development.

This protocol describes an efficient cell-free method for production of high-quality proteoliposome by bilayer-dialysis method using wheat cell-free system and liposomes. This method provides suitable means for functional analysis of membrane proteins, drug targets screening, and antibody development.

Membrane proteins play essential roles in a variety of cellular processes and perform vital functions. Membrane proteins are medically important in drug ...

Sample Preparation using a Lipid Monolayer Method for Electron Crystallographic Studies

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown into highly ordered two-dimensional (2D) crystals under favorable conditions. The quality of the grown 2D crystals is crucial to the resolution of the final reconstruction via 2D image processing. Over the years, lipid monolayers have been used as a supporting layer to foster the 2D crystallization of peripheral membrane proteins as well as soluble proteins. This method can also be applied to 2D crystallization of integral membrane proteins but requires more extensive empirical investigation to determine detergent and dialysis conditions to promote partitioning to the monolayer. A lipid monolayer forms at the air-water interface such that the polar lipid head groups remain hydrated in the aqueous phase and the non-polar, acyl chains, tails partition into the air, breaking the surface tension and flattening the water surface. The charged nature or distinctive chemical moieties of the head groups provide affinity for proteins in solution, promoting binding for 2D array formation. A newly formed monolayer with the 2D array can be readily transfer into an electron microscope (EM) on a carbon-coated copper grid used to lift and support the crystalline array. In this work, we describe a lipid monolayer methodology for cryogenic electron microscopic (cryo-EM) imaging.

Lipid monolayers have been used as a foundation for forming two-dimensional (2D) protein crystals for structural studies for decades. They are stable at the air-water interface and can serve as a thin supporting material for electron imaging. Here we present the proven steps on preparing lipid monolayers for biological studies.

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown ...

High-Resolution Studies of Proteins in Natural Membranes by Solid-State NMR

Membrane proteins are vital for cell function and thus represent important drug targets. Solid-state Nuclear Magnetic Resonance (ssNMR) spectroscopy offers a unique access to probe the structure and dynamics of such proteins in biological membranes of increasing complexity. Here, we present modern solid-state NMR spectroscopy as a tool to study structure and dynamics of proteins in natural lipid membranes and at atomic scale. Such spectroscopic studies profit from the use of high-sensitivity ssNMR methods, i.e., proton-(1H)-detected ssNMR and DNP (Dynamic Nuclear Polarization) supported ssNMR. Using bacterial outer membrane beta-barrel protein BamA and the ion channel KcsA, we present methods to prepare isotope-labeled membrane proteins and to derive structural and motional information by ssNMR.

This work details robust basic routines on how to prepare isotope-labeled membrane protein samples and analyze them at high-resolution with modern solid-state NMR spectroscopy methods.

Membrane proteins are vital for cell function and thus represent important drug targets. Solid-state Nuclear Magnetic Resonance (ssNMR) spectroscopy ...