Characterization of Lipid A Variants by Energy-Resolved Mass Spectrometry: Impact of Acyl Chains
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Lipid A molecules consist of a diglucosamine sugar core with a number of appended acyl chains that vary in their length and connectivity. Because of the challenging nature of characterizing these molecules and differentiating between isomeric species, an energy-resolved MS/MS strategy was undertaken to track the fragmentation trends and map genealogies of product ions originating from consecutive cleavages of acyl chains. Generalizations were developed based on the number and locations of the primary and secondary acyl chains as well as variations in preferential cleavages arising from the location of the phosphate groups. Secondary acyl chain cleavage occurs most readily for lipid A species at the 3′ position, followed by primary acyl chain fragmentation at both the 3′ and 3 positions. In the instances of bisphosphorylated lipid A variants, phosphate loss occurs readily in conjunction with the most favorable primary and secondary acyl chain cleavages.
KeywordsTandem mass spectrometry Collisional activation Lipid A Acyl chain
Gram-negative bacteria are responsible for a variety of food-borne illnesses, respiratory and urinary tract infections, sexually transmitted infections, and highly aggressive immune responses, which can be fatal if proper treatment is unavailable [1, 2, 3, 4]. The outer membrane of gram-negative organisms contains a structurally complex lipopolysaccharide (LPS) responsible for many of their toxic properties [5, 6]. LPS is structurally organized into three unique, covalently linked regions, which extend from the surface of the bacterial membrane . At the exterior is a highly variable polyglycan O-antigen region, connected to a nonrepeating saccharide core, followed by a hydrophobic domain known as lipid A, which is responsible for fastening LPS to the membrane surface [8, 9]. Structural changes in the lipid A domain impact both the ability of the mammalian innate immune system to recognize LPS and play an important role in resistance to key antibiotics (e.g., polymyxins).
The sugar backbone of lipid A is highly conserved in most bacteria, comprised of two pyranosidic hexosamine residues as β(1′,6)-linked glucosamine dimers. Branching from this central motif are a series of highly variable (R)-3-hydroxy acyl groups linked at the O-3′, N-2′, O-3, or N-2 positions by ester or amide bonds . Owing to random mutations along with certain modulating environmental factors, the length and number of acyl chains extending from the sugar backbone of lipid A vary across bacterial species and in response to environmental stress . One or two phosphates are usually found at the 1′ and 4′ carbon positions of the diglucosamine backbone, occasionally with variable substituents, including sugars and phosphoethanolamine groups, the latter reducing the anionic character of the lipid and impacting resistance to interaction with cationic antimicrobial peptides . One example of this structural variability is found in the lipid A of P. aeruginosa isolated from the airways of cystic fibrosis patients [13, 14]. The lipid A present in the clinical isolates incorporated both a palmitate moiety and an aminoarabinose sugar, and retained a C10 fatty acid, modifications specific for P. aeruginosa cystic fibrosis isolates. This finding suggested unique adaptation of the bacterium in a way that may contribute to antimicrobial resistance [13, 14]. Currently, our understanding of the structural changes of lipid A and the way in which they help impart antibiotic resistance and recognition failure by the immune system is limited. As a result, it is imperative that efficient and reliable methods be developed to improve analysis of the diverse structures of lipid A and LPS.
Currently, most mass spectrometry-based lipid analyses rely on MS/MS in order to generate fragmentation profiles for structural characterization. Collision induced dissociation (CID) and high-energy collisional dissociation (HCD) have been particularly useful in generating fragment libraries necessary for accurate profiling of different lipid A variants [13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23]. An alternative to collision-based methods for ion activation/fragmentation is ultraviolet photodissociation (UVPD) [24, 25, 26]. As shown previously, UVPD results in structure-specific glycosidic and cross-ring fragments of lipid A and affords richer fragmentation patterns compared with conventional collisional activation methods [27, 28, 29, 30, 31, 32, 33, 34, 35]. Multi-stage MSn strategies have also been important for structural characterization of lipid A [36, 37, 38]. Recent work by the Goodlett group successfully determined the structures of various lipid A using an automated hierarchical tandem MS algorithm known as HitMS [37, 38]. This profiling method interrogated diagnostic fragment ions and neutral losses across MSn events by aligning each fragment (based on m/z value) with its most probable structure. In order to overcome some of the experimental and data processing obstacles encountered in current lipid A studies, our group recently demonstrated a hierarchical method, UVliPiD, for identification of acyl chain linkages in bis-phosphoryl lipid A structures . Following fragmentation of lipid A by an initial stage of UVPD, the resulting charge-reduced photodetachment products were subjected to secondary activation using CID without isolation of specific intermediates. The rather naïve guesswork of this decision-based approach spurred our interest in refining the method by incorporating specific predictive rules based on monitoring the fragmentation pathways as a function of energy deposition. We envisioned that a combination of MSn and energy-variable MS/MS strategies, ideally coupled with powerful algorithms like HitMS, could further facilitate characterization of lipid A and assignment of structures.
In recent years, energy-resolved mass spectrometry (ERMS) has proven to be a valuable technique for mapping complex oligosaccharides and other biopolymers based on trends in fragmentation relative to internal energy deposited during the activation event [40, 41]. Even for molecules that generate the identical sets of fragment ions upon MS/MS, ERMS may allow an opportunity to differentiate isomers based on variations in the fragmentation trends as a function of energy. Here we use energy-variable HCD, CID, and UVPD and selected MSn experiments to compare the preferred fragmentation pathways and map the genealogies of fragmentation of a series of lipid A molecules. This effort is aimed at developing rules for predicting lipid A fragmentation patterns and to facilitate development of fragment ion assignment algorithms. As shown in the study, HCD proved to be particularly useful for tracking secondary or consecutive fragmentation routes. Further, changes in phosphorylation appeared to cause predictable variations in specific dissociation events.
Synthetic monophosphoryl lipid A (lAA), synthetic monophosphoryl 3-deacyl lipid A (lAC), and detoxified monophosphoryl lipid A from Salmonella minnesota R595 (lAB) were purchased from Avanti Polar Lipids (Alabaster, AL, USA) and used without further purification. E. coli bisphosphorylated lipid A (lAG) was purchased from Sigma-Aldrich (St. Louis, MO, USA) and also used without further purification. Additionally, lipid A variants were isolated and purified from E. coli expressing LpxJ from C. jejuni (lAJ and 1AI), from Acinetobacter baumannii (lAF), from W. succinogenes (lAH), and from E. coli strain BN2 (lAD and lAE), as described previously [33, 42, 43]. HPLC grade chloroform, methanol, and water used during sample preparation were purchased from EMD Millipore (Billerica, MA, USA). Manual determination of fragment ions was accomplished using the ChemDraw software suite (Perkin Elmer, Waltham, MA, USA).
All lipid A samples used in this study were dissolved in 74:23:3 chloroform:methanol:water with final concentrations of 10 μM followed by analysis using both a Thermo Scientific Velos Pro dual linear ion trap and a Thermo Scientific Orbitrap Fusion mass spectrometer (San Jose, CA, USA). Both mass spectrometers were equipped with a 193 nm excimer laser (Coherent, Santa Clara, CA, USA) to perform photodissociation, as previously described [27, 44]. Static emitters were used to transport lipid A molecules into the gas phase via a 1.5 mm o.d. glass capillary pulled to a tip of less than 1 μm by a Sutter Instrument P2000 laser puller (Novato, CA, USA). Solutions of the lipid A molecules were loaded into the pulled glass capillary and a platinum wire was inserted, generating ions when a potential of 1000–1500 V was applied to the wire. All experiments were conducted in the negative ion mode, and CID, HCD, and UVPD were manually stepped to generate ERMS plots for the MS2 spectra. CID and HCD were stepped from NCE 0–50 in increments of 5 NCE, and UVPD was stepped in both energy (0–4 mJ, in 0.5 mJ increments) and pulses (1–9 pulses, in 2 pulse increments). For high resolution data collection on the Orbitrap Fusion mass spectrometer, a resolution of 120 K and a signal-to-noise threshold of 3 was used in analyzing the deconvoluted data in Xtract.
Results and Discussion
ESI of lipid A species resulted in production of singly or doubly deprotonated species depending on whether the lipid A possessed one or two phosphate groups. HCD and UVPD were used to characterize the deprotonated species in an energy-variable manner to obtain information about the dominant fragmentation pathways and their genealogies. ERMS plots were generated via variation of the collision energy for HCD or the number of laser pulses for UVPD. The ERMS plots give insight into the variations in fragment ion distributions as a function of energy deposition, thus providing a convenient way to track the conversion of primary fragment ions into secondary fragment ions and reveal the relative labilities of the acyl chains and the sequential order in which they are lost. For the results shown, fragmentation nomenclature from Domon and Costello  and Morrison et al.  is adopted. All lipid A variants used in this study are shown in Supplementary Figure S1 and listed with their monoisotopic masses and abbreviations used henceforth.
Details about the nature of the consecutive fragmentation pathways (i.e., genealogies) were confirmed by selected MSn experiments. For example, the ion labeled as 3′β + 3α in Figure 1 is isobaric with a product corresponding to cleavages of 3′α + 3β, and thus MS3 experiments were performed to confirm the identity as shown in Supplementary Figure S4. MS3 HCD experiments on the 3α cleavage product ion (Supplementary Figure S4b) reveal sequential cleavages at the 3′ε, 3′β, and 3′α positions. The series of consecutive fragmentation pathways is similarly reflected in the ERMS trends in Figure 1e, for which the abundance of the 3α cleavage product ion decreases at higher collision energies as the 3′α + 3α, 3′ε + 3α, and 3′β + 3α products emerge. MS3 HCD experiments on the 3′α cleavage product ion (Supplementary Figure S4c) demonstrates consecutive cleavage at the 3α position or a cross-ring cleavage at the 0,4A2 position. This experiment confirms that the ion assignment of 3′β + 3α is correct, and there is no evidence for an isobaric 3′α + 3β product.
For each of the monophosphoryl lipid A, cleavages of the 3′ and 3 acyl chains are prominent, either alone at the lower collision energies or in conjunction with additional acyl chain cleavages at higher collision energies. The higher energy onset of the combined loss of the acyl chains at the 3′ and 3 positions suggests that loss of the 3′ chain occurs first, followed by the acyl chain at the 3 position. In contrast to the trends observed for lAA for which there was an abundant ion attributed to combined 3′ε and 3α cleavages (Figure 1e), this pathway is not highly favored for lAC and, instead, cleavage of the 3′ chain (3′ε) in conjunction with cross-ring cleavage of the reducing end sugar (2,4A2) is favored (Figure 3d). This observation is particularly useful for structural elucidation of the acyl chain positions of lipid A. Although the secondary 3′ chain was not present for the other two monophosphoryl lipid A structures, lAD and lAE, they still exhibited combinatorial losses of acyl chains as a function of increasing collisional energy. lAD favored initial cleavage at the 3′α position, whereas lAE favored cleavage at the 2′ε position. Both of these monophosphoryl lipid A isomers exhibited similar 2′ε and 3′α cleavages in conjunction with the acyl chain at the 3 position (Figure 3e, f).
Other correlations and contrasts can be drawn by comparison of the ERMS trends for lipid A species that have identical acyl chain patterns but different number of phosphate groups or ones with the same phosphorylation patterns but different arrangement of acyl chains. For example, lAB and lAG have identical acyl chain patterns, but lAB has a single phosphate group whereas lAG possesses two phosphate groups. In the case of mono-phosphate lAA, the dominant primary and secondary fragment ions observed originated from cleavage of an acyl chain or a cross-ring cleavage occurring (Figure 1). Bisphosphate lAG shows many of these same characteristics, but several of the corresponding pathways are only observed in conjunction with phosphate loss (Figure 5d). For example, the dual cleavage 3′ε + 3α, 3′α + 3α, and 3′β + 3α pathways for lAA occur only with phosphate loss for lAG. Phosphate loss directly from lAA renders the resulting ions nondetectable in the negative mode.
Comparison of lAF and lAH shows the influence of the number of secondary chains on the fragmentation patterns. lAF has two secondary acyl chains present, at the 3′ and 2 positions, whereas lAH has only has a single secondary acyl chain located at the 3′ position. Both molecules favor the 3′ε cleavage (Figure 5c and Figure 6d), and the loss of the additional secondary chain at the 2 position (2ε cleavage) for lAF (Figure 5c) is only observed for one pathway involving multiple acyl chain losses at higher collision energies. In fact, loss of the secondary acyl chain at the 2 position is never a dominant process for lAF (or lAJ).
Energy-resolved mass spectrometry of lipid A molecules enables detailed structural characterization via the genealogical patterns that emerge as specific chains are sequentially cleaved. HCD undertaken in an energy-resolved mode yields the most comprehensive information about the combinatorial cleavage of acyl chains and the preferential cleavage of the chains. Cleavage of a 3′ secondary chain is most favorable for lipid A, followed by cleavages of primary acyl chains at the 3 and 3′ position. A similar pattern, identifying the C3 primary acyl chain and the C3′ secondary acyl chain as the most labile upon collisional activation, has been reported by Sándor et al. for nonphosphorylated lipid A species . For bisphosphorylated lipid A species, loss of a phosphate moiety occurs in conjunction with dominant secondary and primary acyl chain fragmentation. In some instances, cross-ring cleavages of the reducing sugar are observed, but the structural features that modulate this pathway have not yet been deciphered. Recent work presented by Sándor et al. suggests that for nonphosphorylated lipid A species, cross-ring cleavages at the 0,4A2-position generally occur in conjunction with secondary acyl chain fragmentation at the ε-positions . While cross-ring cleavage of the mono- and bis-phosphorylated lipid A species analyzed in the present study did not occur for every lipid, it did occur in conjunction with primary and secondary acyl chain fragmentation at the C3′ positions. Implementation of the full ERMS strategy has limited feasibility on a chromatographic timescale owing to the brief elution profile of each lipid A compared with the number of scan averages to achieve confident characterization. However, a targeted LC experiment during which two or three collisional energies are utilized for each lipid A molecule could allow an efficient means to identify the most labile acyl chains at lower energies and provide total acyl chain content at higher energies.
Funding from the NIH (R01 GM103655 to J.S.B. and RO1s AI064184 and AI076322 to M.S.T.) and the Welch Foundation (F-1155) is gratefully acknowledged.
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