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CRITICAL REVIEWS IN ORAL BIOLOGY & MEDICINE

Lipopolysaccharide Heterogeneity: Innate Host Responses to Bacterial Modification of Lipid A Structure

D.R. Dixon1,2 and R.P. Darveau1,*

1 Department of Periodontics, University of Washington, Health Sciences Center, Box 357444, Seattle, WA 98195, USA; and
2 The United States Army Dental Corps;

Correspondence: * corresponding author,rdarveau{at}u.washington.edu


   ABSTRACT
TOP
 ABSTRACT
(1) INTRODUCTION
 (2) WHAT ARE LIPOPOLYSACCHARIDES...
 (3) HIGH vs. LOW...
 (4) NATURAL LIPID A...
 (5.0) PERSPECTIVES
 REFERENCES
 
The innate host response system is composed of various mechanisms designed to detect and facilitate host responses to microbial components, such as lipopolysaccharides (LPS). To enable this to occur, innate systems contain multiple pattern recognition receptors (i.e., LBP, CD14, and TLRs), which identify certain features within bacterial LPS that are foreign to the host, as well as essential and uniquely specific for bacteria. Innate host identification of unique bacterial components or patterns, therefore, relies on the inability of bacteria to alter these essential or critical components dramatically. Historically, LPS have been viewed as essential outer-membrane molecules containing both a highly variable outer region (O-segment) as well as a relatively conserved inner region (lipid A). However, over the last decade, new evidence has emerged, revealing that increased natural diversity or heterogeneity within specific components of LPS, such as lipid A—resulting in minor to moderate changes in lipid A structure—can produce dramatic host responses. Therefore, examples of natural lipid A heterogeneity, and the mechanisms that control it, represent a novel approach in which bacteria modulate host responses and may thereby confer specific advantages to certain bacterial species under changing environmental host conditions.

Key Words: lipopolysaccharide (LPS) • lipid A • CD14 • TLR • innate immunity


   (1) INTRODUCTION
TOP
 ABSTRACT
 (1) INTRODUCTION
(2) WHAT ARE LIPOPOLYSACCHARIDES...
 (3) HIGH vs. LOW...
 (4) NATURAL LIPID A...
 (5.0) PERSPECTIVES
 REFERENCES
 
Lipopolysaccharides (LPS) are outer membrane molecules essential for virtually all Gram-negative bacteria. They have been historically described as heat-stable, non-proteinaceous, endotoxic microbial cell wall components consisting of highly variable as well as highly conserved segments (Rietschel et al., 1992; Rietschel and Westphal, 1999). Conserved regions of LPS represent critical molecules that are shared between bacterial species, which assist in either the development and/or maintenance of a component or structure that is essential for survival of the bacterium. The variable regions represent segments that are not essential for the bacterium, allowing for evolutionary variation without catastrophic consequences. Changes within these regions can either result in simple modifications of LPS, such as minor alteration in the length of the segment, or can have dramatic effects, like changing the overall chemical configuration, composition, or attached charge groups, which, in turn, can affect overall structure.

LPS are ubiquitous within our environment, in vivo and in vitro, and can express potent bioactivity in extremely small amounts ({eta}g to {rho}g range) (Rietschel and Westphal, 1999). LPS are known to be capable of initiating the morbidity and mortality associated with Gram-negative sepsis, as well as modulation of myriad other host innate inflammatory responses. For example, the ‘classic’ description of the innate host response to LPS is that LPS significantly activate the innate host defense system. Specifically, LPS have been characterized as the ‘prototypical stimuli’ for host activation through myeloid cells (neutrophils, monocytes, macrophages) and non-myeloid cells (fibroblasts, platelets), as well as other innate host defense mechanisms, such as serum complement, as well as specific components within the intrinsic coagulation pathway.

Conservation within structural components, like LPS, serves as the basis for host identification and response by creating ‘molecular signals’ that are unique to prokaryotic cells, thus allowing for host distinction between self and non-self (Medzhitov and Janeway, 2000). However, new insights within the relatively recently discovered Toll-like receptors (TLRs) and co-associated molecules have revealed that even subtle differences in ‘conserved’ LPS structures and their consequent molecular signals can be identified by host cells, resulting in profound host responses. For example, specific synthetic alterations within conserved segments of LPS were shown to be not only recognized by host cells but also capable of blocking or down-regulating the response to other LPS forms normally associated with robust innate cell activation. Furthermore, naturally occurring LPS heterogeneity of specific pathogenic bacteria, resulting from either altered bacterial growth or environmental conditions, was shown to result in differential and altered innate host cell responses in vitro that could potentially represent or confer some inoculation or evasion advantage, resulting in survival and sustained pathogenicity for the bacterium in vivo.

Discoveries such as these have propagated a new way of thinking about bacterial LPS and host interactions that has differed from the ‘classic’ LPS-host interaction dogma. The purpose of this review, therefore, is to introduce to the reader possible reasons for these subtle LPS host-response differences by first reviewing the classic LPS structures normally associated with host activation and discussing identified bacterial LPS heterogeneity and its effect on known endotoxic components.


   (2) WHAT ARE LIPOPOLYSACCHARIDES (LPS)?
TOP
 ABSTRACT
 (1) INTRODUCTION
 (2) WHAT ARE LIPOPOLYSACCHARIDES...
(3) HIGH vs. LOW...
 (4) NATURAL LIPID A...
 (5.0) PERSPECTIVES
 REFERENCES
 
A review of the physical structures located on the outer surfaces of Gram-negative bacteria is essential for understanding the interaction between the host and invading organisms. This section and the following review will focus on one outer membrane molecule in particular, the LPS.

(2.1) Schematic
The majority of Gram-negative bacteria contain various amphiphilic molecules within the outer membrane; however, LPS represent the main surface antigen, possessing both microbiologic and immunologic significance. Although there are some notable exceptions, many species contain a common form or architecture that consists of specific components: an outer or distal segment, the O-chain or O-antigen; the outer core segment; an inner core segment; and, finally, the medial lipid A portion (Luderitz et al., 1981; Rietschel et al., 1992). Fig. 1Go illustrates the major components of LPS and their relative positions on the bacterial outer membrane surface.


Figure 1
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  		Figure 1. General overview of lipopolysaccharide (LPS) on the outer membrane of a Gram-negative bacterium. LPS consists of 3 major components: the highly variable outer O-antigen segment; a more conserved core, which is divided into outer and inner segments; and the bioactive lipid A portion. Variation within the length of the LPS, due to mutational absence of specific structures, not only changes the phenotypic appearance of the bacterium (i.e., smooth [S], semi-rough [SR], or rough [R]), but may also change some bioactive responses by the host to the bacterium itself. (A) Some bacterial species contain an outer capsule that protects the bacterium from host defenses such as complement, lysis, and phagocytosis. (B) Outer lipid bilayer with LPS which is approximately 8 nm in width. (C) Peptidoglycan layer. (D) Inner bilipid membrane. Note: Additional lipoproteins, porin complexes, and additional membrane proteins established within and surrounding the inner and outer membranes have been removed to simplify the diagram (Raetz, 1992; Caroff et al., 2002).

 
(2.2) LPS are Essential Outer Membrane Molecules
Newly synthesized LPS molecules translocate across the periplasm and outer bilayer membranes of Gram-negative bacterial cells, replacing existing phospholipids, thereby representing up to 75% of the total cellular surface (Caroff and Karibian, 2003). In fact, some estimates suggest that there are as many as 106 molecules of LPS for every one bacterium of E. coli (Raetz and Whitfield, 2002; Caroff and Karibian, 2003). Exactly how LPS translocation occurs and accumulates upon the cell surface is still being examined; however, this molecule is known to be essential (with the exception of the LpxA mutant in Neisseria meningitidis) (Steeghs et al., 1998, 2001; Pridmore et al., 2001; van der Ley and Steeghs, 2003) for the in vitro growth and structural integrity of the bacterium. Specific mutations in LPS synthesis (Zhou et al., 1998) or lipid and membrane component transporters (MsbA or other ABC transporters) (Zhou et al., 1998; Doerrler et al., 2001) may result in LPS accumulating beneath the inner membrane, thereby becoming toxic or lethal to the bacterial cell (Zhou et al., 1998; Doerrler et al., 2001; Raetz and Whitfield, 2002). Therefore, the ‘essentialness’ of LPS for the bacterium stems partially from functions shared between certain membrane translocation systems and common outer membrane assembly and maintenance functions utilized by the organism itself (Vaara, 1993a,b; Raetz and Whitfield, 2002).

For the host, the LPS structure then serves as an important molecular pattern or signal that represents the presence of Gram-negative bacteria, as compared with other signals characteristic of other types of organisms or host components (‘self’), thereby allowing the host to identify it as a potentially infectious, microbial, non-self agent (Medzhitov and Janeway, 2000, 2002).

(2.3) Lipid A: ‘Classic’ Lipid A Structure is Represented by E. coli
During the late 1980s, experimentation with LPS and separated lipid A fractions revealed that lipid A was, in fact, the bioactive component of LPS, responsible for the majority of IL-1 induction and immunoregulation in human mononuclear cells within the pg/mL range (Loppnow et al., 1989, 1990). These experiments were based on and correlated with findings from earlier and contemporary experimentation with synthetic lipid A components (Kotani et al., 1985) and Re-mutants strains (Luderitz et al., 1966; Kasai and Nowotny, 1967)—both of which are devoid of any additional polysaccharide (PS) additions—which resulted in full endotoxic activity when compared with the original LPS forms (Kim and Watson, 1967; Kotani et al., 1985; Rietschel and Westphal, 1999). Results from these and other pivotal experiments (Galanos, 1975) confirmed the role of lipid A as the active LPS constituent, relegating other subcomponents of LPS to ‘non-essential for endotoxicity’ status (Rietschel and Westphal, 1999). Therefore, the latter part of this review will focus mainly on the conserved segments and observed heterogeneity within a specific sub-component of LPS, lipid A, and the biologic ramification(s) thereof.


   (3) HIGH vs. LOW BIOACTIVITY
TOP
 ABSTRACT
 (1) INTRODUCTION
 (2) WHAT ARE LIPOPOLYSACCHARIDES...
 (3) HIGH vs. LOW...
(4) NATURAL LIPID A...
 (5.0) PERSPECTIVES
 REFERENCES
 
In vitro and in vivo biologic assays quickly delineate specific differences in LPS-related biologic activity. This section will discuss common host response assays as well as the possible reasons for observed differences in host responses to LPS.

(3.1) A Profile of ‘Classic’ Potent Endotoxic Effects and Assays
Potency of the toxic nature of LPS components has been historically quantified through the use of host response assays. Here, immunostimulatory and/or immunoregulatory systems were developed to facilitate ranking of LPS components based on the specific structural requirements for activation. For example, the three most common assays used to describe potent endotoxic effects are: (1) the chick embryo lethality test (LD50 dosages calculated after intravenous injection and observation of embryo death by intoxication within 48 hrs); (2) the rabbit pyrogenicity assay (positive results have been defined as an increase in rectal temperature of more than 0.6°C after LPS sample challenge); and (3) the local Shwartzman reaction (where two injections of LPS samples are given 24 hrs apart, the first injected dermally and the second intravenously; a positive reaction occurs if a hemorrhagic reaction develops at the dermal [1st] injection site). Additional in vivo and in vitro assays include lethal toxicity in galactosamine-loaded mice (animals are sensitized with an intraperitoneal injection with D-galactosamine HCl, followed immediately by intravenous injection of test material; death by intoxication is usually observed in 24 hrs), induction or enhancement of IL-1, PGE2, O2– production in murine macrophages, and activation of the human complement cascade. A complete review of the biologic activities and structural requirements to induce these specific activities has been discussed by Takada and Kotani (1992).

(3.2) Structures Associated with High Bioactivity
Lipid A structures required to produce the highest or most potent bioactivity, as measured by the above assays, include: a β, (1'-6)-linked D-glucosamine disaccharide backbone; phosphorylation at positions 1 and 4' on the disaccharide unit; an appropriate number (two was determined to have maximum effect) (Shimizu et al., 1987) of 3-acyloxyacyl groups; the position of these 3-acyloxyacyl groups on either the reducing GlcN I or the distal GlcN II residues of the disaccharide (Takada and Kotani, 1992); and an appropriate number (usually 6) of hydrophobic fatty acid chains with a suitable carbon length (C12-C18) (Kumazawa et al., 1988; Nakatsuka et al., 1989), with or without hydroxylation (Fig. 2AGo). Interestingly, these structures, which yield the most potent endotoxic effects, are highly conserved within enteric and non-enteric Gram-negative bacteria, and are often referred to as the ‘canonical lipid A structure’. Furthermore, alteration in any part of the canonical structure—such as changing the number, position, or length of the primary or secondary acyl groups, or subtraction of phosphate or monosaccharide groups—results in dramatic alteration of the biologic effects (Loppnow et al., 1989; Takada and Kotani, 1992; Schumann et al., 1996).


Figure 2
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  		Figure 2. Lipid A chemical structure examples. (A) Representative lipid A structure for E. coli with a mass ion m/z 1798. The mass ion at m/z 1798 accounts for greater than 80% of the lipid A found in E. coli. It consists of a phosphorylated β(1'-6) D-glucosamine disaccharide substituted with hydroxylated and non-hydroxylated fatty acids. LpxA and LpxD transfer the indicated fatty acids to monomers N-acetyl-glucosamine 1 (GlcN 1) and N-acetyl-glucosamine 2 (GlcN 2). Next, the β(1'-6)-linked disaccharide is generated by a disaccharide synthase encoded by lpxB. lpxK is the kinase structural gene responsible for phosphorylating the disaccharide. Two Kdo residues are transferred to the lipid A disaccharide by a bifunctional enzyme encoded by waaA/kdtA. Following the GlcN subunits’ condensing to form a disaccharide, and in the presence of Kdo, lpxL and lpxM transfer the indicated secondary fatty acids to lipid A (Caroff et al., 2002; Raetz and Whitfield, 2002). (B) Chemical structure of the synthetic intermediate lipid A product (labeled Lipid IVA, compound 406, or LA-14-PP), containing low bioactivity and antagonistic properties in certain cell lines (Wang et al., 1990; Takada and Kotani, 1992; Raetz and Whitfield, 2002).

 
(3.3) Synthetic Structures/Preparations
Lipid A research advanced exponentially with the onset of several synthetic preparations in Escherichia which modified or altered specific lipid A components, and the resultant preparations were then characterized for bioactivity (Galanos et al., 1984, 1985; Kotani et al., 1985). The first milestone in this endeavor was the creation of a synthetic lipid A that possessed the same chemical configuration and exhibited the same biologic activity as the wild-type lipid A. This synthetic lipid A preparation—labeled either compound 506 (Kotani et al., 1985) or compound LA-15-PP (Kanegasaki et al., 1986), which contained the same lipid A disaccharides, phosphorylation patterns as well as the 3-hydroxy- and 3-acyloxy-tetradecanoyl groups at the C2, C3 and C2', C3' positions as the E. coli strain (F515)—was shown to exhibit activity equal to or greater than that of the reference E. coli strain, during both the rabbit pyrogenicity experiments and lethal toxicity assays in galactosamine-sensitized mice, as well as the local Shwartzman reaction (Kotani et al., 1985). In addition, it was also noted that chemical alteration of these preparations—either by number or length of the acyl chains, phosphorylation pattern (addition or subtraction) or overall chemical symmetry of the lipid A structure—resulted in intermediate or low biologic activity within the same biologic assays. Interestingly, significant alteration in the acyl pattern—such as those created in compound(s) IVA, LA-14-PP, or 406, which represent intermediate precursors of lipid A biosynthesis and contain only 4, symmetrically placed, (R)-3-hydroxytetradecanoyl (C14-OH) groups (Fig. 2BGo)—either showed very weak activation properties (Kotani et al., 1985) or, in the case of IVA, showed both agonistic properties in mice but antagonistic properties when mixed with LPS in human cells (Raetz et al., 1985; Golenbock et al., 1991). This dual agonistic and antagonistic effect of compounds such as IVA was partially explained by the presence or absence of human Toll-like receptor 4 (hTLR4) and MD-2 expressed within the species-specific target cells being assayed (Lien et al., 2000; Poltorak et al., 2000; Raetz and Whitfield, 2002).

Further explanations of the various bioactivities seen with different lipid A structures could lie within the three-dimensional changes occurring as a result of the presence or absence of specific lipid A structures themselves. Here, experimental evidence has accumulated which suggests that if the aggregate lipid A structure contains an increased cross-sectional hydrophobic region, compared with that of the hydrophilic part, this will yield a conical lipid A shape with increased bioactivity, whereas cylindrical (both regions equal in cross-section) lipid A molecules are biologically inactive (Rietschel et al., 1991; Schromm et al., 2000). Furthermore, residual 3-deoxy-D-manno-oct-2-ulosonic acid (Kdo) groups, if attached to the lipid A structure, have been known to modify or alter lipid A conformation, resulting in higher bioactivity compared with purified (Kdo absent) preparations (Caroff et al., 1986, 2002; Brade et al., 1997; Muroi and Tanamoto, 2002; Caroff and Karibian, 2003). These results not only imply that there is a critical role for symmetry of acyl chain position and overall molecular shape, but also suggest that Kdo molecules and their corresponding charge groups affect responses to lipid A as well (Caroff et al., 1986, 2002; Brade et al., 1997; Muroi and Tanamoto, 2002; Caroff and Karibian, 2003).

(3.4) Enteric Bacteria vs. Oral Bacteria (High vs. Low Bioactivity)
The interest in differential host responses to LPS generated an expansion of research efforts in other areas of non-enteric microbiologic study, yielding some interesting observations. For example, in the field of oral microbiology, early studies revealed (Mansheim et al., 1978; Fujiwara et al., 1990), and more recent studies confirmed (Darveau et al., 1995; Pulendran et al., 2001), the observation that LPS and their isolated lipid A component, obtained from Gram-negative oral anaerobic bacteria, do not elicit host responses in a manner consistent with the responses observed with the classic E. coli-type endotoxin. Early studies examining endotoxin lethality in mice demonstrated that LPS obtained from Gram-negative oral anaerobic bacteria were significantly reduced in potency when directly compared with enterobacterial LPS (Takada and Galanos, 1987; Isogai et al., 1988) and did not induce a Shwartzman reaction (Mansheim et al., 1978; Fujiwara et al., 1990), a systemic innate immune inflammatory response typically associated with E. coli-type endotoxin (Takada and Kotani, 1992). These observations resulted in this type of LPS being initially designated as having no or very low biological activity. However, one explanation for this lowered activity could be due to the number of phosphate groups or altered lengths and positions of the fatty acids present in lipid A from certain oral bacteria compared with those in enteric E. coli, which has been proven to be critical in Shwartzman reaction induction (Takada and Kotani, 1989). In addition, evidence is accumulating to suggest that, although different from E. coli, P. gingivalis LPS do in fact contain significant biologic activity. For example, LPS obtained from P. gingivalis induce inflammatory responses in non-responder LPS mice (Kirikae et al., 1999), subsequently shown to have a mutation in TLR4 (Poltorak et al., 1998; Qureshi et al., 1999), suggesting that these LPS contain, potentially, a TLR4-independent mechanism for cell activation. Subsequently, numerous in vitro studies by our laboratory (Darveau et al., 1995; Cunningham et al., 1996, 1999) and others (Mansheim et al., 1978; Ogawa et al., 1994; Ogawa and Uchida, 1996) have confirmed that P. gingivalis LPS are less potent, but, perhaps more importantly, they elicit a different pattern or repertoire of inflammatory mediators in innate immune cells (Ogawa and Uchida, 1996; Tanamoto et al., 1997), when compared directly with E. coli LPS. Furthermore, it has also been reported that dendritic cell recognition and response to P. gingivalis and E. coli isolated LPS are significantly different, characterized by altered TLR4 signaling, separate cytokine subset production, and induction of separate and distinctly different in vivo adaptive immune responses (Pulendran et al., 2001).

(3.5) Binding Properties and Host Presentation May Partially Explain Low Biologic Reactive LPS
(i) Lipopolysaccharide Binding Protein (LBP) Binding Affinity, the Highs and Lows
Microbial component-LBP binding affinities might explain, in part, low biological reactive LPS. Since the discovery of the LBP/CD14 host activation pathway (Schumann et al., 1990; Wright et al., 1990), it has become increasingly clear that the release of numerous inflammatory mediators and the expression of cell adhesion molecules necessary for inflammation occur in response to LBP and/or CD14 complexed with microbial components (Pugin et al., 1993b; Wright, 1995). It has been shown that aggregate forms of LPS initially bind LBP (an acute-phase serum protein), which then facilitates the transfer of separated, monomeric LPS to either mCD14, a membrane-bound form, or sCD14, a soluble form found in serum (Pugin et al., 1993b; Tobias et al., 1995; Wright, 1995; Yu and Wright, 1996; Heumann and Roger, 2002). mCD14-positive cells (usually cells of myeloid origin) contain mCD14 as a glycosylphosphatidylinositol (GPI)-anchored membrane protein, whereas the sCD14/LPS complex is required for CD14-negative cell (i.e., endothelial and certain epithelial cells) activation by LPS (Pugin et al., 1993a; Heumann and Roger, 2002).

However, LPS obtained from different bacteria have significantly different binding affinities for LBP, and this is one mechanism by which the host may be able to differentiate between LPS molecules (Cunningham et al., 1996). For example, LPS from both Helicobacter pylori and Porphyromonas gingivalis bind LPB 10 to 100 times less efficiently than E. coli (Cunningham et al., 1996). Less efficient LBP binding may represent one mechanism responsible for the low biologic reactivity especially seen in the activation of human monocytes to specific LPS species (Darveau et al., 1995; Cunningham et al., 1996). Consistent with this, reports of LPS activity within in vitro systems have shown that different concentrations of serum (serum being the major source of both endogenous LBP and sCD14), serum from various species, or heat-inactivated serum affect the level of observed activation (Wright et al., 1990; Pugin et al., 1993a; Shapira et al., 1994). The mechanisms by which LPS bind LBP are not fully understood; however, a bactericidal/permeability-increasing protein (BPI), a protein closely related to LBP, has recently been crystallized (Beamer et al., 1997). Analysis of the structural data obtained from this study demonstrates that BPI contains two apolar pockets which interact with the acyl chains of phosphatidylcholine. This information may now potentially serve as a structural model for LBP interaction with LPS acyl chains. As previously mentioned, the number and type of fatty acyl chain and the corresponding interaction with host-derived molecules/cells are key factors influencing the strength of the inflammatory host response to the lipid A component of LPS.

(ii) CD14/LPS Transfer and Presentation to Host Cells
The second potential interaction that may contribute to low reactive LPS is the LPS-LBP-mediated transfer to CD14. After initial LBP-LPS binding, the next step toward host cell activation is the transfer of LBP-LPS complex to either soluble or membrane-bound CD14. Since this occurs through an enzymatic process (Cunningham et al., 1996; Yu and Wright, 1996), comparisons of the ability of different LPS species to interact with CD14 have been normally performed by measurement of the Km as well as the 1/2 Satmax (the concentration of LPS required to obtain half the maximum amount of binding to CD14) (Cunningham et al., 1996). Results from these CD14 transfer comparison experiments, with the use of LPS obtained from E. coli, P. gingivalis, and H. pylori, have revealed significant differences among these bacteria that correlated with the species-specific LPS ability to bind LBP (Cunningham et al., 1996).

An additional site where structural differences in LPS-lipid A composition may be differentially recognized is during the LBP/CD14-dependent presentation of the LPS to the host cell itself. For example, even before the discovery of Toll-like receptors (TLRs), there was evidence that recognition of specific lipid A structural details occurred after CD14 binding (Delude et al., 1995; Kitchens and Munford, 1995). It was demonstrated that inhibition of LPS responses by R. sphaeroides lipid A and synthetic lipid IVA was not due to LPS recognition by CD14 alone (Delude et al., 1995). In this particular study, the authors capitalized on the fact that lipid IVA inhibits human and activates mouse monocyte LPS responses. Here, by transfecting human cells with mouse CD14 and vice versa, they showed that transfectants responded to these LPS analogues according to their cell, and not CD14, type. Furthermore, Akashi et al.(2001) demonstrated that MD-2, a membrane-bound molecule that co-associates with TLR4, selectively conferred responsiveness to lipid A but not to the synthetic precursor lipid IVA, thus influencing the specificity of the TLR and resultant activation pathway. Likewise, moesin, a 78-kDA protein that has recently been identified as another cell-surface molecule on human monocytes, potentially functions as either a TLR4 co-molecule or an independent LPS receptor, capable of transducing CD14-mediated signals itself (Iontcheva et al., 2004).

(iii) Conflicting Roles for Toll-like Receptors (TLRs) in Bacterial LPS Structural Recognition
The discovery of TLRs and their associated co-molecules has generated much excitement in recent years, since it reveals functional cell membrane components that are responsible for host recognition of specifically conserved microbial structures, or ‘PAMPS’ (pathogen-associated molecular patterns) present on invading organisms (Medzhitov and Janeway, 1997). Generating equal excitement is the possibility that this might represent a specific mechanism that can distinguish even subtle structural differences between microbial LPS components, thereby generating a different response to specific bacterial LPS. However, since their discovery, some confusion still remains regarding the exact intracellular pathway, TLR or combination of TLRs, as well as the contributions from associated co-molecules, adhesion molecules (Corbi and Lopez-Rodriguez, 1997; Flaherty et al., 1997; Todd and Petty, 1997; Triantafilou and Triantafilou, 2002), or membrane ion channels (Ishii and Nakae, 1993; Hoang et al., 1997; Blunck et al., 2001) responsible for individual bacterial-mediated host inflammatory activation. For example, early in vitro studies suggested that TLR2 could interact with mCD14—that had previously recognized LBP-bound bacterial LPS (Wright et al., 1990; Ulevitch and Tobias, 1995)—forming an LPS/receptor complex (Yang et al., 1999). Additonally, later studies showed that specific over-expression of TLR2 conferred a much higher LPS responsiveness, via the NF-{kappa}B pathway, with the use of HEK (human embryonic kidney) 293 cells, when compared with the low basal activation seen with the same cells transfected with TLR4 (Kirschning et al., 1998; Yang et al., 1998). However, accumulated evidence from studies with LPS from E. coli and Salmonella suggested that TLR4, and not TLR2, was the Toll receptor responsible for cell activation in the presence of an LPS challenge (Tapping et al., 2000). Additionally, evidence based on positional cloning experiments targeting the Lps gene responsible for C3H/HeJ LPS hyporesponsiveness (Poltorak et al., 1998), TLR4 knock-out mice (Hoshino et al., 1999), or synthetic monosaccharide lipid A analogue preparations (Tamai et al., 2003) allowed researchers to argue that the LPS response observed was, in effect, mediated through TLR4 (Takeuchi et al., 1999). Furthermore, Hirschfeld et al.(2000) revealed that, through added re-purification of LPS preparations, removal of contaminating proteins (which were shown to possess potent bioactivity themselves) eliminated activation signals through both human and murine TLR2. Consequently, conflicting TLR recognition data might have arisen from multiple factors, such as: additional intracellular pathways activated but not yet identified through TLRs, purity of LPS preparations, various amounts or availability of soluble or membrane-bound factors or expression levels of TLRs on the cell lines or primary cells assayed, or the inability of transiently transfected or overexpressed TLR cells in vitro to display accurately the exact role or function displayed by the in vivo, indigenous TLRs (Takeuchi et al., 1999). Therefore, as the numbers of identified TLRs and co-molecules increase, as well as their ability to recognize specific ligands or the biologic effect of TLRs working in combination, exact in vivo TLR function—regarding bacterial recognition and host activation—still remains to be elucidated.


   (4) NATURAL LIPID A HETEROGENEITY
TOP
 ABSTRACT
 (1) INTRODUCTION
 (2) WHAT ARE LIPOPOLYSACCHARIDES...
 (3) HIGH vs. LOW...
 (4) NATURAL LIPID A...
(5.0) PERSPECTIVES
 REFERENCES
 
Biologic assays delineated differences in biologic activity due to structural differences in bacterial lipid A. However, it is now understood that bacteria may contain more than one lipid A structural type, and that environmental conditions can regulate the numbers and types of lipid A species found in a single bacterial population. This section will discuss several examples of how lipid A heterogeneity alters host responses.

(4.1) Natural Yersinia pestis LPS Heterogeneity (Growth Temperature Effects)
One of the most interesting findings in natural lipid A research was the observation that specific bacteria possess the ability to alter or regulate their lipid A form under specific environmental conditions. For example, early bacterial growth and physiology studies indicated that the molecular size of the highly pathogenic Yersinia pestis (the causative agent for bubonic plague) changed when growth temperature shifted from 26 to 37°C (Darveau et al., 1983). In addition, at around the same time, lipid researchers found an increase in the overall amount of an unsaturated fatty acid [hexadecanoic acid (C16:1)] in E. coli when grown at lower temperatures (Van Alphen et al., 1979). This observation led others to propose that specific fatty acid alterations, in relation to environmental temperature changes, might occur in Yersinia pestis (Kawahara et al., 2002), since this same fatty acid (C16:1) was identified within its lipid A structure (Bordet et al., 1977; Dalla Venezia et al., 1985; Aussel et al., 2000). What Kawahara’s group found was that, at 27°C, which is the environmental temperature of its flea vector, Y. pestis displayed lipid A heterogeneity containing a mix of tri-, tetra-, penta-, and hexa-acylated lipid A species. Incubation at 37°C (mammalian host temperature) significantly altered the lipid A species composition, in that the hexa-acylated lipid A species was missing, and the penta-acylated species was significantly reduced, leaving predominantly tri- and tetra-acylated lipid A. Based upon structural analogies with the closely related E. coli lipid A, specific lipid A species modifications, such as predominance of lower acylated lipid A species and down-regulation of higher acylated forms, should result in less potent innate defense activation, consistent with this bacterium’s strategy to evade host defenses. This exact effect was observed in human macrophage cell lines in which activation of TNF-{alpha} was reduced significantly by Yersinia lipid A produced at 37°C, when compared directly with production at 27°C (Kawahara et al., 2002; Rebeil et al., 2004).

(4.2) Environmental Modulation of Two-component Regulatory Systems in Salmonella (PhoP/PhoQ) Results in a Natural Increase of LPS Heterogeneity
Another example of lipid A structure modification in response to environmental conditions comes from research involving two-component regulatory systems within specific bacteria. Salmonellae typhimurium contains a PhoP-PhoQ sensor kinase and transcriptional activator system which regulates genes required for intracellular survival and cationic peptide resistance (Miller et al., 1989). It has been demonstrated that a constitutive S. typhimurium mutant designated PhoPc, which mimics, in part, the intracellular regulation state of the bacteria, contains a modified lipid A species composition (Guo et al., 1997). These modifications resulted in S. typhimurium LPS that contained two additional lipid A species, one modified with an amino-arabinose moiety, and the other containing a 2-OH myristic fatty acid instead of myristic acid (Guo et al., 1997). These additional lipid A species rendered both whole bacteria and isolated LPS less potent activators of E-selectin and TNF-{alpha} when compared with wild-type or another PhoP– mutant (Guo et al., 1997). This was the first description of genetic regulation that resulted in lipid A alterations that modified the innate host response. Another example was found upon examination of the LPS structure found in Pseudomonas aeruginosa strains grown under different cultural conditions and isolated from the lungs of individuals with cystic fibrosis (CF) (Ernst et al., 1999). In this study, it was found that the lipid A species of P. aeruginosa was altered by magnesium concentration in the culture medium under control of a phoP regulator, such that lipid A species now contained an additional palmitic acid as well as amino-arabinose. In this situation, both whole bacteria and isolated LPS preparations were more potent activators of E-selectin and TNF-{alpha} (Ernst et al., 1999). Strains of P. aeruginosa, isolated from the lungs from patients with CF, contained these same modifications and also were demonstrated to be more potent activators of IL-8 production from human endothelial cells (Ernst et al., 1999). Recently, these researchers, utilizing chimeric human or mouse TLR4 proteins, were able to identify the exact site (an 82-amino-acid region within the hypervariable, extracellular domain of human TLR4) that was able to discriminate between these penta- or hexa-acylated LPS forms from P. aeruginosa and alter host response, indicating a direct link between species-specific differences (e.g., mouse vs. human) in bacterial modification recognition and modulation of the innate host response (Hajjar et al., 2002).

(4.3) Outer Membrane Enzymes Affect Lipid A Heterogeneity
In Salmonella typhimurium, once the two-component regulatory system PhoP/PhoQ senses environmental changes, it modulates its lipid A structure, as described above, through the use of two recently identified outer membrane enzymes. PagP, a lipid A palmitoyltransferase, was the first identified example of an outer membrane enzyme under functional PhoP/PhoQ control that was capable of biosynthesizing these hexa-acylated lipid A forms with resistance to cationic anti-microbial peptides (Bishop et al., 2000). Since this discovery, a new enzyme, PagL, determined to be located mainly on the outer membrane, has been identified (Trent et al., 2001). Functional PagL expression was determined to involve deacylase activity, selectively cleaving R-3 hydroxymyristate that may be attached to the C3 position of certain lipid A precursors (Trent et al., 2001). The specific functional ramification of PagL is currently unknown, but it has been suggested that this lipid A alteration might modify host cytokine responses in Salmonella-type infections (Trent et al., 2001).

(4.4) Limited vs. ‘Substantial’ Lipid A Heterogeneity in P. gingivalis
With regard to oral bacteria, one of the most extensively studied lipid A structures comes from Porphyromonas gingivalis. Initially, early studies suggested that P. gingivalis possessed limited lipid A structural heterogeneity, with LPS preparations consisting of only a tri-acylated monophosphorylated form with a negative ion mass of m/z 1195 (Ogawa, 1993). Later, a second study reported multiple P. gingivalis lipid A structural isoforms; however, in this particular study, certain forms predominated, including tetra- and penta-acylated monophosphorylated lipid A’s with molecular mass ions of m/z 1435, m/z 1449, and m/z 1690, respectively (Kumada et al., 1995). These structures are depicted in Fig. 3Go and differ from the canonical E. coli lipid A structure in the number of phosphates, and the numbers, types, and positions of the fatty acid chains. The reasons for the apparent discrepancies in the predominant type of P. gingivalis lipid A found between these two studies were not clear at the time of their publication. Variations in the strain of P. gingivalis used, growth conditions, and isolation procedures were considered as possible mechanisms contributing to the description of different lipid A species. Recently, however, a study from our laboratory has demonstrated that at least three of these P. gingivalis lipid A species (1195, 1435, and 1450) (Bainbridge et al., 2002), as well as others (Darveau et al., 2004), are present in our purified LPS preparations (obtained from multiple laboratory as well as clinical isolates), demonstrating that P. gingivalis, similar to other bacteria (Caroff et al., 2002), contains multiple lipid A species. Additional studies with either phenol and guanidine thiocyanate (Yi and Hackett, 2000) or cold MgCl2 (Darveau and Hancock, 1983) techniques to extract LPS, as well as different lipid A hydrolysis procedures (Caroff et al., 1988; Ogawa et al., 2002), have also found that P. gingivalis contains multiple lipid A moieties, further confirming the presence of heterogeneity within lipid A preparations. These observations, obtained from different laboratories, suggest that the increased heterogeneity within LPS preparations from P. gingivalis might not be a result of specific extraction techniques alone, and efforts have been initiated to determine the in vitro and/or in vivo mechanisms responsible for the observed diversity. Based on structure-activity relationships, discussed previously, it is predicted that these lipid A modifications will elicit specific alterations in innate host response, with potential pathogenic significance.


Figure 3
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  		Figure 3. Heterogeneity of P. gingivalis lipid A may contribute to innate host response modulation. The basic structure of lipid A for P. gingivalis 381 was described by Ogawa (1993) as a monophosphorylated tri-acylated disaccharide with a negative ion FAB MS-MS mass ion located at m/z 1195, which displayed low endotoxic activity. Later, Kumada et al.(1995) reported additional P. gingivalis lipid A moieties (within a clinical isolate) to include lipid A species containing 4 or 5 fatty acid chains, with a negative ion FAB MS-MS mass ion(s) located at m/z 1435, 1449, 1690, and 1770, respectively. The addition or subtraction of specific lipid A components from the parent moiety (m/z 1690 in this example) can be accomplished naturally through enzymatic function. Membrane-associated enzymes (depicted by arrows) that possess this capability, whose function is environmentally or gene-regulated (Bishop et al., 2000; Trent et al., 2001; Kawasaki et al., 2004), might help explain natural heterogeneity within lipid A of P. gingivalis. Characterization of predicted innate host response effects toward these structural differences is currently under way (Ogawa and Uchida, 1996; Ogawa et al., 2002; Darveau et al., 2004).

 
(4.5) Heterogeneity and LPS-complex Co-molecules: Their Effects on Structural Recognition via TLRs
An important observation, one that might help clarify the discrepancy between differing roles of TLRs in bacterial recognition, comes from the requirement of specific co-factors that, by their presence or absence, help regulate specific TLR utilization. Using the well-characterized innate host recognition of E. coli LPS as an example, investigators have determined that a series of essential binding and transfer events must occur between LPS and host cells which require the presence of LPB, CD14, and the MD-2/TLR4 complex (Schumann et al., 1990; Wright et al., 1990). Here, as stated earlier, initial recognition of E. coli LPS—once bound by LBP—was shown to occur through interactions with the host via presentation of [LBP-LPS] to one of two forms of CD14: membrane-bound CD14 (mCD14), which is located on the cell membrane of certain host cells; or a soluble serum form (sCD14) present in serum exudates, such as gingival crevicular fluid (Pugin et al., 1993a; Jin and Darveau, 2001). Next, this interaction (between the LPS-LBP coupled with either mCD14 or sCD14) is transferred to a cell-associated, MD-2-bound TLR4 complex (Poltorak et al., 2000; Akashi et al., 2001; Viriyakosol et al., 2001; Visintin et al., 2003; Saitoh et al., 2004), triggering the inflammatory host cell activation pathways. Later reports, from genetic and evolutionary studies, suggested that LPS signal activation was mediated solely through TLR4 (Beutler et al., 2001), and that previously observed activation through TLR2 might be due to different LPS extraction procedures, sometimes resulting in residual, bioactive protein contamination within LPS preparations themselves (Hirschfeld et al., 2000). However, recent evidence might reveal that actual heterogeneity of specific structural features within highly purified LPS samples might, in fact, modulate activation of individual TLRS like TLR2/TLR4 and/or combinations of TLRs and their associated co-molecules. Modification of Salmonella typhimurium, either by the addition of acyl chains (PagP-induced palmitoylation) or subtraction of acyl chains (PagL 3-O deacylation), resulted in remarkably reduced activation through TLR4 (Kawasaki et al., 2004). In fact, these modifications reduced TLR4-driven NF-{kappa}B activation by 30- to 100-fold, when compared with the unaltered lipid A samples. Furthermore, Darveau et al.(2004), using highly purified P. gingivalis lipid A preparations, showed that extracted preparations enriched in tetra-acylated lipid A forms (m/z 1435/1450) were capable of activation through both human and mouse TLR4, TLR2+TLR1, and even TLR2, when transiently transfected HEK 293 cells co-expressing mCD14 were used. In addition, it was determined from these experiments that MD-2 was required for the functional engagement of TLR4, but not TLR2+TLR1 or TLR2. However, it was also reported that MD-2, which in humans and mice has also been suggested to play a role in species-specific recognition of lipid IVA and taxol (a plant-derived molecule), was not responsible for the observed P. aeruginosa LPS differential recognition in either clinical or laboratory samples, thereby confirming the importance of a newly identified region within human TLR4 in bacterial LPS recognition (Hajjar et al., 2002). Therefore, it is becoming increasingly clear that in vitro detection of subtle changes in LPS structure, ones that are most likely seen in vivo by the host, require the presence of multiple factors (TLRs) and co-factors (LBP, mCD14/sCD14, MD-2, etc.), thus complicating the extrapolation of data from different assay systems as well as different bacterial species and/or preparations. However, it has been proposed that in vivo biologic effects for down-regulation or differential regulation of TLRs, due to heterogeneity in lipid A from bacteria such as Salmonella typhimurium or P. gingivalis, could potentially assist these known pathogens in adaptation to changing host environments, colonization persistence, or pathogenesis (Darveau et al., 2004; Kawasaki et al., 2004).


   (5.0) PERSPECTIVES
TOP
 ABSTRACT
 (1) INTRODUCTION
 (2) WHAT ARE LIPOPOLYSACCHARIDES...
 (3) HIGH vs. LOW...
 (4) NATURAL LIPID A...
 (5.0) PERSPECTIVES
REFERENCES
 
The mechanism by which the innate host defense system detects LPS, specifically the lipid A component, has been fairly well-elucidated over the past 10 years. Lipid A recognition and subsequent innate host responses occur through a series of binding and transfer reactions among different germ-line-encoded host response proteins. This field has been driven mainly by early observations that this microbial component, as obtained from enteric bacteria, elicits a highly potent, often lethal, inflammatory response. However, early studies also identified LPS species (mainly from Gram-negative anaerobic bacteria) that differ structurally from enteric lipid A and did not elicit potent inflammatory responses. Structural differences in lipid A types have been shown to alter the specific binding affinity and transfer properties of the lipid A species which may correlate to reduced inflammatory responses. More recent work has shown that different lipid A structural types may be present in a single bacterial population, and that environmental conditions may alter the types of lipid A molecules present (Fig. 4Go).


Figure 4
Figure 4
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  		Figure 4. Bacterial modification of lipid A leads to changes in innate host response. Panel (A) LPS consist of three distinct regions formerly thought to contain a highly variable outer O-chain, a ‘semi-conserved’ middle core segment, as well as a highly conserved lipid A anchor. However, as more examples of natural heterogeneity become apparent, the once ‘highly conserved’ lipid A segment is being recognized as a site in which bacterial modification can result in multiple forms or moieties of lipid A structure, each with a potentially distinct effect on the innate host response. Examples of natural heterogeneity within lipid A and innate host effects are represented in panel (B). Here, heterogeneity can arise from: temperature changes from flea vector to host, as seen in Yersinia pestis (Kawahara et al., 2002; Rebeil et al., 2004); by PhoP/PhoQ-controlled mechanisms in S. typhimurium and P. aeruginosa (Guo et al., 1997; Ernst et al., 1999; Bishop et al., 2000; Trent et al., 2001); as well as specific environmental conditions and membrane-associated enzymes currently being examined in P. gingivalis. Specific host responses toward these lipid A modifications are described in the text.

 
We are learning that, although lipid A was once thought to be an invariant component of LPS, in some bacterial species it is not. In fact, the structural alterations that occur due to different environmental conditions may have profound effects on the innate host response. It is not surprising, therefore, that bacteria have ‘evolved’ mechanisms to manipulate the ‘pattern recognition’ motif used by the innate host defense system to delay, evade, or eliminate host recognition under certain conditions.

Received for publication August 20, 2004. Accepted for publication February 10, 2005.


   REFERENCES
TOP
 ABSTRACT
 (1) INTRODUCTION
 (2) WHAT ARE LIPOPOLYSACCHARIDES...
 (3) HIGH vs. LOW...
 (4) NATURAL LIPID A...
 (5.0) PERSPECTIVES
 REFERENCES
 

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Journal of Dental Research, Vol. 84, No. 7, 584-595 (2005)
DOI: 10.1177/154405910508400702
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