Open access peer-reviewed chapter

Detection Methods for Lipopolysaccharides: Past and Present

Written By

Loreen R. Stromberg, Heather M. Mendez and Harshini Mukundan

Submitted: 29 April 2016 Reviewed: 03 March 2017 Published: 12 July 2017

DOI: 10.5772/intechopen.68311

From the Edited Volume

Escherichia coli - Recent Advances on Physiology, Pathogenesis and Biotechnological Applications

Edited by Amidou Samie

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Abstract

Lipopolysaccharide (LPS) is the primary component of the outer membrane of Gram‐negativebacteria. LPS aids in protecting bacterial cells, and also defines the unique serogroups used to classify bacteria. Additionally, LPS is an endotoxin and the primary stimulator of innate immune cells in mammals, making it an ideal candidate for early detection of pathogens. However, the majority of methods for detection of LPS focus on detection of the endotoxic component of the molecule, lipid A. Since lipid A is largely conserved among bacterial species and serogroups, these detection approaches are highly nonspecific. Thus, the importance of identifying the O‐polysaccharide antigenic portion of LPS, which confers serogroup specificity, has received a great deal of attention in recent years. However, methods that are highly selective to the O‐antigens are typically less sensitive than those that target the endotoxin. Here we present a history and comparison of the sensitivity of these methods and their value for detecting bacteria in a variety of different sample types.

Keywords

  • lipopolysaccharide
  • endotoxin
  • O‐antigen
  • Serogroup
  • biodetection

1. Introduction

The increasing occurrence of infectious disease is a global issue. Emerging pathogens with increasing levels of drug resistance are a continuing danger to both public health and agriculture. Accurate and rapid detection of pathogens is critical to implement preventative measures to mitigate this problem. Despite this urgent need, conventional methods for bacterial detection require cell culture and serology, which can take several weeks. As new pathogens emerge, it is even more important that our detection technologies evolve to keep pace with the need to discriminate pathogen from host flora. This requires an understanding of pathogen biology, the types of samples they occur in, and their mechanism of immune interaction within the hosts [1].

The innate immune system is able to discriminate pathogens from nonpathogens, and rapidly sense pathogen biomarkers in the complex milieu of the host. Exploiting this recognition via measurement of pathogen signatures, can provide an optimal strategy for discriminatory biodetection. A primary category of such biomarkers is virulence signatures termed pathogen‐associated molecular patterns (PAMPs) [2]. PAMPs are evolutionarily conserved molecules that bind pattern‐recognition receptors in the host, and activate the innate immune response [2, 3], providing a means for both early and specific pathogen detection. Biochemically, PAMPs are a diverse array of proteins, lipopeptides, lipoglycans, peptidoglycans, teichoic acids, and nucleic acids [4]. However, many detection methods have largely focused on proteins and nucleic acids [1, 5], ignoring other categories of PAMPs [2, 68]. Also, their small size, biochemistry, and low concentration in hosts make them difficult to target in detection assays [8, 9].

Classified as a lipogylcan, lipopolysaccharides (LPS) are small amphiphilic molecules that are associated with Gram‐negative bacteria [7, 10]. LPS is an indicator of active infection, is serogroup‐specific [1113], more stable than its protein counterparts, and is released early in infection, making it an ideal candidate for detection and diagnostics. LPS serves as a biomarker that aids in serological discrimination of Gram‐negative bacteria; this allows for identification and characterization of pathotypes that are essential for timely mitigation and treatment of infections. Since LPS is a pathogen‐specific biomarker, it is an indicator of acute infection, which is an advantage over serological assays. In addition to medical diagnostics, LPS detection provides a method for detecting Escherichiacoli in the food‐industry, which is often associated with food‐borne illnesses. Finally, LPS is also a virulence factor whose structure and function determines E. coli serogroup, a factor which has ramifications on vaccine design and therapeutic interventions. While many methods for LPS detection exist, most of them are not optimized for amphiphilic detection in physiological samples. An ideal measurement for LPS should be sensitive enough to detect low concentrations of the amphiphile in aqueous physiological milieu (e.g., blood), and use antibodies or ligands that provide serogroup selectivity [14]. Coupling sensitive detection platforms with surfaces designed to maximize the binding of amphiphilic PAMPs is a potential solution to achieve such an ideal.

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2. Sources of lipopolysaccharides

Bacteria are classified into Gram‐negative and Gram‐positive [15], which release amphiphilic virulence factors such as LPS, lipoarabinomannan (LAM), and lipoteichoic acid (LTA) in the host. Species of pathogenic Gram‐negative bacteria of concern to human health, include Acinetobacter [16], Burkholderia [17], Bordetella [18], Campylobacter [1921], Chlamydia [22, 23],E. coli [20, 24], Helicobacter [25, 26], Hemophilius [27], Klebsiella [28], Legionella [20, 29], Moraxella [30], Neisseria [31], Pseudomonas [32], Proteus [33], Salmonella [20, 34], Shigella [35], Yersinia [36], and others, grouped into the Enterobacteriaceae family. These pathogens are contaminants in food, water, and soil, used as agents of bioterrorism, and can cause nosocomial infections [5]. Detection of these organisms, particularly E. coli, is an important aspect for epidemiology, disease control, and treatment.

Herein, we present a comprehensive description of the structural and biochemical properties of LPS, current methods for its detection, and potential approaches to overcome the current limitations for direct detection of the molecule in physiological matrices.

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3. Lipopolysaccharide structures and conformations

Lipopolysaccharides have been the subject of intense study for over half a century [3739]. LPS is the prototypical lipoglycan with an overall net negative charge [4042], and is the primary component of the outer membrane of nearly all Gram‐negative bacteria [11]. The bacterial membrane of each E. coli cell is composed of approximately 106 lipid A moieties and 107 glycerophospholipid molecules, comprising approximately three‐quarters of the outer membrane [4345]. Thus, there are approximately 62 pg of LPS per cell (for E. coli in log phase growth) [46]. LPS has an amphipathic tripartite structure (Figure 1). Lipid A is the most conserved portion of the LPS molecule, and consists of six, sometimes seven, fatty acid tails(E. coli and Salmonella, respectively), which gives the molecule its hydrophobic properties [10, 43, 45]. Lipid A is also called endotoxin [43], and is responsible for the biological effects of LPS caused by its binding to the mammalian innate immune receptor, toll‐like receptor 4 (TLR4) [11, 44, 47, 48]. Structurally, lipid A is covalently bound to the core polysaccharide, which is further divided into the inner and outer core polysaccharides, with the outer core being less conserved in both sugar moieties and location of glycosidic linkages [45, 49, 50].

Figure 1.

Representative structure of the molecular components of smooth LPS. The hypervariable O‐polysaccharide antigen, core polysaccharide, and the hydrophobic lipid A group. Reprinted with permission from Ref. [74].

There are two main forms of LPS—smooth (S‐form) and rough (R‐form) [42, 45, 46]. The distal end of LPS extends to a long chain O‐polysaccharide antigen (O‐ag(s)) in organisms possessing S‐form, which is an indicator of virulence [51, 52]. R‐form LPS is devoid of the O‐ag [45], but can still induce an immunogenic response [53]. The O‐ag is hyper‐variable, and made up of repeating subunits, each composed of 1–7 glycosyl residues [45, 54]. As many as 40 size variations in subunit repeats of the O‐ag have been reported just for E. coli O111:B4 [55], and 180 O‐ag have been identified overall for E. coli species [47, 54]. The sugars (colitose, paratose, tyvelose, and abequose) that make the O‐ag unique are seldom found elsewhere [54]. Other variations to the polysaccharide are implemented through addition of noncarbohydrate entities, such as acetyl or methyl groups [54]. These variations make discriminative detection of enteric bacteria feasible [56], but complicate antigen characterization. Therefore, LPS serves as an ideal target for early detection and identification of Gram‐negative pathogens.

In aqueous solutions, amphiphiles like LPS can present in a micellar conformation [48, 55, 5759]. This occurs at a concentration specific to the amphiphile [55], and is known as the critical micelle concentration (CMC). At or above the CMC, there is an equilibrium state between monomers, micelles or supramolecular aggregates, depending on environmental conditions [48, 5557, 6063]. This amphiphilic biochemistry and structural variability complicates determination of the exact molecular weight of S‐form LPS. As such, LPS concentrations are reported in weight per volume, or in endotoxin units (EU), a measure of activity. As degree of endotoxicity can vary according to bacterial origin, a rough estimate of 100 pg = 1 EU is used in many cases to facilitate unit conversion [64, 65].

The large oligosaccharide region on S‐form LPS makes the molecule amphipathic [54], which influences the shape of micelles in solution. Lipid A is largely responsible for shaping the LPS micelle [10, 45, 46, 56, 6668], although other factors can also contribute. Lipid A is conserved within species in the number of fatty acid chains and the degree of saturation [44, 66] within those chains [22, 47, 69]. Shapes for LPS micelles include cubic, lamellar, and hexagonal inverted structures [56, 67, 70, 71]. Whether aggregate or monomeric forms (or both) of LPS is required for innate immune activation is debatable [56, 72, 73]. Since this process occurs in aqueous blood, it is unlikely that the molecule is presented as a monomer, unless associated with serum binding proteins.

Variation in LPS micelles [55] modifies presentation of O‐ag‐specific epitopes to antibodies, making detection challenging [74, 75]. This is specifically true when the heterogeneous presentation of linear [76] and conformational epitopes [49, 77] present on LPS molecules are considered. The primary structure of LPS varies in the core polysaccharide, within and between species [47, 55]. Core polysaccharides are primarily made up of common sugars such as heptose and 2‐deoxy‐dmannooctulosonic acid (a.k.a. KDO), which can be functionalized with phosphate or ethanolamine groups [45, 50, 78]. This feature contributes to varying charge distributions and differential size ratio of the hydrophobic to hydrophilic regions which influences micelle assembly [10, 59, 79, 80]. Other factors that contribute to micelle shape [10, 79] are pH [61], ion concentration [8186], and temperature [62]. These biochemical properties drive host‐pathogen interactions and should be considered in the design of detection strategies.

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4. Detection methods for lipopolysaccharides and similar amphiphiles

There have been many efforts to establish rapid and reliable detection methods for LPS in clinical samples [10, 46] and for testing pharmacological products such as infusion fluids, sterile injectables, medical device implants, and others [87]. These methods can be broadly divided into six overlapping categories: invivo and invitro tests, immunoassays and their derivatives; biological, chemical, and cell‐based sensors. These methods span a broad range of sensitivity, but many lack the ability to differentiate between LPS serogroups.

4.1. Limulus amoebocyte lysate assay and the rabbit pyrogen test

The first method approved by the US Food and Drug Administration for LPS detection was called the rabbit pyrogen test [8890], which simply measures the ability of an endotoxin to induce fever in an animal. Any febrile response was attributed to the presence of endotoxin [8991]. The test, clearly, is activity‐based, and nonspecific. In the case of Hepatitis B vaccine manufacturing, the rabbit pyrogen test is still the standard method for determining endotoxin contamination [91], but the test is cost prohibitive and is minimally utilized today, except in some parenteral devices [10].

In 1956, Bang discovered that amoebocytes from Limuluspolyphemus (a.k.a. horseshoe crab) agglutinate upon addition of endotoxin [46], as a result of a protease cascade [10]. Bang and Levin [46, 92] subsequently used this concept to devise a method for endotoxin detection. Since the lysates of amoebocytes were required, it was called the limulus amoebocyte lysate (LAL) assay, and is the gold standard for the detection of lipid A. The LAL assay is prone to variability and can be inhibited through several mechanisms. The United States Pharmacopeia and the Code of Federal Regulations have consequently published guidances for the manufacturing and testing of assays for use on human products [93, 94]. Despite some challenges, the LAL assay is more rapid, cost effective, and reportedly 300 times more sensitive [46] than the rabbit pyrogen test [46].

Variants of the LAL assay use turbidimetric [95], chromogenic [46], or viscosity [10] measurements to determine results [10, 46]. A turbidimetric gel clot has more coagulen, and measures the change in turbidity over time, but does not form a solid clot [46, 95]. The viscosity assay, however, measures the degree of clotting via the change in viscosity. The chromogenic assay can be endpoint or kinetic, and utilizes a p‐nitroaniline substrate, which is cleaved by an LAL proenzyme, providing a colorimetric readout [46]. The sensitivity of LAL assays is dependent on the sample type, processing method and time, as well as the dilution factor [46]. Additionally, the source of the LAL reagent plays a factor, as it is apparent when comparing the different limits of detection (LoD) reported for endotoxin standards. A survey of the relative sensitivities of the LAL assay is shown in Table 1.

DescriptionSampleDetection methodSpeciesSensitivity (ng/mL)*SpecificSource
Rabbit pyrogenPurified endotoxinFebrile responseNo[89]
LALPlasmaGelationMultiple species0.5–5.0No[96]
LALBlood, plasmaGelationE. coli0.5–5.0No[92]
LALSerum plasmaOptical densityE. coli0.025–0.5No[100]
LALUrineGelationE. coli0.5No[204]
LALUrineOptical densityMultiple species2.0No[98]
LALSpinal fluid/ plasmaOptical densityE. coli, Haemophilus influenzae B0.1No[101]
LALAscitesGelationE. coli0.5No[104]
LALCerebral/synovialE. coli1.0No[103]
LALSeawaterOptical densityE. coli2.3No[41]
LALPurified endotoxinGelationE. coli1.0No[95]
LALPurified endotoxinGelationSalmonella minnesota10−11No[111]
LALGround beefGelationEnterobacter aerogenesNo[205]
LALGround beefGelationMultiple species51.0 ng/gNo[108]
LALMilkChromogenicPseudomonas putida0.01No[107]
LALPurified endotoxinGelationE. coli O114100No[206]
LAL‐magnetoelastic sensorPurified LPSresonant frequencyE. coli O111:B40.0105 EU/mLNo[207]
ENDOLisa® (LAL)Purified endotoxinFluorescenceE. coli spp., Salmonella spp.0.05–500 EU/mLNo[129]
ELISAMilkAbs at 405 nmE. coli100–200[165]
LPS pull down‐sandwich ELISAPure culturesAbs at 450 nmE. coli O157[125]
LPS pull down‐sandwich ELISAPurified LPSAbs at 450 nmSalmonella spp. (31 total)1.0Yes[126]
Premier EIA E. coli O157Stool extractSpectro‐photometricE. coli O157Yes[122]
LPS pull downPurified endotoxinRIAE. coli O114300No[206]
LPS pull down‐ion (NTA‐Cu)Purified LPSEISE. coli O55:B50.0001–0.1No[161]
Diaphorase functionalized surfacePurified LPSChemicalE. coli O127:B850Maybe[87]
LPS pull down‐SAMs with synthetic peptidePurified LPSElectro‐chemistryE. coli ATCC 3521821.8 pg/mLNo[189]
LPS pull down‐SAMs with aptamerPurified LPSEISE. coli O55:B50.1–1.0Maybe[159]
LPS pull down‐gold electrode w/ aptamerPurified LPSEIS and cyclic voltammetryE. coli O55:B50.001–1.0No[160]
LPS aptamer sandwichPurified LPSElectro‐chemistry10 fg/mLMaybe[188]
LPS pull down‐gold electrodes w/ PmBPurified LPSEISE. coli O111:B40.2No[162]
Polydiacetylene liposomesPurified LPS (5 groups)Change in AbsE. coli spp, Salmonella spp2.22 mg/mLYes[191]
Impedance enthothelial biosensorPurified LPS in culture medium+Resistivity of cell monolayer500No[169]
Macrophage microarrays on gold electrodesPurified LPS in culture mediumFTIRE. coli O111:B40.1 µg/mLNo[200]
Primary culture HDME cellsPurified LPSFluorescenceE. coli O111:B41.0 µg/mLNo[171]
Engineered cells secrete alkaline phosphatasePurified LPS in culture medium+Electro‐chemistry0.1No[170]
LPS pull down‐PmBPurified LPSEvanescent sensingE. coli O128:B1225No[75]
LPS pull down‐TLR4/MD2 on gold electrodesPurified LPSElectro‐chemistryE. coli O55:B50.0005 EU/mLNo[180]
LPS pull down‐membrane insertionPurified LPS (3 groups)Evanescent sensingE. coli420Yes[74]
LPS pull down‐antibodyPure cultures in ground beefEvanescent sensingE. coli O157Yes[166]
LPS pull down‐proanthocyanidinFITC‐labeled LPSFluorescenceE. coli O55:B5No[192]
Copolythiophene interacts with LPSPurified LPSFluorescenceE. coli O55:B52.5E−5–2.0 µMNo[164]
Polydiacetylene liposomesFluorescence0.1 µMNo[190]
Peptide‐based fluorescencePurified LPSFRET‐increaseE. coli O111:B40.15–2.0 µMNo[194]
Pyrenyl‐derived long‐chain quaternary ammonium probePurified LPSFluorescenceE. coli O55:B5100 nMNo[193]
LPS pull down‐peptide on Graphene OxidePurified LPS (4 groups)FluorescenceSeveral species130 pMNo[195]
LPS pull down‐PmB capturePurified LPS spiked in bloodAcoustic sensingE. coli O55:B51.0No[196]
LPS pull down‐CD14 captureBiotin‐LPSLuminescence10.0No[176]
LPS pull down Polyaniline + ConA lectinPurified LPS and LTAEISE. coli,S. aureus50.0No[208]
Aptamer sandwich on beadsPurified LPSFluorescenceE. coli O55:B50.01Maybe[163]
LPS pull down endotoxin neutralizing proteinPurified LPSCapacitanceE. coli10−13 MNo[181]
LPS pull down CramoLL lectinPurified LPS (4 types)EISE. coli Salmonella Klebsiella Serratia25.0 µg/mLNo[185]

Table 1.

Overview of sensitivities and specificities for LPS detection methods.

*Unless otherwise indicated.

In 1970, Levin discovered that samples tested in whole blood would not render a positive result [92], but if plasma was extracted in chloroform and diluted 1–10%, then endotoxin activity could be detected in the 0.5–5 ng/mL range [92, 96]. Levin correctly assumed that components of whole blood were bound to endotoxin, thereby inhibiting the reaction with the LAL reagent [46, 92, 97], or changing the reaction kinetics [46]. This is evident when the amphiphilic nature of LPS and the aqueous nature of blood are considered. In addition to blood and plasma [46, 92, 96], the LAL assay has been used in urine [46, 98], cerebral spinal fluid, synovial fluid, ascites fluid, vaginal and cervical fluids, broncho‐alveolar lavage samples, seawater [46], bovine milk [99], and beef tissue [100, 101]. Virtually all of these have reported ng/mL LoDs, for endotoxin, but none are serogroup‐specific. Researchers have used heat [46, 102], chemical treatment with chloroform [103], acids [104, 105], alkali [106, 107], or ether [108] to improve sensitivity with some success when using heat or chemical extraction of the endotoxin [46, 109]. However, the results show poor reproducibility between researchers (Table 1). Yin and Galanos [106] reported a sensitivity of 10−11 ng/mL for Salmonella spp., while Cooper et al. [89] reported 1.0 ng/mL for E. coli endotoxin. This disparity leaves a lot of questions and draws attention to the fact that small changes in preparation, heat or chemical treatments, usage of plastics instead of silanized glass, or addition of surfactants can result in altered assay sensitivity. This variation can also be explained by the variable biosynthesis of lipid A, as shown with Salmonella [47, 110]. Additionally, LAL can yield false positives upon reacting with other polysaccharides or β‐(1,3)‐glucans [10, 46] and depends on the source of bacteria, as LPS/endotoxin can vary in toxicity [10, 56, 79, 68], in regards to immune stimulation [41, 46, 111].

Thus, the LAL assay and rabbit pyrogen test, both based on the native immune responses of the horseshoe crab or rabbit, exhibit significant variability in outcomes. Despite these, the LAL is still very useful for quickly detecting contamination. For example, in 1981, Jay [101] used the LAL test to determine both microbial counts and endotoxin load in 153 samples of store bought ground beef with a mean sensitivity of 7.9 µg/mL (endotoxin/beef sample) in 1 h. In 1985, Nachum and Shanbrom [46] used a chromogenic LAL system to detect between 2 and 175 ng/mL of endotoxin in 324 patient urine samples, with the assay taking between 2 and 4 h. Timely detection is valuable to both patient care and product viability. Despite being an ideal test for the presence of endotoxin, determining identity of pathogens still requires culture or enrichment.

4.2. Immunoassays for LPS detection and antibody selection

Developed in 1971 [112, 113], the enzyme‐linked immunosorbent assays (ELISAs) are based on the immune reaction between antigen and antibody, with each assay being tailored for the unique antigen being tested. ELISAs were evaluated for Salmonella O‐ags very early in development [114]. However, ELISAs for lipoglycans such as LPS suffer from low sensitivity and reproducibility [115117]. One of the primary reasons for sensitivity issues is the amphipathic biochemistry of these molecules, leading to inconsistent binding on ELISA plates [118], and variable conformations of epitope binding sites [12, 119].

There exist two primary types of LPS‐ELISAs, which detect either the LPS antigen, or LPS antibody titers. With the former, the plate surface is typically coated with a primary capture antibody specific to LPS, or with the sample to be tested [118]. After antigen capture, an epitope‐specific antibody is used to detect LPS. The detection antibody can be directly labeled with an enzyme [113] or secondary antibody for colorimetric detection [120, 121]. In 1998, Mackenzie et al. [122] reported on the effectiveness of a commercial assay to screen stool samples forE. coli O157 antigens, and found that re‐testing samples provided inconsistent results. It was speculated that this was due to inefficient washing of the microwells, yet the amphiphilic antigen preparation and its presentation to antibodies could have contributed to assay inconsistencies. It was also not considered that LPS is notorious for nonspecific and inconsistent binding on microplates [10, 116, 118]. In clinical samples, the association of LPS with host carrier molecules may affect its ability to adhere to capture surfaces [123, 124], as proteins will preferentially bind to the plate. Some groups have also reported cross reactivity or false positives with LPS sandwich ELISAs [125, 126]. Choi et al. [126] developed a sensitive capture ELISA with 24 species of Salmonella, but cross reactivity was observed. To mitigate cross reactivity, attempts have been made to substitute antibodies with other ligands. Grallert et al. [127, 128] coated microplates with proteins isolated from bacteriophages, which are specific to core polysaccharides in order to capture LPS, followed by detection with Factor C (a component of the LAL assay). This sandwich ENDOLisa®, a microplate assay for direct detection of endotoxin, reports sensitivity between 0.05 and 500 EU/mL. This technology is sold as the Endotoxin Sample Preparation (ESP™) kit, and is one of the few kits available for direct detection of endotoxin in blood or serum [129]. However, the assay is unable to differentiate between serogroups.

The second type of ELISA measures LPS antibody titers to screen for Gram‐negative bacterial infections. Here, the surface of the plate is functionalized with the antigen to pull down antibodies (Immunglobulins A, G, and M (IgG, IgA, IgM)) from serum. Since this method is based on adaptive immunity, there is a lag between initial exposure to the pathogen, and increased antibody titers [130], making early detection difficult. This assay is not specific for active infection, but has been used to monitor population health and track epidemiology of infections. Screening has been used to detect exposure of military personnel to Shigella [131], obstetric patients with Chlamydia spp. [132], patients with Salmonella [133], and other pathogens [114, 134137]. Suthienkul et al. [136] used an indirect ELISA to passively adsorb LPS onto polystyrene plates, and measure associated IgG/IgM titers in cholera patients. The results indicated discrepancies between the titers of IgG and IgM in young versus older patients, which could either be due to the inconsistency of LPS coating or associated with cross reactivity [138]. Suthienkul also acknowledged that antibody levels in infants screened could be inherited from the mother [136, 139].

Functionalizing ELISA plates with amphiphilic LPS is a technical challenge [12], since the surfaces are optimized for protein binding. In the late 1970s, it was discovered that polymyxin B (an antibiotic, PmB) interacted with LPS monomers in a 1:1 ratio [86, 140], and can be used to functionalize surfaces for Gram‐negative detection [119]. However, PmB recognizes the conserved lipid A group of LPS, and does not allow for discriminative detection. Takahashi et al. [118] showed that precoating the plate with high molecular weight poly‐l‐lysine increases surface adsorption and allows for detection of 1 µg/mL LPS, with no cross reactivity. Others have studied the effects of ions such as calcium and magnesium [141], trichloroacetic acid [142], mixing the antigen in chloroform/ethanol, and drying on the plate surface [135], or complexing LPS with a protein such as bovine serum albumin [143] to improve performance and reproducibility. Functionalization of ELISA plates with proteins known to bind LPS, such as high‐ or low‐density lipoproteins (HDL, LDL), chylomicrons, and LPS‐binding protein (LBP) have also been evaluated [123, 124] and offers promise for the reliable detection of LPS antigen in complex samples.

Other limitations for LPS detection include the fact that many LPS antigens have not been isolated [144] and thus are not available for the development of screening assays, limiting accessibility of specific antibodies as well [145150]. However, there is also a need to refine methods for selection of tailored antibodies. While there are variations [10], ELISA plates are typically functionalized with whole dead bacteria to screen monoclonal antibody cultures [145, 146, 148], giving rise to potentially cross reactive clones [10, 144] that are then screened against a multitude of bacterial strains [146, 149, 150]. It is noted that it is impossible to screen clones against all epitopes of LPS, even amongst the many E. coli serotypes. In 2000, Jauho et al. [12] addressed this issue by covalently linking purified LPS O‐ags to polystyrene ELISA plates using anthraquinone and UV irradiation. This technique could prove useful in developing serogroup‐specific antibodies against LPS, as conserved antigens like lipid A and core polysaccharide are absent. Alternative methods for antibody screening have utilized immunoblotting [144, 149, 151153] and flow cytometry [154156]. In addition, ELISAs can suffer from high background due to nonspecific interactions limiting their sensitivity [10, 122, 135, 157]. Particularly, endogenous endotoxin present in reagents, on glassware, or plastics [158], may contribute to false positive results. Factors identified above have to be carefully considered in the development of ELISAs targeting LPS.

4.3. Biological and chemical‐based LPS sensing

Many advanced methods such as electrochemical impedance spectroscopy (EIS) [159161], antimicrobials [75, 162], aptamers [163], synthetic polymers [164], optical immunoassays [122, 125, 165], waveguide technology [75, 166, 167], lipid bilayers [9, 74, 168], and invitro [169171] assays have been applied for LPS detection. These technologies involve functionalizing biosensors with proteins or molecules to pull down LPS from a sample.

LBP [10], a relatively small protein (~60 kDa) that transports LPS in blood, shuttles the antigen to the cluster of differentiation 14 (CD14) protein in the extracellular matrix, or to the membrane of immune cells, such as macrophages [10]. After LPS binds CD14, it is passed to the hydrophobic binding pocket of myeloid differentiation factor 2 (MD‐2) [7, 10], a necessary cofactor for the activation of TLR4. Also, the serum carrier lipoproteins (HDL and LDL), are carriers for LPS in blood. In addition to these, LPS has been demonstrated to bind aptamers [159, 160], various peptides [87, 109, 162, 172], and metal/cation complexes [84, 86, 161, 173175]. Such carrier moieties are exploited in the development of novel detection methods for LPS, as outlined below.

For electrochemical (EC) sensing of LPS, a recognition ligand (similar to ELISA) and a transducer are required to measure the variation in signal [161]. For fluorescence‐based sensing, a receptor captures LPS, while another molecule emits a fluorescent signal when bound to the antigen. Burkhardt et al. [176] used solubilized LBP to transfer LPS to a CD14 functionalized surface, with a LoD of 10 ng/mL using an electro‐chemiluminescent assay. This method enforces the role of LBP as a lipid transfer protein, as demonstrated by Wurfel et al. [177, 178] and shows that CD14 can bind monomeric LPS in the absence of TLR4 [179]. Highly sensitive (LoD = 0.0005 EU) EC sensors have also been developed using a recombinant TLR4 + MD‐2 complex for recognition of LPS [180]. Yet, these assays are unable to discriminate between LPS serogroups. Priano et al. [10] developed a competitive EC assay using recombinant endotoxin‐neutralizing protein (ENP) on a dextran matrix, with a detection range of 1–100 ng/mL. ENP has also been used in a capacitive biosensor with an extremely low LoD (1.0 × 10−13 M) [181]. The sensitivity differences may be due to variations in surface functionalization. Priano et al. [10] used the dextran matrix, and Limbut et al. [181] used self‐assembled monolayers, which provide low background interference [182184]. Inoue and Takano [10] used a recombinant factor C in an EC hybrid LAL biosensor, with a sensitivity range of 5 × 10−4–1.0 EU/mL [10]. Kato [87] and Iijima [10] labeled PmB with ferrocene‐bound LPS in solution, and captured it on a nanocarbon‐film electrode with a detection range of 2–50 ng/mL in 5 minutes [10]. Ding et al. [162] functionalized an electrode with PmB and performed EIS with a detection range of 0.2–0.8 ng/mL which is more sensitive, but has a smaller range. A broader detection range was demonstrated by Rahman et al. [172] who functionalized interdigitated electrodes with PmB and tested 0.1–1000 µg/mL of LPS O111:B4 in food samples, using impedance spectroscopy. Sugar binding proteins, such as lectins and polyaniline coated electrodes, have been used for detecting LPS [10], as with an EIS sensor functionalized with the lectin, cramoLL, with a detection range of 25–200 µg/mL [185].

Several assays have been developed using aptamers as the detection ligand. Su et al. [160, 186, 187] used aptamers attached to gold nanoparticles to detect LPS using EIS, with an impressive detection limit of 0.1 pg/mL [10]. Aptamers have also been used in a magnetic aptasensor to detect LPS in medias containing BSA, sucrose, glucose, or RNA [163], and provide a detection range of 0.01–1.0 × 106 ng/mL (LPS O55:B5) by flow cytometry within 1 minute. Bai et al. [188] developed an EC sensor where aptamers that bind LPS were hybridized with capture probes, which were hybridized to complementary DNA sequences on gold nanoparticles with a very sensitive range (10 fg/mL up to 50 ng/mL). However, multiple aptamer libraries against O‐ag would be essential before this method could be implemented for serogroup discrimination. Modifications to improve sensitivity include use of SAMs to functionalize sensors with peptides [189], PmB [162], antibodies [10], and aptamers [159]. Despite optimal surface capture methods, some of these assays suffer from poor detection limits or range of performance [10, 159].

Investigators have utilized the interaction of LPS with synthetic systems such as copolythiophene copolymers [164] and polydiacetylene liposomes [190, 191]. Johnson et al. [192] demonstrated an endotoxin capture technique by functionalizing a bead matrix with proanthocyanidins and binding with fluorescein isothiocyanate‐labeled LPS [192]. Pyrenyl‐derived quaternary ammonium probes, developed by Zeng et al. [193] exhibited fluorescence when bound to LPS and detected nanomolar concentrations, while fluorescently labeled CD14 synthetic peptides demonstrated an increase in Förster resonance energy transfer when bound to LPS, but were only able to detect µM concentrations [194]. Lim et al. [195] used a functionalized graphene oxide to develop a fluorescence quench‐recovery method for LPS, targeting the lipid A component. Thompson et al. [196] designed a tandem system to both detect (LoD = 1.0 ng/mL)and filter LPS from blood using piezoelectric quartz discs functionalized with PmB.

Other methods have taken advantage of the amphipathic nature of LPS. Harmon et al. [197] demonstrated that disrupting the hydrophobic association of LPS with liposomes increases the sensitivity of the LAL assay. Stromberg et al. [74, 198] were able to detect 4.20 µg/mL of amphiphilic LPS O157 in beef lysates on a waveguide biosensor using a technique called membrane insertion, which has previously been applied to other amphiphiles such as LAM and phenolic glycolipids [8, 9, 199]. Membrane insertion uses the natural association of amphiphiles with a lipid bilayer to facilitate detection and fluorescent detection of a labeled antibody is performed within an evanescent field [168, 199]. Many biosensors report exquisite sensitivity, even down to the picogram [164] and femtomolar [9, 168, 199] range, but very few are capable of physiological presentation of amphiphiles to facilitate discriminative detection of O‐ag groups [74, 167, 198].

4.4. Cell‐based LPS detection systems

Cell systems are ideal for recognizing endotoxin, although interpreting the signal response can be challenging. Bouafsoun et al. [169] functionalized the surface of an impedance biosensor with endothelial cells, and measured the decrease in impedance with LPS binding, with a sensitivity of 500 ng/mL. Veiseh et al. [200] patterned macrophage cells onto gold electrodes to detect LPS concentrations of 0.1–10 µg/mL. However, cells were concurrently stained with necrosis and apoptosis markers in parallel studies, and no staining effect could be seen in cells using concentrations less than 10 µg/mL. This is an interesting effect, as in many invitro studies, cytokine response is induced at much lower concentrations of endotoxin [97, 201, 202]. It can be deduced that Veiseh used serum supplemented media in the experiments, and the lipoproteins and LBP in serum could have a protective effect on cells [46, 202, 203], and attenuated assay sensitivity. The most sensitive cell‐based assay was developed by Inoue et al. [170] with a LoD of 0.1–1.0 ng/mL. Here, cells were engineered to secrete alkaline phosphatase in the presence of LPS, and patterned on the surface of an amperometric biosensor to measure voltage change upon LPS binding. Cell‐based invitro assays are prone to errors and contamination, so developing a robust and fieldable assay based on this technology is not plausible. However, by studying LPS in cell‐based systems, knowledge about interactions with receptors and cell membranes can be gained, which can facilitate better detection methods.

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5. Conclusions

Many novel approaches have been used for the detection of amphiphilic LPS, not all of which are functional in physiological matrices or have the required sensitivity or ease of use. One major reason for this is the failure to incorporate the amphiphilic properties of the antigen into assay design. The presentation, conformation, and host‐interactions of the antigens should be considered for the development of effective assays. While both LAL and EC assays are the most sensitive for testing endotoxicity, identifying O‐ag with a high degree of selectivity remains elusive, and limited to methods that use specific recognition ligands, such as membrane insertion and ELISAs. By far, the greatest limitation has been the lack of sensitive and selective ligands for the serogroup‐specific detection of the antigen. Thus, as repositories of these necessary recognition molecules expand to include more serogroups, so too will our ability to selectively detect LPS.

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Acknowledgments

This work was supported by the Agriculture and Food Research Initiative Competitive Grant no. 2012‐68003‐30155 from the United States Department of Agriculture's National Institute of Food and Agriculture.

References

  1. 1. Love TE, Jones B. Introduction to pathogenic bacteria. Principles of Bacterial Detection: Biosensors, Recognition Receptors and Microsystems. New York, NY: Springer New York; 2008. pp. 3-13
  2. 2. Kumagai Y, Takeuchi O, Akira S. Pathogen recognition by innate receptors. The Journal of Infectious Diseases. 2008;14:86-92
  3. 3. Broz P, Monack DM. Newly described pattern recognition receptors team up against intracellular pathogens. Nature Reviews Immunology. 2013;13:551-565
  4. 4. Miyake K. Innate recognition of lipopolysaccharide by Toll‐like receptor 4‐MD‐2. Trends in Microbiology. 2004;12:186-192
  5. 5. Deisingh AK, Thompson M. Biosensors for the detection of bacteria. Canadian Journal of Microbiology. 2004;50:69-77
  6. 6. Kawai T, Akira S. TLR signaling. Seminars in Immunology. 2007;19:24-32
  7. 7. Kumar H, Kawai T, Akira S. Pathogen recognition by the innate immune system. International Reviews of Immunology. 2011;30:16-34
  8. 8. Sakamuri RM, Price DN, Lee M, Cho SN, Barry CE, Via LE, et al. Association of lipoarabinomannan with high density lipoprotein in blood: Implications for diagnostics. Tuberculosis. 2013;93:301-307
  9. 9. Sakamuri RM, Capek P, Dickerson TJ, Barry CE, Mukundan H, Swanson BI. Detection of stealthy small amphiphilic biomarkers. Journal of Microbiological Methods. 2014;103:112-117
  10. 10. Su W, Ding X. Methods of endotoxin detection. Journal of Laboratory Automation. 2015;20:354-364
  11. 11. Alexander C, Rietschel ET. Bacterial lipopolysaccharides and innate immunity. Journal of Endotoxin Research. 2001;7:167-202
  12. 12. Jauho ES, Boas U, Wiuff C, Wredstrøm K, Pedersen B, Andresen LO, et al. New technology for regiospecific covalent coupling of polysaccharide antigens in ELISA for serological detection. Journal of Immunological Methods. 2000;242:133-143
  13. 13. Munford RS. Sensing Gram‐Negative bacterial lipopolysaccharides: A human disease determinant? Infection and Immunity. 2008;76:454-465
  14. 14. Schreiber A, Humbert M, Benz A, Dietrich U. 3D‐Epitope‐Explorer (3DEX): Localization of conformational epitopes within three‐dimensional structures of proteins. Journal of Computational Chemistry. 2005;26:879-887
  15. 15. Beveridge TJ. Structures of Gram‐negative cell walls and their derived membrane vesicles. Journal of Bacteriology. 1999;181:4725-4733
  16. 16. Joly‐Guillou M‐L. Clinical impact and pathogenicity of Acinetobacter. Clinical Microbiology and Infection, European Journal of Clinical Microbiology & Infectious Diseases. 2005;11:868-873
  17. 17. Wiersinga WJ, van der Poll T, White NJ, Day NP, Peacock SJ. Melioidosis: Insights into the pathogenicity of Burkholderia pseudomallei. Nature Reviews Microbiology. 2006;4:272-282
  18. 18. Mattoo S, Foreman‐Wykert AK, Cotter PA. Mechanisms of Bordetella pathogenesis. Frontiers in Bioscience. 2001;6:E168–E186
  19. 19. Brooks BW, Devenish J, Lutze‐Wallace CL, Milnes D, Robertson, RH, Berlie‐Surujballi G. Evaluation of a monoclonal antibody‐based enzyme‐linked immunosorbent assay for detection of Campylobacter fetus in bovine preputial washing and vaginal mucus samples. Veterinary Microbiology. 2004;103:77-84
  20. 20. Lazcka O, Del Campo FJ, Muñoz FX. Pathogen detection: A perspective of traditional methods and biosensors. Biosensors & Bioelectronics. 2007;22:1205-1217
  21. 21. Ketley JM. Pathogenesis of enteric infection by Campylobacter. Microbiology. 1997;143:5-21
  22. 22. Lüderitz O, Freudenberg MA, Galanos C, Lehmann V, Rietschel ET, Shaw DH. Lipopoly-saccharides of Gram‐Negative Bacteria. Current Topics in Membranes and Transport. Elsevier; 1982. pp. 79-151
  23. 23. Belland RJ, Ouellette SP, Gieffers J, Byrne GI. Chlamydia pneumoniae and atherosclerosis. Cell Microbiology. 2004;6:117-127
  24. 24. Leimbach A, Hacker J, Dobrindt U. E. coli as an All‐Rounder: The Thin Line Between Commensalism and Pathogenicity. Berlin, Heidelberg: Springer; microbiology and immunology. 2013;358:3-32
  25. 25. Moran AP, Lindner B, Walsh EJ. Structural characterization of the lipid A component of Helicobacter pylori rough‐ and smooth‐form lipopolysaccharides. Journal of Bacteriology. 1997;179:6453-6463
  26. 26. Yamaoka Y, Graham DY. Helicobacter pylori virulence and cancer pathogenesis. Future Oncology. 2014;10:1487-1500
  27. 27. Livorsi DJ, Macneil JR, Cohn AC, Bareta J, Zansky S, Petit S, et al. Invasive Haemophilus influenzae in the United States, 1999-2008: Epidemiology and outcomes. Journal of Infection. Elsevier Ltd. 2012;65:496-504
  28. 28. Meatherall BL, Gregson D, Ross T, Pitout JDD, Laupland KB. Incidence, risk factors, and outcomes of Klebsiella pneumoniae bacteremia. American Journal of Medicine. 2009;122:866-873
  29. 29. Newton HJ, Ang DKY, van Driel IR, Hartland EL. Molecular pathogenesis of infections caused by Legionella pneumophila. Clinical Microbiology Reviews. 2010;23:274-298
  30. 30. Laura Perez Vidakovics M, Riesbeck K. Virulence mechanisms of Moraxella in the pathogenesis of infection. Current Opinion in Infectious Diseases. 2009;22:279-285
  31. 31. Stephens DS. Biology and pathogenesis of the evolutionarily successful, obligate human bacterium Neisseria meningitidis. Vaccine. 2009;27:B71–B77
  32. 32. Mahajan‐Miklos S, Tan MW, Rahme LG, Ausubel FM. Molecular mechanisms of bacterial virulence elucidated using a Pseudomonas aeruginosa‐Caenorhabditis elegans pathogenesis model. Cell. 1999;96:47‐56
  33. 33. Mobley HL, Belas R. Swarming and pathogenicity of Proteus mirabilis in the urinary tract. Trends Microbiology. 1995;3:280‐284
  34. 34. Mather AE, Reid S, Maskell DJ, Parkhill J, Fookes MC, Harris S, et al. Distinguishable epidemics within different hosts of the multidrug resistant zoonotic pathogen Salmonella Typhimurium DT104. Science. 2013;341(6153):1514
  35. 35. Caboni M, Pédron T, Rossi O, Goulding D, Pickard D, Citiulo F, et al. An O antigen capsule modulates bacterial pathogenesis in Shigella sonnei. Koehler TM, editor. PLoS Pathogens. 2015;11:e1004749
  36. 36. Fábrega A, Vila J. Yersinia enterocolitica: Pathogenesis, virulence and antimicrobial resistance. Medicina Clinica. SEGO. 2012;30:24-32
  37. 37. Bennett IL, Cluff LE. Bacterial pyrogens. Pharmacological Reviews. 1957;9:427-475
  38. 38. Westphal O, Westphal U. The history of pyrogen research. Microbiology. 1977;1977: 221-238
  39. 39. Beutler B, Rietschel ET. Innate immune sensing and its roots: The story of endotoxin. Nature Reviews Immunology. 2003;3:169-176
  40. 40. Olins AL, Warner RC. Physicochemical studies on a lipopolysaccharide from the cell wall of Azotobacter vinelandii. Journal of Biological Chemistry. 1967;242(21):4994-5001
  41. 41. Schromm AB, Bradenburg K, Loppnow H, Zähringer U, Rietschel ET, Carroll SF, et al. The charge of endotoxin molecules influences their conformation and IL‐6‐inducing capacity. Journal of Immunology. 1998;161:5464-5471
  42. 42. Bishop RE. Fundamentals of endotoxin structure and function. In: Herwald RW, editor. Contributions to Microbiology. Basel: Karger; 2005. pp. 1-27
  43. 43. Meredith TC, Aggarwal P, Mamat U, Lindner B, Woodard RW. Redefining the requisite lipopolysaccharide structure in Escherichia coli. ACS Chemical Biology. 2006;1:33-42
  44. 44. Raetz CRH, Guan Z, Ingram BO, Six DA, Song F, Wang X, et al. Discovery of new biosynthetic pathways: The lipid A story. Journal of Lipid Research. 2009;50(Suppl):S103‐S108
  45. 45. Raetz CRH, Whitfield C. Lipopolysaccharide endotoxins. Annual Review of Biochemistry. 2002;71:635-700
  46. 46. Hurley JC. Endotoxemia: Methods of detection and clinical correlates. Clinical Microbiology Reviews. 1995;8:268-292
  47. 47. Erridge C, Bennett‐Guerrero E, Poxton IR. Structure and function of lipopolysaccharides. Microbes and Infection. 2002;4:837-851
  48. 48. Aurell CA, Wistrom AO. Critical aggregation concentrations of Gram‐negative bacterial lipopolysaccharides (LPS). Biochemical and Biophysical Research Communications. 1998;253:119-123
  49. 49. Plested JS, Makepeace K, Jennings MP, Gidney MA, Lacelle S, Brisson J, et al. Conservation and accessibility of an inner core lipopolysaccharide epitope of Neisseria meningitidis. Infection and Immunity. 1999;67:5417-5426
  50. 50. Silipo A, Molinaro A. The diversity of the core oligosaccharide in lipopolysaccharides. Subcellular Biochemistry. Dordrecht: Springer Netherlands. 2010;53:69-99
  51. 51. Nevola JJ, Stocker BA, Laux DC, Cohen PS. Colonization of the mouse intestine by an avirulent Salmonella typhimurium strain and its lipopolysaccharide‐defective mutants. Infection and Immunity. 1985;50:152-159
  52. 52. Murray GL, Attridge SR, Morona R. Regulation of Salmonella typhimurium lipopolysaccharide O antigen chain length is required for virulence; identification of FepE as a second Wzz. Molecular Microbiology. 2003;47:1395-1406
  53. 53. De Bagüés M, Elzer PH, Jones SM. Vaccination with Brucella abortus rough mutant RB51 protects BALB/c mice against virulent strains of Brucella abortus, Brucella melitensis, and Brucella ovis. Infection and Immunity. 1994;62(11):4990-4996
  54. 54. Stenutz R, Weintraub A, Widmalm G. The structures of Escherichia coli O‐polysaccharide antigens. FEMS Microbiology Reviews. 2006;30:382-403
  55. 55. Goldman RC, Leive L. Heterogeneity of antigenic‐side‐chain length in lipopolysaccharide from Escherichia coli 0111 and Salmonella typhimurium LT2. European Journal of Biochemistry. 1980;107:145-153
  56. 56. Bradenburg K, Schromm AB, Gutsmann T. Endotoxins: Relationship between structure, function, and activity. Structure. Dordrecht: Springer Netherlands. 2004;53:53-67
  57. 57. Bergstrand A, Svanberg C, Langton M, Nydén M. Aggregation behavior and size of lipopolysaccharide from Escherichia coli O55:B5. Colloids and Surfaces B: Biointerfaces. 2006;53:9-14
  58. 58. Santos NC, Silva AC, Castanho MARB, Martins Silva J, Saldanha C. Evaluation oflipopolysaccharide aggregation by light scattering spectroscopy. ChemBioChem. 2003;4:96-100
  59. 59. Israelachvili J. Intermolecular and Surface Forces. 2nd ed. London: Academic Press; 1991
  60. 60. Ruckenstein E, Nagarajan R. Critical micelle concentration. Transition point for micellar size distribution. Journal of Physical Chemistry (American Chemical Society). 1975;79:2622-2626
  61. 61. Din ZZ, Mukerjee P, Kastowsky M, Takayama K. Effect of pH on solubility and ionic state of lipopolysaccharide obtained from the deep rough mutant of Escherichia coli. Biochemistry. 1993;32:4579-4586
  62. 62. Sasaki H, White SH. Aggregation behavior of an Ultra‐Pure lipopolysaccharide that stimulates TLR‐4 receptors. Biophysical Journal. 2008;95:986-993
  63. 63. Moroi Y. CMC range due to polydispersity of micelles. Journal of Colloid and Interface Science. 1991;141:581-583
  64. 64. Krüger D. Assessing the quality of medicinal products containing ingredients obtained by gene technology. Drugs Made in Germany. 1989;32(2):64-67
  65. 65. Hirayama C, Sakata M. Chromatographic removal of endotoxin from protein solutions by polymer particles. Journal of Chromatography B Analytical Technologies Biomedical and Life Sciences. 2002;781:419-432
  66. 66. Netea MG, van Deuren M, Kullberg BJ, Cavaillon J‐M, Van der Meer JWM. Does the shape of lipid A determine the interaction of LPS with Toll‐like receptors? Trends in Immunology. 2002;23:135-139
  67. 67. Seydel U, Schromm AB, Blunck R, Bradenburg K. Chemical structure, molecular conformation, and bioactivity of endotoxins. Chemical Immunology. 2000;74:5-24
  68. 68. Nagarajan R. Molecular packing parameter and surfactant Self‐Assembly: The neglected role of the surfactant tail †. Langmuir. 2002;18:31-38
  69. 69. Rietschel ET, Brade H, Brade L, Kaca W, Kawahara K, Lindner B, et al. Newer aspects of the chemical structure and biological activity of bacterial endotoxins. Progress in Clinical and Biological Research. 1985;189:31-51
  70. 70. Jucker BA, Harms H, Zehnder A. Polymer interactions between five gram‐negative bacteria and glass investigated using LPS micelles and vesicles as model systems. Colloids and Surfaces B: Biointerfaces. 1998;11:33-45
  71. 71. Petsch D, Anspach FB. Endotoxin removal from protein solutions. Journal of Biotechnology. 2000;76:97-119
  72. 72. Mueller M, Lindner B, Kusumoto S, Fukase K, Schromm AB, Seydel U. Aggregates are the biologically active units of endotoxin. Journal of Biological Chemistry. 2004;279:26307-26313
  73. 73. Vasselon T, Hailman E, Thieringer R, Detmers PA. Internalization of monomeric lipopolysaccharide occurs after transfer out of cell surface CD14. Journal of Experimental Medicine. 1999;190:509-521
  74. 74. Stromberg LR, Hengartner NW, Swingle KL, Moxley RA, Graves SW, Montaño GA, et al. Membrane insertion for the detection of lipopolysaccharides: Exploring the dynamics of Amphiphile‐in‐Lipid Assays. Gasset M, editor. PLoS ONE. 2016;11:e0156295
  75. 75. James EA, Schmeltzer K, Ligler FS. Detection of endotoxin using an evanescent wave fiber‐optic biosensor. Applied Biochemistry and Biotechnology. 1996;60:189-202
  76. 76. Kannenberg EL, Perotto S, Bianciotto V, Rathbun EA, Brewin NJ. Lipopolysaccharide epitope expression of Rhizobium bacteroids as revealed by in situ immunolabelling of pea root nodule sections. Journal of Bacteriology. 1994;176:2021-2032
  77. 77. Haselhorst T, Espinosa J‐F, Jiménez‐Barbero J, Sokolowski T, Kosma P, Brade H, et al. NMR experiments reveal distinct Antibody‐Bound conformations of a synthetic disaccharide representing a general structural element of bacterial lipopolysaccharide epitopes. Biochemistry. 1999;38:6449-6459
  78. 78. Hershberger C, Binkley SB. Chemistry and Metabolism of 3‐Deoxy‐d‐mannooctu-losonic acid I. Sterochemical determination. Journal of Biological Chemistry. 1968;243(7):1578-1584
  79. 79. Brandenburg K, Andrä J, Müller M, Koch MHJ, Garidel P. Physicochemical properties of bacterial glycopolymers in relation to bioactivity. Carbohydrate Research. 2003;338:2477-2489
  80. 80. Aurell CA, Hawley ME, Wistrom AO. Direct visualization of Gram‐negative bacterial lipopolysaccharide self‐assembly. Molecular Cell Biology Research Communications. 1999;2:42-46
  81. 81. Adams PG, Lamoureux L, Swingle KL, Mukundan H, Montaño GA. Lipopolysaccharide‐induced dynamic lipid membrane reorganization: Tubules, perforations, and stacks. Biophysical Journal. 2014;106:2395-2407
  82. 82. van Alphen L, Verkleij A, Burnell E, Lugtenberg B. 31P nuclear magnetic resonance and freeze‐fracture electron microscopy studies on Escherichia coli. II. Lipopolysaccharide and lipopolysaccharide‐phospholipid complexes. Biochimica et Biophysica Acta (BBA) ‐ Biomembranes. 1980;597:502-517
  83. 83. Coughlin RT, Haug A, McGroarty EJ. Physical properties of defined lipopolysaccharide salts. Biochemistry. 1983;22:2007-2013
  84. 84. Coughlin RT, Tonsager S, McGroarty EJ. Quantitation of metal cations bound to membranes and extracted lipopolysaccharide of Escherichia coli. Biochemistry. 1983;22:2002-2007
  85. 85. Naumann D, Schultz C, Sabisch A, Kastowsky M, Labischinski H. New insights into the phase behaviour of a complex anionic amphiphile: Architecture and dynamics of bacterial deep rough lipopolysaccharide membranes as seen by FTIR, X‐ray, and molecular modelling techniques. Journal of Molecular Structure. 1989;214:213-246
  86. 86. Schindler M, Osborn MJ. Interaction of divalent cations and polymyxin B with lipopolysaccharide. Biochemistry. 1979;18:4425-4430
  87. 87. Kato D, Iijima S, Kurita R, Sato Y, Jia J, Yabuki S, et al. Electrochemically amplified detection for lipopolysaccharide using ferrocenylboronic acid. Biosensors & Bioelectronics. 2007;22:1527-1531
  88. 88. McClosky WT, Price CW, Van Winkle W, Welch H, Calvery HO. Results of first U. S. P. Collaborative study of pyrogens. Journal of the American Pharmaceutical Association (Scientific edition). 1943;32:69-73
  89. 89. Cooper JF, Levin J, Wagner HN. Quantitative comparison of in vitro and in vivo methods for the detection of endotoxin. Journal of Laboratory and Clinical Medicine. 1971;78:138-148
  90. 90. Roberts KJ. Endotoxins: Pyrogens, LAL testing and depyrogenation. Williams KL, editor. CRC Press, Taylor & Fancis Group LLC; 2007. pp. 261-284
  91. 91. Park C‐Y, Jung S‐H, Bak J‐P, Lee S‐S, Rhee D‐K. Comparison of the rabbit pyrogen test and Limulus amoebocyte lysate (LAL) assay for endotoxin in hepatitis B vaccines and the effect of aluminum hydroxide. Biologicals. 2005;33:145-151
  92. 92. Levin J, Tomasulo PA, Oser RS. Detection of endotoxin in human blood and demonstration of an inhibitor. Journal of Laboratory and Clinical Medicine. 1970;75:903-911
  93. 93. Pharmacopeial Convention US. 85> Bacterial Endotoxins Test. 2011. The United States Pharmacopeial Convention (P)\\usp-netapp2\share\SHARE\USPNF\PRINTQ\pager\xmlIn\AO_20111014080832_M98830.xml
  94. 94. U.S. Department of Health and Human Services, U.S. Food and Drug Administration. Guidance for Industry. 21 CFR; Dec 8, 1987 pp. 1-54.
  95. 95. Novitsky TJ, Roslansky PF. Quantification of endotoxin inhibition in serum and plasma using a turbidimetric LAL assay. Progress in Clinical and Biological Research. 1985;189:181-196
  96. 96. Levin J, Poore TE, Zauber NP, Zauber NP, Oser RS. Detection of endotoxin in the blood of patients with sepsis due to Gram‐negative bacteria. The New England Journal of Medicine. 1970;283:1313-1316
  97. 97. Cavaillon JM, Fitting C, Haeffner‐Cavaillon N, Kirsch SJ, Warren HS. Cytokine response by monocytes and macrophages to free and lipoprotein‐bound lipopolysaccharide. Infection and Immunity. 1990;58:2375-2382
  98. 98. Nachum R, Shanbrom E. Rapid detection of Gram‐negative bacteriuria by limulus amoebocyte lysate assay. Journal of Clinical Microbiology. 1981;13:158-162
  99. 99. Svensson A, Hahn‐Hägerdal B. Comparison of a gelation and a chromogenic Limulus (LAL) assay for the detection of Gram‐negative bacteria, and the application of the latter assay to milk. Journal of Dairy Research. 1987;54:267-273
  100. 100. Jay JM, Margitic S, Shereda AL, Covington HV. Determining endotoxin content of ground beef by the Limulus amoebocyte lysate test as a rapid indicator of microbial quality. Applied and Environmental Microbiology. 1979;38:885-890
  101. 101. Jay JM. Rapid estimation of microbial numbers in fresh ground beef by use of the Limulus test. Journal of Food Protection. 1981;44:275-278
  102. 102. Goto H, Nakamura S. Dry up method as a revised Limulus test with a new technique for gelatin inhibitor removing. The Japanese Journal of Experimental Medicine. 1979;49:19-25
  103. 103. Hartman I, Ziv G, Saran A. Application of the Limulus amoebocyte lysate test to the detection of Gram‐negative bacterial endotoxins in normal and mastitic milk. Research in Veterinary Science. 1976;20:342-343
  104. 104. Reinhold RB, Fine J. A technique for quantitative measurement of endotoxin in human plasma. Proceedings of the Society for Experimental Biology and Medicine. 1971. 334-340
  105. 105. Maxie MG, Valli VE. Studies with radioactive endotoxin. III. Localization of 3H‐labelled endotoxin in the formed elements of the blood and detection of endotoxin in calf blood with the Limulus amebocyte lysate. Canadian Journal of Comparative Medicine‐Revue Canadienne De Medecine Comparee. 1974;38:383-390
  106. 106. Yin ET, Galanos C, Kinsky S, Bradshaw RA, Wessler S, Lüderitz O, et al. Picogram‐sensitive assay for endotoxin: Gelation of Limulus polyphemus blood cell lysate induced by purified lipopolysaccharides and lipid A from Gram‐negative bacteria. Biochimica et Biophysica Acta. 1972;261:284-289
  107. 107. Takagi K, Moriya A, Tamura H, Nakahara C. Quantitative measurement of endotoxin in human blood using synthetic chromogenic substrate for horseshoe crab clotting enzyme: A comparison of methods of blood. Thrombosis Research. 1981;23:51-57
  108. 108. Niwa M, Hiramatsu T, Waguri O. Proceedings: Quantitative aspects of the gelation reaction of horseshoe crab amoebocyte and bacterial endotoxin. Symp. Toxins. jpn j of medical sci biology. 1974;28:98-100
  109. 109. Tsuji K, Martin PA, Gaunnac GL. Recovery of endotoxin from human plasma by acid oxidative treatmtnets as monitored by an automated microtiter plate‐chromogenic substrate limulus amebocyte lysate (LAL) assay method. Progress in Clinical and Biological Research. 1986;231: 443-457
  110. 110. Karibian D, Deprun C, Caroff M. Comparison of lipids A of several Salmonella and Escherichia strains by 252Cf plasma desorption mass spectrometry. Journal of Bacteriology. 1993;175: 2988-2993
  111. 111. Schletter J, Heine H, Ulmer AJ, Rietschel ET. Molecular mechanisms of endotoxin activity. Archives of Microbiology. 1995;164:383-389
  112. 112. Engvall E, Perlmann P. Enzyme‐linked immunosorbent assay, ELISA III: Quantitation of specific antibodies by enzyme‐labeled anti‐immunoglobulin in antigen‐coated tubes. Journal of Immunology. 1972;109:129-135
  113. 113. Van Weemen BK, Schuurs AHWM. Immunoassay using antigen‐enzyme conjugates. FEBS Letters. 1971;15:232-236
  114. 114. Carlsson HE, Lindberg AA, Hammarström S. Titration of antibodies to Salmonella O antigens by enzyme‐linked immunosorbent assay. Infection and Immunity. 1972;6:703-708
  115. 115. Nielsen B, Baggesen D, Bager F, Haugegaard J, Lind P. The serological response to Salmonella serovars typhimurium and infantis in experimentally infected pigs. The time course followed with an indirect anti‐LPS ELISA and bacteriological examinations. Veterinary Microbiology. 1995;47: 205-218
  116. 116. Nalepka JL, Greenfield EM. Detection of bacterial endotoxin in human tissues. Biotechniques. 2004;37:413-417
  117. 117. Reither K, Saathoff E, Jung J, Minja LT, Kroidl I, Saad E, et al. Low sensitivity of a urine LAM‐ELISA in the diagnosis of pulmonary tuberculosis. BMC Infectious Diseases. 2009;9:141
  118. 118. Takahashi K, Fukada M, Kawai M, Yokochi T. Detection of lipopolysaccharide (LPS) and identification of its serotype by an enzyme‐linked immunosorbent assay (ELISA) using poly‐l‐lysine. Journal of Immunological Methods. 1992;153:67-71
  119. 119. Scott BB, Barclay GR. Endotoxin-polymyxin complexes in an improved enzyme-linked immunosorbent assay for IgG antibodies in blood donor sera to Gram-Negative endotoxin core glycolipids. Vox Sanguinis. 1987;52:272-280
  120. 120. Sternberger LA. The unlabeled antibody (PAP) method. Journal of Histochemistry & Cytochemistry. 1979;27:1657-1657
  121. 121. Hsu SM, Raine L, Fanger H. The use of antiavidin antibody and avidin‐biotin‐peroxidase complex in immunoperoxidase technics. American Journal of Clinical Pathology. 1981;75:816-821
  122. 122. Mackenzie A, Lebel P, Orrbine E, Rowe PC, Hyde L, Chan F, et al. Sensitivities and Specificities of premier E. coli O157 and premier EHEC enzyme immunoassays for diagnosis of infection with verotoxin (Shiga‐Like Toxin)‐producing Escherichia coli. Journal of Clinical Microbiology. 1998;36:1608-1611
  123. 123. Vreugdenhil AC, Snoek AM, van ‘t Veer C, Greve JW, Buurman WA. LPS‐binding protein circulates in association with apoB‐containing lipoproteins and enhances endotoxin‐LDL/VLDL interaction. Journal of Clinical Investigation. 2001;107:225-234
  124. 124. Vreugdenhil A, Rousseau CH. Lipopolysaccharide (LPS)‐binding protein mediates LPS detoxification by chylomicrons. Journal of Immunology. 2003;170:1399-1405
  125. 125. Kerr P, Chart H, Finlay D, Pollock DA, MacKie DP, Ball HJ. Development of a monoclonal sandwich ELISA for the detection of animal and human Escherichia coli O157 strains. Journal of Applied Microbiology. 2001;90:543-549
  126. 126. Choi D, Tsang RS, Ng MH. Sandwich capture ELISA by a murine monoclonal antibody against a genus‐specific LPS epitope for the detection of different common serotypes of salmonellas. Journal of Applied Bacteriology. 1992;72:134-138
  127. 127. Schütz M, Meyer R, Grallert H, Miller S. Method for Detecting and for Removing Endotoxin [Internet]. 2010. US Patent Office; US20090017445 A1
  128. 128. Schütz M, Meyer R, Grallert H, Miller S. Method for identifying and for extracting endotoxin. 2009. US Patent Office; 7,585,620
  129. 129. Grallert H, Leopoldseder S, Schuett M, Kurze P, Buchberger B. EndoLISA®: A novel and reliable method for endotoxin detection. Nature Reviews Immunology. Nature Publishing Group. 2011:8:iii‐v
  130. 130. Alberts B. Molecular Biology of the Cell. 5th ed. Garland Science; 2008. pp. 1539-1557
  131. 131. Hyams KC, Malone JD, Bourgeois AL, Hawkins R, Hale TL, Murphy JR. Serum antibody to lipopolysaccharide antigens of Shigella species among U.S. military personnel deployed to Saudi Arabia and Kuwait during Operations Desert Shield and Desert Storm. Clinical and Diagnostic Laboratory Immunology. 1995;2:700-703
  132. 132. Blatz R, Aseffa A, Gedefaw M, Rühle HJ, Forberg J, Christian AR. Prevalence of Chlamydia specific antibodies among obstetric and gynecological outpatients in Gondar, North‐west Ethiopia. Ethiopia Medical Journal. 2001;39:293-303
  133. 133. House D, Wain J, Ho VA, Diep TS, Chinh NT, Bay PV, et al. Serology of typhoid fever in an area of endemicity and its relevance to diagnosis. Journal of Clinical Microbiology. 2001;39:1002-1007
  134. 134. Chaicumpa W, Ngren‐ngarmlert W, Kalambaheti T, Ruangkunaporn Y, Chongsa‐Nguan M, Tapchaisri P, et al. Monoclonal antibody‐based dot‐blot ELISA for the detection of Salmonella in foods. Asian Pacific Journal of Allergy and Immunology. 1995;13:159-166
  135. 135. Freudenberg MA, Fomsgaard A, Mitov I, Galanos C. ELISA for antibodies to lipid A, lipopolysaccharides and other hydrophobic antigens. Infection. 1989;17:322-328
  136. 136. Suthienkul O, Poomchart A, Kositanont U, Siripanichgon K, Vathanophas K. ELISA for seroepidemiological study of exposure to Vibrio cholerae of population in Krabi Province, Thailand. Asian Pacific Journal of Allergy and Immunology. 1992;10:55-60
  137. 137. Chaicumpa W, Ruangkunaporn Y, Burr D, Chongsa‐Nguan M, Echeverria P. Diagnosis of typhoid fever by detection of S typhi antigen in urine. Journal of Clinical Microbiology. 1992;30:2513-2515
  138. 138. Covinsky M, Laterza O, Pfeifer JD, Farkas‐Szallasi T, Scott MG. An IgM lambda antibody to Escherichia coli produces false‐positive results in multiple immunometric assays. Clinical Chemistry. 2000;46:1157-1161
  139. 139. Ben‐Hur H, Gurevich P, Elhayany A, Avinoach I, Schneider D, Zusman I. Transport of maternal immunoglobulins through the human placental barrier in normal pregnancy and during inflammation. International Journal of Molecular Medicine. 2005;16:401-407
  140. 140. Morrison DC, Jacobs DM. Binding of polymyxin B to the lipid A portion of bacterial lipopolysaccharides. Immunochemistry. 1976;13:813-818
  141. 141. Mattsby‐Baltzer I, Jakobsson A, Sörbo J. Endotoxin is angiogenic. International Journal of Experimental Pathology. 1994;75:191-196
  142. 142. Hardy E, Ohlin M, Llano M. Enhanced ELISA sensitivity using TCA for efficient coating of biologically active lipopolysaccharides or lipid A to the solid phase. Journal of Immunological Methods. 1994;176:111-116
  143. 143. Weintraub A, Widmalm G, Jansson PE, Jansson M. Vibrio cholerae O139 Bengal possesses a capsular polysaccharide which may confer increased virulence. Microbial Pathogenesis. 1994;16:235-241
  144. 144. Stromberg LR, Stromberg ZR, Banisadr A, Graves SW, Moxley RA, Mukundan H. Purification and characterization of lipopolysaccharides from six strains of non‐O157 Shiga toxin‐producing Escherichia coli. Journal of Microbiological Methods. 2015;116:1-7
  145. 145. Brooks BW, Lutze‐wallace CL, Blais B, Gauthier M, Deschênes M. Monoclonal antibodies to lipopolysaccharide O antigens of enterohemorrhagic Escherichia coli strains in serogroups O26, O45, O103, O111, O121, and O145. Journal of Food Protection. 2015;78:1252-1258
  146. 146. Westerman RB, He Y, Keen JE, Littledike ET, Kwang J. Production and characterization of monoclonal antibodies specific for the lipopolysaccharide of Escherichia coli O157. Journal of Clinical Microbiology. 1997;35:679-684
  147. 147. Coughlin RT, Bogard WC. Immunoprotective murine monoclonal antibodies specific for the outer‐core polysaccharide and for the O‐antigen of Escherichia coli 0111: B4 lipopolysaccharide (LPS). Journal of Immunology. 1987;139(2):557-561
  148. 148. Rivera‐Betancourt M, Keen JE. Murine monoclonal antibodies against Escherichia coli O4 lipopolysaccharide and H5 flagellin. Journal of Clinical Microbiology. 2001;39:3409-3413
  149. 149. Mutharia LM, Crockford G, Bogard WC, Hancock RE. Monoclonal antibodies specific for Escherichia coli J5 lipopolysaccharide: Cross‐reaction with other Gram‐negative bacterial species. Infection and Immunity. 1984;45:631-636
  150. 150. Bogard WCW, Dunn DLD, Abernethy KK, Kilgarriff CC, Kung PCP. Isolation and characterization of murine monoclonal antibodies specific for gram‐negative bacterial lipopolysaccharide: Association of cross‐genus reactivity with lipid A specificity. Infection and Immunity. 1987;55:899-908
  151. 151. de Jongh‐Leuvenink J, Vreede RW, Marcelis JH, de Vos M, Verhoef J. Detection of antibodies against lipopolysaccharides of Escherichia coli and Salmonella R and S strains by immunoblotting. Infection and Immunity. 1985;50:716-720
  152. 152. Gibb AP, Barclay GR, Poxton IR, di Padova F. Frequencies of lipopolysaccharide core types among clinical isolates of Escherichia coli defined with monoclonal antibodies. Journal of Infectious Diseases. 1992;166:1051-1057
  153. 153. Porat YB, Zan‐Bar I, Ravid A. Quantitative dot‐blot assay for low titer anti‐lipopolysaccharide antibodies in human plasma. Journal of Immunological Methods. 1995;180:213-218
  154. 154. Evans ME, Pollack M, Hardegen NJ, Koles NL, Guelde G, Chia JK. Fluorescence‐activated cell sorter analysis of binding by lipopolysaccharide‐specific monoclonal antibodies to Gram‐negative bacteria. Journal of Infectious Diseases. 1990;162:148-155
  155. 155. Evans ME, Pollack M, Koles NL, Hardegen NJ, Panopoulos D. Lipopolysaccharide heterogeneity in Escherichia coli J5 variants: Analysis by flow cytometry. Journal of Infectious Diseases. 1992;166:803-811
  156. 156. Nelson D. Flow cytometry in bacteriology. Reviews in Medical Microbiology. 1993;4:215-221
  157. 157. Péterfi Z, Kocsis B. Comparison of blocking agents for an elisa for Lps. Journal of Immunoassay. 2000;21:341-354
  158. 158. Gorbet MB, Sefton MV. Endotoxin: The uninvited guest. Biomaterials. 2005;26:6811-6817
  159. 159. Kim S‐E, Su W, Cho M, Lee Y, Choe W‐S. Harnessing aptamers for electrochemical detection of endotoxin. Analytical Biochemistry. Elsevier Inc. 2012;424:12-20
  160. 160. Su W, Lin M, Lee H, Cho M, Choe W‐S, Lee Y. Determination of endotoxin through an aptamer‐based impedance biosensor. Biosensors & Bioelectronics. Elsevier B.V. 2012;32:32-36
  161. 161. Cho M, Chun L, Lin M, Choe W, Nam J, Lee Y. Sensitive electrochemical sensor for detection of lipopolysaccharide on metal complex immobilized gold electrode. Sensors and Actuators B: Chemical. Elsevier B.V. 2012;174:490-494
  162. 162. Ding SJ, Chang BW, Wu CC, Chen CJ. A new method for detection of endotoxin on polymyxin B‐immobilized gold electrodes. Electrochemistry Communications. 2007;9:1206-1211
  163. 163. Zuo M‐Y, Chen L‐J, Jiang H, Tan L, Luo Z‐F, Wang Y‐M. Detecting endotoxin with a flow cytometry‐based magnetic aptasensor. Analytical Biochemistry. 2014;466:38-43
  164. 164. Lan M, Wu J, Liu W, Zhang W, Ge J, Zhang H, et al. Copolythiophene‐derived colorimetric and fluorometric sensor for visually supersensitive determination of lipopolysaccharide. Journal of the American Chemical Society. 2012;134:6685-6694
  165. 165. Mohammed AH, McCallus DE, Norcross NL. Development and evaluation of an enzyme‐linked immunosorbent assay for endotoxin in milk. Veterinary Microbiology. 1988;18:27-39
  166. 166. DeMarco DR, Lim DV. Detection of Escherichia coli O157: H7 in 10‐and 25‐gram ground beef samples with an evanescent‐wave biosensor with silica and polystyrene waveguides. Journal of Food Protection. 2002;65:596-602
  167. 167. Lamoureux L, Adams P, Banisadr A. An optical biosensor for detection of pathogen biomarkers from Shiga toxin‐producing Escherichia coli in ground beef samples. Proceedings of SPIE. 2015; 931004
  168. 168. Mukundan H, Price DN, Goertz M, Parthasarathi R, Montaño GA, Kumar S, et al. Understanding the interaction of Lipoarabinomannan with membrane mimetic architectures. Tuberculosis. Elsevier Ltd; 2012;92:38-47
  169. 169. Bouafsoun A, Othmane A, Jaffrezic‐Renault N. Impedance endothelial cell biosensor for lipopolysaccharide detection. Materials Science and Engineering: C. 2008;28:653-661
  170. 170. Inoue KY, Yasukawa T, Shiku H, Matsue T. Cell‐based electrochemical assay for endotoxin using a secreted alkaline phosphatase reporter system. Electrochemistry. 2008;76:525-528
  171. 171. Unger RE, Peters K, Sartoris A, Freese C, Kirkpatrick CJ. Human endothelial cell‐based assay for endotoxin as sensitive as the conventional Limulus Amebocyte Lysate assay. Biomaterials. 2014;35:3180-3187
  172. 172. Rahman MSA, Mukhopadhyay SC, Yu P‐L, Goicoechea J, Matias IR, Gooneratne CP, et al. Detection of bacterial endotoxin in food: New planar interdigital sensors based approach. Journal of Food Engineering. Elsevier Ltd. 2013;114:346-360
  173. 173. Ganesh V, Bodewits K, Bartholdson SJ, Natale D, Campopiano DJ, Mareque‐Rivas JC. Effective binding and sensing of lipopolysaccharide: Combining complementary pattern recognition receptors. Angewandte Chemie International Edition. 2009;48:356-360
  174. 174. Beveridge TJ, Koval SF. Binding of metals to cell envelopes of Escherichia coli K‐12. Applied and Environmental Microbiology. 1981;42(2):325-335
  175. 175. Langley S, Beveridge TJ. Effect of O‐side‐chain‐lipopolysaccharide chemistry on metal binding. Applied and Environmental Microbiology. 1999;65:489-498
  176. 176. Burkhardt M, LopezAcosta A, Reiter K, Lopez V, Lees A. Purification of soluble CD14 fusion proteins and use in an electrochemiluminescent assay for lipopolysaccharide binding. Protein Expression and Purification. 2007;51:96-101
  177. 177. Wurfel MM, Wright SD. Lipopolysaccharide‐binding protein and soluble CD14 transfer lipopolysaccharide to phospholipid bilayers: Preferential interaction with particular classes of lipid. Journal of Immunology. 1997;158:3925-3934
  178. 178. Fitzgerald KA, Rowe DC, Golenbock DT. Endotoxin recognition and signal transduction by the TLR4/MD2‐complex. Microbes and Infection. 2004;6:1361-1367
  179. 179. Yu B, Hailman E, Wright SD. Lipopolysaccharide binding protein and soluble CD14 catalyze exchange of phospholipids. Journal of Clinical Investigation. 1997;99:315-324
  180. 180. Yeo TY, Choi JS, Lee BK, Kim BS, Yoon HI, Lee HY, et al. Electrochemical endotoxin sensors based on TLR4/MD‐2 complexes immobilized on gold electrodes. Biosensors & Bioelectronics. Elsevier B.V. 2011;28:139-145
  181. 181. Limbut W, Hedström M, Thavarungkul P, Kanatharana P, Mattiasson B. Capacitive biosensor for detection of endotoxin. Analytical and Bioanalytical Chemistry. 2007;389:517-525
  182. 182. Anderson AS, Dattelbaum AM, Montaño GA, Price DN, Schmidt JG, Martinez JS,et al. Functional PEG‐modified thin films for biological detection. Langmuir. 2008;24:2240-2247
  183. 183. Kwon SJ, Kim E, Yang H, Kwak J. An electrochemical immunosensor using ferrocenyl‐tethered dendrimer. Analyst. 2006;131:402-406
  184. 184. Senaratne W, Andruzzi L, Ober CK. Self‐Assembled monolayers and polymer brushes in biotechnology: Current applications and future perspectives. Biomacromolecules. 2005;6:2427-2448
  185. 185. Oliveira MDL, Andrade CAS, Correia MTS, Coelho LCBB, Singh PR, Zeng X. Impedimetric biosensor based on self‐assembled hybrid cystein‐gold nanoparticles and CramoLL lectin for bacterial lipopolysaccharide recognition. Journal of Colloid and Interface Science. 2011;362:194-201
  186. 186. Su W, Kim SE, Cho M, Nam JD, Choe WS, Lee Y. Selective detection of endotoxin using an impedance aptasensor with electrochemically deposited gold nanoparticles. Innate Immunity. 2013;19:388-397
  187. 187. Su W, Cho M, Nam J‐D, Choe W‐S, Lee Y. Aptamer‐Assisted gold Nanoparticles/PEDOT platform for ultrasensitive detection of LPS. Electroanalysis. 2013;25:380-386
  188. 188. Bai L, Chai Y, Pu X, Yuan R. A signal‐on electrochemical aptasensor for ultrasensitive detection of endotoxin using three‐way DNA junction‐aided enzymatic recycling and graphene nanohybrid for amplification. Nanoscale. 2014;6:2902-2908
  189. 189. Zuzuarregui A, Souto D, Pérez‐Lorenzo E, Arizti F, Sánchez‐Gómez S, Martínez de Tejada G, et al. Novel integrated and portable endotoxin detection system based on an electrochemical biosensor. Analyst. 2015;140:654-660
  190. 190. Wu J, Zawistowski A, Ehrmann M, Yi T, Schmuck C. Peptide functionalized polydiacetylene liposomes act as a fluorescent Turn‐On sensor for bacterial lipopolysaccharide. Journal of the American Chemical Society. 2011;133:9720-9723
  191. 191. Rangin M, Basu A. Lipopolysaccharide identification with functionalized polydiacetylene liposome sensors. Journal of the American Chemical Society. 2004;126:5038-5039
  192. 192. Johnson BJ, Delehanty JB, Lin B, Ligler FS. Immobilized proanthocyanidins for the capture of bacterial lipopolysaccharides. Analytical Chemistry. 2008;80:2113-2117
  193. 193. Zeng L, Wu J, Dai Q, Liu W, Wang P, Lee C‐S. Sensing of bacterial endotoxin in aqueous solution by supramolecular assembly of pyrene derivative. Organic Letters. 2010;12:4014-4017
  194. 194. Voss S, Fischer R, Jung G, Wiesmüller K‐H, Brock R. A fluorescence‐based synthetic LPS sensor. Journal of the American Chemical Society. 2007;129:554-561
  195. 195. Lim SK, Chen P, Lee FL, Moochhala S, Liedberg B. Peptide‐assembled graphene oxide as a fluorescent turn‐on sensor for lipopolysaccharide (endotoxin) detection. Analytical Chemistry. 2015;87:9408-9412
  196. 196. Thompson M, Blaszykowski C, Sheikh S, Romaschin A. A true theranostic approach to medicine: Towards tandem sensor detection and removal of endotoxin in blood. Biosensors & Bioelectronics. 2015;67:3-10
  197. 197. Harmon P, Cabral‐Lilly D, Reed RA, Maurio FP, Franklin JC, Janoff A. The release and detection of endotoxin from liposomes. Analytical Biochemistry. 1997;250:139-146
  198. 198. Stromberg LR. Differential interactions of lipopolysaccharides with lipid bilayers: Applications for pathogen detection. [Doctoral Dissertation]. 2016. 1-296
  199. 199. Sakamuri RM, Wolfenden MS, Anderson AS, Swanson BI, Schmidt JS, Mukundan H. Novel optical strategies for biodetection. Proceedings of SPIE. 2013. 881209
  200. 200. Veiseh M, Veiseh O, Martin MC, Bertozzi C, Zhang M. Single‐cell‐based sensors and synchrotron FTIR spectroscopy: A hybrid system towards bacterial detection. Biosensors & Bioelectronics. 2007;23:253-260
  201. 201. Shanmugam A, Rajoria S, George AL, Mittelman A, Suriano R, Tiwari RK. Synthetic Toll like receptor‐4 (TLR‐4) agonist peptides as a novel class of adjuvants. PLoS ONE. 2012;7:e30839
  202. 202. Lamping N, Dettmer R, Schröder NW, Pfeil D, Hallatschek W, Burger R, et al. LPS‐binding protein protects mice from septic shock caused by LPS or Gram‐negative bacteria. Journal of Clinical Investigation. 1998;101:2065-2071
  203. 203. Chien JY, Jerng JS, Yu CJ, Yang PC. Low serum level of high‐density lipoprotein cholesterol is a poor prognostic factor for severe sepsis. Critical Care Medicine. 2005;33:1688-1693
  204. 204. Cooperstock MS, Tucker RP, Baublis JV. Possible pathogenic role of endotoxin in Reye’s syndrome. The Lancet. 1975;1:1272-1274
  205. 205. Jay JM. Limulus lysate endotoxin assay as a test of microbial quality of ground Beef. Journal of Applied Bacteriology. 1977;43:99-109
  206. 206. Kimura H. Measurement of endotoxin. II. Comparison of reactivities measured by radioimmunoassay and with the Limulus test. Acta Medica Okayama. 1976;30:257-270
  207. 207. Ong KG, Leland JM, Zeng K, Barrett G, Zourob M, Grimes CA. A rapid highly‐sensitive endotoxin detection system. Biosensors & Bioelectronics. 2006;21:2270-2274
  208. 208. da Silva JSL, Oliveira MDL, de Melo CP, Andrade CAS. Impedimetric sensor of bacterial toxins based on mixed (Concanavalin A)/polyaniline films. Colloids and Surfaces B: Biointerfaces. 2014;117:549-554

Written By

Loreen R. Stromberg, Heather M. Mendez and Harshini Mukundan

Submitted: 29 April 2016 Reviewed: 03 March 2017 Published: 12 July 2017