Development of a Biological Protocol for Endotoxin

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Sep 3, 2014 - Detection Using Quartz Crystal Microbalance (QCM). E. Pérez-Lorenzo ..... the four channel QCM with dissipation (QCM-D E4) (Q-Sense E4).

Development of a Biological Protocol for Endotoxin Detection Using Quartz Crystal Microbalance (QCM) E. Pérez-Lorenzo, A. Zuzuarregui, S. Arana & M. Mujika

Applied Biochemistry and Biotechnology Part A: Enzyme Engineering and Biotechnology ISSN 0273-2289 Volume 174 Number 7 Appl Biochem Biotechnol (2014) 174:2492-2503 DOI 10.1007/s12010-014-1198-2

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Author's personal copy Appl Biochem Biotechnol (2014) 174:2492–2503 DOI 10.1007/s12010-014-1198-2

Development of a Biological Protocol for Endotoxin Detection Using Quartz Crystal Microbalance (QCM) E. Pérez-Lorenzo & A. Zuzuarregui & S. Arana & M. Mujika

Received: 5 May 2014 / Accepted: 22 August 2014 / Published online: 3 September 2014 # Springer Science+Business Media New York 2014

Abstract In this paper, a biological protocol for endotoxin detection has been developed and optimized by quartz crystal microbalance (QCM). The parameters involved in the formation of the self-assembled monolayer (SAM) have been analyzed, and a study of the pH of the ligand buffer has been performed in order to find the best condition for the ligand immobilization and, in consequence, for the endotoxin detection. The detection limit obtained with the characterized biological protocol corresponds to 1.90 μg/ml. The effectiveness of the optimized biological protocol has been analyzed by cyclic voltammetry analysis. Keywords QCM . Endotoxin . Polymyxin B . SAM . Cyclic voltammetry

Introduction Sepsis is a pathology that causes around 1.400 fatalities per day worldwide and has an annual increase of up to 13 %. This serious disease is characterized by a systemic inflammatory response of the organism to an infection [1]. The main triggers of this inflammatory process are related to the presence in blood of endotoxins or lipopolysaccharides (LPS) [2]. These molecules are an essential component of the surface of Gram-negative bacteria, and minimum amounts of LPS can cause serious effects in the organism like fever, tachycardia, and multiorganic failure [3]. Endotoxins are widespread in the environment, and therefore, it is necessary to ensure that medical implants as well as fluids and drugs for parenteral administration do not exceed certain limits of endotoxin. The maximum accepted concentration ranges from 0.2 to 5 endotoxin E. Pérez-Lorenzo (*) : A. Zuzuarregui : S. Arana : M. Mujika CEIT-IK4 and Tecnun, University of Navarra, Paseo de Manuel Lardizábal 15, 20.018, Donostia-San Sebastián, Spain e-mail: [email protected] E. Pérez-Lorenzo : A. Zuzuarregui : S. Arana : M. Mujika CIC Microgune, Goiru kalea 9 Polo de Innovación Garaia, 20500 Arrasate-Mondragón, Spain Present Address: A. Zuzuarregui CIC NanoGUNE Consolider, Tolosa Hiribidea 76, 20018 Donostia-San Sebastian, Gipuzkoa, Spain

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units (EU)/kg (body weight) for intravenous injections to 2.5–20 EU for medical devices, as established by the European and American Pharmacopoeias. Nowadays, there are three validated methods for endotoxin detection: the Rabbit Pyrogen (RTP) test, the Limulus Amebocyte Lysate (LAL) test, and the In vitro Pyrogen Test (IPT). The RTP test consists in observing a change in body temperature when a pyrogen solution is injected in the rabbits [4]. The LAL test is based on the coagulation cascade that occurs as the LPS interacts with the blood cells of the horseshoe crab, Limulus polyphemus [5, 6]. The IPT measures the interleukin-1β secreted by human blood cells in the presence of LPS [7]. Besides these tests, there are commercialized kits for endotoxin detection such as Endosafe® or EndoLISA® among others. The former system uses LAL reagents to test LPS whereas in the latter kit, the endotoxin content is quantified by means of the recombinant factor C (rFC) and a fluorescent substrate [8]. These techniques are based on colorimetric or fluorometric measurements, and in spite of being sensitive, they are complex and expensive to carry out [9]. As an alternative to these systems, biosensors could be potential methods for endotoxin detection. Biosensors allow real-time analysis of biological reactions and can be applied to nearly any target analyte with an adequate biofunctionalization [10]. Thus, they could constitute a sensing alternative to LAL test. In these devices, the biological compound (bioreceptor) is immobilized on the surface of the transducer. When the immobilized molecule and the analyte interact, a signal is generated which is proportional to the amount of the analyte in the sample. Thus, the bioreceptor and its immobilization protocol are crucial to provide the biosensor with sensitivity and selectivity toward the target analyte. In the case of endotoxin bioreceptors, it is known that some polycationic molecules such as the antibiotic polymyxin B (PmB) bind with high affinity to LPS, which is negatively charged at neutral pH [11]. PmB is a cationic peptide that interacts stoichiometrically with LPS [12]. The amphiphilic nature of the molecule due to the simultaneous presence in the molecule of a polycationic ring and a hydrophobic chain allows not only an ionic interaction with the LPS but also a hydrophobic one [13]. Due to these favorable properties, in this work, the use of PmB as LPS bioreceptor has been investigated. In order to promote and to assure a correct immobilization of bioreceptors onto the surface of the sensors and to decrease the nonspecific adsorption of unrelated compounds, the use of self-assembled monolayers (SAMs) is growing due to their effectiveness to capture the specific molecule [14–16]. SAMs are organized structures of organic molecules that allow the efficient and simple immobilization of different compounds used for biological detection [17]. Chemical modification of the SAM can be achieved by using some activating agents, which have been widely employed for SAM formation in biosensor fabrication [18–20]. Once the SAMs are formed, they remain strongly attached to the surface through their terminal group and provide a well-defined and stable interface for ligand immobilization [18]. The purpose of the organic molecules is to prepare the SAM with efficient properties to achieve an optimum immobilization of the ligand. In the work presented, a biological protocol has been optimized for its implementation onto the gold electrode of a biosensor for the endotoxin detection. On one hand, the influence of several parameters involved in the formation of the SAM has been analyzed, namely, the nature, concentration, and incubating period of the alkanethiol considering two acids: the mercaptopropionic acid (MPA) and the mercaptoundecanoic acid (MUA). Moreover, the effectiveness of SAM activators (EDC and NHS) has been studied to maximize the sensitivity for LPS detection. On the other hand, a study of the PmB buffer pH has been fulfilled to analyze the correct immobilization of the ligand for the endotoxin detection. The characterization of PmB-LPS linkage has been carried out using the quartz crystal microbalance (QCM). This detection method is very effective due to the fact that it allows a

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real-time monitoring of the adsorbed analyte mass and it is not necessary to label the sample. Common applications include measurements on proteins, polymers, surfactants, lipids, and cells [21]. The QCM consists on a piezoelectric quartz crystal disk covered with a gold thin film. This is coupled to an oscillatory circuit that applies an alternating electrical field to the crystal. Directly linked to this circuit, there is a frequency counter which monitors the changes in the crystal frequency [22]. In this way, when a material is deposited onto the crystal surface, a decrease in the amplitude and in its resonance frequency occurs [23]. The adsorbed mass on the crystal surface is given by the Sauerbrey equation, in which the frequency variations are related to the mass deposition following in Eq. (1): Δf ¼

−2Δm f 20 pffiffiffiffiffiffiffiffiffiffi A μq ρ q


where ρq is the quartz density, μq is the crystal shear module, f0 is the crystal fundamental frequency, A is the crystal piezoelectrically active geometrical area (defined by the area of the deposited metallic film on the crystal), and Δm and Δf correspond to the mass and system frequency changes, respectively [24]. When the QCM comes in contact with a solution, there is a decrease in frequency that is dependent upon the viscosity and the density of the solution. Kanazawa and Gordon [25] explained the influence of both the density (ρL) and the viscosity ( L) of the liquid using the mathematical expression: Δ f ¼ −fo 3=2 ½ðρL:ηLÞ=ðπ: ρq:μqÞ1=2


where fo ρq μq ρL ηL

Frequency of oscillation of unloaded crystal Density of quartz Shear modulus of quartz Density of the liquid in contact with the electrode, and Viscosity of the liquid in contact with the electrode

The suitable immobilization of PmB and LPS detection has been obtained with a detailed analysis of all components of the SAM, getting a proper biological stack. The effectiveness of this biological stack has been tested by cyclic voltammetry (CV). The aim of this study is to present a novel biological protocol optimized for the selective detection of endotoxin detection. This protocol is based on the PmB used as ligand and not as a neutralizing molecule.

Materials and Methods Reagents The products that have been purchased from Thermo Scientific were 1-ethyl-3-(3dimethylaminopropyl) (EDC) carbodiimide HCl (Ref. 22980) and N-hydroxysuccinimide (NHS) (Ref. 24500), whereas the 3-MPA has been provided by Fluka (Ref. 63768). The PmB sulfate salt and the 11-MUA (Ref. 450561) have been supplied by Sigma-Aldrich, and the LPS has been purified from Escherichia coli (provided by de Department of Microbiology of The University of Navarra).

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Cleaning Protocol of the Gold Disks The disks of QCM have been cleaned using the following protocol: First, they have been introduced in a UV/Ozone Procleaner (Bioforce Nanoscience) for 30 min. Afterward, they have been cleaned for 5 min at 75 °C with a solution composed by H2O, H2O2 30 % (Ref. 95321, Sigma-Aldrich), and NH3 25 % (Ref. 121129.1611, Panreac). Then, the sensor disks have been rinsed with water and dried with N2, and finally subjected to a second UV/ozone cleaning step for 30 min. Immobilization Protocol The immobilization protocol that has been used for the SAM formation consists in dipping the disks in a solution of MPA or MUA prepared in ethanol purity 99.5 % (Ref. 131085.1211, Panreac). In order to activate the thiolated acid layer, the crystals have been incubated first in EDC and then in NHS. Once the SAM has been activated, first it has been ensured that the corresponding baseline with the PmB buffer (barbital; provided by the Department of Microbiology of the University of Navarra) has stabilized. Then, PmB (100 μg/ml dissolved in barbital) has been immobilized for 2 h and washed once the step is finished to eliminate the unspecific bindings. After that, a baseline with pyrogen-free water has been carried out. Then, 100 μg/ml LPS prepared in pyrogen-free water has been flowed and incubated over the surface of the disks for 30 min, and finally, a cleaning solution of pyrogen-free water has been used to avoid unspecific unions. Before being incubated, LPS samples have been thermally cycled three times between −20 and 56 °C. The assays for endotoxin detection imply three main steps prior to the incubation of the sample containing the analyte. These steps allow the formation of a biofunctionalization layer, which is the key for an optimized performance of a biosensor. Formation of the SAM First of all, and regarding the immobilization conditions of the core of the monolayer, parameters such as the nature, concentration, and incubation period of the thiolated acids have been analyzed and optimized by means of fluorescence measurements. These measurements have been carried out with an epifluorescence microscope (Nikon Ti-E inverted microscope) which implements NIS-elements software to quantify the intensity and calculate the percentage of coverage for the analyzed surfaces. For these assays, amine-coated fluorescent microspheres (F2-XC R04-15, Merck) at a concentration of 100 μg/ml have been used. In order to analyze the influence of different SAM parameters in its immobilization capability, two structures have been considered to study the influence of the nature of the alkanethiol, namely, MPA and MUA. For each of these acids, other two parameters have been kept in mind: acid concentration (1, 10, and 50 mM in ethanol 99,5 %) and the incubating period (1, 2, and 4 h). Two measurements have been done after the incubation period of microspheres: The first one has been carried out after 1 h, and then, the samples have been immersed again in the microspheres solution for 18 h more before the second measurement. The substrates used for this purpose included 16 gold circles (100 μm in diameter) deposited by physical vapor deposition onto oxidized silicon. The fluorescent microspheres used as labels have been incubated overnight before the determination of the final surface coverage. Direct immobilization of the microspheres onto the bare gold surface has served as control assays.

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Activation of the SAM Once the best condition for the acid immobilization has been determined, the two-step thiolated acid activation has been subjected to study. For this reason, the experiments of this section consist of analyzing if it is better to incubate EDC and NHS individually or mixed for 30 min or 1 h, in order to check the best way for the incubation of both compounds. As used for the fluorescence assays, EDC 46 mM and NHS 46 mM have been the activators selected. Once the monolayer has been activated, the bioreceptor (PmB) has been immobilized, and then, the LPS dissolved in pyrogen water has been incubated. The LPS mass (ng/mc2) attached to the PmB has been measured. The immobilization protocol of SAM activators, PmB and LPS, has been performed using the four channel QCM with dissipation (QCM-D E4) (Q-Sense E4). Immobilization of the Bioreceptor Finally, the QCM has been used for studying the effect of PmB buffer pH in the immobilization of this bioreceptor onto the previously optimized SAM. Four different values have been selected for the barbital buffer covering the pH range from 2.6 to 8.6. For these assays, the PmB has been incubated for 2 h and the LPS for 30 min. The SAM parameters used have been the following: 2 h of MPA 1 mM, 1 h of EDC 46 mM, and 1 h of NHS 46 mM. The biofunctionalization protocol involves the immobilization of PmB on a fully formed SAM. To check this protocol, CV has been used for testing purified LPS samples of known concentration (1, 10, and 100 μg/ml). For this purpose, ferro/ferricyanide redox couple has been used since its oxidation and reduction peaks are well defined and well known [26]. The CV conditions are the following: at room temperature (23 °C) and in a 25-mM K3Fe(CN)6 solution. The measurements have been carried out in a range of −0.5 to 0.6 V, with a scan rate of 0.1 V/s. The experiments have been monitored with the Autolab Potentiostat PGSTAT 128N using the Nova 1.6 software version (Eco Chemie).

Results and Discussion For the assessment of the obtained results, surface coverage and detected LPS mass have been the parameters selected for fluorescence and QCM measurements, respectively. In order to test the protocol optimized by QCM, CV measurements have also been performed. The following results describe the optimization of each one of the different steps mentioned above. Development of the Biological Protocol As previously mentioned, a method promoting the correct immobilization of bioreceptors involves their covalent linkage to SAMs formed on the gold electrodes. The parameters optimized in this case have been the following ones: the nature of alkanethiol acids (MPA and MUA), the concentration and incubation period of both thiolated acids. Besides, the acid activation step has been studied to promote a more effective LPS detection.

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Study of Alkanethiol Acids: Nature, Concentration, and Incubation Period of Both Thiolated Acids All the samples have been measured twice, one after 1 h of microsphere incubation and then, 18 h later. It has been observed that the surface coverage varies significantly at 19 h; so, the results presented correspond to the second measurement after 19 h of incubation period. The obtained fluorescence images are shown in Fig. 1. These images correspond to the assays with MPA at different concentrations and incubation times. Based on these images, it can be derived that incubating the thiolated acid during 1 h is not enough to maximize the surface coverage because the MPA cannot reach the entire surface with good uniformity, and in consequence, few microspheres are immobilized onto the surface. On the other hand, a 4-h incubation time is too long, and many MPA chains attach untidily to the surface causing an improper binding and the formation of a nonefficient SAM layer. Thus, it can be concluded that the optimum MPA incubation time takes 2 h. Analyzing the influence of the thiolated acid concentration at 2 h, it can be observed that the lower the concentration, the better the SAM performance. A higher surface coverage could be expected as MPA concentration increases. However, it seems that the formation of clusters between the carbon chains happens, inhibiting a proper immobilization of the alkanethiol onto the surface. Moreover, it could happen that the bindings are less effective; so in the cleaning steps, more unspecific bonds are eliminated, and in consequence, the immobilization is more effective with 50 mM than that with 10 mM. These results confirm the conclusion of Touahir

Fig. 1 Fluorescent microsphere assays. Fluorescence images after a 19-h incubation period of FITC-labeled polystyrene microspheres for the assessment of different SAM formation conditions

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et al. [27] who assert that if the thiolated molecule density is high, it can produce the formation of N-acilurea hindering the activation of the monolayer with the NHS activator. This result explains the differences in the surface coverage among the samples incubated with MPA 1 mM and higher concentrations. Regarding these results, it can be concluded that the optimum concentration for MPA corresponds to 1 mM due to the fact that in all cases, the surface coverage at this concentration is higher than that in the rest of conditions. In the samples with MUA (Fig. 2), it is shown that almost all the values are below the control assays. This is owing to the fact that the mercapto consists of three times longer carbon chains than the MPA and their geometrical distribution does not promote the formation of an active SAM layer [28]. Furthermore, the fact that most of the measured values are below the control demonstrates that the SAM has blocked the active surface for microsphere immobilization [29]. In consequence, it can be stated that the MUA is not a proper acid for this purpose. Results have demonstrated that the highest surface coverage corresponds to MPA, independently of the immobilization time of the fluorescent microspheres. Moreover, the surface coverage with MPA is 45 times higher than that with MUA. The molecules of the MUA form a dense layer onto the gold surface avoiding a specific binding between the SAM monolayer and the labeled molecule. These results are in accordance with the experiments that have been carried out by the group of Briand et al. [29]. According to these results, the best SAM performance has been achieved with MPA in a concentration of 1 mM and incubated for 2 h. Thiolated Acid Activation Once the best concentration and the optimum incubation time for the alkanethiol have been obtained, conditions for the chemical activation of the MPA with EDC and NHS have been studied. The characterization has been carried out with the QCM-D, which has allowed an accurate quantification of the adsorbed mass onto the gold sensor (sensitivity in ng/cm2). The biofuntionalization scheme obtained with the experiments that have been analyzed in these assays has the following structure (Fig. 3): The curve shows the change of the frequency related to the mass deposited onto the sensor along the incubation period and the different steps of the immobilization process: the thiolated

Fig. 2 Surface coverage with MUA at different concentrations and incubation periods. Effect of MUA concentration and incubation time (a) and obtained values (b) using fluorescent microspheres

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Fig. 3 Biofunctionalization scheme by QCM. Different biofunctionalization steps (thiolated acid incubation, activation of thiol molecule, bioreceptor immobilization, and analyte detection) by means of QCM

acid incubation, the activation of the acid, the ligand immobilization, and finally the analyte detection. There are experiments that have been made by other authors [30–32] analyzing separately either the concentrations, the incubation period, pH, or temperature employed in the formation of the biofunctionalization layer of a sensor, but only a few analysis has been focused on the influence of SAM activation parameters. The results obtained with the chemical activators mixed and individually at 30 min and 1 h of incubation period are shown in Fig. 4. As it can be observed, the highest amount of LPS has been detected with the activators incubated during 1 h, both mixed and individually. On the other hand, if the obtained results at 1 h are compared, it can be concluded that the best condition occurs with the chemical activators incubated individually. The lower LPS detection in mixed conditions could be due to the fact that SAM activation is a process in two steps, and if both molecules are incubated at the same time, an interaction could happen between EDC and NHS [33]. In consequence, the EDC molecules could defuse before joining the MPA, entailing a minor activation of components. In summary, derived from the analysis of the best parameters for the formation of the SAM, it can be concluded that the optimum activation protocol implies the incubation of MPA 1 mM

Fig. 4 Analysis of the SAM activation. Thiolated acid activation with EDC and NHS mixed and individually at 30 min and 1 h. Quantification of detected LPS mass by QCM

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for 2 h and its activation by consecutively incubating EDC 46 mM and NHS 46 mM for 1 h each. Analysis of PmB Buffer pH With Optimized SAM Once the SAM components have been analyzed, it has been checked if the developed monolayer allows a correct immobilization of the bioreceptor for a more effective LPS detection. With this aim, the influence of the pH of PmB buffer has been studied using the optimized SAM. As control assays and in order to have a reference to compare with the results of the immobilization with SAM, a study of the barbital pH without SAM has been carried out, i.e., PmB has been adsorbed directly onto the gold surface, and then, LPS has been added. The results are shown in the following figure: As it can be observed in Fig. 5, the highest amount of detected LPS corresponds to pH 3.9. This result is corroborated by the isoelectric point of the PmB, which is around 4.0. In this situation, the molecule carries no electrical charge (the negative and positive charges are equal), and it offers the best condition for serving as receptor. This is in accordance with the result obtained in the study of barbital pH without SAM where the highest mass of LPS is also obtained at pH 3.9. Based on the comparison of the LPS/PmB ratio with and without SAM (Fig. 6) and according to the significantly higher ratio obtained in the former case, it can be asserted that the use of SAM is necessary to assure a more effective and correct binding of the ligand, and in this way, more LPS can be adsorbed per ligand unit. Assays of the Optimized Biological Structure by Cyclic Voltammetry In order to quantify the amount of endotoxin bound to PmB, the anodic peak current (Ipa) before and after addition of LPS has been measured (see inset of Fig. 7). When the peptide is added to the activated SAM, there is an increase in the current, due to the conductive properties of the molecule. Then, the addition of LPS results in a change in the current (a drop in the Ipa) likely reflecting the interaction between PmB and the endotoxin. Specifically, the drop in Ipa is the consequence of both the neutralization of the PmB charge by the LPS and the presence of

Fig. 5 LPS immobilization at different PmB buffer pH values. Detection of LPS mass (ng/cm2) at different barbital pH values of 2.6, 3.88, 6.52, and 8.60

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Fig. 6 LPS/PmB ratio. LPS mass adsorbed per immobilized PmB mass unit with optimized SAM and without SAM

insulating groups in the endotoxin, a molecule with large hydrophobic sections. The results achieved using CV are shown in Fig. 7. To demonstrate the specificity of the biosensors, control assays without SAM and PmB have been carried out, incubating different concentrations of LPS solutions directly onto the gold electrodes. Besides this, further control assays with pyrogen-free water instead of LPS have been performed to establish the detection limit of the biofunctionalization stack. As depicted in the figure, the response of the biosensor to endotoxin, expressed by the decrease in Ipa, is concentration dependent in a logarithmic scale and linear over at least three orders of magnitude. The reduced variability of the results allows discriminating between different concentrations of analyzed endotoxin. In Fig. 7, it can also be observed that the ΔIpa

Fig. 7 Quantitative detection of LPS via CV and corresponding control assays. Variation of the anodic peak current for different LPS concentrations (1, 10, and 100 μg/ml). Control assays are the following: incubation of pyrogen-free water onto the optimized biofunctionalization layer and direct incubation of LPS onto the bare gold disk surface. Upper left inset schematic of the anodic peak current variation on a CV voltammogram

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Table 1 Values of the anodic peak current for different LPS concentrations. Variation of Ipa at LPS concentrations of 1, 10, and 100 μg/ml using K3Fe (CN)6 as electrolyte by means of CV

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LPS (μg/ml)

Δlpa (μA)



10 100

22.4±2.6 32.9±3.2

does not show correlation with the LPS concentration in the case of experiments with nonfunctionalized surface. These results show the low affinity of the gold surface, as compared with the functionalization surface, by the LPS. The detection limit, defined as the concentration that produces a sample equal to the target signal plus three times its standard deviation, that has reached at a concentration of 1 μg/ml LPS corresponds to 1.90 μg/ml. The values obtained for the variation of Ipa with the different concentrations of LPS are shown in the following table (Table 1): The decrease of Ipa in the experiments with the whole functionalization structure has a linear relationship with the analyte concentration. These results indicate that the new detection method developed is suitable for LPS detection in the range of concentrations tested.

Conclusions A biological protocol for LPS detection has been developed and optimized aiming at its application for biosensor biofunctionalization. The characterization of the protocol has been made by means of QCM. Several parameters involved in the SAM monolayer have been tested to find the optimum SAM. Moreover, a study of the PmB buffer has been made for analyzing the best pH condition for the PmB immobilization and, in consequence, for the endotoxin detection. As an overall result, it has been demonstrated that the use of SAM is necessary to improve the quality of the ligand immobilization and, in consequence, the amount of detected LPS mass. On the other hand, the uses of MPA for 2 h, EDC 46 mM for 1 h, and NHS 46 mM for 1 h have been selected as the optimum SAM conditions for the LPS detection, and the best pH of PmB buffer corresponds to 3.9. Finally, a CV analysis has confirmed the effectiveness of the optimized biological protocol for endotoxin detection achieving to quantify an LPS concentration of 1 μg/ml. Acknowledgments The authors acknowledge the University of Navarra for the funding that supported this project and the Department of Microbiology of The University of Navarra. We also acknowledge the Secretary of State of Investigation, Development and Innovation of the Ministry of Economy and Competitivity of Spain for funding this research within the framework of the SIMcell Project DPI 2012-38090-C03-D3.

References 1. 2. 3. 4.

Negredo, J., & Real, M. (2012). September, 2012. Chaby, R. (1999). Drug Discovery Today, 4, 209–221. Cohen, J. (2002). Nature, 420, 885–891. Park, C. Y., Yung, S. H., Bak, J. P., Lee, S. S., & Rhee, D. K. (2005). Biologicals, 33, 145–151.

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5. Harmon, P., Cabral-Lilly, D., Reed, R. A., Maurio, F. P., Franklin, J. C., & Janoff, A. (1997). Analytical Biochemistry, 250, 139–146. 6. Ong, K. G., Leland, J. M., Zeng, K., Barrett, G., Zourob, M., & Grimes, C. (2006). Biosensors & Bioelectronics, 21, 2270–2274. 7. Daneshian, M., Wendel, A., Hartung, T., & von Aulock, S. (2008). Journal of Immunological Methods, 336, 64–70. 8. Grallert, H., Leopoldseder, S., Schuett, M., Kurze, P., Buchberger, B. (2011). Nature Methods, 8, 9. Gee, A. P., Sumstad, D., Stanson, J., Watson, P., Proctor, J., Kadidlo, D., Koch, E., Sprague, J., Wood, D., Styers, D., McKenna, D., Gallelli, J., Griffin, D., Read, E. J., Parish, B., & Lindblad, R. (2008). Cytotherapy, 10(4), 427–435. 10. Bashir, R. (2004). Advanced Drug Delivery Reviews, 56, 1565–1586. 11. Brandenburg, K., David, A., Howe, J., Koch, M. H. J., Andrä, J., & Garidel, P. (2005). Biophysical Journal, 88, 1845–1858. 12. Brandenburg, K., Moriyon, I., Arraiza, M. D., Lewark-Yvetot, G., Koch, M. H. J., & Seydel, U. (2002). Thermochimica Acta, 382, 189–198. 13. Morrison, D. C., & Jacobs, D. M. (1976). Immunochemistry, 13, 813–818. 14. Baldrich, E., Laczka, O., Del Campo, F. J., & Muñoz, F. X. (2008). Journal of Immunological Methods, 336, 203–212. 15. Chen, D., & Li, J. (2006). Surface Science Reports, 61, 445–463. 16. Shankaran, D. R., Gobi, K. V., & Miura, N. (2007). Sensors and Actuators B: Chemical, 121, 158–177. 17. Pillay, J., Agboola, B. O., & Ozoemena, K. I. (2009). Electrochemistry Communications, 11, 1292–1296. 18. Ansorena, P., Zuzuarregui, A., Pérez-Lorenzo, E., Mujika, M., & Arana, S. (2011). Sensors and Actuators B: Chemical, 155, 667–672. 19. Jang, L. S., & Keng, H. K. (2008). Biomedical Microdevices, 10, 203–211. 20. Jiayu, W., Xiong, W., Jiping, L., Wensen, L., Ming, X., Linna, L., Jing, X., Haiying, W., & Hongwei, G. (2009). Archives of Virology, 154, 1901–1908. 21. Schirhagl, R., Latif, U., Podlipna, D., Blumenstock, H., & Dickert, F. (2012). Analytical Chemistry, 84, 3908–3913. 22. Damos, F. S., Mendes, R. K., & Kubota, L. T. (2004). Quim Nova, 27, 970–979. 23. Ogi, H. (2013). Proceedings of the Japan Academy. Series B, Physical and Biological Sciences, 89(9), 401– 417. 24. Sauerbray, G. (1959). Zeitschrift für Physik, 155, 206–222. 25. Kanazawa, K. K., & Gordon, J. (1985). Analytical Chemistry, 57, 1770. 26. García, T., Revenga-Parra, M., Añorga, L., Pariente, F., & Lorenzo, E. (2012). Sensors and Actuators B: Chemical, 161(1), 1030–1037. 27. Touahir, L., Allongue, P., Aureau, D., Boukherroub, R., Chazalviel, J. N., Galopin, E., Gouget-Laemmel, A. C., de Villeneuve, C. H., Moraillon, A., Niedziólka-Jönsson, J., Ozanam, F., Andresa, J. S., Sam, S., Solomon, I., & Szunerits, S. (2010). Bioelectrochemistry, 80, 17–25. 28. Arya, S. K., Solanki, P. R., Datta, M., & Malhotra, B. D. (2009). Biosensors and Bioelectronics, 24, 2810– 2817. 29. Briand, E., Salmain, M., Compère, C., & Pradier, C. M. (2006). Colloids and Surfaces B: Biointerfaces, 53, 215–224. 30. Priano, G., Pallarola, D., & Battaglini, F. (2007). Analytical Biochemistry, 362, 108–116. 31. Skottrup, P. D., Nicolaisen, M., & Justesen, A. F. (2008). Biosensors & Bioelectronics, 24(3), 339–348. 32. Yamasaki, R., Kim, J., Jung, H., Lee, H. Y., & Kawai, T. (2006). Biochemical Engineering Journal, 29, 125– 128. 33. Xia, N., Xing, Y., Wang, G., Feng, Q., Chen, Q., & Feng, H. (2013). International Journal of Electrochemical Science, 8, 2459–2467.

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