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Regenerative Core-shell Nanoparticles for Simultaneous Removal and Detection of Endotoxins Puja Prasad, Siddharth Sachan, Sneha Suman, Girish Swayambhu, and Shalini Gupta Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b00978 • Publication Date (Web): 28 May 2018 Downloaded from http://pubs.acs.org on May 28, 2018

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Langmuir

Regenerative

Core-shell

Simultaneous

Removal

Nanoparticles and

Detection

for of

Endotoxins Puja Prasad, Siddharth Sachan, Sneha Suman, Girish Swayambhu and Shalini Gupta* Dept. of Chemical Engineering, Indian Institute of Technology (IIT) Delhi, New Delhi 110016, India

*Corresponding author: [email protected]

Keywords: Core-shell, Endotoxin, Hydrophobic LPS Capture, LPS Detection, Regeneration, Recyclable, Self Assembled Monolayer.

ABSTRACT. Detection and removal of lipopolysaccharides (LPS) from food and pharmaceutical preparations is important for their safe intake and administration to avoid septic shock. We have developed an abiotic system for reversible capture, removal and detection of LPS in aqueous solutions. Our system comprises of long C18 acyl chains tethered to Fe3O4/Au/Fe3O4 nanoflowers (NFs) that act as solid supports during the separation process. The reversible LPS binding is mediated by facile hydrophobic interactions between the C18 chains and the bioactive lipid A component present on the LPS molecule. Various parameters like pH, solvent, sonication time, NF concentration, alkane chain length and density are optimized to achieve a maximum LPS capture efficiency. The NFs can be reused at least three times by simply breaking the NF-LPS complex in the presence of food grade surfactants, making the entire process safe, efficient and scalable. The regenerated particles also serve as colorimetric labels in dot blot bioassays for simple and rapid estimation of the LPS removed.

Introduction Endotoxins or LPS present on the outer membrane of Gram-negative bacteria provide essential organization and stability to the cell.1 They have, however, undesirable proinflammatory immunogenic effects on humans. Inside the blood, LPS can trigger a systemic 1

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immune response that may lead to endothelial cell destruction, shock, multiorgan dysfunction and death in immune-compromised individuals.2 They are also major contaminants of commercially

available

proteins,

biologically

active

components,

parenteral

drugs,

pharmaceutical products and food items. Therefore, there is a great importance to remove as well as detect the presence of endotoxins in a variety of real systems. The current US Food and Drug Administration (FDA) approved limit of LPS in pharmaceutical drugs is 5 endotoxin units (EU) per kg body weight per hour to ensure the drug’s safe administration via intravenous injection.3,4 To maintain this threshold is a great challenge for the consumer industry. Endotoxin is presently removed by exploiting the electrostatic interactions between the negatively charged LPS and positively charged ligands using methods such as ion exchange chromatography and cationic immobilized adsorbents (histamine, poly-L-lysine etc.).5,6 There also exists a selective LPS removal approach using a polymyxin B sulfate-coated column which is in commercial use, however, the neuro- and nephrotoxicity associated with this antibiotic molecule upon leaching makes it less attractive for medical use.7,8 In addition to removal, monitoring the amount of endotoxin removed is also a costly and time consuming affair. The most popular in vitro test for endotoxin detection is the limulus amoebocyte lysate (LAL) assay which requires expensive reagents, costly equipments (incubators, photometers etc.) and dedicated software. Recent literature also reports the inability of LAL assay to detect LPS because of a “masking effect” caused by detergents mainly in buffer known as “low endotoxin recovery”.9,10 In this study, we demonstrate a facile approach for simultaneous removal and detection of endotoxin in aqueous solvents using hydrophobic interactions. A LPS molecule consists of a hydrophobic component known as lipid A that is mainly responsible for the bioactivity of the endotoxin molecule. This lipid A region consists of six long acyl chains that form a tight hydrophobic packing with the acyl chains of the glycolipids present on the outer membrane of the bacteria.11 The fact that existence of such interactions in bacteria can be mimicked into synthetic analogues motivated us to develop a hybrid nanoparticle (NP) system in which the long acyl chains were incorporated to leverage hydrophobic interactions for entrapping LPS (Figure 1). A few studies in the literature have employed this approach using quantum dots and magnetic NPs (MNPs) to investigate the potent LPS immunomodulatory activity or LPS/TLR4 signaling.12,13 NIPAm-based copolymer beads having C4-C8 hydrophobic groups have also been

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demonstrated to show affinity for molecules having lipid-like domains such as LPS.14 Further, previous work done in our lab has shown that LPS can be immobilized over C18 alkane silanized glass slides via hydrophobic interactions.15 Inspired by these results, we envisioned an extension of this strategy by using hybrid Fe₃O4/Au/Fe3O4 core-shell NPs (also called nanoflowers or NFs) coated with self-assembled monolayers (SAMs) of C18 thiolates. C18-NFs were synthesized to serve as an alternative to biological affinity ligands by hydrophobically entrapping LPS from buffer. These systems are relatively less explored in the literature for any application. The reason for synthesizing gold coated MNPs was that gold eases the surface chemistry for ligand functionalization whereas, Fe3O4 core helps in easy removal of the NF-LPS complexes from suspension, circumventing complicated preprocessing and tedious downstream processes like filtration and centrifugation. Also, Wang and co-workers have shown that NFs not only retain surface plasmon properties but also show 170% increase in magnetic saturation and 23% better conjugation efficacy than only the core-shell due to synergistic and cooperative effects of the layer-by-layer adsorbed NPs.16 All these features make NFs ideal candidates as solid supports for reversible capture and removal of LPS molecules in solution. In addition, the system can be regenerated by breaking the NF-LPS complexes which makes the entire process more efficient and scalable not only for LPS removal but also for their detection. To demonstrate this principle, a simple dot blot bioassay was developed in which the removed LPS was entrapped over a polylysine-coated glass substrate and tagged with regenerated NFs in order to generate a visual response for the naked eyes. In this way, alkane thiol–NFs could be used as versatile probes for endotoxin management.

Experimental Section Materials Purified LPS-AlexaFluor@488 Conjugates (M.W. 10 kDa) extracted from Escherichia coli (E.coli) 055:B5 (Molecular probes, Life Technology); Gold(III) chloride trihydrate (HAuCl4), 1Decanethiol (C10), 1-Octadecanethiol (C18), BSA, Tween 20, silver enhancement kit, Poly-Llysine (PL) (M.W. 30 – 70 kDa) (Sigma-Aldrich); Ellman’s reagent, tetramethylammonium hydroxide (TMAOH), ethylenediaminetetraacetic acid (EDTA), sodium citrate dihydrate, Sodium dodecyl sulfate (SDS) (SRL, India). All endotoxin solutions were prepared in pyrogenfree (PF) water (< 0.001 EU) purchased from GmbH, Germany. All the chemicals were of

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analytical grade and used without further purification. Strict safety guidelines were followed while handling LPS in the laboratory due to its known pyrogenicity and toxicity. Black nunc 96 well microtitre plates were used for recording all fluorescence intensity of Alexa labeled dye.

Methods Synthesis of Fe3O4 MNPs. The cores were prepared by a co-precipitation method.17 Briefly, 3.12 g of FeCl3 was mixed with 2.00 g FeCl2·4H2O (2:1 molar ratio) in 25 mL of degassed and deionized (DI) water in presence of 0.85 mL of 12 N HCl. The above solution was added dropwise to 250 mL of 1.5 N NaOH at 80 °C under vigorous stirring. The Fe3O4 NPs obtained were then separated by a magnetic field and washed with DI water multiple times to finally achieve a clear supernatant. The NPs were finally redispersed in 100 mL of DI water.

Synthesis of Fe3O4/Au/Fe3O4 NFs. 10 ml of the above Fe3O4 NPs and 3 ml of 0.2 N NH2OH·HCl were diluted with 150 ml of 0.01 M TMAOH and heated to 80 °C in a glass bottle. 80 ml of 0.2 wt % HAuCl4 in water was then added dropwise to the above solution under vigorous stirring. A color change from black to reddish brown indicated the reduction of gold salt by hydroxylamine. Further, 200 ml of 15 mM sodium citrate was added incrementally within 2 h to the above reaction mixture. The color of the solution intensified and turned to brick red color. The mixture was stirred for another 3 h after this step. The entire process was carried out at 80 °C. The prepared NFs were then separated via magnetic decantation and washed twice with DI water and ethanol. A further acid wash was performed using 2 N HCl for 1.5 h in order to remove any uncoated Fe3O4 NPs.18 Surface modification of NFs with C18 alkane thiols. The C18-NFs were prepared by reacting 1 mg/ml of NFs in ethanol with C18 alkane thiols.19 The mixture was kept in a rotatory mixer (Scigenics Biotech Orbitek) overnight at room temperature at 50 rpm. The C18-NFs were separated via magnetic decantation and washed twice with ethanol (Figure S3A). The extent of C18 conjugation was quantified using Ellman’s reagent with a slight modification to the approach reported in literature.20,21 During decantation, a 500 µL aliquot was taken from the C18-NF supernatant and mixed with 500 µL of 50 mM tris-HCl buffer (pH 7.8). To this, 100 µL of Ellman’s reagent (4 mg of reagent in 1 mL of buffer) was further added. The whole mixture

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was kept on a rotary shaker for 35 min at 50 rpm. The unbound alkane thiols were quantified using a calibration curve prepared from known concentrations of alkane thiols (Fig. S3B).

NP characterization. The NPs were characterized by various techniques like UV-visible spectroscopy (Shimadzu UV-2600), dynamic light scattering (DLS) and zeta potential measurements (Malvern NanoZS90 Zetasizer), transmission electron microscopy (TEM) and high resolution TEM (HRTEM) (Tecnai G2), energy-dispersive X-ray spectroscopy (EDX) (EDAX AMETEK Materials Analysis Division), X-ray diffraction (XRD) (Rigaku Corporation, MiniFlex600) and total reflection-Fourier transform infrared (FTIR) spectroscopy (Nicolet iS50). The magnetic measurements are performed using a vibrating sample magnetometer (VSM) mode in a physical properties measurements system (PPMS EVERCOOL- II) from Quantum design.

Protocol for LPS capturing. All capturing experiments were performed with alexa fluor labeled fluorescent LPS (λex = 490 nm and λem = 525 nm) in 50 mM tris-HCl buffer:ethanol (8:2 v/v) in the presence of 4 mM EDTA. For this, 100 µg of C18-NFs were mixed with 200 ng of alexa-LPS in 1 mL of buffer:ethanol and the suspension was sonicated for 30 min followed by mild shaking for 90 min. The LPS immobilized over C18-NFs was removed with a magnet for 30 min and visualized using fluorescence microscopy (Olympus BX53F optical microscope fitted with an Orca Flash 4.0 CMOS camera from Hamamatsu) after resuspension in water. The fluorescence image was taken at 100X magnification using an oil immersion objective and an FITC filter (450-480 nm excitation and 515 nm emission). The remaining unbound LPS in the supernatant was quantified as follows (eq. 1) using fluorescence spectroscopy (SpectraMax i3x) in a microtitre plate, % LPS captured by NPs = 1 −

Fluorescence of LPS in the supernatant × 100 (!). Fluorescence of LPS without NPs

For this, a calibration curve for alexa-LPS was prepared in the same 8:2 buffer-ethanol containing EDTA (Figure S4A).

NP regeneration. The alexa-LPS-bound C18-NFs, isolated in the previous step, were resuspended in buffer:ethanol and regenerated in the presence of different concentrations of BSA

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or Tween 20. For this, the mixture was sonicated for 30 min and resubjected to 30 min of magnetization. The regeneration efficiency was again quantified from the supernatant collected, using the fluorescence spectroscopy calibration curve for the best optimized Tween 20 and BSA concentration obtained (Figure S6 (A, B)). The regenerated NFs were washed twice with 500 µl of buffer and further reused for the next cycle of LPS capturing. This whole process was repeated for multiple cycles. It must be noted that in each cycle, fresh 200 ng of LPS was added. The NFs regenerated in each cycle were calculated using following equation (eq. 2),

% NFs regenerated =

Fluorescence of LPS in the supernatant × 100 (%). LPS captured by nanoflowers in that cycle

Bioassay procedure for LPS detection: Microscopic glass slides were cleaned with DI water, acetone and isopropanol alcohol. They were then dried under nitrogen stream and given a plasma treatment for 1 min to activate the glass surface. Microchambers to carry out the assay were prepared by cutting a rectangular piece of Parafilm®M and punching holes in it of 5.5 mm diameter (Figure S7A). The Parafilm sheet was then pressed over the glass slide and heated at 60 °C on a hot plate for 30 to 60 s. 10 µL of 50 µg/mL PL was drop casted into the microchamber and incubated for 1 h at 37 °C in a humid chamber. Excess PL was washed with ultrapure DI water. Next, 10 µL of LPS-spiked sample was inserted and incubated for 1 h. It is important to note that each time prior to insertion, the LPS sample was sonicated for 10 min to break the micelles and convert them into monomers. The excess LPS was then washed twice with PF water. To remove any non-specific interaction, the spots were also treated with BSA blocking agent. Next, the LPS immobilized over glass slide were detected using C18-NFs. The unbound NFs were washed away with copious amounts of ultrapure DI water, and the slide was dried at room temperature. As the spot intensities formed by the gold NFs were difficult to visualize by the naked eye, a quick silver enhancement step was performed at the end of the bioassay to amplify the spot’s output signal intensity by manifold.

Results and Discussion Preparation and characterization of capture probes. The core-shell NPs were prepared in two steps (Figure 2). First, the Fe3O4 cores were synthesized via co-precipitation of FeCl3 and FeCl2

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in an alkaline medium and stabilized by tetramethylammonium hydroxide (TMAOH). Second, a gold salt was reduced on top of these MNPs using hydroxylamine in order to yield a shell. The complete reduction was facilitated by the addition of sodium citrate. TEM characterization showed that the size of Fe3O4 NPs in our system was ~14 nm (Figure 3B) and the gold coated Fe3O4 were ~100 nm (Figure 3C). The gold coated Fe3O4 NPs appeared visibly darker in TEM which was a hallmark of Au domain formation since gold has a higher electron density than iron oxide.22 The images also displayed a unique flower like morphology wherein, the gold-coated core-shells were decorated over their entire surface with NP petals of size ~14 nm. Since the size and intensity of the petals was consistent with that of the Fe3O4 NPs, it led us to believe that the core-shells were actually Fe3O4/Au/Fe3O4 NFs.23 The hydrodynamic radius of the NFs was measured by DLS to be approx. 160 nm which was in agreement with the TEM results (Fig. S1C).

A range of techniques were used to further confirm the formation of Fe3O4/Au/Fe3O4 NFs. HRTEM showed that the spacing of the lattice fringes in the petals was 0.253 nm which corresponded to the (311) plane of Fe3O4 (Figure 3D).24 Similarly, the spacing of the shell lattice fringe appeared at 0.235 nm matching that of the (111) plane of gold.25 The XRD data of Fe3O4 (core) appeared at 35.42°, 43.07°, 57.07°, 62.6°, corresponding to the standard lattice planes of Fe3O4 at (311), (400), (511), (440), respectively (Figure S1A), whereas, the XRD data of NFs showed additional lattice planes of (111), (200), (220) and (311) for gold (along with the standard planes of Fe3O4).26 The UV–visible spectra of Fe3O4 also showed a shoulder-like band at 474 nm (Figure 3A(a)) but exhibited a characteristic peak at 600 nm for NFs (Figure 3A(c)) which confirmed the coating of Au over Fe3O4.27 Finally, a zeta potential study gave us an insight into the possible mechanism of NF formation. The zeta potential of TMAOH-Fe3O4 NPs was found to be negative which attracts Au3+ ions to form Fe3O4-Au3+ hybrid complexes. The hybrid complexes serve as seed to further attract negatively charged Fe3O4 NPs (Figure S1B).28 The Au3+ ions were reduced by hydroxylamine on the surface of Fe3O4 NPs to form Fe3O4/Au/Fe3O4 NFs. The Au coating was completed by further reduction of gold by dropwise addition of sodium citrate.29

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The purification of NFs was initially tried by centrifugation at different rpms to remove the uncoated Fe3O4 NPs but it failed. So, the removal of uncoated Fe3O4 NPs was carried out by acid wash using 2 N HCl. The UV-visible spectra were recorded in order to optimize the washing time period between 30 to 120 min (Figure S2E). It was observed that the characteristic plasmonic peak of gold became prominent and narrower as the acid washing time was increased and got saturated after 90 min. This was also confirmed by TEM imaging (Figure S2A, S2B). Thus, 90 min was set as the optimal time period for removing uncoated magnetite NPs. At higher acid wash time of 6 h, the NFs aggregated irreversibly (Figure S2C) and were difficult to resuspend in water suggesting the complete dissolution of Fe3O4, and thus, poor stability and dispersibility of the core-shells in the absence of any stabilizing agent. EDX performed on the core-shells showed that they contained both iron and gold elements (Figure S2D) proving that the iron oxide within the cores remained intact after this acid wash procedure.30 The NFs were surface modified with SAMs of C18 alkane thiols using simple Au-thiol chemistry. This led to the decrease in their zeta potential value from -28.1 mV to -13.6 mV due to charge shielding by the carbon chains (Figure S1B). The UV-visible spectra of C18-NFs showed a 10 nm blue shift indicating more dispersibility compared to NFs alone (Figure 3A(d)).31 The FTIR data showed asymmetric (υa) and symmetric (υs) stretching vibrations of the CH2 unit in C18 at 2925 and 2851 cm-1, respectively, further confirming the formation of C18NFs (Figure 3F(d)).32 Quantification of the alkane density using Ellman’s reagent showed maximum value when 125 µM alkane thiol was used (Figure S3C). DLS also showed increase in the particle size (180 nm) after conjugation (Figure S1C). The magnetization curves obtained for core and NFs illustrated that the NPs exhibited a typical superparamagnetic behaviour due to no hysteresis, remanence, and coercivity. These properties are essential for applications where magnetic isolation is desired. The saturation magnetization of Fe3O4 and Fe3O4/Au/Fe3O4 NPs was found to be 60 and 30 emu/g, respectively (Figure 3E(a, c)). In this way, the presence of a diamagnetic gold shell (required for easy surface chemical modification) did not drastically reduce the saturation magnetization of NPs and our Au/Fe3O4 core-shell hybrid structures still showed good magnetic potential due to the presence of the Fe3O4 NP coating.23 The magnetization of SAM-C18-NF was found to be 10 emu/g (Fig. 3E(d)). This was sufficient to pull the particles under a magnetic field.

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Endotoxin capture using C18-NFs. Once our capture probes were ready, we performed endotoxin capture via hydrophobic interaction in 3 steps: sonication, shaking and magnetization (Figure 4A). All studies were performed with green fluorescent LPS in order to quantify our results. Since LPS tends to form micelles in aqueous solution, this can quench their fluorescence intensity bringing error to the end result. Therefore, the duration of physical intervention (sonication and/or shaking) and the choice of solvent were extremely important. From the literature, we found 50 mM tris-HCl buffer to be the ideal solvent for dispersing LPS.13 To this, we further added SDS or EDTA to ensure that the LPS micelles were fully broken into monomers.33,34 This led to a 1.5% increase in fluorescence intensity in the presence of EDTA and 1.25% with SDS (in comparison to buffer alone) (Figure S4B). The LPS capturing efficiency achieved was, however, only 16%. To further enhance the performance of our system, we next added ethanol to the EDTA-buffer mix. This led to a significant rise in the capture efficiency (≥ 50%) suggesting that a minimum organic solvent may be required for alkane thiol chains to interact hydrophobically with LPS (Figure 4B). The sonication and shaking time were optimized next as they not only help in disrupting the micellar aggregates but also aid in improved interaction between the LPS and alkyl chains. The sonication time was optimized by keeping the shaking time fixed at 90 min. Shaking alone resulted in ~25% of LPS removal whereas, 30 min of sonication followed by 90 min of shaking resulted in ~55% of LPS capture (Figure 4C). When shaking time was optimized keeping sonication time fixed at 30 min, the maximum capture happened around 90 min of shaking (Figure 4D). Further increase in shaking time had no discernable effect on the capturing efficiency. Therefore, an optimized sequence of 30 min sonication, 90 min shaking and 30 min magnetization protocol was fixed for LPS capturing. At these conditions, the LPS captured by the C18-NFs was clearly visible under an optical microscope (Figure S5). DLS also showed a significant increase in size of the LPS-immobilized NFs (Figure S1C). There are many other parameters that can affect the extent of LPS removal. For instance, the exposed surface area of the hydrophobic tails greatly impacts the hydrophobic interactions. Indeed, when we replaced the C18 alkyl chains with the shorter C10 alkane thiols, the LPS capturing efficiency dropped to as much as ~38% (Figure 5A). Similarly, the alkane thiol ligand density over the NPs also significantly influenced the final outcome. As the concentration of C18 alkane thiols was doubled (for a fixed number of NFs), the %LPS capture first jumped by more

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than twice and then gradually declined (Figure 5B). This non-monotonic behavior was not unexpected as too many tethered chains on the NP surface may lead to local over-crowding and lesser penetration of the LPS chains. The effect of C18-NF concentration was also counterintuitively quite similar (Figure 5C), i.e., increasing the NP concentration did not necessarily lead to more LPS binding. We believe this is due to the fact that NFs start interacting among themselves due to particle-particle chain intercalation leaving less number of sites available for LPS capture (Figure 5C-inset). A good proof of the interactions being driven solely by the hydrophobic groups, and not electrostatic moieties, is the fact that a change in pH in the broad range of 5 to 9 did not at all influence our final results (Figure 5D). This is an additional advantage of our system as it can be applied to a wide range of real analytes. A comparison study had been made in Table S1 comparing endotoxin capturing efficacy of C18-NFs with other system reported in literature.

Regeneration of C18-NFs. NP regeneration is a crucial step for creating an efficient, scalable, and economical system. Several recyclable MNP-based systems have been used in biomedical research using procedures like direct reuse, washing or chemical treatment and high temperature calcinations.35 We tried several different approaches for removing the bound LPS from the C18NF surface including (i) phase separation using hexane/buffer or chloroform/buffer, (ii) vigorous vortexing in buffer:EtOH for up to 10 min, and (iii) sonication for 30 min, but all strategies failed to regenerate the NPs. The highly stable hydrophobic interactions between the LPS and NPs were finally broken in the presence of BSA or Tween 20. Tween 20 (and in general any surfactant above its CMC) is known to facilitate LPS removal by forming micelles (Figure 6A) whereas, BSA acts as a competitive binder of LPS (Figure 6B).36 Thus, addition of these two reagents led to a concentration-dependent release of LPS, with the highest removal efficiency achieved as ~55 ng with 1% w/v BSA and ~37 ng with Tween 20 (Figure 6C). Since particle regeneration was not the only aspect in our study, we also wanted to reuse the particles, the experimental protocol was repeated for up to 5 cycles. The results interestingly showed that in spite of the fact that more LPS was removed by BSA than Tween in the 1st cycle, only 16% LPS was captured with BSA in the 2nd cycle, whereas, Tween gave a steady efficacy of ~50% up to 3 cycles without loss in activity (Figure 6D, S6C). Thus, the overall removal capability of our system in terms of NP capture, regeneration and reusability was found to be better with Tween

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20. A possible reason of this may be that BSA replaces LPS from NF-LPS complex in the first cycle showing good regeneration but in 2nd cycle, LPS is unable to replace BSA from NF-BSA complex. Thus, no LPS capturing was observed in the second cycle whereas, Tween 20 removes LPS from NF-LPS complex by withdrawing endotoxin molecules into its micellar structure through non-polar interactions of alkyl chains of lipid A with the surfactant's tail groups.37 This would make the sites on the NFs free to rebind with LPS in the next cycle.

Detection of LPS with C18-NFs: Finally, a microscopic glass slide dot blot bioassay was developed for the detection of removed LPS using the regenerated C18-NFs. For this, the glass slides were cleaned and treated with plasma to activate their surface with a negative charge. Poly-L-lysine (PL) dissolved in tris-HCl buffer of pH 8 to protonate the amines and imparts positive charge to the surface was then immobilized over the glass slides by electrostatic attraction (Figure 7A). Next, LPS was trapped over the PL in such a way that the hydrophobic acyl groups of LPS were exposed on the surface. To block the surface against any non-specific adsorption in the subsequent steps, various different blocking agents were tested like Tween 20, skimmed milk powder, PEG and BSA. Among them, 2% w/v BSA gave the most promising results (Figure S7C). Finally, the LPS molecules were tagged with reporter C18–NFs via facile hydrophobic interactions between the acyl groups of lipid A chains in LPS and long C18 chains conjugated over the NFs. Throughout this time, the NFs were suspended in DMSO as it is a polar aprotic solvent that provides a better environment for hydrophobic interactions than only buffer (Figure S7B). The intensity of the obtained spots was amplified by adding a simple silver enhancement step in the end to be able to visualize the signal by bare eyes. The silver ions reduce on NFs forming a thick black layer wherever LPS was present yielding sub ng/mL sensitivity. The lower limit of detection (LOD) was 1 ng/mL (Figure 7B). Negative controls in comparison gave clear spots.

Conclusions We have developed a core-shell MNP-based hybrid system of approximately 100 nm size which is capable of removing and detecting molecules with lipid domains such as LPS. The NPs were covered with long C18 acyl chains to remove LPS with ~55% capture efficiency. Reducing the carbon chain length lowered the capture efficacy, whereas, change in pH had no effect on the

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results indicating that the interactions were truly hydrophobically driven. There was an optimum alkane thiol density and NF concentration found at which there was maximum LPS entrapment. We believe higher concentrations result in intra- or interparticle C18 chain intercalation, hindering capture. A sequential protocol of 30 min sonication, 90 min shaking and 30 min magnetization in an EDTA-ethanol-tris-HCl buffer mix yielded the best separation. BSA was established to be the most suitable reagent for maximum LPS recovery (from the C18-NFs) but Tween 20 was overall more efficient as it allowed reversible capture of LPS without loss in activity up to three cycles. The regenerated particles were further used as labels in a dot blot bioassay to report the extent of endotoxin removed per cycle. The lower LOD of this assay was 1 ng/ml. Till date, various biological affinity ligands such as peptides or antibiotics have been explored for detection, capture and tracking of lipid or lipid-like biomolecules. Despite this, difficulty in discovery, development and high reagent cost of biological ligands has greatly hindered their largescale production. Our system offers a new alternative, cost-effective, reusable and scalable platform for efficient LPS management.

Supporting Information: Figure S1 to Figure S7: Particle characterization, Effect of acid wash on nanoparticles Quantification of alkane thiol loading using Ellman’s reagent, Calibration curve of LPS in buffer:ethanol (8:2), Optical micrographs of C18-NFs showing LPS capturing, Fluorescence calibration charts in presence of Tween 20 and BSA in presence of varying amounts of LPS. Regeneration cycle of LPS-bound C18-NFs in the presence of BSA, Optimization of solvent for suspending C18-NFs and Optimization of BSA concentration as blocking agent showed 2% BSA. Table S1: Comparison between C18-NFs and Existing Techniques for Endotoxin removal. ORCID id: Shalini Gupta: 0000-0003-1382-0254

Acknowledgment: We thank Society for Research and Initiatives for Sustainable Technologies and Institutions (SRISTI) for awarding the GYTI research grant to carry out this work. PP acknowledges SERB-DST for funding (NPDF/2016/000087). Authors thank Central Research Facility at IITD for TEM and EDX, Nanoscale Research Facility at IITD for FTIR, Chemical

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Table for Contents/Graphical Abstract

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Main Text Figure

Figure 1. Nanoflowers (NFs) as synthetic analogues of bacterial outer surface

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Figure 2. Schematic diagram showing synthesis of Fe3O4 NPs, Fe3O4/Au/Fe3O4 NFs and C18NFs.

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Figure 3. Full characterization of NPs using (A) UV-Visible spectra, (B & C) TEM, (D) HRTEM, (E) Magnetization curve and (F) FTIR spectroscopy. Here, (a, ̶ ) represents core, (b, ̶ ) NFs before acid wash, (c, ̶ ) NFs after acid wash, and (d, ̶ ) C18-NFs. The TEM micrographs in (C) clearly show Fe3O4 NPs as petals (~ 14 nm). In (F), the frequency peak at 545 cm-1 is for Fe-O IR stretching in magnetite NPs and NFs and C18-NFs. The inset shows the peaks at 2925 and 2851 cm-1 are due to –CH2 alkyl group of alkane thiol in C18-NFs.

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Figure 4. (A) Schematic representation of the steps involved in LPS capture using C18-NFs. (B-D) Optimization of experimental conditions for maximum LPS capture: (B) solvent, (C) sonication time and (D) shaking time. All results are shown for alexa fluor-labeled LPS.

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Figure 5. Effect of system parameters on LPS capture: (A) alkane length, (B) relative particle concentration, (C) unit particle alkane thiol density The inset in (C) indicates phase diagram containing (i) 50 µg/mL (ii) 100 µg/mL and (iii) 200 µg/mL of C18-NFs and (D) pH.

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Figure 6. NP regeneration via competitive binding: (A) LPS recovery from C18-NFs using BSA or (B) Tween 20. (C) Effect of Tween 20 and BSA concentrations on the extent of LPS capture. Data for 2% w/v BSA are not shown as BSA forms a gel at this concentration. (D) Regeneration of LPS-bound C18-NFs in the presence of 1% w/v Tween 20. Here, ‘LPS Captured’ signifies capture efficacy calculated for 200 ng in solution and ‘LPS Released’ implies release efficacy calculated w.r.t LPS bound in the same cycle, and ‘a’, ‘b’, ‘c’ and ‘d’ indicate fresh 200 ng of LPS added before each cycle.

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Figure 7. (A) Schematic diagram of the dot blot bioassay for rapid LPS detection. (B) Digital camera images of silver enhanced spots for LPS detection in spiked aqueous solution containing 1ng/mL LPS. The control contains zero LPS.

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Table of Contents (TOC) 330x233mm (96 x 96 DPI)


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Figure 1. Nanoflowers (NFs) as synthetic analogues of bacterial outer surface 161x126mm (96 x 96 DPI)


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Figure 2. Schematic diagram showing synthesis of Fe3O4 NPs, Fe3O4/Au/Fe3O4 NFs and C18-NFs. 258x77mm (96 x 96 DPI)


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Figure 3. Full characterization of NPs using (A) UV-Visible spectra, (B & C) TEM, (D) HRTEM, (E) Magnetization curve and (F) FTIR spectroscopy. Here, (a, ̶ ) represents core, (b, ̶ ) NFs before acid wash, (c, ̶ ) NFs after acid wash, and (d, ̶ ) C18-NFs. The TEM micrographs in (C) clearly show Fe3O4 NPs as petals (~ 14 nm). In (F), the frequency peak at 545 cm-1 is for Fe-O IR stretching in magnetite NPs and NFs and C18-NFs. The inset shows the peaks at 2925 and 2851 cm-1 are due to –CH2 alkyl group of alkane thiol in C18-NFs. 380x384mm (96 x 96 DPI)


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Figure 4. (A) Schematic representation of the steps involved in LPS capture using C18-NFs. (B-D) Optimization of experimental conditions for maximum LPS capture: (B) solvent, (C) sonication time and (D) shaking time. All results are shown for alexa fluor-labeled LPS. 271x170mm (96 x 96 DPI)


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Figure 5. Effect of system parameters on LPS capture: (A) alkane length, (B) relative particle concentration, (C) unit particle alkane thiol density The inset in (C) indicates phase diagram containing (i) 50 µg/mL (ii) 100 µg/mL and (iii) 200 µg/mL of C18-NFs and (D) pH. 270x198mm (96 x 96 DPI)


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Figure 6. NP regeneration via competitive binding: (A) LPS recovery from C18-NFs using BSA or (B) Tween 20. (C) Effect of Tween 20 and BSA concentrations on the extent of LPS capture. Data for 2% w/v BSA are not shown as BSA forms a gel at this concentration. (D) Regeneration of LPS-bound C18-NFs in the presence of 1% w/v Tween 20. Here, ‘LPS Captured’ signifies capture efficacy calculated for 200 ng in solution and ‘LPS Released’ implies release efficacy calculated w.r.t LPS bound in the same cycle, and ‘a’, ‘b’, ‘c’ and ‘d’ indicate fresh 200 ng of LPS added before each cycle. 388x346mm (96 x 96 DPI)


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Figure 7. (A) Schematic diagram of the dot blot bioassay for rapid LPS detection. (B) Digital camera images of silver enhanced spots for LPS detection in spiked aqueous solution containing 1ng/mL LPS. The control contains zero LPS. 260x186mm (96 x 96 DPI)


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