Antifouling Microparticles To Scavenge ... - ACS Publications

Subscriber access provided by Iowa State University | Library

Article

Antifouling Microparticles to Scavenge Lipopolysaccharide from Human Blood Plasma Mariia Vorobii, Nina Yu. Kostina, Khosrow Rahimi, Silvia Grama, Dominik Söder, Ognen Pop-Georgievski, Adriana Šturcová, Daniel Horák, Oliver Grottke, Smriti Singh, and Cesar Rodriguez-Emmenegger Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.8b01583 • Publication Date (Web): 03 Jan 2019 Downloaded from http://pubs.acs.org on January 4, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Antifouling

Microparticles

to

Scavenge

Lipopolysaccharide from Human Blood Plasma Mariia Vorobii,a Nina Yu. Kostina,a Khosrow Rahimi,a Silvia Grama,b Dominik Söder,a Ognen Pop-Georgievski,b Adriana Sturcova,b Daniel Horak,b Oliver Grottke,c Smriti Singh,a and Cesar Rodriguez-Emmenegger a* a

DWI − Leibniz Institute for Interactive Materials and Institute of Technical and

Macromolecular Chemistry, RWTH Aachen University, Forckenbeckstrasse 50, 52074 Aachen, Germany. b

Institute of Macromolecular Chemistry, Czech Academy of Sciences., Heyrovského nám.

2, 16206 Prague, Czech Republic. c

Department of Anesthesiology, University Hospital RWTH Aachen, Pauwelsstrasse 30,

52074 Aachen, Germany.

KEYWORDS. Photoinduced SET-LRP, microparticles, polymer brushes, polymyxin B, LPS, endotoxins.

ACS Paragon Plus Environment

1

Biomacromolecules 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 30

ABSTRACT

Currently one of the most promising treatments of lipopolysaccharides (LPS)-induced sepsis is based on hemofiltration. Nevertheless, proteins rapidly adsorbed on the artificial surface of membranes which leads to activation of coagulation impairing effective scavenging of the endotoxins. To overcome this challenge we designed polymer-brush-coated microparticles displaying antifouling properties and functionalized them with polymyxin B to specifically scavenge LPS the most common endotoxin. Poly[(N-(2-hydroxypropyl) methacrylamide)-co(carboxybetaine methacrylamide)] brushes were grafted from poly(glycidyl methacrylate) microparticles using photoinduced single-electron transfer living radical polymerization (SETLRP). Notably, only parts-per-million of copper catalyst were necessary to achieve brushes able to repel adsorption of proteins from blood plasma. The open porosity of the particles, accessible to polymerization, enabled to immobilize sufficient polymyxin B to selectively scavenge LPS from blood plasma.


2

Page 3 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

INTRODUCTION

Microorganisms constantly colonize living and nonliving surfaces. Generally, bacteria exist in two type of population: planktonic and sessile known as biofilm.1 Biofilms are a densely packed community of microorganism surrounded with a self-produced matrix of extracellular polymeric substances (EPS).2 EPS mostly consist of polysaccharides, proteins, lipids and nucleic acids. The most important functions of EPS are adhesion to the surfaces, aggregation of bacterial cell, cohesion of biofilm, exchange of genetic information, and creation of protective barrier from specific and nonspecific host defense mechanisms and different antimicrobial agent such as antibiotics.3 All these make treatment of biofilm-related infections very difficult, especially, because of complication in the identification of infection since the routine microbiological tests cannot reveal its presence in standard samples.4 Biofilms formed by Gram-negative bacteria such Escherichia coli, Pseudomonas aeruginosa, Klebsiella pneumoniae and Proteus mirabilis, Grampositive bacteria such Staphylococcus aureus, Staphylococcus epidermidis, Streptococcus viridans and Enterococcus faecalis and yeasts (Candida albicans) pose a special danger to public health due to the colonization of indwelling medical devices and surrounding tissue.5-7 Bacterial biofilm infections on indwelling medical devices and implants (e.g., urinary catheter, peritoneal dialysis catheter, central venous catheters, contact lenses, endotracheal tubes, mechanical heart valves, prosthetic joints, and voice prostheses) may lead to systemic dissemination of the pathogen, tissue destruction, device malfunction, revision surgeries, etc.7 In the most difficult cases it can lead to a life-threatening condition called sepsis.8 Sepsis occurs as a result of uncontrolled activation of the immune system with the release of numerous inflammatory mediators triggered by the present of the pathogenic microorganism or its


3


Page 4 of 30

endotoxins in the blood stream. This can lead to multi–organ failure syndrome, septic shock and death.9-11 Abe et al reported that the level of proinflammatory mediators (C-reactive protein, IL6) were higher in bacteremia induced by Gram-negative bacteria than by Gram-positive ones.12 This can be explained by the presence of lipopolysaccharides (LPS) in their outer membrane wall. The LPS is an endotoxin which causes significant inflammation by binding to multiple receptors such as LPS-binding protein, macrophage receptor, Toll like receptor-4 that leads to activation of intracellular signaling pathways and production of proinflammatory cytokines (TNF-α, IL-1 and IL-6).13-15 The treatment of sepsis is complex and encompasses a manifold of measurements in clinical practice. This include, ameliorating the source of the infection by administering wide spectrum antibiotics, management of inflammation, interfering with the binding of LPS and bioaffinity removal of the endotoxins.16, 11 The use of antibiotics, alone, is usually not enough after the level of endotoxins have increased. Besides, additional killing of bacteria can release even more LPS. Strategies to interfere with the binding of LPS, i.e. those block the mechanism of action, still remain elusive.16 This might be associated with the ability of LPS to self-assemble into micelles. Presumably, this may restrict the targeting of the hydrophobic domain buried in the core of the micellar structure.17, 18 Currently, the most successful strategies for the treatment of sepsis rely on the hemofiltration and affinity capture of remaining LPS19 and inflammatory mediators like cytokines.11 One of the most effective systems is Toraymyxin, an extracorporeal cartridge on which LPS is adsorbed while blood is filtered.20-23 The cartridge consists of polystyrene fibers on which polymyxin B is immobilized.23 Polymyxin B is a polycationic antibiotic that kills Gram-negative bacteria and inactivates endotoxins. The mechanism of inactivation involves the affinity


4

Page 5 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

binding to lipid A, which is the part of LPS associated with its toxicity.24, 25 The binding between polymyxin B and LPS is the result of coulombic and hydrophobic interactions and forms a stable nontoxic complex. However, polymyxin B is not suitable for intravenous use due to its nephrotoxicity and neurotoxicity.26,

27

Thus, polymyxin B must be

immobilized onto insoluble carriers. Various reports deal with development of systems and coatings to (i) detect LPS28, 29 and to (ii) scavenge it.30, 23 Thompson et al introduced a theranostic system based on a bulk acoustic wave sensor and affinity binding of LPS in glass beads coated with a self-assembled monolayer modified with polymyxin B.29 Arguably, the detection limits reached by their sensor are among the best achieved so far. Regretfully, their surface modification was incapable to effectively prevent the fouling from plasma. The contact of blood plasma with their sensors modified with antifouling layers and PMB resulted in protein adsorption as high as 25% of the fouling observed on their uncoated sensors.29 Another example of surface immobilized receptors to bind LPS was introduced by Kang et al. 30 In this system an engineered human opsonin-mannose-binding lectin is immobilized to magnetic nanobeads and was used to cleanse the blood of mice from LPS. 30 The alluded systems are based on exposing large surfaces modified with molecules that have affinity to bind LPS. However, the exposure of artificial surface to blood invariably leads to the rapid adsorption proteins.31-33 This, subsequently activates proteases from the coagulatory system (e.g. factor XII, Kinin-Kalikrein) as well as platelets and leukocytes, which concomitantly results in thrombus formation, clotting of the cartridge and in more severe cases to thrombocytopenia or hypotension.23 Such conditions can be particularly lethal on patient in already critical conditions such as those with sepsis.


5


Page 6 of 30

Inspired by these two examples, we saw the potential of modifying microparticles with antifouling coatings functionalized with polymyxin B. Polymer microparticles have already found various applications in drug delivery as carriers for drugs34-37 or genes,38 in tissue regeneration as scaffolds for tissue regeneration39-41, for imaging,42 diagnostics or biotechnology.43 Their use as a filler of cartridge will increase the contact area enabling to capture higher amount of LPS since it is experimentally proved that one molecule of LPS binds to one molecule of polymyxin B.24 In order to ensure the direct contact of the polymeric microparticles with the biological media, the surface of particles must be engineered in order to suppress the protein fouling from blood plasma. Protein adsorption occurs by the interplay of hydrophobic effect, coulombic interactions, hydrogen bounding, and van der Waals forces.44 The problem is even more complex when dealing with real biological fluids in which competitive adsorption occurs with clear changes with time as described by the Vroman effect.45 In particular, blood plasma has been shown to be the most challenging of all bodily fluids.46 Numerous types of surface modification starting from monolayers of natural components as albumin,47-49 polysaccharides50, 51 or silica52-55 and synthetic polymer as poly(ethylene glycol) (PEG)56-58 have been developed to reduce or prevent the protein fouling.59 The approaches are usually based on increasing the wettability of the surface as well as increasing an entropic barrier by immobilizing long macromolecules. Most of these surface modifications demonstrated excellent results in preventing protein fouling from the main plasma proteins, human serum albumin (HSA) and fibrinogen (Fbg).60-63 But, their resistance to fouling from more challenging media as blood plasma is limited. In such conditions, surface modification based on “grafted-from” polymer brushes demonstrates unmatched results.64 They are capable to prevent fouling from main plasma proteins and to minimize fouling from more challenging media. For example,


6

Page 7 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

oligo(ethylene glycol) methacrylate, oligo(ethylene glycol) methyl ether methacrylate, 2hydroxypropyl methacrylate, and 2-hydroxyethyl methacrylate have reduced the fouling from undiluted blood plasma by 93.6, 92.5, 86.3, and 85.3%, respectively.65, 46 To date only polymer brushes based on carboxybetaines and N-2-hydroxypropyl methacrylamide (HPMA) have shown complete suppression of the fouling.66, 65 However, previous studies demonstrate that activation and biofuctionalization of these brushes lead to an increase of fouling.67 To overcome this problem, our group designed a new non-fouling polymer, where only a small fraction of the side chains could be functionalized. The copolymers consisted of two non-fouling monomers, HPMA and carboxybetaine methacrylamide (CBMAA) statistically copolymerized to obtain a novel nonfouling

polymer;

poly[(N-(2-hydroxypropyl)

methacrylamide)-co-(carboxybetaine

methacrylamide)] (poly(HPMA-co-CBMAA) with a molar ratio of 17:3 of the monomers.

68-71

Only the side groups of CBMAA were functionalized in this polymer brush that allowed us to make minimal changes in the structure and maintain its non-fouling properties. The precise grafting of the polymer brushes requires the use of controlled radical polymerizations.72, 73 However, the polymerization of zwitterionic monomers as well as HPMA – a methacrylamide – is challenging. Reversible addition-fragmentation chain transfer (RAFT) does not provide a high grafting density since the process is not truly surface confined. The process requires the diffusion of macromolecules to the surface to exchange the chain transfer agent.74 Surface-initiated atom transfer radical polymerization (SI-ATRP) does not provide good control over the process. The growth of brushes rapidly reaches a plateau, after which no re-initiation occurs. This is presumably caused by termination reactions at the chain ends.65 Another drawback of ATRP polymerization is the need of high concentrations of copper catalyst which is not suitable for biomedical applications. Unlike to ATRP, SET-LRP75, 76 polymerization requires very low


7


Page 8 of 30

concentration of copper catalyst, have high living nature,77, 78 can be used for a wide range of monomers in different solvents including water,79, 80, 76 and can be performed in non-deoxygenated conditions.81 The SET-LRP mechanism relies on the an activation step mediated by the single electron transfer from Cu(0) to the alkyl halide initiator or polymer ends and the rate of disproportionation of Cu(I)LigandX (X = Br or Cl) species into nascent Cu(0) and Cu(II)LigandX2 the activator and deactivator for the polymerization.82, 83 The disproportionation reaction is highly dependent on the solvent. It requires that the solvent is able to disproportionate effectively Cu(I). Typical solvents include; water, dipolar aprotic, fluorinated and alcohols among other.76 Recently Percec, Lligadas and Monteiro introduced the concept of biphasic systems to extend the repertoire of SET-LRP conditions to poorly disproportionating solvents.84-87 This was achieved by combining water with those solvents or by in-situ generation of the biphasic system.84 In recent time, Anastasaki and others reported of photoinduced SET-LRP.88-90, 79 In this process, an aliphatic tertiary amine (Me6TREN) upon irradiation acts as a photoelectron donor and promotes the polymerization via the outer-sphere single electron transfer into the alkyl halide initiator.88, 91, 77 Our group developed new protocol for photoinduced surface-initiated SET-LRP.92, 93 This protocol requires very low concentrations of copper catalyst, up to ppb level, provides good control of polymerization, and the process can be re-initiated. It has enabled to obtain very thick polymer brushes, up to 1 µm thick in one hour and it suitable for wide range of acrylic and methacrylic monomers.92, 93 Remarkably, this photoinduced SET-LRP protocol was successfully applied for polymerization of HPMA94 and zwitterionic monomers including CBMAA.95 In this study, we introduced an effective system to capture LPS in blood plasma. For that purpose we used porous poly(glycidyl methacrylate) microparticles (μP-GMA) previously functionalized with α-bromoisobutyryl bromide (BIBB). The latter were used to graft non-fouling poly(HPMA-


8

Page 9 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

co-CBMA) brushes from the external and pore’s surface by photoinduced SET-LRP. The brushes were functionalized with polymyxin B. These brushes prevented non-specific adsorption of proteins from blood plasma, a pre-requisite for hemocompatibility while polymyxin B accounted for the efficient removal of LPS spiked in blood plasma. MATERIALS AND METHODS Materials. Oligo(ethylene glycol) methyl ether acrylate (Mn = 480 g mol–1, OEGA), αbromoisobutyryl bromide (98%, BIBB), tris[2-(dimethylamino)ethyl]amine (97%, Me6TREN), Cu2Br (99.999% trace metal basis), 4-(dimethylamino)pyridine (99%, DMAP) triethylamine (99.5%, TEA), anhydrous tetrahydrofuran (99.9%, THF), 2,2′-azobis(2-methylpropionitrile) (AIBN), dibutyl phthalate (DBP), sodium dodecyl sulfate (SDS), phosphate buffer saline tablets (PBS), and human blood plasma were supplied by Sigma-Aldrich, Czech Republic and used as received unless otherwise stated. Styrene (Synthos Kralupy Inc., Czech Republic), glycidyl methacrylate (GMA) (Fluka, Switzerland) and ethylene dimethacrylate (EDMA) (Ugilor, France), were vacuum-distilled before use. Lithium persulfate, lithium hydrogen carbonate, sulfuric acid were purchased from Lach-Ner, Czech Republic. Poly(vinyl alcohol) (PVA) was purchased from Wacker, Germany. Extra-dry (over molecular sieves) toluene (99.85%) and dimethyl sulfoxide (99.7%, DMSO) were purchased from Acros, Czech Republic. Deionized (MilliQ) water was obtained from a Milli-Q system (Merck-Millipore, Czech Republic). Polymyxin B sulfate and 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (for synthesis, EDC) were purchased from Merk Millipore, Germany. N-Hydroxysuccinimide (98%, NHS) was purchased from Sigma Aldrich, Germany. Lipopolysaccharides from Salmonella Minnesota, Alexa Fluor™ 488 conjugate was purchased from Thermo Fisher Scientific, Germany. Cell culture pyrogen-free water was purchased from Biowest, Germany.


9


N-(2-hydroxypropyl)

methacrylamide

(HPMA)

and

Page 10 of 30

(3-methacryloylaminopropyl)-(2-

carboxyethyl)-dimethylammonium (CBMAA) were synthesized according to the literature.96, 97 Synthesis of Poly(Glycidyl Methacrylate) (μP-GMA) and Poly(2,3-Dihydroxypropyl Methacrylate) (μP-OH) Microparticles. Monodisperse macroporous μP-GMA were synthesized by multistep swelling polymerization of polystyrene seeds with DBP and then with a mixture of monomers and porogens, followed by conventional suspension polymerization, using a modified Ugelstad technique.98 Subsequently, μP-GMA microparticles were hydrolyzed with 0.1 M H2SO4 at 60 ⁰C for 5 h under mechanical stirring to yield μP-OH. For detail information of microparticles synthesis refer to the supporting information. Immobilization of Initiator. 2 g of μP-OH were placed in a round-bottomed flask together with anhydrous THF (30 mL), DMAP 0.032 g (0.26 mmol), TEA 1.47 g (14.5 mmol) and BIBB 3.03 g (13.2 mmol) under Ar atmosphere. The reaction was allowed to proceed at room temperature for 16 h on a shaker plate. The modified poly[2,3-di(2-bromo, 2-methyl propionate) propyl methacrylate] (μP-Br) microparticles were then separated by sedimentation, washed with THF, methanol and MilliQ water and dried at 60 ⁰C in a vacuum oven. Grafting of Poly(HPMA-co-CBMAA) and Poly(OEGA) Brushes from μP-Br by Photoinduced SET-LRP 1.314 g (9.18 mmol) of HPMA and 0.392 g (1.62 mmol) of CBMAA (HPMA:CBMAA = 17:3 mol/mol) were dissolved in 6.15 mL DMSO while kept in dark by wrapping it in Al-foil. Then, 273 µL of a freshly prepared stock catalyst solution of CuBr2 (3.9 mM) and Me6TREN (23.4 mM) in DMSO was added. The obtained polymerization solution was degassed by bubbling Ar for 60 min while stirring. Subsequently, 5 mL of the polymerization solution were transferred using a gas-tight syringe under Ar protection to previously degassed


10

Page 11 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

(purging with Ar for 15 min) crimped vial containing 0.2 g of μP-Br. The vial was irradiated inside a UV-reactor (a nail-curing device; four 9 W lamps, λmax = 365 nm) while shaking (150 min-1) and kept at room temperature by fanning with a ventilator. The polymerization was conducted for 40 min and quenched by exposing the reaction mixture to air followed by addition of 2 mL of DMSO. Resulting poly(HPMA-co-CBMAA) microparticles (μP-HPMA-CBMAA) were separated by centrifugation (10 min at 5000 rpm), washed with methanol, MilliQ water, and lyophilized. Poly(OEGA) microparticles (μP-OEGA) were prepared by an analogous procedure, but utilizing 4.8 mL (10.9 mmol) of OEGA and filtered thought basic alumina to remove inhibitor, with irradiation time of 15 min. Physicochemical characterization of microparticles. Chemical composition and physical properties of the particles were investigated using proton nuclear magnetic resonance ( 1HNMR), Fourier-transform infrared spectroscopy (FTIR), Raman spectroscopy, X-ray photoelectron spectroscopy (XPS), thermogravimetric analysis (TGA), differential scanning calorimetry (DSC), field-emission scanning electron microscopy (FESEM), transmission

electron

microscopy

(TEM),

mercury

intrusion

porosimetry

(Hg

porosimetry), and Brunauer–Emmett–Teller surface area analysis (SBET). Detailed information on the techniques could be found in the supporting information. Fouling. The resistance to protein adsorption of the brushes grafted on the microparticles was assessed by contacting the particles (μP-GMA, μP-OEGA and μP-HPMA-CBMAA) with human blood plasma (HBP). A suspension of the particles (18 mg) was thoroughly washed by PBS (1.8 mL; pH 7.4), and centrifuged at 14000 rpm for 10 min. HBP (1% in


11


Page 12 of 30

PBS) was added and the particles were kept in contact with it for 60 min at 25 ⁰C. XPS was used to confirm the resistance to protein adsorption. It was measured before and after contact with blood plasma. This technique allows to probe the top 10 nm of the particle surface and accurately determine if proteins adsorbed, which would be otherwise not accessible by conventional techniques. Activation of carboxylate groups of μP-HPMA-CBMAA. 0.33 g of μP-HPMACBMAA were dispersed in 1 mL of MilliQ water and mixed with 1 mL solution of 1:1 v/v EDC (0.8 M) and NHS (0.2 M). The reaction proceeded for 10 min with continuous shaking. μP-HPMA-CBMAA with activated polymer brushes were separated by centrifugation for 5 min with 10000 rpm and washed two times with 1 mL MilliQ water. Immobilization of Polymyxin B. To a 1 mL suspension of freshly activated μP-HPMACBMAA (330 mg) was added 1 mL solution of polymyxin B (11.2 mg mL-1) in pyrogenfree PBS buffer (pH 7.4) and the mixture was allowed to react for 4 h at room temperature and for 12 h at 4ºC with gentle shaking. The resulting μP-HPMA-CBMAA with immobilized polymyxin B (μP-PMB) were separated by centrifugation at 10000 rpm for 5 min. The microparticles were washed three times with 1.5 mL pyrogen-free PBS buffer and stored at 4ºC in a PBS. LPS removal from blood plasma. Fluorescently-labeled LPS (0.1 mg) was dissolved in 100 µL DMSO. The solution was diluted with pyrogen-free PBS buffer to a concentration of 10 µg·mL-1. HBP (1% in PBS, 100 µL) was spiked with 30 µL of the LPS solution to model septic plasma. Subsequently, 100 µL of μP-PMB suspension (0.6 g·mL-1) were


12

Page 13 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

added to 100 µL of the spiked HBP solution. As a control, the same amount of μP-HPMACBMAA (without polymyxin B) were added to 100 µL of spiked HBP (1% in PBS). The microparticles were incubated for 1 h at 25 ⁰C under shaking. The scavenging of LPS was visualized by CLSM using a Leica SP8 setup (63X/1.40 OIL objective and a 488 nm light laser for excitation). LPS was labeled with Alexa Fluor™. Signals were detected with HyD detector in a wavelength range between 498 – 590 nm. RESULTS AND DISCUSSION The key requirements for a system to effectively scavenge LPS from blood are (i) high specific surface area, (ii) non-fouling properties of the material, and (iii) high affinity to endotoxins. To achieve this properties simultaneously, our approach is based on functionalization of highly porous μP-GMA with poly(HPMA-co-CBMAA) brushes, followed by immobilization of polymyxin B. The selection of polymer brushes was based on their resistance to protein adsorption from biological fluids. Brushes based on HPMA and CBMAA not only withstand non-specific protein adsorption, but also have readily functionalizable carboxylic groups for immobilization of polymyxin B (Scheme 1). Such system was successfully synthesized and fully characterized. The system was benchmarked against the sample particles modified with poly(OEGA) brushes, which are well-known for displaying excellent resistance to protein adsorption.


13


Page 14 of 30

Scheme 1. Chemical route followed to obtain non-fouling μP-PMB able to scavenge LPS. μPGMA are hydrolyzed to yield μP-OH. This is followed by acylation with BIBB. Brushes of OEGA or HPMA and CBMMA were grafted using photoinduced SET-LRP ([M]:[Me6TREN]:[CuBr2] = [10150]:[6]:[1]). The carboxylate groups of CBMMA are used to immobilize polymyxin B. BIBB - α-bromoisobutyryl bromide; TEA – trimethylamine; DMAP - 4-(dimethylamino)pyridine; THF - tetrahydrofuran; DMSO - dimethyl sulfoxide; Me6TREN - tris[2-(dimethylamino)ethyl]amine; UV

-

ultraviolet

lamp;

NHS

-

N-hydroxysuccinimide;

EDC

-

1-ethyl-3-(3-

dimethylaminopropyl)carbodiimide; PMB - polymyxin B; HBP – human blood plasma. Synthesis and characterization. Monodisperse μP-GMA with high porosity were synthesized via a modified Ugelstad technique.98 The microparticles contained 60 wt % GMA and 40 wt % EDMA as a crosslinking agent. Electron micrographs revealed that μP-GMA have very uniform


14

Page 15 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

size distribution (9.0 ± 0.1 μm), possess rough surface and high internal porosity, i.e., high specific surface area (Figure 2-I (a-d)). The size and volume of pores of microparticles was determined by means of Hg porosimetry and BET (Table 1). Interestingly, the microparticles have two distinct ranges of pores – macropores with average size of 115 nm and mesopores with average size of 25 nm. The total pore volume of particles was 0.87 mL g-1 corresponding to ca. 45% porosity and 85.6 m2 g-1 specific surface area. In order to graft polymer brushes from the surface of microparticles, the oxirane rings were hydrolyzed to form hydroxyl groups to which the initiator for polymerization (BIBB) was immobilized. Two types of polymer brushes - poly(OEGA) and poly(HPMA-co-CBMAA) - were grafted by photoinduced SET-LRP (Scheme 1). Importantly, for polymerization we utilized only ca. 8 ppm of copper catalyst. It should be noted, however, that even lower concentrations of Cu catalyst (ca. 8 ppb) could be utilized without compromising the polymerization kinetics (refer to figure S7 in the SI). The successful modification of microparticles with polymer brushes was confirmed by vibrational (Raman, FTIR) microspectroscopy and X-ray photoelectron spectroscopy (XPS). Raman and FTIR spectra of the single μP-GMA, μP-OEGA, and μP-HPMA-CBMAA are displayed in Figure 1-I and 1-II, respectively. Raman spectrum of a single μP-GMA displays the characteristic features of the oxirane rings at 910, 815 and 1260 cm–1 corresponding to the deformation and breathing vibration of the oxirane ring, and at 1135 cm-1 stemming from C–O stretching in ester groups (Figure 1 A and Table S2 in SI).99 Furthermore, the prominent band at 1450 cm-1 and shoulders at 1480 cm-1 and1425 cm-1 correspond to scissoring of CH2, asymmetric deformation of CH3 and twisting of CH2 in the oxirane ring, respectively. The modification with BIBB and grafting of brushes led to the disappearance of the signals associated with the oxirane groups consistent with


15


Page 16 of 30

the full opening of oxiranes. Both brush modifications display a well-resolved band at 1,110 cm–1 stemming from asymmetric stretching of C–O–C in ethylene glycol and of C–O in the secondary alcohol of HPMA (Figure 1-I(b-c)). The bands at 1280 cm–1 and 1245 cm–1 are characteristic of the twisting mode of the methylene groups in oligo(ethylene glycol) and confirm structure of poly(OEGA) (Figure 1-I(b)).100 The most prominent features of poly(HPMA-co-CBMAA) are represented by bands at 1640, 1520, 1600, and 1365 cm -1 in FTIR spectrum corresponding to amide I and II and the asymmetric and symmetric stretching carboxylate groups (Figure 1-II(c) and Table S3 in SI).101, 102 The high resolution XPS spectra provided a more detailed view of the particle topmost interfacial composition (Figure 1-III and Table S1 in the SI). The C 1s envelope of μP-GMA consisted of the contributions from sp2 (C=C) carbon (centered at 284.3 ± 0.1 eV), sp3 carbon (C-C, C-H, centered at 285.0 eV), secondary C*-COO shifts (285.7 ± 0.2 eV), C(=O)-O-C moieties (286.8 ± 0.3 eV), C-O-C oxirane rings (287.5 ± 0.1 eV) and C(=O)-O ester (288.9 ± 0.3 eV) functionalities (Figure 1-III(a)) confirming the presence of oxirane rings. Furthermore, the corresponding O 1s envelope showed similar contributions of C(=O)-O, C-O-C and C(=O)-O moieties centered at 532.0 ± 0.3, 532.9 ± 0.2 and 533.8 ± 0.3 eV (Figure S1 in the SI). The hydrolysis of the oxirane rings led to increase of overall surface concentration of oxygen and a decrease of the contributions in the C 1s spectra originating from the of the C-O-C epoxide moiety. Concomitantly, we observed an increase of the contributions characteristic for the hydroxyl groups at about 286.6 ± 0.1 eV and 533.2 ± 0.3 eV within the C 1s and O 1s high


16

Page 17 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 1. Vibrational spectra of (a) μP-GMA, (b) μP-OEGA, and (c) μP-HPMA-CBMAA. (I) Raman spectra and (II) FTIR spectra; the spectra are scaled and offset for clarity and comparison. (III) High resolution C 1s XPS spectra of (a) μP-GMA, (b) μP-OH, (c) μP-HPMA-CBMAA, (d) μP-OEGA and (e) μP-HPMA-CBMAA.


17


Page 18 of 30

resolution spectra, respectively. The functionalization of the surface of the hydrolyzed particles with BIBB initiator can be clearly evidenced by the appearance of the Br-C contributions in the Br 3d high resolution spectra (Figure S2 in the SI). In addition, the acylation led to increase of aliphatic and ester contributions within the C 1s and O 1s spectra. The polymerization OEGA and HPMA with CBMAA brushes from the surface of initiator-modified microparticles, caused significant changes in the C 1s and O 1s spectra completely masking the underneath μP-OH material. This indicates that the brushes were thicker than the operational sensing depth of XPS (ca. 10 nm). The C 1s and O 1s spectra of μP-OEGA are dominated by contributions at 286.8 ± 0.1 eV and 532.5 ± 0.1 eV, which are highly characteristic for the ethylene oxide side chains of the polymer brush.92 C 1s spectrum of μP-HPMA-CBMAA displayed peaks at 286.4 ± 0.1 eV and 287.8 ± 0.1 eV corresponding to C-N and C(=O)-NH groups. The N 1s spectrum additionally revealed signals from amide (C(=O)-NH) and quaternary amine (-N+(CH3)2) at 399.9 ± 0.1 eV and 402.6 ± 0.1 eV, respectively (Figure S3 in the SI). The high resolution XPS spectra of polymer brushes grown from the surface of μP-Br correspond well with the ones obtained for the polymer brushes grown from initiator molecules immobilized on silicon substrates.92, 69 Analysis of Br 3d spectra of brushes showed that the Br chain-ends were still present after the polymerization suggesting a high degree of livingness of the SET-LRP (Table S1 in the SI) well in line with our previous reports.94, 93 The morphology and distribution of polymer brushes was assessed by a combination of electron microscopy and porosimetry. FESEM micrographs show that the grafting process did not lead to a change in the average size, confirming that the brush thickness were well below 1 µm and that the polymerization was confined to the surface (Figure 2-I). However, notorious differences in the final morphology of the external surface, as well as in the available pore surface, were observed


18

Page 19 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

for both polymer modifications. The most prominent changes in the external morphology were observed after the grafting of poly(HPMA-co-CBMAA).

Figure 2. I. FESEM (a, b, c, e, f, g, i, j and k) and TEM (d, h and l) of μP-GMA (a-d), μP-OEGA (e-h) and μP-HPMA-CBMAA (i-l); (d, h and l) are particle cross-section. II. Thermal analysis of μP-GMA (black), μP-OEGA (red) and μP-HPMA-CBMAA (blue) by (1) DSC and (2) TGA analysis.


19


Page 20 of 30

Significant smoothening of the surface roughness and apparent closing of pores were observed in FESEM. (Figure 2-I(f-h)). TEM micrographs of µP-HPMA-CBMAA confirmed the formation of a dense layer at the periphery of the microparticles. This was accompanied by a reduction in porosity from 43 to 5% (Table 1) and a 98% reduction of the SBET. This indicates that poly(HPMAco-CBMAA) brushes rapidly grew in all surfaces at initial stages, followed by closing the outmost mesopores, preventing further growth in the inner surfaces. In the case of μP-OEGA the porosity and SBET were reduced to 28 and 20%, respectively, of those of the pristine microparticles.

Table 1. Surface analysis of microparticles before and after modification with polymer brushes SBET a (m2/g)

Total pore volumeb (mL·g-1)

Porosityb (%)

μP-GMA 85.6 0.87 43.81 μP-OEGA 16.11 0.43 27.71 μP-HPMA-CBMAA 1.96 0.13 5.33 a b Brunauer–Emmett–Teller surface area analysis, Hg porosimetry;

Pore sizeb (nm)

macropores mesopores 115 24 99 24 86 13

Hydrophobic effect is one of the key intermolecular forces driving protein adsorption. Therefore hydrophilic surfaces are necessary to prevent fouling. DSC and TGA analysis of μP-GMA showed the vaporization of volatiles (0.6% in DSC and 0.3% TGA) at temperature range between 40 – 90ºC. Similarly, DSC and TGA of μP-OEGA showed the vaporization of volatiles and a very low amount (ca. 1-2%) of free water adsorbed from the atmosphere. A different behavior was observed for μP-HPMA-CBMAA. Two endothermic peaks (100 and 125ºC) were observed in the DSC indicating the presence of free and bound water103 while the TGA showed a 2 and 10% weight loss associated with each vaporization. The bound water in μP-HPMA-CBMMA stems from the stronger hydration due to the zwitterionic groups of the betaine monomer.


20

Page 21 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 3. High resolution N 1s XPS spectra of (a) μP-GMA, (b) μP-OEGA and (c) μP-HPMACBMAA before and after exposure to HBP. Protein Fouling. The amount of proteins adsorbed on pristine μP-GMA μP-OEGA and μPHPMA-CBMAA particles was measured using XPS before and after contact with 1% HBP for 60 min at 25 ⁰C (Figure 3). The contact of HBP with μP-GMA was accompanied by significant changes in the C 1s, O 1s and N 1s spectra.While the N 1s spectra of neat particles before contact with blood plasma lacked of any contributions from nitrogen, the particles contacted with blood were characterized by a dominating peak of amide NH—C(=O) at 400.2 ± 0.1 eV and a charged amine peak at 401.6 ± 0.1 eV. This is further corroborated by the appearance of C-N moieties, amide C(=O)-NH at 286.6 ± 0.1 eV and 287.8 ± 0.1 eV in the high resolution C 1s spectrum and


21


Page 22 of 30

the increase of the C(=O)-NH peak at 532.2 ± 0.2 eV in the O 1s spectrum of μP-GMA (Figure S4 and S5 in the SI). Conversely, negligible changes in the XPS spectra of the brush-modified particles were observed. The atomic content calculated from XPS analysis (Table S1 and S4 in the SI) indicates that both brushes, poly(OEGA) and poly(HPMA-co-CBMAA), prevented the adsorption of proteins from blood plasma. Complementary fouling studies were performed using surface plasmon resonance (SPR) in polymer brushes grafted from the gold sensor. SPR allows the facile study of fouling even using undiluted plasma. The brushes of poly(HPMA-co-CBMAA) were able to prevent more than 98% of the fouling from undiluted pooled (5 donors) blood plasma (refer to figure S8 in the SI). Scavenging of LPS from blood plasma. Monodisperse μP-HPMA-CBMAA were selected as a platform to design scavengers for LPS due to their excellent resistance to blood plasma fouling (vide supra) and for the availability of functionalizable groups. The relatively low proportion of these groups was shown to allow the modification without impairment of the antifouling properties.68-71 Polymyxin B was covalently linked to μP-HPMA-CBMAA by amidation to carboxylate groups of CBMMA side groups preactivated by EDC/NHS to yield μP-PMB. Polymyxin B is a potent antimicrobial which can bind LPS endotoxins by hydrophobic and coulombic interactions.24, 25 The immobilization of polymyxin B on poly(HPMA-co-CBMAA) brushes was directly followed in a model surface using SPR. After 50 min of contact the amount of polymyxin B on the brushes could be estimated to be 77 ng·cm-2. Remarkably the functionalization of the brushes did not deteriorate the antifouling


22

Page 23 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

properties of the brushes, which resisted the fouling of blood plasma ranging from 1 to 100% (refer to figure S9 in the SI).

Figure 4. Confocal images (left: green fluorescent channel, right: transmission channel) of (A) μP-HPMA-CBMAA after contact with fluorescently labeled LPS in 1% HBP and (B) μP-PMB after 1 h contact with HBP spiked with LPS. Subsequently, μP-HPMA-CBMAA and μP-PMB were incubated with HBP (1%) spiked with Alexa Fluor™- labeled LPS (C = 1.5 µg mL-1). The binding of LPS to microspheres was visualized by CLSM using a 488 nm light laser for excitation of Alexa Fluor™ fluorophore linked to LPS (Figure 4). μP-PMB become strongly fluorescent, while negligible fluorescence was observed in the surrounding medium. This indicates the efficient and total removal of LPS from blood plasma. Conversely, the CSLM micrographs of μP-HPMA-CBMAA (not functionalized with polymyxin B) suspension showed homogeneous fluorescence indicating a high concentration


23


Page 24 of 30

of LPS in the surrounding media. Centrifugal filtration (EMD Millipore™ Amicon™ Ultra-15, 50 kDa) led to particles without any fluorescence, clearly indicating that any LPS adsorbed in the non-functionalized particles (Figure S6 in the SI). CONCLUSIONS The first example of the grafting of poly(OEGA) and poly(HPMA-co-CBMAA) polymer brushes from the surface of microparticles by photoinduced SET-LRP is reported. The successful modification of microparticles with brushes was confirmed by FTIR, Raman spectroscopy, and XPS. Poly(HPMA-co-CBMAA) brushes completely prevented the fouling from human blood plasma. The excellent resistance to non-specific protein adsorption and high surface area of µPHPMA-CBMAA combined with their functionalization with polymyxin B enabled the scavenging of LPS from blood plasma. This system may be the basis for the development of hemofiltration columns that scavenge LPS with minimal activation of coagulation. ASSOCIATED CONTENT Supporting Information Description of all techniques used for characterization, additional XPS data, assignment of Raman and infrared vibration bands, surface-initiated polymerization kinetic with different concentration of copper catalyst, SPR of immobilization of polymyxin B and protein fouling can be found in the supporting information (PDF).


24

Page 25 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

AUTHOR INFORMATION Corresponding Author *Dr. Cesar Rodriguez-Emmenegger. Email: [email protected] Author Contributions MV, NYK and CR-E conceived the researcher. MV, NYK, KR, SG, OPG and AS perform the experiments. The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes The authors declare no competing financial interest. ACKNOWLEDGEMENT The research was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) via Schwerpunktprogramm “Auf dem Weg zur implantierbaren Lunge” (SPP 2014 Project No. RO 5490/1-1). N.Y.K. acknowledges the Alexander von Humboldt Foundation, O.P.G. acknowledges the support from the Czech Science Foundation (GACR) under Contract No. 16-02702S. C.R.-E. acknowledges support from Prof. Martin Möller.


25


Page 26 of 30

REFERENCES 1. Garrett, T. R.; Bhakoo, M.; Zhang, Z., Prog. Nat. Sci. 2008, 18, (9), 1049-1056. 2. Donlan, R. M.; Costerton, J. W., Clin. Microbiol. Rev. 2002, 15, (2), 167-193. 3. Flemming, H. C.; Wingender, J., Nat. Rev. Microbiol. 2010, 8, (9), 623-33. 4. Hoiby, N.; Bjarnsholt, T.; Moser, C.; Bassi, G. L.; Coenye, T.; Donelli, G.; Hall-Stoodley, L.; Hola, V.; Imbert, C.; Kirketerp-Moller, K.; Lebeaux, D.; Oliver, A.; Ullmann, A. J.; Williams, C.; Werner, Z., Clin. Microbiol. Infect. 2015, 21, S1-S25. 5. Donlan, R. M., Emerg. Infect. Dis. 2001, 7 ( 2), 277–281. . 6. Hall-Stoodley, L.; Costerton, J. W.; Stoodley, P., Nat Rev Microbiol 2004, 2, (2), 95-108. 7. Veerachamy, S.; Yarlagadda, T.; Manivasagam, G.; Yarlagadda, P. K., Proc. Inst. Mech. Eng. H 2014, 228, (10), 1083-99. 8. A.G. Gristina; Costerton, J. W., J. Bone Joint Surg. 1985, 67-A, (2), 264-273. 9. Cohen, J., Nature 2002, 420, 885–891 10. Glauser, M. P.; Zanetti, G.; Baumgartner, J.-D.; Cohen, J., Lancet 1991, 338, (8769), 732736. 11. Rimmelé, T.; Kellum, J. A., Crit. Care 2011, 15, (1), 205. 12. Abe, R.; Oda, S.; Sadahiro, T.; Nakamura, M.; Hirayama, Y.; Tateishi, Y.; Shinozaki, K.; Hirasawa, H., Crit. Care 2010, 14, 27. 13. Cardoso, L. S.; Araujo, M. I.; Goes, A. M.; Pacifico, L. G.; Oliveira, R. R.; Oliveira, S. C., Microb. Cell Fact. 2007, 6, 1. 14. Manocha, S.; Feinstein, D.; Kumar, A.; Kumar, A., Expert Opin Investig Drugs 2002, 11, (12), 1795-812. 15. Saito, N.; Sugiyama, K.; Ohnuma, T.; Kanemura, T.; Nasu, M.; Yoshidomi, Y.; Tsujimoto, Y.; Adachi, H.; Koami, H.; Tochiki, A.; Hori, K.; Wagatsuma, Y.; Matsumoto, H., PLoS One 2017, 12, (3), e0173633. 16. Riedemann, N. C.; Guo, R.-F.; Ward, P. A., Nat. Med. 2003, 9, 517. 17. Gutsmann, T.; B Schromm, A.; Brandenburg, K., The physicochemistry of endotoxins in relation to bioactivity. 2007; Vol. 297, p 341-52. 18. Mueller, M.; Lindner, B.; Kusumoto, S.; Fukase, K.; Schromm, A. B.; Seydel, U., J Biol Chem 2004, 279, (25), 26307-13. 19. Cole, L.; Bellomo, R.; Journois, D.; Davenport, P.; Baldwin, I.; Tipping, P., Intensive Care Med. 2001, 27, (6), 978–986. 20. Kunitomo, T.; Shoji, H., Contrib. Nefrol. 2001, 132, 415-420 21. M. Kodama; K. Hanasawa; Tani., T., Ther. Apher. 1997, 1, (3), 224–227 22. Shimizu, T.; Miyake, T.; Tani, M., Ann. Gastroenterol. Surg. 2017, 1, (2), 105-113. 23. Shoji, H., Ther. Apher. Dial. 2003, 7, (1), 108–114 24. David C. Morrison; Jacobs, D. M., Immunochemistry 1976, 13, (10), 813-818. 25. Ronco, C.; Klein, D. J., Crit. Care 2014, 18. 26. Evans, M. E.; Feola, D. J.; Rapp, R. P., Ann. Pharmacother. 1999, 33, (9 ), 960-967. 27. Sharon R. Snavely; Hodges, G. R., Ann. Intern. Med. 1984, 101, (1), 92-104. 28. Romaschin, A. D.; Harris, D. M.; Ribeiro, M. B.; Paice, J.; Foster, D. M.; Walker, P. M.; Marshall, J. C., J. Immunol. Methods 1998, 212, (2), 169-185. 29. Thompson, M.; Blaszykowski, C.; Sheikh, S.; Romaschin, A., Biosens Bioelectron 2015, 67, 3-10.


26

Page 27 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

30. Kang, J. H.; Super, M.; Yung, C. W.; Cooper, R. M.; Domansky, K.; Graveline, A. R.; Mammoto, T.; Berthet, J. B.; Tobin, H.; Cartwright, M. J.; Watters, A. L.; Rottman, M.; Waterhouse, A.; Mammoto, A.; Gamini, N.; Rodas, M. J.; Kole, A.; Jiang, A.; Valentin, T. M.; Diaz, A.; Takahashi, K.; Ingber, D. E., Nat. Med. 2014, 20, (10), 1211-6. 31. Blaszykowski, C.; Sheikh, S.; Thompson, M., Chem. Soc. Rev. 2012, 41, (17), 5599-5612. 32. Blaszykowski, C.; Sheikh, S.; Thompson, M., Trends Biotechnol. 2014, 32, (2), 61-62. 33. Thompson, M.; Blaszykowski, C.; Sheikh, S.; Rodriguez-Emmenegger, C.; de los Santos Pereira, A., Biological Fluid-Surface Interactions in Detection and Medical Devices Royal Society of Chemistr: United Kingdom, 2016; p 300. 34. K. Tajdaran; M. S. Shoichet; T. Gordon; Borschel, G. H., Biotechnol. Bioeng. 2015, 112, (9), 1948–1953 35. Rahimian, S.; Fransen, M. F.; Kleinovink, J. W.; Amidi, M.; Ossendorp, F.; Hennink, W. E., Biomaterials 2015, 61, 33-40. 36. Shaked, E.; Shani, Y.; Zilberman, M.; Scheinowitz, M., J. Biomed. Mater. Res. B 2015, 103, (6), 1228-1237. 37. Win, K. Y.; Ye, E.; Teng, C. P.; Jiang, S.; Han, M.-Y., Advanced Healthcare Materials 2013, 2, (12), 1571-1575. 38. Bhavsar, M. D.; Amiji, M. M., Expert. Opin. Drug. Deliv. 2007, 4, (3), 197-213. 39. Bock, N.; Dargaville, T. R.; Woodruff, M. A., Prog. Polym. Sci. 2012, 37, (11), 1510-1551. 40. Iwasaki, Y.; Takahata, Y.; Fujii, S., Colloids Surf. B 2015, 126, (Supplement C), 394-400. 41. Salerno, A.; Domingo, C., J. Porous Mater. 2015, 22, (2), 425-435. 42. Shapiro, E. M., Magn. Reson. Med. 2015, 73, (1), 376-389. 43. Wang, W.; Zhang, M.-J.; Chu, L.-Y., Acc. Chem. Res. 2014, 47, (2), 373-384. 44. Vogler, E. A., Biomaterials 2012, 33, (5), 1201-37. 45. Noh, H.; Vogler, E. A., Biomaterials 2007, 28, (3), 405-22. 46. Rodriguez-Emmenegger, C.; Houska, M.; Alles, A. B.; Brynda, E., Macromol. Biosci. 2012, 12, (10), 1413-1422. 47. Gu, Z.; Zuo, H.; Li, L.; Wu, A.; Xu, Z. P., J. Mater. Chem. B 2015, 3, (16), 3331-3339. 48. Horak, D.; Svobodova, Z.; Autebert, J.; Coudert, B.; Plichta, Z.; Kralovec, K.; Bilkova, Z.; Viovy, J. L., J. Biomed. Mater. Res., Part A 2013, 101, (1), 23-32. 49. Jiang, P.; Yu, D.; Zhang, W.; Mao, Z.; Gao, C., RSC Adv. 2015, 5, (51), 40924-40931. 50. Arnon, H.; Granit, R.; Porat, R.; Poverenov, E., Food Chem. 2015, 166, 465-472. 51. Asadinezhad, A.; Novák, I.; Lehocký, M.; Bílek, F.; Vesel, A.; Junkar, I.; Sáha, P.; Popelka, A., Molecules 2010, 15, (2), 1007. 52. Estupiñán, D.; Bannwarth, M. B.; Landfester, K.; Crespy, D., Macromol. Chem. Phys. 2015, 216, (21), 2070-2079. 53. Iqbal, M. Z.; Ma, X.; Chen, T.; Zhang, L. e.; Ren, W.; Xiang, L.; Wu, A., J. Mater. Chem. B 2015, 3, (26), 5172-5181. 54. Konduru, N. V.; Jimenez, R. J.; Swami, A.; Friend, S.; Castranova, V.; Demokritou, P.; Brain, J. D.; Molina, R. M., Part. Fibre Toxicol. 2015, 12, (1), 31. 55. Yue, Q.; Li, J.; Luo, W.; Zhang, Y.; Elzatahry, A. A.; Wang, X.; Wang, C.; Li, W.; Cheng, X.; Alghamdi, A.; Abdullah, A. M.; Deng, Y.; Zhao, D., J. Am. Chem. Soc. 2015, 137, (41), 1328213289. 56. He, Q.; Zhang, J.; Shi, J.; Zhu, Z.; Zhang, L.; Bu, W.; Guo, L.; Chen, Y., Biomaterials 2010, 31, (6), 1085-1092.


27


Page 28 of 30

57. Hlídková, H.; Horák, D.; Proks, V.; Kučerová, Z.; Pekárek, M.; Kučka, J., Macromol. Biosci. 2013, 13, (4), 503-511. 58. Riedel, T.; Riedelová-Reicheltová, Z.; Májek, P.; Rodriguez-Emmenegger, C.; Houska, M.; Dyr, J. E.; Brynda, E., Langmuir 2013, 29, (10), 3388-3397. 59. Rodriguez Emmenegger, C.; Brynda, E.; Riedel, T.; Sedlakova, Z.; Houska, M.; Alles, A. B., Langmuir 2009, 25, (11), 6328-33. 60. Feng, W.; Gao, X.; McClung, G.; Zhu, S.; Ishihara, K.; Brash, J. L., Acta Biomater. 2011, 7, (10), 3692-3699. 61. Ostuni, E.; Chapman, R. G.; Holmlin, R. E.; Takayama, S.; Whitesides, G. M., Langmuir 2001, 17, (18), 5605-5620. 62. Trmcic-Cvitas, J.; Hasan, E.; Ramstedt, M.; Li, X.; Cooper, M. A.; Abell, C.; Huck, W. T. S.; Gautrot, J. E., Biomacromolecules 2009, 10, (10), 2885-2894. 63. Unsworth, L. D.; Sheardown, H.; Brash, J. L., Langmuir 2005, 21, (3), 1036-1041. 64. Hucknall, A.; Rangarajan, S.; Chilkoti, A., Adv. Mater. 2009, 21, (23), 2441-2446. 65. Rodriguez-Emmenegger, C.; Brynda, E.; Riedel, T.; Houska, M.; Subr, V.; Alles, A. B.; Hasan, E.; Gautrot, J. E.; Huck, W. T., Macromol. Rapid Commun. 2011, 32, (13), 952-7. 66. Jiang, S.; Cao, Z., Adv. Mater. 2010, 22, (9), 920-32. 67. Vaisocherova, H.; Sevcu, V.; Adam, P.; Spackova, B.; Hegnerova, K.; de los Santos Pereira, A.; Rodriguez-Emmenegger, C.; Riedel, T.; Houska, M.; Brynda, E.; Homola, J., Biosens. Bioelectron. 2014, 51, 150-7. 68. Riedel, T.; Hageneder, S.; Surman, F.; Pop-Georgievski, O.; Noehammer, C.; Hofner, M.; Brynda, E.; Rodriguez-Emmenegger, C.; Dostálek, J., Anal. Chem. 2017, 89, (5), 2972-2977. 69. Riedel, T.; Surman, F.; Hageneder, S.; Pop-Georgievski, O.; Noehammer, C.; Hofner, M.; Brynda, E.; Rodriguez-Emmenegger, C.; Dostalek, J., Biosens. Bioelectron. 2016, 85, 272-9. 70. Rodriguez-Emmenegger, C.; Surman, F.; Brynda, E.; Riedel, T.; Houska, M.; Lisalova, H.; Homola, J. Copolymer of N-(2-hydroxypropyl) methacrylamide and carboxybetaine methacrylamide, polymer brushes. WO2016177354, November 10, 2016. 71. Vaisocherová-Lísalová, H.; Surman, F.; Víšová, I.; Vala, M.; Špringer, T.; Ermini, M. L.; Šípová, H.; Šedivák, P.; Houska, M.; Riedel, T.; Pop-Georgievski, O.; Brynda, E.; Homola, J., Anal. Chem. 2016, 88, (21), 10533-10539. 72. Edmondson, S.; Osborne, V. L.; Huck, W. T. S., Chem. Soc. Rev. 2004, 33, (1), 14-22. 73. Zhao, B.; Brittain, W. J., Prog. Polym. Sci. 2000, 25, (5), 677-710. 74. Moad, G.; Rizzardo, E.; Thang, S. H., Chem Asian J 2013, 8, (8), 1634-44. 75. Percec, V.; Popov, A. V.; Ramirez-Castillo, E.; Weichold, O., J. Polym. Sci., Part A: Polym. Chem. 2003, 41, (21), 3283-3299. 76. Rosen, B. M.; Percec, V., Chem. Rev. 2009, 109, (11), 5069-5119. 77. Lligadas, G.; Grama, S.; Percec, V., Biomacromolecules 2017, 18, (10), 2981-3008. 78. Samanta, S. R.; Cai, R.; Percec, V., Polym. Chem. 2014, 5, (18), 5479-5491. 79. Jones, G. R.; Anastasaki, A.; Whitfield, R.; Engelis, N.; Liarou, E.; Haddleton, D. M., Angew. Chem. Int. Ed. Engl. 2018, 57, (33), 10468-10482. 80. Nguyen, N. H.; Levere, M. E.; Percec, V., J. Polym. Sci., Part A: Polym. Chem. 2012, 50, (5), 860-873. 81. Nguyen, N. H.; Percec, V., J. Polym. Sci., Part A: Polym. Chem. 2011, 49, (22), 47564765. 82. Anastasaki, A.; Nikolaou, V.; Haddleton, D. M., Polym. Chem. 2016.


28

Page 29 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

83. Anastasaki, A.; Nikolaou, V.; Nurumbetov, G.; Wilson, P.; Kempe, K.; Quinn, J. F.; Davis, T. P.; Whittaker, M. R.; Haddleton, D. M., Chem. Rev. 2015. 84. Grama, S.; Lejnieks, J.; Enayati, M.; Smail, R. B.; Ding, L.; Lligadas, G.; Monteiro, M. J.; Percec, V., Polym. Chem. 2017, 8, 5865-5874. 85. Jezorek, R. L.; Enayati, M.; Smail, R. B.; Lejnieks, J.; Grama, S.; Monteiro, M. J.; Percec, V., Polym. Chem. 2017, 8, 3405-3424. 86. Moreno, A.; Galià, M.; Lligadas, G.; Percec, V., Biomacromolecules 2018, 19, (11), 44804491. 87. Smail, R. B.; Jezorek, R. L.; Lejnieks, J.; Enayati, M.; Grama, S.; Monteiro, M. J.; Percec, V., Polym. Chem. 2017, 8, 3102-3123. 88. Anastasaki, A.; Nikolaou, V.; Zhang, Q.; Burns, J.; Samanta, S. R.; Waldron, C.; Haddleton, A. J.; McHale, R.; Fox, D.; Percec, V.; Wilson, P.; Haddleton, D. M., J. Am. Chem. Soc. 2014, 136, (3), 1141-1149. 89. Anastasaki, A.; Oschmann, B.; Willenbacher, J.; Melker, A.; Van Son, M. H. C.; Truong, N. P.; Schulze, M. W.; Discekici, E. H.; McGrath, A. J.; Davis, T. P.; Bates, C. M.; Hawker, C. J., Angew. Chem. 2017, 56, (46), 14483-14487. 90. Discekici, E. H.; Anastasaki, A.; Kaminker, R.; Willenbacher, J.; Truong, N. P.; Fleischmann, C.; Oschmann, B.; Lunn, D. J.; Read de Alaniz, J.; Davis, T. P.; Bates, C. M.; Hawker, C. J., J. Am. Chem. Soc. 2017, 139, (16), 5939-5945. 91. Lligadas, G.; Grama, S.; Percec, V., Biomacromolecules 2017, 18, (4), 1039-1063. 92. Laun, J.; Vorobii, M.; De Los Santos Pereira, A.; Pop-Georgievski, O.; Trouillet, V.; Welle, A.; Barner-Kowollik, C.; Rodriguez-Emmenegger, C.; Junkers, T., Macromol. Rapid Commun. 2015, 36, (18), 1681-1686. 93. Vorobii, M.; Pop-Georgievski, O.; de los Santos Pereira, A.; Kostina, N. Y.; Jezorek, R.; Sedláková, Z.; Percec, V.; Rodriguez-Emmenegger, C., Polym. Chem. 2016, 7, 6934-6945. 94. Vorobii, M.; de los Santos Pereira, A.; Pop-Georgievski, O.; Kostina, N. Y.; RodriguezEmmenegger, C.; Percec, V., Polym. Chem. 2015, 6, (23), 4210-4220. 95. Obstals, F.; Vorobii, M.; Riedel, T.; de los Santos Pereira, A.; Bruns, M.; Singh, S.; Rodriguez-Emmenegger, C., Macromol. Biosci. 2018, 18, (3). 96. Kostina, N. Y.; Rodriguez-Emmenegger, C.; Houska, M.; Brynda, E.; Michálek, J., Biomacromolecules 2012, 13, (12), 4164-4170. 97. Ulbrich, K.; Šubr, V.; Strohalm, J.; Plocová, D.; Jelıń ková, M.; Řıh́ ová, B., J. Control. Release 2000, 64, (1–3), 63-79. 98. Grama, S.; Plichta, Z.; Trchová, M.; Kovářová, J.; Beneš, M.; Horák, D., React. Funct. Polym. 2014, 77, 11-17. 99. Colthup, N., Introduction to Infrared and Raman Spectroscopy. Academic Press: 2012; p 562. 100. Di Noto, V.; Bettinelli, M.; Furlani, M.; Lavina, S.; Vidali, M., Macromol. Chem. Phys. 1996, 197, (1), 375-388. 101. Smith, B. C., Infrared Spectral Interpretation: A Systematic Approach. CRC Press: 1998; p 288 102. Socrates, G., Infrared and Raman Characteristic Group Frequencies: Tables and Charts. Wiley: 2004; p 366. 103. Leng, C.; Sun, S.; Zhang, K.; Jiang, S.; Chen, Z., Acta Biomater 2016, 40, 6-15.


29


Page 30 of 30

TOC


30

Antifouling Microparticles To Scavenge ... - ACS Publications

Recommend Documents