CN101065134A - Polymeric compositions and related methods of use - Google Patents

Abstract

The present invention provides adhesive polymer compositions that may contain dihydroxyphenyl moieties and derivatives thereof, and related methods of use.

Description

Polymer compositions and related methods of use

Cross References to Related Applications

This application is a continuation-in-part of U.S. Patent Application 10/199,960, filed July 19, 2002, which requires U.S. Patent Application 60, filed July 20, 2001, and April 19, 2002, respectively Priority of /306,750 and 60/373,919. This application also claims priority to US Patent Application 60/548,314, filed February 27, 2004, and US Patent Application 60/549,259, filed March 2, 2004. Each patent is incorporated herein by reference in its entirety.

Statement Regarding Federally Sponsored Research or Development

The United States Government has certain rights in this invention pursuant to grants DE13030, DE12599, and DE14193 to Northwestern University from the National Institutes of Health and grant NCC-1-02097 to Northwestern University from NASA.

Background technique

Mussel adhesive protein (MAP) is a very important adhesion material in water, which forms a strong connection between mussels and the surface on which the mussels reside. During attachment to the surface, MAP is secreted as a liquid and undergoes a cross-linking or hardening reaction to form a solid plaque. One of the unique features of MAP is the presence of L-3,4-dihydroxyphenylalanine (DOPA), a rare amino acid that is believed to be responsible, at least in part, of MAP's resistance to substrates by a mechanism that is not fully understood. bonding. Mussels adhere to a variety of surfaces including metals, metal oxides, polymers, plastics and wood.

For biosensors, medical diagnostic products, all equipment and tests requiring handling of serum and other human/animal body fluids, tissue engineering, localized drug delivery in vivo, implanted medical devices, healing of surgical incisions, tissue adhesion for rehabilitation purposes Controlling the adhesion of cells and proteins to surfaces is crucial, for example in the bonding of bone and cartilage and in nanotechnology (nanoparticle-based therapeutic and diagnostic tools). Controlling the adhesion of cells and proteins to surfaces is also important in many industrial applications. These applications include preventing mussels from adhering to boats, boats, piers and other structures used in marine and fresh water, preventing the growth of algae and bacteria on water lines for industrial and drinking water, and on sensors used to monitor water quality and purity .

In the medical field, physical or chemical immobilization of substances such as poly(alkylene oxide) (PAO), polyethylene glycol (PEG), polyepoxide Ethane (PEO) and PEO-PPO-PEO block copolymers, such as those available under the trade name PLURONICS, and polymers such as PEG/tetraethylene glycol dimethyl ether, poly(methoxyethyl methacrylate) (PMEMA) and poly(methacrylphosphatidylcholine) (polyMPC) (E.W.Merrill, Ann.NY Acad.Sci., 516, 196 (1987); Ostuni et al., Langmuir 2001, 17, 5605-20, incorporated herein by reference) . Current methods for modifying surfaces with polymers must be tailored to each material type and thus require different chemical strategies. For example, the surfaces of noble metals such as platinum, silver, and gold can be modified with molecules containing sulfhydryl groups (-SH), while metal oxides are often modified with silane coupling chemistry. No one surface modification strategy is universal for different types of materials. Moreover, many existing methods rely on expensive equipment or complex synthetic methods, or both.

Summary of the invention

The present invention relates to compositions such as adhesives in a substantially aqueous environment. Preferred compositions generally comprise an adhesive portion and a polymeric portion having the desired surface active effect (or other desired property).

In one aspect, the viscous portion of the composition of the present invention comprises dihydroxyphenyl derivatives, including di(DHPD), wherein the second DHPD is

That is, a methylene derivative of dihydroxyphenyl.

In another aspect, the polymer portion contains poly(alkylene oxide). In a highly preferred embodiment, the adhesive portion comprises DHPD, eg, DOPA (as described herein), and the polymer portion comprises a PEO-PPO-PEO block polymer (as described above).

In another preferred embodiment, the adhesive moiety comprises DHPD comprising side chains containing ethylenic or vinylic unsaturation, eg alkyl acrylates.

Detailed description of the invention

More specifically, the present invention includes dihydroxyphenyl (DHPD) sticky compounds of formula (I),

In the formula, R ₁ and R ₂ may be the same or different, each independently selected from hydrogen, saturated and unsaturated, linear and branched, substituted and unsubstituted C _1-4 hydrocarbon groups;

P is individually and independently selected from -NH ₂ , -COOH, -OH, -SH,

In the formula, R ₁ and R ₂ are as defined above.

single bond, halogen,

In the formula, A ₁ and A ₂ are individually and independently selected from H, a single bond;

protecting group,

roughly poly(alkylene oxide),

where n is 1 to about 3,

A ₃ is

R ₄ is H, C _1-6 lower alkyl, or

Polyalkylene oxide

R ₃ is as defined above, and D is as shown in formula (I).

In one aspect, the poly(alkylene oxide) has the structure,

In the formula, R ₃ and R ₄ are individually and independently selected from H or CH ₃ , the value of m is between 1 and about 250, A ₄ is NH ₂ , COOH, -OH, -SH, -H or a protecting group group.

A highly preferred form of DHPD is

R ₁ , R ₂ and P are as defined above.

A further preferred form of DHPD has the following structure:

In the formula, A ₂ is -OH, and A ₁ is poly(alkylene oxide) of roughly the following structure,

R ₃ , R ₄ and m are as defined in claim 2. Generally, poly(alkylene oxide)s are block copolymers of ethylene oxide and propylene oxide.

A method of the invention comprising adhering a substrate to another substrate comprises the step of providing a DHPD having the following structure:

In the formula, _R and _R are as defined above; the DHPD of the above structure is applied to one or the other or both substrates to be adhered; the substrates to be adhered are brought into contact with the DHPD of the above structure located between them To adhere the substrates to each other, the substrates are optionally repositioned relative to each other by separating the substrates and bringing the substrates into contact with each other again with the DHPD of the above structure located between the substrates.

In preferred methods, _R1 and _R2 are hydrogen.

Definition: For the purpose of this application, the dihydroxyphenyl derivative (DHPD) of the present invention means a dihydroxyphenyl derivative of the following structure:

In the formula, P, R ₁ and R ₂ are as follows, and n is 1 to about 5. In one embodiment, _R1 and _R2 are hydrogen and P itself is dihydroxyphenyl. A preferred DHPD in this embodiment of the invention is L-3,4, dihydroxyphenylalanine (DOPA), (typically),

In the formula, A ₁ and A ₂ are as defined above.

As used herein, "substantially poly(alkylene oxide)" means that the composition is predominantly or predominately alkyl oxide or alkyl ether. This definition takes into account the presence of heteroatoms such as N, O, S, P, etc., functional groups such as -COOH, _-NH2 , -SH and alkenyl or vinyl unsaturations as well as functional groups. It should be understood that these non-alkylene oxide structures can only be present in relative abundance so as not to substantially diminish the desirable overall surface activity, non-toxicity or immune response properties of the polymer.

Brief description of the drawings

Figure 1 shows ^the1H NMR spectra of PLURONIC(R) F127, its carbonate intermediate (SC-PAO7) and DME-PAO7 in _CDCl3 .

Figure 2 provides differential scanning calorimetry graphs of 30 wt% aqueous solutions of DME-PAO7, DOPA-PAO7 and unmodified PLURONIC(R) F127. Arrows indicate the location of the gelation endotherm.

Figure 3 shows the shear storage modulus, G', plotted as a function of temperature for a 22 wt% aqueous solution of DME-PAO7 at 0.1 Hz and 0.45% strain. Shown in the inset is a graph of the rheological profile of unmodified PLURONIC(R) F127 in a 22% by weight aqueous solution as a function of temperature.

Figure 4 shows the shear storage modulus G' plotted as a function of temperature for a 50% by weight aqueous solution of DME-PAO8 at 0.1 Hz and 0.45% strain. Shown in the inset is a graph of the rheological profile of unmodified PLURONIC(R) F68 in a 50% by weight aqueous solution as a function of temperature.

Figure 5 shows the storage modulus plotted as a function of temperature for 45% or 50% by weight aqueous solutions of DME-PAO8 at 0.1 Hz and 0.45% strain, respectively.

Figures 6A to 6B show differential scanning calorimetry maps of (A) DOPA-PAO7 and (B) DME-PAO7 at different concentrations upon heating. Arrows indicate the location of the gelation endotherm observed only at higher polymer concentrations.

Figure 7A-C shows high resolution C(ls) XPS peaks of (A) unmodified Au, (B) m-PEG-OH and (C) m-PEG-DOPA. A significant increase in the ether peak at 286.5 eV in (C) indicates the presence of PEG.

Figures 8A-C provide TOF-SIMS positive spectra showing peaks representing gold binding to catechol. The spectrum was normalized with the Au peak (m/z~197).

Figure 9 provides TOF-SIMS spectra, showing the positive secondary ion peaks of unmodified Au substrate, Au exposed to mPEG-OH, mPEG-DOPA powder or mPEG-DOPA at mass m/z~43.

Figure 10 represents TOF-SIMS spectra showing positive secondary ion peaks for Au substrates chemisorbed with mPEG-DOPA. Gold binding to catechol was observed at m/z~225 (AuOC), 254 (AuOCCO) and 309. Weaker AuO _a C _b peaks were observed at m/z~434, 450, 462 and 478. The periodic triplet observed in the range of m/z 530–1150 corresponds to Au bound to DOPA-( _CH2CH2O ) _n , where the individual _subpeaks are separated by 14 or 16 amu, representing the _CH2 , _{CH2CH2 ,} and _{CH2CH2O .} This pattern was observed for n=1-15.

Figure 11 shows the SPR spectra of proteins (0.1 mg/ml BSA) adsorbed to modified and unmodified gold surfaces. The mPEG-DOPA- and mPEG-MAPd-modified surfaces showed reduced adsorption of proteins compared to bare gold and mPEG-OH-modified surfaces.

Figure 12 shows mPEG-DOPA concentration dependence of antifouling behavior. Gold surfaces were modified in the indicated mPEG-DOPA concentrations for 24 hours and then the density and area of adhered cells were analyzed. ( ^* =p<0.05, ^** =p<0.01, ^*** =p<0.001; black bars=total proj.area, gray bars=surface cell density)

Fig. 13 compares under optimal condition (50mg/ml, 24 hours), the gold surface of bare gold, the gold of mPEG-OH treatment, with the gold surface of mPEG-DOPA 5K, mPEG-MAPd 2K and mPEG-MAPd 5K treatment. Adhesion and diffusion. (Black bars=total protected area, gray bars=surface cell density, ^*** =p<0.001).

Figure 14A-C is a series of SEM micrographs showing the growth of NIH 3T3 fibroblasts on (A) unmodified Au, (B) Au treated with mPEG-OH, and (C) mPEG-DOPA modified Au. form. All treatments were at 50 mg/ml in DCM for 24 hours.

Figure 15 shows the UV/vis absorption spectra of mPEG-DOPA stabilized magnetic nanoparticles suspended in several concentrations of NaCl aqueous solution as indicated. Addition of NaCl did not cause nanoparticle deposition.

Figure 16 shows that addition of salt to untreated Au nanoparticles induces aggregation. Shown are UV/vis scans of 10 nm untreated Au nanoparticles suspended in (indicated concentrations) aqueous NaCl solution. The weakening and shifting of the 520 nm absorption band with increasing NaCl concentration reflects the aggregation of nanoparticles.

Figure 17 shows that addition of salt to mPEG-DOPA stabilized Au nanoparticles does not induce aggregation. Shown are UV/vis scans of 10 nm mPEG-DOPA-stabilized Au nanoparticles suspended in (indicated concentrations) aqueous NaCl. As the NaCl concentration increased, the 520 nm absorption band did not weaken and shift, reflecting the effective stabilization of nanoparticles.

Figure 18 shows the UV/vis absorption spectra of mPEG-DOPA stabilized CdS nanoparticles suspended in (indicated concentrations) aqueous NaCl solution.

Figure 19 shows the XPS survey scans of unmodified TiO ₂ and TiO ₂ treated with mPEG-DOPA _1-3 .

Figure 20 shows long-term resistance to cell adhesion on _TiO2 and mPEG-DOPA _1-3 treated _TiO2 . The duration of the foul-free reaction is proportional to the length of the DOPA peptide anchoring group. Adhered cells were visualized with calcium AM.

Figure 21 shows a high resolution XPS scan of the Cls region of the mPEG-DOPA _1-3 modified _TiO2 substrate. Note the increase in the ether carbon peak (286.0 eV) as the length of the DOPA peptide anchor group increases.

Figure 22 shows a high resolution XPS scan of the Ols region of the mPEG-DOPA _1-3 modified _TiO2 substrate. With the increase of DOPA peptide length, the peak representing polymerization oxygen at 532.9eV increased, while the Ti-OH peak (531.7eV) decreased.

Figure 23 shows a graph of the results of a Robust design experiment on 316L stainless steel.

Figure 24 shows a 4 hour graph of cell adhesion on various surfaces modified with mPEG-DOPA _1-3 for 24 hours at 50°C and the indicated pH values.

Figure 25 shows the percentage of gel conversion plotted against UV exposure time (minutes).

Figure 26 shows the mole fraction of DOPA incorporated versus the mole % of 1 or 7 in the precursor solution.

Figure 27 shows the percentage of gel conversion plotted against 1 or 7 mole % in the precursor solution.

Figure 28 shows X-ray photoelectron spectroscopy XPS analysis of a silicon nitride surface.

Figure 29 shows free monitoring of a functionalized silicon nitride cantilever.

Figure 30 is an entropic elasticity analysis of polyethylene glycol.

Figure 31 is a force assay of side chain modified DOPA.

Figure 32 Possible model for DOPA-T ₁ O ₂ binding mechanism.

Figure 33 is an arrangement of atomic force micrographs.

Figure 34 is the relevant data of strength measurement.

Figure 35 is sticky data.

Figure 36 is the synthesis path and data analysis.

These dihydroxyphenyl derivative ("DHPD") binders function in aqueous environments. To form the polymer composition, usually the DHPD moiety providing the tackiness functionality is coupled to a polymer providing the desired surface active effect. These components will be described in more detail below.

These viscous, polymeric compositions have many uses, including preventing the adhesion of proteins and/or cells to surfaces in various medical, industrial and consumer applications. DHPD can also be used as a substitute for wound sutures to aid in the healing of fractures or cartilage-to-bone injuries. These and other applications are described in more detail below.

Preferred polymer compositions of the present invention have the following structure:

wherein, for each compound of formula (1a), R _{and R} are individually and independently as defined above,

P _{and P are} individually and independently as defined for P in formula (I);

n and m are independently 0 to about 5, provided that at least one of n or m is at least 1;

sticky part

The sticky moiety of the present invention is a dihydroxyphenyl derivative ("DHPD") having the following preferred structure:

wherein R ₁ , R ₂ and P are as defined above, t is 1 to about 10, preferably about 1-5, most preferably 1 to about 3. DHPD bonding works in aqueous environments. Herein, an aqueous environment is any medium that contains water. It includes, but is not limited to, water, including saline and fresh water, cell and bacterial growth media solutions, aqueous buffers, other based aqueous solutions and body fluids. The DHPD moiety can be derivatized. Those skilled in the art understand that such derivatization is limited by the desired adhesion properties to be retained.

polymer component

Those skilled in the art with knowledge of the present invention will be familiar with the various polymer components that provide surface active effects and other desirable properties. The desired surface active effect is associated with reduced particle agglomeration and resistance to biofouling, including resistance to cell and/or protein adhesion. For example, depending on the end use, the polymer component may be water soluble, and/or depending on various other end uses, the polymer component may be capable of forming micelles. Polymers that can be used in the present invention include, but are not limited to, polyethylene glycol (PEG), polyethylene oxide (PEO), polypropylene oxide (PPO), PEO-PPO-PEO block copolymers, polyphenylene oxide , PEG/tetraethylene glycol dimethyl ether, PMEMA, polyMPC and perfluorinated (perfluorunated) polyethers. Polymer compositions can be synthesized in several ways. For example, the polymer composition can be synthesized by general synthetic methods for activating polymer end groups. Carbonate chemistry can be used to activate various polymers or their monomeric components. Specifically, the reaction of succinimide carbonate activated polymer components with DHPD moieties provides stable urethane conjugates. Two of the many possible routes, pathways (a) and (b) in Scheme 1a, 1b below, show coupling to poly(alkylene oxide) in aqueous and non-aqueous solvents without destroying the desired bioadhesion. For example, by forming urethane or amide bonds, DHPD residues can be coupled to polymer components to provide the desired coupling composition. These synthetic routes are shown in Schemes 1a and 1b and described in more detail below.

Option 1a

Option 1b

More specifically, a variety of other functional groups can be used to esterify or derivatize the carboxylic acid groups of the DHPD component if coupled to the polymer component via urethane bond formation. Alternatively, the DHPD component can be coupled to the polymer component (e.g., amidation or esterification can occur depending on the polymer end group, _-NH2 or -OH), providing a variety of known protecting groups that can be used. Derivatized DHPD functional groups, said protecting groups include, but are not limited to, Boc, Fmoc, borate, phosphate, and tributyldimethylsilyl. N-terminal protection of the DHPD component can enable the use of carboxylic acid groups for multifunctional derivatization and/or coupling of higher density polymer components.

Accordingly, as part of the present invention there is also provided a method of incorporating DHPD residues into polymer systems using urethane synthesis. The method comprises (1) providing a polymer component whose terminal is a plurality of monomers, each monomer having a terminal functional group; (2) preparing a carbonate derivative of the polymer component; and (3) the The carbonate derivative is reacted with at least one DHPD moiety to produce a urethane moiety. As noted above, the polymeric components used in the process may include polymeric components having terminal monomeric functionalities that can be reacted with a reagent to provide the desired carbonate derivative and ultimately the polymeric The urethane moiety to which the component is coupled with the DHPD component. Various other coupling agents and/or hydroxyl terminated polymer components may also be used to provide the desired urethane moieties.

It is also part of the present invention to provide methods of using carbonate intermediates to maintain catechol functionality, or to enhance adhesion, of DHPD-incorporated polymer compositions and/or systems. The method comprises (1) providing a polymer component terminated by a plurality of monomers, each monomer having a terminal functional group; (2) reacting the polymer component with a reagent to provide a carbonate intermediate; and ( 3) The carbonate intermediate is partially reacted with at least one DHPD. Without being bound by any theory or embodiment, the method of the present invention can be considered as a method of enhancing the end group reactivity of the polymer component through a suitable carbonate intermediate. Subsequent reactions at the amino nitrogen of the DHPD moiety provided the corresponding conjugates while maintaining the catechol functionality.

According to the present invention, various synthetic routes can be used to couple the DHPD moiety to the carbonate activated intermediate as shown in Scheme 1a. DOPA methyl ester (DME) obtained by reacting DOPA with methanol in the presence of thionyl chloride can be used in an organic solvent. The progress of the reaction can be monitored by TLC and NMR, and the coupling reaction is actually completed within 1 hour (representative conjugates are DME-PAO7 (from PAO PLURONIC(R) F127) and DME-PAO8 (from PAO PLURONIC(R) F68)). After purification with cold methanol, high product yields were obtained.

The free carboxyl form of DOPA can be coupled to the carbonate intermediate in aqueous base. It is well known that the major difficulty in handling DOPA is that it is readily oxidized (to DOPA quinone and other products), which occurs readily in aqueous alkaline solutions. To prevent the undesired oxidation of the catechol side chains of DOPA during the coupling process under alkaline conditions, DOPA can be first added to aqueous sodium borate to form borate-protected DOPA (Scheme 1b). The resulting complexes are very stable in neutral or basic solutions and are easily deprotected under acidic conditions. Using the complexation of DOPA and borate, DOPA was coupled to the ends of several commercially available PAOs under basic conditions to generate DOPA-PAO7 and DOPA-PAO8. Visual inspection of the reaction solution revealed the absence of strongly absorbing DOPA-quinone, indicating that DOPA was not oxidized during the reaction. At the end of the reaction, the DOPA end groups of the block copolymer were deprotected by acidification with HCl.

¹ H NMR spectroscopy and colorimetric measurements confirmed the composition of the succinimide-activated reaction intermediate in Scheme 1 and all four DOPA-modified PAOs. Figure 1 shows the ¹ H NMR spectra of PAO PLURONIC(R) F127, the succinimide carbonate activated intermediate (SC-PAO7) and the corresponding DOPA methyl ester modified PAO (with PLURONIC(R) F127, DME-PAO7). In the ¹ H NMR spectrum of DOPA-containing PAO, the -CH ₂ - proton from the succinimidyl carbonate group forms a sharp peak at ~2.8 ppm, and the -CH ₂ -O- proton (from the activated PAO The sharp peak formed at -4.4 ppm (the only oxirane group adjacent to the carbonate group) completely disappeared, while a series of new peaks appeared due to the partial incorporation of DOPA into the copolymer. A feature of the ¹ H NMR spectrum of DOPA-containing PAOs is the appearance (corresponding to the three protons of the DOPA benzene ring) of a singlet and two doublets in the range 6.5-6.9 ppm. Similar features were also observed in the ¹ H NMR spectrum (not shown) of the DOPA-PAO conjugate synthesized in aqueous solution.

Based on the assumption that there are two succinimidyl carbonate groups present in the corresponding carbonate intermediates SC-PAO7 and SC-PAO8, DOPA methyl ester and DOPA pairs were quantitatively determined by colorimetric analysis. The coupling efficiency of POA was 76%-81% (Table 1). The reported coupling efficiencies are the average of at least 3 replicated syntheses under the same conditions, and no significant increase in coupling efficiency was observed when a greater excess of DOPA was used in the reaction. Similar coupling efficiencies were also observed for DOPA-PAO7 and DOPA-PAO8 prepared from _aqueous solutions, indicating that the hydrolysis of succinimidyl carbonate-activated PAOs was slow in _aqueous alkaline solutions containing _Na2B4O7 .

In contrast to the coupling efficiency, the product yields of selective DOPA-modified PAOs synthesized in aqueous solution (as shown in Figure 1) were lower than those of selective DOPA-modified PAOs synthesized in organic solvents. This may be due to the inefficient extraction of DOPA-modified PAO from water with dichloromethane due to the surfactant properties of the starting PAO material. It should be noted that the free carboxylic acids of DOPA-PAO7 and DOPA-PAO8 can be further functionalized using standard peptide chemistry to tailor the properties of the block copolymers. The four DOPA-modified PAOs in Table 1 can be stored at -20°C, probably without discoloration or changes in properties.

Table 1

COUPLING EFFICIENCY AND PRODUCT YIELD OF DOPA MODIFIED PLURONIC ® Coupling efficiency (%) ^* Product yield (%) DME-PAO7DOPA-PAO7DME-PAO8DOPA-PAO8 78.0±4.080.0±4.076.0±2.081.0±2.0 75.0±5.052.0±3.076.0±4.049.0±2.0

^* As disclosed by Waite and Benedict (Waite, JH & Benedict, CV Determination of dihydroxyphenylalanine (dopa) in invertebrate structural proteins, Methods in Enzymology 107, 397-413 (1984), incorporated herein by reference) Colorimetric assay.

For the performance of biosensors, medical diagnostic products, all equipment and tests that require handling of serum and other human/animal body fluids, tissue engineering, localized drug delivery in vivo, implanted medical devices, healing of surgical incisions, tissues for rehabilitation purposes Adhesion such as that of bone and cartilage, as well as nanotechnology (nanoparticle-based therapeutic and diagnostic tools), controlled adhesion of cells and proteins to surfaces is critical. Controlling the adhesion of cells and proteins to surfaces is also important in many industrial applications. These applications include preventing mussels from adhering to boats, boats, piers and other structures used in marine and fresh water, preventing the growth of algae and bacteria on water lines for industrial and drinking water, and on sensors used to monitor water quality and purity .

The polymer compositions of the present invention are useful as coatings to prevent adhesion of proteins and cells to devices used in medical and research applications, including but not limited to coatings for medical implants, coatings for surgical devices , coatings for devices that process serum and other animal or human-derived substances, coatings for medical diagnostic devices, and coatings for biosensors. Alternatively, the polymer composition may be a tissue-adhesive polymer hydrogel for medical applications such as tissue sealing, gels for preventing surgical adhesion (scar tissue formation), bone and cartilage adhesives, tissue Engineering, site-specific drug efflux, and for research uses such as immobilization of proteins (including antibodies) and small molecule analytes (including drugs). Additionally, these coatings and hydrogels have many uses in industrial and consumer products including, but not limited to, preventing biofouling (seaweed, bacteria, and mussels sticking to underwater surfaces) in marine shipments, preventing runoff to factories such as Bacterial contamination in water streams in power and pharmaceutical plants, prevention of bacterial contamination in drinking water, as dental and denture adhesives, underwater adhesives for delivery indicators, coatings for water purity and detection sensors, for protection against biological Dirt paints, used in cosmetics to adhere desired fragrances and colorants to hair, eyelids, lips, and skin for temporary coloring of the skin, such as tattoos, etc., and in consumer products such as storage bags Resealable adhesive. The method of the invention can be used to prepare a variety of polymer-modified surfaces for medical (diagnostics, devices, nanoparticle-based therapy) and non-medical (paints and other particle dispersions, MEMS, quantum dots, dirt-free surface) technology.

Viscous hydrogels can also be formed using the methods of the invention. The DHPD adhesive is attached to a polymer that can form a hydrogel in vivo or in vitro. These hydrogels can be formed in a number of ways, including using self-assembled polymers that form gels at elevated temperatures, such as the normal temperature of the human body, using polymers that can be cross-linked by enzymatic reactions, using polymers that can be oxidized to form cross-linked hydrogels polymers, and the use of polymers that can be photoactivated to produce cross-linked hydrogels.

Anti-Biofouling Coating

The anti-biofouling coatings of the present invention can be applied to medical devices such as vascular or arterial stents, pacemakers, heart valves, glucose monitors and other biosensors, vascular sheaths, defibrillators, orthopedic and surgical devices, Includes sutures and catheters. The polymer compositions of the present invention are useful as coatings to prevent adhesion of proteins and/or cells to devices used in medical or research applications. These applications include, but are not limited to, use as coatings for medical implants, coatings for surgical devices, coatings for devices handling serum and other animal or human derived substances, medical diagnostic devices, and biosensors. The challenges of modifying biomaterial surfaces with polymers to resist cell adhesion include creating sufficiently high densities of polymers capable of repelling proteins and cells, and creating coatings that completely cover the surface. This is especially problematic in devices that contain components made of multiple different materials. Surfaces may be modified with the polymers of the present invention in any number of ways. For example, the polymer composition, or the DHPD moiety containing the polymerization initiator, can be adsorbed to a surface from which the growth of the polymer begins. In the latter case, a variety of polymerization techniques can be employed including, but not limited to, surface-initiated free-radical polymerization, free-radical polymerization methods, ionic polymerization, ring-opening polymerization, and photopolymerization.

For example, taking advantage of the unique solubility of PEG, the surface can be treated with a PEG solution at a temperature close to the lower critical solution temperature (LCST) or cloud point to increase the density of the polymer. While not being bound by any theory, applicants believe that under the conditions of high ionic strength and elevated temperature used in the present invention, the hydrodynamic radius of the PEG molecule is reduced, allowing, in principle, higher densities than under standard conditions The PEG chains accumulate onto the surface. This method can be used for polymers that exhibit solubility reversal at high temperature and high ionic strength, for example, polyethylene glycol, poly(N-isopropylacrylamide), and other N-substituted poly(acrylamides) that exhibit solubility reversal of polymers.

Resistance to cell and protein adhesion up to 7 days, 14 days, 21 days, 30 days, 60 days, 90 days and 120 days or longer was achieved by modifying the surface with various polymer compositions of the invention. Most of the variation in the resistance of the modified material to cell and/or protein adhesion is due to the number of DHPD moieties in the viscous fraction and the pH of the modification buffer. For surfaces modified by adsorption, the adsorption time and the concentration of the polymer composition have little effect on the variation of the modified material's resistance to cell and/or protein adhesion. The greater the number of DHPD moiety monomers in the adhesive component, the greater the resistance to cell and/or protein adhesion. The density of the polymer composition on the surface correlates strongly with the resistance to cell and/or protein adhesion. The thickness of the coating can be from about 20 A to 100 μm inclusive, depending on the polymer composition used and the pH of the modification buffer.

The concentration of the polymer composition used to modify the surface can be from about .1 mg/ml to 75 mg/ml. The pH of the modification buffer can be about 3-9. The grooming time can be from about 10 minutes to about 72 hours. The temperature used in the modification process may be from about 25°C to 60°C.

As shown in Figure 19, the XPS detection scan of unmodified TiO ₂ has strong peaks at ~458eV (Ti2p) and ~530eV (Ols), which represent the characteristics of the initial oxide, and there is also a small peak at 248.7eV (Cls), which is Alien hydrocarbon contamination. But _TiO2 treated with mPEG-DOPA _1–3 under cloud point conditions showed a significant increase in surface-bound carbon (as shown by Cls), indicating the presence of PEG on the surface. Furthermore, the increase in the Cls peak observed after treatment with mPEG-DOPA _1-3 was directly proportional to the amount of terminal DOPA present. In addition, a small peak at 400 eV (Nls) was observed in the spectra of _the TiO surface modified with mPEG-DOPA _1–3 , representing the amide nitrogen in DOPA.

Quantitative analysis of high resolution XPS data of the substrate surface can provide useful information on the relative amount of PEG bound to the surface. Table 2 shows the atomic composition calculations of titanium, oxygen and carbon in mPEG-DOPA _1–3 modified TiO ₂ . Oxygen signals are further divided into subclasses such as metal oxides (Ti-O-Ti), surface hydroxides (Ti-OH) and organic oxygen and coupled water (CO, H ₂ O).

Table 2

Atomic composition calculations for titanium, oxygen and carbon ^Atomic compositiona (wt%) o surface Ti Ti-O-Ti Ti-OH CO, _H2O C Unmodified TiO ₂ 31.0 47.0 7.1 ^3.9b 11.1 MPEG-DOPA 15.5 24.5 6.2 14.9 38.9 MPEG-DOPA ₂ 12.3 19.4 5.4 18.3 44.6 MPEG-DOPA ₃ 11.5 17.3 4.7 20.6 45.9

^a Ignoring trace amounts of nitrogen.

^b Assumed to be water bound to the surface.

The Ti:Ti-O-Ti ratios of all substrates differed significantly from the theoretical stoichiometric value of 2.0; this difference may be due to the sampling depth exceeding the depth of the surface oxide (3-4 mm). This result was expected considering the Flory radius of 5000 _Mw PEG (2.8 mm) and the typical XPS sampling depth of 5-10 mm. On the mPEG-DOPA _1-3 modified surface, as the DOPA peptide length increased, the Ti:C atomic ratio decreased greatly, corresponding to an increase in the number of adsorbed PEG. The observed ratio of C:organic oxygen (CO) exceeds the theoretical value of 2.0 for pure PEG, indicating that there is still foreign hydrocarbon contamination on the modified surface. These results are shown in Table 3.

table 3

Atomic fraction of adsorbed polymer composition atomic ratio surface C/Ti Ti/Ti-O-Ti C/CO Unmodified TiO ₂ 0.36 1.51 2.84 MPEG-DOPA 2.51 1.58 2.62 MPEG-DOPA ₂ 3.62 1.57 2.43 MPEG-DOPA ₃ 4.01 1.51 2.24

DOPA forms a strong, reversible bond with _TiO2 . The bond has a bond energy of 30.56 kcal/mol and requires a force of about 800 pN at the single molecule level to detach from _TiO2 , which is 4 times stronger than the interaction between avidin and biotin. The DOPA-TiO ₂ interaction force is approximately in between the avidin-biotin interaction force (one of the strongest hydrogen bond based forces in biology (0.1-0.2nN)) and covalent bond (>2nN).

To properly study the adhesion of DOPA, the following conditions were employed: single molecule approach, aqueous environment, platform for immobilizing DOPA. Atomic force microscopy (AFM) was chosen as a research tool because it can satisfy the above 3 conditions and is sensitive enough for detecting the viscoelastic properties of soft materials such as proteins, DNA, and synthetic polymers at the single-molecule level . Amine moieties were incorporated into the cantilever ends (Si3N4), followed by coupling of a mixture of methoxy-poly(ethylene glycol, mPEG) and Fmoc-terminated PEG (Fmoc-PEG) derivatives. (Boc-)DOPA was coupled to the amino group resulting from Fmoc cleavage (Figure 33C). A 5-10 molar excess of mPEG relative to DOPA-PEG was used, allowing isolation of single, unimmobilized DOPA4 PEG. This molecular configuration sterically hinders the molecular dynamics of DOPA-PEG, thus providing an explanation for the subsequent DOPA oxidation test results (Fig. 36). X-ray photoelectron spectroscopy (XPS) was used to monitor the chemical reaction steps, showing successful PEG modification on a flat silicon nitride surface (1×1 cm2) ( FIG. 28 ). The chemical groups introduced to the surface of the silicon nitride tip changed the electrostatic properties, which is also a good indicator of surface modification (Fig. 29-A). It is important to note that a difference in approaching signal was detected between the bare and modified cantilever, representing the resistance due to the molecular layer (Fig. 29-B).

The DOPA-coupled cantilevers exhibited entropic elastic great viscosity with attached PEG chains (FIG. 34). The histogram of the force distribution showed a uni-modal shape, indicating that only one adhesion event occurred, unlike the multivalent protein avidin-12. Force-distance (FD) measurements were collected and statistically analyzed (Figure 34, n=105). At a loading rate of 180nN/s, the average force in water is 785pN. Most importantly, the length of the elongated PEG (36 nm) coincided with the expected contour length of the PEG molecule (37 nm, Figure 30). These data were all used to illustrate one molecular event: monovalent incorporation of DOPA, polydispersity and tip geometry of PEG. A single DOPA was coupled to the end of the polymer chain as a binding unit on the _TiO2 surface (Fig. 33). This monovalent linkage differs from studies on metal-6 histidine (metal-(His)6), where the linked (His)6 provides 3 metal-chelation sites (3×metal/(His)2)13 -15. Furthermore, the "hill-tree"-like configuration of the PEG5 cantilevers could separate two distinct DOPA separation signals due to the polydispersity (tree) and globular tips (hills) of PEG (r = 25 nm).

"d" is defined as the z-shift distance of the piezoelectric device when a single DOPA-PEG molecule is fully extended during retraction (Fig. 34C). The values of 'd', although slightly varied, appeared to be essentially constant over repeated cycles (Fig. 34A). This small variation may be due to DOPA binding to the surface at different angles. An important feature of our assay is that the unbound signal is replicated with the 'same' DOPA molecule. This is in contrast to the traditional single molecule pulling experiment, where a tip randomly lifts a molecule. It also proved that the DOPA bonding chemistry is completely reversible. This reversibility leads us to the conclusion that the weakest chemical bond between the substrate and the tip (TiO2∼Si3N4) is the Ti(surface)-O(DOPA) bond.

This result indicates that at ~0.8 nN, the DOPA-TiO2 interaction is mechanistically at the level of the avidin-biotin interaction (one of the strongest hydrogen-bond-based interactions in biology ( 0.1～0.2nN)) and covalent bond (>2nN). Energy information is learned from force data obtained by varying the loading rate (force applied per unit time). Four orders of magnitude change in loading rate produced four different force distributions to determine the energy profile of DOPA binding. The logarithmic plot of force versus loading rate linearly in Figure 34D provides binding energy and distance, and then removes binding along the direction of force application. The energy barrier of DOPA is 28.1 kcal/mol, and the distance to the maximum activation energy is 1.27 Å (FIG. 34E).

The binding orientation of DOPA is believed to be with the two hydroxyl groups on the aromatic ring pointing downward to the surface. Therefore, identifying the chemical groups responsible for the unimolecular adhesion signal in AFM is important to rule out other binding chemistries caused by different orientations. Two methods were used.

First, covalent chemical modification of the hydroxyl group by tertary butyldimethylsiloxane (TBDMS) resulted in no binding over two hundred approach-retraction cycles (Fig. 31 , the first line). However, deprotection of the TBDMS group regenerated the DOPA binding capacity (Figure 31, bottom two lines).

Second, protection with borate ion complexation also completely inhibited the strong adhesion of DOPA (FIG. 31, n=200). These data clearly identify the dihydroxyl group of DOPA as the true structural source of the strong, reversible binding.

Mussels have developed an interesting way to produce such a strong bond in water by post-translationally modifying tyrosine with tyrosine hydroxylase. Using tyrosine as a substrate, this enzyme catalyzes the addition of a hydroxyl group. Large amounts of this enzyme are found in threads and plaques, and DOPA is also present in threads and plaques. Surprisingly, small post-translational modifications (-OH) appear to produce large changes in adhesion capacity. Therefore, experiments were designed to show the relationship between post-translational modifications and binding capacity.

Cantilevers linked with tyrosine instead of DOPA were prepared and the adhesion of tyrosine on TiO2 was studied. Apart from some very low probability non-specific adhesions, no detectable force signal was observed (Fig. 35A). To rule out the hypothesis that the tips used in this experiment did not contain any tyrosine molecules, the TiO2 surface was replaced with gold. The aromatic ring of tyrosine binds to the gold surface in a surface-parallel orientation through π-π electron interactions, a mechanism well known in surface adsorption chemistry18,19. The same cantilever used in TiO2 produced strong adhesion multiple times on the gold surface (FIG. 35B). Statistical analysis of the force distribution revealed that the π-π electron binding force is 398 (±98) pN, which is about 50% of the DOPA-TiO2 interaction force (FIG. 35C). The force signal from the tyrosine-gold binding also showed the same features as the DOPA-TiO2 interaction shown previously: elastic elongation of PEG with the expected profile length and repetitive signal shape with similar 'd' values (Fig. 33C). In this experiment, it was clearly demonstrated that the post-translational modification mediated by tyrosine hydroxylase greatly increased the binding capacity of DOPA from almost zero to 800pN.

The biological role of DOPA is not just to become sticky after oxidation: it can also cross-link polypeptide chains, creating a much stiffer substance than those found in threads and pads. There are multiple pathways to the cross-linking mechanism, which start from the chemically unstable DOPA quinone structure. Aryl-aryl ring coupling (di-DOPA) has been found in mussel mucin ₂₀ and Michael addition (quinone-alkylamine adduct) products in other species, but this addition product Not found in mussels (Fig. 36A). Therefore, it is also possible that these structures occurred due to oxidation in mussels. This is clear from a cross-linking perspective, but stickiness after ripening (ie, oxidation) is controversial. It has been shown that the DOPA quinone structure is not a major factor in the formation of stickiness. The DOPA quinone-PEG chain is sterically and chemically stabilized by the excess conjugation of methoxy-PEG molecules (5-10 molar equivalents), which is an important molecular configuration to prevent further reaction of DOPA quinone.

Time-resolved monitoring of force signals of unimolecular DOPA quinones elicited by pH increases (=9.7) revealed interesting facts that had been hidden until now. First, the detected AFM signals had two distinct distributions according to the magnitude of the force (high force and low force) (Fig. 36B). Statistical analysis of the data yielded two clear histograms with a weak force of 178 ± 62 pN and a strong force of 741 ± 110 pN (Fig. 36C). The incorporation of quinones can be assigned to the region of weak force, since this region appears only after the onset of oxidation and then becomes more frequent as time progresses (Fig. 36D). The slow kinetics characteristic of DOPA oxidation contribute to the initial high frequency of DOPA signaling. This is the first single-molecule assay to detect changes in the structure of small molecules in response to external stimuli. Based on these results, the possibility that the quinone structure of DOPA is responsible for the high viscosity can be ruled out. Thus, without being bound by any theory, regeneration of the reduced form, ie, the di-hydroxyl groups of DOPA during oxidation is believed to be a very important requirement for maintaining or changing the viscosity of DOPA-containing materials at the interface.

Furthermore, the DOPA anchor system could be a new platform to study other malleable biomacromolecules such as glycans, DNA and proteins. In the studies carried out, the elastic properties of PEG (Mw 3400) have been shown, this PEG length is believed to be the shortest chain length studied so far. This study was only possible due to the use of two defined anchoring methods at both ends: (1) covalent bonds between PEG and the cantilever, and (2) DOPA anchoring between PEG and the substrate. This approach is also in high contrast to conventional single-molecule assays, where the tip "sees" a different molecule with each movement of the cantilever. When _studying the response of molecules to external stimuli, given stimuli that are not 100% effective23 are a major obstacle to conducting research. DOPA-based anchoring systems can be used as an alternative technique to overcome these problems in existing single-molecule pull assays.

Currently, there is no clear answer as to why DOPA acts like a reversible glue similar to a "sticky note." There are two molecular binding models, the mononuclear bidentate chelate (Fig. 32A, right) and the binuclear bidentate chelate (Fig. 32A, left), but neither of these models takes into account the reversible binding of DOPA because these Studies have only focused on the adsorption process rather than the desorption process25. Therefore, it is assumed that the nature of this chemical bond is mainly covalent, independent of factors due to the removal of water molecules after adsorption. A study using FTIR showed that the nature of DOPA-TiO2 binding may be 60% ionic and 40% covalent. Based on this finding, a molecular adsorption model was revised to incorporate reversibility, where multiple hydrogen bonds are formed in water (Fig. 32B).

Figure 33 Experimental design and single-molecule DOPA adhesion

An image depicts how blue mussels (Mytilus Edulis) adhere to metal oxide surfaces. Circles include a plaque in which the rare amino acid DOPA is found.

(B) Two major protein components found in mussel plaques, Mefp-3 and Mefp-5. The content of DOPA in these mussel mucins was high, 27 mole% in Mefp-5 and 21% in Mefp-3. Bold Y (Y): DOPA, italic S (S): phosphoserine, italic R (R): hydroxyarginine.

(C) AFM tip modification. Polymerization of 3-aminopropyltrimethoxysilane (APTMS) introduced amino groups to the Si3N4 tip surface (not shown). The long chains describe PEG molecules terminally coupled to unimolecular (Boc)-DOPA. The DOPA-PEG molecule was stabilized with a mixture of mPEG-NHS (2k) and Fmoc-PEG-NHS (3.4k) in a molar ratio of 5-10:1 (see Supplementary Section for more detailed description).

Figure 34. Single-molecule force measurements and determination of the energetic profile of DOPA bound to a TiO2 surface

Four representative AFM single-molecule DOPA dissociation signals from one cantilever. These 4 signals were not generated consecutively (signals not showing any binding were removed). Although the detection of DOPA binding was rare (5-10%), the detected signal showed similar 'd' values (see Figure 34C).

(B) Histogram depicting force distribution. The average value was 781 ± 151 pN (n = 105) at a loading rate of 180 nN/sec.

(C) Definition of the distance 'd', the z-movement distance of the piezoelectric device when the DOPA-PEG molecule is fully stretched.

(D) Binding force (line) plotted against loading rate (log). The loading rate is the product of the spring constant of the cantilever and the pulling velocity. Four different loading rates were chosen: 1500, 180.7, 28.4 and 2nN/s. The mean force with standard deviation is shown for each given loading rate. The forces are 846.48±157pN (1500nN/s), 781±151pN (180nN/s), 744±206pN (28.4nN/s) and 636.2±150 (2nN/s).

(E) Schematic representation of the energy profile of DOPA binding. The external force tilts the energy distribution and lowers the energy barrier of the reaction complex. The slope (=kBT/xb) of the linear regression plot is 32.31, so that the distance of the activation barrier (xb) is 1.27 Å. The energy barrier height was determined by extrapolation when the loading rate was equal to zero (Eb = 28.1 kcal/mol).

Figure 35. Molecular identification of the sticky origin of DOPA

Monomolecules of tyrosine bind to the surface of TiO2. Apart from initial electrostatic interactions (unavoidable in some cantilevers), no clear binding signal was detected (upper limit represents signal, 700 replicates, n=639). No non-specific adsorption signal (lower limit represents signal, 700 replicates, n=61).

(B) Confirmation of the presence of tyrosine on the tip surface. The π electrons of the tyrosine phenyl group interact specifically with the π electrons of gold.

(C) Force distribution of tyrosine bound to a gold surface. Tyrosine showed an adhesion force of 398±98pN (180nN/sec).

Figure 36. Viscosity change of DOPA (DOPA quinone) after oxidation

Schematic representation of the chemical pathways for the formation and oxidation of DOPA. DOPA is produced by the action of tyrosine hydroxylase, which is then oxidized to DOPA quinone by pH and this enzyme. DOPA quinone is unstable and reactive due to free radical formation. It can be cross-linked with other DOPA molecules (di-DOPA) and can also react with the amino groups of lysine. Arrows are possible reactions found in other species but not in mussel mucin.

(B) Representative force signals (n = 16) with similar 'd' values (~50 nm) as shown in Fig. 1C. They were collected for AFM experiments (1800 repetitions) for 1 hour under alkaline conditions (20mM Tris-Cl, pH 9.7). The top to bottom of the graph represents the passage of time. Red signals represent DOPA-TiO2 and black signals represent DOPA quinone-TiO2.

(C) Histogram of forces obtained after an overall analysis of 1800 F-D curves. The histogram for the weak force region shows 178±82pN (n=143), and the histogram for the strong force region shows 741±110pN (n=51).

(D) Scatterplot of the number of events within the specified time window (10 min). DOPA signal (circles, left y-axis) decreased from 22 events in the first 10 minutes to only 3 events in the last time window. However, the signal for quinones (triangle, right y-axis) increased from one event in the first time window to 42 events in the last time window (50-60 min).

In conclusion, the single-molecule binding of DOPA was successfully detected (~0.8 nN) and its reversible binding chemistry was shown. This strong binding results from post-translational modifications, but oxidation of DOPA to DOPA quinone greatly reduces binding.

The antifouling coatings of the present invention can be substantially permanent, ie, lasting 120 days or more, or biodegradable, depending on the amount of DOPA or DOPA-derived moieties in the adhesive component. Figure 20 shows 3T3 fibroblast adhesion and spreading assay at day 28 on _TiO2 treated with mPEG-DOPA _1-3 . At earlier time points (i.e., less than 7 days), protein and cell adhesion resistance correlated well with the length of the DOPA peptide anchor group, with the order of increasing resistance being mPEG-DOPA < mPEG-DOPA ₂ < mPEG- _DOPA3 . TiO ₂ substrates treated with mPEG-DOPA ₂ and mPEG-DOPA ₃ maintained a reduction in cell adhesion (or cell adhesion resistance) for 21 days.

A Robust design approach was used to determine the effect of DOPA peptide anchor group length and modification conditions (pH, concentration and time) on PEG surface density and antifouling performance on metal, metal oxide, semiconductor and polymer surfaces Impact. The nine tests employed for each substrate are shown in Table 7 below. The length of the DOPA peptide and the pH of the modification buffer resulted in the greatest change in the amount of adsorbed PEG, as measured by XPS and assayed for adhesion and spreading of 3T3 fibroblasts, for almost all surfaces. The experiments summarized in Table 7 allowed us to identify the modified conditions that provided the best resistance to cell adhesion for each material, as detected by the 4 hour cell adhesion assay. After these 9 experiments were performed for each substrate, the data were subjected to Robust design analysis. Large error values are characteristic of this technique when plotting Robust design data, because a single data point at one factor level contains the bias of the other factors averaged over all factor levels.

The polymeric compositions of the present invention can also be used to coat the surfaces of devices and equipment for treating bodily fluids, including serum. The coating on the surface of these devices or equipment prevents proteins from binding to the surface, thereby reducing or eliminating the need for extensive washing or cleaning of the device or equipment between uses. These devices require thorough cleaning to prevent cross-contamination between bodily fluid samples applied to the surface of the device. Currently, cleaning these equipment and devices between uses requires extensive washing with caustic agents such as 50% bleach and/or elevated temperatures. The coating method of the present invention is to circulate a 1 mg/ml aqueous solution of DHPD polymer through the device at room temperature for several hours.

The coatings of the present invention can be used on medical implants for a variety of purposes. For example, these coatings can be used to prevent bacteria from adhering to and thereby preventing bacterial growth on implanted devices, reducing the likelihood of infection at the implant site. These coatings can be used to reduce the amount of acute inflammation that occurs on these devices by reducing protein binding to the device and cell adhesion to the device. The coatings of the invention can also be used as nanoparticles, preventing aggregation of these particles in the presence of serum.

Hydrogels

The polymer compositions of the present invention are also useful as surgical adhesives for medical and dental applications, and as vehicles for drug delivery to mucosal surfaces. The polymer composition may be a tissue-adhesive polymer hydrogel for medical applications such as tissue sealing, a gel for preventing surgical adhesion (scar tissue formation), bone and cartilage adhesives, tissue engineering, Site-specific drug efflux, and for research uses such as immobilization of proteins (including antibodies) and small molecule analytes (including drugs). The polymer compositions of the present invention can also be used as interfacial adhesives, where neat monomer or monomer solution is applied to a surface, as a tissue surface or metal or metal oxide implant/device surface and bulk polymerized Primer or adhesive between objects or polymer hydrogels. By selecting appropriate polymer components (determinable by one skilled in the art), the polymer compositions of the present invention can be injected or delivered in fluid form and harden in situ to form a gel network. Hardening in situ can occur by photocuring, chemical oxidation, enzymatic reactions, or by natural warming after delivery into the body.

In part the present invention provides a method for the non-oxidative gelation of the polymer composition of the present invention. The method comprises (1) providing a polymer composition of the present invention; (2) mixing water and the polymer composition; and (3) increasing the temperature of the mixture sufficiently to gel the polymer composition, thereby The increase in temperature does not oxidize the polymer or residues of DOPA or DOPA-derived moieties incorporated into the composition. Depending on the choice and nature of the polymeric components in the composition, increasing the concentration of the mixture can lower the temperature required to achieve gelation. Depending on the choice and nature of the particular copolymer components in the composition, larger hydrophilic block copolymers can increase the temperature required for gelation of the corresponding composition. Many other structural and/or physical parameters can be varied to tailor gelation, and these changes can be extended to other polymer compositions and/or systems consistent with the larger aspects of the invention.

It is widely recognized that commercially available PLURONIC(R) block copolymers self-assemble in a concentration- and temperature-dependent manner into micelles consisting of a hydrophobic PPO core and a water-swellable corona (composed of PEO segments). At high concentrations, certain PEO-PPO-PEO block copolymers, such as PLURONIC(R) F127 and PLURONIC(R) F68, transform from low viscosity solutions to clear, temperature-reversible gels at elevated temperatures. Without being bound by any theory, it is generally believed that the interaction between micelles at elevated temperature leads to the formation of a gel phase, which is stabilized due to the entanglement of micelles. The micellization and gelation processes depend on factors such as block copolymer molecular weight, relative block size, solvent composition, polymer concentration, and temperature. For example, increasing the length of the hydrophilic PEO block relative to the hydrophobic PPO block results in an increase in micellization and gelation temperature ( _Tgel ).

Different concentration aqueous solutions of DME-PAO7 and DOPA-PAO7 were tested by differential scanning calorimetry (DSC) to detect the aggregation of block copolymers into micelles. The resulting DSC profiles of PLURONIC(R) F127, DME-PAO7 and DOPA-PAO7 were qualitatively similar, characterized by a large endothermic transition corresponding to micelle formation followed by a small endotherm at the T- _gel (Fig. 2 ). The transition temperature of the small peak was found to correlate strongly with the T _gel measured by rheometry and vial inversion (Table 4).

Table 4

22% by weight solutions of DME-PAO7, DOPA-PAO7 and PLURONIC(R) F127 obtained gelation temperature by vial inversion method, rheometry or differential scanning calorimetry. Gel temperature (°C) vial inversion Rheology DSC DME-PAO7 (22% by weight) DOPA-PAO7 (22% by weight) PLURONIC(R) F127 (22% by weight) 22.0±1.022.0±1.017.0±1.0 20.3±0.620.4±0.515.4±0.4 20.9±0.121.7±0.217.5±0.4

Preparation of 10-30% (w/w) aqueous solution of DOPA-PAO7 copolymer and 35-54% (w/w) aqueous solution of DOPA-PAO8 copolymer in which DOPA conjugate is dissolved by cold process In distilled water at about 4°C, stir intermittently until a clear solution is obtained. Thermal gelation of concentrated solutions was initially evaluated by vial inversion. In this method, the temperature at which the solution no longer flows is taken as the gelation temperature.

The gelation temperature was found to be strongly dependent on the copolymer concentration and block copolymer composition (ie, PAO7 vs. PAO8). For example, 22 wt% solutions of DOPA-PAO7 and DME-PAO8 were found to form transparent gels at about 22.0 ± 1.0 °C; reducing the polymer concentration to 18 wt % gave a gelation temperature of about 31.0 ± 1.0 °C. However, DOPA-PAO7 solutions with concentrations less than 17% by weight did not form gels when added to 60°C. DOPA-PAO7 showed a slightly higher gelation temperature than unmodified PLURONIC(R) F127 (17.0±1.0°C). The gelation behavior of DOPA-PAO8 was found to be qualitatively similar, but required higher polymer concentrations to form gels. 54 wt% DOPA-PAO8 solution and DME-PAO8 solution formed gels at 23.0±1.0°C, while 50 wt% DOPA-PAO8 solutions formed gels at 33.0±1.0°C. However, DOPA-PAO8 solutions with concentrations less than 35% by weight did not form gels when heated to 60°C. DOPA-PAO8 exhibited a much higher gelation temperature than unmodified PLURONIC(R) F68 (16.0±1.0°C). These gels were found to not flow for a long period of time. Through this test, we also found that DOPA and DOPA methyl ester derivatives of the same commercially available product PLURONIC® PAO showed almost the same gelation temperature, made at room temperature with a 54% by weight solution of DME-PAO8 or a solution of DOPA-PAO8 The gel is harder than the gel made with 22 wt% DME-PAO7 solution or DOPA-PAO7 solution.

The viscoelastic behavior of the DOPA-modified PLURONIC(R) solutions was further investigated by oscillatory rheometry. Figure 3 shows the elastic storage modulus, G', of unmodified PLURONIC(R) F127 and 22 wt% aqueous solutions of DME-PAO7 as a function of temperature. Below the gelation temperature, the storage modulus G' is negligible, but at the gelation temperature ( _T ), G' increases rapidly, as shown in the graph of G' onset of increase vs. temperature. DOPA-PAO7 (not shown) showed similar rheological characteristics. The T- _gels of 22 wt% DME-PAO7 solution and DOPA-PAO7 solution were found to be the same (20.3±0.6°C), about 5°C higher than the equivalent concentration of unmodified-PLURONIC(R) F127 (15.4±0.4°C). The G' value of DME-PAO7 or DOPA-PAO7 is close to the plateau value of 13 kPa, comparable to that of unmodified PLURONIC(R) F127.

Figure 4 shows the rheological profile of 50% by weight unmodified PLURONIC(R) F68 and DME-PAO8 solutions as a function of temperature. The T- _gel of 50% by weight DME-PAO8 solution is 34.1±0.6°e, while the T _-gel of unmodified PLURONIC(R) F68 at the same concentration is about 18°C lower (16.2±0.8°C). The plateau storage moduli of 50 wt% solutions of DME-PAO8 and unmodified PLURONIC(R) F68 are not very different, approaching plateau values up to 50 kPa. Figure 5 shows the concentration dependence of T- _gel , the rheological profile of two concentrations of DME-PAO8 as a function of temperature. The T _gel of the 45 wt% DME-PAO8 solution is about 12°C higher than that of the 50 wt% DME-PAO8 solution.

Since both DOPA and DOPA methyl ester can be considered hydrophilic, the increase in _gel value of DOPA-modified PLURONIC(R) PAO relative to unmodified PLURONIC(R) PAO may be due to the coupling of DOPA to the end group Increase in the length of the hydrophilic PEO segment. From the data shown in Figures 3 and 4 it is clear that the coupling of DOPA or DOPA methyl ester to the PLURONIC(R) PAO end groups has a greater impact on the T _gel value of PLURONIC(R) F68 relative to PLURONIC(R) F127. This can be explained by the combined molecular weight of F68 (about 8,600) and F127 (about 12,600). Addition of DOPA and DOPA methyl ester to both ends using the chemistry shown in Scheme 1 increased the molecular weight by 446 and 474, respectively. This indicates that the percentage increase in molecular weight is greater for F68 relative to F127 due to the smaller molecular weight base of F68.

The data presented here are consistent with previous calorimetric studies of unmodified PLURONIC(R) PAO, demonstrating that the broad peak at low temperature is due to micelles, whereas the small peak at high temperature observed only in concentrated solutions corresponds to gelation , a process in which there is essentially no heat change. As shown in Figure 5, the micellization onset temperature, maximum heat capacity temperature, and T _gel value of unmodified PLURONIC® F127 were all lower than those of DOPA-PAO7, while the specific enthalpy determined by the area under the transformation (Figure 2 )basically the same. These enthalpies include those due to micellization and gelation. However, since the enthalpy of gelation is small, it is likely that most of the observed enthalpy change is due to micellization.

table 5

Comparison of the micellization onset temperature, maximum heat capacity temperature, enthalpy and gelation temperature of 30% by weight solutions of DME-PAO7, DOPA-PAO7 and unmodified PLURONIC(R) F127 obtained from the differential scanning calorimetry test. Micellization temperature (°C) Maximum heat capacity temperature (℃) ΔH(J/g) Gelation temperature (°C) DME-PAO7 (30% by weight) DOPA-PAO7 (30% by weight) PLURONIC(R) F127 (30% by weight) 5.2±0.24.6±0.21.9±0.3 8.3±0.18.0±0.66.0±0.4 20.3±2.419.3±1.420.6±1.6 14.0±0.414.0±0.210.6±0.6

It was observed that the micellization peak extended above the gelation onset temperature, indicating that other monomers gelled into micelles when the temperature was higher than the gelation temperature. The temperature dependence of the aggregation of DOPA-PAO7 and DME-PAO7 is shown in FIG. 6 . DSC pyrolysis curves showed that the micellization temperature and T _gel decreased with the increase of concentration. At the concentration where gelation does not occur, a broad endothermic peak corresponding to micellization can also be observed in the solution; as the concentration of the copolymer decreases, the characteristic temperature of the broad peak rises linearly, and a small peak and concentrated The gelation temperature of the copolymer is uniform, but disappears when the copolymer concentration is reduced.

As can be seen from the foregoing, a variety of polymer compositions of the present invention can be designed and prepared to provide a variety of micellization and/or thermal gelation properties. Alternatively, or in addition, the polymer compositions of the present invention may be degraded into excretable polymer components and metabolites using, for example, polyethylene glycol and lactic/glycolic acid, respectively. Regardless, the polymer compositions of the present invention provide improved adhesion properties through the incorporation of one or more DHPD residues by coupling a terminal monomer of the polymer component to said residues Achieved.

Another method of non-oxidative gelation of the polymer compositions of the present invention is photocuring. A photocurable monomer containing a DHPD moiety was copolymerized with PEG-DA (PEG-diacrylate) to form a viscous hydrogel by photopolymerization. Photopolymerization can be achieved at any visible or UV wavelength, depending on the monomers used. This can be determined by a person skilled in the art. Photocurable monomers consist of a tacky moiety coupled to a polymerizable monomer with an alkenyl group (such as a methacrylate group, with or without an intervening oligoethylene oxide linker or fluorinated ether linker) .

A UV lamp (365nm) was used to irradiate the photopolymerizable monomer containing the adhesive of the present invention, PEG-DA and 1.5 μL/mL photoinitiator (such as 2,2'-dimethoxy-2-phenyl-benzene Dimethoxy-2-phneyl-acetonephenone (DMPA), camphorquinone/4-(dimethylamino)-benzoic acid (CQ/DMAB), and ascorbic acid/fluorescein sodium salt (AA/ FNa ₂ )) aqueous mixture over 5 minutes. The presence of a binder in the precursor solution was found to affect the free-radical-initiated polymerization process. Catechol binders reduced the extent of gel formation, reduced the percentage of binder incorporation into the gel network, and prolonged gel formation time.

As shown in Figure 25, by measuring the mass with an echo detector, it was determined that the gel conversion rate reached 75% by weight after UV irradiation for 2 minutes, and the gel conversion rate increased to greater than 85% by weight after more than 5 minutes of irradiation. Gelation of PEG-DA occurred in 4 minutes or less (4 minutes for CQ/DMAB and ₃ minutes for AA/FNa2) when visible light initiator was used.

Copolymerization of PEG-DA with compound 1 or 7 (the synthesis of compounds 1 and 7 are shown in Schemes 2 and 3) is qualitatively similar to the polymerization of pure PEG-DA, although compound 1 or 7 was added to the PEG-DA precursor solution This results in a reduction in gel conversion (the gel conversion is dependent on DOPA monomer concentration and initiation system). For example, in the presence of 2.5 mol% or more of 1 or 7, gel conversion was reduced to less than 85% by weight in DMPA initiated UV polymerization. However, the degree of gel inversion of these gels was not statistically different. Similar concentration-dependent inhibition of DOPA was observed for visible light-induced initiators. For the AA/ _FNa2 and CQ/DMAB initiated mixtures, addition of 33.3 mol% of compound 1 increased the gelation time above 8 min. However, solutions containing compounds 1 and 7 were still photocurable even at relatively high DOPA mol%.

Scenario 2

Option 3

For example, the presence of 53.8 mol% of compound 1 or 7 in the precursor solution reduced the gel from 88 wt% to 77 wt%, and only 85 mol% of DOPA was incorporated into the hydrogel. A DOPA colorimetric assay (colorimetric assay developed by Waite and Benedict) performed on photocured hydrogels indicated the presence of the catechol DOPA in the hydrogels. After photocuring, the DOPA-containing gel was dialyzed against 0.5N HCl to extract unreacted DOPA monomer. To quantitatively measure the extent of DOPA incorporation, the dialysate was analyzed according to the DOPA colorimetric test of Waite and Benedict, and the results of this test were used to calculate the amount of DOPA incorporated into the gel network. Figure 26 shows the mole fraction of DOPA incorporated into the gel network as a function of the mole % of monomer 1 and monomer 7 in the precursor solution. There was no significant difference in the fraction of DOPA incorporated in samples containing monomers 1 and 7. The highest DOPA incorporated into the PEG hydrogel was 24.9 μmol/g.

Direct evidence for the presence of DOPA in the gel was obtained by immersing intact dialyzed gels in nitrite reagent followed by NaOH. The initially colorless gel turned bright yellow after the addition of nitrite, and turned red again with the addition of excess base. This color transition is typical of catechols, indicating the incorporation of the unoxidized form of DOPA into the hydrogel by photopolymerization. The intensity of the red color also reflects the concentration of DOPA incorporated into the photocured hydrogel.

Contact mechanical tests were performed on hemispherical photocurable gels to obtain information on the mechanical properties of the gels. The modulus of elasticity (E) was calculated by assuming specific Hertzian mechanics of non-adhesive contact between an incompressible elastic hemisphere and a rigid plane, where the difference between load (P _h ) and displacement (δ _h ) The Hertz relation is:

${\mathrm{<span\; class="notranslate">\; P</span>}}_{\mathrm{<span\; class="notranslate">\; h</span>}}<span\; class="notranslate">\; =</span>\frac{{\mathrm{<span\; class="notranslate">\; 16</span>}\mathrm{<span\; class="notranslate">\; <figure-callout\; id="1"\; label="R"\; filenames="A20058000618000811.png,A20058000618000862.png"\; state="\{\{state\}\}">R</figure-callout></span>}}^{\mathrm{<span\; class="notranslate">\; 1</span>}<span\; class="notranslate">\; /</span>\mathrm{<span\; class="notranslate">\; 2</span>}}\mathrm{<span\; class="notranslate">\; E.</span>}}{\mathrm{<span\; class="notranslate">\; 9</span>}}{\mathrm{<span\; class="notranslate">\; \&delta;</span>}}_{\mathrm{<span\; class="notranslate">\; <figure-callout\; id="5"\; label="h"\; filenames="A20058000618000831.png,A20058000618000861.png"\; state="\{\{state\}\}">h</figure-callout></span>}}{\mathrm{<span\; class="notranslate">\; 5</span>}}^{\mathrm{<span\; class="notranslate">\; 3</span>}<span\; class="notranslate">\; /</span>\mathrm{<span\; class="notranslate">\; 2</span>}}<span\; class="notranslate">\; -</span><span\; class="notranslate">\; -</span><span\; class="notranslate">\; -</span><span\; class="notranslate">\; (</span>\mathrm{<span\; class="notranslate">\; 1</span>}<span\; class="notranslate">\; )</span>$

where R is the radius of the hemispherical gel cantilever. Use equation (1) to fit the load versus displacement data, and the elastic modulus can be calculated according to the proportional coefficient of the fitted curve. As shown in Figure 6, an average Young's modulus (E) of about 50 kPa was obtained for the gel containing DOPA.

Table 6

Average Young's modulus of gels containing DOPA DOPA content ^* (μmol/g gel) Young's modulus (kPa) average standard deviation average standard deviation PEG-DA 72.3 7.69 PEG-DA+1 ⁺ 15.0 0.25 47.4 ^** 8.63 PEG-DA+7 ⁺ 14.5 0.056 51.4 ^** 9.91

⁺ 33 mol% in precursor solution

^* Determined by DOPA test

^** p<0.001 vs. PEG-DA gel

These modulus values were approximately 30% lower than those of the PEG-DA gel, confirming the inhibitory effect of DOPA on free radical-induced photopolymerization. Although the modulus is reduced relative to PEG-DA gels, gels containing DOPA still exhibit moduli suitable for many biomedical applications. A suitable modulus is greater than 500Pa. Another application of viscous hydrogels is topical drug delivery. For example, viscous hydrogels can form on the mucosal surfaces of the mouth or oral cavity. Hydrogels can be loaded with drugs such as antibiotics and facilitate slow drug release over a period of time. These hydrogels can also be loaded with analgesics for pain reduction at local sites. These hydrogels can also be loaded with chemotherapy drugs and inserted into tumor tissue to deliver cancer treatments. These hydrogels can also be loaded with cell proliferation inhibitor therapeutics and used to coat stents or other vascular devices for controlling cell proliferation at the implanted site of the vascular device.

Tissue-adhesive hydrogels that can crosslink in vivo can be used as tissue sealants in place of metal or plastic sutures. Adhesives bond to surrounding tissue at the site of surgery or injury, and the polymer forms an adhesive bond to seal the wound. Hydrogels can also be used to repair fractures and cartilage-to-bone damage.

other uses

These coatings and hydrogels also have many uses in industrial products, including preventing biofouling (seaweed, bacteria and mussels sticking to underwater surfaces) in marine transportation, and preventing bacteria in the water flow of factories such as power and pharmaceutical factories. Pollution, prevention of bacterial contamination in drinking water systems, as dental and denture adhesives, water adhesives for delivery indicators, coatings for water purity and detection sensors, paints for protection against biofouling.

These coatings and hydrogels also have many uses in consumer products and cosmetics, including, but not limited to, in dental and denture adhesives, in cosmetics as hair, skin, and leg adhesives, in cosmetics For example in eyeshadow, lipstick and mascara, in temporary tattoos, and as a resealable adhesive for bags and containers.

Without being limited to any particular synthetic scheme or method of preparation, suitable compositions of the invention may include, but are not limited to, urethane moieties between each terminal monomer and DOPA residues. As described in more detail below, this urethane moiety is an artefact of the substances/reagents used to crosslink DOPA residues and polymer components. A variety of other moieties are contemplated in the broad aspects of the invention, depending on the choice of terminal monomer functionality and coupling agent, as will be appreciated by those skilled in the art with knowledge of the invention.

Example: General Description

The following non-limiting examples and data illustrate various aspects and features related to the compositions and/or methods of the present invention, including the preparation of polymer or co-polymer compositions incorporating one or more DHPD components , known by the synthesis method described in the present invention. Although the use of the present invention has been illustrated with several polymer or co-polymer systems, those skilled in the art will appreciate that similar results can be obtained with a variety of other compositions and/or methods of preparation, which are described in the present invention. within range.

PEO ₁₀₀ PPO ₆₅ PEO ₁₀₀ (PLURONIC® F127, weight average molecular weight = 12,600), PEO ₇₈ PPO ₃₀ PEO ₇₈ (PLURONIC® F68, weight average molecular weight = 8,400), PEG (weight average molecular weight = 8000), pentafluorophenol, 1 , 3-dicyclohexylcarbodiimide (DCC), 4,7,10-trioxa-1,13-tridecanediamine, fluorescein sodium salt (FNa ₂ ), and ascorbic acid (AA) were purchased from Sigma (St. Louis, Missouri). L-DOPA, thionyl chloride, methacryloyl chloride, tert-butyldimethylsilyl chloride (TBDMS-Cl), di-tert-butyl dicarbonate, methacrylic anhydride, 2,2'-dimethoxy-2 -Phenyl-acetophenone (DMPA), acryloyl chloride, 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU), tetrabutylammonium fluoride (TBAF), 4- (Dimethylamino)-benzoic acid (DMAB), 1-vinyl-2-pyrrolidone (VP), N,N-disuccinimidyl carbonate, sodium borate, sodium molybdate dihydrate, sodium nitrite , 4-(dimethylamino)pyridine (DMAP), N-hydroxysuccinimide, N,N-diisopropylethylamine, dimethylamide, and dichloromethane were purchased from Aldrich (Milwaukee, Wisconsin ). Camphorquinone (GQ) was obtained from Polysciences, Inc. (Warrington, PA). Acetone was dried over 4 Å molecular sieves and distilled over P ₂ O ₅ before use. Triethylamine was freshly distilled before use. Other chemical reagents were used according to conventional methods.

L-DOPA methyl ester hydrochloride was prepared according to the method of Patel and Price, J. Org. Chem., 1965, 30, 3575 (incorporated herein by reference). Succinimidyl propionate activated PEG (mPEG-SPA, weight average molecular weight = 5000) was purchased from Shearwater Polymers, Inc. (Huntsville, AL). Ethyl acetate saturated with HCl was prepared by bubbling HCl gas through ethyl acetate (50 mL) for about 10 minutes. Synthesis of 3,4-bis(tert-butyldimethylsilyloxy)L-phenylalanine according to the method of Sever and Wilker, Tetrahedron, 2001, 57, (29), 6139-6146 (incorporated herein by reference) (DOPA(TBDMS) ₂ ) and 3,4-bis(tert-butyldimethylsilyloxy-N-tert-butoxycarbonyl-L-phenylalanine (Boc-DOPA(TBDMS) ₂ ).

Glass coverslips (12 mm in diameter) used in the following examples were immersed in a 5% Contrad70 solution (an anionic and nonionic surfactant sonicated in an allealtine aqueous base (Decon Labs, Inc., Bryn Mawr, PA) emulsion formed in a bath for 20 minutes), then washed with deionized (DI) water, sonicated in DI water for 20 minutes, washed in acetone, sonicated in acetone for 20 minutes, rinsed in hexane, rinsed in hexane Sonicate in medium for 20 min, wash in acetone, sonicate in acetone for 20 min, wash in DI water, and sonicate in DI water for 20 min. The coverslips were then air dried in a HEPA filtered laminar flow hood. To generate pristine gold substrates, clean coverslips were sputtered (Cressington 208HR) with 2nm Cr followed by 10nm Au (99.9% pure).

Titanium dioxide (TiO ₂ ) surfaces were prepared by physical evaporation of TiO ₂ onto silicon (Si) wafers with electron beams and then cleaned in a plasma chamber before testing. 100 nm Ti was coated onto silicon wafers (MEMC Electronic Materials, St. Peters, MO, surface orientation (100)) using an Edwards FL400 e-beam evaporator at <10 ⁻⁶ Torr. The silicon wafer was then sliced into 8 mm x 8 mm pieces and cleaned ultrasonically in the following media: 5% Contrad70, ultrapure water (ultrapure water was deionized and distilled), acetone and petroleum ether. The substrate was then further cleaned in an oxygen plasma chamber (Harrick Scientific, Ossining, NY) at <200 mTorr and 100 W for 3 minutes.

X-ray photoelectron spectroscopy (XPS) was used to characterize the pristine and modified gold surfaces as described below. In the configuration with a monochromated (monochromated) AlKα (1486.8eV) 300-W X-ray source, a 1.5mm-sized circular spot (circular spot), an immersion electron gun for counting charge effects and an ultra-high vacuum (<10 ^-8 Torr XPS data were collected on Omicron ESCALAB (Omicron, Taunusstein, Germany) in ). Takeoff angle is defined as the angle between the substrate normal and the detector, which is fixed at 45°. The substrate was secured to standard sample studs with double-sided tape. All binding energies were normalized to Au(4f _7/2 ) gold peak (84.0eV) or C(ls) carbon peak (284.6eV). The analysis consisted of a broad survey scan (50.0 eV pass energy) and a 10 min high resolution scan for C(ls) 270-300 eV (22.0 eV pass energy). Peak deconvolution and atomic percent calculations were performed with EIS analysis software.

Secondary ion spectra were collected on a TRIFT III ^TM time-of-flight secondary ion mass spectrometer (TOF-SIMS) (Physical Electronics, Eden Prairie, MN) in the mass range 0-2000 m/z. With a Ga ⁺ source, the beam energy is 15keV and the grating size is 100μm. Positive and negative spectra were collected and normalized to a set of low mass ions using PHI software Cadence.

To examine the relative hydrophilicity/hydrophobicity of surfaces, contact angle data were collected using the sessile drop method as described below. Ultrapure water (18.2 MΩ-cm; Barnstead, Dubuque, IA) advancing and receding contact angles (advancing and receding contact angles). For each surface, 4 measurements were made at different locations and the mean and standard deviation are reported.

Surface plasmon resonance (SPR) detection was performed on a BIACORE 2000 (Biacore International AB; Uppsala, Sweden) with bare gold sensor cartridges. Resonant responses were normalized with 0-100 mg/ml NaCl solvent. Dilutions of mPEG-DOPA, mPEG-MAPd and mPEG-OH (0.1 mM in H ₂ O) were injected into the SPR flow cell for 10 minutes before the flow was switched back to pure DI water. In a separate assay to detect protein adsorption to modified substrates, the sensor surface preformed with a PEG membrane was exposed to 0.1 mg/ml bovine serum albumin (BSA) dissolved in 10 mM HEPES buffer (0.15 M NaCl, pH = 7.2). ) solution, followed by pure buffer.

In experiments demonstrating the antifouling effect, NIH 3T3-Swiss albino fibroblasts obtained from ATCC (Manassas, VA) were maintained at 37°C, 10% CO ₂ , and containing 10% (v/v) fetal bovine serum ( FBS) and 100 U/ml penicillin and 100 U/ml streptomycin in Dulbecco's Modified Eagle's Medium (DMEM; Cellgro, Herndon, VA).

RP-HPLC prep was performed with a Waters HPLC system (Waters, Milford, MA) on a Vydac 218TP reverse phase column with a gradient of acetonitrile/0.1% trifluoroacetic acid in water (v/v). ESI-MS analysis was performed on a LCQ LC-MS system (Finnigan, Thermoquest, CA). MALDI-TOF MS analysis was performed on a Voyager DE-Pro mass spectrometer (Perseptive Biosystem, MA). α-cyano-4-hydroxycinnamic acid was used as a matrix. NiTi alloy (10 mm x 10 mm x 1 mm) was purchased from Nitinol Devices & Components (Fremont, CA). Si, _SiO2 (1500 Å thermal oxide) and GaAs wafers were purchased from University Wafer (South Boston, MA).

For the following cell adhesion tests and/or spreading assays, the modified and unmodified substrates were pretreated in 12-well TCPS plates with 1.0 ml of DMEM containing 10% FBS for 30 min at 37 °C, 10% _CO2 . Fibroblasts at passage 12-16 were harvested with 0.25% trypsin-EDTA, resuspended in DMEM containing 10% FBS, and counted with a hemocytometer. Cells were seeded at a density of 2.9×10 ³ cells/cm ² by diluting the suspension to an appropriate volume and then adding 1 ml of the dilution to each well. Substrates were maintained in DMEM containing 10% FBS, 37°C, 10% _CO2 for 4 disappearances, and non-adherent cells were then aspirated. Cells adhered to the substrate were fixed in 3.7% paraformaldehyde for 5 min and then treated with 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl in DMSO Indocarbocyanine perchlorate (1,1'-dioctadecyl-3,3,3',3'-tetramethyl indocarbocyanine perchlorate, DiI; Molecular Probes, Eugene, OR) was treated at 37°C for 30 minutes. The dye was then aspirated, the substrate washed with DMSO for 10 minutes (3x), and mounted on slides with Cytoseal (Stephens Scientific, Kalamazoo, MI) to preserve fluorescence. For statistical purposes, these experiments were performed in triplicate. For electron microscopy, some samples were dehydrated with EtOH after fixation, critical point dried, and sputtered with 3 nm Au.

To quantify cell adhesion, substrates were detected with an Olympus BX-40 (λ _Ex = 549 nm, λ _Em = 565 nm) and color was recorded with a Coolsnap CCD camera (Roper Scientific, Trenton, NJ). For each of the substrates in triplicate, 5 pictures were taken. The resulting pictures were quantified using thresholding in MetaMorph (Universal Imaging, Downington, PA). Statistical significance of data was determined using one-way ANOVA with Tukey's post-hoc test with 95% confidence intervals (SPSS, Chicago, IL). The mean and standard deviation of assay results are reported.

Example 1

Synthesis of Succinimidyl Carbonate PAO, SC-PAO7

PLURONIC(R) F127 (0.60 mmol) was dissolved in 30 mL of anhydrous dioxane. N,N'-disuccinimidyl carbonate (6.0 mmol) dissolved in 10 mL of anhydrous acetone was added. DMAP (6.0 mmol) was dissolved in 10 mL of dry acetone and added slowly under magnetic stirring. Activation was carried out at room temperature for 6 hours, and then SC-PAO7 was precipitated in ether. The disappearance of the starting material in the reaction was followed by TLC (chloroform-methanol (5:1) solvent system). The product was purified by dissolving in acetone and precipitating 4 times with diethyl ether. The product yield was 65%. ¹ H NMR (500MHz, CDCl ₃ ): δppm 0.96-1.68(br, -OCHCH ₃ CH ₂ O-), 2.80(s, -COON(CO) ₂ (CH ₂ ) ₂ ), 3.15-4.01(br,- _{OCH2CH2O- ; -OCHCH3CH2O- ) ,} 4.40 ₍ s, _{-OCH2CH2OCOON} (CO) _{2CH2CH2- )} .

Example 2

Synthesis of DME-PAO7

A slurry of DOPA methyl ester hydrochloride (1.25 mmol) and triethylamine (2.5 mmol) was mixed with SC-PAO7 (0.16 mmol) in 10 mL of chloroform. The disappearance of the starting material in the reaction was followed by TLC (chloroform-methanol-acetic acid (5:3:1) solvent system). After stirring at room temperature for 1 hour, the solvent was evaporated and DME-PAO7 was purified by precipitation with cold methanol three times. DME-PAO7 obtained a positive Arnow's test result, indicating the presence of catechol hydroxyl groups. The product yield was 75%. ¹ H NMR (500MHz, CDCl ₃ ): δppm 0.98-1.71 (br, -OCHCH ₃ CH ₂ O-), 2.83-3.06 (m, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 3.15-4.02 (br, -OCH ₂ CH ₂ O-; -NHCH(CH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 4.05-4.35 (d, -OCH ₂ CH ₂ OCONHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 4.55 (br, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 5.30 (d, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 6.45-6.80 (1s, 2d, - _{NHCHCH2C6H3 ( OH} ) _{2COOCH3 )} .

Example 3

Synthesis of DOPA-PAO7

Under Ar atmosphere, L-DOPA (1.56 mmol) was added to 30 mL of 0.1 M Na ₂ B ₄ O ₇ (pH=9.32) aqueous solution, followed by stirring at room temperature for 30 minutes. SC-PAO7 (0.156 mmol) in 5 mL of acetone was added to the resulting mixture and stirred overnight at room temperature. Sodium carbonate was used to maintain the pH of the solution during the reaction. The disappearance of the starting material in the reaction was followed by TLC (chloroform-methanol-acetic acid (5:3:1) solvent system). The solution was acidified to pH 2 with concentrated hydrochloric acid, then extracted 3 times with dichloromethane. The combined dichloromethane extracts were dried over anhydrous sodium sulfate, filtered and the dichloromethane was removed by evaporation. The product was further purified from cold methanol precipitation. DOPA-PAO7 obtained a positive Arnow's test result, indicating the presence of catechol hydroxyl groups. The product yield was 52%. ¹ H NMR (500MHz, CDCl ₃ ): δppm 0.92-1.70 (br, -OCHCH ₃ CH ₂ O-), 2.91-3.15 (m, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH), 3.20-4.10 ( br, -OCH ₂ CH ₂ O-; -OCHCH ₃ CH ₂ O-), 4.1-4.35 (d, -OCH ₂ CH ₂ OCONHCHCH ₂ C ₆ H ₃ (OH) ₂ COOH), 4.56 (m, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOH), 5.41 (d, -NHCHCH ₂ C ₆ H ₅ (OH) ₂ COOH), 6.60-6.82 (1s, 2d, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOH) .

Example 4

Synthesis of Succinimidyl Carbonate PAO8, SC-PAO8

SC-PAO8 was prepared in a similar manner to that described above for the synthesis and purification of SC-PAO7. The product yield was 68%. ¹ H NMR (500MHz, CDCl ₃ ): δppm 0.95-1.58(br, -OCHCH ₃ CH ₂ O-), 2.80(s, -COON(CO) ₂ (CH ₂ ) ₂ ), 3.10-4.03(br, - _{OCH2CH2O- ; -OCHCH3CH2O-} ), _{4.40 ( s} , _{-OCH2CH2OCOON} (CO) _{2CH2CH2 )} .

Example 5

Synthesis of DME-PAO8

DME-PAO8 was prepared in a similar manner to the synthesis and purification of the DME-PAO7 conjugate described above. The product yield was 76%. ¹ H NMR (500MHz, CDCl ₃ ): δppm 0.98-1.50 (br, -OCHCH ₃ CH ₂ O-), 2.85-3.10 (m, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 3.15-4.01 (br, -OCH ₂ CH ₂ O-; -OCHCH ₃ CH ₂ O-; -NHCH(CH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 4.03-4.26 (d, -OCH ₂ CH ₂ OCONHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 4.55 (m, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 5.30 (d, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOCH ₃ ), 6.45 _-6.77 ( _1s , _2d , _-NHCHCH2C6H3 (OH) _2COOCH3 ).

Example 6

Synthesis of DOPA-PAO8

DOPA-PAO8 was prepared and purified in a similar manner to that described above for the synthesis of DOPA-PAO7 conjugates. The product yield was 49%. ¹ H NMR (500MHz, CDCl ₃ ): δppm 0.92-1.50 (br, -OCHCH ₃ CH ₂ O-), 2.91-3.10 (m, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOH), 3.15-3.95 ( br, -OCH ₂ CH ₂ O-; -OCHCH ₃ CH ₂ O-), 4.06-4.30 (d, -OCH ₂ CH ₂ OCONHCHCH ₂ C ₆ H ₃ (OH) ₂ COOH), 4.54 (m, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOH), 5.35 (d, -NHCHCH ₂ C ₆ H ₅ (OH) ₂ COOH), 6.50-6.80 (1s, 2d, -NHCHCH ₂ C ₆ H ₃ (OH) ₂ COOH) .

Example 7

Colorimetric assay

The coupling efficiency of DOPA methyl ester and DOPA to PLURONICs(R) F127 and F68 was determined by the colorimetric method of Waite and Benedict. Briefly, aliquots of standard or unknown solutions were diluted with 1N HCl to a final volume of 0.9 mL and samples were analyzed in triplicate. Add 0.9 mL of nitrite reagent (1.45 M sodium nitrite and 0.41 M sodium molybdate dihydrate) to the DOPA solution, followed immediately by 1.2 mL of 1 N NaOH. Because absorbance is time-varying, care was taken to ensure that the time interval between addition of NaOH and recording of absorbance was 3 minutes for all standards and samples. The absorbance of all standards and samples was recorded at 500 nm. DOPA was used as a standard for DOPA methyl ester and DOPA conjugates.

Example 8

Rheology

Rheological measurements of the gelation process were performed with a Bohlin VOR rheometer (Bohlin Rheologi, Cranbury, NJ). All measurements were made using a 30 mm diameter stainless steel cone and plate geometry with a cone angle of 2.5°. Control the temperature with a circulating water bath. The samples were cooled in the refrigerator before transferring 0.5 mL of the liquid solution to the instrument. The storage and loss moduli G' and G" were measured at 0.1 Hz in an oscillatory mode with a stress of 0.45%. The heating rate was 0.5 °C/min, but the heating rate was reduced to 0.1 °C/min near the gelation temperature The dependence of the viscoelastic data on the stress amplitude was examined for several samples. Measurements were made only in the linear region, where the modulus does not depend on the stress amplitude. Mineral oil was used around the outer surface of the sample to prevent the measurement process in dehydration.

Example 9

Differential Scanning Calorimetry (DSC)

DSC measurements were performed on a TA Instruments DSC-2920 (TA Instruments, New Castle, DE) calorimeter. Thermal spectra were obtained for 3 samples of each concentration during heating and cooling cycles. A sample volume of 20 ul was used in a sealed aluminum evaporating dish, and scan data were recorded at a heating and cooling rate of 3°C/min, with an empty evaporating dish serving as a control.

Example 10a

Amino-terminated methoxy-PEG, mPEG-NH ₂ (2.0 g, 0.40 mmol, M _w =2,000 or 5,000, Sun-Bio PEGShop), N-Boc-L-DOPA dicyclohexylammonium salt (0.80 mmol), HOBt (1.3 mmol) and _Et3N (1.3 mmol) were dissolved in 20 mL of a 50:50 mixture of dichloromethane (DCM) and DMF. Then HBTU (0.80 mmol) dissolved in 10 mL of DCM was added and reacted for 30 minutes at room temperature under argon atmosphere. Then the reaction solution was washed successively with saturated sodium chloride solution, 5% NaHCO ₃ , dilute HCl solution and distilled water. The crude product was concentrated under reduced pressure and purified by column chromatography on Sephadex(R) LH-20 with methanol as mobile phase. The product mPEG-DOPA was further purified by precipitation 3 times in cold methanol, dried under vacuum at room temperature and stored at -20°C under nitrogen. ¹ H NMR (500MHz, CDCl ₃ /TMS): δ6.81-6.60 (m, 3H, C ₆ H ₃ (OH) ₂ -), 6.01 (br, s, 1H, OH-), 5.32 (br, s , 1H, OH-), 4.22 (br, s, 1H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH(N-)-C(O)N-), 3.73-3.38 (m, PEO), 3.07(m, 2H, PEO-CH2-NH-C(O) _- ), 2.73(t, 2H, _C6H3 (OH) _2- CH2 _- CH(N-)-C(O)N-) , 1.44(s, 9H, ( _CH3 ) _3C- ), 1.25(s, 3H _{, CH3CH2O-} ).

Example 10b

By analogy, the synthesis and related methods of the preceding examples can be extended with other DOPA-containing peptides and oligopeptides (of natural or synthetic origin). Depending on the specific synthetic sequence, an N-terminal protecting group may optionally be employed. As noted above, a variety of other DOPA-like viscous components may also be used and are well known to those skilled in the art having the knowledge of the present invention. For example, B-amino acids and N-substituted glycine DOPA analogs can be used.

Regardless of the particular DHPD viscous component, a variety of polymeric components can be used according to the synthetic techniques and methods described above. The molecular weight of the polymer components can vary, limited only by the corresponding solubility requirements. As noted above, many other polymers can be used for surface antifouling and/or particle stabilization, including, but not limited to, hyaluronic acid, dextran, and the like. Depending on the solubility requirements and the desired surface interaction, the polymer components can be branched, hyperbranched or dendrimeric, and these components are commercially available or can be prepared by known synthetic techniques.

Although the composition of Example 10a is an amidation product of the starting materials described above, it is understood that the N-terminus of the DHPD component can be coupled to the end group of a suitable functionalized natural or synthetic polymer, including the polymers described above. , main chain or side chain to prepare similar polymer-DHPD conjugates. For example and without limitation, as described above, a suitable polymer component terminating in carbonate functionality can be reacted with the N-terminus of the desired DHPD component to provide the desired conjugate.

Example 11a

The blue mussel Mytilus edulis podoprotein 1 (Mefp 1 ) of the consensus decapeptide repeat sequence (Mussin mucin decapeptide, MAPd, _NH2 -Ala-Lys-Pro-Ser-Tyr-Hyp-Thr-DOPA-Lys- _CO2H ). Fmoc was deprotected using 25% piperidine in NMP for 20 minutes. After an initial 10 min preactivation step, amino acid coupling was performed using 2 equivalents of a 1:1:1:1 Fmoc-amino acid:BOP:HOBt:DIEA reaction for 20 min. After completion of the decapeptide, the free amino terminus of the decapeptide was coupled to activated methoxy-PEG- _CO2H (mPEG-SPA, _Mw = 2k or 5k, Shearwater Polymers) using carbodiimide chemistry. The PEG-decapeptide conjugate (mPEG-MAPd, 2k or 5k) was cleaved using 1M TMSBr in TFA with EDT, thioanisole and m-cresol for 2 hours at 0°C. The crude mPEG-MAPd product was precipitated in diethyl ether at 0 °C and purified by preparative HPLC using a Vydac 218TP reverse phase column (220 x 22 mm x 10 μm). The purity of the product was determined to be >90% by analytical HPLC and the structure was confirmed by PerSeptive Biosystem MALDI-TOF-MS.

Example 11b

The synthesis and procedure in Example 11a can be extended analogously to or with variations described in Example 10b. In addition, other conjugates of DOPA-containing polymers can be prepared by enzymatic conversion of the contained tyrosine residues. Other well known techniques in the field of peptide synthesis can also be effectively used to provide other desired protein sequences, peptide conjugates and to achieve adhesion/anti-fouling effects.

Example 12a

The gold surface was modified by adsorption of mPEG-DOPA or mPEG-MAPd (2k, 5k) from DCM or phosphate-buffered saline (PBS; pH = 3, 7.4, and 11) solutions at polymer concentrations of 0.1–75 mg/ml. ongoing. Substrates were placed in vials and immersed in mPEG-DOPA or mPEG-MAPd solutions for up to 24 hours. After removal from solution, the substrate was washed with a suitable solvent (DCM or DI _H2O ) to remove unbound polymer and dried in vacuo. As a comparison, the same surface modification was performed with PEG-monomethyl ether (mPEG-OH, average M _w =5000). Alternatively, incubate a drop of a solution containing mPEG-DOPA or mPEG-MAPd (10 mM in PBS, PEG molecular weight = 2000) on an Au-coated glass coverslip (Au thickness ~10 nm) for 30 min at 37°C , and then wash (3x) the surface of the coverslip with PBS. Analysis of the modified surface by advancing/receding contact angle, XPS and TOF-SIMS revealed the formation of mPEG-DOPA or mPEG-MAPd chemisorbed layers.

Figures 7A-C show the XPS patterns of unmodified, mPEG-OH-modified and mPEG-DOPA-modified surfaces. As expected, the ether peak at 286.5 eV was only minimally maximized after treatment with mPEG-OH, whereas a significant increase was observed after adsorption of mPEG-DOPA, suggesting the abundant presence of ether carbons. Ether peaks of pure PEG with the same binding energy have been reported in the literature. The smaller peak at 285.0 eV in Figure 7 is caused by aliphatic and aromatic carbons in the PEG and DOPA headgroups and some hydrocarbon contamination introduced during preparation/evacuation.

Time-of-flight SIMS data corroborates the findings in XPS. TOF-SIMS analysis was performed on unmodified and mPEG-DOPA-modified Au substrates as well as mPEG-DOPA powder and gold substrates exposed to mPEG-OH. Data collection was performed for approximately 4 minutes for each substrate.

The cation spectrum of unmodified Au showed (C _n H _2n+1 ) ⁺ and (C _n H _2n-1 ) ⁺ peaks typically representing hydrocarbon contaminants (data not shown). Other lesser contaminants _are present, including NH ₄ ⁺ , Na ⁺ and relatively small amounts ^{of CaHbOc+} _{species .} Due to the method used to deposit the Au thin film, a Cr peak was observed at m/z~52 in addition to the Au peak at m/z~196.9. Exposure of the gold surface to mPEG-OH resulted in only a moderate increase in the peak representing the C _a H _b O _c ⁺ PEG fragment, which could be caused by contaminants or nonspecific adsorption of mPEG-OH. This is evidenced by the m/z ~225 (AuOC ⁺ ) and 254 (AuOCCO ⁺ ) peaks, which do not show a significant increase when compared to the substrate modified with mPEG-DOPA. (FIGS. 8A-C).

Dominant in the cation spectrum of _the Au surface modified with mPEG-DOPA is the presence of the _CaHbOc ⁺ peak representing the _adsorbed molecules. As shown in Fig. 9, the relative abundance _{of C2H3O} ⁺ and _C2H5O ⁺ increased relative to the _unmodified and mPEG-OH-modified surfaces. There is also a significant increase in the relative abundance of C ₃ H ₇ ⁺ (m/z ~ 43) and C ₄ H ₅ ⁺ (m/z ~ 53), these peaks may be hydrocarbon contaminants or tert-butyl in the Boc protecting group caused by the base.

Probably the most striking feature in the cation spectrum of the PEGylated Au substrate is the patterned triplet repetition in the high mass range (Figure 10). Each of these triplet clusters corresponds to an Au-DOPA-( _CH2CH2O ) _{n fragment} . When further resolved, each subcluster in the triplet represented the addition of _CH2 , _{CH2CH2 ,} or _CH2CH2O , and these peaks were each separated by about _14-16 amu. The repeating pattern was discernible at n = 0-15, while outside this range the signal was below the detection limit.

In the anion spectrum of the pristine Au surface, except for O ⁻ , HO ⁻ and Au _n ⁻ strong detectable peaks for n=1-3, only a few records were observed (data not shown). Small amounts of hydrocarbon contamination were present at m/z ~ 13 (CH ⁻ ), 24 (C ₂ H ₂ ⁻ ) and 37 (C ₃ H ⁻ ). The anion spectrum of the PEGylated Au surface is dominated by the C ₇ H ₁₁ O ₂ ⁺ peak at m/z ~ 126.893. The presence of this peak at moderate intensity in the spectrum of Au modified with mPEG-OH indicates that it represents a larger ethylene glycol fragment. The most interesting peaks are in the high mass range (>200m/z), representing catechol oxygen coupled to Au. This pattern shows that one Au atom can bind up to 6 oxygen atoms corresponding to 3 DOPA.

Contact angle data showed a strong dependence on the properties of the adsorption solvent used when gold films were modified with mPEG-DOPA (data not shown). Surfaces modified in DCM showed significantly lower θa than unmodified surfaces (p<0.001) and surfaces modified in all aqueous solutions (p<0.05). In general, as the pH of the aqueous solution increases, the hydrophilicity of the treated surface decreases, suggesting a reduced ability to pegylate the surface, possibly due to the tendency of DOPA to be oxidized to its less viscous quinone at increasing pH form, an explanation supported by previous studies showing that the unoxidized catechol form of DOPA plays a major role in viscosity.

Example 12b

Protein adsorption and cell adhesion/spreading onto untreated and treated coverslips were assessed below. Surface plasmon resonance (SPR) experiments showed that DOPA-containing polymers bound rapidly to gold surfaces, while the resulting modified surfaces had enhanced resistance to protein adsorption (Fig. 11). About 70% less protein was adsorbed on the gold modified with mPEG-MAPd(5k) than on the unmodified gold surface. Analysis of fibroblasts cultured on the modified substrates showed that cell adhesion was strongly dependent on the mPEG-DOPA concentration used to prepare the PEG-modified substrates (Figure 12), adsorption solvent and modification time. Surface modification with >25 mg/ml of mPEG-DOPA or mPEG-MAPd for 24 hours showed a statistically significant reduction in cell adhesion and spreading (Figures 12-14). Gold surfaces modified with mPEG-MAPd(5k) showed a 97% reduction in total projected cellular area and a 91% reduction in cell density attached to the surface.

Example 12c

The modifications shown in Example 12a, can optionally be modified with reference to Examples 10b and 11b, and can be extended to other noble metals, including but not limited to: silver and platinum surfaces. As described herein, these applications can also be extended to include surface modification of any bulk metal or alloy with a passivated or oxidized surface. For example, bulk metal oxide and related ceramic surfaces can be modified as described herein. The technique can also be extended to semiconductor surfaces, such as those used in the fabrication of integrated circuits and MEMS devices, which are also described below in the context of nanoparticle stabilization.

Example 13

Silicate glass surfaces (glass coverslips) were modified by adsorption of mPEG-MAPd(2k) from 10 mM aqueous solution using the method described in Example 12a. Cell density of NIH 3T3 cells adhered to modified and unmodified glass surfaces was assessed as described above. The cell density on the glass surface modified with mPEG-MAPd for 24 hours showed a 43% decrease compared to the unmodified glass surface (cell density (cells/mm ² ): 75.5+ on unmodified glass /-6.5; 42.7 +/-9.8 on glass modified with mPEG-MAPd).

Example 14a

To illustrate the stabilization of metal oxides (and specifically metal oxide nanoparticles), 50 mg mPEG-DOPA (5k) was dissolved in water (18 MΩ-cm, Millipore) and 1 mg magnetic powder (Fe ₃ O ₄ ) was added. Similar preparations were made with mPEG- _NH2 (5k) (Fluka) and mPEG-OH(2k) (Sigma) as controls. Each aqueous solution immersed in a 25°C bath was sonicated for 1 hour using a Branson Ultrasonics 450 probe sonicator. The frequency of the probe is 20kHz, the length is 160mm, and the tip diameter is 4.5mm. The samples were then removed and allowed to sit overnight at room temperature to allow any unmodified magnetite to precipitate out of solution. Suspensions prepared with control polymers (mPEG- _NH2 and mPEG-OH) precipitated rapidly to give a brown solid and a clear, colorless supernatant. In samples prepared using nanoparticles stabilized with PEG-DOPA, the samples were clear brown. The clear brown supernatant was separated and dialyzed against Spectra/Por ^(R) membrane tubing (MWCO: 15,000) in water for 3 days. After dialysis, samples were freeze-dried and stored under vacuum at room temperature until use.

Example 14b

The nanoparticles stabilized with mPEG-DOPA were characterized by transmission electron microscopy (TEM), thermogravimetric analysis (TGA), Fourier transform infrared spectroscopy (FTIR) and UV/visible spectroscopy. TEM results showed that most of the nanoparticles were 5-20 nm in diameter (data not shown). TGA analysis of 0.4 mg of magnetite stabilized with mPEG-DOPA indicated that the particles contained 17 wt% mPEG-DOPA (data not shown). FTIR performed on untreated magnetite showed a small absorption in the wavelength range of 4000-400 ^cm , while FTIR of nanoparticles treated with mPEG-DOPA showed 800-1600 ^cm and 2600-3200 cm ^-1 absorption band, which confirms the existence of mPEG-DOPA.

Example 14c

Dried magnetite nanoparticles stabilized with PEG-DOPA are readily dispersed in aqueous and polar organic solvents (eg, dichloromethane) to obtain clear brown suspensions that are stable for months without significant precipitate formation. Nanoparticles stabilized with mPEG-DOPA were prepared by dispersing 1 mg of magnetite treated with mPEG-DOPA in 1 ml of water (18 MΩ-cm filtered with a ^Millex® AP 0.22 μm filter (Millipore)), DCM or toluene. Suspensions in different solvents. Place the suspension in a bath sonicator for 10 min to disperse the nanoparticles. All three solutions were stable for at least 6 months at room temperature, while control suspensions of untreated magnetite and magnetite stabilized with mPEG-OH or mPEG- _NH2 in each solvent were stable in less than 24 hours Precipitation occurs.

Example 14d

Suspensions of nanoparticles stabilized with mPEG-DOPA were also found to be stable in the presence of physiological concentrations of salt. To determine whether mPEG-DOPA could inhibit salt-induced nanoparticle aggregation, 0.3 mg of magnetite treated with mPEG-DOPA was placed in a quartz cuvette and 0.7 ml of water (18 MΩ-cm filtered with a 0.25 μ filter) was added. Aliquots of saturated NaCl solution (5 μl, 10 μl, 20 μl, 50 μl, 100 μl) were successively added to cuvettes and allowed to stand for 10 minutes before UV-VIS spectroscopy ( FIG. 15 ). The absorption spectra of nanoparticles stabilized with mPEG-DOPA suspended in solutions containing increasing NaCl concentrations were almost the same, which indicated that mPEG-DOPA effectively stabilized nanoparticles and prevented aggregation. The peak centered at 280 nm is indicative of the catechol side chain of DOPA.

Example 14e

Those skilled in the art with knowledge of the present invention will understand that the steps and techniques illustrated in Examples 14a-14d can be extended to various other metal oxide or ceramic nanoparticles. Likewise, these applications of the invention may further include polymer-DHPD conjugates for a wide range of compositions and variants similar or identical to those described in Examples 10b and 11b. As explained below, in the preparation of semiconductor compositions, in-situ stabilization of metal oxide or ceramic nanoparticles can be performed in the presence of the polymer-DHPD conjugates of the present invention during formation.

Example 15a

To demonstrate the stability of the metal nanoparticles, a commercially available gold colloidal suspension (Sigma, particle size 5 or 10 nm) was placed in a dialysis tube (molecular weight cut-off of 8000 was used for 5 nm and 15000 for 10 nm) , and dialyzed against ultrapure water for 2-3 days to remove sodium azide present in commercially available preparations. The dialyzed suspension was then placed in a vial and mPEG-DOPA (10 mg/ml) was added. The samples were left to stand at room temperature for about 2 days, and then the samples were dialyzed again to remove excess mPEG-DOPA. Untreated 10 nm Au nanoparticles were unstable and aggregated in the presence of NaCl (Figure 16), while the treated Au nanoparticles remained in stable suspension in the presence of NaCl aqueous solution (Figure 17).

Example 15b

Various other metal nanoparticles including, but not limited to, silver, platinum, etc. can be stabilized as described in the previous examples. While stability was demonstrated using representative coupling compositions of the invention, various other compositions can also be prepared in a manner similar to or identical to the alternative embodiments described in Examples 10b and 11b. Comparable results can be obtained by in situ formation of stabilized nanoparticles synthesized from the corresponding metal precursors in the presence of a suitable adhesive coupling polymer of the invention.

Example 16a

The data in this example demonstrate the stability of semiconductor nanoparticles. CdS nanoparticles (quantum dots) were prepared using a standard method based on slow mixing of dilute Cd(NO ₃ ) ₂ solutions with dilute Na ₂ S solutions. Fresh stock solutions (2 mM) of Cd(NO ₃ ) ₂ and Na ₂ S were prepared in nanopure water. Using a gas-tight syringe, slowly inject the Na ₂ S solution into the 50 ml Cd(NO ₃ ) ₂ solution at a rate of 20 μl s ⁻¹ . With the addition of _Na2S , the solution turned yellow, and after injecting ₂ mL of Na2S, a yellow precipitate appeared due to the aggregation of CdS nanoparticles. The CdS precipitate was isolated and dried for future use. Using the method described above for magnetite, the dried CdS powder was dispersed in the mPEG-DOPA solution by sonication to obtain a clear yellow solution. The above yellow aqueous suspension was stored in the dark at room temperature for several months, no precipitate formation was observed. Control experiments performed in the absence of polymer but in the presence of mPEG-OH or mPEG- _NH2 gave a yellow precipitate and a clear colorless supernatant. CdS nanoparticles stabilized with mPEG-DOPA remained in stable suspension in the presence of aqueous NaCl ( FIG. 18 ).

Example 16b

The results of this example show the in situ formation of stabilized semiconductor nanoparticles. CdS nanoparticles (quantum dots) were prepared by slowly mixing dilute methanolic solutions of Cd( _NO3 ) ₂ and _Na2S in the presence of mPEG-DOPA. Fresh stocks (2 mM) of Cd( _NO3 ) ₂ and _Na2S were prepared in methanol. 25 mg of mPEG-DOPA (PEG molecular weight = 2000) was dissolved in 5 ml of 2 mM Cd(NO ₃ ) ₂ methanol solution, and then 5 ml of 2 mM Na ₂ S solution was slowly added with a syringe at a rate of 20 μl s ⁻¹ . The solution gradually turned yellow during the addition. No yellow precipitate was observed, and dynamic light scattering showed that the particles had an average diameter of 2.5 nm. Control experiments performed without polymer or in the presence of mPEG-OH gave a yellow precipitate and a clear colorless supernatant. Those skilled in the art will appreciate that various other inorganic particulate substrates can be prepared in the presence of adhesive compositions of the type described herein, depending on the materials selected and the corresponding ion substitution or exchange reactions.

Example 16c

The polymeric coupling compositions of the present invention can also be used for the stabilization of a variety of other semiconducting materials. For example, core-shell nanoparticles can be surface stabilized as described herein.

Example 17

The optimized experiments in Examples 17-20 were performed with mPEG-DOPA-5K. Several parameters were determined to optimize the adsorption of mPEG-DOPA on gold from solution, including the type and pH of solvent, adsorption time and mPEG-DOPA solution concentration. Cell adhesion and spreading did not differ greatly using the adsorption solvent. The number of cells on the substrate and their total protruding area were not significantly different between DCM and the three different aqueous solutions. The adsorbed substrates in neutral, alkaline, and organic mPEG-DOPA solutions all had significantly enhanced antifouling properties compared to unmodified substrates (p<0.01). Although no differences were observed in cell adhesion and spreading between solutions, the contact angle data support the use of organic solvents as a way to reduce catechol oxidation in optimizing modification strategies. Furthermore, only surface modifications performed in DCM were shown to have significantly fewer cells on the surface and a lower total protruding cell area.

Example 18

Cell adhesion and spreading showed a strong dependence on the concentration of mPEG-DOPA solution (Figure 12). Cell adhesion and spreading were significantly reduced on substrates modified with more than 25 mg/ml mPEG-DOPA compared to the original gold surface (p<0.001) and the surface modified with 10 mg/ml solution (p<0.05). There was no difference in cell adhesion and spreading at concentrations below 10 mg/ml compared to unmodified substrates. No differences were observed between the modified surfaces in cell adhesion and spreading in 25-75 mg/ml mPEG-DOPA solution.

Example 19

Prolonged mPEG-DOPA adsorption time period also only observed less fibroblast adhesion and spreading. While substrate modification as short as 5 minutes appeared to reduce cell adhesion and spreading, an adsorption time of 24 hours resulted in significantly less cell adhesion and spreading on PEGylated substrates than on unmodified substrates (p<0.001) and Substrates treated for a short time (p<0.05).

Example 20

The morphology of fibroblasts cultured on unmodified and PEG-modified surfaces was observed by electron microscopy (Hitachi 3500 SEM). Fibroblasts on unmodified Au and mPEG-OH-modified Au usually spread out and spread well, while those cultured on mPEG-DOPA-modified Au spread far less ( Figure 14A-C). It should be noted that the number of cell protrusions, which play a role in cell adhesion and local adhesion by integrins, was observed on the mPEG-DOPA surface compared to other surfaces. Fig. 1 3 shows that fibroblasts were treated with mPEG-DOPA 5K, mPEG-MAPd 2K or mPEG-MAPd 5K in bare Au, Au treated with mPEG-OH and under optimized conditions (50mg/ml, treated for 24 hours). Adhesion and diffusion differences on treated Au. Cell adhesion and spreading were significantly less on surfaces modified with DOPA-containing conjugates than on either of the other two surfaces. While mPEG-MAP5K modification resulted in a 97% reduction in total neurite cell area and a 91% reduction in cell density on the surface, this reduction was much greater than that achieved with mPEG-DOPA 2K modification.

Differences in cell adhesion and spreading between surfaces modified with DOPA- and MAPd-conjugated PEG in Figure 13 can be attributed to the physical properties of the associated PEG adhesion layer. Analysis of the SPR results indicated that thicker and stronger adhesion layers with higher PEG concentrations were formed with MAPd-PEG than equivalent molecular weight PEGs immobilized with DOPA. Thicker adhesion layers made with MAPd-mediated PEGylation were more successful in inhibiting protein adsorption and thus cell adhesion.

Example 21

Synthesis of Boc-DOPA(TBDMS) ₂ -Osu

N-Hydroxysuccinimide (NHS) (0.110 g, 0.95 mmol) was added to a solution of Boc-DOPA(TBDMS) ₂ (0.500 g, 0.95 mmol) in dry dichloromethane (DCM) (8.0 mL). The solution was stirred on an ice bath, and 1,3-dicyclohexylcarbodiimide (DCC) (0.197 g, 0.95 mmol) was added under nitrogen atmosphere. The reaction was stirred at 0°C for 20 minutes, then allowed to warm to room temperature and stirred for an additional 4 hours. The reaction mixture was filtered to remove urea by-product, which was then evaporated to 1/5 of its original volume. The solution was cooled to 4°C, allowed to stand for 2 hours to precipitate residual urea by-product, filtered and evaporated to give Boc-DOPA(TBDMS) ₂ -OSu (0.567 g, 96% yield) as white foam.

Example 22

Synthesis of Boc-DOPA ₂ (TBDMS) ₄

Boc-DOPA(TBDMS) ₂ -OSu (0.567 g, 0.91 mmol) was dissolved in dry dimethylformamide (DMF) (2.5 mL), then DOPA(TBDMS) ₂ (0.405 g, 0.95 mmol). The mixture was stirred on an ice bath, and diisopropylethylamine (DIEA) (158 μL, 0,91 mmol) was added dropwise by syringe. After 20 minutes, the reaction was warmed to room temperature, stirred for 17 hours, filtered (if necessary), diluted with ethyl acetate (EtOAc) (40 mL), transferred to a separatory funnel, and rinsed with 5% aqueous HCl. Extract from the aqueous layer with EtOAc. The organic layers were combined and washed with 5% aqueous HCl (3x), _H2O (1x), dried over _MgSO4 , and evaporated to give Boc- _DOPA2 (TBDMS) ₄ (0.83 g, 98% yield) as a white foam.

Example 23

Synthesis of Boc-DOPA ₂ (TBDMS) ₄ -Osu

The procedure of Example 21 was repeated using Boc-DOPA ₂ (TBDMS) ₄ to obtain Boc-DOPA ₂ (TBDMS) ₄ -Osu.

Example 24

Synthesis of Boc-DOPA ₃ (TBDMS) ₆

The procedure of Example 22 was repeated using Boc-DOPA ₂ (TBDMS) ₄ -OSu to obtain Boc-DOPA ₃ (TBDMS) ₆ .

Example 25

Synthesis of DOPA ₂

Boc- _DOPA2 (TBDMS) ₄ (0.5 g, 0.54 mmol) was dissolved in saturated HCl/EtOAc (3 mL), and the solution was stirred under nitrogen. After 5 hours, additional HCl gas was bubbled gently through the solution for 25 minutes. The reaction was allowed to stand overnight before being concentrated to  of its original volume. The resulting precipitate was collected by centrifugation, washed with cold EtOAc (3x), and dried to obtain _DOPA2 (0.15 g, 74% yield) as a white powder. The product was further purified by preparative RP-HPLC and characterized by ESI-MS.

Example 26

Synthesis of DOPA ₃

Boc- _DOPA3 (TBDMS) ₆ (1.06 g, 0.79 mmol) was dissolved in saturated HCl/EtOAc (3 mL), and the solution was stirred under nitrogen. After 12 hours, additional HCl gas was gently bubbled through the solution for 30 minutes, then the reaction was continued for 40 hours. Bubble the solution again with more HCl gas for 30 min, stop stirring. The resulting precipitate was collected by centrifugation, washed with cold EtOAc (3x), and dried to give _DOPA3 (0.424 g, 96% yield) as a white powder. The product was further purified by preparative RP-HPLC and characterized by ESI-MS.

Example 27

Synthesis of mPEG-DOPA _1-3

0.1 M borate buffer (50 mL, pH 8.5) was degassed with argon for 20 minutes and L-DOPA (0.197 g, 1.0 mmol) was added. After the solution was stirred for 15 minutes, methoxy-terminated PEG-SPA (mPEG-SPA) 5K (0.5 g, 0.1 mmol) was added in portions and the reaction was stirred for 3 hours. The resulting clear solution was then acidified to pH 1-2 with aqueous HCl and extracted with DCM (3x). The combined organic layers were washed with 0.1M HCl, dried over _MgSO4 , and concentrated. The remaining residue was dissolved in DCM and precipitated 3 times with ethyl acetate to give rmPEG-DOPA (0.420 g, 84% yield) as a white powder. The product was characterized by MALDI-MS and ¹ H NMR spectroscopy.

Example 28

Surface Modification

Solid metal substrates (Al, 316L stainless steel, and NiTi) were ground and polished, and finally 0.04 m colloidal silica (Syton, DuPont) was used. 20nm TiO ₂ or 10nm TiO ₂ /40nm Au was vapor-deposited onto the Si wafer at <10 ^-6 Torr using an Edwards FL400 electron beam evaporator, and then cut into 8mm x 8mm pieces. All substrates were ultrasonically cleaned for 20 minutes in each of the following: 5% Contrad70 (Fisher Scientific), ultrapure _H2O , acetone, and petroleum ether. The surface was then further cleaned by exposure to _O2 plasma (Harrick Scientific) at 150 mTorr and 100 W for 5 min. To prevent the formation of a gold oxide (Au ₂ O ₃ ) layer, some Au substrates were not exposed to the O ₂ plasma. To create a surface similar to biopolymers, glass coverslips (Fisher Scientific) were cleaned as described above and soaked in 0.01% poly-L-lysine (Sigma) solution for 5 min with ultra- Wash with pure _H2O and dry under nitrogen.

In order to investigate various modification conditions with a minimum number of samples, a 9-element Robust Design approach was used. Substrates were modified by soaking in mPEG-DOPA _1-3 in 0.6 M _K2SO4 solution buffered with 0.1 M _MOPS at 50 °C under cloud point conditions. The pH of the buffer, modification time and mPEG-DOPA concentration were modified as shown in Table 1. The modified substrates were then washed with ultrapure _H2O and dried under nitrogen flow.

Table 1 test anchor group buffer pH Adsorption time polymer concentration 1 -DOPA 3.0 1h 0.5mg/mL 2 -DOPA 6.0 4h 1.0mg/mL 3 -DOPA 9.0 24h 3.0mg/mL 4 -DOPA ₂ 3.0 4h 3.0mg/mL 5 -DOPA ₂ 6.0 24h 0.5mg/mL 6 -DOPA ₂ 9.0 1h 1.0mg/mL 7 -DOPA ₃ 3.0 24h 1.0mg/mL 8 -DOPA ₃ 6.0 1h 3.0mg/mL 9 -DOPA ₃ 9.0 4h 0.5mg/mL

Example 28a

_TiO2 substrate

Under cloud point conditions, the _TiO2 substrate was modified by soaking the substrate in mPEG-DOPA _1-3 with 0.1M N-3-morpholinopropanesulfonic acid (MOPS) at 50 °C. Buffered 0.6M K ₂ SO ₄ solution for 24 hr. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28b

Under cloud point conditions, 316L stainless steel (Goodfellow, Devon PA) was modified as follows: at 50°C, the substrate was soaked in mPEG-DOPA _1-3 with 0.1M N-3-morpholinopropanesulfonate acid (MOPS) buffered 0.6M K ₂ SO ₄ solution for 24 hours. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28c

Under cloud point conditions, Al ₂ O ₃ (Goodfellow, Devon PA) was modified by soaking the substrate in mPEG-DOPA _1-3 with 0.1M N-3-morpholinyl at 50°C. propanesulfonic acid (MOPS) buffered 0.6M K ₂ SO ₄ solution for 24 hours. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28d

Under cloud point conditions, SiO ₂ (1500 Å thermal oxide, University Wafer, South Boston, MA) was modified by soaking the substrate in mPEG-DOPA _1-3 with 0.1 MN-3-morpholinopropanesulfonic acid (MOPS) buffered 0.6M _{K2SO4 solution} for 24 hours. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28e

NiTi alloys (10 mm × 10 mm × 1 mm) were obtained from Nitinol Devices & Components (Fremont, CA), and were modified under cloud point conditions by soaking the substrate in mPEG-DOPA _1-3 0.6 M K ₂ SO ₄ solution buffered with 0.1 M N-3-morpholinopropanesulfonic acid (MOPS) for 24 hours. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28f

Under cloud point conditions, Au (deposited by electron beam evaporation on Si wafers from University Wafer) was modified by immersing the substrate in mPEG-DOPA _1-3 with 0.1 MN-3-morpholinopropanesulfonic acid (MOPS) buffered 0.6M _{K2SO4 solution} for 24 hours. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28g

Au ₂ O ₃ (the Au sample described in Example 28f was exposed to an oxygen plasma to form Au ₂ O ₃ ) was modified under cloud point conditions by soaking the substrate at 50°C mPEG-DOPA _1-3 in 0.6M _K2SO4 buffered with 0.1M N-3-morpholinopropanesulfonic acid ( _MOPS ) for 24 hours. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28h

Under cloud point conditions, GaAs (University Wafer, South Boston, MA) was modified as follows: at 50°C, the substrate was soaked in mPEG-DOPA _1-3 with 0.1M N-3-morpholino propanesulfonic acid (MOPS) buffered 0.6M K ₂ SO ₄ solution for 24 hours. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 28i

The pL-Lys surface was prepared as follows: glass coverslips (Fisher Scientific) were soaked in 0.01% poly-L-lysine (pL-Lys, Sigma) for 5 min, washed with ultrapure _H2O , and incubated under N ₂ strokes to dry. Then, under cloud point conditions, they were modified by soaking the substrate in mPEG-DOPA _1-3 buffered with 0.1M N-3-morpholinopropanesulfonic acid (MOPS) at 50°C. 0.6 M K ₂ SO ₄ solution for 24 hr. The modified substrates were washed with ultrapure _H2O and dried under nitrogen flow.

Example 29

cell adhesion

Passage 12-16 3T3 Swiss albino fibroblasts (ATCC, Manassas, VA) were cultured at 37°C and 5% CO in _media supplemented with 10% fetal bovine serum (FBS) (Cellgro, Herndon, VA) Dulbecco's Modified Eagle's Medium (DMEM) (Cellgro, Herndon, VA) with , 100 g/mL penicillin and 100 U/mL streptomycin. Prior to cell adhesion assays, fibroblasts were harvested with 0.25% trypsin-EDTA, resuspended in growth medium, and counted using a hemocytometer.

General steps of the 4-hour analysis

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4-h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min and then treat with 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetraformaldehyde Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Acquired with a Leica epifluorescent microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) by acquiring 9-16 images (depending on substrate size) at arbitrary positions on each substrate. Quantitative cell adhesion data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29a

_TiO2 substrate (4 hour analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4-h cell adhesion assays, fix and recover cells in 3.7% paraformaldehyde for 5 min, followed by 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl Indocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29b

_TiO2 substrate (long-term research)

For long-term studies of _TiO , twice-weekly reseeding was performed on the substrate at the same density as for the 4-h analysis. At periodic intervals, non-adherent cells were removed by aspirating medium from each well.

Example 29c

316L stainless steel substrate (4 hour analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4 h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min, followed by 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29d

Al ₂ O ₃ substrate (4 hour analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4-h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min and then treat with 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetraformaldehyde Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29e

_SiO2 substrate (4 hour analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4 h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min, followed by 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29f

NiTi substrate (4 hours analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4 h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min, followed by 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29g

Au substrate (4 hours analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4-h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min and then treat with 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetraformaldehyde Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29h

Au ₂ O ₃ substrate (4 hour analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4 h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min, followed by 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 29i

GaAs substrate (4 hours analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4 h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min, followed by 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assays are reported.

Example 29j

p-L-Lys substrate (4 hour analysis)

Prepare test substrates in 12-well tissue culture polystyrene plates filled with 1.0 mL of DMEM containing FBS for 30 min at ₃₇ °C and 5% CO. Seed the cells on the substrate at a density of 2.9 × ¹⁰³ cells/ ^cm2 and maintain in DMEM containing 10% FBS for 4 h at 37 °C and 5% _CO2 . For 4 h cell adhesion assays, fix adherent cells in 3.7% paraformaldehyde for 5 min, followed by 5 μM 1,1'-dioctadecyl-3,3,3',3'-tetramethyl Indolocarbocyanine perchlorate (DiI) (Molecular Probes, Eugene, OR) in DMSO was stained at 37°C for 45 minutes.

Quantitative cell adhesion was obtained using a Leica epifluorescence microscope equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI) to acquire 9-16 images (depending on substrate size) at arbitrary positions on each substrate. attached data. The total protruding cell area of the resulting images was quantified using thresholding in MetaMorph. (Universal Imaging Corporation ^(TM) , a subsidiary of Molecular Devices Corporation, Downington, PA). The mean and standard deviation of the assay results are reported.

Example 30

24-hour modification of substrate and 4-hour analysis

Each surface was modified in 1.0 mg/mL mPEG- _DOPA3 (or mPEG-OH as a control) at the pH values shown in Figure 24 for 24 hours at 50°C. 4 hour cell adhesion and spreading assays were performed as described in Example 9 above. The results are shown in Figure 24. The cell adhesion resistance of all substrates treated with mPEG-DOPA ₃ was compared. Cell adhesion and spreading on mPEG-OH-treated substrates were indistinguishable from unmodified surfaces (data not shown).

Example 31

Surface and Surface Preparation

_TiO2 (20 nm) was coated on silicon wafers (WaferNet GmbH, Germany) by reactive magnetron sputtering (PSI, Villigen, Switzerland) by physical vapor deposition. The metal oxide coated wafers were then cut into 1 cm x 1 cm pieces for ex-situ ellipsometry. The optical waveguide plate used for OWLS measurements was purchased from Microvacuum Ltd. (Budapest, Hungary), and consisted of an AF45 glass substrate (8 × 12 × 0.5 mm) and a 200-nm-thick Si _0.25 Ti _{0.75 O} waveguide surface layer. An 8 nm layer of _TiO2 was deposited on top of the waveguide layer using the same conditions as described above for the silicon wafer. Before polymer modification, silicon wafers and waveguides coated with _TiO2 were sonicated in 2-propanol for 10 min, rinsed in ultrapure water, and dried under nitrogen flow before exposing them to O ₂ plasma (Harrick Scientific, Ossining, USA) for 3 minutes to remove all organic components from the surface. After the OWLS measurement, the waveguide was regenerated for reuse as follows: the waveguide was sonicated in a cleaning solution (300 mM HCl, 1% detergent; Roche Diagnostics, Switzerland) and then rinsed with ultrapure water to remove Adsorbed substance.

Surface Modification.

Using cloud point buffer (CP buffer: 0.6M K ₂ SO ₄ buffered to pH=6.0 with 0.1M MOPS), in the temperature range of 25-50°C, with mPEG-DOPA _1-3 prepared according to Example 27 The surface was modified for 24 hours using a polymer concentration of 1.0 mg/ml. Substrates were rinsed with water after modification, dried under a flow of _N2 , and immediately analyzed as described below.

X-ray Photoelectron Spectroscopy (XPS) Determination

Acquired on a SAGE 100 (SPECS, Berlin, Germany) using a standard (non-monochromatizing) AlK _α X-ray source operating at 325 W (13 kV, 25 mA) and a take-off angle of 0° Measurement results and high-resolution spectra, , The off-angle is defined as the angle between the photoelectron detector and the surface normal. The pass energies for measurement and high-resolution spectroscopy were 50 eV and 14 eV, respectively. During data acquisition, the pressure in the analysis chamber was kept below 2×10 ⁻⁸ Pa. All XPS spectra are referenced to the aliphatic hydrocarbyl component in the Cls signal at 287.4 eV. With CasaXPS software, Shirley background subtraction and the sum of 90% Gaussian function and 10% Lawrence function were used for curve fitting. The atomic sensitivity factor is used to convert the measured intensities (peak areas) into normalized intensities, from which the atomic composition of the surface can be calculated. Average values for triplicate replicate substrates are reported in Tables 7-8. Standard deviations are typically less than 10% of the mean and are omitted for clarity.

Table 7

Quantitative analysis of XPS data for mPEG-DOPA modified _TiO2 surface surface Atomic concentration (atomic%) Ti o C Ols _A TiO ₂ Ols _B TiOH Ols _C CO, H ₂ O Cls _A CC, CH Cls _B CO Cls _C NHC (=O) Clean TiO ₂ mPEG-DOPA ₁ mPEG-DOPA ₂ mPEG-DOPA ₃ 24.317.911.17.4 50.936.822.515.9 14.15.23.20.5 4.113.120.225.3 4.52.83.74.0 1.223.137.343.7 0.91.22.13.2

Table 8

Atomic ratios calculated from XPS data of mPEG-DOPA-modified _TiO2 surface surface Atomic ratio ^a C/Ti O _A /Ti O _B /O _A C _B /C _A C _C /C _B C _C /C _A Cleaned TiO ₂ mPEG-DOPAmPEG-DOPA ₂ mPEG-DOPA ₃ 0.271.523.896.87 2.102.062.032.14 0.280.140.140.03 0.338.4210.0711.00 0.420.0520.0550.074 0.200.420.560.80

^a Contributions A, B and C are defined in Table 7.

Spectroscopic Ellipsometry

For the ELM determination, Si substrates sputtered with _TiO2 were ex situ modified as described above, and the temperature of the modification solution was varied between 25–50 °C. After modification, the substrate was washed with H ₂ O, incubated in 10 mM HEPES buffer (pH=7.4) at room temperature for 48 hours, washed with H ₂ O again, and dried with N ₂ . To detect protein resistance, modified and unmodified substrates were exposed to pure human serum for 15 min, washed with water, and dried in a flow of _N2 . ELM was performed on an M-2000D spectroscopic ellipsometer (JAWollam Co., Inc., Lincoln, USA) at 65°, 70° and 75° before and immediately after modification, after HEPES incubation and after serum exposure Determination, the wavelength used is 193-1000nm. In the WVASE32 analysis software, the optical properties of the generalized Cauchy polymer layer (A _n =1.45, B _n =0.01, C _n =0) were used to fit the ELM spectrum with a multi-layer model to obtain the Adsorbed PEG and the "dry" thickness of the serum adhesion layer. (“Dry” or dehydrated thickness refers to the thickness measured under ambient conditions after drying with _N2 ). The average thickness of triplicate determinations is reported in Tables 9-10.

Table 9

Effect of Adsorption Temperature on the Thickness of PEG Adhesive Layer on TiO ₂ ^a Adsorption temperature (°C) Thickness (A) 2528313437404550 10.6±1.813.9±0.316.4±1.119.4±1.919.6±3.522.5±2.024.4±2.033.8±4.6

^a TiO ₂ surfaces were exposed to mPEG-DOPA ₃ (1 mg/ml) for 2 hours, then each surface was washed in HEPES for 48 hours.

Table 10

Apparent thickness (A) of the organic adhesion layer on _TiO2 determined by spectroscopic ellipsometry deal with ^a mPEG-DOPA ₃ adsorption time (minutes) 0 1 30 60 240 1080 Before exposure to serum After exposure to serum -60.2 5.123.4 24.224.7 27.6< ^28.1b 31.8< ^32.3b 35.0< ^35.5b

^a TiO ₂ surface was exposed to mPEG-DOPA ₃ (1 mg/ml) for 0–1080 min, washed with water, incubated in HEPES for 48 h, and then exposed to serum for 15 min.

^b The net increase in the thickness of the adsorbed layer after exposure to serum is less than 0.5 Å, which is close to the resolution of the ELM technique.

Optical Waveguide Lightmode Spectroscopy (OWLS).

The TiO2 _- coated waveguides were cleaned with 2-propanol and _O2 plasma as described above. The cleaned waveguide was mounted on the assay head of OWLS110 (Microvacuum Ltd.) and stabilized at room temperature in cloud point buffer (CP buffer: 0.6M _K2SO4 buffered to pH= _6.0 with 0.1M MOPS) Leave for at least 48 hours. The stabilization phase allows the ion exchange on the _TiO2 surface to reach equilibrium and achieve a stable baseline. To monitor polymer adsorption, mPEG-DOPA in CP buffer was injected in stop-flow mode, followed by injection of CP buffer to remove unbound PEG, after which the signal was stabilized. The incoupling angles - α _TM and α _TE - were recorded and converted to refractive indices (N _TM , N _TE ) by the software provided by the manufacturer. The real-time change in the sensor's effective refractive index was converted into adsorbed mass using the de Feijter formula. Nostalgia for reference to be included [de Feijter, 1978 #14] Adding the refractive index of each mPEG-DOPA polymer using linear interpolation between 0.13 cm ³ /g for pure PEG and 0.18 cm ³ /g for pure polyamino acid - dn/dc is calculated. For the protein adsorption assay, the temperature of the assay head was equilibrated at 37 °C until the signal stabilized, and then the serum was injected for 15 min before the injection buffer. The mass of adsorption did not change significantly with the prolongation of serum exposure time.

Example 32

Synthesis of N-methacryloyl 3,4-dihydroxy-L-phenylalanine

7.1d(1H，-NH-)；6.6-6.8(3H，C₆H₃(OH)₂-)；5.68s(1H，CHH＝)；5.632s(unknownpeak)；5.33s(1H，CHH＝)；4.67m(1H，-CH-)；2.93-3.1m(2H，CH₂-)；1.877s(3H，-CH₃)。1.15 g (5.69 mmol) Na ₂ B ₄ O ₇ was dissolved in 30 ml water. The solution was degassed with Ar for 30 minutes, then 0.592 g (3.0 mmol) of L-DOPA was added and stirred for 15 minutes. Then 0.317g (3.0mmol) Na ₂ CO ₃ was added, the solution was cooled to 0°C, and 0.3ml (3.0mmol) methacryloyl chloride was slowly added with stirring. During the reaction, the pH of the solution was maintained at 9 _{with Na2CO3} . After stirring at room temperature for 1 hour, the solution was acidified to pH 2 with concentrated HCl. The mixture was extracted 3 times with ethyl acetate. After washing with 0.1N HCl, drying over anhydrous _MgSO4 , the solvent was removed in vacuo to obtain a light brown crude solid. The product was further purified by eluting from a silica gel column with dichloromethane (DCM) and methanol (95:5). After removal of the solvent by evaporation, a white sticky solid was obtained in 35% yield. ¹ H NMR (500MHz, acetone-d ₆ ):

7.1d(1H, -NH-); 6.6-6.8(3H, _C6H3 (OH) _2- ); 5.68s(1H, CHH=); 5.632s ₍ unknownpeak); 5.33s(1H, CHH=) ; 4.67m (1H, -CH-); 2.93-3.1m (2H, _CH2- ); 1.877s (3H, _-CH3 ).

Example 33

Synthesis of 3,4-bis(tert-butyldimethylsilyloxy)-L-phenylalanine

3.60 g (24.0 mmol) of TBDMS-Cl were dissolved in 18 ml of anhydrous acetonitrile. 1.60 g (8.0 mmol) of L-DOPA were added to the solution, the suspension was stirred and cooled to 0° C., then 3.6 ml of DBU (24.0 mmol) were added. The reaction mixture was then stirred at room temperature for 24 hours. Cold acetonitrile was added to the reaction solution to obtain a colorless precipitate. The precipitate was filtered and washed several times with cold acetonitrile, then dried in vacuo. A white powder was obtained in 78% yield. ¹ H NMR (500MHz, methanol-d); 6.7-6.9 (eH, C ₆ H ₃ (O-Si-) ₂ -); 3.72 (m, 1H, -CH-); 2.82-3.2 (m, 2H, -CH ₂ -); 1.0 (d, 18H , -C( _CH3 )); 0.2(d, 12H, Si- _CH3 ).

Example 34

Synthesis of 3,4-bis(tert-butyldimethylsilyloxy)-N-tert-butoxycarbonyl-L-phenylalanine

6.68-6.81(3H，C₆H₃(O-Si-)₂-)；4.28(m，1H，-CH-)；2.78-3.08(m，2H，-CH₃-)；1.4(s，9H，-O-C(CH₃)₃)；1.0(d，18H，-Si-C(CH₃)₃)；0.2(d，12H，Si-(CH₃)₂)。1.60 g (3.77 mmol) of 3,4-bis(tert-butyldimethylsilyloxy)-L-phenylalanine was added to 10 ml of water containing 0.34 g (4.05 mmol) of NaHCO ₃ . A solution of 0.96 g (4.30 mmol) of di-t-butyl dicarbonate in 10 ml of tetrahydrofuran was added and the reaction mixture was stirred at room temperature for 24 hours. After evaporation of the tetrahydrofuran, 10 ml of water were added to the residue. The solution was acidified to pH 5 with dilute HCl and extracted ₃ times with ethyl acetate. After drying over anhydrous MgSO, the solvent was removed in vacuo. The crude product was purified by column chromatography (silica gel; eluent; 10% methanol in DCM solution). After evaporating off the eluting solvent, a white solid was obtained with a yield of 70%. ¹ H NMR (500MHz, methanol-d);

6.68-6.81(3H, C ₆ H ₃ (O-Si-) ₂ -); 4.28(m, 1H, -CH-); 2.78-3.08(m, 2H, -CH ₃ -); 1.4(s, 9H , -OC(CH ₃ ) ₃ ); 1.0 (d, 18H, -Si-C(CH ₃ ) ₃ ); 0.2 (d, 12H, Si-(CH ₃ ) ₂ ).

Example 35

Synthesis of 3,4-bis(tert-butyldimethylsilyloxy)-N-tert-butoxycarbonyl-L-phenylalanine pentafluorophenyl ester

6.65-6.81(3H，C₆H₃(O-Si-)₂-)；4.85(m，1H，-CH-)；3.05-3.2(m，2H，-CH₃-)；1.41(s，9H，-O-C(CH₃)₃)；1.0(d，18H，-Si-C(CH₃)₃)；0.2(d，12H，Si-(CH₃)₂)。1g (1.90mmol) 3,4-bis(tert-butyldimethylsilyloxy)-N-tert-butoxycarbonyl-L-phenylalanine and 0.351g (1.90mmol) pentafluorophenol (pentafluorophenol) Dissolve in a solvent mixture consisting of 24 ml of dioxane and 1 ml of DMF, and add 0.432 g (2.10 mmol) of DCC at 0°C. The solution was stirred at 0°C for 1 hour, then at room temperature for 1 hour, after which the solution was filtered to remove dicyclohexylurea and evaporated in vacuo. Product 4 was purified by column chromatography (silica gel; eluent; hexane/ethyl acetate=11.2). After removal of the eluent, a pure white sticky solid was obtained in 55% yield. ¹ H NMR (500MHz, CDCl ₃ );

6.65-6.81(3H, C ₆ H ₃ (O-Si-) ₂ -); 4.85(m, 1H, -CH-); 3.05-3.2(m, 2H, -CH ₃ -); 1.41(s, 9H , -OC(CH ₃ ) ₃ ); 1.0 (d, 18H, -Si-C(CH ₃ ) ₃ ); 0.2 (d, 12H, Si-(CH ₃ ) ₂ ).

Example 36

N-(13'-amino-4',7',10'-trioxatridecyl)-tert-butoxycarbonyl-3',4'-bis(tert-butyldimethylsilyloxy)- Synthesis of L-phenylpropanamide

7.38(m，1H，-CONH-)；6.60-6.80(3H，C₆H₃(O-Si-)₂-)；5.26(m，1H，-CONH-)；4.30(m，1H，-CH-)；3.4-3.8(m，12H，-CH₂O-；3.03-3.4(m，4H，-CH₂-NH-，-CH₂-NH₂)；2.78-3.02(m，2H，-CH₂-)；2.0(m，2H，-CH₂-)；1.7(m，2H，-CH₂-)；1.39(s，9H，-O-C(CH₃)₃)；1.0(d，18H，Si-C(CH₃)₃)；0.2(d，12H，Si-C(CH₃)₂)。At 0°C, 0.869g (1.26mmol) of 3,4-bis(tert-butyldimethylsilyloxy)-N-tert-butoxycarbonyl-L-phenylalanine pentafluorophenyl ester in 10ml DCM solution was added dropwise to a mixture of 2.07ml (9.44mmol) 4,7,10-trioxa-1,13-tridecanediamine and 1.32ml (9.44mmol) Et ₃ N in 30 minutes. 1ml of the resulting mixture in DMF. The solution was stirred for an additional 2 hours at room temperature, then the solvent was removed in vacuo. The crude product was loaded on silica gel and eluted with DCM, 5% methanol in DCM, 10% methanol in DCM and 15% methanol in DCM. The solvent was removed under vacuum to afford compound 5 as a white solid. The yield was 63%. ¹ H NMR (500 MHz, acetone-d ₆ );

7.38(m, 1H, -CONH-); 6.60-6.80(3H, C _{6 H 3} (O-Si-) ₂ -); 5.26(m, 1H, -CONH-); 4.30(m, 1H, -CH -); 3.4-3.8(m, 12H, -CH ₂ O-; 3.03-3.4(m, 4H, -CH ₂ -NH-, -CH ₂ -NH ₂ ); 2.78-3.02(m, 2H, -CH ₂ -); 2.0 (m, 2H, -CH ₂ -); 1.7 (m, 2H, -CH ₂ -); 1.39 (s, 9H, -OC (CH ₃ ) ₃ ); 1.0 (d, 18H, Si -C(CH ₃ ) ₃ ); 0.2(d, 12H, Si—C(CH ₃ ) ₂ ).

Example 37

N-(13-(N'-tert-butoxycarbonyl-L-amino-3',4'-bis(tert-butyldimethylsilyloxy)-4,7,10-trioxatridecyl )-Methacrylamide Synthesis

6.60-6.80(3H，C₆H₃(O-Si-)₂-)；6.40(m，1H，-CONH-)；5.71s(1H，CHH＝)；5.30s(1H，CHH＝)；5.096(m，1H，-CONH-)；4.21(m，1H，-CH-)；3.2-3.65(m，16H，-CH₂O，-CH₂-NH--CH₂-NH₂)；2.80-2.99(m，2H，-CH₂-)；1.96s(3H，-CH₃)；1.81(m，2H，-CH₂-)；1.68(m，2H，-CH₂-)；1.40(s，9H，-O-C(CH₃)₃)；1.0(d，18H，Si-C(CH₃)₃)；0.2(d，12H，Si-C(CH₃)₂)。0.57g (0.79mmol) of N-(13'-amino-4',7',10'-trioxatridecyl)-tert-butoxycarbonyl-3',4'-bis(tert-butyldimethyl Methylsilyloxy)-L-phenylpropanamide and 0.166ml (1.18mmol) of Et ₃ N were dissolved in 5ml of anhydrous chloroform, and 0.176ml (1.18mmol) of methacrylic anhydride was added thereto. The solution was stirred at room temperature for 3 hours, then the solvent was removed under vacuum. Pure compound 6 was obtained by column chromatography in 61% yield as a white viscous solid (silica gel; eluent: ethyl acetate). ¹ H NMR (500MHz, CDCl ₃ );

6.60-6.80 (3H, C ₆ H ₃ (O-Si-) ₂ -); 6.40 (m, 1H, -CONH-); 5.71s (1H, CHH=); 5.30s (1H, CHH=); 5.096 (m, 1H, -CONH-); 4.21 (m, 1H, -CH-); 3.2-3.65 (m, 16H, -CH ₂ O, -CH ₂ -NH--CH ₂ -NH ₂ ); 2.80- 2.99 (m, 2H, -CH ₂ -); 1.96s (3H, -CH ₃ ); 1.81 (m, 2H, -CH ₂ -); 1.68 (m, 2H, -CH ₂ -); 9H, -OC(CH ₃ ) ₃ ); 1.0 (d, 18H, Si-C(CH ₃ ) ₃ ); 0.2 (d, 12H, Si-C(CH ₃ ) ₂ ).

Example 38

Synthesis of N-(13-(N'-t-Boc-L-3',4'dihydroxyphenylalaninyl)-4,7,10-trioxatridecyl)-methacrylamide

7.90(m，1H，-CONH-)；7.23-7.40(d 2H，C₆HH₂(OH)₂-)；6.56-6.76(3H，C₆HH₂(OH)₂-；5.930(m，1H，-CONH-)；5.71s(1H，CHH＝)；5.30s(1H，CHH＝)；4.20(m，1H，-CH-)；3.1-3.60(m，16H，-CH₂O，-CH₂-NH-，-CH₂-NH₂)；2.70-2.95(m，2H，-CH₂-)；1.96s(3H，-CH₃)；1.78(m，2H，-CH₂-)；1.65(m，2H，-CH₂-)；1.39(s，9H，-O-C(CH₃)₃)。Add 0.344 g (0.433 mmol) N-(13-(N'-tert-butoxycarbonyl-L-amino-3',4'-bis(tert-butyldimethylsilyloxy) to a 10 ml round bottom flask -4,7,10-trioxatridecyl)-methacrylamide, 3 mL THF and 0.137 g (0.433 mmol) TBAF. The solution was stirred at room temperature for 5 minutes, then 3 ml 0.1N HCl was added. The solution was extracted 3 times with DCM , and the solvent was evaporated in vacuo. Compound 7 was obtained as a white solid in 63% yield by column chromatography (silica gel; eluent: 7% methanol in DCM) ^{. 1} H NMR (500 MHz, acetone-d ₆ );

7.90 (m, 1H, -CONH-); 7.23-7.40 (d 2H, C ₆ HH ₂ (OH) ₂ -); 6.56-6.76 (3H, C ₆ HH ₂ (OH) ₂ -; 5.930 (m, 1H , -CONH-); 5.71s (1H, CHH=); 5.30s (1H, CHH=); 4.20 (m, 1H, -CH-); 3.1-3.60 (m, 16H, -CH ₂ O, -CH ₂ -NH-, -CH ₂ -NH ₂ ); 2.70-2.95 (m, 2H, -CH ₂ -); 1.96s (3H, -CH ₃ ); 1.78 (m, 2H, -CH ₂ -); 1.65 (m, 2H, _-CH2- ); 1.39 (s, 9H, -OC( _CH3 ) ₃ ).

Example 39

Synthesis of PEG-diacrylate (PEG-DA)

40 g (5 mmol) of PEG were dried by azeotropic evaporation in benzene, then dissolved in 150 mL of DCM. 4.18 mL (30 mmol) of Et ₃ N and 3.6 mL (40 mmol) of acryloyl chloride were added to the polymer solution. The mixture was refluxed with stirring for 5 hours and allowed to cool at room temperature overnight. Diethyl ether was added to the mixture to form a dark yellow precipitate. The crude product was then dissolved in saturated NaCl solution and heated to 60 °C to form two layers. Add DCM to the upper layer, and add _MgSO to remove moisture. After removing _MgSO4 by filtration, the volume of solvent was reduced under vacuum and the sample was precipitated in diethyl ether. The final product was dried under vacuum and stored at -15 °C. The yield was 75%. ¹ H NMR (500MHz, D ₂ O): δ6.47(d, 1H, CHH=C-); 6.23(m, 1H, C=CH-C(=O)-O-); 6.02(d, 1H , CHH=C-); 4.35 (m, 2H, -CH ₂ -OC(=O)-C=C); 3.23-3.86 (PEG CH ₂ )

PEG-DA photopolymerization

PEG-DA, compound 1, compound 7 and photoinitiator precursor solution were prepared and mixed immediately before photopolymerization. Stock solutions of PEG-DA (200 mg/mL) and compound 1 (40 mg/mL) were dissolved in _N2- purged phosphate-buffered saline (PBS, pH 7.4), while compound 7 (60 mg/mL ) in 50:50 PBS/95% ethanol pre-purged with _N2 . To prepare the final polymeric compound, compound 1 or 7 was combined with PEG-DA to obtain PEG-DA and DHPD derivatives at a final concentration of 150 mg/mL. Then 100 μL of the mixture was added to a disc mold (100 μL, diameter=9 mm, depth=2.3 mm, Secure Seal ⁽ R) SA8R-2.0, Grace Bio Lab, Inc., OR), and a UV lamp (Black Ray ^(R) Lamp, 365 nm, model UVL-56, UVP, CA) or blue light (VIP ^(R) , 400-500 nm, BISCO Inc., IL) irradiation for up to 20 minutes. For UV-induced photocuring, DMPA (600 mg/mL in VP solution) was added to the polymerization solution to obtain a final concentration of 34 mM. CQ (100 mg/mL in VP solution, final concentration = 150 mM) containing DMAB (30 mg/mL in VP solution, final concentration = 151 mM), or AA (100 mg/mL in PBS solution, final concentration = 17 mM) of FNa ₂ (188 mg/mL in PBS, final concentration = 2 mM) as a photoinitiator. Adjust the final concentration of VP to 135-300 mM.

After irradiation, the gel was blotted with filter paper to remove the liquid surface layer and weighed. The gel weight was then divided by the weight of 100 μL of the starting solution to give the percent gel conversion.

Example 41

Determination of DOPA binding

The amount of DOPA incorporated into the photopolymerized gel was determined using a modified colorimetric DOPA assay developed by Waite and Benedict. The photocrosslinked gel was stirred in 3 mL of 0.5N HCl to extract DOPA monomers not incorporated into the gel network. Add 0.9mL nitrite reagent (1.45M sodium nitrite and 0.41M sodium molybdate dihydrate) and 1.2mL 1M NaOH to 0.9mL extraction solution, within 2-4 minutes of adding NaOH, use Hitachi U-2010 The absorbance (500 nm) of the mixture was measured by UV-Vis spectrophotometer. A standard curve was constructed using known concentrations of compounds 1-7.

Example 42

mechanical testing

Hemispherical hydrogels were formed by loading 25 μL of the polymerization mixture onto glass slides treated with 1H,1H,2H,2H-perfluorooctyltrichlorosilane. Gels were irradiated for 10 min, dialyzed against 0.15 M HCl for at least 24 h to extract unbound DOPA monomer, and then equilibrated in PBS for more than 15 min before testing. To determine the modulus of the gel, a hemispherical gel cover was attached to one end of a steel graduated cylinder (diameter = 6 mm, length = 30 mm) with superadhesive. The other end of the graduated cylinder was connected to a piezoelectric stepper motor (IW-701-00, Burleigh Instruments, NY), which was connected in series with a 50 g load transducer (loadtransducer) (FTD-G-50, Schaevitz Sensors, VA). The movement of the steel column in the axial direction was measured with a fiber optic displacement sensor (RC100-GM2OV, Philtec, Inc., MD). The Si wafer coated with _TiO2 was placed under the hydrogel, and the _Tio2 surface was flooded with PBS to maintain the hydration state of the gel. The indenter advances at 5 μm/s until a maximum compressive load of 4 mN is measured.

The modulus of elasticity is calculated by assuming the specific case as a Hertzian mechanism of non-adhesive contact between an incompressible elastic hemisphere and a rigid plane, in which case the Hertzian relationship between load (P _h ) and displacement (δ _h ) becomes for:

${\mathrm{<span\; class="notranslate">\; P</span>}}_{\mathrm{<span\; class="notranslate">\; h</span>}}<span\; class="notranslate">\; =</span>\frac{\mathrm{<span\; class="notranslate">\; 16</span>}{\mathrm{<span\; class="notranslate">\; <figure-callout\; id="1"\; label="R"\; filenames="A20058000618000811.png,A20058000618000862.png"\; state="\{\{state\}\}">R</figure-callout></span>}}^{\mathrm{<span\; class="notranslate">\; 1</span>}<span\; class="notranslate">\; /</span>\mathrm{<span\; class="notranslate">\; 2</span>}}\mathrm{<span\; class="notranslate">\; E.</span>}}{\mathrm{<span\; class="notranslate">\; 9</span>}}{{\mathrm{<span\; class="notranslate">\; \&delta;</span>}}_{\mathrm{<span\; class="notranslate">\; <figure-callout\; id="3"\; label="h"\; filenames="A20058000618000811.png,A20058000618000861.png"\; state="\{\{state\}\}">h</figure-callout></span>}}}^{\mathrm{<span\; class="notranslate">\; 3</span>}<span\; class="notranslate">\; /</span>\mathrm{<span\; class="notranslate">\; 2</span>}}$

where R and E are the radius of curvature and elastic modulus of the hemispherical gel, respectively. The radius of curvature of the gel was determined from the height and width measured from photographs of the gel.

Example 43

Chemical oxidation of PEG-DOPA to hydrogel

4-arm-PEG-amine (PEG-(NH ₂ ) ₄ , M _w =10,000) was purchased from SunBio, Inc. (Walnut Creek, CAv), while linear PEG-bis-amine (PEG-(NH ₂ ) ₂ , _Mw = 3,400) and methoxy-PEG-amine (mPEG- _NH2 , _Mw = 5,000) were purchased from Shearwater Polymers, Inc. (Huntsville, AL). Sephadex ^(R) LH-20 was obtained from Fluka (Milwaukee, WI). N-Boc-L-DOPA dicyclohexyl ammonium salt, sodium periodate (NaIO ₄ ), mushroom tyrosinase (MT, EC 1.14.18.1) and horseradish peroxidase (HRP, EC 1.11.1.17) Obtained from Sigma Chemical Company (St. Louis, MO). Triethylamine ( _Et3N ), hydrogen peroxide (30 wt%, _H2O2 ), sodium molybdate dihydrate _, and sodium nitrite were purchased from Aldrich Chemical Company (Milwaukee, WI). L-Dopa was purchased from Lancaster (Windham, NH). 1-Hydroxybenzotriazole (HOBt) was obtained from Novabiochem Corp. (La Jolla, CA), while hexafluorophosphate O-(benzotriazol-1-yl)-N,N,N′,N′-tetrafluorophosphate Methyluronium (HBTU) was obtained from Advanced ChemTech (Louisville, KY).

Synthesis of PEG modified with DOPA

PEGs modified with linear and branched chain DOPA containing up to 4 DOPA end groups were synthesized using standard carbodiimide coupling chemistry as described below. The structure of PEG modified with 4 DOPAs is shown in FIG. 1 .

PEG-(N-Boc-DOPA) ₄ , Synthesis of PEG-(NH ₂ ) ₄ (6.0 g, 0.60 mmol) with 50:50 dichloromethane (DCM) and dimethylformamide (DMF) in 60 mL N-Boc-L-DOPA dicyclohexyl ammonium salt (4.8 mmol), HOBt (8.0 mmol) and _Et3N (8.0 mmol) were reacted in a mixture. Then a solution of HBTU (4.8 mmol) in 30 mL DCM was added and the coupling reaction was carried out at room temperature under argon for 1 hour. The solution was washed sequentially with saturated sodium chloride solution, 5% NaHCO ₃ , dilute HCl solution and distilled water. The crude product was concentrated under reduced pressure and purified by column chromatography on a Sephadex ^(R) LH-20 column with methanol as the mobile phase. The product was further purified by three precipitations in cold methanol, dried under vacuum at room temperature and stored at -20°C under nitrogen. ¹ H NMR (500MHz, CDCl ₃ /TMS): δ6.81-6.77 (m, 2H, C ₆ HH ₂ (OH) ₂ -), 6.6 (d, 1H, C ₆ H ₂ H(OH) ₂ -) , 6.05(br,s,1H), 5.33(br,s,1H), 4.22(br,s,1H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH(N-)-C(O)N -), 3.73-3.41(m, PEO), 3.06(m, 2H, PEO-CH ₂ -NC(O)-), 2.73(t, 2H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH- ), 1.44(s, 9H, ( _CH3 ) _3C- ). GPC-MALLS: _Mw = 11,900, _Mw / _Mn = 1.01.

Synthesis of PEG-(DOPA) ₄ , II 3.0 g of compound I (0.25 mmol) was dissolved in 15 mL of DCM at room temperature. 15 mL of TFA was added to the mixture for 30 minutes under argon. After evaporating the solvent in a rotary evaporator, the product was precipitated 3 times with cold methanol, dried under vacuum at room temperature, and stored at -20°C under nitrogen. ¹ H NMR (500MHz, D ₂ O): δ6.79(d, 1H, C ₆ H ₂ H(OH) ₂ -), 6.66(s, 1H, C ₆ H ₂ H(OH) ₂ -), 6.59 (d, 1H, C ₆ H ₂ H(OH) ₂ -), 4.00 (t, 1H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH(N-)-C(O)N-), 3.70 -3.34(M, PEO), 3.24(m, 2H, PEG-CH ₂ -NC(O)-), 3.01-2.88(m, 2H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH(N- )-C(O)N-). GPC-MALLS: _Mw = 11,400, _Mw / _Mn = 1.02.

PEG-(N-Boc-DOPA) ₂ , Synthesis of III. PEG-(NH ₂ ) ₂ (5.0g, 1.5mmol), N-Boc-L-DOPA dicyclohexyl ammonium salt (5.9mmol), HOBt ( 9.8 mmol) and _Et3N (9.8 mmol) were dissolved in 50 mL of a 50:50 mixture of DCM and DMF. A solution of HBTU (5.9 mmol) in 25 mL of DCM was then added and the reaction was carried out at room temperature under argon for 30 minutes. The product was recovered and purified as described for compound I. ¹ H NMR (500MHz, CDCl ₃ /TMS): δ6.81-6.77 (m, 2H, C ₆ HH ₂ (OH) ₂ -), 6.59 (d, 1H, C ₆ H ₂ H(OH) ₂ -) , 6.05(br,s,1H), 5.33(br,s,1H), 4.22(br,s,1H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH(N-)-C(O)N -), 3.73-3.42(M, PEO), 3.06(m, 2H, PEO-CH ₂ -NC(O)-), 2.74(t, 2H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH( N-)-C(O)N-), 1.44 (s, 9H, ( _CH3 ) _3CO- ). GPC-MALLS: _Mw = 4,600, _Mw / _Mn = 1.02.

Synthesis of Methoxy-PEG-(N-Boc-DOPA), IV mPEG-NH ₂ (2.0g, 0.40mmol), N-Boc-L-DOPA dicyclohexyl ammonium salt (0.80mmol), HOBt (1.3 mmol) and _Et3N (1.3 mmol) were dissolved in 20 mL of a 50:50 mixture of DCM and DMF. Then a solution of HBTU (0.80 mmol) in 10 mL of DCM was added and the reaction was carried out at room temperature under argon for 30 minutes. The product was recovered and purified as described for compound I. ¹ H NMR (500MHz, CDCl ₃ /TMS): δ6.81-6.60 (m, 3H, C ₆ H ₃ (OH) ₂ -), 6.01 (br, s, 1H, OH-), 5.32 (br, s , 1H, OH-), 4.22 (br, s, 1H, C ₆ H ₃ (OH) ₂ -CH ₂ -CH(N-)-C(O)N-), 3.73-3.38 (m, PEO), 3.07(m, 2H, PEO- _CH2- NH-C(O)- ₎ , 2.73(t, 2H, _C6H3 (OH) _2- CH2 _- CH(N-)-C(O)N- ), 1.44 (s, 9H, (CH ₃ ) ₃ C-), 1.25 (s, 3H, CH ₃ CH ₂ O-). GPC-MALLS: M _w =6,100, M _w /M _n =1.02.

Determination of DOPA content

The DOPA content in DOPA-modified PEG was determined by integration of the relevant peaks in the 1H NMR spectrum and by colorimetric DOPA analysis. In the NMR method, the DOPA content is determined by comparing the Boc methyl proton integral value at δ = 1.44 with the PEG methylene proton integral value at δ = 3.73-3.38. The DOPA assay was based on the aforementioned Waite and Benedict method. Briefly, aqueous PEG-DOPA was treated with nitrite reagent (1.45M sodium nitrite and 0.41M sodium molybdate dihydrate), followed by the addition of excess NaOH solution. Within 2-4 minutes of adding NaOH, the absorbance (500 nm) of the mixture was recorded with a Hitachi U-2010 UV/vis spectrophotometer. A standard curve was constructed using solutions of known DOPA concentrations.

Formation of PEG-DOPA hydrogel

To form PEG-DOPA hydrogel, sodium periodate (NaIO ₄ ), horseradish peroxidase and hydrogen peroxide (HRP/H ₂ O ₂ ) or mushroom tyrosinase and oxygen (MT/O ₂ ) Add PEG-DOPA (200 mg/mL) to a solution of phosphate-buffered saline (PBS, pH 7.4). For gelation induced by MT, flush PBS with air for 20 min before adding MT. The gel time was characterized as the time at which the mixture ceased to flow as determined by inverting the vial containing the liquid of the mixture.

Oscillatory rheometry

Oscillatory rheometry is used to monitor the gelation process and to determine the mechanical properties of hydrogels. The crosslinker was added to the PEG-DOPA aqueous solution, and the well-mixed solution was loaded on a Bohlin VOR rheometer. The analysis was carried out under the following conditions: a frequency of 0.1 Hz, a strain of 1%, a cone with a diameter of 30 mm and a plate fixed at a cone angle of 2.5°.

Spectroscopic Evaluation of DOPA Oxidation

PEG modified with DOPA was dissolved _in 10 mM PBS solution (argon sparged for HRP/ _H2O2 and _NaIO4 , air sparged for MT assay). After adding the oxidant, monitor the time-dependent UV/vis spectrum of the solution at a wavelength of 200–700 nm at a scan rate of 800 nm/min. For all samples, PBS buffer was first used as a blank, and then recorded with a Hitachi U-2010 UV/vis spectrophotometer at room temperature.

Molecular weight analysis

Molecular weight was determined using the following conditions: GPC-MALLS, on DAWN EOS (Wyatt Technology), using a Shodex-OH Pak column, in an aqueous mobile phase (50 mM PBS, 0.1M NaCl, 0.05% NaN ₃ ; pH=6.0), and using Optilab DSP (Wyatt Technology Refractive Index Detector. For molecular weight calculations, the experimentally determined dn/dc value of Compound IV (0.136) was used.

Example 44

Materials and methods

Tip modification

Silicon nitride (Si ₃ N ₄ ) tips were subjected to a 3-minute cleaning step using an O ₂ plasma instrument (the name of a machine) before surface modification, and then they were transferred into a piranha solution (sulfuric acid:H ₂ O ₂ =8:2) for 30 minutes. After washing with _H2O , they were transferred to 20% (v/v) 3-aminopropyltrimethoxysilane in toluene for 30-60 min to functionalize with amines. Two polyethylene glycol (PEG) derivatives were selected for PEGylation on the AFM tip: mPEG-N-hydroxysuccinimide (NHS) (Mw 2000) and Fmoc-PEG-NHS (Mw 3400) ( Nektar Inc.). A mixture of PEG (Fmoc-PEG-NHS:mPEG NHS=1 :5-10, 5 mM) was prepared in a mixture of 50 mM sodium phosphate buffer, 0.6 M _K2SO4 , pH _7.8 , and chloroform. The PEGylation reactions were performed sequentially as follows: first in sodium phosphate buffer at 40°C, then in chloroform, each step for 3 hours. The reason for using the PEG mixture is to prevent multiple DOPAs from binding on the _TiO2 . Fmoc-PEG-NHS provides an amine for Boc-DOPA conjugation after Fmoc cleavage. Deprotect Fmoc with piperidine (20% v/v in NMP) for 5 min, then transfer the cantilever to BOP/HOBt/DOPA (1:1:1 molar ratio, final 8 mM NMP solution) containing 10 μL DIPEA ) solution. The same procedure was also used for tyrosine modification.

AFM test

All data were collected from an AFM instrument (AsylumResearch, Santa Barbara, CA) placed on top of a Nikon inverted microscope. By applying the equipartition theorem to the thermal noise spectrum (S1), the spring constant of each cantilever (around 45, 100 and 300 pN/nm according to the information provided by the manufacturer) was calculated. Apply 1 drop of water to the pre-cleaned (sonication in organic solvent and using _O2 plasma) _TiO2 surface. A Force-distance curve containing PEG elasticity and profile length was selected for further statistical analysis. For DOPA quinone assays, all assays were performed in 20 mM Tris, pH 9.8.

Dynamic test

Loading rate-dependent force assays revealed the energy landscape of DOPA binding (17). The slope (=kBT/xb) of the linear plot of [Force Ratio over 1n(Rate of Lift)] determines the distance of the energy barrier xb along the axis of applied force. The combined energy barrier was calculated by taking the log intercept force at zero lift load rate and xb from the slope for the force transition that occurs from a change in traction rate. Silicon nitride AFM cantilevers (Bio-Levers, Olympus, Japan) were used due to their small spring constants (~5 pN/nm and ~28 pN/nm). The lowest lift-up rate of 2 nN/s in this study was obtained by using a pulling rate of 400 nm/s with a cantilever (~5 pN/mm). The highest rate of loading loading (1500 nN/s) was generated by operation of the piezoelectric device at 5 μm/s and using a stiff cantilever (300 pN/nm, Veeco). Surface Characterization Surfaces were analyzed by X-ray photoelectron spectroscopy (XPS) (Omicron, Taunusstein Germany) equipped with a monochromatic Al Kα (1486.8eV) 300W X-ray source and an electron gun for eliminating charge build-up. Silicon nitride surfaces (0.7 x 0.7 cm ² ) fabricated in a high temperature chamber (custom made by Keun Ho) were cleaned and modified using the same procedure as described in AFM tip modification. The photoelectric signal from _the carbon 1s orbital is the main indicator of the surface modification involving all species rich in Si, O and N on the _Si3N4 surface.

While the principles of this invention have been described with reference to specific embodiments, it should be clearly understood that these descriptions are included by way of example only, and are not intended to limit the scope of the invention in any way. For example, the present invention can enhance the adhesive properties of a wide variety of polymeric compositions, whether or not they are hydrogelable. Likewise, the present invention may use various other synthetic techniques well known to those skilled in the art to functionally modify specific polymer components for subsequent conjugation and preparation of corresponding DOPA conjugates. Other advantages, features, and benefits will be apparent from the claims appended hereto, along with the scope of their reasonable equivalents, as will be apparent to those skilled in the art.

Claims (25)

1. A dihydroxyphenyl (DHPD) sticky compound of formula (I),

In the formula, R ₁ and R ₂ may be the same or different, each independently selected from hydrogen, saturated and unsaturated, branched and linear, substituted and unsubstituted C _1-4 hydrocarbon groups; P is individually and independently selected from -NH ₂ , -COOH, -OH, -SH,

In the formula, R ₁ and R ₂ are as defined above, single bond, halogen,

where _A and _A are individually and independently selected from H, a single bond, protecting group, roughly poly(alkylene oxide),

where n is 1 to about 3, A ₃ is

R ₄ is H, C _1-6 lower alkyl, or polyalkylene oxide R ₃ is as defined above, and D is as shown in formula (I). 2. The compound of claim 1, wherein the poly(alkylene oxide) has the structure: In the formula, R ₃ and R ₄ are individually and independently selected from H or CH ₃ , the value of m is between 1 and about 250, A ₄ is NH ₂ , COOH, -OH, -SH, -H, DHPD or protecting group. 3. The compound of claim 1, wherein the DHPD has the following structure: 4. The compound of claim 1, wherein the DHPD has the following structure:

5. The compound of claim 1, wherein the DHPD has the following structure:

where _A2 is -OH, and _A1 is roughly poly(alkylene oxide) of the following structure

R ₃ , R ₄ and m are as defined in claim 2 . 6. The DHPD of claim 5, wherein the poly(alkylene oxide) is a block copolymer of ethylene oxide and propylene oxide. 7. A method of adhering a substrate to another substrate, the method comprising the steps of: providing a DHPD of the following structure,

Wherein R ₁ and R ₂ are as defined above; applying a DHPD of the above structure to one or the other substrate or both substrates to be adhered; contacting the substrates to be adhered with the DHPD of the above structure located therebetween to adhere the substrates to each other; and optionally translocating the substrates relative to each other by separating the substrates and bringing the substrates into contact with each other again with the DHPD of the above structure located between the substrates. 8. The method of claim 7, wherein _R1 and _R2 are hydrogen. 9. The method of claim 7, wherein one or the other of the DHPD moieties has the following structure:

10. The method of claim 7, wherein both DHPDs have the structure:

11. A binder containing a compound shown in formula (II):

In the formula, R ₁ , R ₂ and P are as defined in claim 1. 12. The adhesive of claim 11, wherein the adhesive bonds in an aqueous environment. 13. The adhesive of claim 11, wherein the compound of formula (II) is DOPA. 14. A composition containing a compound of formula (III):

Wherein, for each compound of formula (1a), R ₁ and R ₂ are individually and independently as defined in claim 1; For each compound of formula (1a), P _{and P} are individually and independently as defined for P in claim 1, and n and m are independently from 0 to about 5, provided that one of m or n is at least 1. 15. The composition of claim 14, wherein R _and R are _both hydrogen. 16. The composition of claim 14, wherein at least one of P ₁ or P ₂ contains approximately poly(alkylene oxide). 17. The composition of claim 14, wherein at least one of _P1 or _P2 contains at least one site of ethylenic unsaturation. 18. The composition of claim 14, wherein at least one of _P1 or _P2 comprises PEG. 19. The composition of claim 14, wherein at least one of (1a) is tris-DOPA and P is PEG. 20. The composition of claim 14, wherein _P1 and _P2 are coupled to each other. 21. A coated medical device comprising: device, and A coating comprising the composition of claim 14. 22. The coated medical device of claim 20, wherein said device is selected from the group consisting of a stent and a pacemaker. 23. The coated medical device of claim 14, wherein the coating is biodegradable. 24. A method of preventing adhesion of cells or proteins to a surgical incision site comprising the step of coating said site with the composition of claim 14. 25. A force intermediate in strength between a hydrogen bond and a covalent bond, said force forming, breaking and reforming substantially reversibly.

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