Patent Application
20180118916
The present invention relates to surface-modified, core-shell, scintillant-doped nanoparticles for sensitive detection of radioisotope activity, referred to herein as scintillant nanoparticles (SNPs). In one embodiment, the surface modification is a covalent attachment of binding ligands or a specific binding of radiolabeled target molecules or a lipid membrane coating.
Radionuclides, such as 3H, 14C, 33P, and 35S, are commonly used as bioanalytical labels and tracers in a wide range of biological, chemical and environmental assays due to the prevalence of H, C, P and S. However, these radioisotopes are challenging to quantify due to their low decay energies (Emax≤300 keV) and short β-particle penetration depths (≤0.6 mm) in aqueous media. Unlike fluorescent or fluorogenic probes and fluorescent protein tags, radionuclides do not significantly increase the size or mass of the labeled component, and therefore have minimal effects on binding, conformational changes, diffusion and active transport, etc. In addition, lower background signals are typically obtained for radioassays compared to fluorescence assays, where background signal arises from the inherent fluorescence of many biomolecules within the sample. Radiolabeling techniques have been used to quantify antigens at sub-μM concentrations, drugs such as buprenorphine at sub-nM concentrations, and enzyme activity.
β-particle emission from radiolabeled analytes is commonly quantified using liquid scintillation analysis (LSA). Scintillation occurs when the energy of a particle (α, β or photon) emitted during radioactive decay is absorbed by absorbing molecules within range, promoting them to an electronically excited state. Relaxation to the ground state results in emission of a photon or the transfer of energy to another molecule, which may then emit a photon. For example, the energy from radioactive decay of radioisotopes is absorbed by aromatic compounds and converted to detectable light, and is ultimately converted to current by a photomultiplier tube (PMT) detector. The energy from radioactive decay is measured in Ci, which is equal to 3.7×1010 disintegrations per second or 3.7×1010 Bq.
Typically, aqueous samples are dispersed in scintillation cocktails (LSCs), which are mixtures of aromatic absorbing hydrocarbon molecules (e.g., benzene, toluene, xylene, diisopropylnaphthalene, alkylbezenes), dispersing agents such as surfactants that accommodate the aqueous samples utilized in biological assays, and scintillant fluorophores (molecules that emit photons when excited by energy emitted during radioactive decay, e.g., 2,5-diphenyloxazole or 1,4-Bis(4-methyl-5-phenyl-2-oxazolyl)benzene). An approximation of the efficiency of liquid scintillation cocktails is about 10 photons released per 1 keV of emitted beta particles.
One disadvantage of LSC is that it is limited to uses for measurement of ex vivo bulk biological samples due to the cytotoxicity and hydrophobicity of its components. Experiments utilizing LSCs can result in liter or greater volumes of radioactive mixed waste that must be collected and disposed according to state and federal regulations. The toxicity and volatility of the primary solvent components of many LSC formulations complicate their transport, storage, and disposal. When disposing of the waste generated from LSCs, the waste is often classified as mixed waste containing a large quantity of organic solvent, markedly adding to the cost and complexity of disposal. In addition, surfactants can have a negative impact on non-mammalian aquatic organism populations, and may also help spread other pollutants throughout water systems.
Scintillation counting is not limited to samples in LSC, but can also be used for solid scintillators held within a liquid scintillation vial. These solid scintillators can be used instead of liquid scintillation cocktail to reduce toxic effects. Solid scintillators, like the organic scintillant fluorophores described above, absorb energy and emit photons, but are polymers (e.g., polyvinyltoluene or polystyrene) or inorganic crystals (e.g., CdWO4, CeF3, yttrium silicate (YSi), or yttrium oxide (YOx)). Inorganic crystalline scintillators have several advantages over polymer-based scintillators, as their high density and high atomic numbers means they can efficiently absorb energy, leading to more sensitive assays. Inorganic scintillants can also absorb higher energy radiation (gamma and X) than polymer-particle based scintillants. However, the density of inorganic crystalline scintillators causes them to settle in storage and sample containers, making it difficult to accurately dispense equal numbers of particles per volume of solution. In addition, samples must be continuously agitated in order to maintain the particles in suspension throughout the time period of the assay.
Although they usually exhibit lower quantum efficiencies than inorganic crystalline scintillators, polymer-based scintillators are less dense and therefore more easily dispensed and dispersed. The molecular structure of polymers limits their use to lower energy radiation such as 3H β-particle emission and Auger electrons emitted by 125I, which are considered safer due to their reduced penetration distances in air, water, and tissue. Further, polymer scintillators can incorporate scintillants to which the energy absorbed by the polymer can be transferred, resulting in photon emission at characteristic wavelengths. However, a drawback to the solid scintillant materials is that the majority of the scintillant is unused due to the low penetration depth for many β-particles. The low penetration depth proves particularly problematic for 3H β-particles, which are difficult to detect prior to energy dissipation.
To overcome this limitation, scintillation proximity assays (SPAs) particles are utilized for detection of low energy β-particles from radiolabeled analytes via conjugation of specific binding elements (i.e. radiolabeled analytes) to a particle surface of a solid scintillator, which increases the probability of energy absorption by the scintillant. When the β-particle of the bound analyte is emitted close to the surface (within the penetration distance), the efficiency of detection is markedly increased. SPA is particularly useful for β-particles with low penetration depths (approximately 0.5 μm for β particles emitted from 3H in water) and results in an increased number of emitted photons upon analyte binding. SPA also eliminates the need to separate bound from unbound analytes in radioimmunoassays, markedly enhancing the throughput and simplicity of the assay. Consequently, SPA lends itself to the monitoring of binding kinetics under steady state conditions, as well as to quantification of radiolabeled analytes using automation and high-throughput screening methods.
Modern SPA particle formats utilize scintillant-doped polyvinyltoluene (PVT) or polystyrene (PS) particles, or YSi or YOx particles, to which receptors have been covalently attached. SPAs have been successfully used to measure binding and/or binding kinetics of enkephalins, thyroxin, morphine, inositol phosphates, and many other analytes. However, current SPA platforms are in the micron size regime and thus the majority of scintillant encapsulated within the particle is not utilized due to the sub-micron penetration depths. Further, the lower solubility of polymer microspheres coupled with the more difficult surface modification chemistries complicate utilization and increase the price of the particles. Finally, the large size (5-10 μm diameters) and high density of SPA particles, relative to cells, prevents utilization for intracellular radioisotope detection.
Currently available SPA assays have no mechanism for integration of membrane proteins, many of which are implicated in variety of diseases and are critical to the development of drug treatments, but are unstable and inactive if removed from a cell membrane-like environment. Lipids have been previously used to coat silica particles and lipid membranes can be stabilized by incorporating polymerizable groups into the lipid structure itself, by crosslinking hydrophobic monomers within the inner leaflet of the membrane, or by combining a polymerizable lipid with a hydrophobic crosslinker. Stabilization of the membrane could reduce coating degradation during storage or use under adverse conditions (centrifugation, surfactant, sonication, drying etc). Hence, there exists a need for scintillant materials that are non-toxic and easily modifiable when used in used measuring low-energy radionuclide activity.
Any feature or combination of features described herein are included within the scope of the present invention provided that the features included in any such combination are not mutually inconsistent as will be apparent from the context, this specification, and the knowledge of one of ordinary skill in the art. Additional advantages and aspects of the present invention are apparent in the following detailed description and claims.
It is an objective of the present invention to provide nanosensors for dynamic intracellular imaging based on scintillation proximity assays (nanoSPA). The nanoSPA particles and their geometry possess unique and inventive technical features that can overcome several key limitations in radioisotope detection via commercially available LSC, SPA and solid scintillants. These advantages over existing scintillation technologies include, but are not limited to, the following:
1) enhanced compatibility with aqueous samples;
2) the ability to recover from solution following the assay to reduce the waste disposal volume and composition;
3) improved detection efficiency of low-energy β-particle emission using nanoSPA particles having a high surface area to volume ratio;
4) ease of production and modification;
5) an easily modified surface for attachment of biomolecules and other chemical species;
6) a smaller size and high signal output facilitate intracellular detection, which is an unprecedented assay;
7) compatibility with real-time, intracellular radioisotope detection; and
8) use of membrane coatings to reduce non-specific adsorption of protein and other molecules, and to maintain the stability and functionality of membrane proteins/membrane associated binding elements.
Without wishing to limit the invention to a particular theory or mechanism, the highly innovative nanoSPA approach described herein incorporates all key elements for SPA into a small, self-contained nanostructure for real-time studies.
None of the presently known prior references or work has the unique inventive technical features of the present invention. Although SPA beads made of polymers and inorganic crystals are commercially available, they are several μm in diameter (nanoSPA can be <1 μm), which can prohibit intracellular application, and have no mechanism for incorporation of membrane proteins or membrane-specific elements (such as gangliosides like GM1). Other techniques involve the use of solubilized cell membranes, which introduces detergents to the SPA sample.
According to one embodiment, the present invention utilizes polystyrene as the core material due to its low cost and the ease with which scintillant fluorophores, which tend to be hydrophobic, can be entrapped therein. The polystyrene matrix facilitates energy absorption and transfer from the radioisotope to the scintillant dye and provides a hydrophobic matrix for maximal scintillation efficiency. A thin (ca. 10-50 nm) silica shell is deposited onto the outside of the polystyrene core of variable diameter (ca. 100-1000 nm). The addition of silica shells to the radioisotope responsive scintillant-doped polystyrene particles increases solubility in aqueous samples and makes the particles more hydrophilic without using polyhydroxy films or surfactants, which are often required for the dispersion of PVT and PS particles in aqueous samples, thus reducing the aggregation commonly seen with polystyrene nanoparticles. Further still, the silica shell provides an easily modified surface for the covalent attachment of binding ligands or specific binding of radiolabeled target molecules.
According to another embodiment, the present invention features a multifunctional nanosensor platform for dynamic quantification of radioisotope-labeled, membrane protein/membrane associated ligands. The nanoSPA particle may comprise the polystyrene core of variable diameter into which radioisotope-responsive scintillant fluorophores are doped, and on which a silica shell is deposited onto the outside of the polystyrene core. Again, addition of the silica shell increases solubility in aqueous samples, and provides an easily modified surface. The silica shell is also essential for the deposition of a lipid membrane coating on the nanoparticle surface, which not only significantly reduces the non-specific adsorption of proteins and other biomolecules, but is also necessary for the incorporation of membrane proteins or other membrane-associated binding molecules.
Due to their high surface area to volume ratio high surface area to volume ratio, the nanoparticles can have greater dynamic ranges and higher efficiency than microwell plate-based designs as well as commercially available scintillant particles, which are often 5 to 10 microns in diameter. Furthermore, the density of the core-shell particles will allow for the particles to remain dispersed in a sample for a longer period of time than YSi particles, while still providing for easy recovery by centrifugation, thereby simplifying waste disposal and facilitating possible reuse.
AAPH or AIBA, 2,2′-azobis-2-methyl-propanimidamide dihydrochloride;
APTES or APTS, 3-aminopropyltriethoxysilane;
biotin-NHS, biotin-N-hydroxysuccinimide;
Bq, Becquerel (SI unit of radioactivity);
CE-UV/FS, Capillary electrophoresis-absorbance or fluorescence detector;
Ci, Curie (non-SI unit of radioactivity);
ConA, concanavalin A;
Cys, Cysteine; CySS, Cystine;
DBS, 2,4-dinitrobenzenesulfonyl;
DLS, Dynamic light scattering;
DMPOPOP or dimethyl POPOP, 1,4-Bis(4-methyl-5-phenyl-2-oxazolyl)benzene;
DOPC, 1,2-dioleoyl-sn-glycero-3-phosphocholine
DTDP, 4,4′-dithiodipyridine;
DTNB, 5,5′-dithiobis-(2-nitrobenzoic acid), also known as Ellman's reagent;
DTT, Dithiothreitol;
Emax, Maximum energy of beta particle;
EDAC, N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride
FRET, Förster Resonance Energy Transfer;
hCys, Homocysteine;
HPLC-EC, High-performance liquid chromatography with electrochemical detector;
HPLC-MS, High-performance liquid chromatography with mass spectrometry detector;
HPLC-UV/FS, High-performance liquid chromatography-absorbance or fluorescence detector;
LSA, Liquid scintillation analysis;
LSC, Liquid scintillation cocktail;
MES, 2-(N-morpholino)ethanesulfonic acid hydrate
MPTS, 3-mercaptopropyltrimethoxysilane;
NEM, N-ethylmaleimide;
NHS, N-hydroxysuccinimide;
NP, Nanoparticle;
NPM, N-(1-pyrenyl)maleimide;
PMT, Photomultiplier tube;
PS, Polystyrene;
PS-APTS, Polystyrene NPs with 3-aminopropyl triethoxysilane coating;
PS-MPTS, Polystyrene NPs with 3-mercaptopropyl trimethoxysilane coating;
pTP, Para-terphenyl;
PVT, Polyvinyltoluene;
RI, Radioisotope;
SAmax, Maximum specific activity of radioisotope;
SBD-F, 7-fluorobenzo-2-oxa-1,3-diazole-4-sulfonate;
SBH, Sodium borohydride;
SDS, Sodium dodecylsulfate;
SHE, Standard hydrogen electrode;
SNP, Scintillating nanoparticles;
SPA, Scintillation proximity assay;
SSA, Solid scintillation analysis;
TCEP, Tris(2-carboxyethyl)phosphine;
TEM, Transmission electron microcopy;
TEOS, Tetraethoxysilane;
THF, Tetrahydrofuran.
This patent application contains at least one drawing executed in color. Copies of this patent or patent application publication with color drawing(s) will be provided by the Office upon request and payment of the necessary fee.
The features and advantages of the present invention will become apparent from a consideration of the following detailed description presented in connection with the accompanying drawings in which:
The term “nano” when referring to the average particle size or diameter refers, for example, to average particle sizes of from about 1 nm to about 1000 nm, as understood by one ordinarily skilled in the art. Likewise, the term “micron” when referring to the average particle size refers, for example, to average particle sizes of from about 1 μm to about 500 μm.
Referring now to
In some embodiments, the polymer matrix core may be comprised of polystyrene or related aromatic polymers. In preferred embodiments, the scintillation nanoparticle is free of surfactants.
In other embodiments, the scintillator may comprise at least two scintillant fluorophores. The scintillant fluorophores may each absorb and/or emit light at different wavelengths. Without wishing to limit the invention to a particular theory or mechanism, the scintillant fluorophores primarily function to shift the emitted light into a wavelength range that is more easily detectable. For instance, the first fluorophore may emit light at a wavelength of about 300 nm. In another embodiment, the second fluorophore emits light at a wavelength of about 450 nm. In other embodiments, the scintillant fluorophores may enable the scintillator to detect different compounds with the same radioisotopes simultaneously. In some embodiments, the scintillator may comprise para-terphenyl (pTP), 1,4-Bis(4-methyl-5-phenyl-2-oxazolyl)benzene (DMPOPOP), or a combination thereof. However, the scintillators are not limited to the aforementioned examples, and may be any suitable fluorophore. In further embodiments, the scintillator may comprise about 3 to 6 different types of scintillant fluorophores.
In preferred embodiments, a surface of the silica shell is modified with functional groups. For example, the surface of the silica shell may be modified with amine or thiol functional groups. In one embodiment, the amine functionality may be obtained from APTS. In another embodiment, the thiol functionality may be obtained from MPTS. However, the surface functionality may be any other suitable or equivalent moiety obtained from a source that would be known to one of ordinary skill in the art.
In some embodiments, the surface modifier may comprise surface-attached receptors bound to the functionalized silica shell. The surface-attached receptors may be covalently or non-covalently bound to the silica shell. For example, surface-attached receptors may be covalently linked to the amine or thiol functional groups. Examples of these surface-attached receptors, include, but are not limited to, proteins, nucleic acid aptamers, small molecules or DNA oligomers. Without wishing to limit the invention to a particular theory or mechanism, each particle composition is specific for an individual analyte based on the type of receptor immobilized onto the particle surface. If there is no surface functionalization, then there is no specificity for isotopes.
According to other embodiments, the surface modifier may comprise a lipid membrane substantially covering a surface of the core-shell particle. As used herein, the term substantially can mean covering at least 50 of the core-shell particle surface. For example, the lipid membrane may cover about 50%-75% or 75%-90% or 90% to 100% of the core-shell particle surface.
In one embodiment, the lipid membrane may comprise a lipid bilayer. In another embodiment, the lipid membrane may further comprise receptors embedded in the lipid bilayer. Without wishing to limit the invention to a particular theory or mechanism, each particle composition is specific for an individual analyte based on the type of receptor. If there is no surface functionalization, then there is no specificity for isotopes. Examples of said receptors include, but are not limited to, membrane protein receptors, growth factor receptors, G-protein coupled receptors, ion channels, lipid-derived receptors, glycoprotein receptors, glycolipids, phospholipids, or a combination thereof.
Without wishing to be bound by theory, the lipid membrane may reduce non-specific adsorption of receptors, and maintain stability and functionality of said receptors and other membrane associated binding elements.
In some embodiments, the lipid bilayer may comprise polymerizable lipid monomers and functionalized lipid monomers. The polymerizable lipid monomers may be sorbyl- or dienoyl-containing lipid monomers such as, for example, 1,2-bis(octadeca-2,4-dienoyl)-sn-glycero-3-phosphocholine or 1,2-bis[10-(2′,4′-hexadieoyloxy)decanoyI]-sn-glycero-2-phosphocholine. The functionalized lipid monomers may be amine-functionalized lipid monomers such as amino(polyethylene glycol) (NH2-PEG).
In other embodiments, the lipid bilayer may comprise non-polymerizable lipid monomers and polymerized, hydrophobic non-lipid monomers. The non-polymerizable lipid monomers may be cell membrane fragments, 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-diphytanoyl-sn-glycero-3-phosphocholine monomers, naturally occurring lipids, or synthetic lipids. In one embodiment, each polymerized, hydrophobic non-lipid monomer may comprise a methacrylate, a styrene, or a combination thereof, and a cross-linking agent. In another embodiment, the methacrylate may be an aliphatic methacrylate, or an aromatic methacrylate such as benzyl methacrylate or naphthyl methacrylate. In yet another embodiment, the cross-linking agent may be a dimethacrylate such as, for example, ethylene glycol dimethacrylate.
In still other embodiments, the lipid bilayer may be a non-polymerizable naturally-occurring lipid bilayer, a synthetic lipid bilayer, or a combination thereof.
In some embodiments the diameter of the polymer core can range from about 50-1000 nm. For example, the diameter of the polymer core may be about 100-300 nm or about 300 nm to 500 nm or about 500 nm to 1000 nm. In other embodiments, the silica shell may have a thickness ranging from about 10-500 nm, such as about 10-50 nm or about 50-100 nm or about 100-300 nm or about 300-500 nm.
Another embodiment of the present invention features a method of detecting radioisotope activity in a sample. The method may comprise providing a scintillant material comprising a plurality of scintillation nanoparticles; combining the scintillation material and the sample in a medium such that radioactive decay of the radioisotopes in the sample generate energetic particles that interact with the scintillation material resulting in the emission of photons; and counting the photon emissions generated by the radioactive decay of the radioisotopes in the sample. Without wishing to limit the present invention to a particular theory or mechanism, the scintillant material is effective for detecting different compounds with the same radioisotopes simultaneously. In preferred embodiments, the scintillation nanoparticles may be any scintillation nanoparticle described herein.
In some embodiments, the energetic particles can be β-particles. In other embodiments, the medium is an aqueous solution. In yet other embodiments, the medium may comprise biological cells. Without wishing to limit the present invention to a particular theory or mechanism, the scintillation material may function as a cellular or intracellular imaging probe.
According to another embodiment, the present invention features a surfactant-free method of producing scintillation nanoparticles for detection of radioisotope activity. In one embodiment, the method may comprise adding monomers to an aqueous solution, polymerizing the monomers to form polymer core nanoparticles in solution, dissolving scintillators in an organic solvent, adding the scintillators in the organic solvent to the polymer core nanoparticle solution, agitating the mixture of the scintillators in the organic solvent and the polymer core nanoparticle solution, thereby doping the polymer core nanoparticles with the scintillators to form scintillant-doped polymer nanoparticles, removing the organic solvent from the mixture, thereby forming a concentrated solution of scintillant-doped polymer nanoparticles, redispersing the concentrated solution of scintillant-doped polymer nanoparticles in a second solvent having a base, and mixing silica precursors into the scintillant-doped polymer nanoparticles dispersed in the second solvent. Without wishing to limit the present invention to a particular theory or mechanism, the silica precursors can form a functionalized silica shell that encapsulates each scintillant-doped polymer nanoparticle, thereby forming the scintillation nanoparticles.
In preferred embodiments, the mixture may be agitated by sonication. However, other techniques of agitation as known to one of ordinary skill in the art may be utilized.
In a preferred embodiment, the step of removing the organic solvent from the mixture may comprise evaporating a portion of the organic solvent, agitating (i.e. by sonication) the remaining mixture, and repeating said steps for a number of iterations. The number of iterations can range from about 5 to 15. Without wishing to limit the present invention to a particular theory or mechanism, this method of removing the organic solvent advantageously provides for improved loading by increasing the contact of the scintillators with the polymer core nanoparticles as the organic solvent is gradually removed.
In one embodiment, the monomers may be styrene monomers that polymerize to form polystyrene core nanoparticles. In another embodiment, the monomers may be any aromatic monomers that polymerize to form aromatic polymer core nanoparticles.
In some embodiments, the scintillator may comprise at least two scintillant fluorophores that can each absorb and/or emit light at different wavelengths. Without wishing to limit the invention to a particular theory or mechanism, the scintillant fluorophores can shift the emitted light into a wavelength range that is more easily detectable. In one embodiment, the first fluorophore may emit light at a wavelength of about 300 nm. In another embodiment, the second fluorophore emits light at a wavelength of about 450 nm. Further still, the scintillant fluorophores may enable the scintillator to detect different compounds with the same radioisotopes simultaneously. In yet another embodiment, the scintillator may comprise para-terphenyl (pTP), 1,4-Bis(4-methyl-5-phenyl-2-oxazolyl)benzene (DMPOPOP), or a combination thereof. However, the scintillators are not limited to the aforementioned examples, and can be any suitable fluorophore. In further embodiments, the scintillator may comprise about 3 to 6 different types of scintillant fluorophores.
In one embodiment, the second solvent may comprise a base that has a pH ranging from 8 to 12. A non-limiting example of said base is ammonium hydroxide. Without wishing to limit the present invention to a particular theory or mechanism, the base may be effective for tuning the thickness of the silica shell.
In some embodiments the diameter of the polymer core can range from about 50-1000 nm. For example, the diameter of the polymer core may be about 100-300 nm or about 300 nm to 500 nm or about 500 nm to 1000 nm. In other embodiments, the silica shell may have a thickness ranging from about 10-500 nm, such as about 10-50 nm or about 50-100 nm or about 100-300 nm or about 300-500 nm.
In one embodiment, the silica precursor may comprise tetraethylorthosilicate (TEOS). In another embodiment, the silica precursor may further comprise functionalized silanes. The amount of functionalized silanes may range from about 5% to 15% volume of the silica precursor. For instance, the silica precursor may comprise about 90% vol of TEOS and about 10% volume of a functionalized silane. In some embodiments, the functional silane may be an aminosilane such as APTS, or a thiol-functionalize silane such as MPTS. In preferred embodiments, a surface of the silica shell is modified with functional groups. Without wishing to limit the present invention to a particular theory or mechanism, the functionalized silanes are effective for modifying the outer surface of the silica shell with functional groups. In one embodiment, the amine functionality may be obtained from APTS. In another embodiment, the thiol functionality may be obtained from MPTS. However, the surface modification may be other suitable or equivalent moieties obtained from sources that would be known to one of ordinary skill in the art.
In some embodiments, the method may further comprise depositing a surface modifier on the surface of the functionalized silica shell. In one embodiment, the step of depositing a surface modifier on the surface of the functionalized silica shell may comprise attaching receptors to the outer surface of the functionalized silica shell. The receptors can be covalently or non-covalently bound to the surface. For example, these surface-attached receptors may be covalently linked to the functional groups of the silica shells, such as the amine or thiol functional groups. Without wishing to limit the invention to a particular theory or mechanism, the type of receptor makes the scintillation nanoparticle specific for an individual analyte. Examples of these surface-attached receptors, include, but are not limited to, proteins, nucleic acid aptamers, small molecules or DNA oligomers.
In another embodiment, the step of depositing a surface modifier on the surface of the functionalized silica shell may comprise depositing a lipid membrane on an outer surface of the scintillation nanoparticle such that the outer surface is substantially covered by the lipid membrane. For instance, the lipid membrane may be deposited on the outer surface of scintillation nanoparticle using vesicle fusion techniques. Vesicle fusion techniques are known to one of ordinary skill in the art. In some embodiments, the lipid membrane may cover about 50%-75% or 75%-90% or 90% to 100% of the outer surface.
In some embodiments, the lipid membrane may comprise a lipid bilayer. In other embodiments, the lipid membrane may further comprise receptors embedded in the lipid bilayer. Again, without wishing to limit the invention to a particular theory or mechanism, the type of receptor makes the scintillation nanoparticle specific for an individual analyte. Examples of said receptors include, but are not limited to, membrane protein receptors, growth factor receptors, G-protein coupled receptors, ion channels, lipid-derived receptors, glycoprotein receptors, glycolipids, phospholipids, or a combination thereof. Preferably, the lipid membrane may reduce non-specific adsorption of receptors, and maintain stability and functionality of said receptors and other membrane associated binding elements.
In one embodiment, the lipid bilayer may comprise polymerizable lipid monomers and functionalized lipid monomers. The polymerizable lipid monomers may be sorbyl- or dienoyl-containing lipid monomers such as, for example, 1,2-bis(octadeca-2,4-dienoyl)-sn-glycero-3-phosphocholine or 1,2-bis[10-(2′,4′-hexadieoyloxy)decanoyI]-sn-glycero-2-phosphocholine. The functionalized lipid monomers may be amine-functionalized lipid monomers such as amino(polyethylene glycol) (NH2-PEG).
In another embodiment, the lipid bilayer may comprise non-polymerizable lipid monomers and polymerized, hydrophobic non-lipid monomers. The non-polymerizable lipid monomers may be cell membrane fragments, 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-diphytanoyl-sn-glycero-3-phosphocholine monomers, naturally occurring lipids, or synthetic lipids. In one embodiment, each polymerized, hydrophobic non-lipid monomer may comprise a methacrylate, a styrene, or a combination thereof, and a cross-linking agent. In another embodiment, the methacrylate may be an aliphatic methacrylate, or an aromatic methacrylate such as benzyl methacrylate or naphthyl methacrylate. In yet another embodiment, the cross-linking agent may be a dimethacrylate such as, for example, ethylene glycol dimethacrylate.
In yet another embodiment, the lipid bilayer may be a non-polymerizable naturally-occurring lipid bilayer, a synthetic lipid bilayer, or a combination thereof.
Without wishing to limit the present invention to a particular theory or mechanism, the method of producing the scintillation nanoparticles described herein can advantageously provide for scintillation nanoparticles that are functionalized and have improved dye or fluorophore loading. Further still, the method provides for nanoparticles that suitably sized to function as cellular or intracellular imaging probes.
The following examples are presented for illustrative purposes only, and are not intended to limit the present invention in any way.
The following is a non-limiting example of producing polystyrene core-silica shell nanoparticles. Equivalents or substitutes are within the scope of the present invention.
Materials
The primary scintillant p-terphenyl (pTP) and secondary scintillant 1,4-bis(4-methyl-5-phenyl-2-oxazolyl)benzene (dimethyl-POPOP) were obtained from Acros Organics (Geel, Belgium). Styrene, biotin-N-hydroxysuccinimide (biotin-NHS), 2,2′-azobis (2-methylpropionamidine) dihydrochloride (AAPH), tetraethylorthosilicate (TEOS), and 3-aminopropyltriethoxysilane (APTS) were obtained from Sigma Aldrich (St. Louis, Mo.). 3H acetic acid (500 mCi/mmol) and 3H glutamic acid (49.6 Ci/mmol) were obtained from MP Biomedicals (Solon, Ohio) and Perkin Elmer (Boston, Mass.) respectively. Sodium dodecylsulfate (SDS) was obtained from Fisher (Pittsburgh, Pa.).
Fabrication of Scintillant-Doped Polystyrene Core nPs
Scintillant-doped polystyrene (PS) core nPs were fabricated via a surfactant-free emulsion polymerization process by combining 6.0 g (58 mmols) of styrene containing 540 mM (3.6 mmols) dimethyl POPOP and 54 mM pTP (360 mmols) for a pTP: dimethyl POPOP ratio of 100:1, 30 mM (0.2 mmols) dimethyl POPOP and 300 mM (2.0 mmols) pTP for a pTP: dimethyl POPOP ratio of 10:1, or 30 mM (2 mmols) dimethyl POPOP and no pTP, with 100 mL of degassed nanopure water in a 500 mL round-bottomed flask equipped with a magnetic stir bar. Polymerization was initiated by adding 6 mg of AAPH (220 mM final concentration). The water/styrene combination was stirred briskly under a gentle stream of argon and maintained at 70-80° C. in a water bath for 12 hours. Unpolymerized styrene and some water were removed under reduced pressure. The nPs were rinsed several times with water, isopropanol, and water again. The average diameter of the PS nPs was ca. 175 nm as determined by dynamic light scattering (DLS, Brookhaven Instruments, BI-200 with a BI-DS detector and BI-800AT autocorrelator software, non-negatively constrained least squares, multiple pass calculation). The concentration of nPs in the PS core stock suspension was estimated by freezing and then lyophilizing 4 mL of the suspension, and was found to be approximately 30 mg/mL. PS nPs without scintillants were made according to the same procedure by simply omitting dimethyl POPOP and pTP from the reaction flask.
Addition of Silica Shells to PS Core nPs
Silica shells were added to the PS core nPs by diluting 0.05 mL (1.5 mg) of the PS core stock suspension with 3.95 mL water and 20 mL isopropanol in a 50 mL round bottomed flask. The mixture was stirred briskly using a stir bar and 0.50 mL NH4OH (28%) was added. After several minutes, 100 mL of a TEOS:APTS combination (90% TEOS, 10% APTS by volume) was added dropwise to the flask. Stirring was continued for 1 hour, after which PS-core silica-shell nPs were collected by centrifugation. The nPs were washed by repeatedly dispersing them in ethanol, centrifuging the ethanol at 14,000 rpm, and removing the supernatant liquid. Ethanol rinsing cycles were followed by three water rinsing cycles. Shells synthesized with TEOS only (no APTS) were fabricated using a similar procedure in which the NH4OH volume was reduced to 0.25 mL. The average diameter of the PS-core silica-shell nPs was ca. 300 nm as determined by DLS.
Transmission Electron Microscopy
As shown in
Zeta Potential Measurements
Zeta potential measurements were made in disposable folded capillary cells using a Zetasizer Nano (Malvern Instruments Ltd., Worcestershire UK). PS, silica, and PS-core silica-shell nPs were dispersed in 100 mM NaCI at varying pH (from 3 to 10) immediately prior to measurement. Cells were flushed with water and then with 100 mM NaCI at the appropriate pH between samples. Zeta potential was calculated using the Smoluchowski approximation (1.5) as the solution for the Henry equation for all samples.
Comparison of Scintillation Efficiencies for PS Cores with and without pTP
The effect of including pTP along with dimethyl POPOP in PS core nPs was investigated by comparing the scintillation response of PS cores with a ratio of 10:1 pTP (300 mM):dimethyl POPOP (30 mM) to the response of PS cores fabricated with the same concentration of dimethyl POPOP but no pTP. Both the mass-dependent and the activity-dependent responses of the PS core nPs were tested with 3H acetic acid (labeled at the a carbon) using a liquid scintillation counter (Beckman LS 6500, Beckman Coulter, Brea, Calif.). In the mass-dependent response study, 1 mCi of 3H acetic acid was diluted to 10 mL with water. The nP mass was then increased from 0 to 13 mg (0 to 1 mg/mL) by adding volumes of nP stock suspensions to the vials and mixing the solution. The scintillation response in counts per minute was measured for each sample after each addition of nPs. In the activity-dependent study, a fixed mass (10 mg, 1 mg/mL final concentration) of nPs was suspended in 10 mL of water. The activity of the samples was then increased from 0 to 2000 nCi (0 to 190 nCi/mL) by sequential additions of 10 mCi/mL 3H acetic acid. Samples were mixed by aspirating the solution with a pipette and the scintillation response in counts per minute was measured after each addition.
Assay of PS Core and PS-Core Silica-Shell nPs with 3H Acetic Acid
The mass-dependent and activity-dependent scintillation responses of PS core (with dimethyl POPOP and pTP at a ratio of 100:1), PS (without dimethyl POPOP and pTP) and PS-core silica-shell (with dimethyl POPOP and pTP at a ratio of 100:1 in the PS core) were tested with 3H acetic acid. In both studies, all nP samples were dispersed in 1 mL 10 mM SDS solution. The scintillation vials for the Beckman LS 6500 liquid scintillation counter can hold approximately 20 mL, and when only 1 mL sample volumes are used the liquid sample only fills the bottom 1/20 of the vial. Such low volumes in 20 mL scintillation vials may decrease measurement efficiency, as photons emitted from the sample at a the distal edges may not be detected as well as photons emitted from the more central regions of the vial volume. This problem could be alleviated by using larger sample volumes; however, fabricating the amount of nPs needed to complete experiments using larger volumes is impractical. Instead, smaller volume (2 mL) glass autosampler vials were glued to the interiors of 20 mL scintillation vials in such a way as to ensure that 1 mL samples would be contained within a region close to the center of the scintillation vial volume.
In the mass-dependent response study, where mass refers to the mass of PS present rather than the combined mass of both PS and silica, 1 mL aliquots of 10 mM SDS solution were added to modified scintillation vials, along with 33 mL 10 mCi/mL 3H acetic acid (approximately 300 nCi final activity). The nP mass was then increased from 0 to 5 mg (0 to 4 mg/mL) by adding volumes of nP suspensions to the vials and mixing the solution. The scintillation response in counts per minute was measured for each sample after each addition of core-shell nPs.
In the activity-dependent study, a fixed mass (4 mg, based on the mass of PS present) of nPs was added to 1 mL 10 mM SDS solution in modified scintillation vials. The activity of the samples was increased from 0 to 1250 nCi (0 to 890 nCi/mL) by sequential additions of 10 mCi/mL 3H acetic acid. Samples were mixed by aspirating the solution with a pipette and the scintillation response in counts per minute was measured after each addition. Data for both the mass-dependent and activity-dependent studies were normalized to pTP fluorescence intensities at 345 nm (300 nm excitation) in order to account for differences in PS-core silica-shell NP versus PS core NP concentrations.
The response of a fixed mass of scintillant-doped polystyrene-core silica-shell particles to increasing activity of 3H acetic acid (
Further investigation into the significance of the interactions between the silica surfaces of core-shell NPs and 3H acetic acid was accomplished by comparing the scintillation efficiencies of PS-core silica-shell nPs (10:1 pTP:dimethyl POPOP) made with TEOS and APTS (10% APTS) and PS-core silica-shell NPs made with TEOS only. Equivalent amounts (based on the fluorescence intensity of entrapped pTP excited with 300 nm light) of PS-core TEOS and APTS silica-shell NPs and PS-core TEOS-only silica-shell NPs were diluted to 10 mL with water in unmodified plastic scintillation vials. 3H acetic acid was then incrementally added to each sample to increase the activity from 0 to 70 mCi (0 to 7 mCi/mL). Samples were mixed by aspirating the solution with a pipette and the scintillation response in counts per minute was measured after each addition.
Biotin-Streptavidin Binding Model SPA
Scintillant-doped PS-core silica-shell nPs (ca. 60 mg nPs) were suspended in 5 mL 20 mM pH 8.3 borate buffer with 0.6 mM (3.0 mmols) biotin-NHS for 2.5 hrs. After thorough rinsing and redispersion in water, nPs were treated with 570 nM streptavidin (2.8 nmols). The nP/streptavidin solution was shaken at 120 rpm for 2.5 hrs followed by further rinsing with water. Streptavidin functionalized scintillant-doped PS-core silica-shell nP were then diluted to 33 mL with water. Biotinylated 3H glutamic acid was prepared by diluting 15 mL (15 mCi) 49.6 Ci/mmol 3H glutamic acid with 15 mL 20 mM pH 8.3 borate buffer and 3 mL 20 mM biotin-NHS (60 nmols). 3H glutamic acid and biotin-NHS were allowed to react for 12 hours. A solution containing the same activity of 3H glutamic acid and same concentration of biotin was prepared as a control.
For SPA experiments, 1 mL aliquots of streptavidin-functionalized scintillant-doped PS-core silica-shell nP solution (1.9 mg, or ca. 1.0×1011 nPs each) were placed in modified scintillation vials with 100, 250, 500, 1000, or 3000 nCi biotinylated 3H glutamic acid or 3H glutamic acid and biotin mixture. Water was added to each sample to bring the final sample volume to 1.3 mL. Samples were mixed and allowed to react for 30 minutes before the scintillation response was measured.
The following is a non-limiting example of producing polystyrene-core silica-shell scintillant nanoparticles (nanoSCINT) for low-energy radionuclide quantification in aqueous media. Equivalents or substitutes are within the scope of the present invention.
Materials
Styrene, alumina, p-terphenyl (pTP), and 1,4-Bis(4-methyl-5-phenyl-2-oxazolyl)benzene (dimethyl-POPOP) were purchased from Acros Organics. Tetraethylorthosilicate (TEOS), and 2,2′-azobis(2-methylpropionamidine) dihydrochloride (AIBA), Triton X-100,cycloheaxane, sodium citrate (tribasic) hydrate, sodium tetraborate hydrate, and 2-(N-morpholino)ethanesulfonic acid hydrate (MES) were obtained from Sigma Aldrich. Sodium chloride, sodium phosphate hydrate (monobasic), isopropanol and ammonium hydroxide were obtained from EMD Millipore. Hexanol was purchased from Alfa Aesar. BioCount Liquid scintillation cocktail was acquired from Research Products International. 3H labelled acetic acid (as sodium acetate) was purchased from Perkin Elmer. All chemicals except styrene were used as received. Inhibitor was removed from styrene by passing the styrene through a 3 cm long alumina column immediately prior to use.
Preparation of NanoSCINT Particles
About 3 g of styrene without inhibitor was added to 100 mL degassed water in an argon-flushed 500 mL round-bottomed flask heated to 70° C. in an oil bath. Polymerization was initiated by adding 10 mg of AIBA dissolved in approximately 200 μL of water to the reaction flask. The water/styrene mixture was stirred rapidly throughout the polymerization process, which was allowed to continue for at least 6 hours. Excess styrene and some water were removed from the nanoparticle solution using a rotary evaporator. Polystyrene nanoparticles were doped with scintillant fluorophores by dissolving 53 mg (135mmoles) of dimethyl POPOP and 262 mg (1.14 mmoles) of pTP in 20 mL of a 10% isopropanol, 90% chloroform solvent mixture. The scintillant fluorophores in solvent were then added directly to the aqueous polystyrene nanoparticle solution in a 500 mL round-bottomed flask. The nanoparticle solution was sonicated in a bath sonicator for several minutes to disperse organic solvent droplets throughout the water. A small amount of the isopropanol and chloroform was then removed using a rotary evaporator. Not all solvent was removed; instead multiple sonication and evaporation cycles were used to allow solvent droplets contact the polystyrene as much as possible and remove the solvent slowly. Once the solvent was removed, the scintillant fluorophore doped polystyrene nanoparticle solution was stored at room temperature until use. A small volume of the solution was removed and lyophilized to determine the weight per volume of nanoparticles.
Silica shells were added to scintillant fluorophore doped polystyrene nanoparticles by dispersing 2 mL of the polystyrene nanoparticle stock solution (approximately 56 mg of nanoparticles) in 200 mL isopropanol with 38 mL water and 5 mL ammonium hydroxide. The dispersion was stirred briskly for several minutes while 1 mL TEOS was added dropwise. Stirring was continued for 1 hour before nanoSCINT particles were collected by centrifugation and rinsed several times with water.
For comparison in zeta potential measurements, silica nanoparticles without polystyrene cores were prepared by dispersing 1.8 mL Triton X-100 in a mixture of 8.0 mL cyclohexane and 1.8 mL hexanol in a 50 mL round bottom flask. The mixture was stirred briskly with a stir bar for several minutes before 550 pL water, 60 μL ammonium hydroxide and 100 μL TEOS were added. Stirring was continued for approximately 12 hours. Silica particles were collected by adding several mL of acetone to cause the particles to flocculate at the bottom of the flask. Particles were rinsed three times with 1.5 mL aliquots of ethanol and then another three times with 1.5 mL aliquots of water. The silica particles were found to be approximately 140 nm diameter by dynamic light scattering.
Transmission Electron Microscopy
Transmission electron micrographs of polystyrene and of nanoSCINT particles were obtained by drying small amount of nanoparticle solutions on carbon films on copper grids. Samples were observed using a Technai G2 Spirit transmission electron microscope (FEI).
Zeta Potential Measurements
Zeta potential measurements were made in disposable folded capillary cells using a Zetasizer Nano (Malvern Instruments Ltd., Worcestershire UK). Polystyrene, silica, and nanoSCINT particles were dispersed in 100 mM NaCl at varying pH (from 3 to 10) immediately prior to measurement. Cells were flushed with water and then with 100 mM NaCl at the appropriate pH between samples. Zeta potential was calculated using the Smoluchowski approximation as the solution for the Henry equation for all samples.
Scintillation Efficiency Measurements and NanoSCINT Particle Recovery
The scintillation efficiency of nanoSCINT was tested by combining fixed amounts of nanoSCINT particles in water with 3H-labeled acetic acid. 2 mL of nanoSCINT particle stock solution was added to three 7 mL polyethylene scintillation vials. 3H labeled acetic acid (150 mCi/mmol) was sequentially added to the nanoSCINT samples, as well as to three vials containing only 2 mL each of water and three vials containing 2 mL each of liquid scintillation cocktail. The scintillation efficiency, in counts per minute, was measured for each sample after every addition of 3H acetic acid with a Beckman LS 6000IC liquid scintillation counter. After the final addition of 3H labeled acetic acid, nanoSCINT particles were recovered from each sample by centrifugation. Each recovered nanoSCINT sample was rinsed once by re-suspending the particles in 5 mL fresh water and then collecting them by centrifugation at 16,000 g for 5 minutes. The particles were then suspended again in 2 mL of water and treated with increasing activities of 3H labeled acetic acid as before.
The possible effects of pH and increased salt concentration on nanoSCINT efficiency were explored by dispersing fixed amounts of nanoSCINT particles in 10 mM buffer (sodium citrate at pH 3.0, MES at pH 5.5, sodium phosphate at pH 7.0, and sodium tetraborate at pH 9.5) and measuring scintillation before and after the addition of 3H-acetic acid. Scintillation was measured again after the concentration of NaCl was increased to 100 mM by adding a volume of concentrated NaCl to the same samples.
Liquid scintillation cocktails typically employ the same general process of energy absorption followed by energy transfer and photon emission, although the chemical formulations, which are proprietary, may vary depending on application. The primary component of cocktails is an energy absorbing solvent such as toluene or xylene, or the less flammable and less toxic diisopropylnaphthalene, phenylxylylethane, or dodecylbenzene. Additives such as surfactants are used to facilitate the dispersion of aqueous samples into the organic solvent. Scintillant fluorophores, to which energy absorbed by the solvent is transferred, are included to shift photon emission to wavelengths more readily detected by photomultipliers and CCDs. In the case of solid polymer scintillation particles, the polymer (e.g. polystyrene or polyvinyltoluene) replaces the organic solvent base as the primary absorber of energy from β-particle emission. Although the π-orbital electrons of polystyrene itself are readily excited by β-particle emission, the radiative quantum yield of polystyrene is low (ca. 7%). As with liquid scintillation cocktails, scintillant fluorophores (pTP and dimethyl POPOP) were used to both improve total radiative quantum yield of polystyrene core nanoparticles, and redshift the emission wavelength.
Scintillant-doped polystyrene core nanoparticles were fabricated via surfactant-free emulsion polymerization using the cationic initiator 2,2′-azobis(2-methylpropionamidine) dihydrochloride (AIBA), as illustrated in
Plots showing the response of nanoSCINT particles and liquid scintillation cocktail in counts per minute as measured by a liquid scintillation counter versus activity can be seen in
The proximity of individual nanoSCINT particles to 3H labeled molecules affects scintillation response and is also important to consider when comparing nanoSCINT to liquid scintillation cocktail. In 1 mL of a solution containing 4×1012 nanoSCINT particles, the particles will occupy less than 1% of the sample volume, necessitating the use of either high nanoSCINT concentrations or high radionuclide activities to achieve the same scintillation response as LSC. In contrast, aqueous sample droplets dispersed in LSC will be completely surrounded by the cocktail, which is a more efficient geometry for absorbing energy from isotropic emission. This concept is depicted in the illustration shown in
The surface charge characteristics of nanoSCINT were examined by comparing the zeta potentials of nanoSCINT particles to the zeta potentials of silica and polystyrene core nanoparticles (
The importance of measuring zeta potential of nanoSCINT particles at varying pH and low and high salt concentrations lies in the stability of an aqueous nanoSCINT dispersion. Although silica is already considered lyophilic, electrostatic repulsions between particles (which are directly related to the fraction of deprotonated silanols and the ions present at the particle surface) at least partially dictate whether particles resist flocculation/aggregation or not. The zeta potential plot shown in
The following is a non-limiting example of a polystyrene-core silica-shell nanoparticle-based SPA platform (nanoSPA). Equivalents or substitutes are within the scope of the present invention.
In a preferred embodiment, a core-shell nanoparticle based scintillation proximity assay platform for the detection of 3H labeled analytes features scintillant fluorophores incorporated into polystyrene core particles surrounded by functionalized silica shells. The functional groups of the silica shells then allow for the covalent attachment of specific binding moieties such as proteins, small molecules, or DNA. The utility of the SPA platform has been demonstrated in two model assays, one in which biotin-functionalized nanoSPA particles are used to measure 3H-labeled Neutravidin, and another in which DNA-oligomer-functionalized nanoSPA particles are used to detect a 3H-labeled complementary strand via hybridization. In both models, nanomole and sub-nanomole amounts of the targets were detected. The nanoSPA platform not only facilitates measurement of 3H-labeled analytes in bulk aqueous solutions, but due to the small diameter of the particles and the protection of the hydrophobic polymer scintillant core by an easily modified silica shell, nanoSPA particles may be used as cellular or intracellular imaging probes.
As previously described, the polystyrene cores of nanoSPA particles were prepared in a surfactant-free emulsion polymerization, into which scintillant fluorophores can be incorporated either before polymerization (dissolved in the monomer, styrene) or after polymerization by means of swelling the particles in solvent. pTP was chosen as a primary scintillant fluorophore, and dimethyl POPOP as a wavelength-shifting secondary scintillant fluorophore. Amine or thiol functionalized silica shells were then added to the cores by combining 10% of functional siloxanes (APTS or MPTS) with tetraethylorthosilicate during shell synthesis. As shown in
The presence of amine or thiol groups on the silica shells allowed for covalently attaching binding elements to the nanoSPA particle surfaces. Biotin-functionalized nanoSPA particles were used to measure 3H-labeled Neutravidin, and DNA-oligomer-functionalized nanoSPA particles were used to detect a 3H-labeled complementary strand via hybridization.
The results of the DNA oligomer hybridization nanoSPA model assay are illustrated in
Both the Neutravidin and the complementary DNA oligomer were labeled using the well-studied N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDAC) and N-hydroxysuccinimide (NHS) coupling reaction, demonstrating that because this method of 3H labeling can be done in-house, nanoSPA could be used even for analytes without 3H labeled versions available for commercial purchase.
NanoSPA particles avoid many of the problems associated with traditional liquid scintillation counting techniques as the scintillator is solid and protected by a hydrophilic silica shell. The easy attachment of recognition elements to the functionalized silica surfaces of the particles makes them versatile for separation-free SPA applications in aqueous samples. In addition, because nanoSPA particles are also small in diameter (<300 nm) and are composed of biologically inert materials, they may potentially serve as sensitive extra- or even intracellular imaging probes.
The following is a non-limiting example of a lipid membrane coated polystyrene-core silica-shell nanoSPA particle. Equivalents or substitutes are within the scope of the present invention.
Materials
Styrene, alumina, p-terphenyl (pTP), and 1,4-Bis(4-methyl-5-phenyl-2-oxazolyl)benzene (dimethyl-POPOP) were purchased from Acros Organics. Tetraethylorthosilicate (TEOS), (3-aminopropyl)triethoxysilane (APTS), (3-mercaptopropyl)triethoxysilane (MPTS), and 2,2′-azobis(2-methylpropionamidine) dihydrochloride (AIBA) were obtained from Sigma Aldrich. Ammonium hydroxide and tetrahydrofuran (THF) were obtained from EMD Millipore. 3H-labeled acetic acid (as sodium acetate) was purchased from Perkin Elmer. Lipids were purchased from Avanti Polar Lipids. All chemicals except styrene were used as received. Inhibitor was removed from styrene by passing the styrene through an alumina column immediately prior to use.
Preparation of Lipid Membrane Coated NanoSPA Particles
3 g of styrene without inhibitor was added to 100 mL degassed water in an argon-flushed 500 mL round-bottomed flask heated to 70° C. in an oil bath. Polymerization was initiated by adding 10 mg of AIBA dissolved in approximately 200μL of water to the reaction flask. The water/styrene mixture was stirred rapidly throughout the polymerization process, which was allowed to continue for at least 6 hours. Excess styrene and some water were removed from the nanoparticle solution using a rotary evaporator. Polystyrene nanoparticles were doped with scintillant fluorophores by dissolving 53 mg (135 mmoles) of dimethyl POPOP and 262 mg (1.14 mmoles) of pTP in 20 mL of a 10% isopropanol 90% chloroform solvent mixture, or in THF. The scintillant fluorophores in solvent were then added directly to the aqueous polystyrene nanoparticle solution in a 500 mL round-bottomed flask. The nanoparticle solution was sonicated in a bath sonicator for several minutes to disperse organic solvent droplets throughout the water. A small amount of the isopropanol and chloroform was then removed using a rotary evaporator. Not all solvent was removed; instead multiple sonication and evaporation cycles were used to allow solvent droplets contact the polystyrene as much as possible and remove the solvent slowly. Once the solvent was removed, the scintillant fluorophore doped polystyrene nanoparticle solution was stored at room temperature until use. A small volume of the solution was removed and lyophilized to determine the weight per volume of nanoparticles.
Silica shells were added to scintillant fluorophore doped polystyrene nanoparticles by dispersing 2 mL of the polystyrene nanoparticle stock solution (approximately 56 mg of nanoparticles) in 200 mL isopropanol with 38 mL water and 5.0 mL or 7.5 mL (depending on the silica precursors used) ammonium hydroxide. The dispersion was stirred briskly for several minutes while, depending on the desired surface functionality, 2 mL TEOS, TEOS with APTS, or TEOS with MPTS was added dropwise. Stirring was continued for 1 hour before nanoSPA particles were collected by centrifugation and rinsed several times with water.
Lipid membranes were deposited on the silica shells of nanoSPA particles by vesicle fusion techniques. To prepare the vesicles, 3 mg of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) in chloroform was aliquoted into a vial. The chloroform was removed by evaporation under vacuum overnight. The dried lipid was re-suspended in 1.5 mL nanopure water (18.3 MO) to yield a solution with lipid concentration of 2 mg/mL. The lipid solution was sonicated for 10 minutes with 10 minute rest intervals until it became transparent. NanoSPA particles (0.9 mg) were combined with 450 μL of freshly prepared vesicle solution, then sonicated for 5 minutes and stirred overnight to allow lipid vesicles to fuse onto the silica surfaces of the particles. Lipid membrane coated nanoSPA particles were centrifuged (3000 rpm×15 minutes) and re-suspended in buffer (0.1 M sodium phosphate, 0.15 M sodium chloride, pH=7.4) to yield a solution with 0.3 mg/mL particle concentration.
The reduction in non-specific adsorption of protein for lipid membrane coated nanoSPA was evaluated by incubating particles with 3H-labeled bovine serum albumin and then measuring the scintillation response of the particles (
The utility of lipid membrane coated nanoSPA has been demonstrated by incorporating the ganglioside GM1 (which has two alkyl chain tails that insert into the leaflet of a lipid bilayer as it is naturally found in animal cell membranes) into the nanoSPA lipid membrane coating for a binding assay with 3H-labeled cholera toxin B, a known GM1 ligand (
The following is a non-limiting example of thiol-responsive scintillation proximity assay core-shell nanoparticles as turn-on biosensors. Equivalents or substitutes are within the scope of the present invention.
Small thiols, such as cysteine (Cys), participate in metabolism, regulation of enzymes, cellular antioxidant defense, signal transduction, protein folding, and coordination of metals. Cys is a regulating factor of thiol-disulfide exchange. It plays a key role in biological activity of S100 proteins and regulates glutathione (GSH) synthesis. Intracellular thiols and disulfides exist primarily in the form of Cys and its dimer cystine (CySS). Cys is the precursor of GSH, concanavalin A(ConA), and taurine. Other biothiols include homocysteine (hCys), small peptides such as cysteinyl-glycine and GSH, and proteins with Cys residues. The free thiol group of Cys makes it one of the most reactive amino acids. Elevated levels of this non-essential amino acid can lead to cardiovascular diseases, neurotoxicity and hypoglycemic brain damage, whereas its deficiency is related to many health problems such as liver damage, muscle and fat loss, and lethargy. Elevated level of Cys is observed in patients with diseases such as Alzheimer's, Parkinson's, cystinuria, and cystinosis and decreased concentration of Cys and CySS is reported in HIV-infected patients. Cys undergoes a reversible redox reaction that can impact protein structure, activity, and interactions with DNA and other proteins. Cys can also reduce metals, such as iron, which can subsequently initiate the redox cycle of iron in the presence of H2O2 and lead to DNA damage.
Disulfides are important members of intracellular media and they play key roles in protein folding and function. Disulfide bond formation is mediated by auto-oxidation of thiols, and occurs in cells in specific organelles such as endoplasmic reticulum. Thiol-disulfide exchange is a dynamic process in cells. Thiolate anions, which are the reactive form of thiol, approach disulfide bonds to initiate thiol-disulfide exchange by an SN2 displacement mechanism. The thiol-disulfide ratio is important in maintaining cellular homeostasis and antioxidant defense; otherwise, redox imbalance and oxidative stress will affect the structure and activity of many proteins and cell signaling, which will lead to a variety of diseases such as Alzheimer's, chronic lung disease, atherosclerosis, and age-related macular degeneration.
In order to measure disulfides, thiols that are already present must be protected by blocking agents. N-ethylmaleimide (NEM) is a thiol blocking agent that is commonly used to alkylate thiols through Michael addition, as known to one of ordinary skill in the art. Then, a reducing reagent, such as sodium borohydride (SBH), can be used to cleave the disulfides to thiols for measurement. SBH is a reducing agent that can be used to convert the disulfides to thiols without affecting the NEM-blocked thiols. It also deactivates any excess NEM to prevent the further alkylation of thiols that are newly reduced from the disulfides.
Current optical methods for thiol measurement (HPLC-UV/FS, CE-UV/FS, etc.) are based on derivatization and separation. Fluorimetry detection has been widely used as a sensitive and versatile probe for biothiols. UV-VIS spectrophotometry-based sensitive and selective probes of derivatized biothiols have also been reported. Thiol-specific labels (e.g. 4,4′-dithiodipyridine (DTDP), 7-fluorobenzo-2-oxa-1,3-diazole-4-sulfonate (SBD-F), N-(1-pyrenyl)maleimide (NPM)), 2,4-dinitrobenzenesulfonyl (DBS), and Ellman's reagent (5,5′-dithiobis-(2-nitrobenzoic acid) or DTNB)) are commonly used to label thiols. However, the derivatized thiol might be unstable and light sensitive. Most of these methods that take advantage of nucleophilic interaction of thiol groups with electrophile centers in the derivatizing agent are not selective and the separation step after derivatization is a big drawback. Generally, the derivatizing agents are costly and they require relatively expensive instrumentation for detection. More importantly they can only be employed for in vitro analyses.
Electrochemical measurement and HPLC-MS are other sensitive and selective physical techniques used for the analysis of Cys and other biothiols and do not require derivatization. However, the detector in HPLC-EC is hard to stabilize and MS is an expensive and destructive technique. Enzyme-based techniques are only possible for homocysteine (hCys).
As previously stated, radionuclides are widely used in bioanalytical experiments to tag small molecules lacking optical/electrochemical activity. The mass difference introduced by radiolabels to the target analyte is negligible, as compared to other labeling methods such as fluorescent labeling. Thus, radiolabeling minimizes the perturbation on analyte properties, such as diffusion coefficient, binding kinetics, etc. Beta particles are emitted from 3H, 35S, 14C, and 32P, where the maximum energy (Emax) of emitted beta particles is 18 keV for 3H decay, 156 keV for 14C decay, 167 keV for 35S decay, and 1710 keV for 32P decay. Radiolabeled biomolecules have very similar chemical, physical, and biological properties to the unlabeled molecules. 35S-Cys, for example, undergoes thiol-disulfide exchange reactions similar to unlabeled Cys with a slightly different rate because of the isotope effect.
Because of the high prevalence of hydrogen in biomolecules, using 3H-labeled analytes is very advantageous. Beta particles emitted from 3H have lower penetration depth (maximum ca. 6 μm in water and 0.5 cm in air), making it a relatively safe isotope. However, it is more challenging to detect this radioisotope due to its low energy beta particles. The 35S isotope, which is a medium-energy isotope, has higher decay energy and therefore its beta particles travel a longer distance (maximum ca. 300 pm in water and 25 cm in air), making 35S a better tracer for sensitive detection of sulfur-containing analytes. Although 35S emits higher energy beta particles, the penetration depth is short enough that this radioisotope is also considered to be safe to work with, and it is commonly used in biological systems. As compared to 3H, 35S is also more convenient to work with because its shorter half-life (87.4 days) allows for easier disposal. LSA is a very efficient technique for measurement of 35S. However, LSCs is not practical or feasible for the same reasons as previously discussed.
Due to the limitations of LSCs, solid scintillation analysis (SSA) may be a better choice for intracellular analysis of the radionuclides with low energy and short range emission. SSA facilitates intimate contact of the radionuclides with the scintillants which may be separated from the analytes for reuse. SPA is used for binding measurements and works based on the conversion of energy released from radionuclides bound to the sensor to detectable visible light. This conversion of energy is done using reporter organic fluorophores and inorganic crystals that transfer the energy via Förster Resonance Energy Transfer (FRET). The reporter fluorophores can be doped into solid matrices such as microplates or micro/nanoparticles. The binding moiety in SPA makes this procedure more sensitive, as compared to SSA.
A sensor with high cell-permeability and low possible interference that can measure intracellular thiols and disulfides and their ratio is desirable. The ideal sensor should have quick and dynamic binding capability so that thiol-disulfide homeostasis is not perturbed over the measurement time.
Material and instruments
Cystine, L [35S] dissolved in 0.01 N HCl (contains both monomer and dimer) was received from American Radiolabeled Chemicals (St. Louis, Mo.). Ultrapure L-Cys was obtained from Fluka (Milwaukee, Wis.). TCEP, NEM, DTT, TEOS (98%), APTS (99%), MPTS (97%), AAPH (97%) and sodium hydrogen phosphate (99%) were purchased from Sigma Aldrich (St. Louis, Mo.). Styrene (99%), pTP (99%), and DMPOPOP (98%) were purchased from ACROS Organics (Geel, Belgium). SBH was a product of Mallinckrodt (St. Louis, Mo.). ACS-grade isopropyl alcohol was purchased from Merck (Kenilworth, N.J.). Biocount LSC was obtained from Research Product International (Mt. Prospect, Ill.). ACS-grade ammonium hydroxide was purchased from EMD Millipore (Billerica, Mass.). Sodium hydroxide (98.9%) was purchased from Avantor Performance Materials (Center Valley, Pa.). Angeli's salt (Na2N2O3) was synthesized as described in Smith &Hein (“The alleged role of nitroxyl in certain reactions of aldehydes and alkyl halides”. Journal of the American Chemical Society 1960, 82 (21), 5731-5740.) Nanopure milliQ water (18 MΩ) was obtained using Easy Pure UV/UF Barnstead.A Tecnai G2 Spirit 20-120 kV transmission electron microscope (TEM) was used to characterize the NPs. A Beckman Coulter LS 60001C was used for scintillation measurements.
Nanosensor Fabrication and Characterizations
Polystyrene/silica core-shell NPs doped with red-shifted scintillants were fabricated to convert the emitted low energy beta particles from the labeled biomolecule to a visible light via FRET. First, inhibitors were removed from styrene by passing 3 grof liquid styrene through an alumina column. Styrene was then polymerized using a surfactant-free emulsion polymerization method in the presence of scintillant fluorophores (pTP and DMPOPOP with mole ratio of 10:1) in order to produce reporter fluorophore-doped NPs. A cationic initiator, AAPH, was utilized to trigger the polymerization of styrene in 50 mL degassed nanopure water overnight. Then any excess unpolymerized styrene was removed from the flask by rotary evaporation. The concentration of PS NPs was obtained by freeze-drying. PS NPs were covered with a silica shell which protects the assembly and increases its hydrophilicity. The silica shell is modifiable so that the surface of the NPs can be decorated with a variety of functional groups that covalently bind to the analyte of interest. Amine and thiol functionalized NPs were prepared using 10% APTS and MPTS, respectively, in addition to 90% TEOS. About 60 mg PS NPs were dispersed in a mixture of 200 mL isopropanol and 40 mL water. 7 or 5 mL NH4OH was added to adjust the pH for MPTS or APTS, respectively. Then 2 mL of a mixture of 10% MPTS or APTS and 90% TEOS were added dropwise to the PS NPs to obtain PS-MPTS or PS-APTS, respectively. Transmission electron microscopy with accelerating voltage of 100 kV was used to explore the NPs' size and surface morphology on the polystyrene core, and the polystyrene-silica core-shell NPs. TEM samples were prepared on cupper grids of 300-mesh coated with carbon. A drop of diluted suspension was applied to the surface of the grid and left for 10 minutes. Then the residual sample was removed using a piece of filter paper.
Binding Experiment
Three trials were prepared for all samples in all of the following experiments.
Phosphate buffer (100 mM) pH 7 was prepared and degassed with argon for 20 minutes. 35S-CySS dissolved in 0.01 N HCl that was frozen was thawed and 40 μL of the stock was diluted in 1060 μL buffer. The volumes of reagents were chosen such that the pH stayed buffered after dilution. Solutions of 0, 25, 50, and 100 μL 35S-Cys was added to 1 mL PS-MPTS NPs with concentration of 1 mg/mL. 35S-Cys concentration in these samples ranges from 0 to about 4 nM. A set of control samples with PS-APTS NPs was also prepared to investigate the binding of 35S-Cys to NPs with amine functional groups. All reagents were freshly made and used except for the NPs that are stable over a long time.
Disulfide Cleavage/Exchange
Excess TCEP and DTT (a few mM) were added to the mixture of 35S-Cys and PS-MPTS NPs to cleave the formed bonds. Excess unlabeled Cys (a few mM) was added to the mixture of 35S-Cys and PS-MPTS NPs for disulfide exchange. Data collection was repeated over time to follow the cleavage/exchange reaction.
pH-Dependent Binding
Binding of 35S-Cys to PS-MPTS NPs in three different buffers was performed. For each pH NPs were dispersed in the corresponding buffer and 35S-Cys was diluted in the same buffer. MES (pH 5), PBS (pH 7), and carbonate (pH 9) were chosen for this experiment. About 22 μL of 35S-Cys was diluted in 2 mL of buffer and 650 μL of this solution was added to 1.5 mL of PS-MPTS NPs dispersed in buffer (conc.=1 mg/mL). SPA was measured at different pH values.
Thiol-Blocking by NEM
35S-Cys (˜5 nM) in phosphate buffer pH 7 and PS-MPTS NPs (1 mg/mL) dispersed in the same buffer were separately mixed with excess NEM (˜16 mM) in order to block the thiols. Then the thiol-blocked reagents (35S-Cys and PS-MPTS NPs) were mixed to follow the scintillation inhibition due to binding inhibition. Thiol blocking on both reagents (35S-Cys and PS-MPTS NPs) as well as only one of them were tested. In the NEM concentration-dependent experiment, only the thiols on 35S-Cys (-1 nM) were blocked. A control sample was also prepared without NEM to observe the scintillation due to the specific binding of 35S-Cys to PS-MPTS. NEM treatment for thiol blocking was performed for 10 minutes and then the thiol-blocked 35S-Cys and PS-MPTS were mixed. The control sample did not have any NEM and 35S-Cys was added to PS-MPTS at the same time as the NEM-treated reagents.
Thiol-Blocking by Nitroxyl (HNO)
35S-Cys (˜5 nM) in phosphate buffer pH 7 and PS-MPTS NPs (1 mg/mL) dispersed in the same buffer were separately mixed with excess Angeli's salt (˜2 mM). Then they were mixed to follow the scintillation inhibition due to binding inhibition. Thiol blocking on both reagents (35S-Cys and PS-MPTS NPs) as well as only one of them were tested. In the HNO concentration-dependent experiment, only the thiols on 35S-Cys (˜1 nM) were blocked.
Thiol Generation by SBH
Sample preparation was performed similar to thiol-blocking by NEM. 35S-Cys (˜2.5 nM) in phosphate buffer pH 7 and PS-MPTS NPs (1 mg/mL) dispersed in the same buffer were separately mixed with excess NEM (˜30 mM) in order to block the thiols. Then the thiol-blocked reagents (both 35S-Cys and PS-MPTS NPs) were mixed to follow the scintillation inhibition due to binding inhibition. A control sample was also prepared without NEM to observe the scintillation due to the specific binding of 35S-Cys to PS-MPTS. After reading SPA, excess SBH (˜15 mM) was added to the sample with NEM, to specifically cleave disulfide bonds of 35S-CySS and reduce it to more thiols and observe scintillation due to new bonds formed between NPs and newly reduced 35S-Cys.
Disulfide Generation by Metals
35S-Cys (2.5 nM) was treated with few representative metals (100 ppm) prior to addition to1.5 mL PS-MPTS NPs (1mg/mL). Then 35S-Cys was added to NPs to read scintillation response due to binding. Thiol-disulfide ratio is the key factor in scintillation response, because scintillation occurs on PS-MPTS NPs only due to binding. A set of control samples was prepared with the addition of the same metal-treated 35S-Cys samples to 1 mL LSC. Scintillation response in LSC is not dependent on binding and should stay the same for any thiol-disulfide ratio.
Characterization of Thiol-Functionalized NPs
TEM was performed on PS NPs before and after silica coating to measure their size and explore the surface morphology of the NPs.
Disulfide Binding of 35S-Cys to PS-MPTS NPs
To illustrate that thiol-functionalized NPs (PS-MPTS) could be used for SPA-based detection of 35S-labeled analytes, the binding of a model compound, 35S-Cys, was examined and the scintillation was quantified. Cys has a thiol group which can bind to another thiol from other molecules to form a disulfide bond. This covalent bond is stable unless a reducing agent is added. In order to investigate the binding of 35S-Cys to thiol-functionalized NPs (PS-MPTS), a binding experiment was performed.
Disulfide Exchange/Cleavage
In order to test for the reversibility of disulfide binding on PS-MPTS NPs, disulfide exchange/cleavage was investigated. Disulfide bonds formed between 35S-Cysand PS-MPTS can be cleaved by reducing agents, such as TCEP and DTT. TCEP and DTT were both used as reducing agents because they have different reducing potential and chemistry. Also, disulfide exchange using unlabeled Cys can dynamically displace bound radiolabeled Cys molecules from the surface of the PS-MPTS NPs, which decreases the scintillation intensity.
The circles in
pH-Dependent Binding
CySS is almost 40% of total Cys equivalent in plasma and Cys is known to undergo a pH-dependent dimerization to CySS. The Binding of 35S-Cys to PS-MPTS NPs at different pH values was examined to evaluate the effect of pH on scintillation response. The binding affinity is a function of the fraction of amino acid in its active monomer form and that is a function of pH as shown in
Data presented in
Thiol-Blocking by NEM Inhibits Binding of 35S-Cysteine to PS-MPTS NPs.
NEM is a highly reactive cyclic alkene, which can react with thiols as an alkylating agent to form a thioether. This N-substituted maleimide is commonly used in protein chemistry as a thiol-blocking reagent, which benefits from its small size. In this experiment, NEM is used to block the thiol groups on both PS-MPTS NPs and 35S-Cys.
There is no specific binding expected from thiol-blocked reagents (
The data illustrates successful blocking of thiol groups on 35S-Cys and PS-MPTS NPs, which is another strong evidence for specific binding of 35S-Cys to PS-MPTS NPs via disulfide binding. Thiol blocking by NEM was also performed on a fixed concentration of 35S-Cys by adding different concentrations of NEM. More binding inhibition was observed with higher concentration of NEM, as shown in
Thiol Generation by Reduction of Disulfides using SBH
NEM can be used to block the thiol groups on 35S-Cys to inhibit binding to PS-MPTS NPs, as explained above. In order to measure the remaining part of the amino acids that are in disulfide form, a reducing agent can be used to cleave the disulfide bonds and create new thiols to bind to the NPs. SBH (NaBH4) is a reducing agent that can cleave the disulfide bonds of CySS and create new Cys molecules. It also inactivates any excess blocking agent (NEM) which prevents blocking of newly formed thiols and thus allows for new bindings to the NPs. This experiment was performed for 35S-Cys at pH 7 on PS-MPTS NPs. The first column of
Thiol-Blocking by Nitroxyl (HNO) Partially Inhibits Binding of 35S-Cysteine to PS-MPTS NPs.
Angeli's salt decomposes to variety of nitrogen oxide species depending on pH. Nitric oxide (NO) is the dominant product of the decomposition of Angeli's salt at a pH less than 4. HNO is produced in the pH range 4-8 which then dimerizes and decomposes to nitrous oxide (N2O). The protonation of N in Angeli's salt produces HNO which has shown promising characteristics as a treatment for cardiovascular disorders. HNO is also a potential drug for breast cancer and is used as a treatment for alcoholism. HNO produced from Angeli's salt decomposition is a thiophil which rapidly reacts with cysteine and blocks thiols to yield sulfinamide, as shown in the following scheme:
ONNOO2−+H+→HNO+NO2−
HNO+RSH→RSONH2
Therefore, HNO can be indirectly measured with the biosensor presented in this report by blocking the thiols and following the decrease in scintillation response as a function of HNO concentration. Similar to NEM treatment, both 35S-Cys and PS-MPTS NPs were treated with Angeli's salt separately to block all thiol groups, and then mixed to check for the inhibition of specific binding. Disulfide bond formation was inhibited due to the formation of sulfinamide. However,
Thiol blocking by HNO was also performed on a fixed concentration of 35S-Cys by adding different concentrations of Angeli's salt. More binding inhibition was observed with higher concentration of HNO that is the product of Angeli's salt decomposition, as shown in
Disulfide Generation using Oxidative Metals
Transition metals can oxidize amino acids via Fenton reaction by forming hydroxyl radicals.10Representative metals were used in this work to investigate the oxidation of thiol groups of 35S-Cys to disulfides. Consequently, the scintillation intensity decreased due to the decreased binding of 35S-Cys to PS-MPTS NPs. In the control samples that were prepared by LSC, no difference was expected between the vials with no metals and those with added metals because the scintillation in LSC does not occur as a result of binding. Therefore, a constant intensity of scintillation is expected from LSC control samples due to the same amount of radiolabeled amino acid (monomer or dimer).
NanoSPA is a potentially strong biosensor candidate for real-time intracellular analyses because of the advantages discussed above. Feeding cells with radiolabeled analyte of interest leads to selective analysis in a pool of molecules with similar structures and properties. The present invention advantageously provides for a selective and specific biosensor for 35S-Cys which can detect this amino acid in a concentration range as low as sub-nanomolar at physiological pH and does not require any light source. Specific binding of 35S-Cys to the NPs occurs in seconds and can be useful for intracellular studies of thiol containing compounds. PS-MPTS NPs may also be potential biosensors for other biothiols such as hCys. These NPs may also be used for indirect measurement of thiol-reactive agents.
The following is a non-limiting example of scintillating core-shell nanoparticles as turn-on nanosensors for selective detection of 33P, a lower energy phosphorus isotope, to analyze kinase activity. Equivalents or substitutes are within the scope of the present invention.
Kinases are enzymes that catalyze phosphate transfer from adenosine triphosphate (ATP) to their target substrates to produce adenosine diphosphate (ADP) and phosphorylated substrates. Phosphates play important roles in transfer of energy (ATP) and information (DNA and RNA) in biology and phosphorylation is one of the most important post translational modifications which contribute to cell signaling cascades. Referring to
Various techniques are used to study kinases due to their importance as therapeutic targets for cancer treatment. Enzyme-linked immunosorbent assay (ELISA) requires multiple expensive reagents (e.g. antibodies) and many washing steps. Therefore, ELISA is a costly and low throughput technique for kinase studies. Fluorescence based assays are useful for kinase studies, but they are susceptible to interference and artifacts. Radioactive assays require radiolabeled materials but they are robust and less prone to artifacts as compared to fluorescence based assays and provide a better indicator of kinase activity. ATPγ33P is used in radioactive assays for kinase studies with radioisotope 33P on the γ-phosphate group that is transferred to the substrate of a kinase. LSA is used in phosphocellulose filter paper-based kinase studies and involves many washing steps that make it tedious and not environmentally friendly due to the high volume of radioactive and toxic LSC waste. SPA microbeads are used for separation-free analysis of kinases. SPA works based on the interaction of ionizing beta particles emitted from decaying 33P-radioisotope with scintillating fluorophores that are doped into SPA beads. The commercial SPA beads are dense and large (at least 1 pm) which makes them hard to disperse in samples. Additionally, SPA microbeads require specific coupling functionalities on the kinase substrates and on the beads, which adds to the cost and complexity of the analysis.
The nanoparticles of the present invention provide a cost-effective and high throughput technique for kinase studies. Polystyrene-silica core-shell nanoparticles with reporter fluorophores doped in the polystyrene core were fabricated to perform nanoSPA. The core-shell NPs are hydrophilic and easy to disperse in aqueous samples, as opposed to SPA microbeads. More importantly, nanoSPA is a separation-free or homogeneous technique, as opposed to other techniques such as ELISA. Only radiolabeled molecules bound to nanoSPA (i.e. 33P-phosphorylated substrate) generate a significant signal. Therefore, it is unnecessary to separate excess unbound radiolabeled reagents (i.e. ATPγ33P) from the mixture of NPs because the radioactive energy from unbound ATPγ33P dissipates in the solution before it reaches the NPs. The homogeneity of nanoSPA minimizes the waste disposal and complexity of the analysis, which is beneficial for high throughput screening of therapeutics. The analysis based on nanoSPA is performed by combining the kinase mixture (substrate+ATPγ33P±kinase) with NPs and measuring the scintillation signal, which makes this technique simple, robust, and easy to automate. Moreover, nanoSPA is useful for intracellular analysis of kinases due to the smaller size of the NPs as compared to commercial SPA microbeads. In some embodiments, NPs may be incorporated in live cells that are fed with ATPγ33P for intracellular study of kinases by optical imaging.
Materials and Methods
ATPγ33P was purchased from American Radiolabeled Chemicals Inc. (St. Louis, Mo.). 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) lipid was purchased from Avanti Polar Lipids, Inc. (Alabaster, Ala). SRC kinase kit (kinase, substrate, buffer, DTT, and MnCl2) was purchased from Promega Corporation (Madison, Wis.). TEOS (98%), APTS (99%), MPTS (97%), AAPH (97%) and sodium hydrogen phosphate (99%) were purchased from Sigma Aldrich (St. Louis, Mo.). Styrene (99%), pTP (99%), and DMPOPOP (98%) were purchased from ACROS Organics (Geel, Belgium). ACS-grade isopropyl alcohol was purchased from Merck (Kenilworth, N.J.). Biocount LSC was obtained from Research Product International (Mt. Prospect, Ill.). ACS-grade ammonium hydroxide was purchased from EMD Millipore (Billerica, Mass.). Sodium hydroxide (98.9%) was purchased from Avantor Performance Materials (Center Valley, Pa.). Purified water (18 MΩ) was obtained using Easy Pure UV/UF Barnstead. A Beckman Coulter LS 6000IC was used for scintillation measurements. A Tecnai G2 Spirit 20-120 kV transmission electron microscope (TEM) was used to characterize the NPs.
NanoSPA Particle Fabrication and Characterization
PS core NPs doped with scintillant fluorophores were prepared in three steps. First, inhibitors were removed from styrene by passing 3 g of styrene through an alumina column. Second, styrene was polymerized using a surfactant-free emulsion polymerization method. 3 g styrene was dispersed in 50 mL degassed nanopure water by stirring. Polymerization was initiated at 80° C. using AAPH. The emulsion was stirred continuously for 12 hrs. Excess styrene was removed from the flask by evaporation at reduced pressure. The concentration of PS NPs was obtained by freeze-drying a known volume of the NP solution and weighing the dry NPs. Third, primary and secondary scintillants, pTP (1.14 mmoles) and DMPOPOP (0.135 mmoles) dissolved in 20 mL of a 90:10 mixture of chloroform and isopropyl alcohol, were used for ex-situ doping of NPs by swelling the PS core. The organic mixture of scintillants was added to 500 mg PS NPs dispersed in 100 mL water and sonicated and stirred continuously for 2 hours. Excess organic phase was removed from the mixture by evaporation at reduced pressure.
The scintillant-doped PS core NPs were then coated with silica shells to obtain silica-coated (PS-TEOS) or thiol-functionalized (PS-MPTS) core-shell NPs. 110 mg PS core NPs were dispersed in a mixture of 200 mL isopropanol and 40 mL water. 7 mL NH4OH was added to adjust the pH of the mixture. Then 2 mL TEOS or 2 mL of a mixture of 90% (v/v) TEOS and 10% (v/v) MPTS was added dropwise to the PS NPs to obtain PS-TEOS or PS-MPTS, respectively. The mixture was stirred for an hour and then the core-shell NPs were obtained after centrifugation and rinsing with water.
PS-MPTS NPs were tested for non-specific adsorption of 33P-phosphorylated SRC kinase substrate by electrostatic attraction of positively charged SRC kinase substrates to negatively charged silane groups on nanoSPA. To specifically bind 33P-phosphorylated SRC kinase substrate to nanoSPA, PS-MPTS NPs were further functionalized with a cross linker (Mal-PEG2-NHS ester) by thiol-ene coupling of the maleimide end group of the linker to thiol groups on the surface of PS-MPTS NPs in citrate buffer (pH 6.5) for 30 minutes, to obtain PS-MPTS-NHS NPs. The NHS end of the linker on PS-MPTS-NHS NPs was used to facilitate peptide coupling of SRC kinase substrate through the amine groups on its lysine amino acid residues.
PS-TEOS NPs were coated with DOPC lipid to illustrate the minimization of non-specific adsorption of 33P-phosphorylated SRC kinase substrate to lipid-coated nanoSPA. Lipid coating was done by using 6 mg DOPC for 6 mg PS core NPs (6 samples). 6 mg DOPC (250 μL, 25 μg/μL) was obtained from the freezer and dried with argon to remove excess chloroform. The residual chloroform was removed by lyophilizing for 4 hours. 3 mL nanopore water was added to the dried DOPC and mixed. The multilamellar vesicles of DOPC that was formed in the suspension of DOPC in water were broken into vesicles by sonication of the lipid suspension in a glass vial. Sonication was performed for 10 minutes and stopped for 10 minutes to avoid heating the sample. The sonication/rest cycle was repeated four times. The suspension of DOPC vesicles was divided into 6 plastic scintillation vials and 500 μL DOPC (2 mg/mL) was obtained in each vial. 1 mg PS-TEOS NPs in 25 μL water was added to each vial to preserve the concentration of DOPC. The mixture of PS-TEOS NPs and DOPC vesicles was incubated at room temperature overnight (at least 12 hours) to obtain lipid-coated NPs (PS-TEOS-DOPC) after centrifugation for 15 minutes at 3000 rpm. There was no rinsing step or sonication after the NPs were collected.
Analysis of SRC Kinase Activity
SRC kinase was used to phosphorylate its peptide substrate (KVEKIGEGTYGVVYK-amide) with ATPγ33P or a mixture of ATP and ATPγ33P. All samples were prepared in SRC kinase buffer (pH 7). Kinase activity analyses by nanoSPA were performed by combining a pre-incubated kinase mixture (substrate+ATPγ33P±kinase) with NPs and measuring the scintillation signal using LSC instrument. 1 mL NPs (1 mg core/mL) was used for all samples in nanoSPA experiments.
Analysis of SRC Kinase Activity in Free Solution
ATP-mix was prepared by combining 1 μL ATPγ33P (8.72 μCi) with 20 μL 0.13 mM ATP. A mixture of 4 μL 2 μg/μL SRC kinase and 10 μL 0.1 μg/μL SRC kinase substrate (KVEKIGEGTYGVVYK-amide) was prepared. Then 10 μL ATP-mix was added to the kinase mixture (24 μL total volume). A negative control sample was prepared using the mixture of SRC kinase substrate and ATP-mix, with no SRC kinase enzyme. 4 μL buffer was added to the negative control sample to compensate for the volume. Table 1 shows the amount of kinase mixture components for positive and negative control samples. Kinase mixtures were incubated at room temperature for 30 minutes for phosphorylation reaction on SRC kinase substrate. Phosphocellulose filter paper-based LSA was performed for measurement of phosphorylation. The kinase mixtures were deposited on filter papers and washed three times with 0.85% phosphoric acid and once with acetone to remove excess ATPγ33P. The filter papers were submerged in 3 mL LSC to monitor the scintillation due to immobilized 33P-phosphorylated SRC kinase substrates on the filter paper.
Referring to
Analysis of SRC Kinase Activity by NanoSPA (Specific Binding and Non-Specific Adsorption)
Kinase mixtures were prepared in a set of separate experiments, see Table 2, using ATP-mix (Hot-Cold ATP), ATP (Cold ATP), andATPγ33P (Hot ATP), as described above. ATP-mix was prepared for Hot-Cold ATP samples by combining 6 μL ATPγ33P (50.91 μCi) with 114 μL 0.13 mM ATP. The kinase mixtures were incubated for 30 minutes in case of Cold ATP and Hot-Cold ATP samples. However, Hot ATP samples were incubated overnight because of much lower concentration of ATP to provide enough time for the phosphorylation reaction.
To evaluate SRC kinase activity and the sensitivity of nanoSPA to ATP, thiol-functionalized NPs, doped with pTP and DMPOPOP, were fabricated and used to immobilize 33P-phosphorylated SRC substrate on the surface of nanoSPA. NanoSPA functionalized with a sulfhydryl-to-amine crosslinker, i.e. PS-MPTS-NHS, was used for specific binding of 33P-phosphorylated SRC substrate. NanoSPA without the sulfhydryl-to-amine crosslinker, PS-MPTS, was used for non-specific adsorption of 33P-phosphorylated SRC kinase substrates to the surface of NPs.
To improve the signal to background ratio, a similar experiment was performed using only ATPγ33P. The kinase mixtures were prepared by mixing ATPγ33P, SRC kinase substrate, and with or without SRC kinase. Incubation of kinase mixtures overnight facilitated phosphorylation over a longer time to compensate for the low concentration of ATPγ33P.
Analysis of SRC Kinase Activity by Lipid-Coated NanoSPA (Inhibition of Non-Specific Adsorption)
Kinase mixtures were prepared using ATPγ33P, as previously described, and incubated overnight, see Table 3. To demonstrate the reduction of non-specific adsorption of 33P-phosphorylated SRC kinase substrates to nanoSPA, a set of samples were prepared using lipid-coated NPs, PS-TEOS-DOPC to monitor scintillation. PS-TEOS NPs were used for non-specific adsorption of 33P-phosphorylated SRC kinase substrates to nanoSPA to compare the scintillation counts on lipid-coated NPs to NPs without lipid coating.
DOPC coating on the NPs led to decreased scintillation counts by inhibition of non-specific adsorption of 33P-phosphorylated SRC substrate to the surface of the NPs.
Analysis of SRC Kinase Activity by NanoSPA at Varying Concentrations of ATPγ33P
To demonstrate the sensitivity of nanoSPA to ATP, kinase mixtures were prepared with constant concentration of SRC kinase and SRC substrate at varying concentrations of ATPγ33P, see Table 4. PS-MPTS-NHS NPs were used to monitor phosphorylation by specifically binding to 33P-phosphorylated SRC kinase substrates. A negative control sample was prepared at the highest concentration of ATPγ33P with no SRC kinase added. Kinase mixtures were incubated overnight to provide enough time for the phosphorylation reaction. These samples were then mixed with PS-MPTS-NHS NPs and the scintillation signal was collected by LSC.
While the SPA particles of the present invention have been shown to function as sensors for thiol-disulfide measurements and for intracellular analysis of kinases, it is to be understood that these are non-limiting examples, and that SPA may be used in other applications for analyzing various molecular species.
As used herein, the term “about” refers to plus or minus 10% of the referenced number.
Various modifications of the invention, in addition to those described herein, will be apparent to those skilled in the art from the foregoing description. Such modifications are also intended to fall within the scope of the appended claims. Each reference cited in the present application is incorporated herein by reference in its entirety.
Although there has been shown and described the preferred embodiment of the present invention, it will be readily apparent to those skilled in the art that modifications may be made thereto which do not exceed the scope of the appended claims. Therefore, the scope of the invention is only to be limited by the following claims. Reference numbers recited in the claims are exemplary and for ease of review by the patent office only, and are not limiting in any way. In some embodiments, the figures presented in this patent application are drawn to scale, including the angles, ratios of dimensions, etc. In some embodiments, the figures are representative only and the claims are not limited by the dimensions of the figures. In some embodiments, descriptions of the inventions described herein using the phrase “comprising” includes embodiments that could be described as “consisting of”, and as such the written description requirement for claiming one or more embodiments of the present invention using the phrase “consisting of” is met.
This application is a non-provisional and claims benefit of U.S. Provisional Patent Application No. 62/477,638, filed Mar. 28, 2017, and U.S. Provisional Patent Application No. 62/414,557, filed Oct. 28, 2016, the specification(s) of which is/are incorporated herein in their entirety by reference.
This invention was made with government support under Grant No. R21 EB019133 awarded by NIH. The government has certain rights in the invention.
| Number | Date | Country | |
|---|---|---|---|
| 62477638 | Mar 2017 | US | |
| 62414557 | Oct 2016 | US |
