SOLID PHASE MICROEXTRACTION COATING
The present disclosure provides an extraction coating for an SPME sampling instrument, where the extraction coating includes a sorptive material immobilized in a fluorocarbon polymer that is compatible with thermal-assisted desorption techniques, solvent-assisted desorption techniques, or both. The disclosure also provides SPME sampling instruments, methods of making an SPME sampling instrument, and methods of extracting an analyte from a sample matrix using the SPME coating.
This application is a continuation of U.S. patent application Ser. No. 15/446,972, filed Mar. 1, 2017, which claims the benefit of priority of U.S. Provisional Patent Application No. 62/302,710 filed Mar. 2, 2016, which is hereby incorporated by reference.
FIELDThe present disclosure relates to solid phase microextraction coatings.
BACKGROUNDThe following paragraphs are not an admission that anything discussed in them is prior art or part of the knowledge of persons skilled in the art.
Solid phase microextraction (SPME) is an approach for sample preparation that may be used in various analytical methods. SPME devices include an extraction coating present on a support. The extraction coating includes adsorptive or absorptive particles, which may have different geometries. Exposure of the SPME device directly into a matrix or into its headspace, for a certain period of time, extracts and enriches analytes contained in the sample matrix.
The SPME process is governed by the partitioning of analytes from the matrix onto or into the extraction phase, and extraction efficiency of an analyte depends on the analyte's affinity toward the adsorptive or absorptive particles present in the extraction coating.
After extraction and enrichment of the analytes onto or into the extraction phase, the SPME device may be placed in an analytical device where the analytes are desorbed and analyzed. Alternatively, the analytes may be desorbed separately and analyzed separately.
INTRODUCTIONThe following introduction is intended to introduce the reader to this specification but not to define any invention. One or more inventions may reside in a combination or sub-combination of the apparatus elements or method steps described below or in other parts of this document. The inventors do not waive or disclaim their rights to any invention or inventions disclosed in this specification merely by not describing such other invention or inventions in the claims.
SPME integrates sampling and sample preparation into a single step. Because of this, the SPME coating must be compatible with the sample matrix, the analytical device, and the desorption process used by or before the analytical device. It is also desirable for the SPME device to be capable of being used multiple times without degrading, without accumulating components of the matrix that could generate errors or incorrect readings in the analytical device, or both.
It is desirable to develop an SPME coating that addresses or ameliorates one or more shortcomings associated with known SPME coatings. For example, it is desirable to develop an SPME coating that is: (a) compatible with a plurality of different sample matrixes, (b) compatible with a sample matrix that was previously incompatible with SPME devices, (c) compatible with a plurality of different analytical devices, (d) compatible with an analytical device that was previously incompatible with SPME devices, (e) compatible with a plurality of different desorption methods, (f) compatible with a desorption method that was previously incompatible with SPME devices, or (g) any combination thereof.
Desorption of the analytes may be followed by a detection step (such as mass spectrometry), with an optional separation step (such as chromatography) in between. In some examples, the analyte may be directly desorbed from the SPME coating and transferred to the detector (such as a mass spectrometer) while omitting a separation step. Analytes that are adsorbed or absorbed onto or into the SPME coating may be desorbed using thermal-assisted desorption, solvent-assisted desorption, or both. The method of desorption may be determined by the separation/detection system, by the SPME coating, or both. For example, an SPME coating may not be compatible with thermal desorption if the coating is thermally labile; and/or may not be compatible with organic solvent-assisted desorption if the chemical composition of the coating changes on exposure to the organic solvent. An SPME coating that may be used in both thermal desorption and solvent desorption may be used, for example, in both liquid- and gas-chromatographic applications, thus broadening the amount of chemical information that can be obtained from a sample matrix.
An SPME coating according to the present disclosure may be compatible with a biological sample that includes macromolecules and/or other biological components. Such an SPME coating may reduce or eliminate accumulation of the macromolecules and/or matrix components on their surface. SPME coatings with such matrix compatibility features may provide greater extraction and/or more accurate analysis of medium- to low-volatile analytes, such as analytes from in-tissue or in vivo samples. For example, such an SPME coating may be useful in metabolomics or untargeted analysis, where it is desirable for the coating to extract a broad range of medium- to low-volatile analytes, while also reducing or eliminating accumulation of the macromolecules and/or matrix components on the coating surface, and being compatible with different desorption strategies in order to identify the different analytes.
The present disclosure provides an SMPE coating that attempts to address or ameliorate one or more shortcomings involved with known SPME coatings. SPME coatings according to the present disclosure include an extraction coating that has particulate sorptive material immobilized in a fluorocarbon polymer that is compatible with thermal-assisted desorption techniques, solvent-assisted desorption techniques, or both.
Fluorocarbon polymers are typically characterized as non-interactive with non-fluorocarbon material. However, the authors of the present disclosure surprisingly determined that fluorocarbon polymers according to the present disclosure were able to anchor non-fluorocarbon based particulate sorptive material to a support. Further, fluorocarbon polymers are hydrophobic. Accordingly, the fluorocarbon polymers were not expected to properly embed particulate sorptive material having polar character and anchor them to the support. Even more surprisingly, the authors determined that polar particulate sorptive material could be embedded.
In another aspect, the present disclosure provides a SPME sampling instrument that includes a support, and an extraction coating covering at least a portion of the support. The extraction coating includes particulate sorptive material immobilized in a fluorocarbon polymer that is compatible with thermal-assisted desorption techniques, solvent-assisted desorption techniques, or both.
In yet another aspect, the present disclosure provides a method of making an SPME coating. The method includes mixing a fluorocarbon polymer and a particulate sorptive material in a solvent; applying the mixture to a support to form a substantially uniform SPME coating layer; and removing the solvent. The solvent may be a fluorocarbon-based fluid. The fluorocarbon polymer may be soluble in the fluorocarbon-based fluid, which allows the coating layer to be formed without curing the fluorocarbon polymer.
In still another aspect, the present disclosure provides a method of extracting an analyte from a sample, such as a small molecule analyte. The method includes exposing an SPME sampling instrument to the analyte, where the SPME sampling instrument includes a support, and an extraction coating covering at least a portion of the support. The extraction coating includes particulate sorptive material immobilized in a fluorocarbon polymer that is compatible with thermal-assisted desorption techniques, solvent-assisted desorption techniques, or both. The authors of the present disclosure determined that exemplary sampling instruments were capable of extracting analytes (with a broad range of polarity and/or molecular weight) from biological matrixes, which contain macromolecules such as proteins and cells, without acting as a barrier for the analytes.
The authors of the present disclosure determined that the combination of exemplary hydrophilic-lipophilic-balance (HLB) particles and at least one exemplary fluorocarbon polymer could be used in gas chromatography, liquid chromatography, capillary electrophoresis, and direct coupling to an analytical instrument (such as a mass spectrometer) by omitting chromatographic separation.
In one aspect, the present disclosure provides a solid-phase micro-extraction (SPME) coating that includes a sorptive particulate material immobilized in a fluorocarbon polymer that is compatible with a thermal-assisted desorption technique, a solvent-assisted desorption technique, or both.
The fluorocarbon polymer may comprises a polymer formed from the polymerization of a fluorocarbon monomer or a mixture of monomers that include a fluorocarbon monomer. The fluorocarbon monomer may be: vinyl fluoride (VF1), vinylidene fluoride (VDF), tetrafluoroethylene (TFE), hexafluoropropylene (HFP), perfluoropropylvinylether (PPVE), perfluoromethylvinylether (PMVE), or chlorotrifluoroethylene (CTFE); or the mixture of monomers comprises: vinyl fluoride (VF1), vinylidene fluoride (VDF), tetrafluoroethylene (TFE), hexafluoropropylene (HFP), perfluoropropylvinylether (PPVE), perfluoromethylvinylether (PMVE), chlorotrifluoroethylene (CTFE), or a combination thereof, and optionally ethylene (E) and/or propylene (P).
The fluorocarbon polymer may be a polymer of, or a copolymer that includes: a polytetrafluoroethylene (PTFE), a fluorinated ethylene-propylene (FPE), a fluoroelastomer [tetrafluoroethylene-propylene] (FEPM), or any combination thereof.
The fluorocarbon polymer may be a copolymer of tetrafluoroethylene and 2,2-bistrifluoromethyl-4,5-difluoro-1,3-dioxole, such as a copolymer in the ratio of 13:87 or a copolymer in the ratio of 35:65.
The fluorocarbon polymer may have: a thermal stability up to 320° C.; a chemical stability towards a non-fluorinated solvent such as: a short chain alcohol, a hydrocarbon (for example an aliphatic, a cyclic, or an aromatic hydrocarbon), water, a chlorinated solvent, an ester, an ether, a nitrile, or any combination thereof; a chemical stability towards a solvent having a pH from 0.01 to 14; or any combination thereof.
The fluorocarbon polymer may be a mixture of polymers.
The fluorocarbon polymer may have carbon-fluorine (C—F) bonds and carbon-hydrogen (C—H) bonds, wherein fewer than 5% of all the C—F and C—H bonds in the fluorocarbon polymer are C—H bonds. In some examples, the fluorocarbon polymer may have substantially no C—H bonds.
The sorptive material may be a porous material having meso-, macro-, or micro-pores. At least 80% of the pores may have a diameter from about 10 Å to about 10,000 Å.
The sorptive material may be a porous material having pores with diameters from about 100 Å to about 180 Å. At least 80% of the pores may have a diameter from about 100 Å to about 180 Å.
The sorptive material may have a surface area of about 10 m2/g to about 3000 m2/g, such as about 200 m2/g to about 800 m2/g.
The sorptive material may include particles, nanosheets, nanotubes, or any combination thereof. The sorptive material may be inorganic, organic, a hybrid inorganic/organic material, or a mixture of both inorganic and organic materials.
The particles may have diameters from about 1 nm to about 100 μm, such as from about 3 μm to about 10 μm, or from about 3 μm to about 7 μm. The nanosheets may have thicknesses from about 1 nm to about 100 nm. The nanotubes may have diameters from about 1 nm to about 10 nm, such as from about 4 nm to about 6 nm.
The sorptive material may include: normal-phase silica particles, C-1/silica particles, C-4/silica particles, C-6/silica particles, C-8/silica particles, C-18/silica particles, C-30/silica particles, reverse-phase amide silica particles, HS-F5/silica particles, phenyl/silica particles, cyano/silica particles, diol/silica particles, ionic liquid/silica particles, molecular imprinted polymer particles, hydrophilic-lipophilic-balance (HLB) particles, carboxen 1006 particles, carbowax particles, divinylbenzene (DVB) particles, octadecylsilane particles, nanoparticles, processed mineral based particles, carbon nanotubes, functionalized-carbon nanotubes, graphene, graphene oxide, functionalized-graphene, quantum dots, organic polymeric particles optionally functionalized with an organic moiety (such as a carbon chain, a strong cation moiety, a weak cation moiety, a strong anion moiety, or a weak anion moiety), inorganic polymeric particles optionally functionalized with an organic moiety (such as a carbon chain, a strong cation moiety, a weak cation moiety, a strong anion moiety, or a weak anion moiety), or any combination thereof.
In some examples, the sorptive material may include hydrophilic-lipophilic-balance particles, such as particles comprising the copolymerization polymer product of N-vinylpyrrolidinone and divinylbenzene.
The fluorocarbon polymer may be a copolymer of tetrafluoroethylene and 2,2-bistrifluoromethyl-4,5-difluoro-1,3-dioxole in the ratio of 13:87; and the sorptive material may include hydrophilic-lipophilic-balance particles comprising the copolymerization polymer product of N-vinylpyrrolidinone and divinylbenzene.
The coating may have an average thickness from about 5 μm to about 500 μm.
The coating may have a homogeneous coating surface.
In another aspect, the present disclosure provides a solid-phase micro-extraction (SPME) sample instrument that includes: a support; and an extraction coating as discussed above covering at least a portion of the support.
The support may be a metal support, a metal alloy support, a fused silica support, a plastic support, a fluoro-plastic support, or a carbon material support. The carbon material support may be a carbon fiber fabric. The support may include stainless steel, titanium, or a nickel-titanium alloy such as nitinol.
The support may be shaped in the form of a needle, a mesh fabric, a metallic mesh, or a blade.
In another aspect, the present disclosure provides a method of making a solid-phase micro-extraction (SPME) sample instrument. The method includes: mixing a fluorocarbon polymer and a sorptive material in a solvent; applying the mixture to a support to form a substantially uniform SPME coating layer on at least a portion of the support; and removing the solvent.
The solvent may include a fluorocarbon-based fluid. The fluorocarbon-based fluid may include perfluorohexane, perfluoro(2-butyl-tetrahydrofurane), perfluorotripentylamine, or a combination thereof.
The solvent may include a perfluoro-polyether.
The solvent may include a non-fluorocarbon-based fluid. The fluorocarbon-based fluid and the non-fluorocarbon-based fluid may be in sufficient amounts to form an emulsion capable of acting as a porogen for the fluorocarbon polymer coating layer.
The method may include successively applying a plurality of layers of the mixture to form the SPME coating layer.
The sorptive material may include normal-phase silica particles, C-1/silica particles, C-4/silica particles, C-6/silica particles, C-8/silica particles, C-18/silica particles, C-30/silica particles, reverse-phase amide silica particles, HS-F5/silica particles, phenyl/silica particles, cyano/silica particles, dial/silica particles, ionic liquid/silica particles, molecular imprinted polymer particles, hydrophilic-lipophilic-balance (HLB) particles, carboxen 1006 particles, carbowax particles, divinylbenzene (DVB) particles, octadecylsilane particles, nanoparticles, processed mineral based particles, carbon nanotubes, functionalized-carbon nanotubes, graphene, graphene oxide, functionalized-graphene, quantum dots, organic polymeric particles optionally functionalized with an organic moiety (such as a carbon chain, a strong cation moiety, a weak cation moiety, a strong anion moiety, or a weak anion moiety), inorganic polymeric particles optionally functionalized with an organic moiety (such as a carbon chain, a strong cation moiety, a weak cation moiety, a strong anion moiety, or a weak anion moiety), or any combination thereof.
In some examples, the sorptive material may include hydrophilic-lipophilic-balance particles, such as particles comprising the copolymerization polymer product of N-vinylpyrrolidinone and divinylbenzene.
In particular examples, the fluorocarbon polymer may be a copolymer of tetrafluoroethylene and 2,2-bistrifluoromethyl-4,5-difluoro-1,3-dioxole in the ratio of 13:87; and the sorptive material may include hydrophilic-lipophilic-balance particles comprising the copolymerization polymer product of N-vinylpyrrolidinone and divinylbenzene.
The solvent may be perfluorohexane, perfluoro(2-butyl-tetrahydrofurane), perfluorotripentylamine, or any combination thereof.
In another aspect, the present disclosure provides a method of solid-phase micro-extraction (SPME), where the method includes exposing an extraction coating as discussed above to a sample matrix that includes at least one analyte; and desorbing the extracted analyte.
The desorbing may include exposing the SPME coating to a thermal-assisted desorption temperature, such as a temperature up to 300° C., and the method optionally further comprises gas chromatography or direct coupling to a spectroscopic technique suitable for detection of a thermally stable analyte, such as mass spectrometry.
The desorbing may include exposing the SPME coating to a solvent-assisted desorption solvent, and the method optionally further comprises liquid chromatography, gas chromatograph, capillary electrophoresis, or any spectroscopic technique suitable for determination of a solvent stable analyte.
The desorption may include electrothermal vaporization, arc and spark ablation, laser ablation, glow discharge, matrix-assisted laser desorption/ionization (MALDI), or desorption electrospray ionization (DESI), and the method optionally further comprises a spectroscopic technique, such as gas chromatography or direct coupling to mass spectrometry, suitable for detection of the analyte.
The analyte may be a polar compounds, such as a water-soluble compound. The analyte may be a non-polar compound. The analyte may be a non-fluorocarbon polymer.
The sample matrix may be a biological matrix or an environmental sample, such as a sludge or soil.
The SPME coating may be used more than once to extract and desorb the analyte from the sample matrix or its headspace.
Embodiments of the present disclosure will now be described, by way of example only, with reference to the attached Figures.
Generally, the present disclosure provides an extraction coating for an SPME sampling instrument, where the extraction coating includes a particulate sorptive material immobilized in a fluorocarbon polymer that is compatible with thermal-assisted desorption techniques, solvent-assisted desorption techniques, a solid sample desorption method, or a combination thereof. The extraction coating may be referred to as “an SPME coating”.
The particulate sorptive material may be material that can be used in solid phase extraction or partition chromatography (such as liquid chromatography). The particulate material may be, for example: particles, nanosheets, and/or nanotubes. As noted above, the authors of the present disclosure surprisingly determined that some exemplary fluorocarbon polymers were able to anchor particulate sorptive material to a support. Even more surprisingly, they determined that polar particulate sorptive material could be anchored to a support. The sorptive material may be adsorptive or absorptive, and may be referred to as “the adsorptive or absorptive material”.
A fluorocarbon polymer according to the present disclosure refers a polymer having carbon-fluorine (C—F) bonds and carbon-hydrogen (C—H) bonds where fewer than 5% of all the C—F and C—H bonds in the fluorocarbon polymer are C—H bonds. Some particular examples of fluorocarbon polymers according to the present disclosure have substantially no C—H bonds.
Thermal- and solvent-assisted desorption techniques contemplated by the present disclosure include techniques that may be used to introduce a solid sample into an analytical instrument, such as to a gas chromatography instrument, a liquid chromatography instrument, or a capillary electrophoresis instrument.
An extraction coating that is “compatible” with a thermal-assisted desorption technique refers to a coating that loses less than 5% of its weight when exposed to the thermal desorption conditions. An extraction coating that loses less than 5% of its weight when exposed to a temperature of up to about 310° C. may be considered to be compatible with common thermal-assisted desorption techniques for SPME.
An extraction coating that is “compatible” with a solvent-assisted desorption technique refers to a coating that does not swell when exposed to a solvent or a mixture of solvents. An extraction coating that does not swell when exposed to commonly used for liquid chromatography may be considered to be compatible with common solvent-assisted desorption techniques. Exemplary solvents may have pHs from about 0.01 to about 14 and may include short chain alcohols, hydrocarbons (aliphatic, cyclic, aromatic), water, chlorinated solvents, esters, ethers, nitriles, and combinations thereof.
The extraction coating may also be compatible with one or more other well-known solid sample desorption methods, such as: electrothermal vaporization, arc and spark ablation, laser ablation, glow discharge, matrix-assisted laser desorption/ionization (MALDI), or desorption electrospray ionization (DESI). An extraction coating that is “compatible” with one of these methods refers to a coating that releases the extracted analyte efficiently with the application of the method.
The fluorocarbon polymer may be selected based on a desired stability for a desorption method, a desired biocompatibility with a sample matrix, desired immobilization characteristics for the adsorptive or absorptive material, or any combination thereof. The fluorocarbon polymer may include a polymer formed from the polymerization of a fluorocarbon monomer or of a mixture of monomers that include a fluorocarbon monomer. The monomer may be, for example: vinyl fluoride (VF1), vinylidene fluoride (VDF), tetrafluoroethylene (TFE), hexafluoropropylene (HFP), perfluoropropylvinylether (PPVE), perfluoromethylvinylether (PMVE), or chlorotrifluoroethylene (CTFE). The mixture of monomers may include, for example: VF1, VDF, TFE, HFP, PPVE, PMVE, CTFE, or any combination thereof, and may optionally include ethylene (E) and/or propylene (P) so long as fewer than 5% of all the C—F and C—H bonds in the fluorocarbon polymer are C—H bonds.
In particular examples, the fluorocarbon polymer may be a polymer of, or a copolymer that includes: polytetrafluoroethylene (PTFE), fluorinated ethylene-propylene (FPE, tradename Teflon®), fluoroelastomer [tetrafluoroethylene-propylene] (FEPM, tradename AFLAS®), or any combination thereof. For example, the fluorocarbon polymer may be a Teflon Amorphous Fluoroplastic (Teflon AF) formulation, such as Teflon AF 2400 (which has a Tg of 240° C., and is also referred to as PTFE AF2400), or Teflon AF 1600 (which has a Tg of 160° C., and is also referred to as PTFE AF1600). PTFE AF 2400 is a copolymer of tetrafluoroethylene (13%) and 2,2-bistrifluoromethyl-4,5-difluoro-1,3-dioxole (87%). PTFE AF 1600 is a copolymer of tetrafluoroethylene (35%) and 2,2-bistrifluoromethyl-4,5-difluoro-1,3-dioxole (65%). These exemplary fluorocarbon polymers are particularly beneficial, in comparison to non-fluorocarbon polymers, when it is desirable to reduce or avoid the possibility that sample matrix will stick to the extraction coating.
The adsorptive or absorptive material may be selected based on the features of the intended analyte being extracted, and the intended desorption strategy. The adsorptive or absorptive material may be considered an extractive material in that it adsorbs or absorbs analytes for extraction and removal from the sample matrix. In some examples, the sorptive material may be a porous material having meso- macro- or micro-pores. In some examples, the pores may have diameters from about 10 Å to about 10,000 Å. Such pores allow the analyte to be extracted onto or into the adsorptive or absorptive material. The pores may preferably be from about 100 Å to about 180 Å. These pore sizes provide sufficient surface area to adsorb or absorb enough analytes to be detectable, while also being an appropriate size to allow desorption so that micro-condensation is reduced or avoided. Larger pore sizes may be desirable when the target analyte is a large biomolecule, such as a protein or a peptide. The surface area of the particulate sorptive material may be from about 10 m2/g to about 3000 m2/g. For example, the surface area of hydrophilic-lipophilic-balance (HLB) particles may be, from about 200 m2/g to about 800 m2/g, while the surface area of carbon nanotubes and nanosheets may be up to 3000 m2/g.
The adsorptive or absorptive material may be particles, nanosheets, and/or nanotubes that are suspended in the fluorocarbon polymer. Particles which may be used in coatings according to the present disclosure may be from about 1 nm to about 20 μm in diameter. Preferably, the particles may be from about 2 μm to about 20 μm in diameter, and more preferably from about 3 μm to about 10 μm in diameter. Even more preferably, the particles may be from about 3 μm to about 7 μm in diameter. Particles from about 3 μm to about 7 μm in diameter may be particularly useful as smaller sized particles have less of an effect on coating thickness and coating homogeneity than larger particles. Depending on the thickness of the coating and the relative size differences between the particles, using smaller particles may result in a coating that is more uniform in thickness and has a more homogeneous particle distribution than a coating made with larger particles. A “uniform thickness” should be understood to refer to a thickness that varies by less than 5% over the support surface. The particles may be spherical, or substantially spherical. Nanosheets which may be used in coatings according to the present disclosure may be about 1 nm to about 100 nm in thickness. Nanotubes which may be used in coatings according to the present disclosure may have a diameter from about 1 nm to about 10 nm. Preferably, the nanotubes have a diameter from about 4 nm to about 6 nm.
With some adsorptive or absorptive material, a homogeneous coating surface may be obtained when the largest dimension of the adsorptive material is less than about half the thickness of the coating. In the context of the present disclosure a “homogeneous coating surface” should be understood to refer to a coating surface having a substantially uniform distribution of the adsorptive or absorptive material and the fluorocarbon polymer on the support surface.
The adsorptive or absorptive material may be selected based on its compatibility with the intended desorption method. The adsorptive or absorptive material may be inorganic (for example a silica-based material or a metal oxide-based material), organic (for example a carbon-, carboxen- or divinylbenzene-based material), an inorganic/organic hybrid (for example a silica and organic polymer), or a mixture of inorganic and organic materials. In particular examples, the adsorptive or absorptive material may be: normal-phase silica particles, C-1/silica particles, C-4/silica particles, C-6/silica particles, C-8/silica particles, C-18/silica particles, C-30/silica particles, reverse-phase amide silica particles, HS-F5/silica particles, phenyl/silica particles, cyano/silica particles, diol/silica particles, ionic liquid/silica particles, molecular imprinted polymer particles, hydrophilic-lipophilic-balance (HLB) particles, carboxen 1006 particles, carbowax particles, divinylbenzene (DVB) particles, octadecylsilane particles, nanoparticles, processed mineral based particles, carbon nanotubes, functionalized-carbon nanotubes, graphene, graphene oxide, functionalized-graphene, quantum dots, or any combination thereof. The functionalized-carbon nanotubes and functionalized-graphene may be functionalized with a polar or non-polar functionality. The polar functionality may be ionic. The functional group may be used to tune the extraction capability of the adsorptive or absorptive material towards a targeted compound, or a targeted class of compounds. For example, a cationic compound may be targeted for extraction by using an anionic-functionalized-carbon nanotube, HLB particles may have a specific surface area from about 700 m2/g to about 900 m2/g; average pore diameters from about 70 angstroms to about 90 angstroms; total pore volumes from about 1.15 cm /g to about 1.45 cm3/g; and/or an average particle diameter from about 5.2 to about 5.7 μm. An exemplary HLB particle is made by Waters and sold under the trade name Oasis HLB. The Oasis HLB particles are made from copolymerization of two monomers, the hydrophilic N-vinylpyrrolidinone and the lipophilic divinylbenzene. The resulting HLB particles include both hydrophilic groups and lipophilic groups, and have a specific surface area of about 762 m2/g, an average pore diameter of about 79 angstroms, a total pore volume of about 1.18 cm3/g and an average particle diameter of about 5.47 μm.
Although any of the above adsorptive or absorptive materials may be used for disposable coatings, some adsorptive or absorptive materials are suitable for use in coatings that are used more than once. For example, organic or inorganic polymeric particles functionalized by organic moieties (such as carbon chains from C-1 to C-30, strong and weak cation moieties, or strong and weak anion moieties) may be used more than once in solvent-assisted desorption methods. In other examples, carbon based sorbents, inorganic sorbents, divinylbenzene-based particles, and hydrophilic-lipophilic-balance particles may be used more than once with thermal- and/or solvent-assisted desorption methods.
The extraction coating may be loaded with a standard prior to exaction of any analytes of interest from the sample matrix. The internal standard may help to account for variations in the sample preparation, coating thickness, instrument response, extraction conditions, desorption of the extracted analyte, ionization of the desorbed analyte, or any combination thereof. The internal standard may be loaded on the extraction coating by exposing the coating to a known concentration (for example 50.0 μg/L) of a known compound (for example codeine-D3, 6-acetylmorphine-D3, or cocaine-D3).
In another aspect, the present disclosure provides a SPME sampling instrument that includes a support, and an extraction coating covering at least a portion of the support, where the extraction coating is as discussed above.
The support may be formed from any acceptable material that would be amenable for the deposition of the SPME coating, and capable of being used with an intended matrix. The SPME sampling instrument may use a different support for matrices of different natures or viscosities. The support may include, for example: a metal, a metal alloy, fused silica, a polymer (e.g. polybutylene terephthalate), a plastic, a fluoro-plastic, glass wool fibers, or a carbon material (e.g. a carbon fiber fabric). The fabric may have a unit weight of about 100 g/m2 to about 300 g/m2. The fabric may have a thickness of about 400 to about 900 microns. The metal or metal alloy may be, for example: stainless steel, titanium, a nickel-titanium alloy, or any other metal or metal alloy known to a person of skill in the art. In particular examples, the support may be a flexible, inert, biocompatible nickel-titanium alloy, such as Nitinol. The support may be, for example, a metal with shape memory properties that enable the metal to maintain straightness, even after being inserted into tissue or being use in high-speed agitation. Nitinol is an example of a metal alloy with shape memory properties. The geometry of the support may be selected based on the sample matrix to be analyzed, the desorption technique to be used, or both. For example, a support that is to be used as a probe into a tissue, such as an in vivo tissue, may be shaped in the form of a needle or a blade, using a metal or metal alloy. In comparison to a probe having a large, blunt geometry, a needle-like support would facilitate insertion of the sampling instrument into the tissue while reducing the invasiveness of the sampling instrument. In another example, a support that is to be used as a probe in a liquid sample (such as a urine test for banned doping agents) may be shaped in the form of a carbon mesh fabric, a metallic mesh, a blade, or a thin-film. Such a support would increase the surface area available for extraction, which may result in a better detection limit.
Homogenous coating surfaces may be from about 2 μm to about 1000 μm in thickness. Coatings of this thickness may provide a suitable balance between the total amount of analyte capable of being adsorbed by the coating, and the speed and efficiency of extraction and desorption (which is based on the thickness of the extraction coating). Coating thickness may be selected based on the intended application for the sampling instrument.
In yet another aspect, the present disclosure provides a method of making an SPME sampling instrument. In some examples, the method includes mixing a fluorocarbon polymer and a sorptive material in a solvent; applying the mixture to a support to form a substantially uniform SPME coating layer; and removing the solvent. Methods where the fluorocarbon polymer is dissolved in a solvent, and the solvent is removed to leave a solid fluorocarbon polymer may be beneficial in situations where it is desirable to avoid thermal curing (such as when it is desirable to avoid in situ polymerization methods, in situations where the adsorptive or absorptive material includes chemical functionalities that could react with a monomer, or both).
Mixing the fluorocarbon polymer, the sorptive material, and the solvent may be performed in any order, or may be mixed all together at the same time. For example, the fluorocarbon polymer may be mixed with the solvent, followed by mixing the sorptive material; or the fluorocarbon polymer may be mixed with the sorptive material, following by mixing with the solvent; or the solvent and the sorptive material may be mixed together, followed by mixing the fluorocarbon polymer.
The fluorocarbon polymer may be dissolved in the solvent, such as a fluorocarbon-based fluid or a mixture that includes a fluorocarbon-based fluid. Examples of fluorocarbon-based fluids include: perfluorohexane (tradename Fluorinert™ FC-72), perfluoro(2-butyl-tetrahydrofurane) (tradename Fluorinert™ FC-75), and perfluorotripentylamine (tradename Fluorinert™ FC-70). FC-72 has a boiling point of 56° C. and may be selected for methods where it is desirable to coat the support rapidly and it is desirable to avoid thermal conditioning. FC-75 has a higher boiling point and slower evaporation rate, and may be selected for methods where it is desirable to coat the support more slowly since the coating slurry will be at substantially the same viscosity during the whole coating procedure. Thermal conditioning may be used to help remove higher boiling fluorocarbon-based fluids.
Fluorocarbon polymers are preferentially soluble in fluorocarbon-based fluids. Adding an additional solvent, such as a halogenated solvent, to the fluorocarbon-based fluid may reduce the solubility of the fluorocarbon polymer, or may generate an emulsion that may act as a porogen. Including a porogen in the solvent may be used to change the permeability of the dried fluorocarbon polymer and change the selectivity of extraction for a class of compounds. For example, FC-72, FC-75, FC-70, or any combination thereof may further include a perfluoro-polyether, such as Krytox 157-FSH (DuPont, average molecular weight of about 7,000-7,500 g/mol). Krytox 157-FSH can act as a porogen, generating pores in the dried fluorocarbon polymer that more preferentially allow certain classes of compounds (for example: pyridines) to be extracted from the sample matrix. The Krytox 157-FSH emulsion remains a part of the coating as the fluorocarbon-based fluid is evaporated, but is subsequently removed since it does not react with the polymer or the adherent material. Once removed, it leaves behind pores in the fluorocarbon polymer.
The fluorocarbon polymer may be dissolved in the solvent at a weight percentage of about 1% to about 20% wt/wt. In particular examples, the weight percentage may be about 2.6 wt % fluorocarbon polymer to solvent. Using about 2.6 wt % fluorocarbon polymer to solvent allows for both coating robustness and homogeneity.
In particular examples, the SPME coating may be prepared by covering a solid substrate with a suspension of adsorptive or absorptive particles (e.g. C-18/silica, divinylbenzene, carboxen 1006, or HLB), nanosheets (e.g. graphene, graphene oxide, or mesoporous carbon) or nanotubes (e.g. carbon nanotubes) in a solution of fluorocarbon polymer (e.g. PTFE, FPE, or FEPM) dissolved in fluorocarbon solvent (e.g. FC-72, FC-70, or FC-75).
In more particular examples, the fluorocarbon polymer is PTFE, such as PTFE AF 2400. When using PTFE AF 2400 and adsorptive or absorptive particles, such as HLB particles, the PTFE AF 2400 and the adsorptive or absorptive particles may be applied to the substrate in a ratio from about 0.5:1 to about 2:1 (w/w) based on the weight of the PTFE AF 2400 and the adsorptive or absorptive particles before dissolution in a solvent. More preferably, the PTFE AF 2400 and the adsorptive or absorptive particles are in a ratio of about 1:1 (w/tier). The PTFE AF 2400 may be mixed with sufficient solvent to result in a ratio of about 1:100 to about 4:100 (w/w) of PTFE AF 2400 to solvent. More preferably, the ratio of PTFE AF 2400 to solvent is from about 2:100 to about 3:100 (w/w). Even more preferably, the PTFE AF 2400 and solvent may be mixed to form a solution of about 2.6 wt % PTFE AF 2400. As noted above, the fluorocarbon polymer may be dissolved in a fluorocarbon-based fluid. The PTFE AF 2400 may be dissolved in FC-72, FC-75, FC-70, or any mixture thereof. Preferably, the PTFE AF 2400 is dissolved in perfluorohexane (Fluorinert™ FC-72).
The fluorocarbon polymer/sorptive material mixture may be applied to the support in any manner that results in a coating of substantially consistent thickness. For example, the mixture may be applied onto the support by: dipping, spreading, brush painting, spraying, spin coating or electrospinning. For a fabric support, a coating thickness from about 5 to about 250 micrometer may be obtained with a single layer application by controlling the speed of a bar coater (varying from 5 mm/sec to 150 mm/sec). A fabric support may be coated on both sides, for example by using two bar coater applications.
The mixture may be applied to the support in a plurality of layers, such as a sufficient number of layers to provide a desired coating thickness. Applying the mixture in a plurality of layers may result in a coating that has improved bonding to the substrate, more uniform coating thickness across the coated area, or both.
Since the fluorocarbon polymer is already polymerized before being applied to the support, it is not necessary to cure the polymer and multi-layered coatings may be produced substantially faster than in methods that require monomers to be polymerized or crosslinked. The coating is formed on the support by evaporation of the solvent. Depending on what solvent is used, the solvent may be evaporated at an elevated temperature, at a reduced pressure, through flow of a drying gas (such as nitrogen) over the mixture, by leaving the coated device for a sufficient period of time, or any combination thereof. In some examples, the solvent may be evaporated over a length of time that is from about 30 seconds to about 2 minutes. With particular solvents, the evaporation time may be about 1 minute. Perfluorohexane, for example, has a boiling point of 56 ° C. and a vapor pressure of 27 kPa at 25 ° C. and may be evaporated from the mixture if it is left at room temperature for about 1 minute.
The fluorocarbon polymer-based coating may be further coated with a biocompatible layer, such as a layer of a fluorocarbon polymer that lacks sorptive material or a non-fluorocarbon based biocompatible polymer. Examples of biocompatible non-fluorocarbon based polymers include: polyacrilonitrile, polydimethylsiloxane (PDMS), and polypyrrole (PPY). Such an additional biocompatible layer may result in the SPME coating having improved compatibility with an intended sample matrix.
In some other examples, the method of making an SPME sampling instrument includes polymerizing, on a support, a coating of a fluorocarbon-based monomer, or a mixture of monomers that includes a fluorocarbon-based monomer, around the sorptive material. The coating may be formed, for example: by mixing the monomer or mixture of monomers with the sorptive material and coating the support with the reaction mixture. The monomer may be, for example: vinyl fluoride (VF1), vinylidene fluoride (VDF), tetrafluoroethylene (TFE), hexafluoropropylene (HFP), perfluoropropylvinylether (PPVE), perfluoromethylvinylether (PMVE), or chlorotrifluoroethylene (CTFE). The mixture of monomers may include, for example: VF1, VLF, TFE, HFP, PPVE, PMVE, CTFE, or any combination thereof, and may optionally include ethylene (E) and/or propylene (P) so long as fewer than 5% of all the C—F and C—H bonds in the fluorocarbon polymer are C—H bonds.
Methods of making an SPME sampling instrument may additionally include one or more support processing steps. A support processing step may: clean a surface of the support, roughen a surface of the support, or both, where the processed surface is a surface to be coated with the SPME coating. A support processing step may improve attachment of at least a portion of the SPME coating to the support. Processing the support material may include cleaning the support by sonication in water, an organic solvent, or a mixture thereof. The organic solvent may be, for example: methanol, acetonitrile, isopropyl alcohol, or any mixture thereof. Roughening the surface to be coated may include: using an abrasive (such as sand paper or any other type of sanding device), etching in hydrochloric acid (for example for 5 to 60 min) if the support material includes a metallic substrate, or applying a voltage across the support (for example for 30 seconds at a voltage of 3.5 V in a solution of water saturated with sodium chloride) if the support material includes a metallic substrate. After a roughening step, the substrate may additionally be cleaned before being coated with the SPME coating. The cleaning may include sonication in a solution of water and methanol 50:50 (v/v) for 10 minutes.
In still another aspect, the present disclosure provides a method of extracting an analyte from a sample. The method includes exposing an SPME sampling instrument according to the present disclosure to the analyte. As discussed above, the SPME sampling includes an extraction coating covering at least a portion of the support. Exposing the SPME sampling instrument to the analyte may include contacting the sample with the extraction coating, or placing the extraction coating in a headspace sufficiently close to the sample. The sampling may be, for example, in vitro or in vivo sampling. Contacting the sample with the extraction coating may include immersing the extraction coating in a sample fluid, or penetrating the extraction coating into a viscous or solid sample.
The method may optionally include rinsing the extraction coating after extracting the analyte from the sample. The rinsing may be performed with an aqueous solvent, or a mixture of aqueous and organic solvents. The rinsing may be performed under static or agitating conditions. The rinsing may reduce, during a subsequent analyte detection step, interferences due to loosely attached matrix components on the coating.
The analyte may be desorbed directly from the coating to an analytical instrument for detection, such as by mass spectrometry. Alternatively, the method may also include desorbing the analyte from the extraction coating, such as via a thermal- or solvent-based desorption, and separating the desorbed analytes by gas- or liquid-chromatography prior to detection. Thermal-base desorption may include exposure of the coating to a temperature of up to 300° C., such as about 250° C. Solvent-based desorption may include exposure of the coating to a solvent that would extract the analyte from the coating, without dissolving or swelling the coating polymer. Examples of suitable solvents may have pHs from about 0.01 to about 14 and may include short chain alcohols, hydrocarbons (aliphatic, cyclic, aromatic), water, chlorinated solvents, esters, ethers, nitriles, and combinations thereof.
The method may be implemented as part of a high-throughput automated sample preparation method.
An SPME coating according to the present disclosure may be used for direct microextraction of biological molecules (such as proteins or peptides) or small molecules from a biological, environmental, or food matrix, such as fluids and tissues. The small molecules may be hydrophobic or hydrophilic, and should preferably weigh less than 10,000 atomic mass units. Examples of such small molecules include contaminants, drugs, biomarkers and metabolites.
EXAMPLES Example 1 SPME Sampling Fiber Prepared by Multiple Dipping CyclesStainless steel wires of 200 μm diameter were etched by immersion in a solution of water saturated with sodium chloride and application of a voltage of 3.5 V. Before the coating procedure, the etched substrate was sonicated in a solution water/methanol 50:50 (v/v) for 10 minutes. A coating slurry was prepared by suspending 60 mg of HLB particles (5 μm diameter) in 1.5 ml of a 2.6% (w/w) solution of PTFE AF 2400 in FC-72. FC-72 has a density of 1.68 g/cm3. Accordingly, the slurry includes about 3.8 g of FC-72, about 0.10 g of PTFE AF 2400, and 60 mg of HLB particles. The ratio of PTFE AF 2400 to particles is 1.7:1 (w/w).
The slurry was deposited on the metallic support by dipping and slowly retracting the support from the slurry. Seven application cycles were used to prepare the extraction coating. After each cycle, the fiber was left at room temperature for 1 minute to allow solvent evaporation. The resulting extraction coating was about 1 cm in length and about 90 μm in thickness, and is shown in
A coating slurry was prepared as described in Example 1. A carbon mesh fabric having a unit weight of 115 g/m2, a thickness of 406 microns, and a carbon content of 99%, which was not-pretreated before coating, was immobilized on the surface of a bar-coater in order to keep the fabric straight on the surface. The immobilized fabric was coated with the slurry using the film applicator at a constant spreading speed to result in a carbon mesh fabric having a thin, homogeneous coating layer thereon. The coated fabric was left to dry for a minute in order to evaporate the FC-72 solvent. The coating volume per SPME instrument is greater with the coated fabric compared to the coated wires of Example 1 and the authors of the present disclosure expect that the coated fabric will have a lower analyte detection threshold.
Example 3 Analysis of Fruit Metabolites in an Aqueous Matrix Using PTFE AF2400/HLB Fiber and GC-MSUltrapure water was spiked with a mixture of all the GC-amenable analytes listed in Table 1, having diverse chemical functionalities and physical chemical characteristics. The water was spiked at concentrations ranging from 8.3 to 664.4 μg of compound per liter, keeping the organic solvent volume below 1% (v/v). The spiked water sample allowed for testing extraction of all the analytes simultaneously.
The samples were extracted using the SPME sampling fiber described in Example 1 by directly exposing the coating to the sample matrix for 8 hrs at a temperature of 35° C. while shaking the sample at 500 rpm.
The desorption, separation, and detection of the analytes was performed by an Agilent 6890/5973 GC-MS equipped with a GERSTEL CIS septumless PTV injector. The adsorbed analytes were thermally desorbed at 250° C. for 15 minute using ultrapure helium as a carrier gas and at a flow velocity of 1.5 ml/min. The capillary column used for the chromatographic separation was an Agilent J&W HP-5 (30 m, 0.25 mm i.d., 0.25 μm film thickness). The column temperature program was initially set at 35° C. for 6 min, ramped at 10° C./min to 140° C. then, ramped at 20° C./min to 270° C., and held at 270° C. for 2 min, resulting in a total run time of 24 min. Helium (purity level 99.999%) was used as carrier gas and its flow set at 1.2 ml/min. The mass spectrometer working conditions were: electron ionization (El) 70 eV, mass range 50-350 m/z, ion source temperature: 230° C., quadrupole temperature: 150° C., and transfer line temperature: 280° C.
The same set of experiments and analysis was performed using an SPME sampling instrument having a DVB/Carboxen/PDMS (30μm 50μm) coating in order to compare extraction efficiency (fc values) at equilibrium conditions. When an SPME coating includes adsorptive particles, the adsorption mechanism of extraction requires that the extraction phase surface concentration (Se) of adsorbed analytes is considered rather than the extraction phase concentration. Therefore, the calculation of fibre coating/sample distribution constant (Kfs) for SPME adsorbents requires the determination of Se values or the knowledge of Sa constants (since Se can be expressed as the ratio of amount extracted and the active surface of the fibre coating (Sa)). Since Sa is tedious to determine experimentally, a new constant, termed “fibre constant” (fc) representing the products Kfs*Sa for adsorptive materials, is alternatively used for the estimation of SPME enrichment factors at equilibrium.
The DVB/Carboxen/PDMS coating was obtained from Supelco Sigma Aldrich and had a 30 μm inner layer of carboxen embedded in PDMS and a 50 μm outer layer of DVB embedded in PDMS. These results are shown in Table 1 and confirm that the PTFE AF 2400/HLB coating prepared as discussed in Example 1 provides results that are comparable to, or superior to, those achieved using the DVB/Carboxen/PDMS coating. Moreover, superior reproducibility, expressed as relative standard deviation (RSD %), was obtained using the PTFE AF 2400/HLB coating as compared to the DVB/Carboxen/PDMS coating (Table 2). Reproducibility was measured by performing extractions for 30 min at 600 rpm and 30° C. Reproducibility was tested under pre-equilibrium conditions (30 min) because at shorter extraction times any error made on the extraction times reflects in higher errors in amount extracted. Reproducibility under equilibrium conditions does not vary with extraction time. Reproducibility was calculated by relative standard deviation for each analyte extracted, after a series of at least three replicate extractions, using the same coating.
A phosphate buffer (pH 7.4) was spiked with a mixture of all the LC-amenable compounds listed in Table 3. The phosphate buffer was spiked at concentrations ranging from 0.1 and 1000.0 μg of compound per liter, keeping the organic solvent volume below 1% (v/v). Codeine-D3, 6-acetylmorphine-D3 and cocaine-D3 were used as internal standards (IS) at a concentration of 50.0 μg/L.
Prior to extraction, the SPME sampling fiber described in Example 1 was conditioned for 15 min in 1.8 mL of ACN/MeOH/H2O (40:40:20, v/v/v) which was acidified to contain 0.1% (v/v) formic acid (FA), under agitation at 1200 rpm. The conditioning step was followed by a rapid rinsing in 1.8 mL of water for 2 seconds.
Extraction experiments were performed at 30 minutes in the spiked phosphate buffer using 1.8 mL of sample, under agitation at 1200 rpm. Following the extraction, a 2 second static washing step, employing 0.1 mL of water, was carried out.
The adsorbed analytes were desorbed from the fiber for 30 minutes, under agitation at 1200 rpm, using 0.1 mL of ACN/MeOH/H2O (40:40:20, v/v/v) acidified to contain 0.1% (v/v) formic acid.
In addition, using the abovementioned conditions, experiments were repeated three times in a same day and in three separate days in order to calculate intra-day and inter-day reproducibility. Moreover, using the same experimental conditions longer extraction time (660 min) was tested.
The extracted samples were run in an Exactive™ benchtop Orbitrap mass analyzer system (Thermo Scientific, San Jose, USA). Optimum conditions for each compound were determined by doing direct infusion of standards. Chromatographic separation of the analytes was achieved using a Discovery HS F5 column (100 mm×2.1 mm i.d., 3 μm; Supelco, Bellefonte, Pa., USA) using 0.3 mL/min flow rate in a ternary gradient elution with water (solvent A) and acetonitrile (solvent B) and methanol (solvent C), all acidified to contain 0.1% (v/v) formic acid. The elution employed the following gradient: the initial solvent was 5% B and 5% C and 90% A, and this solvent mixture used for the first 0.5 min of elution; over 6.5 min, the solvent mixture was raised to 50% B and 50% C; over the following 5 min, the solvent mixture was decreased to 25% B and increased to 75% C; this solvent mixture was maintained for 3.5 min; over 0.2 min the solvent mixture was returned to the initial solvent conditions; the initial solvent conditions were maintained for 2 min to ensure re-equilibration of the column before any subsequent analyte injection.
The injection volume for all standards and samples was 10 μL in full loop mode. The tray temperature of the autosampler was maintained at 5° C., the column oven compartment temperature was set at 35° C. Other conditions were set as follows: spray voltage=4000 V, vaporizer temperature=300° C., sheath gas=45 units, auxiliary gas=10, and capillary temperature=300° C. Data processing and acquisition was done using the software Xcalibur (2.0.7 SP1). Tables 3 and 4 present the figures of merit obtained for the investigated analytes in phosphate buffer solution.
The biocompatibility of the SPME coating described in Example 1 was investigated in terms of inertness to biofouling. The SPME coated fiber was exposed to consecutive adsorption/rinsing/solvent desorption cycles (which is typical of the routinely applied procedures for SPME in LC applications) using blood, urine, grape juice, human serum, and saliva as model matrices. As a quality control, extraction from PBS spiked with analytes were used. Prior to exposure to the complex matrices, 3 QC extractions from the spiked PBS were performed to show the initial performance of the coatings. The fibers were then exposed to matrices for 10 extraction/desorption cycles, and were subsequently again exposed to QC extraction in PBS in order to investigate any changes of the coating performance after matrix exposure compared to the initial performance. This cycles of PBS QC extractions and matrix extractions were performed alternately for total amount of 50 matrix exposures.
Prior to extraction, fibers were pre-conditioned in 300 μL of MeOH/ACN/H2O 40:40:20 v:v:v acidified to contain 0.1% FA. Following the pre-conditioning step, a short rinsing with 300 μL ultrapure water was applied under static conditions to remove traces of organic solvents. Following the rinsing, the matrixes were extracted using the SPME fibers for 30 min at 1500 rpm agitation. The sample volumes of matrix and PBS (as QC) were kept at 300 μL and 1500 μL, respectively. The fibers were subsequently rinsed for 10 sec using 1500 μL of ultrapure water at 1500 rpm agitation. The fibers were then desorbed in 100 μL of a solution MeOH/ACN/H2O 40:40:20 v:v:v acidified to contain 0.1% FA. The fibers were desorbed for 30 min or 15 min, for PBS or matrix extracts, respectively. Following the first desorption, the fibers were sequentially desorbed in 300 μL of a dicloromethane/MeOH 2:1 v:v solution for 15 min at 1500 rpm agitation. In case of blood, extractions were performed at 500 rpm in an orbital shaker and subsequently 3 rinsing cycles with ultrapure water were carried out. All other steps followed the abovementioned procedure.
Extracts from the PBS samples were analyzed by LC-MS/MS for the determination of the coating performances toward extraction of the target analytes.
The coating was inspected under microscope in order to identify any deposition of matrix components on the coating surface, at the beginning of the evaluation and before every QC extraction. There was no evidence of accumulation of biofouling after the first 50 extractions, suggesting that the coating would be suitable as a reusable probe for blood and animal tissues sampling.
Repetitive extraction results obtained for each matrix are summarized in Tables 5-9 for blood, saliva, serum, urine, and grape juice, respectively. After 50 extractions in each matrix, the SPME coating substantially maintained its initial extractive properties. Although the coatings sometimes showed a difference in extraction for the coating before matrix exposure and after the 5th extraction, repetitive exposures of the coating to the matrices after the 5th extraction did not show substantive changes to the performance of the coating. This suggests that coating is conditioned with the matrix in the first five exposures.
The biocompatibility of the SPME coating described in Example 1 was investigated in terms of its capability to avoid co-extraction of matrix components that may contribute to ion suppression or enhancement in LC-ESI-MS analysis. The SPME coating was investigated in terms of absolute matrix effect assessment using whole blood, grape juice, saliva, human serum, and urine as sample matrices.
Ion suppression or enhancement resulting from co-eluting constituents was investigated using nicotine, cocaine, diazepam, malathion, myclobutanyl, diazinon, fentanyl, and methadone, which are compounds that possess a wide range of polarities, as target analytes.
Prior to extraction, fibers were pre-conditioned in 500 μL of MeOH/ACN/H2O 40:40:20 v:v:v acidified to contain 0.1% FA. Following the pre-conditioning step, a short rinsing with 300 μL ultrapure water was applied under static conditions to remove traces of organic solvents. Exactions were performed for 30 min in 500 μL of sample matrix. Following extraction, the fibers were rinsed with 1.5 mL of ultrapure water for 10 second. After the rinsing step, the extracted analytes were desorbed for 30 min in 500 μL of ACN/MeOH/Water (40:40:20, v/v/v), acidified to contain 0.1% formic acid. The first desorption solution was preserved for absolute matrix effect investigation. The fiber was subjected to a second desorption step, performed using 500 μL dichloromethane/MeOH 2:1 v:v for 15 min. All steps were performed at 850 rpm agitation using manual Concept 96 system.
For the testing using whole blood, the fibers were treated with a pre-conditioning step prior to extraction. The fibers were pre-conditioned for 30 min in 500 μL of ACN/MeOH/Water 40:40:20 v:v:v, acidified to contain 0.1% formic acid. Following the pre-conditioning step, the fibers were washed for 10 seconds with ultrapure water in order to remove traces of organic solvent from the coating surface. Exactions were performed in an orbital shaker at 500 rpm for 30 min in 500 μL of sample matrix. After extraction, three sequential washing steps were performed using 500 μL ultrapure water for 10 second in a vortexer. After the third washing step, the analytes were desorbed for 30 min in 500 μL of ACN/MeOH/Water 40:40:20 v:v:v, acidified to contain 0.1% formic acid. The first desorption solution was preserved for absolute matrix effect investigation. Following the first desorption step, the fiber was subjected to a second desorption step, performed using 500 μL dichloromethane/MeOH 2:1 v:v for 15 min. The conditioning, first washing, and two desorption steps were all performed in mechanical agitator.
The above-mentioned procedures were repeated 50 times for each matrix in order to investigate the cumulative effect of any possible matrix constituent deposition and consequential absolute matrix effect.
Afterwards, the extracts were spiked to contain 10 or 100 ng/mL of nicotine, cocaine, diazepam, malathion, myclobutanyl, diazinon, fentanyl, and methadone. In parallel, as controls, aliquots of neat desorption solvents were also spiked to contain 10 or 100 ng/mL of the mixed analytes. Spiked matrix extracts and spiked neat solvents were injected to LC-MS/MS instrument and ratio of (i) the response of peak area of each analyte in the spiked extract to (ii) the response of peak area of each analyte in the spiked neat solvent, was used to calculated absolute matrix effect for each analyte.
The absolute matrix effects for each tested analyte are provided in Tables 10 to 25. The Teflon-HLB fibers discussed in Example 1 provided good matrix compatibility for all the matrices tested.
The capability of the SPME coating described in Example 1 was investigated in terms of its ability to maintain its extraction capability constant after extraction in complex matrices and thermal desorption into the GC injection port. Grape juice, serum, urine and saliva were used as model matrices.
The SPME coatings were exposed to the model matrixes for 30 minutes at 300 rpm and 35° C. The coatings were then rinsed in ultrapure water (30 sec at 500 rpm). Afterwards, desorption was performed by inserting the coating in the GC-injector port for 15 minutes at 270° C. The coatings were then washed in a mixture MeOH/H2O for 1 minute at 500 rpm. This extraction/desorption procedure was repeated for up to 50 extractions. The extraction performance of the coating was evaluated by QC extractions of known concentrations of diazepam, methadone, malathion, diazinon, cocaine, and myclobutanil from a nanopure water solution. The QC extractions were performed for 30 minutes at 500 rpm and 40° C. The coating was desorbed in the GC-injector port for 15 minutes at 270° C. The QC analyses were carried out at the beginning of the evaluation, and after every ten extraction from the sample matrices. No evidence of accumulation of biofouling was observed after the first 50 extractions.
The repetitive extraction results obtained for each matrix are summarized in Tables 26-29 for grape juice, urine, saliva, and serum, respectively.
Although some residual attachment of matrix may occur on the coating surface, the tables indicate that the coating extraction capability does not drop below 80% when exposed to the exemplary matrixes. The response can be corrected by internal standard.
In the preceding description, any discussion of a range of values should be understood to disclose all possible values within the range and all possible ranges falling with the range. For example, a discussion of “from about 1 to about 100” should be understood to be a disclosure of every value from about 1 to about 100 (for example 2, 10.7, 50, 80.5, and 92) and every range that falls between about 1 and about 100 (for example 10-20, 5-95, 75-80.5, and 24.3-47.5).
In the preceding description, for purposes of explanation, numerous details are set forth in order to provide a thorough understanding of the examples. However, it will be apparent to one skilled in the art that these specific details are not required. Accordingly, what has been described is merely illustrative of the application of the described examples and numerous modifications and variations are possible in light of the above teachings.
Since the above description provides examples, it will be appreciated that modifications and variations can be effected to the particular examples by those of skill in the art. Accordingly, the scope of the claims should not be limited by the particular examples set forth herein, but should be construed in a manner consistent with the specification as a whole.
Claims
1. A solid-phase micro-extraction (SPME) coating comprising:
- a sorptive particulate material immobilized in a fluorocarbon polymer that comprises polyvinylidene fluoride (PVDF).
2. The SPME coating according to claim 1, wherein the fluorocarbon polymerconsists of PVDF.
3. The SPME coating according claim 1, wherein the fluorocarbon polymer has carbon-fluorine (C—F) bonds and carbon-hydrogen (C—H) bonds, wherein fewer than 5% of all the C—F and C—H bonds in the fluorocarbon polymer are C—H bonds.
4. The SPME coating according to claim 1, wherein the sorptive material is a porous material having meso-, macro-, or micro-pores.
5. The SPME coating according to claim 1, wherein the sorptive material has a surface area of about 10 m2/g to about 3000 m2/g.
6. The SPME coating according to claim 5, wherein the sorptive material has a surface area of about 200 m2/g to about 800 m2/g
7. The SPME coating according to claim 1, wherein the sorptive material comprises particles, nanosheets, nanotubes, or any combination thereof.
8. The SPME coating according to claim 7, wherein the sorptive material is inorganic, organic, a hybrid inorganic/organic material, or a mixture of both inorganic and organic materials.
9. The SPME coating according to claim 1 wherein the sorptive material comprises: normal-phase silica particles, C-1/silica particles, C-4/silica particles, C-6/silica particles, C-8/silica particles, C-18/silica particles, C-30/silica particles, reverse-phase amide silica particles, HS-F5/silica particles, phenyl/silica particles, cyano/silica particles, diol/silica particles, ionic liquid/silica particles, molecular imprinted polymer particles, hydrophilic-lipophilic-balance (HLB) particles, carboxen 1006 particles, carbowax particles, divinylbenzene (DVB) particles, octadecylsilane particles, nanoparticles, processed mineral based particles, carbon nanotubes, functionalized-carbon nanotubes, graphene, graphene oxide, functionalized-graphene, quantum dots, organic polymeric particles optionally functionalized with an organic moiety, inorganic polymeric particles optionally functionalized with an organic moiety, or any combination thereof.
10. The SPME coating according to claim 9, wherein the organic moiety is a carbon chain, a strong cation moiety, a weak cation moiety, a strong anion moiety, or a weak anion moiety.
11. The SPME coating according to claim 9, wherein the sorptive material comprises hydrophilic-lipophilic-balance (HLB) particles, divinylbenzene (DVB) particles, C-18/silica particles, or any combination thereof.
12. A solid-phase micro-extraction (SPME) sample instrument comprising:
- a support; and
- an extraction coating according to claim 1 covering at least a portion of the support.
13. The SPME sample instrument according to claim 12, wherein the support is a metal support, a metal alloy support, a fused silica support, a plastic support, a fluoro-plastic support, or a carbon material support.
14. A method of making a solid-phase micro-extraction (SPME) sample instrument, the method comprising:
- applying a mixture comprising vinylidene fluoride and a sorptive material on at least a portion of a support; and
- curing the mixture to form a substantially uniform SPME coating layer on the support.
15. A method of solid-phase micro-extraction (SPME) comprising:
- exposing the SPME coating according to claim 1 to a sample matrix that comprises at least one analyte; and
- desorbing the extracted analyte.
16. The method according to claim 15, wherein the desorbing comprises exposing the SPME coating to: (i) a thermal-assisted desorption temperature, and the method optionally further comprises gas chromatography or direct coupling to a spectroscopic technique suitable for detection of a thermally stable analyte; (ii) a solvent-assisted desorption solvent, and the method optionally further comprises liquid chromatography, gas chromatograph, capillary electrophoresis, or any spectroscopic technique suitable for determination of a solvent stable analyte; or (iii) electrothermal vaporization, arc and spark ablation, laser ablation, glow discharge, matrix-assisted laser desorption/ionization (MALDI), or desorption electrospray ionization (DESI), and the method optionally further comprises a spectroscopic technique suitable for detection of the analyte.
17. The method according to claim 16, wherein the thermal-assisted desorption temperature is a temperature up to 300° C.
18. The method according to claim 16, wherein the spectroscopic technique suitable for detection of a thermally stable analyte is mass spectrometry.
19. The method according to claim 16, wherein the spectroscopic technique suitable for detection of the analyte is gas chromatography or direct coupling to mass spectrometry.
Type: Application
Filed: Dec 5, 2019
Publication Date: Apr 9, 2020
Inventor: Janusz B. PAWLISZYN (Waterloo)
Application Number: 16/704,426
