CN116710747A

CN116710747A - Precoat for biocompatible solid phase microextraction device

Info

Publication number: CN116710747A
Application number: CN202180081542.4A
Authority: CN
Inventors: Y·陈
Original assignee: Sigma Aldrich Co LLC
Current assignee: Sigma Aldrich Co LLC
Priority date: 2020-12-03
Filing date: 2021-12-02
Publication date: 2023-09-05
Also published as: CN116547489A

Abstract

An improved apparatus for solid phase microextraction includes a plastic substrate, a precoat layer on the plastic substrate, and an SPME coating layer on the precoat layer. SPME coatings are often incompatible with plastic substrates due to differences between the surface energy of the substrate and the coating, resulting in uneven coating and poor adhesion. The addition of the precoat provides increased SPME coating uniformity and stronger SPME coating adhesion to the plastic substrate. Also provided are methods of coating an SPME coating on a plastic substrate, the methods comprising the steps of pre-coating the plastic substrate to provide a pre-coated substrate and then coating the pre-coated substrate with the SPME coating, wherein the pre-coating provides improved coating uniformity and adhesion of the SPME coating to the plastic substrate.

Description

Precoat for biocompatible solid phase microextraction device

Cross Reference to Related Applications

The present application claims priority from U.S. provisional patent application Ser. No. 63/121,050, filed on 3 months 12 and 2020, 63/121,035, filed on 3 months 12 and 2020, and 63/121,071, each of which is incorporated herein by reference in its entirety.

Background

There is a need for a disposable biocompatible solid phase microextraction (BioSPME) device for use in immersion extraction. Inexpensive materials such as plastics are preferred in order to keep costs low. However, the use of inexpensive plastics (e.g. polyolefins) presents problems when applying BioSPME extract phases. Useful extraction phases, such as C18 silica in Polyacrylonitrile (PAN), do not adhere sufficiently to plastic substrates.

The surface energy of the plastic is low. For example, polypropylene has a surface energy of about 30mJ/m ² . The surface energy of the SPME coating slurry is much higher than that of the plastic, which means that the plastic surface cannot be wetted by the SPME coating slurry. In other words, without any pretreatment of the plastic surface, the SPME coating cannot be uniformly applied to the plastic surface, and the SPME coating does not firmly adhere to the plastic surface.

Traditionally, three methods can be used to treat plastics. First, mechanical methods such as sand blasting, barreling, and grinding with a power tool; second, physical methods such as flame, corona discharge, plasma; third, chemical methods, such as acid etching, anodizing. However, even though the adhesion of the coating to the plastic surface may be improved using these methods, the adhesion is still not strong, especially in the area of, for example, the tips, making these methods insufficient for use with devices having tips, such as SPME devices.

Thus, new methods of adhering coatings (e.g., biocompatible SPME coatings) to plastic surfaces are needed. Such a method should be inexpensive to keep the cost of the device low while producing strong, uniform adhesion to the plastic substrate of the device, even along the edges or tips.

Disclosure of Invention

An apparatus for Solid Phase Microextraction (SPME) is provided having a plastic substrate, a precoat layer on the plastic substrate, and an SPME coating layer on the precoat layer.

In one embodiment, an apparatus for SPME includes a plastic substrate having a first surface energy, a precoat layer on the plastic substrate, and an SPME coating layer on the precoat layer having a second surface energy, wherein the first surface energy is lower than the second surface energy, the precoat layer is firmly adhered to the plastic substrate, and the biocompatible coating layer is firmly adhered to the precoat layer.

In another embodiment, an apparatus for SPME includes a plurality of plastic pins, a precoat layer on the plastic pins, wherein the precoat layer is PAN or X18, and optionally includes particles, such as silica, titania, sodium carbonate, or a polymer resin, and an SPME coating on the precoat layer that includes a binder and a sorbent. In this embodiment, the binder is selected from the group consisting of Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane (PDMS), polyacrylate, polytetrafluoroethylene (PTFE), and polyaniline. The adsorbent is selected from the group consisting of functionalized silica, carbon, polymeric resins, and combinations thereof.

Methods for improving the adhesion of SPME coatings on plastic substrates are also provided. The method includes coating a plastic substrate with a precoat layer and then coating the precoated substrate with an SPME coating layer to provide a device, wherein the SPME coating layer adheres better to the precoated substrate than to an untreated plastic substrate. The precoat is selected from polyacrylonitrile and X18. In embodiments where the precoat is X18, the precoat may further comprise silica or other particles, such as titanium dioxide, sodium carbonate, or a polymer resin.

Drawings

FIG. 1A is a microscopic image of a pin device coated with a BioSPME coating on a silica pre-coat; fig. 1B is a microscopic image of a pin device coated with BioSPME coating over PAN precoat.

Fig. 2 is a microscopic image showing the non-uniform coating edges resulting from the direct coating of BioSPME coating on plastic substrates.

Fig. 3 is a microscopic image showing a roughened surface produced by a conventional pretreatment method.

FIG. 4A is a microscopic image showing the weak adhesion of the BioSPME coating at the tip of the pin-shaped substrate from the side; the same pin-shaped substrate is shown from the edge as in fig. 4B.

FIG. 5A is a microscopic image showing a silica pre-coat with a ratio of 7:1 (w/w); FIG. 5B shows a pre-coating of x18:silica with a 5.8:1 (w/w) ratio; FIG. 5C shows a pre-coat of x18:silica with a 5:1 (w/w) ratio; and FIG. 5D shows a pre-coating of X18: silica with a 3.5:1 (w/w) ratio.

FIG. 6A shows a 16 pin device coated with an X18:silica precoat and a PAN/C18BioSPME coating; FIG. 6B shows a 96 pin device coated with X18: silica pre-coat and PAN/C18BioSPME coat, and FIG. 6C shows 384 pins coated with X18: silica pre-coat and PAN/C18BioSPME coat.

FIG. 7 shows the relative standard deviation percentages of caffeine, carbamazepine, and tranquilization extracted from the same pin multiple times at 1000 ng/mL.

Figure 8 shows the percent of relative standard deviation of peg-to-peg for caffeine, carbamazepine, and stabilized at 1000ng/mL extracted multiple times from the same device.

Figure 9 shows the percentage of device-to-device relative standard deviation of caffeine, carbamazepine, and tranquilization extracted from multiple devices at 1000 ng/mL.

FIG. 10 is a representative chromatogram of albumin, extracted pins, and 2.5 μg/mL standard.

FIG. 11 shows TIC chromatograms of protein precipitated samples, bioSPME extracted samples, and phospholipids in desorption solutions. The chromatograms were adjusted to the same relative counts.

Figure 12 depicts the extraction step (left) to remove free analyte from plasma and buffer, and the analyte released into the desorption solution (right).

FIG. 13 shows a comparison of protein binding values between RED and SPME methods.

FIG. 14 shows representative chromatograms of phospholipids in control samples (precipitated acetonitrile proteins) and BioSPME samples.

Detailed Description

The present inventors have found that heretofore incompatible SPME coatings can satisfactorily adhere to plastic substrates by using a pre-coating to improve the compatibility of the plastic substrate with the SPME coating without the need to use conventional pre-treatment methods on the plastic substrate prior to coating. Although the terms SPME and BioSPME are used throughout this specification, the precoats described herein may also be used in other devices, including, for example, solid Phase Extraction (SPE) devices.

As previously mentioned, the surface energy of inexpensive plastics that are ideally used to produce, for example, multi-pin SPME devices is much lower than that of typical SPME coatings, resulting in incompatibility of the coatings with the substrate, as illustrated in fig. 2, which illustrates the problem of poor wettability due to the different surface energies. Some approximate surface energies are shown in the table below.

Table 1. Surface energies of materials used to make SPME devices.

Material	Surface energy (mJ/m) ² )
		Polypropylene (PP):	about 30
Polyacrylonitrile (PAN):	about 39 to about 50
		Dimethylformamide (DMF):	about 37
C18 silica:	about 73 of

Several conventional methods are known to promote adhesion of coatings to substrates having different surface energies. Such methods include mechanical methods such as sand blasting, barreling and grinding with power tools; physical methods including flame, corona discharge, and plasma; and chemical methods such as acid etching and anodization. The principle behind these methods is to increase the surface energy and contact area so that the coating slurry can be uniformly applied to the plastic surface and the coating can adhere firmly to the plastic. However, such a method creates a rough surface, as shown in fig. 3, which may result in poor adhesion, as shown by the edges of the pins in fig. 4.

As described herein, it has been found that a new approach to promote coating uniformity and adhesion is to use a pre-coat or primer layer on the substrate prior to coating with the SPME coating. The precoat layer acts as a buffer between the plastic surface and the SPME coating. It adheres strongly to the plastic substrate and provides a surface on which a top coating, such as an SPME coating, can be uniformly applied and adheres strongly, as shown in FIG. 1.

The methods provided herein allow coating of plastic substrates with SPME coatings without any conventional pretreatment steps. Some non-limiting examples of suitable plastic substrates include polyolefin, polyamide, polycarbonate, polyester, polyurethane, polyvinylchloride, polytetrafluoroethylene (PTFE), polyetheretherketone (PEEK), polysulfone, and polybutylene terephthalate substrates. In some preferred embodiments, the plastic substrate is polypropylene or polyethylene. The precoat layer may be applied directly to the untreated plastic substrate using the same coating method as used to apply the SPME or BioSPME coating. When using BioSPME coatings, the pre-coating should improve the adhesion of the SPME coating while maintaining the biocompatibility of the SPME coating. That is, the precoat must be compatible with the biological sample of interest and should not negatively interfere with the adsorptive properties of the SPME coating or otherwise cause interference in sampling or analysis. Although the terms SPME and BioSPME are used throughout this specification, the precoats described herein may also be used in other devices, including, for example, solid Phase Extraction (SPE) devices.

The methods described herein can be used to coat any plastic device useful for SPME, including, for example, fibers, blades, tubes, screens or nets, columns, and pins. As used herein, the term "pin" includes a thin sheet of plastic having a pointed end at one end. Such pins may be cylindrical, rod-shaped, conical, frustoconical, pyramidal, truncated pyramidal, rectangular, square, etc. The pins described herein preferably have a solid closed surface. When a pin is referred to as a "solid pin" or "wherein the pin is solid," it means that the surface of the pin is solid. A solid pin as defined herein may be distinguished from a design having an opening in the tip, such as may be used as a housing to hold SPE or SPME fibers, where typical metal fibers are substrates coated with SPE or SPME coatings. The surface of the pin was coated with an SPME coating. Since only the coated outer surface of the pin is in contact with the sample, it is not important that the inner surface is solid or hollow, as neither the coating nor the sample is in contact with the inner surface. The tip or tip (point) of the pin may be flat, rounded or may be a point. In some embodiments, the SPME device may include a single pin, while in other embodiments, the device may include multiple pins. Particularly preferred pin arrangements are described in co-pending international publication number WO 2019/036414, the entire contents of which are incorporated herein by reference.

Preferably, the diameter of the pin is in the range of about 0.2mm to about 5mm. In a preferred embodiment, the diameter of the pin is in the range of about 0.5mm to about 2 mm. In a particularly preferred embodiment, the diameter of the pin is about 1mm. The length of the pin may be varied, for example, to accommodate various sample volumes and hole depths. The length of the pin is preferably in the range of about 0.2mm to about 5cm. In some embodiments, the length may be about 0.5mm to about 2.5cm. In other embodiments, the length may be about 1mm to about 1cm.

The coatings, precoats, and SPME coatings described herein are applied to the ends of pins that will contact the sample of interest. In some embodiments, the pin is coated with a precoat and SPME coating for about half the length. In other embodiments, the pin is coated with a precoat and SPME coating for about one-quarter of its length. In various embodiments, the precoat and SPME coating may cover some portion of the length of one or more pins, for example, 1/10, 1/5, 1/4, 1/3, or 1/2 of the length of one or more pins. In other embodiments, the coating may be measured from the tip of the pin (i.e., the end of the pin that will contact the sample). In some embodiments, the pre-coat and coating may cover 1mm pins, in other embodiments, the pre-coat and coating may cover 1.5mm pins, and in other embodiments, the pre-coat and coating may cover 2mm pins. In an embodiment of a 1cm pin, the pre-coat and coating may cover 0.5mm, 0.6mm, 0.7mm, 0.8mm, 0.9mm, 1mm, 1.1mm, 1.2mm, 1.3mm, 1.4mm, 1.5mm, 1.6mm, 1.7mm, 1.8mm, 1.9mm, 2mm, 2.1mm, 2.2mm, 2.3mm, 2.4mm, 2.5, 2.6mm, 2.7mm, 2.8mm, 2.9mm, 3mm, 3.1mm, 3.2mm, 3.3mm, 3.4mm, 3.5mm, 3.6mm, 3.7mm, 3.8mm, 3.9mm, 4.1mm, 4.2mm, 4.3mm, 4.4mm, 4.5mm, 4.6mm, 4.7mm, 4.8mm or 5mm from the end of the pin. In other embodiments, other suitable coating coverage may be readily determined based on the length, shape, and diameter of the pin.

When the device comprises more than one pin, for example 4 pins, 8 pins, 12 pins, 24 pins, 48 pins, 96 pins, 384 pins or 1536 pins, it is preferred that the coating covers similar parts of each pin. In one embodiment, the pins of the multi-pin device are coated simultaneously using a dip coating process. In such a method, the plastic multi-pin device is first immersed in a precoat, removed and allowed to dry, then immersed in an SPME coating, removed and dried. Only the portion of the pin to be coated is contacted with the coating formulation or slurry. Such a coating method may ensure consistent coating on all pins in the device. Alternatively, other coating methods, such as spraying, may be used. In both single and multi-pin embodiments, dip coating is the most preferred method of applying the pre-coat and SPME coating to the plastic substrate/pin.

In some embodiments, when the pre-coat is applied, the plastic substrate is untreated. In a preferred embodiment, the use of a pre-coating provides better adhesion of the SPME coating than when using other pre-treatment methods (e.g., mechanical, physical, or chemical methods).

In some embodiments, the plastic substrate may be pretreated using conventional pretreatment methods prior to application of the precoat.

The precoat provided herein improves both the surface uniformity and adhesion of the SPME coating. Two particularly suitable precoats include Polyacrylonitrile (PAN) and X18, a proprietary low viscosity one-component primer available from Master Bond, inc., hackensack, NJ 07601. When the precoat is X18, silica or other solid particles (e.g., titanium dioxide, sodium carbonate, or a polymer resin) may be added to the precoat slurry to adjust the viscosity and modify the surface properties of the precoat. Although the addition of silica or other particles is not necessary, this does improve uniformity relative to X18 alone. In some embodiments, it is preferred to add silica to X18.

When the precoat is PAN, the precoat thickness may be in the range of 0.5 μm to 200 μm. In some embodiments, the PAN pre-coat thickness may be 0.4 μm, 0.5 μm, 0.6 μm, 0.7 μm, 0.8 μm, 0.9 μm, 1 μm, 1.5 μm, 2 μm, 2.5 μm, 3 μm, 3.5 μm, 4 μm, 4.5 μm, 5 μm, 5.5 μm, 6 μm, 6.5 μm, 7 μm, 7.5 μm, 8 μm, 8.5 μm, 9 μm, 9.5 μm, 10 μm, 10.5 μm, 11 μm, 11.5 μm, 12 μm, 12.5 μm, 13 μm, 13.5 μm, 14 μm, 14.5 μm, 15 μm, 15.5 μm, 16.5 μm, 17 μm, 17.5 μm, 18 μm, 19 μm, 20 μm. In still other embodiments, the PAN precoat layer thickness may be, for example, 5 μm, 10 μm, 20 μm, 25 μm, 30 μm, 35 μm, 40 μm, 45 μm, or 50 μm. In still other embodiments, the PAN precoat thickness may be, for example, 10 μm, 20 μm, 30 μm, 40 μm, 50 μm, 60 μm, 70 μm, 80 μm, 90 μm, 100 μm, 110 μm, 120 μm, 130 μm, 140 μm, 150 μm, 160 μm, 170 μm, 180 μm, 190 μm, or 200 μm. In a preferred embodiment, the thickness of the PAN precoat is in the range of 0.5 μm to 15 μm.

When the precoat is X18, X18 may be used alone or may be combined with particulate SiO ₂ 、TiO ₂ 、Na ₂ CO ₃ A solid polymer resin or other solid particles. When silica or other particles are added to the X18 precoat, the particle size may range from nano-particles to micro-particles. In a preferred embodiment, the particles used for the X18-based precoat are silica having a particle size in the range of nanoparticles to 10 μm. The type of silica or other particles added to the precoat (e.g., porosity, dispersibility, etc.) is not particularly important. Without being bound by theory, it is believed that the function of the particles in the precoat is to adjust the viscosity of the precoat slurry, adjust the surface properties of the precoat, and help improve both the uniformity of the surface and the adhesion of the SPME coating to the substrate. The silica or other particles in the precoat layer are believed to have no effect in the function of the SPME coating.

The ratio of X18 to particles is preferably greater than 3:1 (w/w). In various embodiments, the ratio of X18 to particles is in the range of 12:1 to 3:1 on a weight/weight basis. In certain embodiments, the ratio of X18 to particles (w/w) is 7:1 to 3:1. In various other embodiments, the ratio of X18 to silica (w/w) may be, for example, 11:1, 10.5:1, 10:1, 9.5:1, 9:1, 8.5:1, 8:1, 7.5:1, 7:1, 6.5:1, 6:1, 5.5:1, 5:1, 4.5:1, 4:1, 3.5:1, or 3:1. The preferred range of X18 particles (w/w) may include any of these.

In a preferred embodiment, the particles are silica. The ratio of X18 to silica is preferably greater than 3:1 (w/w). In some embodiments, the ratio of X18 to silica is 12:1 on a weight/weight basis; in various other embodiments, the ratio of X18 to silica may be (w/w) e.g. 11:1, 10.5:1, 10:1, 9.5:1, 9:1, 8.5:1, 8:1, 7.5:1, 7:1, 6.5:1, 6:1, 5.5:1, 5:1, 4.5:1, 4:1, 3.5:1 or 3:1. In some embodiments, the ratio of X18 to silica is in the range of 10:1 to 3:1 (w/w). In certain embodiments, the ratio of X18 to silica ranges from 7:1 to 3.5:1 (w/w). In a preferred embodiment, the ratio of X18 to silica is in the range of 8:1 to 5:1 (w/w).

When particles (i.e., silica, titania, sodium carbonate, or polymer resin) are added to the X18 precoat, the precoat thickness may be, for example, 0.4 μm, 0.5 μm, 0.6 μm, 0.7 μm, 0.8 μm, 0.9 μm, 1 μm, 1.5 μm, 2 μm, 2.5 μm, 3 μm, 3.5 μm, 4 μm, 4.5 μm, 5 μm, 5.5 μm, 6 μm, 6.5 μm, 7 μm, 7.5 μm, 8 μm, 8.5 μm, 9 μm, 9.5 μm, 10.5 μm, 11 μm, 11.5 μm, 12.5 μm, 13 μm, 13.5 μm, 14.5 μm, 15 μm, 15.5 μm, 16 μm, 16.5 μm, 17.5 μm, 18 μm, 19 μm, or 20 μm. In still other embodiments, the X18 and particle precoat thicknesses may be, for example, 5 μm, 10 μm, 20 μm, 25 μm, 30 μm, 35 μm, 40 μm, 45 μm, or 50 μm. In still other embodiments, the X18 and particle precoat thicknesses may be, for example, 10 μm, 20 μm, 30 μm, 40 μm, 50 μm, 60 μm, 70 μm, 80 μm, 90 μm, 100 μm, 110 μm, 120 μm, 130 μm, 140 μm, 150 μm, 160 μm, 170 μm, 180 μm, 190 μm, or 200 μm. In a preferred embodiment, the X18 and particle precoat thicknesses are preferably in the range of 0.5 μm to 15 μm.

SPME coatings are coatings useful for solid phase microextraction applications, and typically include binders and adsorbents. In some applications, the binder and the adsorbent are biocompatible. By "biocompatible" is meant that the coating is compatible with the biological sample of interest and that the biological sample does not negatively interfere with the adsorptive properties of the SPME coating or otherwise cause interference in sampling or analysis.

Some non-limiting examples of binders that can be used for SPME include Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane (PDMS), polyacrylate, polytetrafluoroethylene (PTFE), and polyaniline. For some applications, the adhesive should also be biocompatible. Particularly suitable biocompatible binders include Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, and polyamide. In a preferred embodiment, the adhesive is a biocompatible adhesive. In a particularly preferred embodiment, the biocompatible adhesive is PAN.

Adsorbents useful in the SPME devices described herein include microspheres, such as functionalized silica spheres, functionalized carbon spheres, polymer resins, mixed mode resins, and combinations thereof. In general, microspheres useful for liquid chromatography (i.e., affinity chromatography) and microspheres useful for Solid Phase Extraction (SPE) and Solid Phase Microextraction (SPME) are preferred for the coatings described herein.

In particular, the adsorbent may comprise functionalized silica microspheres, such as C18 silica (silica particles derivatized with a hydrophobic phase containing octadecyl), C8 silica (silica particles having a bonding phase containing octyl groups), RP-amide-silica (silica having a bonding phase containing palmitoylaminopropyl groups) or HS-F5-silica (silica having a bonding phase containing pentafluorophenyl-propyl groups).

Some other non-limiting examples of suitable adsorbents include: normal phase silica, C1 silica, C4 silica, C6 silica, C8 silica, C18 silica, C30 silica, phenyl/silica, cyano/silica, diol/silica, ionic liquid/silica, titan ^TM Silica (Millipore Sigma), molecularly imprinted polymer microparticles, hydrophilic-lipophilic-balance (HLB) microparticles (particularly those disclosed in co-pending U.S. patent application Ser. No. 16/640,575 published as US 2020/0197907),1006 (millipore sigma), divinylbenzene, styrene and poly (styrene-co-divinylbenzene). Mixtures of adsorbents can also be used in the coating. The adsorbents used in the coatings described herein can be inorganic (e.g. silica), organic (e.g. +. >Or divinylbenzene) or inorganicOrganic mixtures (e.g. silica and organic polymers). In a preferred embodiment, the adsorbent is C18 silica, C8 silica or mixed mode functionalized silica. In a particularly preferred embodiment, the adsorbent is C18 silica.

The diameter of the adsorbent particles or microspheres may be in the range of about 10nm to about 1 mm. In some embodiments, the spherical particles have a diameter in the range of about 20nm to about 125 μm. In certain embodiments, the diameter of the microspheres is in the range of about 30nm to about 85 μm. In some embodiments, the spherical particles have a diameter in the range of about 10nm to about 10 μm. Preferably, the spherical particles have a narrow particle size distribution.

In some embodiments, the surface area of the sorbent particles is about 10m ² /g to 1000m ² In the range of/g. In some embodiments, the porous spherical particles have a surface area of about 350m ² /g to about 675m ² In the range of/g. In some embodiments, the surface area is about 350m ² /g; in other embodiments, the surface area is about 375m ² In other embodiments, the surface area is about 400m ² /g; in other embodiments, the surface area is about 425m ² /g; in other embodiments, the surface area is about 450m ² /g; in other embodiments, the surface area is about 475m ² /g; in other embodiments, the surface area is about 500m ² /g; in other embodiments, the surface area is about 525m ² /g; in other embodiments, the surface area is about 550m ² /g; in other embodiments, the surface area is about 575m ² /g; in other embodiments, the surface area is about 600m ² /g; in other embodiments, the surface area is about 625m ² /g; in other embodiments, the surface area is about 650m ² /g; in still other embodiments, the surface area is about 675m ² /g; and in still other embodiments, a surface area of about 700m ² /g。

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In preparation for coating, a slurry of the adsorbent in the binder is prepared. The adsorbent, binder and solvent are weighed into a vessel. If desired, larger pieces or agglomerates of the adsorbent are crushed, for example with a doctor blade or mixer. The binder is dissolved in a solvent. Sonication and mixing may also be used to ensure a homogeneous distribution of particles in the binder solution. If desired, the slurry may be degassed prior to coating the substrate.

In the dip coating process, the substrate is lowered into the SPME coating slurry, then removed and allowed to dry and cure. In some embodiments, the drying step may be performed in air or under nitrogen, and may be performed at an elevated temperature. In a preferred embodiment, the drying may be performed in air or under nitrogen in a temperature and humidity controlled environment as disclosed in co-pending international patent application entitled "Drying Process for BioSPME Coatings" filed by applicant Sigma-Aldrich co.llc at month 12 of 2021. In another preferred embodiment, the coating may be treated in an immersion precipitation process as disclosed in co-pending international patent application entitled "Preparation of Solid Phase Microextraction (SPME) Coatings Using Immersion Precipitation" filed by the applicant Sigma-Aldrich Co.LLC at 12, month 2 of 2021.

The coating thickness of the SPME coating may be varied to achieve the desired properties. In various embodiments, the coating thickness may be in the range of about 0.1 μm to about 200 μm. In a preferred embodiment, the coating thickness is in the range of about 2 μm to about 50 μm. In other embodiments, the coating thickness may be, for example, about 1 μm, about 2 μm, about 3 μm, about 4 μm, about 5 μm, about 6 μm, about 7 μm, about 8 μm, about 9 μm, about 10 μm, about 15 μm, about 20 μm, about 25 μm, about 30 μm, about 35 μm, about 40 μm, about 45 μm, about 50 μm, about 55 μm, about 60 μm, about 65 μm, about 70 μm, about 75 μm, about 80 μm, about 90 μm, about 100 μm, about 110 μm, about 120 μm, about 130 μm, about 140 μm, about 150 μm, about 160 μm, about 170 μm, about 180 μm, about 190 μm, or about 200 μm. In some embodiments, the coating thickness is in the range of about 2 μm to about 50 μm, in other embodiments, the coating thickness is in the range of about 2 μm to about 40 μm, in still other embodiments, the coating thickness is in the range of about 5 μm to about 30 μm, in still other embodiments, the coating thickness is in the range of about 10 μm to about 100 μm. In a preferred embodiment, the coating thickness is in the range of about 10 μm to about 50 μm. For example, by performing the coating step multiple times, the coating thickness can be varied. For example, thinner coatings may be used when the sample size is very small or when rapid extraction equilibration is desired, however, thinner coatings may limit the amount of analyte that can be extracted. For a multi-pin arrangement, it is preferred that the coating thickness be uniform over all pins.

The embodiments described herein are particularly suitable for SPME coatings, which include biocompatible SPME coatings on plastic substrates. The SPME device may include a single plastic pin, or may include multiple plastic pins, for example on a device configured to extract multiple samples simultaneously. Such a device is particularly useful in an automatic sampling system.

In a first embodiment, an apparatus for Solid Phase Microextraction (SPME) includes a plastic substrate, a precoat layer on the plastic substrate, and an SPME coating layer on the precoat layer.

In this embodiment, some non-limiting examples of plastic substrates include polyolefin, polyamide, polycarbonate, polyester, polyurethane, polyvinylchloride, polytetrafluoroethylene (PTFE), polyetheretherketone (PEEK), polysulfone, and polybutylene terephthalate substrates. In some preferred embodiments, the plastic substrate is polypropylene or polyethylene.

In some embodiments, the pre-coat comprises Polyacrylonitrile (PAN). In some embodiments, the pre-coat is PAN. When the precoat comprises PAN or when the precoat is PAN, the thickness of the precoat is preferably in the range of 0.5 to 200 microns. In some embodiments, the thickness of the precoat layer is in the range of 0.5 microns to 50 microns. In still other embodiments, the thickness of the precoat layer is in the range of 0.5 microns to 20 microns. In a particularly preferred embodiment, the thickness of the precoat layer is in the range of 0.5 microns to 15 microns.

In other embodiments, the pre-coat comprises X18. In some embodiments, when the pre-coat layer comprises X18, it further comprises particles selected from silica, titania, sodium carbonate, a polymer resin, or a combination thereof. Particles may be added to X18 to alter the viscosity of the precoat slurry, adjust the surface properties of the precoat, improve coating uniformity, and improve adhesion of the SPME coating to the substrate. The size of the silica or other particles may be in the range of nanoparticles or microparticles. In a preferred embodiment, the silica or other particles have a diameter of 10 microns or less. In some embodiments, the pre-coat is a combination of X18 and silica. In other embodiments, the pre-coat is a combination of X18 and titanium dioxide, sodium carbonate, or a polymer resin. In a preferred embodiment, the precoat is X18 and silica. When the precoat layer includes X18 and silica, or when the precoat layer is X18 and silica, the thickness of the precoat layer is preferably in the range of 0.5 microns to 200 microns. In some embodiments, the thickness of the precoat layer containing X18 and silica is in the range of 0.5 microns to 50 microns. In still other embodiments, the thickness of the precoat layer comprising X18 and silica is in the range of 0.5 microns to 20 microns. In a particularly preferred embodiment, the thickness of the precoat layer containing X18 and silica is in the range of 0.5 microns to 15 microns.

When the pre-coat layer comprises both X18 and silica, the weight ratio of X18 to silica (X18: silica (w/w)) is preferably greater than 3:1. In some embodiments, the ratio of X18 to silica is greater than 5:1 (w/w). In still other embodiments, the ratio of X18 to silica is in the range of 10:1 to 3:1 (w/w). In still other embodiments, the ratio of X18 to silica is in the range of 8:1 to 5:1 (w/w). It will be appreciated that other particles, such as titanium dioxide, sodium carbonate or polymer resins, may be added in the same ratio.

In this first embodiment, the SPME coating or BioSPME coating includes a binder and an adsorbent. Some non-limiting examples of binders that may be used in this embodiment include Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane (PDMS), polyacrylate, polytetrafluoroethylene (PTFE), polyaniline, and combinations thereof. In a particularly preferred embodiment, the binder is PAN.

The adsorbent in this embodiment includes any adsorbent that can be used in SPME or BioSPME. Such adsorbents include functionalized silica, carbon, polymeric resins, and combinations thereof. Many suitable adsorbents are discussed above. Some preferred silica adsorbents include C18 silica, C8 silica, and mixed mode functionalized silica. Some preferred polymeric resins include HLB resins, divinylbenzene resins, styrene resins, poly (styrene-co-divinylbenzene) resins, and combinations thereof.

In this first embodiment, the plastic substrate is pin-shaped. Preferably, the pin is a solid pin. The length of the pin may be any length suitable for the device. The coatings (i.e., precoat and SPME coatings) are applied to the tip of the substrate (i.e., the portion of the substrate or the portion of the pin that will contact the sample to be analyzed). In some embodiments, the device includes a plurality of pins, allowing for simultaneous sampling of a plurality of different samples. Such a multi-pin device is particularly suitable for interfacing with an automatic sampling system.

In a second embodiment, an apparatus for SPME includes a plastic substrate having a first surface energy, a precoat layer on the plastic substrate, and an SPME coating layer on the precoat layer having a second surface energy, wherein the first surface energy is lower than the second surface energy. In this embodiment, the pre-coat coats the plastic substrate, thus providing a more compatible surface energy with the SPME coating, allowing the SPME coating that is otherwise incompatible with the plastic substrate to be uniformly coated on the plastic substrate and firmly adhered to the plastic substrate.

In preferred embodiments, the plastic substrate is a polyolefin, while in other embodiments, the plastic substrate may be a polyamide, polycarbonate, polyester, polyurethane, polyvinyl chloride, polytetrafluoroethylene, polyetheretherketone, polysulfone, or polybutylene terephthalate.

In this embodiment, the precoat may comprise PAN or X18. In some embodiments, the pre-coat is PAN or X18. In embodiments where the precoat comprises X18 or X18, the precoat may further comprise silica. As with the first embodiment, when the pre-coat layer comprises silica, the size of the silica particles may be in the nanoparticle or microparticle range. In a preferred embodiment, the silica particles have a diameter of 10 microns or less.

The thickness of the precoat layer is preferably in the range of 0.5 to 200 microns except when the precoat layer is X18 and does not include silica. In some embodiments, the thickness of the precoat layer is in the range of 0.5 microns to 50 microns. In still other embodiments, the thickness of the precoat layer is in the range of 0.5 microns to 20 microns. In a particularly preferred embodiment, the thickness of the precoat layer is in the range of 0.5 microns to 15 microns.

As in the previous embodiments, the SPME coating includes a binder and an adsorbent. The binder may be selected from Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane, polyacrylate, polytetrafluoroethylene, and polyaniline. The adsorbent may be selected from the group consisting of functionalized silica, carbon, polymeric resins, and combinations thereof. In a preferred embodiment, the plastic substrate is polypropylene, the binder is PAN, and the adsorbent is functionalized silica.

A third embodiment provided by the present invention is an apparatus for SPME, where the apparatus includes a plurality of pins, for example 4 pins, 8 pins, 12 pins, 24 pins, 48 pins, 96 pins, 384 pins, or 1536 pins. The pins of the multi-pin device may be made of polyolefin, polyamide, polycarbonate, polyester, polyurethane, polyvinyl chloride, polytetrafluoroethylene, polyetheretherketone, polysulfone, and polybutylene terephthalate. In a preferred embodiment, the pin is a solid plastic pin. In another preferred embodiment, the pin is a solid polypropylene pin.

In this embodiment, each pin in the device comprises a precoat layer, the precoat layer comprising PAN or X18, and when the precoat layer comprises X18, it may further comprise particles, such as silica, titania, sodium carbonate, or a polymer resin. In some embodiments, the pre-coat is PAN or X18, optionally comprising silica. When the precoat layer includes X18 or X18 and further includes particles (e.g., silica), the size of the silica or other particles may be in the nanoparticle or microparticle range. In a preferred embodiment, the particles have a diameter of 10 microns or less.

The thickness of the precoat layer is preferably in the range of 0.5 microns to 200 microns except when the precoat layer is X18 and does not include silica or other particles. In some embodiments, the thickness of the precoat layer is in the range of 0.5 microns to 50 microns. In still other embodiments, the thickness of the precoat layer is in the range of 0.5 microns to 20 microns. In a particularly preferred embodiment, the thickness of the precoat layer is in the range of 0.5 microns to 15 microns.

The SPME coating on the precoat on each pin includes a binder and a sorbent. Suitable binders include Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane, polyacrylate, polytetrafluoroethylene, and polyaniline. Suitable adsorbents include functionalized silica, carbon, polymeric resins, and combinations thereof.

Methods for improving the adhesion of SPME coatings on plastic substrates are also provided. According to the method, a plastic substrate (e.g., one or more pins) is coated with a pre-coat to provide a pre-coated substrate, which is then coated with an SPME coating. According to this method, the adhesion of the SPME coating to the pre-coated substrate is better than its adhesion to the untreated plastic substrate.

The method is applicable to a variety of plastic substrates that may be used in SPME, including but not limited to polyolefins, polyamides, polycarbonates, polyesters, polyurethanes, polyvinylchlorides, polytetrafluoroethylene, polyetheretherketones, polysulfones, and polyterephthalates. In a preferred embodiment, the plastic substrate is a polyolefin, such as polypropylene or polyethylene. In a particularly preferred embodiment, the plastic substrate is one or more pins, for example in a multi-pin device. In some embodiments, a plastic substrate is used without any pretreatment. In other embodiments, the substrate may be subjected to mechanical, physical, or chemical pretreatment prior to coating with the precoat.

The precoat according to the method preferably comprises Polyacrylonitrile (PAN) or X18. In some embodiments, the pre-coat is PAN or X18. In embodiments where the precoat comprises X18, the precoat may further comprise particles, such as silica, titania, sodium carbonate, or a polymer resin. When the precoat layer includes X18 or X18 and further includes particles, the size of the particles may be in the range of nanoparticles or microparticles. In a preferred embodiment, the particles have a diameter of 10 microns or less.

According to this method, the SPME coating includes a binder and an adsorbent. Suitable binders for this process include Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane, polyacrylate, polytetrafluoroethylene, and polyaniline, and suitable adsorbents include functionalized silica, carbon, polymeric resins, and combinations thereof.

In a preferred embodiment, the precoat is prepared as a slurry and coated onto a plastic substrate by dip coating, which is then dried. SPME coatings were also prepared as slurries. The pre-coated substrate was then coated again with SPME coating by dip coating. In other embodiments, other coating techniques (e.g., spray coating) may be used for one or both of the pre-coating and SPME coating steps. The coated SPME device may then be dried, cured or otherwise processed in a conventional manner.

Pre-coating procedure with X18 of particles. The particles (silica, titania, sodium carbonate or polymer resin) and X18 are weighed into a vessel and added to a solvent. If desired, the particles may be broken up with a scraper, for example, due to agglomeration of the particles. The mixture is sonicated for a sufficient time to form a homogeneous slurry. After sonication, the slurry was mixed for an additional period of time, then degassed in an sonicator and cooled to room temperature. The X18/particle slurry was mixed until ready for use. The substrate is coated by dip coating or other coating method, and then the pre-coat is cured. The pre-coat may be cured by heating (e.g., by heating to 110 ℃ for 1-4 minutes) and then cooling the coated cured substrate to room temperature.

Precoating for PAN. PAN and a suitable solvent (e.g. DMF) are weighed into a container. Any larger pieces of PAN may be broken into small pieces using a doctor blade or mixer. The PAN-solvent mixture is heated to dissolve PAN. As with the X18/silica precoat, the substrate is coated by dip coating or other coating method, and the precoat is then cured. The pre-coat may be cured by heating (e.g., by heating to 110 ℃ for 1-4 minutes) and then cooling the coated cured substrate to room temperature.

Application procedure of PAN/C18 BioSPME coating. PAN and a suitable solvent (e.g. DMF) are weighed into a container. Any large pieces of PAN are broken into smaller pieces. The solution was heated to dissolve PAN. The adsorbent was weighed out, added to the PAN solution, and thoroughly mixed. The slurry was sonicated to prepare a homogeneous slurry. The slurry may be degassed and should be mixed until ready for coating. For coating, the pre-coated substrate is dip coated, removed and cured. For the comparative example, the SPME coating was dip coated directly onto a plastic substrate.

The coating was visually observed using a microscope. The coatings were tested for asperity (ruggeddess) and adhesion as follows: (a) By finger rubbing over the cured coating, and (b) by blue tape adhesion test. The blue tape adhesion test was performed as follows: a blue paint tape (medium tack) was applied to the coated cured SPME device and allowed to stand in place for 90 seconds, then the tape was removed at an angle of 180 degrees relative to the device. Adhesion was observed visually using a microscope.

The precoat described herein shows improved uniformity and adhesion of the SPME coating on plastic substrates, providing improved results without any loss of biocompatibility. As shown in fig. 2, conventional PAN/C18 SPME applied directly onto the plastic pins via dip coating resulted in edge non-uniformity due to poor wettability caused by the surface energy difference between the plastic substrate and the coating. The weak adhesion of the PAN/C18 coating to the plastic substrate without the precoat is further shown in fig. 4, showing both the sides and tips of the coated pin.

Advantageously, the devices provided herein maintain a high level of biocompatibility, as will be more fully demonstrated in the examples below.

Examples

Example 1. A precoat slurry was prepared using 33g of x18, 7g of silica and 11g of mesitylene. Conventional PAN/C18 BioSPME coating slurries were prepared. The polypropylene multi-pin SPME device was coated as follows. A thin layer of precoat was formed on the pin by dip-coating the pin in the precoat slurry and drying at 110 ℃ for 4 minutes. After cooling the pre-coated pin device to room temperature, the PAN/C18SPME coating was dip coated onto the pre-coated pin tool and dried. A uniform thin layer of PAN/C18 is formed on the pins.

Example 2. A PAN pre-coat slurry was prepared by dissolving 5g PAN in 35g DMF. Conventional PAN/C18 BioSPME coating slurries were prepared. The polypropylene multi-pin SPME device was coated as follows. A thin layer of precoat was formed on the pin by dip-coating the pin in the precoat slurry and drying at 110 ℃ for 4 minutes. After cooling the pre-coated pins to room temperature, the PAN/C18SPME coating was dip coated onto the pre-coated pin tool and dried. A uniform thin layer of PAN/C18 is formed on the pins.

The PAN/C18SPME coatings in examples 1 and 2 were uniformly coated and firmly adhered to the plastic substrate as shown in fig. 1.

Example 3. Comparative example. The polypropylene multi-pin device is subjected to a plasma treatment. (AST Products, inc., billerica, MA). Contact angle of plasma treated PP: 60-80 (ref: 90-120). Conventional PAN/C18 SPME coatings were dip coated onto pre-coated pin tools and dried. A thin layer of PAN/C18 is formed on the pins. No significant improvement in the adhesion of PAN/C18 on PP surfaces was observed after plasma treatment compared to untreated PP substrates.

Example 4. Preparation of silica pre-coat slurry at the following X18: silica ratio: (A) 7:1, (B) 5.8:1, (C) 5:1, and (D) 3.5:1, are (w/w). The image of each is shown in fig. 5.

Example 5 the multi-pin device (A) 16 pins, (B) 96 pins, and (C) 384 pins shown in FIG. 6 were pre-coated with X18: silica pre-coat and PAN/C18 BioSPME coat.

Example 6 analytical method validation and device reproducibility. 16 pin devices were prepared with X18/silica precoat and PAN/C18 BioSPME coating. Caffeine, carbamazepine and diazepam were extracted multiple times at 1000ng/mL per pin. Carbamazepine and stable individual% RSD less than 5%. The% RSD of caffeine is more variable and ranges from 2.7% to 16.3%. The results are shown in fig. 7.

Example 7. 16 pin devices were prepared with an X18/silica precoat and a PAN/C18 BioSPME coating. Caffeine, carbamazepine and diazepam were extracted from the same apparatus multiple times at 1000 ng/mL. Carbamazepine and stable% RSD less than 5% and caffeine% RSD less than 12%. The results are shown in fig. 8.

Example 8. Three multi-pin devices were prepared with an X18/silica precoat and a PAN/C18 BioSPME coating. Caffeine, carbamazepine and diazepam were extracted at 1000ng/mL using multiple devices and compared to relative standard deviations. The RSD percentages were consistent among the three devices and less than 5% precision in the carbamazepine and stable devices. The inter-device accuracy indicates that caffeine is similar to the% RSD of carbamazepine, but that the stable% RSD is slightly higher than the in-device accuracy. The results are shown in fig. 9.

Example 9 biocompatibility testing. The effectiveness of the pin tool with the precoat was checked. 96 pin devices were prepared with X18/silica precoat and PAN/C18 BioSPME coating.

To determine the amount of phospholipids remaining in the samples after extraction using the 96 pin tool, they were compared to the phospholipids remaining after acetonitrile-assisted protein precipitation. Briefly, the pin tool was conditioned in isopropanol followed by a brief rinse in water. At this point, the pin tool is ready for extraction. After extraction, the pin tool is briefly rinsed to remove any protein that may remain on the pin surface before the analyte is desorbed, and then ready for analysis.

Protein precipitation was performed by using 100 μl human plasma and mixing with 300 μl acetonitrile. The mixture was stored at 4℃for 20 minutes and then centrifuged at 5,000rpm for 10 minutes. The supernatant was transferred and dried under a stream of nitrogen at 45℃under 10 PSI. The sample was then resuspended in 200 μl of the starting mobile phase.

Five samples from both methods were analyzed on an AB Sciex-3200Q Trap mass spectrometer with Agilent 1290LC using the methods described in Table 2. The phospholipids monitored are listed in table 3.

TABLE 2 LC-MS/MS conditions for monitoring phospholipids

Table 3. Phospholipids monitored.

Analyte(s)	Precursor(s)	Product(s)	Residence time (millisecond)	DP	CE
						Choline choline	184.1	104.1	40	120	80
LPC 16:0	496.4	184.1	40	120	80
						LPC 18:0	524.4	184.1	40	120	80
PC 30:1	704.4	184.1	40	120	80
						PC 34:2	758.4	184.1	40	120	80
PC 36:2	786.4	184.1	40	120	80
						PC 38:6	806.4	184.1	40	120	80
LPC 18:2	520.4	184.1	40	120	80
						LPC 18:1	522.4	184.1	40	120	80
PC 36:1	788.4	184.1	40	120	80
						PC 38:5	804.4	184.1	40	120	80
PC 34:1	760.4	184.1	40	120	80
						PC 36:3	784.4	184.1	40	120	80

LPC-lysophosphatidylcholine

PC-phosphatidylcholine

Protein (albumin) is removed. Using NanoOrange ^TM The kit determines the amount of protein retained in the extracted sample via non-specific retention on the pin. The pins (eight) were conditioned in 800 μl isopropanol in an orifice plate under static conditions for 15 minutes. The pin was then washed in 800 μl of water for 10 seconds. Extraction of pooled human plasma (800 μl) from 96-well plates occurred while shaking with a thermal adapter at 1200rpm at 37 ℃ setting. After extraction, the pins were washed in water for 1 minute.

As described in the product specification, 1mL of working solution (dye for protein staining) was added to the appropriate wells of the well plate to prepare another well plate. The pin for BSA extraction was exposed to the working solution and allowed to react at 90-96 ℃ for 10 minutes while shaking at 300 rpm. The well plate is covered with foil to protect the sample from light. The sample was then cooled to room temperature.

Samples were analyzed on Thermo Scientific Dionex HLPC using a fluorescence detector with direct flow (no column), see table 4. The samples were quantified using peak heights, using an external calibration in the range of 0.1-5.0 μg/mL BSA.

TABLE 4 LC fluorescence conditions for monitoring fluorescence signals from labeled proteins

Overall sample cleanliness. The cleanliness of the samples was determined by collecting the TICs of the three conditions. These conditions were a control of 80:20 desorption solution, an extracted spiked plasma sample, and an acetonitrile protein precipitation sample.

Acetonitrile precipitated samples were prepared as follows. The remaining spiked plasma, corresponding to the plasma used in the extracted samples, was diluted with acetonitrile 3 x. The sample was then centrifuged at 10,000rpm for 10 minutes at 4 ℃. After completion, the supernatant was removed and dried at 10PSI under nitrogen and resuspended in desorption solution to keep solvent effects to a minimum and better reflect sample cleanliness. All three samples were analyzed as described in Table 2, with Q1 scan between 100-900m/z using 2. Mu.L injection. Multiple methanol injections were performed for each sample of interest to remove any entrainment between the samples.

As a result. The amount of phospholipids in the BioSPME prepared samples was compared to the amount of phospholipids in the acetonitrile-assisted protein precipitation prepared samples. Less than 0.1% of the phospholipids remained in the final extracted samples from BioSPME compared to the acetonitrile protein precipitated control. Sample chromatograms comparing the two conditions are shown in fig. 14.

According to NanoOrange ^TM The pin tool was studied to accumulate about 1.2 μg of protein on the pin surface, quantified as BSA. A representative chromatogram from the pin can be seen in fig. 10, compared to the calibrator. Albumin accounts for half of the total protein in plasma (between 35mg/mL and 50 mg/mL) (Merlot, a., kalinowski, d.,&Richardson,D.(2014).Unraveling the mysteries of serum albumin-more than just a serum protein (elucidating mystery of serum albumin-not just serum proteins). Frontier in Physiology,1-7, https:// doi.org/10.3389/fphys.2014.00299). This value relates to less than 0.01% protein in the final extracted sample in the 8 pins tested.

To demonstrate that the samples proved cleaner compared to standard preparations, protein precipitated plasma samples, bioSPME extracted plasma samples and complete TIC of the desorption solution were collected (see fig. 11). As can be seen, bioSPME extracted samples were significantly closer to the desorption solution than acetonitrile precipitated samples. The peaks observed in the desorption solution correspond to the deuterated carbamazepine present.

Example 10 protein binding by BioSPME was studied. 96 pin devices were prepared with X18/silica precoat and PAN/C18 BioSPME coating. Human plasma and buffer were spiked at therapeutically relevant concentrations and incubated at 37℃for 1 hour while shaking at 300 rpm. After incubation, 200 μl of plasma and buffer were loaded into separate columns onto extraction well plates (n=8). Protein binding was determined by automated robotic methods using BioSPME C18 pin tools. Briefly, the pin tool was statically conditioned in isopropanol for 20 minutes, then transferred to a new well plate in water for 10 seconds (washing step). This is followed by an extraction step. The pin tool is transferred to the preloaded extractor plate described previously. Here, the pin tool extracts the analyte while shaking at 1200-1250rpm for 15 minutes at 37 ℃. The pin tool was returned to the aqueous solution for 60 seconds and finally transferred to the stripper plate. The desorption liquid is 80:20 methanol/water, and the desorption is carried out for 20 minutes under static conditions. Samples were analyzed using the methods described in tables 5 and 6.

Extraction plates used in this study included both plastic and glass coated plates. The choice of plate depends on the nature of the compound and how well the compound behaves in the buffer solution. More hydrophobic compounds (e.g., ketoconazole and imipramine) were found to exhibit non-specific binding to plastics and extraction efficiency from glass coated 96-well plates was better. Extraction of erythromycin and propranolol was also performed from glass coated plates, as higher extraction efficiency values were obtained from glass compared to extraction from plastic plates.

Determination of protein binding by Rapid Equilibrium Dialysis (RED)

RED was performed as directed in the specification. Briefly, 200 μl of therapeutically relevant concentrations of "spiked" human plasma and 400 μl of Phosphate Buffered Saline (PBS) were loaded into the respective chambers in at least triplicate. Dialysis was performed for at least 4 hours while covering and shaking on an Eppendorf shaker at 300rpm and 37 ℃. At the end of dialysis, 50 μl of spiked plasma was mixed with 50 μl of clean (undoped) PBS, and 50 μl of dialysate (buffer chamber) was mixed with 50 μl of clean plasma. This is done to ensure matrix consistency. Next, 300. Mu.L of ice-cold acetonitrile was added to each sample, followed by centrifugation at 5,000rpm at 4℃for 10 minutes. Finally, the supernatant was transferred to a glass vial for analysis by LC-MS/MS, as described in tables 5 and 6, using AB Sciex 6500 and Agilent 1290LC, using matrix-matched external calibration (in desorption solution).

Table 5 LC-MS/MS conditions for monitoring analytes for free fraction determination.

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Removal of phospholipids (matrixing)

To determine the amount of phospholipids remaining between the different methods, samples treated by flash equilibrium dialysis or BioSPME, respectively, were compared to the phospholipids remaining by acetonitrile-assisted protein precipitation. Protein precipitation was performed by using 100 μl human plasma and mixing with 300 μl acetonitrile. The mixture was stored at 4℃for 20 minutes and then centrifuged at 5,000rpm for 10 minutes. The supernatant was transferred and dried under a stream of nitrogen at 45℃under 10 PSI. The sample was then resuspended in 200 μl of the starting mobile phase.

Five samples from the three methods were analyzed on an AB Sciex-3200Q Trap mass spectrometer with Agilent 1290LC using the methods described in Table 7. The phospholipids monitored are listed in table 8.

TABLE 7 LC-MS/MS conditions for monitoring phospholipids

TABLE 8 Phospholipids monitored

LPC-lysophosphatidylcholine

PC-phosphatidylcholine

% free fraction was determined by BioSPME (F _U )

The BioSPME method determines the free concentration of analyte in plasma by comparing it to the analyte extracted from the buffer sample, where 100% of the analyte is considered to be free of protein binding.

The free or unbound percentages are determined in equation 1:

equation 1./>

Where the free concentration represents the unbound concentration of the analyte in the matrix (in this case, plasma) and the total concentration represents the total concentration of the analyte. The amount of extraction is unit-independent and preferred amounts (e.g. nanograms or moles) M can be used _{Free form} And extraction volume of plasma V _{Plasma of blood} To be applied. The concentration of analyte in the desorption solution is quantified by an external calibration curve and if the desorption volume is equal to the plasma and buffer extraction volume, the concentration from the desorption will be equal to the concentration of the extraction, as shown in equation 2.

Equation 2.

Equation 3.

Binding score F _B Can be determined from the concentration of the extraction as shown in equation 6.

Equation 4. Bond score (F _B ) =100% -free fraction (F _U )

Equation 5.

Equation 6.

Equation 7.

Fig. 12 depicts the extraction step (left) to remove free analyte from plasma (pink) and buffer (blue) and the analyte released into the desorption solution (right). The amount of extraction does not greatly affect the concentration of free analyte, which is referred to as non-depleting. Since the buffer solution is considered to be 100% free, bioSPME will extract more from the buffer than from the plasma.

When consumption of compounds from plasma is significant (extraction over 5% of total spiked analyte) at BioSPME extraction, correction to calculated binding scores is required as follows:

equation 8

Wherein B and P represent the amounts extracted from buffer B and plasma P, respectively. B (B) ⁰ Indicating the concentration of the initially spiked sample. Equation 8 considers the concentration in the solution after extraction on the fiber; analytes from samplesIs not limited. Equations 6 and 7 do not take this into account. However, when the extraction amount is less than 5%, they provide accurate values.

Comparison of RED versus BioSPME

Using equations (equations 5 and 6), the values of analyte-protein binding in table 9 were determined by BioSPME extraction. These values are very consistent with the values determined using a rapid equilibrium dialysis device (RED) and reported literature values. These values are compared graphically in fig. 13.

In addition, table 10 shows the amount of phospholipids remaining compared to standard protein precipitation methods. A chromatogram of the BioSPME sample versus the acetonitrile protein precipitated sample is shown in fig. 14. As shown in table 10, bioSPME removed more than 99% of the phospholipids in the processed samples. This is in sharp contrast to RED devices which leave about 50% phospholipid. The amount is reduced compared to the representative value as explained by dilution of the centrifuged sample with clean buffer or plasma according to the compartment being tested.

BioSPME technology also provides time savings of over 50%, as shown in table 11, the longest step in BioSPME method is initial incubation of analyte with plasma (60 minutes). This is much shorter than the minimum 4 hour incubation time required for RED devices.

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FIG. 13 shows a comparison of protein binding values between RED and SPME methods. Blue lines indicate protein binding literature value intervals. At physiological pH, the asterisked compound is charged.

TABLE 10 phospholipids retained in analytes (per method)

Method	Sample #	Average% of remaining phospholipids	RSD
				BioSPME	5	<0.1	<0.01
Fast equilibrium dialysis	5	56	7.8

TABLE 11 comparison of time requirements by method

RED method	Step time (minutes)	BioSPME method	Step time (minutes)
				Preparation of samples	5	Preparation and incubation of samples	60
Dialysis	240	Conditioning	20
				Post sample preparation	5	Washing	0.2
Centrifuging	10	Extraction	15
				Transfer to vials for analysis	5	Washing	1
		Desorption of	15
				Totals to	265	Totals to	111.2

The BioSPME C18 technique provides a 50% time savings for protein binding assays when compared to the Rapid Equilibrium Dialysis (RED) method, and it is used via a fully automated robotic method. The BioSPME protein binding values were well compared to those of the rapid equilibrium dialysis method, as demonstrated with these 10 compounds with log P in the range of 1-5. In addition, bioSPME also provided cleaner samples compared to samples from RED devices.

The examples included herein are for illustrative purposes only and are not intended to limit the scope of the invention, which is defined by the claims.

Claims

1. An apparatus for Solid Phase Microextraction (SPME) comprising

A plastic substrate having a surface with a plurality of holes,

a precoat layer on said plastic substrate, and

SPME coating on the precoat layer.

2. The device of claim 1, wherein the plastic substrate is selected from the group consisting of polyolefin, polyamide, polycarbonate, polyester, polyurethane, polyvinylchloride, polytetrafluoroethylene (PTFE), polyetheretherketone (PEEK), polysulfone, and polybutylene terephthalate.

3. The device of any one of claims 1 or 2, wherein the plastic substrate is selected from the group consisting of polypropylene and polyethylene.

4. The device of any one of claims 1-3, wherein the pre-coat comprises Polyacrylonitrile (PAN).

5. The device of any one of claims 1-3, wherein the pre-coat comprises X18 and optionally particles selected from the group consisting of silica, titania, sodium carbonate, polymer resins, and combinations thereof.

6. The device of claim 5, wherein the pre-coating comprises X18 and particles, preferably silica particles, the particle size of the particles being in the range of 1 nm to 10 microns, and X18: particles being in the range of 8:1 to 3:1 (w/w).

7. The device of any one of claims 1-6, wherein the SPME coating comprises a binder selected from the group consisting of Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane (PDMS), polyacrylate, polytetrafluoroethylene (PTFE), and polyaniline, and

an adsorbent selected from the group consisting of functionalized silica, carbon, polymeric resins, and combinations thereof.

8. The device of claim 7, wherein the adsorbent is selected from the group consisting of C18 silica, C8 silica, mixed mode functionalized silica, HLB resin, divinylbenzene resin, styrene resin, poly (styrene-co-divinylbenzene) resin, and combinations thereof.

9. The device of any one of claims 1-8, wherein the plastic substrate comprises one or more pins.

10. The device of claim 8, wherein the one or more pins are solid.

11. The device of any one of claims 1-10, wherein

The plastic substrate has a first surface energy and,

the SPME coating has a second surface energy that is higher than the surface energy of the plastic substrate,

wherein the pre-coat layer is firmly adhered to the plastic substrate, and

Wherein the SPME coating adheres strongly to the precoat layer.

12. A method for improving the adhesion of SPME coatings on plastic substrates, the method comprising the steps of:

a plastic substrate is provided and is provided with a plastic substrate,

coating the substrate with a pre-coating layer to provide a pre-coated substrate, and

coating the pre-coated substrate with an SPME coating,

wherein the pre-coat is selected from the group consisting of polyacrylonitrile and X18; wherein if the precoat is X18, the precoat may further comprise particles selected from the group consisting of silica, titania, sodium carbonate, a polymer resin, and combinations thereof, and

wherein the SPME coating adheres better to the pre-coated substrate than to an untreated plastic substrate.

13. The method of claim 12, wherein the plastic substrate is selected from the group consisting of polyolefin, polyamide, polycarbonate, polyester, polyurethane, polyvinylchloride, polytetrafluoroethylene (PTFE), polyetheretherketone (PEEK), polysulfone, and polybutylene terephthalate.

14. The method of any one of claims 12 or 13, wherein

The binder is selected from Polyacrylonitrile (PAN), polyethylene glycol (PEG), polypyrrole, derivatized cellulose, polysulfone, polyacrylamide, polyamide, polydimethylsiloxane (PDMS), polyacrylate, polytetrafluoroethylene (PTFE), and polyaniline, and

The adsorbent is selected from the group consisting of functionalized silica, carbon, polymeric resins, and combinations thereof.

15. The method of claim 14, wherein the binder is PAN and the adsorbent is C18 functionalized silica.