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The aim in this paper was to design an experiment using LC-MS/MS that had enough sensitivity to be able to detect THC-COOH. The approach chosen was a chemical derivatization using isotope dilution LC-MS/MS (IDMS) with ESI in positive mode  .
Drug standards of THC and THCA were used, with internal standards of THC-d3 and THCA-d3. These were chosen as they are isotopes of THC and THCA, but are deuterated so have a slightly higher mass number, so when detected on the mass spectrum they have a peak slightly higher than THC and THCA. The samples were pretreated as described1.
A stock solution of methanol with the dissolved compounds was made up to the concentration of 1 Î¼g/mL, and further diluted with methanol to the required concentrations. A 250 Î¼L sample for negative control, positive control and authentic specimens were mixed with the internal standards, at 100 pg/mL for THC-d3 and 25 pg/mL for THCA-d3. Samples were votexed for 10 seconds, 500 Î¼L acetonitrile added, vortexed for 30 seconds and then centrifuged at 14,000 rpm for 10 minutes. The supernatant was then evaporated to dryness under nitrogen at 45Â°C. This was dissolved in 100 Î¼L sodium bicarbonate buffer (pH made to 10.5) and vortexed for 60 seconds. 100 Î¼L 5-(dimethylamino)-1-naphthalenesulfonyl chloride solution (1 mg/mL in actone) was added to each sample and vortex-mixed for 60 seconds. The mixture was extracted twice with 500 Î¼L of hexane for 3 minutes and evaporated to dryness. The sample was reformed in 50 Î¼L of methanol an 20 Î¼L of this was used to inject into the LC-MS/MS system. Oral fluid samples taken from drug-free volunteers were added to 100 pg/mL THC and 20 pg/mL THCA and used for the positive control and method development1. WHY HEXANE
Dansylated drug standards (DC-THC and DC-THCA) and dansylated deuterated internal standards (DC-THC-d3 and DC-THCA-d3) were made from 100 ng of the original internal standards. They were danslyated, extracted with hexane and evaporated to dryness. The samples were reformed in 1 mL of methanol and stored at 4Â°C until ready to use. The reason why these danslyated compounds were used was because the required sensitivity for drug testing in oral fluid cannot be reached without using these chemical derivatives. THC and THCA are poorly ionized by ESI so not easily detectable, yet the derivatives are easily detectable by LC-MS/MS. Derivatization can also improve the fragment characteristics therefore enhancing sensitivity when using MS/MS. The phenol group in the THC and THCA is an active functional group for dansylation so enables both THC and THCA to be derivatized. pH 10.5 was used as the prescence of a dimethylamino group in the dansylated compounds meant that under acidic conditions it was susceptible to protonation1. Figure 2 shows the derivatization reaction scheme.
Figure 2 - The chemical derivatization of THC and THCA by dansyl chloride (DC). Adapted from reference1.
A gradient chromatographic separation was performed by reverse-phase HPLC using a Gemini c18 column (3 Î¼m particle size, 50 mm x 2 mm). The oven temperature for the column was set at 50Â°C, and the autosampler cooling unit was set at 10Â°C. Originally a mobile phase system using water-methanol for separation was used but due to the highly hydrophobic nature of DC-THC and DC-THCA, the analysis time was too long (20 mins) meaning that the peaks obtained were too broad or distorted. For this reason it was changed to 2 mobile phases, mobile phase A consisting of water with 0.5% formic acid an mobile phase B consisting of THF with 0.5% formic acid. 20 Î¼L sample was injected and the gradient programme used was 0 - 0.5 min with 40% mobile phase B, 0.5 - 1.5 min 40-90% mobile phase B, and 1.5-2 min, 90% mobile phase A. This changed the chromatographic elution behaviour and improved the separation efficiency, reducing the analysis time to 2 mins, producing a sharp peak for both DC-THC and DC-THCA. However, shorter separation times often cause a severe matrix effect. This was assessed by postcolumn immersion of DC-THC and DC-THCA during LC-MS analysis of methanol and the matrix extracted from the blank oral fluid. Taking this into account, a slower gradient elution was used: 0 - 0.5 min with 30% mobile phase B, 0.5 - 2.5 min 30-90% mobile phase B, and 2.5-3.5 min, 90% mobile phase A, increasing the time to 5 mins, and therefore reducing the matrix effect1. Figure 3 shows the peaks obtained.
Figure 3 - Chromatogram obtained from a single injection of 250 ÂµL of extracted preserved oral fluid. Copied from reference1
The detector used was an Applied Biosystems SCIEX API 3000 triple-quadrupole mass spectrometer with a TurboIonSpray interface (ESI) and a linearly accelerating high-pressure collision cell1. All analyses were performed in MRM mode and positive ESI mode, the peak width settings for Q1 and Q3 were unit resolution. The ion signals were maximised by using a semiautomated ramp procedure. The ions detected are summarised in the table below.
Precursor ion (m/z)
Product ion (m/z)
Table 1 - Table showing the m/z values detected and which compounds they relate to. Adapted from reference1.
Figure 4 - Proposed fragmentation of DC-THC and DC-THCA. Copied from reference1.
Figure 5 - Product ion spectra of DC-THC and DC-THCA. Copied from reference1.
Linearity of sample response was investigated in the range 5-500 pg/mL for THCA and 0.2-20 ng/mL for THC. Calibration curves for six levels of calibration from low to high concentrations for THC by adding 0.05, 0.1, 0.2, 0.4, 2, and 5 ng of standards, and for THCA adding 1.25, 2.5, 5, 10, 50 and 125 pg of standards, including 50 and 25 pg of their deuterated internal standards to 250 ÂµL of drug-free oral fluid. The linearity of the method was found by calculation of the regression line by the method of least squares and expressed by the squared correlation coefficient. The peak area ratios of target analysis and their internal standards were used to carry out a correlation analysis. The squared correlation coefficient versus analyte concentration was reported. It was found that at low control concentration, the mean recoveries were slightly lower compared to a high control concentration. The method was shown to be highly precise with the use of deuterated internal standard, good sensitivity and linearity were obtained for the analytes.
In this paper they have opted to go for the determination of THC in oral fluids by polymer monolith microextraction (PMME) combined with gas chromatography-mass spectrometry (GC-MS). The aim was to find a rapid method that had enough sensitivity to be able to detect THC in oral fluids. PMME was chosen as the structure has a larger surface area compared to other methods (such as SPME) allowing for a higher extraction capacity. Also, the polymer sorbent has hydrophilic carboxylic acid groups in the hydrophobic bone structure, which are biocompatible with complicated sample matrix such as oral fluid. After using this extraction process, the sample can be directly injected into the GC-MS system without derivatization which can sometime be time consuming.
THC standard solution was used and THC-d3 was used as an internal standard. Saliva samples were collected from drug-free volunteers and stored at -20Â°C until ready to use. They were centrifuged for 5 mins at 12,000 rpm at 4Â°C. 200 ÂµL of drug-free samples were collected and 10 ÂµL of internal standard solution (0.4 Âµg/mL in methanol) was added. After standing for 30 minutes to allow for thorough mixing, 1 mL phosphate butter (20 mM, pH 7) was added then filtered with 0.45 Âµm nylon membrane filter. The polymer monolith could collapse under high pressures (~300 psi) so the saliva samples were diluted with phosphate solution before extraction to reduce the viscosity of the sample and to keep it working under lower pressures. pH
The sample solution was then ready for extraction, using a p(MAA-co-EGDMA) monolith in fused silica capillary (2cm x 530 Âµm). The order for the extraction process was preconditioning, sorption, clean-up and desorption. A syringe infusion pump was used for the delivery of the sample. For preconditioning, 0.3 mL methanol in the syringe was ejected via the monolithic capillary at 0.15 mL/min, and then the monolithic capillary was washed with 0.3 mL phosphate buffer (20 mM, pH 7) at 0.15 mL/min. For sorption, 1 mL of sample was pumped through the capillary at 0.08 mL/min, then 0.2 mL of the phosphate buffer was put through at the same velocity to rid the residual matrix to avoid interference for separation. Flow rate was shown to have no significant influence on the extraction efficiency so 0.08 mL/min was chosen due to short extraction time with acceptable pressure of the monolith column. For desorption 0.05 mL acetone was ejected by the monolithic capillary at 0.04 mL/min and the eluate was collected for analysis by GC-MS. This extraction process was done in less than 20 minutes.
GC-MS analysis was performed using a Shimadzu GC-MS QP2010plus with an AOC-20i+s autosampler. The separation was done on a fused silica capillary column (HP-5MS, 30 m x 0.25 mm i.d., film thickness 0.25 Âµm). The oven temperature was programmed at 120Â°C for 2 mins, increased to 175Â°C at 20Â°C/min and held for 2 mins and finally increased to 295Â°C at 15Â°C/min and held for 10 mins. Splitless injection mode was used, with the splitless time at 1 min, the carrier gas used was helium at a flow rate of 1.2 mL/min. The injection port, ion source and interface temperatures were 260, 200 and 280 respectively.
The electron impact (EI) mass spectra of the analytes were recorded by SCAN mode (range 50-450 m/z) to conclude characteristic mass fragments and retention times. For analysis, the chosen characteristic mass fragments were monitored in the selected-ion monitoring (SIM) mode: for THC, m/z 299, 314, and 231, for THC-d3 m/z 302,317 and 234. SIM mode was used to achieve suitable sensitivity.
Internal calibration was done by plotting peak area ratios (THC/internal standard) against concentration ratios of THC and THC-d3. Stock solution was added to the sample solutions to get final concentrations of THC at 3, 10, 20, 50, 100 and 300 ng/mL, and THC-d3 at 20 ng/mL for calibration curves.