OR WAIT null SECS
Automated solid-phase extraction (SPE) has been used extensively with liquid chromatography–tandem mass spectrometry (LC–MS-MS) to facilitate high-throughput analysis in the pharmaceutical, diagnostic, and forensic toxicology areas. In this work, we demonstrate the use of a systemized approach to SPE method development and LC–MS-MS analysis. This approach provides dramatic savings in analysis time and takes advantage of new innovations in high performance liquid chromatography (HPLC) columns to provide the cleanest extracts for LC–MS injection.
The use of automated solid-phase extraction (SPE) has been shown to reduce sample preparation time by at least 50% in comparison with manual SPE (1). A further 50% improvement in throughput can be achieved by avoiding evaporation and reconstitution steps and direct injecting the SPE extracts into the liquid chromatography–mass spectrometry (LC–MS) system (1). An additional advantage with direct injection of SPE extracts is elimination of analyte loss during evaporation, as observed for compounds such as isoniazid (2) and indomethacin (3).
For the cleanup of basic drugs, associated SPE methods commonly utilize strong cation exchange sorbents. However, even acidic and neutral compounds can be cleaned up effectively with these strong cation exchange sorbents, including hydrophobic compounds such as THC-carboxylic acid or very polar molecules like acrylamide, γ-amino butyric acid analogs, and antivirals like penciclovir (4–7).
The preference for employing strong cation exchange sorbents also stems from the fact that proteinaceous and endogenous components of biological matrices can be washed out very efficiently with a strong organic solvent before elution of basic analytes. Strong organic washes furnish very clean elution extracts, displaying insignificant ion suppression and enhancing lower detection and quantitation limits in analyses (8). A vast majority of drugs are basic and fit this scenario perfectly.
The typical elution conditions used during SPE of biological samples with strong cation exchange sorbents consist of an organic solvent containing 2–5% ammonium hydroxide (9,10).
The nature of the strong organic elution solvent and the high pH of these extracts have made such elutions incompatible with traditional column chemistries and reversed-phase solvent conditions. As such, the common practice is to evaporate off the ammonia along with the solvent and reconstitute the analyte or analytes in an aqueous organic solvent amenable to reversed-phase LC–MS analysis.
The recent development of LC columns that have increased pH stability gives method developers the option to inject these high-pH extracts directly into the LC–MS-MS system. Studies during the last decade clearly show that LC–MS employing high-pH mobile phases on the novel stationary phases, resistant to degradation/dissolution under such conditions, furnishes several advantages (11,12). Most attractive are the enhancement in sensitivity and resolution, along with improved retention for polar basic analytes. These investigations also show that mixtures of neutral, acidic, and basic analytes can be analyzed efficiently under these high-pH mobile phase LC–MS conditions, which when extrapolated to drug–metabolite combinations are highly desirable for obtaining excellent resolution and sensitivity for very low abundance metabolites.
Materials and Reagents
A multisorbent strata-X method development plate and single-sorbent strata-X-C 96-well plates (both with 10-mg bed masses) were obtained from Phenomenex (Torrance, California). All the analytes studied were obtained from Sigma (St. Louis, Missouri) and used as such. Human plasma was obtained from Valley Biomedical (Winchester, Virginia).
A pH-stable 50 mm × 2.0 mm Gemini NX C18 column was obtained from Phenomenex. MS detection was done using an ABI 3000 mass spectrometer with electrospray ionization in the positive ion mode at a temperature of 425 °C (Applied Biosystems, Foster City, California).
All reagents and solvents used were also from Sigma and were used without further purification.
Preparation of Samples
Stock solutions of atenolol, toliprolol, bunitrolol, and bupranolol were made by dissolving each drug in methanol at a concentration of 1 mg/mL. The plasma samples were spiked with appropriate volumes of these solutions corresponding to concentrations of 0.5–500 ng/mL of plasma. The concentration of nadolol (internal standard) was 40 ng/mL.
Automated SPE Method Development Protocol
Step 1: Method development with the strata multisorbent well plate:
The strata-X method development plate was conditioned with 400 μL of methanol followed by 400 μL of water. The plasma solutions were loaded under the three sets of conditions specified below:
After the initial wash (wash-1) at one specific pH, the second wash (wash-2) was done with 400 μL of 70% methanol before elution. The plate was dried for 1 min at 10 in. of Hg before elution.
Step 2: Method validation with the protocol developed in Step 1:
The single-sorbent well plate was conditioned with 400 μL methanol followed by 400 mL of 0.1 M acetic acid. Human plasma with disodium EDTA as anticoagulant was spiked with analytes (studied for a wide calibration range for precision and accuracy) and internal standard (nadolol). Samples were diluted (1:2) with 0.1 M acetic acid before loading on the plate. Washing was done in two steps, first with 400 μL of 0.1 M acetic acid, followed by 400 μL of methanol. Before eluting with 300 μL of 5% ammonium hydroxide in methanol, in two increments, the sorbent was dried for 60 s under high vacuum of 10 in. of Hg. The elution was injected directly into the LC–MS-MS system.
For HPLC, a 50 mm × 2.0 mm, 3-μm dp Gemini NX column was used under gradient conditions, using a mobile phase of 5 mM ammonium bicarbonate (pH 10.0)–acetonitrile from 90:10 to 25:75 in 2.5 min followed by equilibration for 1.5 min with a flow rate of 0.5 mL/min. The injection volume was 5 μL.
The acidic mobile phase runs utilized the standard mobile phase conditions consisting of aqueous 0.1% formic acid and acetonitrile, with the same gradient conditions as for the basic analysis.
Figure 1 gives the structures of the analytes and internal standard used in this study, along with their pKa and log P values. Of all the analytes, atenolol is the most polar, though all of them display about the same basicity. All analytes are secondary amines.
Figure 1: Structures and physicochemical parameters of the probes studied.
The first step in an overall analytical method development process for an analyte from any matrix, and a biological matrix in particular, is to develop an appropriate sample preparation protocol. The purpose of sample preparation is to concentrate the analyte and to clean it up from matrix constituents that can interfere with its analysis. Sometimes, a selective separation–enrichment is a bonus, as with low-abundance metabolites in large quantities of drugs.
To simplify the method development process, we designed an automated SPE method development protocol using a multisorbent 96-well plate in tandem with an automatic liquid handler (13). This well plate consists of four different functionalized polymeric sorbents, a neutral sorbent (strata-X), a strong cation exchanger (strata-X-C), a weak cation exchanger (strata-X-CW), and a weak anion exchanger (strata-X-AW). These four collectively cover a wide range of polarities and cater to all possible modes of hydrophobic and polar interactions (including ion exchange interactions) that can be displayed by any analyte.
The Step-1 generic protocol was designed for screening all four sorbents simultaneously using three sets of conditions, as outlined in the Experimental section. After this Step-1 protocol is completed, a method developer is able to identify the most appropriate sorbent, as well as the load, wash, and elution conditions (pH and solvent) for the subsequent validation step (Step 2).
In Step 1, a 70% methanol wash step was utilized after an initial water or buffer wash. From the work of Chambers and colleagues (8), it is evident that methanol is an excellent solvent for the removal of phospholipids and other endogenous materials from plasma, even at concentrations of 60–80%.
Aimin Tan and coworkers (14) report a protocol that takes advantage of the fact that basic analytes undergo a dramatic reduction in the degree of ionization at or near their pKa values, which enables the hydrophobic mechanism to operate more effectively with neutral sorbents that permit a stronger organic wash. The results from Step-1 protocol (Figure 2) demonstrate that even without employing any such basic aqueous–organic wash conditions, the neutral sorbent provides recoveries of around 60% for the strongly polar basic analyte atenolol when a wash-2 with 70% methanol is used. No loss of atenolol was observed when 30% methanol was used for wash-2 with the neutral sorbent.
Figure 2: Recovery of atenolol from a strata multisorbent 10-mg 96-well plate.
Even though decent recoveries were obtained with the neutral sorbent under the NN conditions, the results of the Step-1 program indicated that the strong cation exchange sorbent (strata-X-C) yielded the best results of any of the sorbents using the AB conditions. Recoveries of 98% were achieved when samples were subjected to a 70% methanol wash (Figure 2). The same near-quantitative recoveries were obtained with 100% methanol wash, allowing for efficient removal of phospholipids and other endogenous materials, which are responsible for ion suppression during LC–MS-MS. However, the matrix effects were insignificant with the 100% methanol wash.
Step 2 incorporates the optimized conditions developed in Step 1, except that 100% methanol wash was used and a 96-well strong cation exchange plate is used for validation of the method. Of particular interest in this analysis was the use of the AB conditions. Under these conditions, the eluates will be in 5% ammonium hydroxide in methanol. It is common practice to evaporate off the ammonia along with the solvent and reconstitute the residue in a formic acid–acetonitrile mobile phase for LC–MS-MS analysis, as reported in all documented literature using strong cation exchange sorbents for sample preparation.
This evaporation–reconstitution for 96 samples involves an additional 2.5–3.0 h that laboratories must spend to prepare their samples. However, if it is possible to avoid these steps and inject the ammonia–organic extract directly into the LC–MS-MS system, this time could be saved. Due to 100% methanol wash, the eluate from the strong cation exchange material is extremely clean, so there is less potential for contamination. An additional advantage of the direct injection option is that basic drugs will be in either neutral or near-neutral form (depending upon their pKa values), and this would enhance hydrophobic interactions and hence, retention under reversed-phase high performance liquid chromatography (HPLC) conditions.
However, such direct injection onto traditional silica-based reversed-phase columns is not feasible because of stability problems. The recently introduced Gemini and Gemini-NX columns allow method developers to use pH conditions from 1–12. Therefore, handling the high-pH mobile phases in the present work with Gemini-NX was not problematic.
Previous work from our laboratory has demonstrated that when performing LC–MS-MS analysis using these new materials in high-pH mobile phases, analyte sensitivity can be increased (11). The increase in sensitivity achieved by performing chromatographic separations at high pH is due to multiple factors, including extended retention, excellent peak shapes, and good efficiency. However, this increase in sensitivity is observed not only for the basic analytes, but for some acidic and neutral analytes as well.
Figure 3: Linearity of toliprolol and atenolol over a 1000-fold concentration range.
The data collected from Step 2 through LC–MS-MS analysis under mobile phase conditions outlined in the experimental section is summarized in Figures 3–5. Figure 5 shows a representative chromatogram for these drugs in both acidic and basic mobile phase conditions. In comparison with data from the same extracts run under acidic conditions, employment of the high-pH mobile phase showed about 1.5–4.0 times higher sensitivity for the four analytes studied. This increase in sensitivity helped compensate for the fact that we did not concentrate extracts through evaporation and reconstitution steps. Also interesting was the fact that the peak shape of all compounds, including atenolol, was not affected adversely due to the high organic concentration of the injection solvent, under either basic or acidic conditions. More will be published on these results at a later date.
Figure 4: Linearity of bunitrolol and bupropranolol over a 1000-fold concentration range.
Figures 3 (atenolol and toliprolol) and 4 (bunitrolol and bupranolol) show the linearity of the method over the concentration range studied, 0.5–500 ng/mL for these analytes from the Step 2 protocol. The low limit of quantitation, broad dynamic range, and precision obtained suggest that this method would be suitable for most pharmaceutical and other analytical needs.
Figure 5: Comparison of sensitivity between high pH (a) and low pH (b) mobile phases for atenolol, nadolol, and bunitrolol.
The use of a structured SPE method development protocol quickly identified the best sorbent for the extraction procedure. The use of the strong cation exchange material allowed for 100% methanol wash conditions, resulting in extremely clean extracts. Using the Gemini-NX high pH stable HPLC column allowed the basic eluate to be injected directly onto the LC–MS-MS system. The high-pH chromatographic conditions resulted in higher MS response. By eliminating time-consuming steps throughout the process, the resulting method is perfectly suited to the high-throughput environment.
Although this work demonstrated the advantages of combining automated sample preparation with direct injection and high-pH LC–MS-MS analysis for a small set of basic probes, this universal protocol is applicable to any analyte, neutral, acidic, or basic, from any kind of matrix.
Liming Peng, Shahana Wahab Huq, Krishna M. Kallury, and Terrell Mathews are with Phenomenex, Torrance, California.
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