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Comparison of LC–MS and GC–MS for the Analysis of Pharmaceuticals and Personal Care Products in Surface Water and Treated Wastewaters 8Blake Rushing, Ashley Wooten, Marcus Shawky, and Mustafa I. Selim

This study of a selected group of PPCP contaminants in eastern North Carolina demonstrates the advantages and disadvantages of LC–MS and GC–MS as well as SPE and liquid–liquid extraction.

Simultaneous Quantitation of Buprenorphine and Its Metabolites Using LC–MS 15Anand A. Joshi, Neha V. Maharao, and Phillip M. Gerk

An LC–MS method for simultaneous quantification of buprenorphine and three metabolites: norbuprenorphine, buprenorphine glucuronide, and norbuprenorphine glucuronide

The Applicability of Field-Portable GC–MS for the Rapid Sampling and Measurement of High-Boiling-Point Semivolatile Organic Compounds in Environmental Samples 20Tai Van Truong, Nathan L. Porter, Edgar D. Lee, and Robert J. Thomas

A look at the use of field-portable GC–MS with solid-phase microextraction, purge-and-trap, thermal desorption, and heated headspace sampling techniques to provide a fast response for in-field analysis of SVOCs in a wide variety of environmental-type samples including potable waters, tea, plants, and road gravel

Sensitive, Rapid Estimation of Moxidectin in Cattle Hair by LC–MS-MS 27P. Sambasivarao, Raman Batheja, N. Subbarao, S. Ashma, K. Ashwini, and M. Mupeksha

Validation of this rapid bioanalytical method for the determination of moxidectin in cattle hair demonstrated that the method is

accurate, reliable, and reproducible.

DepartmentsReview of the 64th Conference on Mass Spectrometry and Allied Topics . . . . . . . . . . . . . . . . 32

ASMS Product Profiles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34

Calendar . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

Ad Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

Cover image courtesy of Anirut Kongsorn 2015/Getty Images

July 2016

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www.spec t roscopyonl ine .com8 Current Trends in Mass Spectrometry July 2016

Blake Rushing, Ashley Wooten, Marcus Shawky, and Mustafa I. Selim

Water samples were obtained from the Tar River and a local water treatment plant in eastern North Carolina in spring 2013 and fall 2015 to monitor the presence of a panel of pharmaceutical and personal care products (PPCPs). Samples were extracted by solid-phase extraction (SPE) or liquid–liquid extraction and analyzed for parent PPCPs and their metabolites by high performance liquid chromatography–time-of-flight mass spectrometry (HPLC–TOF-MS) and gas chromatography–mass spectrometry (GC–MS). Both extraction and detection methods were compared by their recoveries and detection limits for each compound. Many parent PPCPs and their metabolites were detected including: carbamazepine, iminostillbene, oxcarbazepine, epiandrosterone, loratadine, β-estradiol, triclosan, and others. Liquid–liquid extraction was found to provide overall superior recoveries. Furthermore, HPLC–TOF-MS yielded lower detection limits than GC–MS. Library searching of addi-tional peaks identified further compounds with biological activity. Additionally, the effectiveness of the treatment plant on the removal of the compounds of interest is discussed.

Comparison of LC–MS and GC–MS for the Analysis of Pharmaceuticals and Personal Care Products in Surface Water and Treated Wastewaters

Pharmaceuticals and personal care products (PPCPs) have been found as contaminants in drinking and wastewater worldwide (1–4) and can pose a toxicological risk to hu-

mans as well as wildlife (5–8). Although water treatment plants in-corporate a wide variety of methods to remove these compounds, many PPCPs have been shown to persist posttreatment, allowing for their accumulation in the environment (9–12). Because of the ubiquitous nature of these contaminants and their wide variety of effects on biological organisms, detection and tracking of PPCPs has become an area of increasing research interest.

Statistics from the Center of Disease Control and Prevention show that Eastern North Carolina has the highest occurrence of stroke and heart disease compared to other regions in the United

States (8). Although this phenomenon may be attributable to fac-tors such as socioeconomic status, ethnic distribution, or dietary trends, exposure to contaminants like PPCPs may be a large con-tributing factor based on the wide range of health effects that they can impart. Data on the prevalence of PPCPs in this region is currently limited to only one study detecting a total of four PPCPs using gas chromatography (GC) (13).

Chromatography and mass spectrometry (MS) are com-monly used analytical techniques to identify and quantify water contaminants such as PPCPs. Previous studies have used liquid chromatography–MS (LC–MS) (3,4,10,14–17), GC–MS (1,18,19), or both (2,9,20,21) to detect compounds within this class. In terms of sample preparation, many stud-

www.spec t roscopyonl ine .com July 2016 Current Trends in Mass Spectrometry 9

ies use solid-phase extraction (SPE) (1–4,10,14,15,17–19,21); however, some studies use liquid–liquid extraction (20), or both (16). The PPCPs analyzed in these studies vary greatly not only with respect to their biological mechanisms of action, but also to their chemical–physical properties such as acid–base properties, volatilities, thermal stabili-ties, and polarities. These differences, combined with the matrices of surface and wastewaters, influence the efficien-cies of these extraction methods. PPCP contaminants are commonly present in the nanogram-per-liter range, so low extraction recoveries can often lead to many undetected compounds. Fur-thermore, liquid and gas chromatog-raphy can possess drastically different detection limits based on the types of compounds being analyzed, which can exacerbate the issue of undetected con-taminants. Data comparing these two instrumental and extraction techniques on PPCPs is lacking, making it unclear which combination of methods would be best suited for this type of analysis. Figure 1: Representative TIC chromatograms of 12 selected PPCPs using (a) GC–MS and (b) HPLC–TOF-MS.

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The goal of this study is to uncover the advantages and disadvantages of LC–MS and GC–MS as well as SPE and liquid–liquid extraction on a selected group of PPCP contaminants in Eastern North Carolina. These data would un-cover the advantages and disadvantages to these commonly used techniques, revealing some insight about which in-strumental and extraction techniques are most suitable for these PPCPs. Also,

this study aims to use this informa-tion to uncover some of the additional emerging PPCPs in Eastern North Carolina and determine their relation-ship to any underlying diseases in the area as well as any seasonal variations in detected compounds. It is expected that these findings will be translatable to similar studies detecting PPCPs in other geographical regions, improving the detection of these compounds.

MethodsChemicals

Solvents for standard preparation and instrumental analysis (acetonitrile, methylene chloride, methanol, and water) were LC–MS-grade and were pur-chased from Sigma-Aldrich. ENVI-Disk C18 SPE disks (47 mm in diameter) were purchased from Supelco. β-estradiol (≥98%), caffeine (≥98.5%), ketoprofen (≥98%), naproxen (≥98%), ibuprofen (≥98%), triclocarban (≥99%), diphenyl-hydramine (≥98%), and napropamide (≥98%) were purchased from Sigma Aldrich. Carbamzepine (≥99%), oxcar-bazepine (≥97.5), and loratadine (≥98%) were purchased from Fluka. Iminostil-bene (97%) was purchased from Chem Service. Epiandrosterone (≥99%) was purchased from MP Biomedicals.

Instrumental Conditions

The GC–MS system consisted of an Agilent 7890A gas chromatograph set in splitless injection mode and an Agilent 5975B mass spectrometer with an elec-tron ionization source. A 30 m × 0.25 mm, 0.5-μm df DB-5MS column (J&W Scientific) was used in this study. The carrier gas (helium) was kept at a flow of 0.8 mL/min. The injection volume for all samples was 1 μL using a 7683B series Agilent liquid injector. Temperature pro-gramming was as follows: Hold initially at 150 °C for 5 min, ramp at 10 °C/min to 300 °C, and hold at 300 °C for 10 min.

Table I: Selected PPCPs and their characteristics. Compounds were quantified by GC–MS using the most abundant fragment (target) and verified using two qualifier ions (Q1 and Q2). Compounds analyzed via HPLC–TOF-MS were quantified using [M+H]+ values.

CompoundGC HPLC

DescriptionTarget Q1 Q2 [M+H]+

β-Estradiol 272 213 160 273.1849 Contraceptive—estrogenic activity

Caffeine 194 109 67 195.0877 Stimulant

Carbamazepine 193 192 236 237.1022 Anticonvulsant

Diphenylhydramine 58 73 165 167.0876* Antihistamine

Epiandrosterone 290 107 108 291.2319 Androgenic hormone

Ibuprofen 91 161 163 207.1380 NSAID

Iminostilbene 193 194 192 194.0964 Metabolite of carbamazepine

Ketoprofen 105 177 209 255.1016 NSAID

Loratadine 266 245 280 383.1521 Antihistamine

Naproxen 185 230 141 231.1016 NSAID

Oxcarbazepine 180 209 252 253.0972 Anticonvulsant

Triclocarban 127 161 163 314.9853 Antibacterial

*Diphenylhydramine was quantified off of a more abundant characteristic mass on HPLC–TOF-MS – mass 167.0876.

Figure 2: Comparison of recoveries obtained from extracting PPCPs using liquid–liquid extraction and SPE. Recovery values were quantified using HPLC–TOF-MS.

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The source temperature was kept at 230

°C, and the electron multiplier detector was set at a voltage of 2012 V, absolute. For quantitative analysis, selected ion monitoring (SIM) was used for the mea-surement of each compound. For each compound, the most abundant m/z frag-ment was used for quantification and two additional characteristic ions were used as qualifiers. Data were collected and analyzed using MSD ChemStation E.02.00.493 software.

The high performance liquid chro-matography–time-of-flight mass spec-trometry (HPLC–TOF-MS) system consisted of an Agilent 1200 series HPLC system coupled to an Agilent 6220 TOF-MS system with a dual elec-trospray ionization (ESI) interface set to positive ion mode. The HPLC column used was a 150 mm × 2.1 mm, 3.5-μm df Agilent Zorbax Eclipse Plus C18 col-umn. Mobile-phase A was HPLC-grade water with 1% formic acid, and mobile-phase B was acetonitrile with 1% formic acid. The injection volume used for all samples was 3 μL. Solvent program-ming was as follows: 20–80% B over 20 min, hold at 80% B for 5 min. The col-umn temperature was set to 35 °C. The drying gas (nitrogen) temperature was set to 335 °C at a rate of 10 L/min with the nebulizer pressure kept at 35 psig. The capillary voltage was set to 3300 V with the fragmentor voltage at 185 V and the skimmer voltage at 30 V. Iden-

tification and quantification of each compound was based off of [M+H]+ values. Agilent MassHunter Worksta-tion software version B.02.01 was used to collect and analyze data.

Comparison of Detection Limits

A mixture containing 10 μg/mL each of β-estradiol, caffeine, carbamazepine, diphenylhydramine, epiandrosterone, ibuprofen, iminostilbene, ketoprofen, loratadine, naproxen, oxcarbazepine, and triclocarban was prepared in aceto-nitrile. Compounds were selected based on preliminary screening of local water sites (data not shown). Calibration curves were constructed for each com-pound by making dilutions between 5 μg/mL to 25 ng/mL. Each dilution was analyzed by GC–MS and HPLC–TOF-MS. For compounds that were detect-able at 25 ng/mL, 3(S0) was used as the limit of detection, where S0 = standard deviation of zero concentration. For compounds that were not detected at

25 ng/mL, the lowest dilution in which they were detected was set as the detec-tion limit.

Comparison of Extraction Methods

Standard Mixture Preparation

A mi x ture of 1 μg /mL each of β-estradiol, caffeine, carbamazepine, diphenylhydramine, epiandrosterone, ibuprofen, iminostilbene, ketoprofen, loratadine, naproxen, oxcarbazepine, and triclocarban was prepared in ace-tonitrile. Then 1 mL of this standard mixture was spiked into 500 mL of de-ionized water and mixed by shaking.

Solid-Phase Extraction

The SPE disks were conditioned using 10 mL of acetonitrile, 10 mL of metha-nol, and 10 mL of deionized water with 2 min of equilibration time in between each solvent. The 500 mL of spiked water (described above) was loaded onto the conditioned SPE disk at 0.15–0.2 mL/min and then dried under vacuum for

Table II: Detection limits of all 12 PPCPs as determined by GC–MS and HPLC–TOF-MS

CompoundHPLC D.L.

(ppb)GC D.L. (ppb)

β-Estradiol 250 250

Caffeine 45 25

Carbamazepine 24 25

Diphenylhydr-amine

18 25

Epiandros-terone

234 1000

Ibuprofen 20 250

Iminostilbene 20 25

Ketoprofen 21 1000

Loratadine 15 1000

Naproxen 12 25

Oxcarbazepine 20 100

Triclocarban 246 1000

Table III: Additional contaminants found in influent or effluent samples. Compounds were identified using NIST Mass Spectral Search software that was connected to the GC–MS.

Compound Description

1,4,6-Androstatriene-3,17-dioneAromatase inhibitor, metabolite of

boldenone

3-DeoxypregnenloneNeurosteroid, metabolite of

pregnenelone

Androstadiendione Prohormone, precursor to boldenone

Androstane-11,17-dione, 3-hydroxy-, (3α,5β)- Androgenic activity

Epiandrosterone Androgenic activity, metabolite of DHEA

PrednisoloneGlucocorticoid, treats inflammatory and

autoimmune conditions

Pyrrobutamine Antihistamine

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Dehydroabietic acid Resin acid

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20 min. For elution, 10 mL of acetoni-trile was added to the disk, allowed to sit for 5 min, and then eluted at 0.1 mL/min. The eluate was collected in a gradu-ated 15-mL conical test tube and then dried under nitrogen to a volume less than 1 mL. This process was repeated a total of three times and the extracts were analyzed using both GC–MS and HPLC–TOF-MS.

Liquid–Liquid Extraction

The 2000 mL of spiked water (described above) was loaded into a separatory fun-nel along with 100 mL of methylene chloride. The solution was shaken by

hand for 1 min and then the methy-lene chloride layer was collected into a round-bottom flask. This process was repeated a total of three times, giving a final volume of 300 mL of methylene chloride. The methylene chloride extract was combined in a 500-mL round-bot-tom flask, which was then loaded onto a rotary evaporator where the extract was concentrated to a few milliliters. The concentrated extract was quanti-tatively transferred to a 15-mL gradu-ated test tube and the final volume was adjusted to 1 mL, by repeated washing, transfer, and evaporation. This process was repeated a total of three times and

the extracts were analyzed using both GC–MS and HPLC–TOF-MS. Analyte recoveries for both liquid–liquid extrac-tion and SPE were calculated using the following formula:

(Analyte mass in extract/Analyte mass in spiked water) × 100 = Analyte recovery [1]

Analysis of PPCPs in

Surface and Wastewater

Sample Collection

Two sampling sites within the Greenville, North Carolina, wastewater treatment plant were chosen for analysis of PPCPs: influent wastewater and effluent water (cleaned end-product). At each of those sites, 4-L grab samples were collected dur-ing the spring of 2013 and the fall of 2015. Samples were transported back to the laboratory on ice within 30 min of collec-tion. Amber, 4-L HPLC solvent bottles that were previously cleaned using methanol and acetonitrile were used for sample col-lection and storage. For samples that were not immediately analyzed, the pH was adjusted to 2 using hydrochloric acid to prevent bacterial growth and subsequent breakdown of PPCPs. Samples were then stored at 4 °C until analysis.

Extraction and Measurement of PPCPs

Samples from 2013

First, 2 L of sample material was filtered using a 0.45-μm glass fiber filter and ex-tracted using the liquid–liquid extraction method detailed earlier. Extracts were analyzed using HPLC–TOF-MS and origi-nal sample concentrations were calculated based off of determined recovery values. Both influent and effluent sites were ana-lyzed in duplicate. GC–MS was also used to analyze extracts to qualitatively identify additional PPCPs using the National Soft-ware Reference Library (NIST) Mass Spec-tral Search Program software version 2.0. Peaks were identified based on the com-pounds with the highest matching spectral score given by the software.

Samples from 2015

First, 500 mL of sample material was fil-tered through a 0.45-μm glass fiber filter and spiked with 1 mL of napropamide as an internal standard. PPCPs were ex-tracted from 500-mL aliquots using SPE disks according to the method outlined

Figure 3: PPCPs detected using HPLC–TOF-MS in influent and effluent waters in a local wastewater treatment plant in Greenville, North Carolina. (a) Samples were collected in the spring of 2013 and extracted using liquid–liquid extraction. (b) Samples were collected in the fall of 2015 and extracted using SPE.

Influent

4500

4000

3500

3000

2500

2000

Co

nce

ntr

ati

on

(n

g/L

)C

on

cen

tra

tio

n (

ng

/L)

1500

1000

500

0

15,000

20,000

25,000

30,000

35,000

(a)

(b)

10,000

5000

0

Effluent

Influent

Effluent

β-Es

trad

iol

Caffe

ine

Carbam

azep

ine

Dip

henylhyd

ram

ine

Epia

ndroster

one

Ibupro

fen

Imin

ostilb

ene

Ketopro

fen

Lora

tadin

e

Nap

roxe

n

Oxc

arbaz

epin

e

Triclo

carb

an

β-Es

trad

iol

Caffe

ine

Carbam

azep

ine

Dip

henylhyd

ram

ine

Epia

ndroster

one

Imin

ostilb

ene

Ketopro

fen

Lora

tadin

e

Nap

roxe

n

Oxc

arbaz

epin

e

Triclo

carb

an

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earlier (n = 5 and n = 6 for influent and effluent samples, respectively). Extracts were analyzed using HPLC–TOF-MS and quantified using the internal stan-dard peak.

Results and DiscussionTable I displays a panel of 12 PPCPs that were selected based on preliminary screen-ing of water samples (data not shown). For GC–MS analysis, compounds were quan-tified using a target ion that was the most abundant m/z value, and the next two most abundant fragments were used as qualifier ions (Q1 and Q2). For HPLC–TOF-MS, compounds were quantified using their [M+H]+ adduct masses, with the excep-tion of diphenylhydramine, which has a much more abundant characteristic mass of 167.0876 that was produced by the frag-mentor. Standard mixtures were prepared and analyzed using both GC–MS and HPLC–TOF-MS with the methods detailed above. Representative total ion current (TIC) chromatograms showing chromato-graphic separation of these 12 compounds are shown in Figures 1a (GC–MS) and 1b (HPLC–TOF-MS). Of the selected com-pounds, triclocarban and ketoprofen were unable to be detected using the GC–MS method; however, the HPLC–TOF-MS method could detect all 12.

Detection limits were calculated to further compare the suitability of each instrument for the selected compounds. Because PPCPs are typically present in nanogram-per-liter concentrations in surface water and wastewater, detection limits are often one of the greatest chal-lenges to overcome when analyzing these samples. As shown in Table II, HPLC–TOF-MS achieved lower detection limits for all compounds with the exception of caffeine and β-estradiol. Caffeine’s detec-tion limit was 1.8 times lower on the GC–MS and β-estradiol had the same detec-tion limit between both instruments. In general, more polar compounds are more amenable for liquid chromatography, so superior detection limits were expected with HPLC. Less-polar compounds like caffeine and β-estradiol are typically well-suited for GC analysis, which was reflected in our results. Overall, HPLC–TOF-MS was able to detect lower amounts of the selected compounds and was therefore chosen to quantify all samples for the re-

mainder of the study.Both liquid–liquid extraction (using

methylene chloride) and C18 SPE were compared on their ability to recover each of the 12 analytes. As shown in Figure 2, liquid–liquid extraction produced higher recoveries for all 12 compounds as com-pared to SPE. In this scenario, the use of SPE for surface and wastewater samples may lead to a reduced ability to detect PPCPs because of lower recovery rates, particularly with iminostilbene and diphe-nylhydramine. Although more sample loss was observed, SPE produced less variation than liquid–liquid extraction. This in-creased precision combined with higher throughput may allow SPE to be more advantageous than liquid–liquid extrac-tion in circumstances where these prop-erties are favored. Another key difference between the two extraction methods is that liquid–liquid extraction had a higher capacity for sample volume. Amounts greater than 500 mL led to sample break-through on the SPE disks, whereas liquid–liquid extraction could potentially handle volumes greater than the 2000 mL used in this study. Higher sample capacity could assist in detecting trace analytes by allow-ing for a larger degree of concentration.

The two extraction methods, liquid–liq-uid extraction and SPE, were next com-pared on their ability to extract PPCPs from surface and wastewater field samples. Samples were taken from the wastewater treatment plant in the city of Greenville, North Carolina in the spring of 2013 (ex-tracted using liquid–liquid extraction) and the fall of 2015 (extracted using SPE). This also allowed us to compare seasonal differ-ences in the amounts of PPCPs detected. The treatment plant involved three main methods of treatment to produce clean drinking water. The primary treatment involves the use of screens to remove physical objects (for example, plastic, wood, sand, and so forth). The secondary treatment uses a microorganism chamber to break down organic matter and remove nitrogen as well as phosphorus. Finally, a tertiary treatment uses a deep-bed sand fil-ter and UV light to further disinfect and purify. Samples were collected from the in-fluent as well as effluent water to not only determine PPCPs present in untreated water, but also to see the effectiveness of these treatment methods on removal of

these compounds.Using HPLC–TOF-MS as the preferred

instrumental technique for quantification, extracts from influent and effluent waters were analyzed for all 12 analytes. Figure 3a shows that in the spring of 2013, all compounds were detected in the influent water with carbamazepine as the most prominent contaminant at 3.47 μg/L using liquid–liquid extraction. Carbamazepine has been found at the microgram-per-liter concentration level in a number of other studies, particularly in European countries (22–26). Carbamazepine has been found to be particularly resilient toward removal by water treatment methods (27,28) as well as having reduced biodegradability (29). The high rates of seizures in the population of this region may be an additional explana-tion as to why this drug is found at such high levels along with its metabolite, imi-nostilbene, as well as oxcarbazepine. Addi-tionally, these data show that the treatment plant was effective at reducing the levels of all of these compounds, with β-estradiol and epiandrosterone being undetectable in effluent samples. Overall, samples col-lected in the fall of 2015 also showed a de-crease in PPCP levels in influent samples as compared to effluent samples after extraction by SPE with carbamazepine as the most prominent contaminant at 27.0 μg/mL (Figure 3b). Two exceptions to this decrease were carbamazepine and β-estradiol, which both had increased levels in effluent samples. It is well known that many PPCPs heavily partition into suspended solids in wastewater samples (30–38), which can heavily influence their detectable concentration in the aqueous phase. Differences in solid composition and amount may suppress the detectable concentration in the aqueous phase using this method. Additionally, the liquid–liq-uid extraction method may have been bet-ter able to extract adsorbed compounds in any residual solid material left in the water samples. When comparing the two sets of samples, water from the fall of 2015 had higher amounts of all PPCPs except for ibuprofen and iminostilbene (ibuprofen was unable to be detected in either influent or effluent samples from 2015). Differences in the two sample sets may be due to varia-tions in prescribed drugs at both times, city population, as well as any differences in water treatment processes. These results


highlight the degree of variation of PPCPs within surface and wastewaters over time.

In addition to the 12 compounds se-lected for this study, other contaminants were identified in the spring 2013 samples using NIST library searching that was coupled to the GC–MS. Using this soft-ware, unidentified peaks could be quali-tatively identified, further characterizing the compounds that were detected in these samples using these methods. Table III lists the major additional compounds detected across all samples along with a brief description of their class and any no-table biological activity. Many of these con-taminants had endocrine activities or were plasticizers or surfactants. Future studies may apply the methods used here on these additional compounds or other PPCPs.

ConclusionsThe data provided in this study provide evidence for the advantages of different combinations of instrumental and extrac-tion techniques for the analysis of PPCPs in surface and wastewaters. For the com-pounds chosen in this study, HPLC–TOF-MS was overall more sensitive and allowed for the detection of more compounds than GC–MS. Additionally, liquid–liquid ex-traction achieved higher recoveries for the measured PPCPs than SPE, although SPE showed less variation. Also, the method of water treatment used in this study was overall shown to be effective at reducing or eliminating this panel of PPCPs. The results provided in this study highlight the importance of selecting the appropri-ate extraction and analysis techniques to best analyze PPCPs water samples.

AcknowledgmentsThe authors would like acknowledge the help of the Mr. Jeff Camp and other admin-istrative personnel from Greenville Utilities, for facilitating sample collection for this project. Funding support for this work was provided by East Carolina University, Of-fice of the Vice President for Research, East-West Project. We also would like to thank Dr. Siddhartha Mitra for providing some of the analytical standards used in this work.

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Blake Rushing, Ashley Wooten, Marcus Shawky, and Mustafa I. Selim are with the Department of Pharmacology and Toxicology in the Brody School of Medicine at East Carolina University in Greenville, North Carolina. Direct correspondence to: [email protected] ◾


Anand A. Joshi, Neha V. Maharao, and Phillip M. Gerk

A liquid chromatography–mass spectrometry (LC–MS) method has been developed for simultaneous quantification of buprenorphine and its three metabolites, namely norbuprenorphine, buprenorphine glucuronide, and norbuprenorphine glucuronide. Chromatographic separation was achieved on a C18 column with a gradient of acetonitrile over ammonium acetate buffer (25 mM, pH 6.6). The method run time was 7.5 min. Quantification was performed by selected ion monitoring of [M+H]+ ions of norbu-prenorphine glucuronide (590), norbuprenorphine (414), buprenorphine glucuronide (644), and buprenor-phine (468). Naloxone (328) (328 ng/mL) was used as an internal standard. The samples were processed by protein precipitation and extraction recovery was *95% with minimal observed matrix effects ()11%). Linear calibration curves were obtained over a range of 25 or 100 ng/mL to 4000 ng/mL, depending on the analyte. The lower limit of quantitation for buprenorphine and norbuprenorphine was 25 ng/mL and for buprenorphine glucuronide and norbuprenorphine glucuronide it was 100 ng/mL. The intraday and interday precision and accuracy determinations were <15% coefficient of variation and )15% bias, respec-tively. The method was successfully applied to in vitro enzymatic metabolism studies of buprenorphine.

Simultaneous Quantitation of Buprenorphine and Its Metabolites Using LC–MS

Buprenorphine (BUP) is a partial μ-opioid receptor agonist and κ-opioid receptor antagonist that displays 25–40-fold potent analgesic activity than morphine (1). It appears to

display lower potential for addiction and lesser respiratory de-pression compared to morphine (2). While BUP displays good oral absorption in the gastrointestinal tract, it suffers from very low oral bioavailability because of extensive presystemic metabo-lism in the gut and the liver (3,4). The majority of BUP metabo-lism is through oxidation by cytochrome P450 (CYP) 3A4 and to a lesser extent by CYP2C8 to form norbuprenorphine (NBUP) (Figure 1) (5–7). In addition, BUP and NBUP are also metabo-lized by phase II pathways such as glucuronidation to produce BUP-glucuronide (BUPG) and NBUP-glucuronide (NBUPG), respectively (Figure 1) (8–11). BUP in combination with nalox-one (NX) (Suboxone, Indivior UK Limited) was approved in the United States for the treatment of opioid dependence in 2002.

Early analytical methods for quantitation of BUP included analysis by high performance liquid chromatography (HPLC) coupled to fluorescence detection (12), ultraviolet (UV) detection (13), or use of gas chromatography (GC) (14). In addition, sev-

eral liquid chromatography–mass spectrometry (LC–MS) meth-ods have also been reported for quantitation of BUP in plasma (15–17), serum (18), whole blood (19), and urine (16,20,21). In the previous reports, reversed-phase HPLC has been extensively used for chromatographic separation of BUP and its metabo-lites (16,18–25). Previously developed LC–MS-based methods for quantifying BUP and its metabolites have been reviewed elsewhere (26,27).

A benchtop mass spectrometer similar to a single-quadrupole detector was used in this study. While several methods developed in the past have used reversed-phase HPLC for quantitation of BUP and its metabolites (as stated above), reports for quantitation of BUP and its metabolites using reversed-phase HPLC coupled to this detector have not been reported. To the best of our knowl-edge, we report for the first time the use of reversed-phase HPLC coupled to this benchtop MS detector for quantifying BUP and its three metabolites: NBUP, BUPG, and NBUPG. We believe that as this benchtop MS detector grows in popularity it would be worthwhile and beneficial to report a validated LC–MS method for BUP and its metabolites using it.


The aim of the present study was to de-velop and validate a LC–MS method for simultaneous quantitation of BUP, NBUP, BUPG, and NBUPG using reversed-phase HPLC coupled to a benchtop mass spec-trometer. The method was successfully developed and validated and had a run time of 7.5 min. Further, the method was successfully applied to in vitro micro-somal metabolic studies of BUP.

Experimental Materials

LC-grade solvents such as acetonitrile were purchased from Pharmaco and Aaper. Reference standard solutions of BUP, NBUP, BUPG, and NBUPG were purchased from Cerilliant. NX and BUP for metabolism studies were purchased from Medisca. Ammonium acetate was obtained from EMD Chemicals Ltd. A 100 mm × 4.6 mm, 3-μm dp C18 Alltech Alltima HP column for LC–MS analysis was obtained from Grace Discovery Sci-ences. Human liver microsomes (HLM) for metabolism studies were obtained from Xenotech. NADPH and MgCl2 were purchased from Akron Biotechnology LLC and Fisher Scientific, respectively. Bovine serum albumin (BSA) was pur-chased from Gemini Bio-Products.

Preparation of Calibration Standards

and Quality Control Samples

Commercially available stocks were ob-tained for NBUP, BUPG, and NBUPG at 100 μg/mL in methanol and BUP at 1 mg/mL in methanol. The stocks were stored at -20 °C until the day of the experiment. A solution of 0.2 mg/mL of HLM in 0.1 M potassium phosphate buffer with 0.05% BSA was used as a blank matrix. Blank matrix was used as a diluent for preparing

calibration standards of appropriate con-centrations from the methanolic stocks. The calibration standards for BUP and NBUP were 25, 50, 100, 200, 500, 1000, 2000, and 4000 ng/mL and 100, 200, 500, 1000, 2000, and 4000 ng/mL for BUPG and NBUPG. The quality controls (QC) were also prepared similar to the cali-bration standard solutions. Three levels of QCs with final concentration of 100 (low-QC), 2000 (mid-QC), and 4000 ng/mL (high-QC) for NBUP and BUP and 500 (low-QC), 2000 (mid-QC), and 4000 ng/mL (high-QC) for BUPG and NBUPG were prepared.

Sample Preparation

The samples from the enzymatic me-tabolism studies of BUP (100 μL) were mixed with an equal volume of aceto-nitrile (100 μL) containing NX (328 ng/mL) as the internal standard. The samples were thoroughly vortexed to precipitate protein. Subsequently, the samples were

centrifuged to separate the supernatant. A part of the supernatant (100 μL) was mixed with blank potassium phosphate buffer (no BSA) (100 μL) and the result-ing solution was injected (50 μL) into the LC–MS. The samples were maintained at 5 °C during the analysis.

LC–MS Method Development

HPLC Conditions

The LC system consisted of a model 2695 HPLC system (Waters). All four of the an-alytes (BUP, NBUP, BUPG, and NBUPG) were retained on the C18 column. The LC method included a gradient of 99% A (10% acetonitrile, 90% aqueous, 25 mM ammonium acetate, pH 6.6, 5 μL glacial acetic acid per liter) and 1% B (neat aceto-nitrile) for the initial first min (0–1 min) followed by a gradient from 1% to 50% B over 1.5 min (from 1 to 2.5 min). Subse-quently, the gradient was ramped from 50% to 90% B over 0.5 min (from 2.5 to 3 min) and further, 90% B was maintained

Figure 1: Structures of analytes and internal standard.

HO

OH

N

O

O

O

O

H OH

NHN

OO

O

HO

O

O

O

OH

H

NNH

Glucuronic acid

Glucuronic acid

OH OHOH

HO

BUP BUPG

NBUPG NX (IS)

NBUP

Table I: Gradient Conditions

Time (min) %A* %B† Flow (mL/min)

0.0 99 1 1.0

1.0 99 1 1.0

2.5 50 50 1.0

3.0 10 90 1.0

6.0 10 90 1.0

6.1 99 1 1.0

7.5 99 1 1.0

*25 mM ammonium acetate buffer (pH 6.6) + 10% acetonitrile; †acetonitrile

Table II: Summary of LC–MS parameters

Analyte or IS SIR Transition Cone Voltage (V) tR (min)

BUP (A) 468.60 15 6.91

BUPG (A) 644.70 15 4.51

NBUP (A) 414.50 15 4.53

NBUPG (A) 590.60 15 3.76

NX (IS) 328.38 15 4.66

A = analyte; IS = internal standard


for 3 min (from 3 to 6 min). This was fol-lowed by a re-equilibration to 1% B for 1.5 min (until 7.5 min). The flow rate was constant at 1 mL/min. The column tem-perature during the analysis was 30 °C. The eluent was diverted to waste for the initial 3 min. The total run time was 7.5 min (Table I).

MS Conditions

The MS system was an Acquity QDa mass spectrometer (Waters), which is similar to a single-quadrupole system. The eluent from the analytical column was diverted to the waste for initial 3 min. The eluent from 3 to 7.5 min was injected into the mass spectrometer. The capillary posi-tive voltage was set at 0.8 kV and the probe temperature was 600 °C. Instrument con-trol, acquisition, and data processing were performed using Empower 3 software (Waters). The single ion recording (SIR) and cone voltages for each analyte are noted in Table II. The source, ion block, and sample cone were cleaned weekly.

LC–MS Method Validation

Linearity and Lower

Limit of Quantitation

Each calibration curve consisted of eight different levels of concentrations of cali-bration standards in duplicate for BUP and NBUP and six levels for BUPG and NBUPG. The calibration curves were fit-ted to nonlinear regression using a qua-dratic equation and 1/y2 as the weighting factor. The regression equation was:

y = b0 + b1x + b2x2 [1]

The calibration parameters were calcu-lated by using GraphPad Prism 5 software and the nonlinear regression equation was used to determine the sample concentra-tions. The lowest concentration of the calibration curves was the lower limit of quantitation (LLOQ) for all four analytes.

Precision, Accuracy, Extraction

Recovery, Matrix Effects, and

Autosampler Stability

The method was evaluated for intra-day precision and accuracy for all four of the analytes by analyzing at least five replicates at three concentration levels of QC samples and at LLOQ as well and thereby calculating the percent coefficient

of variation (%CV) and percent bias, re-spectively. Interday precision and accu-racy was evaluated by calculating %CV and %bias, respectively, and analyzing samples at three levels of QCs that were run on four days. Interday precision and accuracy was also calculated at LLOQ on three days. The extraction recovery of the analyte was also measured at three concentration levels of QC sample in at least triplicates by comparing mean peak area obtained from blank matrix solutions spiked with the analyte before protein pre-

cipitation to the mean peak area obtained by spiking with analyte after protein pre-cipitation. The matrix effect was also cal-culated at the three levels of QC sample in at least triplicate by comparing the mean peak area obtained from post protein pre-cipitation spiked blank matrix solutions to that with the mean peak area of external analyte solution. Carryover was evaluated by injecting high QC sample followed by a blank solvent used for sample preparation. The autosampler stability of the analytes was assessed following 14 h of storage in

Figure 2: Representative chromatograms of analytes and internal standard.

(a)

(b)

(c)

(d)

(e)

NBUPG,(M+H)+ = 590.6,t

R = 3.76 min

NBUP,(M+H)+ = 414.5,t

R = 4.53 min

BUPG,(M+H)+ = 644.7,t

R = 4.51 min

BUP,(M+H)+ = 468.6,t

R = 6.91 min

NX,(M+H)+ = 328.4,t

R = 4.66 min

Peak from blank

Inte

nsi

ty

Inte

nsi

tyIn

ten

sity

Inte

nsi

tyIn

ten

sity

Time (min)

Time (min)

Time (min)

Time (min)

Time (min)


the autosampler at 5 °C. The analytes were evaluated at four levels of concentrations: 100, 500, 2000, and 4000 ng/mL followed by a calculation of %CV.

Enzymatic Study

The reaction mixture comprised of 0.2 mg/mL HLM, 1 mM NADPH, 2.5 mM UDPGA, 12.5 mM MgCl2, 8.1 mM sac-charolactone, 31.25 μg/mL alamethicin, 8 μM BUP, and 0.1 M potassium phosphate buffer (pH 7.4) containing 0.05% (w/v) BSA. The microsomes were preactivated by incubation with alamethicin on ice for 20 min. The reaction was initiated by adding UDPGA and NADPH followed by incubation at 37 °C for 30 min. An equal volume of cold acetonitrile containing the

internal standard (328 ng/mL NX) was used to stop the reactions. Subsequently, protein precipitate was separated from the supernatant by centrifugation and the supernatant was stored at -20 °C until further analysis. The incubation was per-formed in triplicates.

Results Optimization of LC–MS Conditions

The chromatographic conditions were optimized on an Alltech Alltima HP C18 column using a mobile phase contain-ing acetonitrile and 25 mM ammonium acetate buffer. All of the analytes were eluted in the 3.6–7 min range. The chro-matographic gradient was optimized such that there was sufficient time for the buf-fer salts to be eluted to the solvent waste (initial 3 min of the run time) followed by a sequential elution of all four analytes in-cluding the internal standard until 7 min. These chromatographic gradient condi-tions produced a quantifiable and repro-ducible peak response for all the analytes and internal standard. The selected ion recording (SIR) used for each analyte was as follows: BUP = 468.6, NBUP = 414.5, BUPG = 644.7, NBUPG = 590.6, and NX = 328.3 (Table II and Figure 2a–e). Blank

samples containing matrix with and with-out an internal standard did not display background peaks for all four analytes. Baseline resolution was achieved between all the analytes except for between NBUP and BUPG. Although baseline resolution was not considered necessary for quan-tification of metabolites because of mass selective detection, we also performed a short study to verify whether there was any bleeding of NBUP into the BUPG channel and thus, whether NBUP con-tributes to the chromatographic peak area of BUPG. We were prompted to perform this study because of the presence of a mass peak of NBUP (m/z = 414) following integration of the chromatographic BUPG peak in the BUPG channel. To investigate this m/z 414 peak, a high concentration solution (4000 ng/mL) of NBUP was in-jected into the LC–MS and the BUPG channel was monitored for the presence of a NBUP chromatographic peak. How-ever, no peak was observed. Further, a low-concentration solution of BUPG (400 ng/mL) along with and without a high-concentration solution (4000 ng/mL) of NBUP were injected onto the LC–MS system. The chromatographic peak areas of BUPG peaks were similar (less than 5%

Table IV: Results of the metabolism study

Analyte Amount (pmol)*

BUP 27.8 ± 3.54

BUPG 133 ± 1.26

NBUP 101 ± 4.40

NBUPG < LLOQ†

*The amounts of analytes are follow-ing 30 min reaction time and in 100 μL of reaction mixture. †LLOQ for NBUPG is 17 pmol for 100 μL of injection volume.

Table III: Summary of precision, accuracy, recovery, and matrix effects for all of the analytes

Analyte

NC Intraday MC Intraday Intraday Interday MC Interday Interday ER ME

ng/mLMean ± SD

(ng/mL)CV (%) Bias (%)

Mean ± SD (ng/mL)

CV (%) Bias (%) (%) (%)

NBUP

25 (LLOQ) 27.8 ± 2.23 8.05 11.2 21.4 ± 2.16 10.1 -14.4

100 90.2 ± 3.01 3.34 -9.80 97.3 ± 4.28 4.40 -2.73 113 8.97

2000 1800 ± 73.2 4.07 -10.0 2020 ± 56.5 2.80 1.00 107 -1.07

4000 3710 ± 153 4.13 -7.25 3960 ± 127 3.21 -1.00 107 -0.60

BUP

25 (LLOQ) 26.1 ± 1.68 6.45 4.4 21.9 ± 1.91 8.73 -12.4

100 95.5 ± 4.33 4.54 -4.50 87.7 ± 8.66 9.88 -12.3 95 6.22

2000 2300 ± 111 4.82 15.0 2130 ± 159 7.46 6.50 95 2.39

4000 4170 ± 219 5.25 4.25 3700 ± 189 5.11 -7.50 101 2.07

NBUPG

100 (LLOQ) 85.9 ± 12.5 14.6 -14.1 92.9 ± 12.2 13.1 -7.11

500 478 ± 15.6 3.25 -4..40 463 ± 61.2 13.2 -7.36 103 11.0

2000 1990 ± 126 6.33 -0.50 2020 ± 80.5 3.99 -1.00 110 -1.02

4000 4040 ± 176 4.35 1.00 3890 ± 115 2.96 -2.75 109 -1.08

BUPG

100 (LLOQ) 91.6 ± 14.9 16.3 -8.40 104 ± 0.55 0.53 3.62

500 464 ± 32.1 6.91 -7.20 498 ± 28.8 5.78 -0.42 97 8.84

2000 1830 ± 81.9 4.47 -8.50 2040 ± 51.0 2.50 2.00 103 -1.03

4000 3630 ± 146 4.02 -9.25 3940 ± 67.4 1.71 -1.48 106 -0.53

NC = nominal concentration, MC = measured concentration, CV = coefficient of variation, ER = extraction recovery, ME = matrix effect. ER of NX = 119% and matrix effect less than ± 17%


difference) with and without the presence of NBUP, indicating that NBUP contrib-utes minimally to the chromatographic peak area of BUPG.

Selection of an Internal Standard

The ideal internal standard for LC–MS analysis is the isotopically labeled ana-lyte. While the use of an isotopically la-beled analyte NBUP-d3 was attempted, a spillover was observed into the NBUP channel, possibly because of an isotopic effect. While the spillover was minimal, NX also displayed a retention time in the 3.6–7 min time period and displayed good sensitivity. Hence, an NX concentration of 328 ng/mL that produced a quantifiable peak response was used as the internal standard for the analysis.

LC–MS Method Validation

Linearity and LLOQ

The calibration curves for the BUP and NBUP displayed good linearity over the concentration range of 25–4000 ng/mL and 100–4000 ng/mL for BUPG and NBUPG. The correlation coefficients (r2) for all calibration curves were greater than 0.99, except one calibration curve for BUP that displayed a correlation coefficient of 0.985. All the calibration curves were run in du-plicate for all the analytes. The calibration curves for NBUP, BUPG, NBUPG, and BUP were fitted by nonlinear regression to a quadratic equation. The LLOQ of NBUP and BUP is 25 ng/mL and that for NBUPG and BUPG is 100 ng/mL. The %CV and %bias for LLOQ was less than 20%.

Precision, Accuracy, Extraction

Recovery, Matrix Effects,

and Autosampler Stability

The precision and accuracy for all four an-alytes were evaluated at three levels of QC and also at the LLOQ. For all four analytes, while the intraday and interday precision ranged from 3.25% to 16.3% and 0.53% to 13.2%, respectively (Table III), the intra- and interday accuracy ranged from -14.1% to 15.0% and -14.4% to 6.50%, respectively (Table III). The matrix effect for all the ana-lytes was less than or equal to 11% (Table III). The recoveries of all analytes following protein precipitation ranged from 95% to 113% (Table III). All of the analytes dis-played a minimal carryover (<1%). The %CV determined at the four concentration

levels in the autosampler stability test was less than 15% for all of the analytes.

Metabolism Study of BUP

BUP concentrations at the end of 30 min were consistent with almost complete conversion to its metabolites. Of the BUP metabolites, the highest concentrations of BUPG were observed followed by NBUP whereas NBUPG was below the LLOQ (Table IV). Successful quantitation of BUP and its metabolites BUPG and NBUP was performed using this analytical method.

ConclusionsAn LC–MS method was developed and validated for the simultaneous quantitative determination of BUP and its three metab-olites (BUPG, NBUP, and NBUPG) with a run time of 7.5 min. The method displayed good linearity and acceptable inter- and intraday precision and accuracy. All of the analytes displayed more than 95% extrac-tion recoveries and matrix effects that were less than or equal to 11%. This method was further successfully applied for the analy-sis of BUP and its metabolites from micro-somal metabolism studies. By diverting to waste for the initial part of the run, no sam-ple extraction was necessary for mass de-tection, and equipment maintenance was minimal. We believe this method would serve as a good reference for laboratories using the benchtop MS detector.

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Anand A. Joshi, Neha V. Maharao, and Phillip M. Gerk are with the Department of Pharmaceutics in the School of Pharmacy at Virginia Commonwealth University in Richmond, Virginia. Direct correspondence to: [email protected] ◾


Tai Van Truong, Nathan L. Porter, Edgar D. Lee, and Robert J. Thomas

In this study, we report on the use of field-portable gas chromatography–mass spectrometry (GC–MS) with solid-phase microextraction, purge-and-trap, thermal desorption, and heated headspace sampling techniques to provide a fast response for in-field analysis of semivolatile organic compounds (SVOCs) in a wide variety of environmental-type samples including potable waters, tea, plants, and road gravel. We demonstrate that this field-portable approach can provide the required sensitivity and selectivity for the effective analysis of SVOCs with very high boiling points such as polycyclic aromatic hydrocarbons (PAHs), pesticides, phenolic compounds, and phthalate esters in a number of different field-based samples, in less than 10 min.

The Applicability of Field-Portable GC–MS for the Rapid Sampling and Measurement of High-Boiling-Point Semivolatile Organic Compounds in Environmental Samples

Over the years, many types of analytical instruments have been reduced to a portable or handheld for-mat to be used in the field, including X-ray f luo-

rescence (XRF), laser induced breakdown spectroscopy (LIBS), Raman, Fourier transform infrared (FT-IR), and near-infrared (NIR) analyzers. However, shrinking a gas chromatography–mass spectrometry (GC–MS) system to a field-portable configuration, while maintaining laboratory analytical performance, is a much greater challenge. Most of the previous attempts have used “point-and-shoot” ap-proaches, which have not required any type of sample prepa-ration or sample introduction accessories. For that reason, the practical value of a field-portable instrument is reduced significantly if it necessitates complex sample preparation or delicate procedures are required to introduce the sample into the gas chromatograph.

In this study, we describe results from field-portable GC–

MS analysis of a wide variety of environmental samples (gas, liquid, solid), including high-boiling, semivolatile organic compounds (SVOCs). The analyses performed includedt�2VBOUJGZJOH�B�NJYUVSF�PG�UFSQFOFTt�%FUFDUJOH�HFPTNJO�JO�QPUBCMF�XBUFSTt�"OBMZ[JOH�QPMZDZDMJD�BSPNBUJD�IZESPDBSCPOT�1")T�JO�BT-

phalt and coal tar–based gravelt�$IBSBDUFSJ[JOH�B�TVJUF�PG�PSHBOPDIMPSJOF�QFTUJDJEFT�JO�

black teat�4DSFFOJOH�GPS�QIFOPMJD�DPNQPVOET�BOE�QIUIBMBUF�FTUFST�

in waterThe rapid sampling techniques used include solid-phase

NJDSPFYUSBDUJPO�41.& �B�OFFEMF�USBQ�GPS�HBTFPVT�TBNQMFT �purge-and-trap and thermal desorption sampling for aque-ous samples, and heated headspace sampling for solid sam-ples. The combination of field-portable GC–MS with rapid sample preparation and introduction techniques enables a


wide variety of field-based assays, in-cluding quantitative studies, and pro-vides actionable results for nonspecial-ist operators in the field.

ExperimentalThe system used in this study was a Torion T-9 portable GC–MS system 1FSLJO&MNFS�XJUI�B�DPNQBDU �CBUUFSZ�operated, rugged, fieldable sampling ac-cessory. The original system and its ap-plicability for field-based analysis have been described previously in the open literature (1,2). However, a number of recent improvements have been made by replacing the conventional capillary column with a low-thermal-mass col-umn bundle that uses direct-contact electrical resistive heating. This column provides identical heat distribution, but nearly eliminates cooler spots of tradi-tional column technology, thus improv-ing the chromatographic separation for SVOCs at the high temperature GC runs required for high-boiling-point compounds.

The mass spectrometer uses a toroi-dal ion-trap configuration, which is

well-suited for miniaturization com-pared to other designs (3). The novel configuration allows for large trap-ping volumes, which result in high ion counts, low noise levels, and good spectral quality. The ion-trap mass ana-lyzer is heated to ~175 °C and operates under vacuum, which results in the electrodes staying clean for long peri-ods of time and reduces the need for frequent maintenance.

Instrumental Conditions

The mass spectrometer operating condi-tions for this investigation are shown in Table I. The GC separating conditions

are described in each subsection on the methodology for each different type of sample matrix.

MethodologyLet’s now take a more detailed look at the methodology for analysis of a suite of different SVOCs, with a wide range of boing points.

The Analysis of Terpenes

Terpenes are a large class of organic compounds, produced by a variety of plants, including conifers, hops, and cannabis with a typical boiling point range of 150–180 °C. They are the pri-

Figure 1: The four terpene analytes were extracted by half–half SPME (PDMS–DVB 65 μm fibers) at room temperature (22 °C) for 15 min, before being injected into the GC–MS system.

Figure 2: Total ion chromatogram of four terpene compounds. Peaks: 1 = (+)-α-pinene, 2 = myrcene, 3 = (+)-α-limonene, 4 = isolongifolene.

1/2

1/2

4000

1

2

3

4

2000

TIC

0

20 40 60 80 100 120 140 160

Table I: Mass spectrometer parameters

Mass spectrometer Toroidal ion trap

Ionization source Electron capture

MS operating temperature 175 °C

Mass range 41–500 amu

Resolution <1 amu at 300 amu

MS scan rate 10–15 scans/s

Detector Electron multiplier

Table II: The chromatographic separation conditions for terpenes (gas chromatographic separation conditions)

Sample delivery Coil wire filament injection

Injection type Split–splitless

Injector temperature 300 °C

Transfer line temperature 280 °C

Trap temperature 200 °C

Column (Restek)5 m × 0.1 mm, 4-μm df MXT-5 (low-polar-ity phase diphenyl dimethyl polysiloxane )

Initial temperature and hold time 50 °C for 10 s

Temperature ramp rate 2 °C/s

Final temperature and hold time 280 °C for 50 s


mary constituents of the essential oils of many types of plants and f lowers widely used as fragrances in perfum-ery, as well as for medicinal purposes.

Synthetic variations and derivatives of natural terpenes are also used for a va-riety of aromas and flavors used as food additives. Therefore, to exemplify the

capability of this technology, four ter-pene compounds were spiked into 200 mL of 0.6% sodium chloride in water. The analytes were then extracted using IBMGoIBMG�41.&�QPMZEJNFUIZMTJMPYBOFoEJWJOZMCFO[FOF�1%.4o%7#��N�fibers at room temperature (22 °C) for 15 min without shaking or vibrating. With this sampling approach, the fiber is placed half in the head space and half immersed into the liquid phase of the sample, as shown in Figure 1.

This sample was then injected into the GC–MS system using the chro-matographic separating conditions shown in Table II.

The total ion chromatogram (TIC) of the four terpenes ([+]-α-pinene, myrcene, [+]-α-limonene, and isolon-gifolene), is shown in Figure 2.

"�GPVS�QPJOU�DBMJCSBUJPO�HSBQI�XBT�then generated for the four terpene compounds. The concentrations of the standards and the respective calibra-tion plots with correlation coefficients (R2) are shown in Figure 3. It should be noted that the estimated detection limit for the four compounds was 20 ppt, which was based on the statistical analysis of multiple replicates of the lowest standard (sample 1).

Analysis of Geosmin

in Drinking Water

Geosmin is an organic compound pro-duced by a variety of microorganisms and bacteria. It has a distinct earthy flavor and aroma and is responsible for the earthy taste of beets and the strong scent that occurs in the air when rain falls after a dry spell of weather. Geos-min is produced by several classes of microbes, including cyanobacteria and actinobacteria, and is released when these microbes die. Communities whose water supplies depend on surface water can periodically experience epi-sodes of unpleasant-tasting water when a sharp drop in the population of these bacteria releases geosmin into the local water supply (9). Chemically, it is a bicy-clic alcohol with a formula of C12H22O, and a derivative of decahydronaphtha-lene, commonly known as decalin. Its boiling point is ~270 °C (10).

For this study, 20 ppt of geosmin was spiked into 500 mL of a water sample.

Figure 3: Calibration plots of the four terpene compounds.

R2

Concentration (ppb)Name

Alpha pinene

Alpha limonene

Myrcene

Isolongfolene

Table IV: GC system operating conditions for the separation of PAHs in gravel samples

Sample delivery Needle trap

Injection type Splitless with pre-run split closed






Table III: The chromatographic separation conditions for geosmin in water (GC separa-tion conditions)

Sample delivery Coil wire filament injection


Split injection times:

10:1 split on 20 s

10:1 split off 40 s

50:1 split on 40 s

50:1 split off 80 s




Column (Restek)5 m × 0.1 mm, 4-μm df MXT-5 (low-polar-ity phase diphenyl dimethyl polysiloxane )




Carrier gas inlet pressure 26 psi


Without any pretreatment step, the water sample was then passed through QPMZEJNFUIZMTJMPYBOF�1%.4�QBSUJDMFT��o��N�TJ[F�QBDLFE�JO�B�EFBDUJ-vated stainless steel solid-phase extrac-UJPO�41&�EFTPSQUJPO�UVCF�BU�BNCJFOU�temperature using a flow rate of 25–35 mL/min delivered by a vacuum pump. The target analyte was then transferred JOUP�B�1%.4�OFFEMF�USBQ�VTJOH�UIF�JO-strument’s thermal desorber system. The desorption step was carried out at 200 °C at 6 mL/min for 10 min, using helium carrier gas. Sample introduc-tion into the GC–MS system using the needle trap was conducted at 270 °C for ��T��"�TDIFNBUJD�PG�UIF�TBNQMF�EFMJWFSZ�approach is shown in Figure 4.

The chromatographic separation conditions are shown in Table III. The TIC of the separation is shown in Fig-ure 5, together with the extracted ion chromatogram (RIC), showing the parent molecular ion and the associ-ated fragments of geosmin, which is confirmed by the National Institute of Standards and Technology (NIST) ref-erence mass spectrum underneath it. Figure 6 shows the deconvoluted chro-matogram and mass spectrum, dem-onstrating that the 20 ppt geosmin is well-separated using the instrument’s deconvolution algorithm. Based on the statistical analysis of the geosmin cali-bration, it was estimated that the de-tection limit was in the order of single-digit parts-per-trillion levels.

PAHs in Asphalt and Coal

Tar-Based Gravel Samples

Road and parking lot surfaces are typica l ly made from asphalt and coal tar products that contain high levels of carbonaceous compounds. For this reason it is very important to know the composition of the poly-cyclic aromatic hydrocarbon levels in the gravel samples used in the road TVSGBDF�QSFQBSBUJPO�QSPDFTT��1")T�JO�these types of samples typically range from napthalene up to dibenz[a,h]an-thracene with boiling points between 220 °C and 525 °C. It is well recog-nized that high-temperature program methods are normally required for the determination of high-boiling-point semivolatile analytes such as

1")T�BOE�QFTUJDJEFT�JO�WBSJPVT�FO-vironmental sample matrices, which can make it extremely diff icult to separate these compounds with good spectral quality (4–8).

So for this study, 40 g of the gravel samples was spiked with stock stan-dard solutions to make calibration standards of 0.05, 0.25, 0.5, and 1.0 QQN�PG�UIF�1")�BOBMZUFT��5IF�TBNQMFT�were then extracted with a mixture of dichloromethane (5 mL) and water

(~15 mL) by hand shaking for about 2–3 min. The liquid phase was then transferred to another vial to let the two phases separate out. For some of the samples, preconcentration was necessary to improve the detection. This preconcentration was achieved by placing 1 mL of the organic phase into a 2-mL vial and allowing the solvent to evaporate to get a suitable WPMVNF�GPS�UIF�NFBTVSFNFOU��"��-�aliquot of the sample in the organic

Figure 4: The sampling procedure and thermal desorption step for the analysis of geosmin by GC–MS.

PTFE-siliconeseptum

O-ring

PDMS tube

SPE holder

500-mLcontainer

1000-mLcontainer

Vacuum

Sample

Table VI: The GC separation conditions for a suite of organochlorine pesticides in tea


Injector temperature 290 °C, 20 s splitless with prerun split closed



Column (Restek)5 m × 0.1 mm, 4-μm df MXT-5 (low-polarity

phase diphenyl dimethyl polysiloxane )




Table V: Concentration ranges of the organochlorine pesticides used in this study

No. NameSample A

Concentration, (ppb)

Sample B Concentration,

(ppb)

Sample C Concentration,

(ppb)

1 Benfluralin 4 40 200

2 Lindan 2 20 100

3 Heptachlor 2 20 100

4 Chlorthal dimethyl 4 40 200

5 Heptachlor epoxide 2 20 100

6 Endrin 2 20 100

7 Methoxychlor 2 20 100


phase was then introduced into the glass tube using a syringe and the sol-vent was eliminated using a vacuum pump or air compressor. The target analytes then were transferred into UIF�1%.4�OFFEMF�USBQ�VTJOH�B�TBNQMF�displacement approach at 300 °C for 5 min with a purging f low rate of 30 mL/min. The GC conditions for the separation are shown in Table IV. The TIC of the separation is seen in Figure 7 and clearly shows that high-molecular-weight, high-boiling-point 1")T �TVDI�BT�CFO[P<ghi]perylene and benzo[b]f luoranthene, have been sep-

arated and detected.

Organochlorine Pesticides in Tea

For this investigation, 20 g of dry black tea was steeped in 500 mL of hot water and left for four days at 22 oC. Then a 15 mL sample was spiked at three DPODFOUSBUJPO�MFWFMT�" �# �$�PG�TFWFO�different organochlorine insecticides shown in Table V, with boiling points ranging between 275 °C and 425 °C.

The sample preparation was per-GPSNFE�VTJOH� 41.&� G JCFST� DPOEJ-tioned at 220 °C for 60 min, and the immersion extraction process was

carried out at 22 °C for 10 min by stirring with a bar mixer at 300 rpm. The fibers were then rinsed after the extraction with deionized water for 10 s without vibration. The sample was injected at 270 °C for 40 s (15 s split-MFTT��"GUFS�JOKFDUJPO�UIF�GJCFST�XFSF�washed with deionized water for 30 s and conditioned at 270 °C for 30 s (in the GC injector). The GC separation conditions are shown in Table VI.

The TIC of sample B is shown in Figure 8.

General Screening Tool for SVOCsThis portable GC–MS approach can also be used as a general screening tool for SVOCs in water using micro l iquid extraction and a coi l wire filament. The experiments were car-ried out using tap water spiked with SVOCs at concentrations from low-parts-per-billion to sub-parts-per-NJMMJPO�MFWFMT��"�TNBMM�BNPVOU��o0.5 mL) of suitable solvent, such as dichloromethane, hexane, pentane, or acetone, is used for extraction. Manual shaking and salting-out may be applied using sodium chloride at 0.5–3% to speed up the extracting process. The extraction is performed for a few minutes and the solvent containing the analytes is then ap-plied on to the coil, or if necessary, concentrated by letting the solvent evaporate after transferring it to a small vial. Sample introduction using the coil is performed after solvent on the coil is evaporated. The screen-ing tests were carried out with mix-UVSFT�PG�1")T �QIFOPMJD�DPNQPVOET �phthalate esters, organochlorine, organophosphorus, and pyrethroid pesticides and herbicides. However, because we have previously shown UIF�TFQBSBUJPO�PG�1")T�BOE�WBSJPVT�pesticides, we only show representa-tive data for the phenolic compounds and the phthalate esters here. The chromatographic separation condi-tions for the phenols and phthalate esters are shown in Table VII.

Figure 9 shows the TIC of the sepa-ration of all the phenolic compounds in water, with phenol (C 6H5OH) identified with the bold red arrow. The RIC of phenol is shown on the

Figure 5: Total ion (TIC) and extracted ion chromatograms (RIC) of geosmin and its MS fragments in a water sample, identified and confirmed by the mass spectrum from the NIST reference library.

Table VII: The chromatographic separation conditions for the screening of nine phenolic compounds and six phthalate esters

GC Parameter Phenols Phthalate Esters

Sample deliveryCoil wire filament

injectionCoil wire filament

injection

Injection type Split–splitless Split–splitless

Injector temperature 290 °C 300 °C

Transfer line temperature 270 °C 280 °C

Trap temperature 200 °C 200 °C

Initial temperature and hold time 50 °C for 10 s 50 °C for 10 s

Temperature ramp rate 2 °C/s 2 °C/s

Final temperature and hold time 290 °C for 60 s 300 °C for 60 s

Note: An additional 14 general pesticides, six herbicides, and 10 insecticides were screened using similar conditions


right with the reference mass spec-trum from the NIST library below it. The phenols identified from left to right are phenol, 4-methylphe-nol, 2-nitrophenol, 3,5-dichloro-phenol, 4-chloro-3-methylphenol, 2,4,6-trichlorophenol, 4-nitrophe-nol, 2-methyl-4,6-dinitrophenol, and pentachlorophenol.

The group of phthalate esters is shown in Figure 10, with dimethyl phthalate (C10H10O4) shown with the bold red arrow. The RIC of dimethyl phthalate is shown on the right, with the reference mass spectrum from the NIST library below it. The phthalate esters identified from left to right are dimethyl phthalate, diethyl phthal-ate, dibutyl phthalate, benzyl butyl phthalate, diisooctyl phthalate, and di-n-octyl phthalate.

The total running time for these screening tests for both phenols and phthalate esters was less than 3 min. Ion molecule chemistry occurred to some degree on both types of sam-ples, so absolute identification was confirmed using the NIST library TFBSDI�DBQBCJMJUZ��"MUIPVHI�UIF�QFBL�capacities are relatively low for these separations, the deconvolution algo-rithm helped to separate and identify the analytes with greater accuracy. %ZOBNJD�SBOHFT�BOE�EFUFDUJPO�MJN-its in real samples will be determined and presented in a future study.

ConclusionThere is a growing demand for the analysis of trace levels of volatile and semivolatile organic compounds in air, water, and solid matrix samples under harsh conditions in remote, field-based locations. This study has demonstrated that it is now possible to achieve laboratory-grade perfor-mance with portable GC–MS com-bined with rapid sample preparation or introduction techniques. This combination enables a wide variety of environmental-based assays for both quantitative and qualitative screen-ing purposes, which can provide fast, actionable data for nontechnical and inexperienced operators in the field.

It has been demonstrated that the approach used in this study has

detected SVOCs relevant to ter-penes, plant protection chemicals, and polycyclic aromatic hydrocar-

CPOT� 1")T �XJUI�WFSZ�IJHI�CPJM-ing points (up to 550 °C), at low parts-per-trillion concentrations in

Figure 7: TIC of a 250 ppb spiked sample of PAHs.

1.3e5

1.2e5

1.1e5

1.0e5

0.9e5

0.8e5

0.7e5

0.6e5

0.5e5

0.4e5

0.3e5

0.2e5

0.1e5

0.0

50 100 150 200 250

Ind

en

o[1

,2,3-cd

]pyre

ne

Dib

en

z[ah

]an

thra

cen

eB

en

zo

[ghi]

pyery

len

e

Ben

zo

[a]p

yre

neA

nth

race

ne

Naphthalene

Acenaphthene

Fluorene

Acenaphthylene

Phenanthrene

Benz[a]anthracene

Pyrene

FluoroantheneIS

6ChryseneIS 5IS

4IS 3

IS 2

IS 1

Ben

zo

[b]fl

uo

ran

then

e/B

en

zo

[k]fl

uo

ran

then

e

TIC

Figure 6: The deconvoluted chromatogram and mass spectrum demonstrating that the geosmin is well-separated using the instrument’s deconvolution algorithm.

Figure 8: The TIC of the seven organochlorine pesticides in sample B (identity and concentration of pesticide shown in Table V).

9000

8000

7000

6000

5000

TIC

4000

3000

2000

1000

60 80 100 120 140 160 180 200 220 240 260 280

1

2

3

4

5 6 7


under 10 min total analysis time. It has also been shown that the detec-tion of natural compounds such as geosmin can be detected in water at low parts-per-tril lion levels. In addition, the screening of phenolic compounds and phthalate esters in drinking water can be carried out at MPX�QBSUT�QFS�CJMMJPO�MFWFMT��"T�B�SF-sult, the use of portable GC–MS and associated sampling techniques pro-

vide the required sensitivity, selectiv-ity, and speed of analysis for the ef-fective analysis of high-boiling-point SVOCs in the field.

References(1) J.A. Contreras et al., Journal of Amer-

ican Society of Mass Spectrometry, 19(10), 1425–1414, (2008).

(2) T.V. Truong et al., Scientia Chromato-graphica, 6(1), 13–26, (2014).

(3) Product Note: Torion T-9 portable GC/MS, PerkinElmer Inc., Shelton, CT: http://torion.com/fileadmin/media/documents/brochures/To-rion_T9_GCMS_ProductNote.pdf.

(4) SW-846, Test Method 8275A: Semi-volatile Organic Compounds in Soil/Sludges and Solid Wastes Using Thermal Extraction/Gas Chromatog-raphy/Mass Spectrometry (TE/GC/MS): United States Environmental Protection Agency, December, 1996: https://www.epa.gov/hw-sw846/sw-846-test-method-8275a-semivolatile-organic-compounds-soilsludges-and-solid-wastes-using.

(5) Test Method 8141A: The Analysis of Organophosphorus Pesticide Compounds by GC Capillary Column Technology, United States Environ-mental Protection Agency, Septem-ber, 1994: https://www3.epa.gov/wastes/hazard/testmethods/sw846/pdfs/Method%208141A,%20Revi-sion%201%20-%201994.pdf.

(6) U.S. Environmental Protection Agency, Compendium Method TO-13A, Determination of Polycyclic Aromatic Hydrocarbons (PAHs) in Ambient Air Using Gas Chromatog-raphy/Mass Spectrometry (GC/MS). Office of Research and Develop-ment, Cincinnati, OH. March, 1999: https://www3.epa.gov/ttnamti1/files/ambient/airtox/to-13arr.pdf.

(7) R. Provost, L. Marotta, and R. Thomas, LCGC North Am. 32(10), 810–818 (2014).

(8) L. Marotta, S. Varisco, M. Snow, T. Kwoka, and R. Thomas, LCGC North Am. 34(3), 214–220 (2016).

(9) N.N. Gerber and H A. Lechevalier, Appl. Microbiol. 13(6), 935–938 (1965).

(10) T. Manickum and W. John, Hydrol.: Curr. Res. 2(3), 1–10 (2012).

Tai Van Truong and Nathan L. Porter are senior scientists for portable MS and GC–MS at PerkinElmer in American Fork, Utah. Edgar D. Lee is a director of research for portable MS and GC–MS at PerkinElmer. Robert J. Thomas is the principal consultant at Scientific Solutions in Gaithersburg, Maryland. Direct correspondence to: [email protected] ◾

Figure 9: The TIC of the separation of all the phenolic compounds in water with phenol (C6H5OH) identified with the bold red arrow. The XIC of phenol is shown on the right, with the reference mass spectrum from the NIST library below it. The full suite of phenols going from left to right includes phenol, 4-methylphenol, 2-nitrophenol, 3,5-dichlorophenol, 4-chloro-3-methylphenol, 2,4,6-trichlorophenol, 4-nitrophenol, 2-methyl-4,6-dinitrophenol, and pentachlorophenol.

Time (s)60 80

22,000

20,000

18,000

16,000

14,000

TIC 12,000

10,000

8,000

6,000

4,000

2,000

0100

Figure 10: The TIC of the separation of a group of phthalate esters with dimethyl phthalate (C10H10O4) shown with a bold red arrow. The XIC of dimethyl phthalate is shown on the right with the reference mass spectrum from the NIST library below it. The full suite of phthalate esters going from left to right includes dimethyl phthalate, diethyl phthalate, dibutyl phthalate, benzyl butyl phthalate, diisooctyl phthalate, and di-n-octyl phthalate.

3.5e5

3.0e5

2.5e5

2.0e5

1.5e5

1.0e5

0.5e5

80 100 120

Time (s)

140

TIC


P. Sambasivarao, Raman Batheja, N. Subbarao, S. Ashma, K. Ashwini, and M. Mupeksha

Moxidectin formulations help reduce hair loss and irritation because of parasite worms in animals. The estimation of moxidectin in hair is important to evaluate therapeutic levels, distribution, and accumulation. The estimation is also useful to evaluate harm to birds when they eat animal hair. Hence, moxidectin estimation is required for pharmacokinetic as well as environmental exposure studies. The objective of the present work is to develop a rapid, selective method for the estimation of moxidectin in cattle hair by liquid chromatography–tandem mass spectrometry (LC–MS-MS).

Sensitive, Rapid Estimation of Moxidectin in Cattle Hair by LC–MS-MS

M oxidectin is a semisynthetic derivative of nemadectin which is produced by fermentation of Streptomyces

cyano-griseus. The molecular formula for moxidectin is C37H53NO8 and its molecular weight is 639.819 g/mol. The chemi-cal structure of moxidectin is presented is Figure 1. Moxidectin is an anthelmintic drug that kills parasitic worms (helminths), and is used for the prevention and control of heartworm and intestinal worms. It can be found in treatments prescribed for animals such as dogs, cats, horses, cattle, and sheep. Application methods for moxidectin vary by treatment, and include oral, topical, and inject-able solutions (1). Moxidectin is also used in products to treat horses for large and small strongyles, encysted cyathostomes, ascarids, pinworms, hair worms, largemouth stomach worms, and horse stomach bots (2). Moxidectin is not expected to have an adverse effect on hair-eating birds. In one day, hair-eating birds would have to consume many times their weight in cattle hair with moxidectin residues to be exposed to potentially toxic levels of moxidectin (3). Moxidectin 10% long-acting formulations are injected subcutane-ously, at a concentration of 0.5 mL/50 kg body weight. Moxidectin is absorbed following subcutaneous injection with maximum blood concentrations being achieved 24–48 h post injection. The drug is distributed throughout the body tissues including hair, but because of its lipophilicity it is concentrated mainly in the fat. Moxidectin undergoes limited biotransformation by hydroxylation in the body. The only significant route of excretion is via the feces.

Moxidectin formulations help to reduce hair loss and irritation because of parasite worms in animals. So the estimation of mox-idectin in hair is important to evaluate therapeutic levels, distribu-tion, and accumulation. The estimation is also useful to evaluate harm to birds when they eat animal hair. Hence, moxidectin esti-

mation is required for pharmacokinetic as well as environmental exposure studies.

Moxidectin is used extensively for animals to treat parasitic worms, and many manufacturers are creating moxidectin formula-tions. Formulators should evaluate the pharmacokinetic properties of moxidectin in plasma as their primary objective and the distri-bution and accumulation of moxidectin in body tissues including hair as a secondary objective. The primary objective, evaluation of the pharmacokinetic properties, will provide the bioavailability of the formulation whereas the secondary objective will explain how the drug is distributed and its accumulation status.

Many methods are available for the determination of moxidec-tin in plasma (4–8). These methods seem so simple because the industry has hands-on-experience for the determination of drugs in plasma by liquid chromatography–tandem mass spectrometry (LC–MS-MS) and the methods do not require special reagents, in general. Hair analysis, on the other hand, is not a primary focus except in forensics or environmental studies in which analysts use traditional instrumentation such as ultraviolet (UV) spectroscopy, colorimetry, and gas chromatography (GC). Method development for hair analysis involves many challenges, including endogenous interferences, high matrix effects, complex processing steps like hair weighting, hair cutting to micrometer-size pieces, digestion with special reagents, incubation, selection of extraction solvents, as well as preparation of calibrators and controls. The work described in this article is the first ever bioanalytical method for the determina-tion of moxidectin in cattle hair. The first challenge in this work is the selection of a matrix for the preparation of calibration curve standards and quality control samples; because direct hair cannot be used as the matrix, one should opt for surrogate procedures.


Therefore, required equivalent spiking solutions were added to preweighed blank hair, then methanol was added for sonica-tion, followed by evaporation to dryness. The second challenge was the selection of a buffer concentration and volume for the digestion of the hair samples; extensive work was conducted to optimize Soren-son’s buffer concentration and the volume per sample used. The third challenge was the selection of the temperature for incuba-tion, and the last challenge was the selection of an extraction solvent. The incubator time

and temperature were optimized for greater recovery and the extraction solvent was op-timized to reduce interferences without los-ing recovery.

We successfully achieved our target method by overcoming all challenges one by one. In the current study, we report a method for the estimation of moxidectin in hair by LC–MS-MS which is selective and rapid. This method facilitates fast analysis. The method can be applied to hair analysis for the evaluation of moxidectin concentra-tion levels in hair.

Experimental Material

Working standards were obtained from VerGo Pharma Research Labs. Purified water was from taken from an in-house Milli-Q gradient water purification sys-tem (EMD Millipore). Methanol, methyl tert-butyl ether (MTBE), ammonium formate, dibasic sodium phosphate, and monobasic potassium phosphate were purchased from Rankem. Blank cattle matrix was supplied by RLS.

Stock Solutions, Calibration

Curve Solutions, and Quality

Control Spiking Solutions

Two 1-mg/mL stock solutions for mox-idectin and one for oxcarbazepine were prepared by accurately weighting work-ing standards on a microbalance. The standards were dissolved in methanol and stored at refrigerator maintained at 2–8 °C. A solution of 80% methanol in water was used as further diluent. Cali-bration curve standard spiking solutions were prepared in the range of 2.560–107.145 ng/mL using diluent along with four quality control levels at the lower limit of quantification quality con-trol (LLOQQC), lower quality control (LQC), middle quality control (MQC), and high quality control (HQC). Spiking solutions were stored in a refrigerator for long-term storage.

Preparation of Final Standards

The screened blank cattle hair was weighed and spiked with the required spiking solution prepared in diluent. The spiking was performed in weighted blank hair to get final calibration curve standard concentrations of 0.026 to 1.071 ng/mg, LLOQQC of 0.028 ng/mg, LQC of 0.076 ng/mg, MQC of 0.444 ng/mg, and HQC of 0.766 ng/mg. Then 0.250 mL of methanol was added for proper vortexing of the content. The content was vortexed for 5.0 min and sonicated for 5 min, centrifuged at 4000 rpm for 10 min at 5.00 °C, and evaporated for approximately 20 min at 40.0 °C until dryness. Dried samples were stored at -70 °C until analysis.

Sample Processing

The sample processing steps are de-scribed in Table I.

Figure 1: Moxidectin chemical structure.

Figure 2: Gradient chart.

O

O

O

OO

O

OH

OH

N

50

40

30

20Pu

mp

B

Time (min)

10

01 2 3 4 5 6 7

Figure 3: Sample chromatogram of blank.

65

60

55

50

45

40

35

30

25

20

15

10

Inte

nsi

ty (

cps)

5

00.5

0.24 0.430.96

1.11 1.381.54

1.73 1.912.062.22

2.392.662.74

2.94

3.27

3.08

3.57 3.67 3.944.06 4.19

1.0 1.5 2.0 2.5

Time (min)

3.0 3.5 4.0


LC–MS Conditions

The LC conditions are described in Table II. A gradient chart is presented in Figure 2. The mass spectrometer conditions are described in Table III.

Results and Discussion Selectivity of the Method

The selectivity of the method was evaluated using six hair blanks of individual cattle. Blank and LLOQ were processed from each hair lot. Interferences at analyte and internal standard retention times were compared with that of respective LLOQs. No sig-nificant endogenous interferences were observed at the retention times of either the analyte or internal standard. Interference is acceptable up to 20% of LLOQ area for analytes and 5% for the internal standard. Blank sample and LLOQ sample chromato-grams are presented in Figures 3 and 4.

Internal Standard Normalized Matrix Factor

The internal standard matrix factor was evaluated for the method by calculating the ratio of analyte and internal standard response at LQC and HQC levels. The internal standard nor-malized matrix factor was found between 0.85 to 1.15 and %CV for the internal standard matrix factor was 5.4 at LQC and 6.7 at HQC levels. Six post-extraction blank samples for each LQC and HQC were spiked with respective QC level spiking solution (mixture of analyte and internal standard); these samples were analyzed along with equivalent aqueous samples. The experi-ment is considered acceptable when the %CV for the internal standard normalized matrix factor is less than 15%.

Linearity

The linearity of the calibration curve was evaluated using three calibration curve sets (eight standards) on two different days. Au-tomatic integrations were conducted on Analyst 1.6.2 software for linear regression using 1/x2 weighting factor; back calcula-tions were done using the formula in equation 1:

y = mx + c [1]

where x is an unknown sample concentration, y is the area ratio of analyte versus the internal standard, m is the slope of the curve, and c is the intercept of the curve. The correlation coefficient, r2, was calculated for each curve and found more than 0.98 regression value each time. A sample calibration curve is presented in Figure 5.

Precision and Accuracy

Intra- and interday precision and accuracy were evaluated using four levels (LLOQ, LQC, MQC, and HQC) of quality control samples covering the calibration curve range. The six replicates of each QC level sample were processed under a fresh calibra-tion curve. Ruggedness of the method was tested for analyst and analytical column change. Interday precision ranged from 0.9% to 11.7% whereas interday accuracy found to be between 90.7% to 105.5%. Calibration curve standard or quality control samples are acceptable when the %nominal is within ±15% to the nomi-nal concentration except LLOQ where it is ±20%. Precision is accepted when the %CV is less than 15% for all standards and

Table I: Sample processing procedure

Step Number Procedure

1Retrieve required samples from deep freezer and keep on bench to attain room tempera-ture

2Weigh 25 mg sample and transfer into the prelabeled tubes

3Add 20 μL of internal standard working solution to all the samples except blank sample, to which 20 μL of diluent should be added and vortexed

4Add 500 μL Sorenson’s buffer to all samples and vortex

5Keep the samples at 40.0 °C for 90 min in the laboratory oven

6Remove the samples from the laboratory oven and add 2.500 mL of MTBE to all samples and vortex for 5.00 min on a shaker

7Centrifuge the samples for 10.00 min at 5.0 ± 2.0 °C at 4000 rpm

8Withdraw and transfer 1.500 mL of the super-natant organic liquid into the prelabeled tube

9Dry the sample using a nitrogen evaporator at 40.0 ± 2.0 °C for ~15 min or until dry

10Reconstitute the samples with 0.500 mL of reconstitution solution and vortex it for 10 s

11Transfer the sample into prelabeled HPLC vial for analysis

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samples except LLOQ where it is ±20%. A calibration curve is accepted when 75% of a standard meeting the acceptance criteria

including LLOQ and ULOQ, no two con-secutive standards are excluded, and r2 is greater than 0.98.

Stability of Analyte

The six LQC and six HQC samples are subjected to different storage and pro-cessing conditions including three freeze–thaw cycles for hair samples, 6 h at ambi-ent temperature for hair samples, 1 h at 40

°C for processed samples, 2 h at ambient temperature for processed samples, and 24 h at 5 °C for reconstituted samples. Samples were processed and analyzed under a newly prepared calibration curve prepared from freshly prepared stock solution. Back-calculated quality control values were used to calculate %nominal for each level and were found to be within acceptable limits. Stability will be accept-able when the %nominal is found to be ±15% to the nominal concentrations.

ConclusionA rapid, selective method for the esti-mation of moxidectin in cattle hair by LC–MS-MS using oxcarbazepine as an internal standard has been developed and validated. Hair samples were subjected to liquid–liquid extraction using Sorenson’s buffer followed by MTBE. Multiple reac-tion monitoring (MRM) detection in posi-tive mode at unit resolution was opted and separation was achieved on a C18 column with a methanol–10 mM ammonium for-mate mobile-phase gradient. The calibra-tion curve for the range 0.026–1.000 ng/mg was linear with regression greater than 0.98. The method was selective for analytes and internal standards from any endogenous interferences. The matrix fac-tor was found to be between 0.85 and 1.15 for all tested blank lots. Other validation parameters revealed that the method is re-liable, reproducible, and accurate. Stability evaluation proved that moxidectin is stable for different storage and processing condi-tions. The method was applied to estimate moxidectin in six cattle hair samples.

References(1) https://en.wikipedia.org/wiki/Moxidec-

tin, accessed on April 19, 2016.(2) Moxidectin question and answers for

pet owners by Bayer health care posted on www.Moxidectin facts.com, ac-cessed on April 19, 2016.

(3) Environmental assessment - Cydectin Moxidectin 0.5% pour-on for cattle, June 1997, Z154314.

Figure 5: Sample calibration curve.

0.24

0.22

0.20

0.18

0.16

0.14

0.12

0.10

0.08

0.06

0.04

0.02

0.000.05 0.10 0.15 0.20 0.25 0.30 0.35 0.40 0.45 0.50 0.55 0.60

Analyte concentration/IS concentration

An

aly

te a

rea

/IS a

rea

0.65 0.70 0.75 0.80 0.85 0.90 0.95 1.00

Table II: LC conditions

Column 100 mm × 4.6 mm, 5-μm Kinetex EVO C18 100A

Flow rate 1.000 mL/min

Injection volume 5 μL

Column temperature 40.0 °C

Autosampler temperature 5.0 °C

Mobile phase A: methanol; B: 10 mM ammonium formate

Mobile-phase gradient program

Module Time Event Parameter Ratio

Controller 0.01 Start — 60:40

Pumps 1.50 Pump B conc. 40 60:40





Controller 4.50 Stop — 60:40

Retention timeMoxidectin: 3.31 min

Oxcarbazepine: 1.36 min

Figure 4: Sample chromatogram of LLOQ.

1400

1300

1200

1100

1000

900

800

Inte

nsi

ty (

cps)

700

600

500

400

300

200

100

00.5 1.0 1.5 2.0 2.5

Time (min)3.5

3.26

4.03.0


(4) J. Dupuy, J.F. Sutra, and M. Alvineirie, Veterinary Parasitology 147(3-4), 252–257 (2007). DOI: 10.1016/j.vet-par.2007.05.002.

(5) M. Alvinerie, J.F. Sutra, M. Badri, and P. Galtier, J. Chromatogr B Biomed. Appl. 674(1), 119–124 (1995).

(6) M. Alvinerie, E. Escudero, J.F. Sutra, C. Eeckhoutte, and P. Galtier, Vet. Res. 29(2), 113–118 (1998)

(7) D.C. Hughes, K. Fraser, C.M. Miller, and D.M. Leathwick, Proceedings of the New Zealand Society of Animal Production 73, 180–182 (2013).

(8) Determination of Ivermectin, Doramec-tin, and Moxidectin by HPLC, United States Department of Agriculture, http://www.fsis.usda.gov/wps/wcm/

connect/ 87680e50-d76b-407b-9d94-d2ecc37b3cd0/CLG_AVR_04.pdf?MOD=AJPERES.

(9) A. Lifschitz, G. Virkel, F. Imperiale, J.F. Sutra, P. Galtier, C. Lanusse, and M. Alvinerie, J. Vet. Pharmacol. Ther. 22(4), 266–273 (1999).

(10) J. Sallovitz, A. Lifschitz, F. Imperiale, A. Pis, G. Virkel and C. Lanusse, The Veteri-nary Journal 164, 47–53 (2002).

P. Sambasivarao, Raman Batheja, N. Subbarao, S. Ashma, K. Ashwini, and M. Mupeksha are with the bioanalytical department at VerGo Clinicals in Corlim, India. Direct correspondence to: [email protected] ◾

Table IV: Precision and accuracy results

Mean (ng/mg) 0.0229 0.0650 0.4746 0.8178

SD 0.00144 0.00766 0.02207 0.03704

%CV 6.29 11.78 4.65 4.53

%Nominal 91.60 103.17 103.40 101.09

Nominal concentration (ng/mg) 0.025 0.063 0.459 0.809

Table III: Mass spectrometer conditions

Description Q1 Mass (m/z) Q3 Mass (m/z) Dwell Time (ms) DP (v) EP (v) CE (v) CXP (v)

Moxidectin 640.45 528.5 + 498.5 200 59.4 10 13.8 17.5

Oxcarbazepine 253.10 180.1 200 80.0 10 43.0 11.0

Source Parameters

CUR (psi) CAD (psi) Ion Spray Voltage (kv) Temp. (oC) Gas 1 (psi) Gas 2 (psi)

20.00 3.00 5000.00 500.00 50.00 50.00

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Cindy Delonas, Spectroscopy, Associate Editor

We present a brief review of this year’s ASMS conference, which took place June 5–9, 2016, in San Antonio, Texas.

Review of the 64th Conference on Mass Spectrometry and Allied Topics

T he 64th Conference on Mass Spectrometry and Allied Topics took place June 5–9 in San Anto-nio, Texas, at the Henry B. González Convention

Center. Short courses preceded the conference open-ing, starting Saturday, June 4, and continuing through Sunday, June 5.

Opening the conference program on Sunday were tu-torial sessions and a plenary lecture. The first tutorial, by Facundo Fernandez of the Georgia Institute of Tech-nology (Atlanta, Georgia), and Glen Jackson of West Virginia University (Morgantown, West Virginia), was titled “Forensic Mass Spectrometry: Tell Me Something I Don’t Know.” Lars Konermann of the University of Western Ontario (London, Ontario, Canada) followed Fernandez and Jackson with a tutorial titled “An Ana-lyte’s Journey from Solution into the Gas Phase.”

ASMS Vice President for Programs Vicki H. Wysocki of the Ohio State University (Columbus, Ohio) officially welcomed attendees. She was followed by Erica Ollmann Saphire of the Scripps Institute (La Jolla, California), who gave the plenary lecture, titled “A Molecular Ar-senal Against Ebola.” After Saphire’s talk, a welcome reception took place in the poster and exhibit hall.

Three more plenary events took place during the week, two of which were award lectures, given by Scott A. McLuckey on Monday, and by Kristina Håkansson on Tuesday (details below). On Thursday, William Bi-alek of Princeton University (Princeton, New Jersey)

gave a plenary lecture titled “More than the Sum of its Parts: Collective Phenomena in Living Systems, from Single Molecules to Flocks of Birds.”

Continuing in the tradition of previous ASMS confer-ences, oral sessions, poster sessions, and exhibits took place throughout the day, Monday through Thurs-day, and workshops took place daily, Monday through Thursday.

AwardsThis year’s conference included several award presenta-tions. The 2016 Award for a Distinguished Contribution in Mass Spectrometry was presented on Monday after-noon to Scott A. McLuckey of Purdue University (West Lafayette, Indiana) for his pioneering contributions to the understanding of the gas-phase ion-ion reactions of polyatomic molecules and their applications in ana-lytical mass spectrometry. McLuckey and his cowork-ers initiated and sustained a line of research employing electrospray and ion traps that has revealed a wide and expanding array of ion–ion reactions that significantly expand the scope of tandem mass spectrometry, particu-larly in biological mass spectrometry.

Krist ina “Kicki” Håkansson of the University of Michigan (Ann Arbor, Michigan) was presented with the 2016 Biemann Medal on Tuesday afternoon for her work in developing and elucidating the mechanisms of electron-based activation methods, including electron


capture dissociation, electron de-tachment dissociation, and electron induced dissociation. She has ap-plied these electron-based activation methods to identify and characterize biological molecules from a number of classes, including peptides, oligo-nucleotides, and oligosaccharides.

Also presented on Tuesday af-ternoon were t he 2016 Research Awards, funded by Thermo Fisher Scientific and Waters Corporation, in the amount of $35,000 each. This year’s awardees were Ronghu Wu of Georgia Institute of Technology, and Etienne Garand of the University of Wisconsin-Madison.

The 2016 Ron A. Hites Award for an outstanding research publication in The Journal of the American So-

ciety for Mass Spectrometry (JASMS) was presented on Wednesday after-noon to Kevin Pagel, of the Max Planck Society in Berlin, Germany, and coauthors Waldemar Hoffmann and Johanna Hofmann for their paper “Energy-Resolved Ion Mobil-

ity-Mass Spectrometry: A Concept to Improve the Separation of Iso-meric Carbohydrates.” The award recognizes an outstanding publica-tion of original research, based on a paper’s innovative aspects, techni-cal and presentation quality, likely stimulation of future research, and impact on future applications. The award is named in honor of Profes-sor Ron Hites of Indiana Univer-sity, who led the creation of JASMS in 1988 while he was president of ASMS. The award includes $2,000 and a certificate for each author.

Also presented on Wednesday af-ternoon were the 2016 Postdoctoral Awards. Four awards in the amount of $10,000 each went to John Cahill of the Oak Ridge National Labora-tory, Andrew DeBlase of Purdue University, Catherine Going of Stan-ford University, and Pengyuan Liu of The Wistar Institute. The awards are intended to promote the profes-sional career development of post-doctoral fellows in the field of mass

spectrometry. Activities funded by these awards include conference and workshop attendance, travel to other mass spectrometry laboratories, and the purchase of books and software.

A closing event took place Thurs-day evening at the Briscoe West-ern Art Museum. The museum was named in honor of the late Texas Governor, Dolph Briscoe, Jr. His wife, Janey, preserves and interprets the art, history, and culture of the American West though engaging ex-hibitions, education programs, and public events ref lective of the region’s rich traditions and shared heritage.

ASMS 2017The 65th Annual ASMS Conference will be held June 4–8, 2017, in India-napolis, Indiana. For more informa-tion, visit www.asms.org in the com-ing months. ◾

Cindy Delonas is the Associate Editor for Spectroscopy. Direct correspondence to: [email protected] ◾

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Cholesteryl-bonded core-shell reversed-phase HPLC columnThe COSMOCORE Cholester is a core-shell reversed-phase HPLC column with cholesteryl-bonded stationary phase. It has the same hydrophobicity as C18, and consequently it retains molecules in similar ways. The COSMOCORE Cholester differs by having superior shape selectivity. This is a powerful tool for the separation of epimers, cis-trans isomers, and other structurally similar compounds. For example, Vitamin D2 and D3 isocratic separation is achieved under 3 min using 100% MeOH. In another example, 25(OH) Vitamin D2 and D3 metabolites and their C-3 epimers can also be baseline separated.

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ASMS PRODUCT PROFILESDry scroll vacuum pumpAnest Iwata Corporation is the original developer of the dry, air cooled, scroll vacuum pump. Our inherently balanced design provides longer tip seal life, sustained pumping performance, and reliability. We offer pump down speeds from 2–42 CFM and have an ultimate vacuum of 10-3 Torr. Ideal for backing turbo pumps in LC–MS and GC–MS applications. Our dry (oil free) design means reduced maintenance cost, quiet cool operation, and no risk of oil contamination in your laboratory!

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Portable mass spectrometerBaySpec Portability is a compact lightweight (<10 kg) mass spectrometer specifically designed for field applications. It uses a miniature linear ion trap mass analyzer, which provides high sensitivity and low detection limits. The instrument is equipped with two independent inlets, one for electron ionization and another for atmospheric pressure ionization. It is compatible with electrospray, thermal-desorption electrospray, APCI, and most of the ambient ionization techniques such as DART and DESI. The Portability is ideal for trace in situ analysis in many demanding applications, such as process control, detection of explosives and warfare agents, or screening for pesticides and other environmental contaminants.

BaySpec, Inc.San Jose, CAwww.bayspec.com

Tandem ionization for mass spectrometry Tandem Ionization® allows both soft and classical electron ionization mass spectra to be acquired simultaneously across an entire GC or GC×GC run. This provides analysts the capability to acquire both soft EI spectra and conventional 70 eV spectra at the same time in an integrated workflow. This means that a single GC or GC×GC run can provide all the information needed to fully characterize a sample for both target compounds and unknowns. Tandem ionization, unique to Markes, generates two spectra from a single peak by rapid switching between “soft” ionization (typically 10–16 eV) and conventional “hard” ionization (70 eV), and is fully automated by Markes’ TOF-DS software for BenchTOF.

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ASMS PRODUCT PROFILES

Automated real-time VOC analysis at parts per trillion (PPT) levelsThe Selected Ion Flow Tube Mass Spectrometry (SIFT-MS) system delivers comprehensive, instant gas analysis to trace pptv levels. Now available with high-throughput continuous sample analysis, the Voice200ultra from Syft Technologies utilizes direct, ultra-soft chemical ionization. Using eight instantly switchable mass-selected reagent ions enable detection of a very wide range of challenging compounds like ammonia, formaldehyde, hydrogen chloride, and hydrogen sulfide. The system combines extremely high selectivity with push-button simplicity—all with-out requiring sample preparation. Value-added financing plans, including rent, lease, or zero down and zero interest financing, are available.

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LC–MS front end componentsPeeke Scientific is a source for a variety of LC–MS front end components. HPLC and UHPLC columns are available for small molecule, peptide, and protein separations in both nano and analytical formats. To complement columns, there are traps, guard cartridges, and in-line filters available with wrench-tight, finger-tight, and quick disconnect configurations. For fluidics control, they supply tubing, fittings, syringes, valves, splitters, metering pumps, and HPLC and UHPLC pump replacement parts.

Peeke ScientificNovato, CAwww.peekescientific.com

Torion portable GC–MSPerkinElmer’s Torion T-9 Portable GC–MS is the smallest instrument available for GC–MS analyses outside of the laboratory in the field. The system rapidly screens chemicals, including environmental volatiles and semivolatiles (VOCs and SVOCs), explosives, chemical warfare agents, and hazardous substances. It can also be used in food safety and industrial applications. The system is fully self-contained and weighs 32 pounds with rechargeable battery operation.

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OptocouplersChoose from 10 kV, 15 kV, or 50 kV optocouplers used to prevent ground loops in noisy environments where instruments are used. The linear reverse leakage current output, when the optocoupler is reverse-biased, is useful in control circuitry, feedback loops, or when used as a high voltage switch or op amp.

The 10 kV and 15 kV OC100/150 series feature high gain, long-term gain stability, and high isolation.

The 50 kV OC500 is designed for a push-pull configuration. It has two legs, each capable of withstanding 25 kV reverse voltage, controlled independently via low voltage LED connections.

Voltage MultipliersVisalia, CAwww.VoltageMultipliers.com

ADVERTISEMENT36 Mass Spectrometry

The compound 4-methylimidazole (4-MEI) is formed as

a by-product in some foods and beverages. Caramel

coloring Type III and Type IV in beverages is one of the

ingredients which may contain 4-MEI and it may be

found in products such as certain colas, beers, soy sauces,

breads, coffee, ammoniated livestock feed, and others.

There has been an increase in concern lately about 4-MEI

being a suspected carcinogen. Europe has regulated

the amount of 4-MEI allowed in coloring used in food

products. The state of California has added 4-MEI to its

proposition 65 list of known carcinogens. California now

requires products with an exposure potential of >29 μg/

day to carry warning labels.

T his work will demonstrate a simple methodology using

automated solid-phase extraction (SPE) and HPLC coupled

with mass spectrometric detection.

Th e SmartPrep® Cartridge Extractor was used with a Waters

Oasis® MCX 3 cc (60 mg) cartridge. Th e extraction method was

optimized using a series of conditions and collecting fractions on

the SmartPrep (1).

Th e samples for this experiment were common soft drinks con-

taining caramel coloring. Th e point of purchase and original product

production site are variables in 4-MEI levels. Some of the beverages

were purchased in New England and others in California, in common

supermarkets and their bottling origin is listed with the results.

Results and Discussion

A sensitive method for 4-MEI detection was developed using

HPLC–MS, showing adequate measurement below the California

Prop 65 requirements of 29 μg per day. Solid-phase extraction

was shown to adequately concentrate the 4-MEI from the soft

drinks analyzed and provide clean-up. Two samples were shown

to exceed the daily exposure limits in Prop 65 with one serving.

References

(1) “A Simple Analysis of 4-Methylimidazole Using Automated Solid

Phase Extraction and High Performance Liquid Chromatography with

MS-MS and MS–SIM Detection,” AN1111606_01, available from

www.horizontechinc.com.

A Simple Analysis of 4-Methylimidazole Using Automated Solid-Phase Extraction and High Performance Liquid Chromatography with MS-MS and MS–SIM DetectionDavid Schiessel*, Paul Monroy*, Chris Shevlin†, and William Jones†, *Babcock Laboratories, Inc., and †Horizon Technology, Inc.

Horizon Technology, Inc.16 Northwestern Drive, Salem, NH 03079

tel. (603) 893-3663

Website: www. horizontechinc.com

Figure 1: Mexican soda sample, S/N = 383 for quantitation ion.

Table I. Results for 4-MEI in selected samples from various

geographic regions

Serving Size Result

SmartPrep Origin fl oz mL μg/L μg/serving

Diet cola 1 New England 12 355 55.2 19.6

Diet cola 2 New England 12 355 6.5 2.3

Orange cola blend Germany 11.2 331 9.3 3.1

Apple cola Mexico 12 355 84.1 29.8

Sangria soda Mexico 12 355 22.4 8.0

Malt drink (non-alcoholic) Jamaica 12 355 84.7 30.1

ADVERTISEMENT Mass Spectrometry 37

The Diablo 5000A Real-Time Gas Analyzer (RTGA) is a

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Visualize Your Process Streams in Real-Time with MS for Rapid Process DevelopmentTerry Ramus, PhD, Diablo Analytical, Inc.

Diablo Analytical, Inc.5141 Lone Tree Way, Antioch CA 94531

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Figure 1: Real-time monitoring of starting materials, addition of cata-lyst at 9 min showing reaction products; catalyst lasts only about 5 min.

changes in seconds. But the process returns to the original reagent

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Figure 2: RTGA interface design to MS. Shows process stream sampling to detection by Agilent MSD.

38 Current Trends In Mass Spectrometry July 2016 www.spec t roscopyonl ine .com

Calendar of EventsAugust20–26 21st International Mass Spectrometry Conference (IMSC 2016) Toronto, Canada www.imsc2016.ca/

September12–15 25th ICP-MS User Meeting & 12th Symposium Mass Spectrometric Methods of Trace AnalysisSiegen, Germany icpms-anwendertreffen.de/

12–15 Mass Spectrometry: Applications to the Clinical Lab (MSACL) 2016 EU 3rd Annual Congress & Exhibition Salzburg, Austriawww.msacl.org

13–15 37th British Mass Spectrometry Society (BMSS) Annual Meeting 2016 Eastbourne, England www.bmss.org.uk/bmss2016/bmss2016.shtml

18–22 15th Human Proteome Organization World CongressTaipei, Taiwan www.hupo2016.org/index.html

18–23 SciX 2016Minneapolis, MN www.scixconference.org

18–23 Forensic Isotope Ratio Mass Spectrometry (IRMS) Conference 2016Auckland, New Zealand www.forensic-isotopes.org/2016.html

27–30 13th Symposium on the Practical Applications of Mass Spectrometry in the Biotechnology Industry San Diego, CA www.casss.org/?MS1600

October4–6 Cannabis Science Conference (CANNCON) Portland, OR www.cannabisscienceconference.com/

5–7 SFC 2016 10th International Conference on Packed Column SFCVienna, Austria www.greenchemistrygroup.org/

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