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Liquid chromatography–tandem mass spectrometry method for the bioanaly-
sis of N,N-dimethyltryptamine (DMT) and its metabolites DMT-N-oxide and
indole-3-acetic acid in human plasma
Dino Luethi, Karolina E. Kolaczynska, Severin B Vogt, Laura Ley, Livio
Erne, Matthias E. Liechti, Urs Duthaler
PII: S1570-0232(22)00439-1
DOI: https://doi.org/10.1016/j.jchromb.2022.123534
Reference: CHROMB 123534
To appear in: Journal of Chromatography B
Received Date: 28 September 2022
Revised Date: 7 November 2022
Accepted Date: 7 November 2022
Please cite this article as: D. Luethi, K.E. Kolaczynska, S.B. Vogt, L. Ley, L. Erne, M.E. Liechti, U. Duthaler,
Liquid chromatography–tandem mass spectrometry method for the bioanalysis of N,N-dimethyltryptamine
(DMT) and its metabolites DMT-N-oxide and indole-3-acetic acid in human plasma, Journal of Chromatography
B (2022), doi: https://doi.org/10.1016/j.jchromb.2022.123534
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© 2022 Published by Elsevier B.V.
1
Liquid chromatography–tandem mass spectrometry method for the
bioanalysis of N,N-dimethyltryptamine (DMT) and its metabolites DMT-N-
oxide and indole-3-acetic acid in human plasma
Dino Luethia, Karolina E. Kolaczynskaa, Severin B Vogta, Laura Leya, Livio Ernea, Matthias E.
Liechtia,*, Urs Duthalera,*
Affiliations
aDivision of Clinical Pharmacology and Toxicology, Department of Biomedicine and Department of
Pharmaceutical Sciences, University Hospital Basel and University of Basel, Basel, Switzerland
*To whom correspondence may be addressed. Email: matthias.liechti@usb.ch
*To whom correspondence may be addressed. Email: urs.duthaler@unibas.ch
Word count: abstract: 242 words; main text: 4252 words; references: 29; figures: 6, tables: 2;
supplementary tables: 4
Key words: ayahuasca; psychedelic; metabolism; bioanalysis; pharmacokinetics; HPLC
2
Abstract
The indole alkaloid N,N-dimethyltryptamine (DMT) induces psychedelic effects in humans. In
addition to ceremonial and recreational use, DMT is subject to clinical investigations. Sensitive
bioanalytical methods are required to assess the pharmacokinetics of DMT and its metabolites in human
plasma. Here, a high performance liquid chromatography–tandem mass spectrometry (LC–MS/MS)
method for the quantification of DMT and its major metabolites indole-3-acetic acid (IAA) and DMT-
N-oxide (DMT-NO) was developed and validated. As IAA is an endogenous component of human
plasma, 13C6-IAA was used to determine IAA concentrations. After simple protein precipitation with
methanol, analytes were separated on a pentafluorophenyl column. A gradient consisting of 0.1% (v/v)
formic acid in a methanol-water mixture was applied for analyte separation. The analytes were detected
by positive electrospray ionization followed by multiple reaction monitoring. The calibration range of
the assay was 0.25–250 ng/mL for DMT, 0.1–100 ng/mL for DMT-NO, and 25–25,000 ng/mL for 13C6-
IAA. The intra- and inter-assay accuracy was 93–113% for all analytes at all quality control levels, with
coefficient of variation ≤ 11%. All analytes were stable under storage conditions relevant for the analysis
of large batches of study samples. The validated method was capable of assessing pharmacokinetic (PK)
parameters of DMT and its metabolites in study participants intravenously perfused with 1 mg/min DMT
for 90 min. Overall, the developed method is easy-to-use, has short run times, and qualifies for PK and
metabolism studies of DMT in clinical settings.
1 Introduction
The serotonergic psychedelic N,N-dimethyltryptamine (DMT) is a psychoactive ingredient in
Ayahuasca, a brew prepared from DMT-containing plants, such as Psychotria viridis, in combination
with Banisteriopsis caapi vines; the latter contain β-carboline alkaloids that inhibit monoamine oxidase
(MAO), thereby preventing rapid breakdown of DMT [1]. Ayahuasca has a long history of being used
in rituals by indigenous people of South America [1-3]. DMT induces psychedelic effects mediated by
non-selective 5-HT2A receptor agonism and displays low toxicity [4, 5]. In addition to ceremonial and
recreational use, DMT has recently gained increased interest as a potential therapeutic agent for the
treatment of mental disorders and drug dependence [6-10]. Currently, different and mostly intravenous
DMT dosage regimens are under investigation in several clinical studies [11]. Acute high doses of DMT
may produce a delirium in users, for which reasons intravenous perfusions may be preferred to gradually
move individuals into states of altered consciousness [10]. Reliable bioanalytical assays are thus crucial
to accurately assess the pharmacokinetics of DMT and its metabolites. In humans, DMT is primarily
metabolized to indole-3-acetic acid (IAA) via MAO-A [12, 13]. Independent of MAO-A, DMT is
additionally metabolized to DMT-N-oxide (DMT-NO), the second most abundant metabolite [12, 13].
N-methyltryptamine and 2-methyl-tetrahydro-betacarboline are formed to a lesser extent [12, 13]. A
number of analytical methods for the bioanalysis of DMT have been described so far [14-21]. However,
3
long run times, extensive sample preparation, or the lack of sensitive metabolite detection illustrate the
need for sensitive but easy-to-use and rapid methods. Recently, Eckernäs and colleagues developed a
highly sensitive LC–MS/MS method to quantify DMT and its two major metabolites in human plasma
[22]. Another approach with simplified sample preparation and reduced run time is described herein,
which also enabled the quantification of DMT-NO in clinical study samples. IAA is endogenously
present at relatively high levels in plasma, thus precluding the preparation of IAA calibration lines in
plasma. Therefore, the applicability of 13C6-IAA to calculate IAA concentrations was established with
the aim of avoiding less reliable baseline subtraction or laborious standard addition methods.
2 Experimental section
2.1 Chemicals, reagents, and reference compounds
DMT (99.8% HPLC purity) was purchased from Lipomed (Arlesheim, Switzerland). DMT-NO
(98.0% HPLC purity) and IAA sodium salt (100.0% HPLC purity) were obtained from Cayman
Chemical (Ann Arbor, MI, USA). 13C6-IAA (phenyl-13C6, 99%;) was bought from Cambridge Isotope
Laboratories, Inc. (Tewksbury, MA, USA). The deuterated internal standard DMT-d6 hemifumarate
(99% D; 99.4% HPLC purity) was synthesized by Reseachem (Burgdorf, Switzerland). IAA-d2 (98%
D; 98% HPLC purity) was purchased from Sigma-Aldrich (Buchs, Switzerland). LC–MS grade water
and methanol were obtained from Merck (Darmstadt, Germany). Formic acid (99–100%) was bought
from VWR Chemicals (Radnor, PA, USA). Dimethyl sulfoxide (DMSO) was a product of Sigma-
Aldrich. Human blood was and obtained from the blood donation center of the University Hospital Basel
and collected in lithium heparin coated S-Monovette tubes (Sarstedt, Nümbrecht, Germany). To obtain
plasma, blood was centrifuged (Eppendorf 5810 R centrifuge; Hamburg, Germany) at 1,811 × g for 10
min at room temperature.
2.2 Calibration and quality control sample preparation
Separate analyte stock solutions were prepared for calibrator (CAL) and quality control (QC)
samples. The analytes were weighed in on a XP 26 microbalance (Mettler Toledo, Columbus, USA) and
dissolved in DMSO at a final concentration of 10 mg/mL. All stock solutions were stored at -20 °C. A
pool of blank human plasma derived from 8 donors was spiked with the CAL and QC working solutions
in DMSO at a ratio of 1:100 (v/v). The CAL line in plasma was prepared using 10 different
concentrations in the range from 0.25 ng/mL to 250 ng/mL for DMT, 0.1 ng/mL to 100 ng/mL for DMT-
NO, and 25 ng/mL to 25,000 ng/mL for 13C6-IAA. Four QC levels were prepared in plasma at the
following concentrations: 0.25 ng/mL DMT, 0.1 ng/mL DMT-NO, and 25 ng/ mL 13C6-IAA (LLOQ);
0.5 ng/mL DMT, 0.25 ng/mL DMT-NO, and 50 ng/ mL 13C6-IAA (low QC; LQC), 5 ng/mL DMT, 5
ng/mL DMT-NO, and 500 ng/mL 13C6-IAA (medium QC; MQC), and 100 ng/mL DMT, 100 ng/mL
DMT-NO, and 10,000 ng/mL 13C6-IAA (high QC; HQC).
4
2.3 LC–MS/MS instrumentation and settings
A modular ultra-high performance liquid chromatography (UHPLC) system (Shimadzu, Kyoto,
Japan) consisting of four pumps was used to separate the analytes. The UHPLC system was connected
to a API 4000 QTRAP tandem mass spectrometer (AB Sciex, Ontario, Canada). The analytes were
separated on a Luna PFP(2) analytical column (3.0 μm, 100 Å, 2 × 50 mm, Phenomenex, Torrance, CA,
USA), which was kept at 45 °C. Water and methanol, both supplemented with 0.1% formic acid, were
used as mobile phase A and B, respectively. In order to improve interactions of the methanolic sample
with the column, 10 μL of the sample were first mixed in a T-union with mobile phase A, delivered by
pump C. The initial flow rate of pump C was 0.6 mL/min, which was turned off after 0.5 min of each
run. Concurrently, pump A and B loaded the sample onto the analytical column using 15% mobile phase
B. The flow rate of pump A and B was kept at 0.1 mL/min for the first 0.5 min of each run and afterwards
set to 0.5 mL/min until the end of the run (0.5–4.5 min). In order to elute the analytes, mobile phase B
concentration was linearly increased to 30% between 0.5 min and 2.5 min and then increased to 95%
between 2.5 min and 3 min. Afterwards, the column was washed for 1 min with 95% mobile phase B
and finally re-conditioned for 0.5 min with 15% mobile phase B. Between each sample injection, the
autosampler port was washed with a water-methanol-acetonitrile-isopropanol (1:1:1:1, v/v) solution.
The gradient program resulted in a retention time of 1.68 min for DMT and DMT-d6, 2.21 min for DMT-
NO, and 3.31 min for IAA, 13C6-IAA, and IAA-d2 (Fig. 1). Scheduled multiple reaction monitoring
(sMRM) was used with a detection window of 60 s and a target cycle time of 0.4 s. Ion transitions and
analyte specific settings are indicated in Table 1. The electrospray ionization source was operated in
positive mode at an ion spray voltage of 5,500 V and a temperature of 500 °C. Nitrogen was employed
as curtain (10 psi), ion source (gas 1: 60 psi; gas 2: 50 psi), and collision gas (4 psi).
2.4 Sample extraction
Human plasma (50 μL) was transferred into 0.75 mL Matrix™ Blank Storage Tubes (Thermo
Fisher Scientific, Waltham, MA, USA) and precipitated with 150 μL internal standard (ISTD) solution
consisting of 5 ng/mL DMT-d6 and 500 ng/mL IAA-d2 in methanol. The tubes were subsequently
vortexed on a Multi Tube Vortexer (VX-2500, VWR, Radnor, PA, USA) for 30 s and centrifuged for
30 min at 10 °C and 3,220 × g to produce supernatant free of plasma proteins. After extraction, samples
were kept in the autosampler at 10 °C until analysis.
2.5 Method validation
The method was validated according to the Bioanalytical Method Validation Guidance for
Industry of the United States Food and Drug Administration (FDA) [23] and the Guideline on
bioanalytical method validation of the European Medicines Agency (EMA) [24]. Linearity, accuracy
and precision, selectivity and sensitivity, extraction recovery and matrix effect, and analyte stability
under various storage conditions were validated on an API 4000 QTRAP LC–MS/MS system.
5
2.5.1 Linearity
Two separate CAL lines were included in each analytical run, one measured at the beginning
and one at the end of the acquisition batch. A CAL line comprised a blank plasma sample, spiked plasma
at ten concentration levels, and a double blank sample. The spiked plasma samples were arranged in
ascending order; concentration levels are shown in Supplementary Table 1. The blank sample was
precipitated with ISTD solution; the double blank sample was processed with methanol and measured
directly after the highest CAL sample to determine the degree of analyte carry-over. The upper limit for
analyte carry-over was set at 20% of the LLOQ peak area. A linear regression line was calculated in
MultiQuant by plotting the nominal CAL concentrations against the ratio of the analyte peak area to the
ISTD peak area. A weighing factor of 1/x2 was used to reduce the error at low concentrations. The
difference between measured and nominal concentration was calculated to determine the accuracy of
the method. CAL points with 85–115% (LLOQ: 80–120%) accuracy were used to plot CAL lines. At
least 75% of all CAL points as well as one LLOQ and ULOQ sample were required to fulfill the accuracy
criteria. Only CAL lines with correlation coefficient (R) of ≥ 0.99 were accepted.
2.5.2 Intra- and inter-assay accuracy and precision
The intra- and inter-assay accuracy and precision were determined in three independent
validation runs on separate days. Each validation run included two CAL lines and seven QC replicates
per level (LLOQ, LQC, MQC, and HQC). The accuracy was determined as difference of the measured
concentration to the nominal concentration. The coefficient of variation (CV, relative standard
deviation) was used to assess intra- and inter-assay precision. Intra-assay (n=7) and inter-assay (n=21)
accuracy of 85–115% (LLOQ: 80–120%) and CV ≤ 15% (LLOQ: ≤ 20%) was the required validation
criteria. Furthermore, ≥ 50% of QC samples per level and ≥ 67% of all QC samples had to meet the
accuracy criteria for a validation run to succeed. This requirement was set in addition to the
recommendations of the FDA and EMA [23, 24] in order to further ensure the reliability of the method.
2.5.3 Selectivity and sensitivity
The selectivity and sensitivity of the analytical method was assessed by analyzing individual
double blank, blank, and LLOQ samples of seven subjects. Each individual plasma was extracted in
triplicate. To ensure method sensitivity, the LLOQ analyte response (peak area) had to be at least five
times larger than the noise signal in the blank and double blank samples. Chromatograms of blank and
double blank plasma extracts were overlaid with their corresponding LLOQ samples to exclude that
method selectivity is affected by co-eluting plasma components. In addition, the accuracy of ≥ 50%
individual LLOQ samples and mean accuracy had to be within 80–120% for the method to be considered
sufficiently selective.
6
2.5.4 Extraction recovery and matrix effect
Extraction recovery and matrix effect were examined at different QC levels (LLOQ, LQC,
MQC, and HQC). To assess the extraction recovery, plasma of seven separate subjects was spiked with
equal amounts of analyte before and after extraction. For each subject and QC level, the analyte peak
area of samples spiked before extraction was compared to blank plasma samples spiked after extraction
at corresponding concentrations. By definition, the recovery of samples spiked after extraction was
100%. The matrix effect was determined by comparing the analyte peak area of plasma spiked after
extraction with matrix free samples (containing water and ISTD solution; 1:3, v/v). Inter-batch matrix
variability was investigated by assessing plasma samples of seven separate subjects. Overall, the
extraction recovery and matrix effect had to be consistent between plasma batches and QC levels, with
CV ≤ 15% (LLOQ: ≤ 20%).
2.5.5 Stability
Analyte stability was examined under several different storage conditions, which were
compared with freshly prepared CAL lines. Specifically, QC samples (LLOQ, LQC, MQC, and HQC)
were analyzed after storage at room temperature for 8 h (benchtop stability) and storage at -20 °C for 1
and 3 months (short- and medium-term stability). Short- and medium-term stability at -20 °C also covers
stability at colder temperatures [23]. Additionally, stability after three freeze and thaw cycles was
analyzed (freeze-thaw stability). In a freeze and thaw cycle, samples were frozen at -20 °C for at least
24 h and then thawed unassisted at room temperature. A deviation of the measured QC samples of ≤
15% (LLOQ: ≤ 20%) from the nominal concentration indicated sample stability under the particular
storage condition. To established whether processed samples can be reinjected in case of equipment
failure, stability of samples stored in the autosampler overnight at 10 °C was investigated by reinjecting
validation run samples on the following day [24].
2.6 Method application
To test the applicability of the method, study samples from five healthy volunteers participating
in a clinical phase 1 study (ClinicalTrials.gov Identifier: NCT04353024) were analyzed. The study was
approved by the ethics committee of Northwest and Central Switzerland (BASEC ID: 2020-00376) and
was conducted according to the Declaration of Helsinki and the International Conference of
Harmonization for Good Clinical Practice guidelines. The study participants were intravenously
perfused with 1 mg/min DMT for 90 min. Blood samples (7.5 mL) were drawn into lithium heparin
coated S-Monovettes (Sarstedt) 5 min prior to treatment and 2, 5, 10, 15, 20, 30, 40, 50, 60, 70, 80, 90,
95, 100, 105, 110, 120, 130, 140, and 150 min after starting the perfusion. The blood samples were
centrifuged at 1,811 × g for 10 min to obtain plasma (approximately 3 mL per time point). Plasma
samples were stored as 1 mL aliquots at -80 °C until analysis. The 105 study samples were analyzed in
one analytical run together with two CAL lines and four sets of QC samples (LLOQ, LQC, MQC, and
7
HQC), which were stored at -20 °C. In addition, IAA samples (2,500, 5,000, 10,000, and 25,000 ng/mL)
were included in the run to ensure the applicability of the 13C6-IAA CAL lines to calculate IAA
concentrations. The QC samples were distributed equally over the acquisition batch, whereas CAL lines
were placed at the beginning and the end of the analysis. Phoenix WinNonlin Software (Version 8.1.0,
Certara, Princeton, NJ, USA) was employed to calculate pharmacokinetic parameters. The maximal
concentration (Cmax) and the time to reach Cmax (Tmax) were directly derived from the plasma
concentration-time curves. The elimination constant (λ) was the slope of the log concentration-time
regression determined in the terminal elimination phase. The elimination half-life (t1/2, h) was the ratio
of ln(2) and the elimination constant. The linear trapezoidal rule from timepoint zero to the timepoint of
the last quantifiable concentration was employed to calculate the area under the plasma concentration-
time curve (AUC0–150 min). Plasma concentrations below the LLOQ were set to 0 ng/mL.
3 Results and discussion
3.1 Method development
Clinical research with DMT has recently gained new momentum. Reliable and easy-to-use
analytical methods are essential to ensure smooth and efficient analysis of clinical study samples. The
aim of this study was the development of a method with simple sample preparation and short run time.
In addition, enabling sensitive quantification of metabolites benefits the investigation of metabolic
differences due to genetic polymorphisms or drug-drug interactions [25]. When DMT is applied without
MAO inhibition, most of the parent compound is metabolized to IAA and only a minor part is
transformed into DMT-NO [12, 13]. High method sensitivity is therefore important when measuring
samples of studies in which DMT is applied as sole compound, leading to low DMT-NO plasma
concentrations. The analysis of IAA in human plasma poses a challenge due to its high endogenous
concentrations, which preclude linear calibration lines. One way to overcome this issue is to subtract the
peak area ratio of a blank sample from each calibration sample [22]. The standard addition method is
another approach; however, it is very laborious and requires large amounts of material as every sample
has to be extracted multiple times with increasing amounts of analyte. Here, an alternative approach was
chosen to minimize time efforts and potential error sources. Specifically, IAA concentrations were
calculated using 13C6-IAA, which exhibits very similar physicochemical properties as the unlabeled
analyte but is not present in human plasma (Fig. 2).
Analytes were first infused into the mass spectrometer to obtain their fragmentation pattern and
to optimize ionization parameters. DMT, DMT-NO, and DMT-d6 were ionized in the positive ionization
mode. Ionization of IAA, 13C6-IAA, and IAA-d2 was tested in positive and negative mode due to their
acid moieties. Both ionization modes yielded satisfactory results; hence, positive mode ionization was
chosen for all analytes to avoid polarity switching within the analytical run. The same approach was
applied in another recent study using the same type of mass spectrometer [22]. The two predominant
fragments of DMT (m/z 189.1 58.1 and m/z 189.1 144.1) were chosen as quantifier and qualifier,
8
respectively. For DMT-NO, m/z 205.1 144.0 was the most abundant fragment, which was therefore
chosen as quantifier. The less abundant fragment m/z 205.1 115.0 was included as qualifier. For
DMT-d6, fragments corresponding to the fragments of DMT were selected (i.e., m/z 195.0 64.1 and
m/z 195.0 144.0). For 13C6-IAA, the most abundant fragment m/z 182.0 136.1 was included as
quantifier and the second most abundant fragment m/z 182.0 109.0 as qualifier transition.
Corresponding fragments were chosen for IAA (m/z 176.0 130.1 and m/z 176.0 103.0) and IAA-
d2, (m/z 178.0 132.1 and m/z 178.0 105.1). The quantifier fragments for DMT, DMT-NO, and
IAA were identical with fragments monitored by the method of Eckernäs and colleagues [22].
Due to the extensive metabolism of DMT to IAA, the ULOQ for 13C6-IAA was set to 25 µg/mL.
At this concentration, peak intensities were outside the linear detection range. The signal strength for
IAA and 13C6-IAA was therefore quenched by increasing the collision energy. By this approach, the
sensitivity of the other analytes remained unaffected in contrast to reduction of the injection volume or
increased sample dilution. An overview over the mass spectrometric settings is provided in Fig. 1 and
Table 1. Based on previous experience with the analysis of psilocybin's major metabolites [26], different
C18 columns were tested. The Luna PFP(2) column was tested due to the aromatic moiety and polarity
of the analytes; it yielded good peak separation for DMT, DMT-NO, and endogenous interferences. The
mobile phase gradient was adjusted to achieve sufficient separation of DMT and DMT-NO. Formic acid
was added to the mobile phase to facilitate positive ionization. At low IAA-d2 concentration in the ISTD
solution (20 ng/mL), interferences with the IAA signal were observed. This issue was circumvented by
increasing the IAA-d2 concentration in the ISTD solution to 500 ng/mL. Due to the high IAA-d2
concentration in the ISTD solution, the analyte signal was quenched by adapting the collision energy.
At identical collision energy, the peak area of 13C6-IAA was slightly higher than the IAA peak area.
Therefore, the collision energy of IAA was adapted in a way that both signals matched at identical
concentrations.
Simple protein precipitation was used for sample preparation, allowing for fast and
straightforward analysis of study samples [26-29]. A plasma volume of 50 µL renders multiple run
repetitions possible, even if only a small amount of plasma is available. To improve the retention of the
polar analytes on the stationary phase, the methanolic sample was diluted online with mobile phase A
before reaching the analytical column. Finally, plasma samples of study participants dosed with 1
mg/min DMT during 90 min were measured to determine the calibration ranges for all analytes. These
preliminary measurements were only used to determine the calibration range, not for PK analysis. The
ULOQ was set sufficiently high in order to avoid additional study sample dilution, thereby facilitating
the analysis of large batches of clinical samples.
3.2 Method validation
3.2.1 Method linearity and carry-over
9
The method was linear in the range of 0.25–250 ng/mL for DMT, in the range of 0.1–100 ng/mL
for DMT-NO, and in the range of 25–25,000 ng/mL for 13C6-IAA. The correlation coefficient (r) was
0.997 or higher for all CAL lines. Method linearity for all analytes is shown in Supplementary Table 1.
Carry-over signals in double blank samples injected directly after the ULOQ were 1.2–2.3%, 2.4–13.9%,
and 4.5–14.3% of the LLOQ signals for DMT, DMT-NO, and 13C6-IAA, respectively. The low carry-
over (≤ 20% LLOQ signal) indicates that no additional blank samples need to be included in the analysis
after measuring high concentration samples.
3.2.2 Accuracy, and precision
Intra- and inter-assay accuracy and precision were assessed at four different QC levels (LLOQ,
LQC, MGC, and HQC), measured in septuplicate on three separate days (Fig. 3 and Supplementary
Table 2). The intra-assay accuracy was 96–104% for DMT, 93–113% for DMT-NO, and 99–105% for
13C6-IAA. The inter-assay accuracy was 98–101% for DMT, 95–110% for DMT-NO, and 99–102% for
13C6-IAA. Intra- and inter-assay CV was ≤ 8% for DMT and ≤ 11% for DMT-NO and 13C6-IAA at all
QC levels. All of the 84 measurement points for DMT and 13C6-IAA met the validation criteria, whereas
79 of the 84 measurement points for DMT-NO met the criteria (Fig. 3); this result indicates a high
reliability of the method.
3.2.3 13C6-IAA validation
13C6-IAA was employed as surrogate to calculate IAA plasma concentrations. Fig. 2A
demonstrates that the signal intensity of both analytes in water matches over the entire CAL range (25–
25,000 ng/mL). However, plasma concentrations correspond only between 2,500 and 25,000 ng/mL
because of the large baseline IAA plasma concentration (Fig. 2B). The accuracy of 83 of the 84 IAA
QC plasma samples was between 89 and 115 %, calculated based on 13C6-IAA CAL lines (Fig. 2C).
Only one sample was outside the accepted range with accuracy of 116%. This result demonstrates that
13C6-IAA can be used to reliably determine IAA plasma concentrations. The approach allows to quantify
even low IAA plasma concentrations as the linear range of 13C6-IAA is two orders of magnitude larger
than the one of IAA. Overall, the use of a stable isotope labeled reference as surrogate for IAA is more
robust than baseline subtraction methods and less time consuming compared to standard addition
approaches.
3.2.4 Selectivity and sensitivity
The noise signal in double blank and blank plasma samples of seven separate donors was < 1%
of the LLOQ peak area for DMT and 13C6-IAA, and < 4% of the LLOQ peak area for DMT-NO (Fig. 4
and Supplementary Table 3). Moreover, the noise signal in double blank plasma samples was < 1% of
the average ISTD signal in CAL and QC samples (data not shown). This corroborates method selectivity
and sensitivity. To further assess method selectivity, plasma from 7 individual donors was each spiked
10
with the LLOQ and quantified. The accuracy of LLOQ peak areas was 87–113% for each plasma sample
and analyte, with average accuracies of 93–103%. Inter-subject CV was ≤ 6% for all analytes, affirming
method selectivity (Supplementary Table 4).
3.2.5 Recovery and matrix effect
Extraction recovery and matrix effect are shown in Table 2. Simple protein precipitation with
methanol resulted in extraction recoveries of 96–110% for all analytes and QC levels, with inter-subject
variation (CV) ≤ 10%. The matrix effect was 97–110% for all analytes and QC levels, with inter-subject
variation of ≤ 9%. Extraction recovery and matrix effect were consistent over all QC levels and were
not affected by plasma batch, ensuring consistent analysis. The simple extraction method resulted in
higher analyte recovery compared to a previously reported approach that included an evaporation step
[22]. Eckernäs and colleagues furthermore reported a high matrix effect for DMT-NO at low
concentrations, whereas the matrix effect for DMT was negligible [22]. Compared to the aforementioned
method, the method described herein therefore yields higher extraction recovery and a low matrix effect
that is consistent over the whole calibration range.
3.2.6 Stability
Samples of DMT, DMT-NO, and 13C6-IAA were stable for 8 h at room temperature, 3 months
at -20 °C, and after 3 freeze-thaw cycles. (Fig. 5). Moreover, no stability issues were observed for
validation samples re-injected after storage in the autosampler at 10 °C overnight. For DMT and DMT-
NO, only 2 out of 140 stability samples were outside the required range of 85–115% (LLOQ: 80–120%)
accuracy. For 13C6-IAA, 3 out of 140 stability samples did not meet the required accuracy. This confirms
previous reports of DMT, DMT-NO, and IAA sample stability for 24 h in the autosampler (8 °C), 3
months at -80 °C, and after 3 freeze-thaw cycles [22].
3.2.7 Clinical application
To test the applicability of the method, plasma samples of five subjects intravenously perfused
with 1 mg/min DMT during 90 min were analyzed (Fig. 6). QC samples included in the run were in line
with required accuracy and precision specifications. The following pharmacokinetic parameters (± SD)
were calculated: Cmax of DMT was 44.5 ± 12.7, Tmax was 80 ± 10 min, and AUC0–150 min was 2,498 ± 767
ng × min/mL. Different half-lives were observed for DMT, indicating multi-compartment kinetics.
Initial t1/2 was 4.4 ± 0.7 min, whereas terminal t1/2 was 16.7 ± 3.3 min. For DMT-NO, Cmax was 1.3 ±
0.3 ng/mL, Tmax was 89.0 ± 5.5 ng/mL, t1/2 was 42.5 ± 11.5 min, and AUC0–150 min was 105 ± 25 ng ×
min/mL. Substantially higher metabolite formation was measured for IAA: Cmax was 4,337 ± 1,045
ng/mL, Tmax was 95.0 ± 3.5 ng/mL, t1/2 was 312 ± 187 min, and AUC0–150 min was 412,444 ± 90,073 ng
× min/mL.
11
4 Conclusion
The validated method permits reliable and sensitive quantification of DMT and its two major
metabolites DMT-NO and IAA. Whereas DMT and DMT-NO were directly measured, concentrations
of the endogenously occurring metabolite IAA were calculated using linear 13C6-IAA CAL lines.
Advantages compared to other bioanalytical assays are the low amount of plasma required, simple
sample preparation by protein precipitation, and short run times. Accuracy and precision of the method
were in line with international regulatory guidelines. Complete extraction recovery and no significant
matrix effect was observed. The bioanalytical assay was successfully tested to measure DMT levels and
metabolite formation in plasma of study participants intravenously perfused with DMT. To conclude,
this method will facilitate DMT analysis in pharmacokinetics and drug interaction studies due to its high
sensitivity and simplicity.
CRediT author statement
Dino Luethi: methodology, validation, formal analysis, investigation, data curation, writing –
original draft, writing – review & editing, visualization, funding acquisition. Karolina E. Kolaczynska:
methodology, investigation, writing – review & editing. Severin B. Vogt: resources, writing – review
& editing. Laura Ley: resources. Livio Erne: resources. Matthias E. Liechti: writing – review &
editing, supervision, project administration, funding acquisition. Urs Duthaler: conceptualization,
methodology, validation, formal analysis, data curation, writing – review & editing, visualization,
supervision, project administration.
Funding
This work was supported by the Swiss National Science Foundation (SNSF, Grant No.
P5R5PM_206796 to D.L. and Grant No. 32003B_185111 to M.E.L.) and Mind Medicine Inc.
Declaration of competing interest
M.E.L. is a consultant for Mind Medicine, Inc. All other authors do not have any conflicts of
interest to declare for this work. Knowledge and data associated with this work are owned by the
University Hospital Basel and were licensed by Mind Medicine Inc. Mind Medicine Inc. had no role in
planning or conducting the study or writing the publication.
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15
Figure legends:
Fig. 1. Chromatographic analyte separation in human plasma. Concentrations of analytes in plasma
were as follows: DMT: 100 ng/mL; DMT-NO: 100 ng/mL; 13C6-IAA: 10,000 ng/mL. The ISTD solution
contained 5 ng/mL DMT-d6 and 500 ng/mL IAA-d2. The IAA signal is the result of endogenous IAA in
the plasma sample and corresponds to 296 ng/mL IAA, calculated using the 13C6-IAA CAL line.
Fig. 2. 13C6-IAA CAL lines can be used to calculate IAA concentrations. A) IAA CAL lines are
linear in water but not plasma due to endogenous IAA. B) Ratio of IAA to 13C6-IAA signal increases at
low concentrations in plasma due to endogenous IAA. The shaded area shows the range of 0.85–1.15.
C) 13C6-IAA CAL lines can be used to calculate IAA concentrations in plasma. The plasma was spiked
with IAA at different concentrations; blank IAA signals were subtracted from the total signals and IAA
concentrations were calculated using 13C6-IAA CAL lines. Data points colored in red were outside the
accepted accuracy range of 85–115%.
Fig. 3. Accuracy in human plasma at different QC levels. Inter-assay accuracy and precision of three
independent assays. Data points outside the accepted accuracy range of 85–115% (LLOQ: 80–120%)
are colored in red.
Fig. 4. Selectivity of DMT, DMT-NO, and 13C6-IAA in human plasma. An overlay of seven LLOQ
chromatograms per analyte (colored lines) is compared to double blank and blank signals (black lines).
Each LLOQ and blank signal was measured in plasma originating from different donors. The blank
plasma signal intensity was < 20% of the LLOQ signal for each analyte.
Fig. 5. Analyte stability under various storage conditions. All analytes were sufficiently stable when
reinjected after storage in the autosampler at 10 °C overnight, at room temperature for 8 h, or at -20 °C
for up to 3 months. Furthermore, three freeze-thaw cycles did not affect analyte stability. Signals that
did not meet the required accuracy of 85–115% (LLOQ: 80–120%) are colored in red.
Fig. 6. Human plasma concentrations of DMT, DMT-NO, and IAA. Five healthy subjects were
intravenously perfused with 1 mg/min DMT. Concentration-time curves are shown individually in linear
scale and combined in logarithmic scale. Data points indicate mean plasma levels + SD, n=5.
16
Tables
Table 1. Mass spectrometric settings for the quantification of the analytes and internal standards.
Analyte
MRM (m/z) (Q1 → Q3)
RT (min)
DP (V)
EP (V)
CE (V)
CXP (V)
DMT
189.1 → 58.1
quantifier
1.68
56
10
27
10
189.1 → 144.1
qualifier
1.68
56
10
27
10
DMT-NO
205.1 → 144.0
quantifier
2.21
66
10
21
10
205.1 → 115.0
qualifier
2.21
66
10
59
6
13C6-IAA
182.0 → 136.1
quantifier
3.31
56
10
65
8
182.0 → 109.0
qualifier
3.31
56
10
28
18
IAA
176.0 → 130.1
quantifier
3.31
56
10
61
8
176.0 → 103.0
qualifier
3.31
56
10
28
18
DMT-d6
195.0 → 64.1
ISTD quantifier
1.68
31
10
27
12
195.0 → 144.0
ISTD qualifier
1.68
31
10
29
10
IAA-d2
178.0 → 132.1
ISTD quantifier
3.31
56
10
47
8
178.0 → 105.1
ISTD qualifier
3.31
96
10
37
18
CE, collision energy; CXP, collision cell exit potential; DP, declustering potential; EP, entrance potential; m/z, mass-to-charge ratio; MRM, multiple reaction
monitoring; Q1, quadrupole 1; Q3, quadrupole 3; RT, retention time; V, volt.
17
Table 2. Recovery and matrix effect at different QC levels.
Analyte
QC level
%Recovery (%CV)
%Matrix effect (%CV)
DMT
LLOQ
99.0 (7.1)
104 (3.3)
LQC
95.5 (7.4)
102 (4.9)
MQC
99.6 (5.6)
102 (4.0)
HQC
97.9 (2.9)
103 (2.4)
all QC levels
97.6 (1.8)
105 (4.2)
DMT-NO
LLOQ
110 (9.5)
110 (8.5)
LQC
98.3 (4.1)
110 (3.3)
MQC
102 (2.3)
105 (2.4)
HQC
102 (3.4)
105 (2.3)
all QC levels
103 (4.4)
107 (2.6)
13C6-IAA
LLOQ
102 (9.3)
109 (6.0)
LQC
98.7 (8.2)
96.6 (4.7)
MQC
100 (4.1)
98.6 (2.7)
HQC
101 (5.1)
98.6 (2.7)
all QC levels
102 (2.9)
102 (6.8)
18
CRediT author statement
Dino Luethi: methodology, validation, formal analysis, investigation, data curation, writing – original draft, writing – review & editing, visualization,
funding acquisition. Karolina E. Kolaczynska: methodology, investigation, writing – review & editing. Severin B. Vogt: resources, writing – review & editing.
Laura Ley: resources. Livio Erne: resources. Matthias E. Liechti: writing – review & editing, supervision, project administration, funding acquisition. Urs
Duthaler: conceptualization, methodology, validation, formal analysis, data curation, writing – review & editing, visualization, supervision, project administration.
19
Declaration of competing interest
M.E.L. is a consultant for Mind Medicine, Inc. All other authors do not have any conflicts of interest to declare for this work. Knowledge and data associated
with this work are owned by the University Hospital Basel and were licensed by Mind Medicine Inc. Mind Medicine Inc. had no role in planning or conducting the
study or writing the publication.
20
Highlights:
-An easy-to-use LC-MS/MS method for DMT and its two major metabolites was developed
-The method allows sensitive analyte quantification and rapid sample measurement
-The endogenously occurring metabolite IAA was quantified using 13C6-IAA calibrations
-Only small volumes of plasma (50 μL) are required for sample analysis
21
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23
24
25
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