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Title: doi:10.1016/j.chroma.2007.08.049

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Journal of Chromatography A, 1167 (2007) 195–201

Quantitative determination of 4-ethylphenol and 4-ethyl-2-methoxyphenol
in wines by a stable isotope dilution assay
Sierra Rayne, Nigel J. Eggers ∗
Chemistry, Earth & Environmental Sciences, Irving K. Barber School of Arts & Sciences, The University of British Columbia at Okanagan,
3333 University Way, Kelowna, British Columbia V1V 1V7, Canada
Received 20 March 2007; received in revised form 17 August 2007; accepted 23 August 2007
Available online 26 August 2007

Abstract
The deuterium-labelled standards 4-ethylphenol-d3 and 4-ethyl-2-methoxyphenol-d3 were synthesized and utilized in a rapid, sensitive, and
accurate stable isotope dilution assay for 4-ethylphenol and 4-ethyl-2-methoxyphenol in wine. For a 5-mL sample of a Merlot wine, quantitation
was reliable down to 500 ng/L for 4-ethylphenol and 100 ng/L for 4-ethyl-2-methoxyphenol at estimated signal-to-noise ratio of 3:1, respectively.
The concentrations of 4-ethylphenol and 4-ethyl-2-methoxyphenol were also measured in 54-barrelled red commercial wines from the Okanagan
Valley in British Columbia. The results indicate significantly different internal standard recoveries for the two analytes, demonstrating the need for
individual stable isotope derivatives to reliably quantitate 4-ethylphenol and 4-ethyl-2-methoxyphenol in wines.
© 2007 Elsevier B.V. All rights reserved.
Keywords: 4-Ethylphenol; 4-Ethyl-2-methoxyphenol; Brettanomyces; Stable isotope dilution analysis; Aroma compound; Wine

1. Introduction
Brettanomyces is wild yeast implicated in the spoilage of
wine and has long been associated with European wines, but
in recent years is also considered to occur in wines from the
New World. Two compounds that are widely considered to
be primarily responsible for the Brettanomyces aroma in wine
are 4-ethylphenol and 4-ethyl-2-methoxyphenol, although other
volatile phenols [1], acetic acid [2], isovaleric (3-methylbutyric)
acid [3], and tetrahydropyridines [4] have been linked to
Brettanomyces-sourced organoleptic defects. Recent work even
suggests that isovaleric acid may be the dominant odorant in
some high-Brettanomyces wines [3,5].
Coupled with this complexity, not all strains of Brettanomyces (or replicates of the same strain) produce equivalent
amounts of 4-ethylphenol and 4-ethyl-2-methoxyphenol given
equal starting conditions of nutrients and other growth factors
[5]. The relationship between Brettanomyces numbers and levels
of 4-ethylphenol is problematic, as a number of leading research
groups have noted poor or absent correlations (see, e.g., ref.



Corresponding author. Tel.: +1 250 807 9575; fax: +1 250 807 8004.
E-mail address: nigel.eggers@ubc.ca (N.J. Eggers).

0021-9673/$ – see front matter © 2007 Elsevier B.V. All rights reserved.
doi:10.1016/j.chroma.2007.08.049

[6]). At low concentrations, these compounds are thought by
some to add to wine complexity, but at high concentrations the
wine is considered spoiled and has been described as ‘animal’,
‘barnyard’, ‘stable’, ‘phenolic’, and ‘mousy’. Odor thresholds
for the two compounds have some variation, although in general,
4-ethylphenol is perceived in model wines at about 500 ␮g/L,
whereas 4-ethyl-2-methoxyphenol can be distinguished at levels
about an order of magnitude lower near 50 ␮g/L [7].
Brettanomyces (the asexual, nonsporulating form) and
Dekkera (its sexual, sporulating form) are ubiquitous in the vineyard and winery and are likely to be present in the water, soil,
grapes and must, and throughout the winery, their presence can
be monitored but not controlled [6]. Once this yeast is established in a winery, it is difficult to eliminate. Spoilage of wine
by Brettanomyces can be devastating and wineries have had to
shut down to remove this contaminant. Maintaining appropriate sulfur dioxide (SO2 ) levels, lower temperatures (generally
<20 ◦ C), filtration, excluding oxygen ingress into the wine during topping up of barrels (and keeping the barrels topped up),
and general winery hygiene have all been noted as methods to
keep Brettanomyces problems minimized [6]. However, infected
barrels cannot effectively be sterilized due to their large internal surface areas and porosity, whether it be by washing with
sulfite water, shaving and firing, or ozone treatment [6,8], with

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S. Rayne, N.J. Eggers / J. Chromatogr. A 1167 (2007) 195–201

Brettanomyces found as deep as 8 mm into the oak wood [9].
This suggests that to truly minimize future problems in a winery that has already experienced Brettanomyces contamination,
previously infected barrels may need to be discarded.
For these reasons, rigorous analytical methods are required
for the two most well-known Brettanomyces metabolites, 4ethylphenol and 4-ethyl-2-methoxyphenol, in order to facilitate
reliable in-depth studies targeted at better understanding the levels, distributions, and factors governing the production of these
compounds in wine. Most studies to date have attempted to
quantify 4-ethylphenol and 4-ethyl-2-methoxyphenol in wines
using non-isotopically labelled internal standards such as 3,4dimethylphenol (see, e.g., ref [7,10–17]), 2-octanol [14,18],
3-octanol [19,20], 4-decanol [21], ␥-hexalactone [22], and 4methylphenol [23]. Because of the complexity of the wine matrix
and differences between individual samples, these methods are
unlikely to give equal recoveries between the internal standard
and the analyte of interest, and may be unreliable.
On the other hand, stable isotope dilution analysis (SIDA) is
an accurate means for quantifying low concentrations of odorants, and its advantages have been outlined previously [24,25].
In principle, addition of an isotopically labelled odorant to a
sample results in a more absolute quantitative measurement of
this odorant, as the isotopomer of the respective analyte is used
to more fully correct for losses of analytes during extraction
and cleanup. Relatively few studies have attempted to develop
SIDA methods for 4-ethylphenol and 4-ethyl-2-methoxyphenol
in wines. Pollnitz et al. have previously synthesized an aromatic
ring labelled 4-ethylphenol-d4 derivative from the acetate of
polydeuterated phenol via the Fries rearrangement followed by
a Wolff–Kischner reduction [8], and subsequently used this in a
series of studies for the quantitation of both 4-ethylphenol and
4-ethyl-2-methoxyphenol in wines [26,27]; other groups also
used the 4-ethylphenol-d4 standard from ref. [8] in their work
[28–30].
However, no studies to date have used a SIDA method for
4-ethyl-2-methoxyphenol, thus preventing accurate determination of both important Brettanomyces metabolites in the same
sample. For this reason, we sought to synthesize stable isotope
derivatives of both 4-ethylphenol and 4-ethyl-2-methoxyphenol,
develop a SIDA method for wines using these labelled surrogate
standards, and then validate and test the method on a suite of real
wine samples.
2. Experimental
2.1. General
All solvents were reagent grade and obtained from Caledon
Laboratories (Georgetown, ON, Canada). All other chemicals
were from Sigma–Aldrich (Milwaukee, WI, USA) and of purity
>99% (checked by GC–MS where practical). Prior to use, all
glassware was washed with commercial AlconoxTM detergent
in a laboratory dishwasher, rinsed three times with sequential
washes of tap water and E-Pure deionized water, followed by
rinses with acetone, toluene, and dichloromethane (in that order).
Following cleaning, glassware was then baked in a vented oven

at 350 ◦ C for a minimum of 24 h prior to use. Nuclear magnetic
resonance (NMR) spectra were obtained using a Varian Mercury
Plus 400 NMR (Varian NMR Systems; Palo Alto, CA, USA) (1 H
at 400.1 MHz, 13 C at 100.6 MHz). All samples were dissolved
in deuterochloroform and the solvent (chloroform-d) was used
as the internal standard (7.25 ppm in 1 H and 77.0 ppm in 13 C).
Mass spectra, to aid in structural confirmation of the synthesized isotopically labelled internal standards, were obtained
using a Varian 3800 gas chromatograph (GC) coupled to a Varian
saturn 2000 mass spectrometer (MS) operating in the positive ion
electron impact (EI) ionization mode at 70 eV. The GC column
was a Hewlett packard HP-5MS brand (5% phenyl–95% methylsiloxane) with dimensions of 30 m × 0.25 mm × 0.50 ␮m. The
carrier gas was ultra high-purity helium at a constant flow rate
of 1.5 mL/min. A liquid sample of 1 ␮L (dichloromethane as
solvent) was manually injected (20:1 split ratio) onto the column. The MS trap temperature was 170 ◦ C, the MS manifold
temperature was 45 ◦ C, and the GC–MS transfer line temperature was 230 ◦ C. MS scans were obtained in the full-scan (auto)
mode at unit resolution with an emission current of 50 ␮A and
a scan time of 0.5 s over the mass range m/z 40–400. The GC
injector temperature was constant at 220 ◦ C over the course of a
sample run, with the oven temperature held at 40 ◦ C for 1 min,
ramped to 220 ◦ C at 2 ◦ C/min, and held at 220 ◦ C for 30 min for
a total run time of 121 min. Under this temperature program, 4ethylphenol-d3 eluted at a Kovat’s retention index (RI) of 1170,
and 4-ethyl-2-methoxyphenol-d3 eluted at a RI of 1285.
2.2. Synthesis of 4-ethylphenol-d3 and
4-ethyl-2-methoxyphenol-d3
Dry tetrahydrofuran (10 mL) was combined with NaH
(2.00 g, 83.3 mmol) and deuterium oxide (9.1 mL, 500 mmol) in
a pressure tube. The solution was stirred for 10 min, after which
1-(4-hydroxyphenyl)ethanone (2.82 g, 20.7 mmol) was added to
the tube. The pressure tube was sealed and placed in a 60 ◦ C
sand bath for 72 h, after which the tube was removed from the
heat source and cooled in an ice bath. Once cooled, the reaction
mixture was transferred to a 50 mL Erlenmeyer flask and the
reaction was quenched with 3 M acetic acid. Litmus paper was
used to determine the endpoint. The organic layer was extracted
with CH2 Cl2 (3 × 10 mL) and the combined organic layers were
washed with water (5 mL) to remove excess acetic acid. The
organic layer was dried over MgSO4 and the solvents were
removed by rotary evaporation. 1-(4-Hydroxyphenyl)ethanoned3 (1.69 g, 58.8%) was recovered as a yellow-orange solid and
was sufficiently pure for use in the next step.
Methanol (100 mL) was combined with palladium on carbon (100 mg) in a 250 mL two-necked round-bottom flask.
The solution was stirred for 15 min, after which 1-(4hydroxyphenyl)ethanone-d3 (425 mg, 3.1 mmol) was added to
the flask. The flask was evacuated via a water vacuum while
stirring for 5 min, after which H2 gas was bubbled through the
solution at atmospheric pressure (1 bar) for 10 min. While sparging with H2 gas, the flask was sealed to the ambient atmosphere
and the gas flow stopped. Following isolation from the surroundings, the flask was stirred for 12 h, and then the H2 sparging was

S. Rayne, N.J. Eggers / J. Chromatogr. A 1167 (2007) 195–201

197

Fig. 1. Reaction sequence used in the synthesis of 4-ethylphenol-d3 and 4-ethyl-2-methoxyphenol-d3 .

repeated for 15 min. This 15 min sparge–12 h isolation sequence
was repeated a total of six times over the course of 72 h.
The resulting solution was filtered into a 250 mL beaker
using rinsings of methanol, and the solvent allowed to evaporate overnight. The crude product was purified by atmospheric
pressure silica gel column chromatography using 1:1 (v/v)
hexanes:diethyl ether as the eluant to give 4-ethylphenol-d3
(265 mg, 68.4%) as a light yellow oil (Fig. 1): 1 H NMR (CDCl3 ,
400 MHz) ␦2.61 (s, 2 H, benzylic CH2 ), 5.48 (br, s, 1 H,
exchangeable with D2 O, phenolic OH), 6.74 (XX protons of
AA XX , 2 H, JAB = 8.2 Hz, ring H adjacent to –OH group),
7.05 (AA protons of AA XX , 2 H, JAB = 8.2 Hz, ring H adjacent
to ethyl group). 13 C NMR (CDCl3 , 100.6 MHz) ␦15.8 (terminal CH3 ), 28.0 (benzylic CH2 ), 115.4 (aromatic CH ortho to
phenolic group), 129.0 (aromatic CH meta to phenolic group),
153.1 (aromatic C attached to phenolic group). MS m/z (ion,
relative % intensity) 125 (M+ , 46), 107 ([M-CD3 ]+ , 100), 93
([M-CD3 CH2 ]+ , 2), 77 ([M-CD3 CH2 O]+ , 2).
Dry tetrahydrofuran (10 mL) was combined with NaH
(0.97 g, 40.4 mmol) and deuterium oxide (5.2 mL, 290 mmol) in
a pressure tube. The solution was stirred for 10 min, after which
1-(4-hydroxy-3-methoxyphenyl)ethanone (2.12 g, 12.8 mmol)
was added to the tube. The pressure tube was sealed and placed
in a 60 ◦ C sand bath for 72 h after which the tube was removed
from the heat source and cooled in an ice bath. Once cooled,
the reaction mixture was transferred to a 50 mL Erlenmeyer
flask and the reaction was quenched with 3 M acetic acid. Litmus paper was used to determine the endpoint. The organic
layer was extracted with CH2 Cl2 (3 × 10 mL) and the combined
organic layers were washed with water (5 mL) to remove excess
acetic acid. The organic layer was dried over MgSO4 and the
solvents were removed by rotary evaporation. 1-(4-Hydroxy-3methoxyphenyl)ethanone-d3 (0.51 g, 24.0%) was recovered as
a yellow oil and was sufficiently pure for use in the next step.
Methanol (50 mL) was combined with palladium on carbon (50 mg) in a 125 mL two-necked round-bottom flask. The
solution was stirred for 15 min after which 1-(4-hydroxy-3methoxyphenyl)ethanone-d3 (250 mg, 1.5 mmol) was added to
the flask. The flask was evacuated via a water vacuum while
stirring for 5 min, after which H2 gas was bubbled through the
solution at atmospheric pressure (1 bar) for 10 min. While sparging with H2 gas, the flask was sealed to the ambient atmosphere
and the gas flow stopped. Following isolation from the surroundings, the flask was stirred for 12 h, and then the H2 sparging was
repeated for 15 min. This 15 min sparge–12 h isolation sequence
was repeated a total of six times over the course of 72 h.

The resulting solution was filtered into a 150 mL beaker
using rinsing of methanol, and the solvent allowed to
evaporate overnight. The crude product was purified by atmospheric pressure silica gel column chromatography using 1:1
(v/v) hexanes:diethyl ether as the eluant to give 4-ethyl-2methoxyphenol-d3 (128 mg, 55.1%) as a dark brown oil: 1 H
NMR (CDCl3 , 400 MHz) ␦2.58 (s, 2 H, benzylic CH2 ), 5.65 (br,
s, 1 H, exchangeable with D2 O, phenolic OH), 6.68–6.85 (m, 3
H, aromatic ring). 13 C NMR (CDCl3 , 100.6 MHz) ␦15.8 (terminal CH3 ), 28.7 (benzylic CH2 ), 55.8 (methoxy), 110.4 (aromatic
CH para to phenolic group), 114.3, 120.2, 136.2, 143.5 (aromatic C attached to methoxy group), 146.3 (aromatic C attached
to phenolic group). MS m/z (ion, relative % intensity) 155 (M+ ,
78), 137 ([M-CD3 ]+ , 100), 122 ([M-CD3 CH3 ]+ , 3).
The high isotopic purity of the synthesized surrogate
standards was indicated by 1 H NMR and MS. Confirmation of the successful incorporation of three deuterium into
4-ethylphenol-d3 and 4-ethyl-2-methoxyphenol-d3 was demonstrated by the absence of the three proton 1 H NMR singlets
at ␦1.20 and 1.21 that are present in the unlabelled native
analogs 4-ethylphenol and 4-ethyl-2-methoxyphenol, respectively. As well, the benzylic –CH2 protons at ␦2.61 and
2.58 in the 1 H NMR of 4-ethylphenol-d3 and 4-ethyl-2methoxyphenol-d3 , respectively, are singlets with an integration
of 2, indicating no significant presence of neighbouring protonated carbons. In the 1 H NMR of both 4-ethylphenol-d3
and 4-ethyl-2-methoxyphenol-d3 , small unresolved peaks with
relative integration values of 0.006 and 0.011 were present
in the purified product at ␦1.20 and 1.21. The residual proton peaks indicate approximately 99.8% and 99.6% deuterium
incorporation into the desired terminal methyl position for 4ethylphenol-d3 and 4-ethyl-2-methoxyphenol-d3 , respectively.
For 4-ethylphenol, the observed relative intensities in the
molecular ion cluster (m/z 120 (4%), 121 (32%), 122 (100%),
and 123 (11%)) closely matched those of 4-ethylphenol-d3
(123 (4%), 124 (26%), 125 (100%), and 126 (15%)). For
4-ethyl-2-methoxyphenol, the observed relative intensities in
the molecular ion cluster (m/z 150 (3%), 151 (21%), 152
(100%), and 153 (13%)) were similar to those of 4-ethyl-2methoxyphenol-d3 (153 (5%), 154 (36%), 155 (100%), and
156 (32%)). It is important to note that the [M-3]+ ions of 4ethylphenol-d3 (m/z 122) and 4-ethyl-2-methoxyphenol-d3 (m/z
152) have relative intensities only 1% that of the M+ ion. Thus,
there is negligible contamination of the molecular ion channels
for 4-ethylphenol and 4-ethyl-2-methoxyphenol used for native
analyte quantitation by interfering ions from the corresponding

198

S. Rayne, N.J. Eggers / J. Chromatogr. A 1167 (2007) 195–201

labelled standards. In addition, the native and labelled standards
do not fully coelute on the GC program, further minimizing any
interference in lower mass channels by the labelled standards.
Fragmentation of both 4-ethylphenol-d3 and 4-ethyl-2methoxyphenol-d3 involves loss of the terminal –CD3 label (18
mass units) as expected to yield the stabilized benzylic radical
cation as the base peak (m/z 107 for 4-ethylphenol-d3 and 137
for 4-ethyl-2-methoxyphenol-d3 ). The native analytes similarly
lose the terminal –CH3 group (15 mass units) during MS fragmentation. Since both the native and surrogate analytes have the
same base peak, this ion ([M-CD3 ]+ for 4-ethylphenol-d3 and
4-ethyl-2-methoxyphenol-d3 and [M-CH3 ]+ for 4-ethylphenol
and 4-ethyl-2-methoxyphenol) cannot be used for quantitation,
but is used for qualifying purposes to aid in identity confirmation. Thus, the molecular ions of all analytes (m/z 122 and
125 for 4-ethylphenol and 4-ethylphenol-d3 , respectively, and
m/z 152 and 155 for 4-ethyl-2-methoxyphenol and 4-ethyl-2methoxyphenol-d3 , respectively) were selected for quantitative
analysis in wines. Both labelled analytes are stable as ethanolic solutions when kept at room temperature, but do decompose
slightly over time when stored as oils, necessitating periodic
repurification.
2.3. Sample collection and processing
A selection of 2005 vintage Okanagan Valley (British
Columbia, Canada) barrelled red wines were obtained from local
wineries operating at the small, medium, and large production
scales. Samples were collected from 225 L oak barrels using
a 50 mL glass volumetric pipette that had been sterilized with
neat ethanol prior to use and placed in 50 mL amber glass jars for
transport and storage prior to analysis. Samples were stored at
4 ◦ C prior to analysis, with storage time ranging from <24 h–30
days.
A sample of wine (5 mL) was pipetted into a 10 mL test
tube containing 2 g of sodium chloride. The sample was spiked
with 20 ␮L of a (101.5 mg/L, in ethanol) 4-ethylphenol-d3 and
(118.0 mg/L, in ethanol) 4-ethyl-2-methoxyphenol-d3 standard
solution using a calibrated microliter syringe (25 ␮L) (Hamilton; Reno, NV, USA). These internal standard additions (2.03 ␮g
4-ethylphenol-d3 and 2.36 ␮g/L 4-ethyl-2-methoxyphenol-d3 )
correspond to concentrations of 406 ␮g/L 4-ethylphenol-d3 and
472 ␮g/L 4-ethyl-2-methoxyphenol-d3 in the 5 mL wine sample
to be analyzed. The solution was shaken for 10 s to ensure mixing, and diethyl ether (2 mL) was added to the vial and shaken
again for 20 s. The sample was centrifuged until two distinct layers formed (typically 2 min). A portion (ca. 1 mL) of the organic
phase was transferred directly from the test tube to a 3 mL vial,
capped, and the extract injected directly into the GC–MS under
the instrumental conditions described below.
2.4. Sample analysis
Standard solutions were prepared by dissolving known
amounts of the unlabelled native and deuterium-labelled surrogate analytes in ethanol in volumetric flasks to provide the
desired concentrations. Wine extracts were analyzed using a

thermo electron trace GC coupled to a thermo electron DSQ
dual-stage quadrupole MS operating in the positive ion electron impact (EI) ionization mode at 70 eV. The GC column
was an Agilent/J&W DB-1701 ((14% cyanopropyl-phenyl)methylpolysiloxane) with dimensions of 30 m length × 0.25 mm
inside diameter × 0.25 ␮m film thickness. The carrier gas was
ultra high-purity helium at a constant flow rate of 1.2 mL/min
without vacuum compensation. A liquid sample of 2 ␮L (diethyl
ether as solvent) was injected using a thermo electron AI
3000 autosampler. The autosampler syringe was rinsed both
pre- and post-injection with methanol (first three rinses) and
dichloromethane (second three rinses), and was rinsed once during pre-injection with the sample solution of interest. Pre- and
post-injection dwell times for the syringe were 2 s.
The GC injector was operated in the split/splitless mode, with
the splitless time of 1 min followed by a split flow of 50 mL/min.
The GC injector temperature was constant at 220 ◦ C over the
course of a sample run, with the oven temperature held at 40 ◦ C
for 1 min, ramped to 260 ◦ C at 8 ◦ C/min, and held at 260 ◦ C
for 1 min, for a total run time of 30 min (with an equilibration
time of 0.5 min prior to injection). The MS ion source temperature was 200 ◦ C and the GC–MS transfer line temperature was
250 ◦ C. A 2 min solvent delay was included on the GC–MS
program, and the MS filament was turned off after 11 min.
MS scans were obtained in the selected ion monitoring (SIM)
mode operating at unit resolution with an emission current of
100 ␮A and a dwell time of 100 ms at each of the following
masses: m/z 107, 122, 125, 137, and 155. Under this temperature program, the analyte retention indices were as follows:
4-ethylphenol, RI = 1170; 4-ethylphenol-d3 , RI = 1175; 4-ethyl2-methoxyphenol, RI = 1285; and 4-ethyl-2-methoxyphenol-d3 ,
RI = 1291.
Samples were prepared for GC–MS analysis by placing 1 mL
of solution in a 3 mL clear glass vial covered with a plastic
septa and cap. All samples were analyzed with the extract at
room temperature (ca. 25 ◦ C) for at least 2 h prior to injection. Samples were processed in batches of 16 (two separate
batches of eight samples), containing 14 unknown extracts, a
procedural blank, and one of the calibration standards. One
batch of eight samples contained seven unknown extracts and
the procedural blank (which checks for background contamination during the method using deionized water as the solution to
be extracted and analyzed), and the next batch of eight samples
contained seven unknown extracts and a calibration standard.
The calibration standard concentration as part of the ongoing
in-house quality assurance–quality control (QA–QC) program
was continuously varied among the five available solutions
(containing 0–3000 ␮g/L of 4-ethylphenol and 0–660 ␮g/L 4ethyl-2-methoxyphenol) to ensure no drift in the method was
occurring in any region of the calibration range. We also considered potential artifactual generation of the analytes on the
elevated injector block temperatures. The analytical method was
performed using injector temperatures of 150, 200, 220, and
250 ◦ C on five wine samples analyzed in triplicate. We did not
observe any significant difference (p < 0.05 using single-factor
ANOVA) in the determined concentrations of the analytes as a
function of injector temperature.

S. Rayne, N.J. Eggers / J. Chromatogr. A 1167 (2007) 195–201

The target analytes (both native compounds and surrogate deuterium-labelled internal standards) were identified only
when the GC–MS data satisfied all of the following criteria: (1) two ions from each of the individual analytes were
detected by their masses (m/z 107 as qualifying ion for both 4ethylphenol and 4-ethylphenol-d3 , m/z 137 as qualifying ion for
both 4-ethyl-2-methoxyphenol and 4-ethyl-2-methoxyphenold3 , and the following target and quantification ions by analyte:
4-ethylphenol, m/z 122; 4-ethylphenol-d3 , m/z 125; 4-ethyl-2methoxyphenol, m/z 152; and 4-ethyl-2-methoxyphenol-d3 , m/z
155) with the mass spectrometer operating at unit resolving
power during the entire chromatographic run, (2) the retention
time of the specific peaks was within 5 s to the predicted time
obtained from analysis of authentic compounds in the calibration standards, (3) the peak maxima for both characteristic ions
(qualifying plus target) of a specific analyte coincided within
3 s, (4) the observed ratio of the two ions monitored per analyte
were within 15% of the ratio found during instrument calibration runs, and (5) the signal-to-noise ratio resulting from the
peak response of the two corresponding ions was ≥3 for proper
quantification of the analyte.
Concentrations of identified compounds and their method
detection limits (MDLs) were calculated by the internal standard isotope-dilution method using the mean relative response
factors (RRFs) determined from calibration standard runs as
described below. All post-processing for analyte identification
and quantitation (including determination of peak areas) was
done manually using the Thermo Electron Xcalibur software (v.
1.4 SR1).
Calibration curves were generated from integrated peak
area ratios (peak areas of (4-ethylphenol m/z 122 peak)/(4ethylphenol-d3 m/z 125 peak) and (4-ethyl-2-methoxyphenol
m/z 152 peak)/(4-ethyl-2-methoxyphenol-d3 m/z 155 peak)
and plotted against the mass of each respective native analyte injected within 2 ␮L of the corresponding standard
solution (0–15 ng for 4-ethylphenol and 0–3.3 ng for 4-ethyl-2methoxyphenol). These native analyte mass ranges correspond
to those obtained by 2 ␮L injections of 2 mL diethyl ether
extracts containing 100% extraction efficiency, 0–3000 ␮g/L
4-ethylphenol, and 0–660 ␮g/L 4-ethyl-2-methoxyphenol from
a wine sample. Three replicate injections were performed at
each concentration. Quantitation was reliable down to 500 ng/L
for 4-ethylphenol and 100 ng/L for 4-ethyl-2-methoxyphenol at
estimated signal-to-noise ratios of 3:1, respectively, for a Merlot
wine.
The stable isotope dilution assay was validated by a standard
addition experiment using 12 individual 5 mL samples of a Merlot wine. The samples were processed and analyzed by stable
isotope dilution analysis with 4-ethylphenol-d3 and 4-ethyl-2methoxyphenol-d3 as the internal standards using the method as
described above for unknown wines. Duplicate extractions were
performed at each concentration of native analyte added.
3. Results and discussion
Using 4-ethylphenol-d3 and 4-ethyl-2-methoxyphenol-d3 as
internal standards, 4-ethylphenol and 4-ethyl-2-methoxyphenol

199

could be quantified from triplicate 5 mL samples of wine at concentrations down to 500 ng/L and 100 ng/L, respectively, at a
signal-to-noise ratio of 3:1. With a larger sample of wine (5 mL
was used in the present study), it may be possible to extend
the method detection limit for both analytes to near 1 ng/L. The
reproducibility of the method was also assessed by triplicate
5 mL wine samples, and for both analytes, the relative standard
deviations decreased from about 7–8% at levels near 5 ␮g/L to
4% at about 100 ␮g/L.
The method was validated by a series of duplicate
standard additions of 4-ethylphenol (0–530 ␮g/L) and 4-ethyl-2methoxyphenol (0–564 ␮g/L) to an Okanagan Valley Merlot red
wine. The standard addition curve obtained was linear throughout the concentration range, with a coefficient of determination
(r2 ) of 0.989 and linear regression equation y = 1.05x + 121 for
4-ethylphenol, and a r2 = 0.991 and linear regression equation
y = 1.11x + 67.6 for 4-ethyl-2-methoxyphenol. This Okanagan
Merlot showed a 4-ethylphenol concentration of 115 ␮g/L and a
4-ethyl-2-methoxyphenol concentration of 61 ␮g/L. The regression slopes greater than unity for each target analyte indicate that
less of the deuterium-labelled surrogate standard was recovered
during each extraction than for the corresponding native analyte (5% and 11% greater recovery of the native analyte than the
labelled analog for 4-ethylphenol and 4-ethyl-2-methoxyphenol,
respectively). These higher recoveries of the native analyte versus the labelled analog are in the range previously reported for
4-ethylphenol (7–21%) [8].
The analytical procedure was tested by determining 4ethylphenol and 4-ethyl-2-methoxyphenol in 54 duplicate
barrelled red commercial wines from the Okanagan Valley in
British Columbia (Table 1). Concentrations detected in the 54
wines analyzed ranged from <0.5 to 586 ␮g/L 4-ethylphenol
(average 58.2 ␮g/L) and from 4.3 to 411 ␮g/L 4-ethyl-2methoxyphenol (average 86.6 ␮g/L), with the percent range
of concentrations from duplicate analyses between 0.1 and
9.7%. In comparison with concentrations reported from other
worldwide winemaking regions, the levels of 4-ethylphenol and
4-ethyl-2-methoxyphenol we found in the Okanagan barrelled
reds are quite low, although there is considerable variation in
concentrations of these two compounds reported in the literature. Chatonnet et al. [7] analyzed 137 red and white wines
from France, and found that white wines contained an average of 3 ␮g/L 4-ethylphenol (range 0–28 ␮g/L) compared to
red wines with an average of 440 ␮g/L (range 1–6047 ␮g/L).
As well, in a comprehensive survey of barrelled and bottled
red wines from Australia, Pollnitz et al. reported barrelled
4-ethylphenol ranges of 385–680 ␮g/L and bottled ranges of
2–2660 ␮g/L (mean 795 ␮g/L) [8,27]. The corresponding ranges
of 4-ethyl-2-methoxyphenol were 28–45 ␮g/L in barrelled reds
and 1–437 ␮g/L (mean 99 ␮g/L) in bottled reds from Australia.
As noted above, previous work [8,26–30] has used a SIDA
for 4-ethylphenol, and then applied the recovery of the 4ethylphenol-d4 standard to determine 4-ethyl-2-methoxyphenol
concentrations. Such an approach is premised on the assumption that the recovery of the 4-ethylphenol-d4 standard in
each sample would be equivalent to the hypothetical recovery
of an isotopically labelled 4-ethyl-2-methoxyphenol stan-

200

S. Rayne, N.J. Eggers / J. Chromatogr. A 1167 (2007) 195–201

Table 1
Determination of 4-ethylphenol and 4-ethyl-2-methoxyguaiacol concentrations
in 54 commercial 2005 vintage barrelled Okanagan Valley red wines
Variety

4-Ethylphenol (␮g/L)
(%)

4-Ethyl-2-methoxyphenol
(␮g/L) (%)

Blend of Cabernet
Sauvignon/Merlot/
Cabernet Franc
Cabernet Franc
Cabernet Sauvignon
Cabernet Sauvignon
Cabernet Sauvignon
Cabernet Sauvignon
Cabernet Sauvignon
Cabernet Sauvignon
Cabernet Sauvignon
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Merlot
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Pinot Noir
Syrah
Syrah
Syrah
Syrah
Syrah
Syrah
Syrah
Syrah
Syrah

16.3 ± 0.7 (49.1)

43.7 ± 0.9 (94.4)

<0.5 (88.5)
<0.5 (56.4)
135.1 ± (82.4)
<0.5 (55.0)
147.5 ± 12.3 (42.0)
144.0 ± 2.8 (73.7)
7.3 ± 0.4 (93.6)
57.1 ± 1.4 (84.5)
<0.5 (52.8)
<0.5 (92.3)
29.6 ± 0.9 (87.8)
39.5 ± 1.8 (80.0)
106.7 ± 0.4 (67.7)
119.8 ± 6.4 (59.6)
18.2 ± 0.4 (59.6)
<0.5 (77.1)
16.2 ± 0.9 (69.0)
<0.5 (63.2)
84.9 ± 6.6 (45.0)
199.7 ± 5.8 (78.1)
55.2 ± 3.6 (78.1)
74.9 ± 0.3 (57.6)
<0.5 (55.0)
22.1 ± 1.7 (56.1)
0.6 ± 0.1 (43.6)
21.4 ± 0.1 (55.9)
5.8 ± 0.3 (82.8)
<0.5 (78.4)
54.6 ± 0.1 (44.7)
3.9 ± 0.4 (67.5)
80.9 ± 3.8 (50.6)
1.0 ± 0.2 (43.5)
23.6 ± 0.7 (84.0)
51.2 ± 2.4 (88.5)
31.7 ± 2.9 (98.1)
91.0 ± 2.3 (92.9)
39.5 ± 0.6 (85.1)
586.2 ± 10.1 (43.1)
33.9 ± 2.0 (93.7)
19.7 ± 1.4 (64.1)
72.9 ± 4.2 (87.7)
<0.5 (55.9)
117.4 ± 0.5 (68.5)
2.9 ± 0.3 (64.7)
200.6 ± 13.7 (91.2)
92.7 ± 5.5 (93.4)
39.3 ± 1.2 (90.7)
125.4 ± 0.2 (55.8)
44.7 ± 1.3 (80.7)
18.8 ± 1.7 (54.4)
28.0 ± 0.6 (65.9)
56.4 ± 2.7 (61.2)
23.1 ± 0.0 (76.5)

24.0 ± 0.8 (63.2)
343.5 ± 14.3 (43.0)
120.3 ± 7.1 (36.8)
56.5 ± 3.6 (17.4)
42.2 ± 3.4 (35.9)
29.1 ± 1.0 (12.1)
39.2 ± 3.3 (22.7)
34.4 ± 2.6 (57.0)
130.3 ± 4.3 (19.4)
78.6 ± 7.3 (19.7)
128.1 ± 1.7 (23.2)
94.8 ± 5.2 (31.5)
57.1 ± 3.3 (47.3)
35.0 ± 3.2 (70.9)
44.1 ± 2.6 (22.3)
45.7 ± 1.7 (35.3)
102.8 ± 8.3 (33.7)
24.4 ± 0.4 (26.4)
61.7 ± 4.0 (25.6)
35.1 ± 3.1 (108.1)
188.0 ± 10.0 (40.1)
47.4 ± 1.4 (21.7)
4.3 ± 0.1 (16.0)
21.7 ± 1.6 (25.0)
100.1 ± 6.5 (64.0)
36.4 ± 2.4 (14.7)
57.0 ± 2.6 (13.8)
21.0 ± 0.2 (36.3)
125.7 ± 4.2 (14.2)
94.7 ± 3.3 (9.8)
148.1 ± 11.4 (19.3)
32.4 ± 1.1 (12.8)
81.4 ± 3.1 (13.1)
45.1 ± 2.7 (19.6)
68.7 ± 5.0 (35.3)
111.0 ± 7.5 (13.8)
67.9 ± 7.0 (11.9)
410.5 ± 19.8 (19.0)
53.5 ± 3.5 (10.0)
96.9 ± 3.5 (12.8)
35.6 ± 2.0 (19.6)
131.7 ± 12.6 (8.2)
400.3 ± 35.8 (10.3)
184.3 ± 16.0 (6.8)
24.8 ± 1.2 (67.1)
55.9 ± 1.8 (69.6)
44.4 ± 1.8 (12.1)
93.3 ± 7.4 (75.3)
46.2 ± 1.2 (22.3)
43.6 ± 2.7 (23.7)
19.2 ± 0.7 (19.1)
96.0 ± 5.7 (15.2)
119.2 ± 12.0 (21.2)

Concentrations shown are the average ± range of duplicate analyses with the
percent recovery of the isotopically labelled internal standard given in parentheses.

dard (which was not included in these studies). However,
in our work using stable isotope derivatives for both 4ethylphenol and 4-ethyl-2-methoxyphenol, we found different
recoveries of the corresponding internal standards in a number of samples (Table 1). In general, percent recoveries
of 4-ethyl-2-methoxyphenol-d3 in the present work were
about fourfold lower than 4-ethylphenol-d3 . Thus, normalizing 4-ethyl-2-methoxyphenol concentrations to recovery of a
4-ethylphenol-d3 internal standard may result in estimated levels
significantly lower than where 4-ethyl-2-methoxyphenol concentrations are normalized to the recovery of the more reliable
4-ethyl-2-methoxyphenol-d3 standard.
Our standard additions validation of the 4-ethylphenol-d3 and
4-ethyl-2-methoxyphenol-d3 methods indicate that differences
in recoveries between the native and surrogate analytes, while
being non-zero, are only on the order of 10%, far below the average 350% difference in recoveries between 4-ethylphenol-d3
and 4-ethyl-2-methoxyphenol. We cannot comment quantitatively on the use of entirely non-SIDA based approaches
(see, e.g., ref. [7,10–23]) for determining 4-ethylphenol and
4-ethyl-2-methoxyphenol in wines using such internal standards as unlabelled alkylphenols and straight chain alcohols,
but based on the current study, it is reasonable to suggest
some previous reports on 4-ethylphenol, and especially 4-ethyl2-methoxyphenol, levels in wines may be unreliable. Such
potential uncertainty in the historical 4-ethylphenol and 4ethyl-2-methoxyphenol dataset has implications for the use of
4-ethylphenol:4-ethyl-2-methoxyphenol concentration ratios as
reliable indicators of Brettanomyces contamination as well as in
assessing whether 4-ethylphenol or 4-ethyl-2-methoxyphenol is
the dominant contributor to a Brettanomyces aroma based on
comparisons to their respective odor thresholds. The current
results thus demonstrate the need for individual stable isotope
derivatives to reliably quantitate 4-ethylphenol and 4-ethyl-2methoxyphenol in wines.
Acknowledgments
Thanks to the British Columbia Wine Institute (BCWI),
the Investment Agriculture Foundation of British Columbia
(IAFBC), and the Western Diversification Program (WDP) for
funding of this research, and the support of wineries who generously donated expertise and samples.
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