Studies on the formation of methylglyoxal from dihydroxyacetone in Manuka (Leptospermum scoparium) honey

Article abstract of DOI:10.1016/j.carres.2012.07.025

Dihydroxyacetone (DHA) and methylglyoxal (MGO) are unique carbohydrate metabolites of manuka honey. A method for the reliable quantification of DHA in honey samples was established, based on derivatization with o-phenylenediamine (OPD) and subsequent RP-HPLC with UV detection. The previously unknown reaction product of DHA and OPD was identified as 2-hydroxymethylquinoxaline by spectroscopic means. DHA was exclusively determined in 6 fresh manuka honeys originating directly from the beehive as well as 18 commercial manuka honey samples, ranging from 600 to 2700 mg/kg and 130 to 1600 mg/kg, respectively. The corresponding MGO contents varied from 50 to 250 mg/kg in fresh and 70 to 700 mg/kg in commercial manuka honey samples. A good linear correlation between DHA and MGO values in commercial manuka honeys was observed, resulting in a mean ratio of DHA to MGO of 2:1. In contrast to this, the DHA-to-MGO relation was much higher in fresh manuka honeys but approximated to a ratio of 2:1 while honey ripening. Heating experiments revealed that MGO formation based on thermal treatment as a consequence, for example, of caramelization in honey does not occur. DHA and MGO can serve as suitable unique quality parameter for manuka honey.

Full text of DOI:10.1016/j.carres.2012.07.025

Carbohydrate Research 361 (2012) 7–11
Contents lists available at SciVerse ScienceDirect
Carbohydrate Research
journal homepage: www.elsevier.com/locate/carres
Studies on the formation of methylglyoxal from dihydroxyacetone in Manuka
(
Leptospermum scoparium) honey
⇑
Julia Atrott, Steffi Haberlau, Thomas Henle
Institute of Food Chemistry, Technische Universität Dresden, D-01062 Dresden, Germany
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 22 May 2012
Received in revised form 17 July 2012
Accepted 31 July 2012
Available online 8 August 2012
Dihydroxyacetone (DHA) and methylglyoxal (MGO) are unique carbohydrate metabolites of manuka
honey. A method for the reliable quantification of DHA in honey samples was established, based on deriv-
atization with o-phenylenediamine (OPD) and subsequent RP-HPLC with UV detection. The previously
unknown reaction product of DHA and OPD was identified as 2-hydroxymethylquinoxaline by spectro-
scopic means. DHA was exclusively determined in 6 fresh manuka honeys originating directly from
the beehive as well as 18 commercial manuka honey samples, ranging from 600 to 2700 mg/kg and
Keywords:
130 to 1600 mg/kg, respectively. The corresponding MGO contents varied from 50 to 250 mg/kg in fresh
Manuka honey
Carbohydrate metabolism
Dihydroxyacetone
Methylglyoxal
Quinoxaline
and 70 to 700 mg/kg in commercial manuka honey samples. A good linear correlation between DHA and
MGO values in commercial manuka honeys was observed, resulting in a mean ratio of DHA to MGO of 2:1.
In contrast to this, the DHA-to-MGO relation was much higher in fresh manuka honeys but approximated
to a ratio of 2:1 while honey ripening. Heating experiments revealed that MGO formation based on ther-
mal treatment as a consequence, for example, of caramelization in honey does not occur. DHA and MGO
can serve as suitable unique quality parameter for manuka honey.
Glycation
Ó 2012 Elsevier Ltd. All rights reserved.
1
. Introduction
The presence of 1,2-dicarbonyl compounds such as 3-deoxyglu-
course of caramelization. In a recent report, Adams et al.11 identi-
fied dihydroxyacetone (DHA) as direct precursor for MGO forma-
tion in manuka honeys. In this study, the authors had
determined DHA in nectar and fresh honey derived from manuka
plants and proved that DHA is non-enzymatically transferred to
MGO during honey storage. However, the origin of DHA stills re-
mains enigmatic.
cosone (3-DG), glyoxal (GO), and methylglyoxal (MGO) in honey
was first described in 2004 by Weigel et al.1 as a consequence of
sugar degradation or caramelization. Over a wide range of different
honey types, 3-DG, which is the precursor for 5-hydroxymethyl-
furfural (HMF), could be determined in high amounts, varying be-
tween 80 and 1270 mg/kg and correlating with heating or storage
DHA is a well-known degradation product of carbohydrates and
was detected in caramelized mixtures next to other compounds as
1
–3
12
conditions.
Contents of GO and MGO generally were very low
a result of the Maillard reaction. DHA was also found in naturally
1
3
(
up to 5 mg/kg). However, surprisingly high amounts of MGO (up
aged wine samples. Moreover, its presence was described in rela-
2
14
to 760 mg/kg) were found exclusively in manuka honey. This hon-
ey is derived from the flowers of the manuka tree (Leptospermum
scoparium) in New Zealand. It was clearly demonstrated that the
pronounced antibacterial activity of New Zealand manuka honey
tion to fermentation processes of selected microorganisms. Fur-
thermore, DHA is discussed as food additive, for example to
1
5
16
enhance browning in baked foods, or for food preservation. In
certain cosmetic products, namely self-tanning creams, DHA is
the functional ingredient.
2
17
directly originates from MGO. The amount of MGO correlates with
the antibacterial activity in manuka honey,4 and, therefore, can be
used as a tool for labelling the bioactivity of manuka honey.
Manuka honey was shown to inhibit a wide range of microor-
ganisms, including multiresistant strains.6 Furthermore, wound
dressings containing manuka honey seem to be useful supports
in clinical applications for wound healing.9
,5
Knowledge about the amount of DHA and MGO in honey is of
prime importance for manufacturers as well as for labelling pur-
pose. In the present study, a new method for the reliable quantifi-
cation of DHA via RP-HPLC using pre-column derivatization with o-
phenylenediamine (OPD) is reported. The previously unknown
derivative resulting from DHA and OPD is identified by spectro-
scopic means. DHA and MGO was measured for a number of com-
mercially available and freshly produced manuka honeys in order
to obtain information about the ratio between DHA and MGO
and the transformation of DHA to MGO during storage. Further-
more, the influence of a thermal treatment on MGO content in
–8
,10
The origin and in particular the high concentration of MGO in
manuka honey cannot be explained by sugar degradation in the
⇑
Corresponding author. Fax: +49 351 463 34138.
E-mail address: Thomas.Henle@chemie.tu-dresden.de (T. Henle).
0
008-6215/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved.
http://dx.doi.org/10.1016/j.carres.2012.07.025

8
J. Atrott et al. / Carbohydrate Research 361 (2012) 7–11
manuka honey was studied in order to clarify whether the bioac-
tive compound can be produced artificially in the final product.
ketoamine (V) according to the conversion of Amadori products.
A second dehydration results in formation of an intermediate
(
VI), which is converted into the quinoxaline (VII), putatively by
oxidation. Compared to dicarbonyl compounds like MGO or 3-
DG, complete derivatization of DHA with OPD takes more time
for development of the quinoxaline. Therefore, it can be supposed
that the formation of the keto group in (V) via enaminol (IV) is the
rate determining step of the reaction. The oxidation of intermedi-
ate (VI) to the quinoxaline cannot be proven yet, but it may due
to the oxidative properties of the reaction mixture, for example
the influence of oxygen.
Kinetic studies with varying incubation time, temperature, pH
value, and molar ratio of DHA to OPD were performed in order to
optimize derivatization of DHA to compound (VII) (data not
shown). Finally, it could be shown that best results are obtained
after incubation for 16 h at 37 °C using an acetate buffer with a
pH value of 4.0 for sample preparation. Due to these differences
in methodical conditions, reliable simultaneous quantification of
DHA together with MGO was not possible. At pH value of 4.0,
acid-induced formation of the MGO–quinoxaline was observed in
carbohydrate solutions, and hence, MGO values would be overesti-
mated. Otherwise, reaction of DHA and OPD is too slow at neutral
pH, and therefore, DHA does not react completely with OPD under
these conditions. Consequently, to reach optimal quinoxaline for-
mation and best sensitivity for both analytes, two independent
sample preparations and two chromatographic runs per sample
are necessary.
2
2
. Results and discussion
.1. RP-HPLC analysis of DHA and characterization of DHA
quinoxaline
The analysis of 1,2-dicarbonyl compounds such as 3-desoxy-
glucosone (3-DG) or methylglyoxal (MGO) in honey is generally
achieved by RP-HPLC of the corresponding quinoxalines resulting
from derivatization with o-phenylenediamine (OPD). Under condi-
tions applied for optimal derivatization of these compounds (phos-
phate buffer pH 6.5, incubation at room temperature),
dihydroxyacetone (DHA) also formed an OPD-derivative eluting
in the chromatogram at about 13 min (Fig. 1A). This reaction prod-
uct was isolated using semi preparative RP-HPLC. The molar mass
of the isolate was determined with 160 g/mol by LC-ESI-TOF-MS.
The UV maximum at 319 nm was observed, varying only slightly
when compared to the absorption maximum of 312 nm generally
reported for quinoxalines. Based on the NMR data, the structure
was identified as 2-hydroxymethylquinoxaline (2-quinoxaline-
methanol, compound VII in Fig. 2).
2
-Hydroxymethylquinoxaline has already been described as
reaction product formed during incubation of sugars like glucose
or ribose, respectively, with OPD.1
8,19
The identified quinoxalines
derived from this reaction mixture were explained by the buffer-
induced degradation of hexoses to 1,2-dicarbonyl fragments.
However, the authors did not propose DHA being involved in quin-
oxaline formation. Up to now, it was generally accepted that DHA
does not react with OPD to a stable derivative, and, therefore, can-
not be quantified by means of RP-HPLC following derivatization
1
9
Performing DHA determination under these conditions, it be-
came clear that notable amounts of DHA are also formed from
the honey matrix itself, namely from glucose and fructose break-
down, during derivatization (Fig. 1B and C). This reproducible basis
level of DHA has to be taken into account by using an artificial hon-
2
1
2
0
ey matrix as blank value. Calibration of (VII) using this sugar ma-
with OPD.
trix as 10% solution in acetate buffer showed a good linearity
A possible reaction mechanism for the formation of 2-hydrox-
ymethylquinoxaline from DHA and OPD is shown in Figure 2. First,
a nucleophilic addition of one amino group of OPD (I) to the car-
bonyl carbon of DHA (II) occurs. The iminol (III) is formed by dehy-
dration and is able to convert via the enaminol (IV) into the
2
(
regression equation y = 0.15x + 5.66, R = 0.9944) in the range
from 0.18 to 2.20 mmol/L DHA in sample solution, equivalent to
DHA concentrations between 165 and 1980 mg/kg in honey. Quan-
tification of DHA in manuka honey (Fig. 1D) was finally performed
Figure 1. RP-HPLC after derivatization with OPD of (A) DHA standard solution in water, (B) solution of fructose and glucose simulating honey matrix, (C) rape honey, (D)
manuka honey.

J. Atrott et al. / Carbohydrate Research 361 (2012) 7–11
9
Figure 2. Proposed reaction mechanism for the reaction of DHA (II) with OPD (I) to 2-hydroxymethylquinoxaline (VII).
by matrix calibration using the above mentioned artificial honey
matrix. LOD (limit of detection) and LOQ (limit of quantification)
of DHA in test solutions were estimated with 0.01 and
0
.12 mmol/L, respectively, representing a LOD of 10 mg/kg and
LOQ of 110 mg/kg in honey while using 10% sample solution.
Recovery of DHA added to the artificial honey matrix was deter-
mined with a mean of 95.3%. The relative repeatability for DHA
quantification was assessed by repetitive analysis of one manuka
honey sample and was at 7.8%.
2
.2. Quantification of DHA and MGO in manuka honey
Within this work, 18 commercial as well as 6 fresh manuka
honey samples obtained directly from the beehive without further
processing were analysed for their contents of DHA and MGO. The
results are shown in Figure 3. In the fresh manuka honeys, DHA
varied from 611 to 2724 mg/kg (median 1622 mg/kg). These fresh
honey samples contained MGO in concentrations from 50 to
2
60 mg/kg (median 129 mg/kg), whereas in commercial manuka
honeys, MGO contents were ranging from 66 to 684 mg/kg with
a median value of 356 mg/kg, confirming the data in the litera-
2
,5
ture. DHA in commercial samples varied from 127 to 1563 mg/
kg (median 735 mg/kg).
A good linear correlation (regression equation y = 0.47x + 11.57,
R = 0.8977) between the corresponding concentration of DHA and
MGO in commercial manuka honeys was observed (Fig. 4, black
squares), resulting in a mean ratio of DHA to MGO of 2:1. Com-
pared to this, concentrations of DHA exceeded the values of MGO
in fresh manuka honeys seven to sixteenfold (Fig. 4, open squares).
Figure 4. Relation of DHA to MGO in 18 commercial manuka honeys (dark squares),
compared with 6 fresh manuka honeys before (opened squares) and after storage at
37 °C for 6 weeks (grey triangles).
2
During storage of these fresh samples at 37 °C for 6 weeks, the
amount of DHA decreased considerably due to formation of MGO
(Fig. 4, grey triangles). For stored fresh manuka honeys, a ratio of
DHA to MGO ranging from 5 to 9 was calculated. A longer honey
storage would be followed by a further increase of MGO content till
there is reached a plateau which can be also observed for data re-
1
1
ported by Adams et al. This conversion can be additionally char-
acterized by the ratio of DHA to MGO which is approximating
factor 2 in the course of honey ripening. Consequently, the discrim-
ination between fresh and ‘ripened’ manuka honeys on the basis of
defined DHA-to-MGO ratios seems to be a suitable tool for classifi-
cation of manuka honeys and for monitoring reactions occurring
during ripening. The simultaneous presence of DHA and MGO in
certain relation to each other is a unique quality parameter for
manuka honey.
The findings of this study are in agreement with the results of
1
1
Adams et al. These authors identified DHA as precursor for
MGO formation by illustrating curves of DHA decrease and MGO
increase while honey storage. Authors observed a similar relation
of DHA to MGO after honey incubation. Adams et al.,11 however,
had determined higher values of DHA and MGO in fresh manuka
honeys, which can be due to the quantification method used by
these authors.
Other floral or honeydew honeys which were analysed in this
study did not contain any DHA above LOD (Fig. 1C). The peak
Figure 3. Concentrations of DHA and MGO in 6 fresh and 18 commercial manuka
honeys (box plot, Whiskers indicate min/max values, ꢀ = median value).

1
0
J. Atrott et al. / Carbohydrate Research 361 (2012) 7–11
detectable for DHA in Figure 1C is due to the small amount of DHA
formed from the honey matrix during derivatization (see above).
Confirming the data of Weigel et al.,1 only small amounts of
MGO ranging from 2 to 28 mg/kg were detected in these honeys.
In contrast, other honeys from Leptospermum species sources like
jellybush honey from Australia (Leptospermum polygalifolium) can
also contain MGO and DHA in high amounts.20 Two samples of this
honey type analysed in this study revealed an analogous DHA-to-
MGO ratio of 2:1. By evaluation of the data of Windsor et al. in
the same way, it can be suggested that a similar conversion oc-
curred in these honeys which can be illustrated by calculated
DHA-to-MGO relation ranging from 18.5 till 0.7. However, these
authors analysed partially very high values of MGO up to
not possible. In general, treatment of honey with high tempera-
tures is followed by massive changes in sensory properties like col-
our, smell, and taste, respectively. Moreover, a remarkable increase
of 3-DG, resulting from degradation of glucose and fructose, and, in
a consequence, of 5-hydroxymethylfurfural (HMF) will occur due
to the heating processes, making such honeys inacceptable.
2
.4. Conclusion
2
0
Reliable analysis of DHA and MGO in honey was achieved by
pre-column derivatization with OPD, followed by RP-HPLC with
UV-detection. DHA is transformed to MGO during ripening of man-
uka honey, resulting in a constant DHA-to-MGO ratio of 2:1, which
can be used as a quality index to monitor changes during storage
and to classify commercial products. DHA and MGO are unique
and naturally occurring constituents of manuka honey.
1
723 mg/kg by doing a simultaneous determination of DHA and
MGO, raising doubts concerning a possible overestimation of
MGO as a result of ‘neoformation’ during sample preparation,
which cannot be dispelled, as several methodical details are miss-
2
0
ing in this paper.
3. Materials and methods
3.1. Chemicals
The phenomenon that DHA is not completely transferred to
MGO during honey storage and that a constant ratio between
DHA and MGO is obtained, might be due to the water content of
honey being the limiting factor. MGO formation occurs by dehy-
dration of DHA. A back reaction presumably does not occur since
this transformation of DHA to MGO was described to be irrevers-
Methylglyoxal (MGO, 40% in water) and dihydroxyacetone
(DHA) were obtained from Sigma–Aldrich (Steinheim, Germany)
and the derivatizing agent o-phenylenediamine (OPD) from Fluka
(Munich, Germany). Sodium acetate, sodium dihydrogenphosphate
and disodium hydrogenphosphate were purchased from Grüssing
(Filsum, Germany). Methanol (HPLC grade) and acetic acid were or-
dered from VWR Prolabo (Leuven, Belgium). Water used for buffer
preparations and HPLC solvents was obtained using a Purelab plus
purification system (USFilter, Ransbach-Baumbach, Germany).
2
2
ible. The water content of 10 commercial manuka honeys was
determined varied between 16.0% and 19.3% with a median value
of 18.1%. This is in agreement with legislative regulations, which
give very narrow limits for an acceptable water content in honey
2
3
(
maximum 20%).
2
.3. Influence of heat treatment on MGO levels in manuka
honey
3.2. Honey samples
Honey is not allowed to be heated.23 However, speculations
Manuka honeys were kindly provided by Manuka Health Ltd, Te
Awamutu, New Zealand. In total, 18 commercial samples with
varying methylglyoxal content and 6 samples of fresh manuka
honey from different regions originating directly from the beehives
without any further treatment were obtained. Other honey sam-
ples (rape, acacia, sunflower, bush flower, heather, chestnut, leath-
erwood, thyme, pine, and forest, in total 17 samples) were
obtained from local supermarkets, or from Neuseelandhaus, Bergk-
amen, Germany (2 samples of jellybush honey), respectively.
Water content of honey samples was measured refractrometrically
concerning a fraudulent heat-treatment to increase the MGO con-
tent in order to obtain manuka honey with ‘optimized’ bioactivity,
are conceivable. MGO may also increase while heating due to a po-
tential release of ‘sugar-bound’ MGO.24
To study the effect of thermal treatment on MGO content, two
commercial samples of manuka honey with different MGO con-
tents and one sample of a rape honey were incubated at 70 °C for
1
0 min (resembling conditions of pasteurization) and up to 24 h
for long-term heating. The results of these analyses, given in Ta-
ble 1, show unambiguously that heat treatment of commercial
rape honey did not lead to MGO formation, whereas the concentra-
tion of MGO in manuka honey remained constant while short heat-
ing and decreased markedly during long-term incubation at 70 °C.
Moreover, the long-term storage of honey for 12 weeks at 37 °C re-
sulted in a decrease of MGO in manuka honey, which was more
pronounced in the sample which had been pre-heated for 10 min
at 70 °C. These results clearly show that a thermal generation of
MGO as a consequence of caramelization of sugars in honey is
1
according to Weigel et al. Additionally, an artificial honey matrix
consisting of 46.5% fructose, 34.0% glucose, 1.5% sucrose, and 18.0%
2
1
water was used.
3.3. Honey storage and heat treatment
To simulate a long-term storage, the 6 fresh manuka honeys
were incubated at 37 °C for 6 weeks. To investigate the effect of
thermal treatment, one rape honey and two manuka honeys were
heated at 70 °C for 10 min as well as for 8 h and 24 h. Furthermore,
samples heated for 10 min at 70 °C were incubated afterwards at
3
7 °C for 12 weeks to assess the combination of heating and stor-
Table 1
age of honey compared to a non-heated sample.
Concentration of MGO after thermal treatment of manuka and rape honeys at 70 °C,
partially followed by storage at 37 °C for 12 weeks
3
.4. Sample preparation
Incubation conditions
MGO concentration (mg/kg)
The determination of MGO was performed according to Mavric
et al. with slight modifications. For this, 1 mL aliquots of 10% (w/v)
honey solutions in 0.5 M phosphate buffer (pH 6.5) were mixed
7
0 °C
37 °C
Manuka 1
Manuka 2
Rape
2
0
0
1
1
8
2
min
min
0 min
0 min
h
—
100.7
90.6
107.9
67.4
68.7
35.9
307.2
268.1
329.1
241.8
237.8
132.8
2.3
5.3
3.1
5.5
3.8
3.9
+12 weeks
—
+12 weeks
—
—
with 300
1% in phosphate buffer pH 6.5). Samples were incubated at room
temperature overnight and membrane filtered (0.45 m). For anal-
ysis of DHA, 1 mL aliquots of 10% honey solutions in 0.5 M acetate
lL phosphate buffer (pH 6.5) and 300 lL OPD solution
(
l
4 h

J. Atrott et al. / Carbohydrate Research 361 (2012) 7–11
11
buffer (pH 4.0) were mixed with 300
00 L OPD solution (1% in acetate buffer pH 4.0), followed by
incubation for 16 h at 37 °C and membrane filtration (0.45 m).
l
L acetate buffer (pH 4.0) and
ESI-TOF mass spectrometer (PerSeptive Biosystem, Framingham,
USA). The chromatographic conditions were the same as described
for analytical HPLC, 50 lL sample solution was injected. Electro-
3
l
l
spray ionization was used in the positive ionization mode. Full scan
mass spectra were measured in mass range 100–1000 m/z in the
tic-mode. For data acquisition the software Data Explorer Version
4.0.0.1 (Applied Biosystems, Foster City, USA) was used.
H NMR spectrum was recorded on a Bruker DRX 500 instru-
ment (Rheinstetten, Germany) at 500 MHz. For this, 5 mg of the
3
.5. Analytical RP-HPLC
HPLC analyses were performed using an Äkta basic system from
Amersham Pharmacia Biotech (Uppsala, Sweden) with a pump P-
00 and an online degasser K-5004 (Knauer, Berlin, Germany) as
1
9
well as an UV detector UV-900 and an auto sampler A-900. Peak
evaluation was managed using the software UNICORN 4.11. The
separation of quinoxalines was realized on a stainless steel column
purified and lyophilized isolate was dissolved in 750 lL deuterium
oxide. All chemical shifts are given in parts per million (ppm) rel-
ative to the internal HOD signal (4.70 ppm).
filled with Eurospher 100 RP18 material (250 mm ꢀ 4.6 mm, 5
lm
Data of the isolated quinoxaline of DHA were as follows. 1
H
particle size, integrated pre column; Knauer, Berlin, Germany). The
mobile phase were 0.075% acetic acid in water (solvent A) and a
mixture of 80% methanol and 20% solvent A (solvent B). The gradi-
ent started with 40% solvent B for 1 min and then was elevated lin-
early to 100% B over a period of 20 min, was changed back to 40% B
in 4 min and was held there for 7 min. The flow rate was 0.8 mL/
2
NMR (500 MHz, D O), d [ppm]: 8.74 (s, H-3), 7.81–7.87 (m, H-5,
0
0
H-8), 7.68–7.73 (m, H-6, H-7), 4.82 (s, H-1 A, H-1 B). Mass spectros-
+
copy gave a m/z of 161 for [MH] . UV spectroscopy gave a kmax of
319 nm.
Acknowledgements
min, the separation was done at 30 °C, 20 lL sample solution
was injected and peaks were detected by measurement of UV
absorbance at 312 nm. Quantification was achieved by external
calibration with standard solution for MGO or by matrix calibra-
tion using an artificial honey matrix according to Wahdan for
DHA, respectively.
The authors thank Dr. Uwe Schwarzenbolz, Institute of Food
Chemistry, for his help during the LC-ESI-TOF-MS measurements,
and furthermore, the members of the Institute of Organic Chemis-
try, namely Dr. Margit Gruner and Anett Rudolph, for recording the
NMR data.
2
1
The limits of detection (LOD) and quantification (LOQ) of DHA
were calculated from blank values using artificial honey matrix
References
2
5
(
measurement of 10 independent blank values on one day). For
1
2
.
.
Weigel, K. U.; Opitz, T.; Henle, T. Eur. Food Res. Technol. 2004, 218, 147–151.
Mavric, E.; Wittmann, S.; Barth, G.; Henle, T. Mol. Nutr. Food Res. 2008, 52, 483–
determination of repeatability, 10 samples of a manuka honey
were derivatized on different days. The recovery of DHA was esti-
mated by spiking 10% solutions of artificial honey with varying
concentrations of DHA and analysing as described above.
489.
3. Marceau, E.; Yaylayan, V. A. J. Agric. Food Chem. 2009, 57, 10837–10844.
4
.
.
Adams, C. J.; Boult, C. H.; Deadman, B. J.; Farr, J. M.; Grainger, M. N.; Manley-
Harris, M.; Snow, M. J. Carbohydr. Res. 2008, 343, 651–659.
Atrott, J.; Henle, T. Czech. J. Food Sci. 2009, 27, 163–165.
5
3
.6. Isolation and identification of quinoxaline resulting from
6. Cooper, R. A.; Molan, P. C.; Harding, K. G. J. Appl. Microbial. 2002, 93(5), 857–
63.
8
DHA and OPD
7.
Molan, P. Honey: Antimicrobial Actions and Role in Disease Management. In
New Strategies combating bacterial infection, Ahmad, I., Aqil, F., Eds.; Wiley-VCH:
Weinheim, Germany, 2008; pp 229–253.
Semipreparative HPLC was performed with a system consisting
of two pumps K-1001 with mixing chamber, automatically driven
valves for injection and fractionation, online degasser and UV
detector K-2501 (all from Knauer, Berlin, Germany). Data handling
was managed using the software Eurochrom 2000. The separation
was realized on a stainless steel column filled with Eurospher 100-
8. George, N. M.; Cutting, K. F. Wounds 2010, 19, 231–236.
Molan, P. C. Int. J. Low. Extrem. Wounds 2006, 5, 40–54.
0. Simon, A.; Traynor, K.; Santos, K.; Blaser, G.; Bode, U.; Molan, P. eCAM 2009, 6,
65–173.
11. Adams, C. J.; Manley-Harris, M.; Molan, P. C. Carbohydr. Res. 2009, 344, 1050–
053.
2. Severin, T.; Hiebl, J.; Popp-Ginsbach, H. Z. Lebensm. Unters. Forsch. 1984, 178,
84–287.
13. Laurie, V. F.; Waterhouse, A. L. J. Agric. Food Chem. 2006, 54, 4668–4673.
9.
1
1
1
1
8
C18 (33 mm ꢀ 8 mm with integrated pre column; Knauer, Berlin,
2
Germany). A DHA standard solution in water (2500 mg/L) after
derivatization with OPD (1% in 0.5 M acetate buffer pH 4) for
1
1
1
4. Green, S. R. U.S. 2948658, 1960 Aug 09.
5. Kirk, D. L. U.S. 3220850, 1965 Nov 30.
6. Oborsh, E. V.; Barkate, J. A.; Wesu, C.; Owen, T. M. CA 1054434 A1, 1979 May
15.
1
6 h at 37 °C was used for fractionation. The derivative of DHA
was separated with same solvents as mentioned above for analyt-
ical separations. The gradient started with 40% B for 1.2 min, ele-
vated on 86% B in 14.2 min, on 95% in further 3 min, and on 100%
in 1.8 min, then gradient changed back on 40% B in 3 min and
was held there for further 7.8 min. The flow rate was set to
1
1
7. Monfrecola, G.; Prizio, E. Compr. Ser. Photosci. 2001, 3, 489–493.
8. Takagi, M.; Mizutani, M.; Tsuchiva, K. Bull. Univ. Osaka Prefecture, Ser. B: Agric.
Life Sci. 1972, 24, 43–48.
19. Rizzi, G. P. J. Agric. Food Chem. 2004, 52, 953–957.
20. Windsor, S.; Pappalardo, M.; Brooks, P.; Williams, S.; Manley-Harris, M. J.
Pharmacog. Phytother. 2012, 4, 6–11.
1
1
.6 mL/min. Injection volume was 2 mL. The eluate collected from
6.4 to 19.4 min was evaporated to dryness, taken up in 2 mL
21. Wahdan, H. A. L. Infection 1998, 26, 30–35.
22. Fedoronko, M.; Koenigstein, J. Collect. Czech. Chem. Commun. 1969, 34, 3881–
water and lyophilized.
3894.
2
2
3. Codex Alimentarius Commission: Revised Codex Standard for honey, Codex
STAN 12-1981, Rev.1 (1987), Rev.2 (2001).
4. Stephens, J. C.; Schlothauer, R. C. WO 2010082845 A1, 2010 Jul 22.
Absorption spectrum of the isolate in water was recorded by a
Specord S100 diode array spectrophotometer (Carl Zeiss, Jena, Ger-
many). LC–MS measurement was realized by using a LC system
25. Mocak, J.; Bond, A. M.; Mitchell, S.; Scollary, G. Pure Appl. Chem. 1997, 69, 297–
28.
3
1
100 Series (Agilent Technologies, PaloAlto, USA) with Mariner