Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus

Article abstract of DOI:10.1016/j.phytochem.2014.12.013

This study describes the identification of regioisomers and enantiomers of triacylglycerols of C20 polyunsaturated fatty acids (PUFAs) in the alga Trachydiscus minutus cultivated at different temperatures using reversed- and chiral-phase liquid chromatogr

Full text of DOI:10.1016/j.phytochem.2014.12.013

Phytochemistry xxx (2015) xxx–xxx
Contents lists available at ScienceDirect
Phytochemistry
journal homepage: www.elsevier.com/locate/phytochem
Temperature dependence of production of structured triacylglycerols
in the alga Trachydiscus minutus
Tomáš Rezanka a, , Jaromír Lukavsk y´ , Karel Sigler , Linda Nedbalová , Milada Vítová
⇑
b
a
c
d
ˇ
a
Institute of Microbiology, Academy of Sciences of the Czech Republic, Víde nˇ ská 1083, 142 20 Prague, Czech Republic
Institute of Botany, Academy of Sciences of the Czech Republic, Biorefinery Res. Centre of Competence, Dukelská 135, 379 82 T rˇ ebo nˇ , Czech Republic
Department of Ecology, Faculty of Science, Charles University in Prague, Vini cˇ ná 7, 128 44 Prague 2, Czech Republic
b
c
d
Institute of Microbiology, Academy of Sciences of the Czech Republic, Laboratory of Cell Cycles of Algae, Centre Algatech, Opatovick y´ ml y´ n 237, 379 81 T rˇ ebo nˇ , Czech Republic
a r t i c l e i n f o
a b s t r a c t
Article history:
This study describes the identification of regioisomers and enantiomers of triacylglycerols of C20 polyun-
saturated fatty acids (PUFAs) in the alga Trachydiscus minutus cultivated at different temperatures using
reversed- and chiral-phase liquid chromatography–mass spectrometry. The use of the two different
phases contributes to ready identification, both qualitative and semiquantitative, of regioisomers and
enantiomers of triacylglycerols containing eicosapentaenoic and arachidonic in the molecule. The ratio
of regioisomers and enantiomers of triacylglycerols (TAG) depends on the temperature of cultivation;
with lowering temperature the proportion of the achiral TAG increases and the enantiomer ratio diverges
from 1:1.
Received 21 October 2014
Received in revised form 10 December 2014
Available online xxxx
Keywords:
Alga
Trachydiscus minutus
Eustigmatophyceae
Temperature
Ó 2014 Elsevier Ltd. All rights reserved.
Structured triacylglycerols
Chiral-LC/APCI-MS
1
. Introduction
Unicellular alga Trachydiscus minutus (Bourr.) Ettl, strain
explored over the past few decades (Bajpai and Bajpai, 1993; Cao
et al., 2012). Many species have the ability to produce substantial
amounts (e.g. 20–50% dry cell weight) of TAGs as a storage lipid
(Hu et al., 2008). One-fifth of algal strains in the Culture Collection
of Algae at Goettingen University (SAG) contain EPA. Most of these
strains belong to classes Chlorophyceae, Trebouxiophyceae and
Rhodophyceae.
The highest proportions of EPA and arachidonic acid (ARA)
(more than 1/3 of total fatty acids) were found in algae of the gen-
era Cryptomonas, Heterococcus, Nannochloropsis and/or Porphyridi-
um (Lang et al., 2011).
Lukavsk y´ et P rˇ ibyl 2005/1 (former Pseudostaurastrum minutum)
belongs to Eustigmatophyceae as described by Pribyl et al.
(
2012). It was isolated from cooling pools of the Temelín nuclear
power plant (Czech Republic). It contains chlorophyll a, violaxan-
thin, vaucheriaxanthin ester, and b-carotene. No other chlorophylls
were determined. It has a broad temperature adaptability with an
optimum growth temperature at 25 °C (Gigova et al., 2012). The
life cycle of T. minutus includes darkness induced zoosporogenesis
(
Pribyl et al., 2012).
T. minutus is an interesting organism from the biochemical
The effect of temperature on the composition of phospholipid
membranes and content of FAs in algae has also been intensively
studied (Morgan-Kiss et al., 2006). The lipid composition of the
green alga Botryococcus was studied at three different cultivation
temperatures: suboptimal (18 °C), optimal (25 °C), and supraopti-
mal (32 °C). The content of polyenoic fatty acids was found to
decrease with increasing temperature, e.g. with 18:3nꢀ3 to less
than half (Kalacheva et al., 2002). Similar results were found also
in Chlorella vulgaris and Botryococcus braunii, where culturing was
performed at 18 °C, 25 °C, and 32 °C. The content of linolenic acid
also decreased when the cells were cultured at 32 °C compared
to culturing at 18 °C while the amount of 18:2nꢀ6 was almost dou-
bled (from 14.3% to 28.1% of total FAs) (Sushchik et al., 2003). The
effect of temperature on EPA production was studied in the
unicellular red algae Porphyridium purpureum (Nuutila et al.,
point of view, because it contains high levels of polyunsaturated
fatty acids (PUFAs), especially eicosapentaenoic acid (EPA)
(
Iliev et al., 2010; Rezanka et al., 2010; Lukavsk y´ et al., 2010).
Currently it is being extensively studied for biotechnological
purposes. Cultivation conditions play a key role in biomass
production and PUFA composition (Alexandrov et al., 2014;
Cepák et al., 2014).
The use of algae as an alternative and renewable source of lipid-
rich biomass feedstock for oil production has been intensively
⇑
Corresponding author. Tel.: +420 241 062 300; fax: +420 241 062 347.
E-mail address: rezanka@biomed.cas.cz (T. Rˇ ezanka).
http://dx.doi.org/10.1016/j.phytochem.2014.12.013
031-9422/Ó 2014 Elsevier Ltd. All rights reserved.
0
ˇ
Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013

ˇ
T. Rezanka et al. / Phytochemistry xxx (2015) xxx–xxx
2
1
997) and Porphyridium cruentum (Akimoto et al., 1998). In. P.
two different phases contributes to ready identification, both qual-
itative and semiquantitative, of regioisomers and enantiomers of
triacylglycerols containing EPA and ARA in the molecule. Our
atmospheric pressure chemical ionization mass spectrometry
(APCI-MS) method is well suited for direct determination of TAG
profiles and for comparing the production of lipids after a change
in microorganism cultivation conditions.
cruentum, ARA content was found to increase with increasing tem-
perature while EPA content decreased (from 13.5% at 20 °C to 3.1%
at 32 °C). Another study (Rezanka et al., 1987) of the effect of tem-
perature on the content of PUFAs and specific growth rate of P.
cruentum showed that the EPA content decreased with increasing
temperature whereas the ARA content showed only an insignifi-
cant change. Cultivation of T. minutus at low (15 °C) and high
(
40 °C) temperatures was found to increase the lipid content (up
2
. Results and discussion
to 35.2% dry weight) relative to the cultivation at 25 °C, which is
optimal for growth (Gigova et al., 2012).
2.1. Identification and production of fatty acids
Structured triacylglycerols (STAGs) can be defined as triacylgly-
cerols (TAGs) that have been modified or restructured from natural
oils and fats. The STAGs are synthesized for nutritional use or spe-
cific purposes, such as human milk-fat substitutes, low-caloric fats
and oils enriched usually by essential fatty acids (FAs) (e.g. linoleic
Methyl esters were identified based on their compliance with
the retention characteristics of commercially obtained standards
by GC–MS and comparison with previously obtained results
(Rezanka et al., 2010). The fatty acids were those listed in Table 1.
and
(
a-linolenic acid) or long-chain x-3 polyunsaturated fatty acids
PUFAs) (EPA and ARA). During the last 25 years, a range of animal
The major acids in control cultivation as well as in the two temper-
ature variations were myristic (M), palmitic (P), palmitoleic (Po),
linoleic (L), a-linolenic (Ln), arachidonic acid (A) and eicosapenta-
and human studies have been conducted using STAGs; they consti-
tute the background for many of the clinical studies performed
subsequently (Akoh, 2006).
enoic (E) acids. It is generally accepted that the content of polyun-
saturated fatty acids (PUFAs) decreases with increasing
temperature while the amount of saturated or monoenoic FAs
increases (Guschina and Harwood, 2006, 2009). Our results
describing growth/biomass production (DW g/L), as well as the
total fatty acid content (% DW) and the volumetric content (mg/L
of culture) for EPA and ARA are presented in Table 1.
We chose several representative articles to compare the influ-
ence of culture temperature on the fatty acid content of the algae.
When culturing T. minutus in the temperature range 15–40 °C, it
was found that the production of total lipids is lowest at the opti-
mum temperature, i. e. 25 °C. When culturing T. minutus at both
limit temperatures (15 and 40 °C) the lipid production was higher,
at 15 °C by nearly two thirds (it increased from 21.8% to 35.2%)
STAGs, i.e. modified or restructured TAGs from natural oils and
fats, irrespective of whether they are synthesized or biosynthe-
sized in vivo or in vitro’’ (Dey and Maiti, 2013; Dey et al., 2014)
can be produced by different methods. Chemical interesterification
with a catalyst like sodium methoxide results in TAGs with random
distribution of the fatty acids within the glycerol backbone. This
randomization distributes the fatty acids equally in all three posi-
tions of the TAG. Another possibility is a stereospecific organic syn-
thesis which allows the stereospecific esterification of positions sn-
1
, sn-2 and sn-3 of glycerol (Kristinsson et al., 2014).
STAGs can be also produced by enzymatic interesterification
using sn-1,3 specific lipases, which results in specific replacement
of the fatty acids in the sn-1,3 positions of the TAG. The outcome
of this process depends primarily on the substrates and the ratio
between them, the choice of enzyme, and the process technology
(Gigova et al., 2012). Conversely, the biomass production was high-
est at the optimum temperature, i.e. 5 times higher than at 15 °C
and more than 4 times higher than at 40 °C. The same trend in bio-
mass production was also described by Cepák et al. (2014),
wherein the yield of biomass production was over 10 g/L at 28 °C
and only 2 g/L and 1 g/L, respectively, at 20 and 33 °C. Similar con-
clusions were reached earlier, e.g. by Sukenik (1991) in the genus
Nannochloropsis, who used the lowest cultivation temperature of
(Akoh, 2006). A promising method of preparation and even produc-
tion of STAGs is the use of different conditions during the cultivation
of microorganisms, either algae or yeast (Rezanka et al., 2012, 2013).
Fats and oils having similar fatty acid compositions do not nec-
essarily have the same fatty acid distributions in their TAGs. For
instance, EPA and ARA are mainly located in the sn-1,3-positions
of TAGs from marine mammals whereas they are enriched in the
sn-2-position of TAGs from fish oil (Nagai et al., 2013). Evidence
is accumulating that both the overall fatty acid profile and the
intramolecular structure of dietary fats are of importance when
considering the nutritional effects of a given fat. Studies on the
effects of dietary fatty acids as related to their position on TAGs
have been published (Mu and Porsgaard, 2005).
2
5 °C to produce the highest EPA levels. As already mentioned by
Cepák et al. (2014), the highest proportion of EPA was observed
at low culture temperature (20 °C), but the production in g/L/day
was negligible, mainly due to low biomass production. Similar con-
clusions were reached in the analysis of the effect of temperature
Table 1
Another important factor that affects the digestion and absorp-
tion of oils and fats is their physical state, i.e. the presence of a solid
TAG phase. This phenomenon has an effect on the digestion,
absorption and metabolism of dietary lipids. It limits the enzy-
matic hydrolysis of TAG and then their absorption (Michalski
et al., 2013).
In contrast to the temperature dependence of FAs composition,
similar studies concerning lipids are much fewer (Chen et al., 2008;
Olofsson et al., 2012) and, to our mind, the effect of temperature on
the ratio of regioisomers and enantiomers of triacylglycerols has
not yet been studied at all, unlike studies describing commonly
available animal and plant oils (e.g. Lisa et al., 2009; Gotoh et al.,
Fatty acid content (%) of triacylglycerols from T. minutus cultivated at different
temperatures.
FA
10 °C
25 °C
0.2
24.8
10.4
10.9
0.4
32 °C
La
M
P
Po
S
Lauric
0.1
12.3
8.5
6.3
0.2
0.3
29.3
13.1
9.8
Myristic
Palmitic
Palmitoleic
Stearic
0.8
O
L
Oleic
Linoleic
0.3
2.9
0.5
5.3
0.9
6.9
Ln
A
E
a
-Linolenic
6.3
7.2
55.7
0.2
5.8
4.1
37.0
0.6
4.9
1.6
31.5
0.9
Arachidonic
Eicosapentaenoic
Behenic
B
2
011; Lisa and Holcapek, 2013).
Dry weight (g/L)
Total FAs (% dw)
EPA (mg/L)
1.8
36.1
361.9
46.8
9.6
33.2
1179.3
130.7
4.7
25.8
382.0
19.4
This study describes the identification of regioisomers and
enantiomers of triacylglycerols of C20 PUFAs in the alga T. minutus
cultivated at different temperatures using reversed- and chiral-
phase liquid chromatography–mass spectrometry. The use of the
ARA (mg/L)
ˇ
Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013

ˇ
3
T. Rezanka et al. / Phytochemistry xxx (2015) xxx–xxx
on other, more or less related algae. For example the linolenic acid
Table 2
Relative quantities (%), retention time (RT, min), equivalent carbon number (ECN) and
acyl carbon number (ACN) of molecular species of TAGs identified in alga T. minutus.
content in green algae of the genus Chlorella and Botryococcus
decreases to half when culture temperature rises from 18 to
3
2 °C (Sushchik et al., 2003). Similarly, the hexadecatrienoic acid
RT
TAG
ECN
ACN
10 °C
25 °C
32 °C
content in Chlorella decreased from 11.4% to 3.2%, i.e. almost 4
times, on temperature change from 13 to 38 °C, whereas the con-
tent of myristic acid was essentially unchanged, 2.8 versus 3.1%,
upon a temperature change from 4 to 30 °C (Teoh et al., 2013).
Analogously, the hexadecatetraenoic acid content in the genus
Chlamydomonas decreased from 16.5 to 3.2% for a temperature
change from 13 to 33 °C and the stearic acid content increased
from 8.6% to 24.9%. The spectrum of fatty acids was studied in
the unicellular red alga P. cruentum grown at 21, 24 and 28 °C.
The proportion of unsaturated fatty acids (L, A and E) increased
with decreasing cultivation temperature while the production of
saturated acids (P) declined (Rezanka et al., 1987).
Table 1 shows the fatty acid content of the alga T. minutus cul-
tivated in our experiments. The differences in percent proportion
between our results and those of an important study published
by Cepák et al. (2014) are mainly caused by two distinct basic
experimental conditions – different culture temperatures (10, 25
and 32 °C versus 20, 28 and 33 °C), and the fact that Cepák et al.
21.9
25.8
EEE
LnEE
AEE
PoEE
MEE
AELn
AAE
30
32
32
34
34
34
34
34
34
34
36
36
36
36
36
36
36
36
36
36
36
36
38
38
38
38
38
38
38
38
38
38
38
38
38
38
38
38
38
40
40
40
40
40
40
40
40
40
40
40
40
40
40
40
40
40
40
42
42
42
42
42
42
42
42
42
42
42
42
42
42
42
42
60:15
58:13
60:14
56:11
54:10
58:12
60:13
58:12
56:11
56:11
60:12
58:11
56:10
58:11
56:10
56:10
56:10
56:10
54:9
54:9
54:9
52:8
58:10
56:9
56:9
56:9
56:9
54:8
54:7
54:8
54:8
54:8
52:7
52:7
52:7
52:7
50:6
50:6
48:5
56:8
56:8
54:7
54:7
54:7
54:7
52:6
52:6
52:6
52:6
52:6
50:5
50:5
50:5
50:5
48:4
48:4
46:3
52:5
54:6
50:4
52:5
54:6
52:5
52:5
52:5
50:4
50:4
50:4
48:3
48:3
46:2
48:3
44:1
3.9
2.2
2.2
2.3
2.7
0.6
0.6
2.3
2.3
1.3
1.2
1.6
2.4
1.4
1.4
0.7
1.4
2.3
0.7
1.4
1.8
1.8
0.7
0.6
0.8
1.5
1.4
0.6
1.5
0.9
1.1
1.4
0.7
1.5
1.2
1.9
1.2
1.9
2.0
0.5
0.6
0.5
0.5
0.5
1.4
0.6
0.6
0.6
1.1
1.5
0.6
1.1
1.0
1.9
1.1
1.6
1.3
0.6
0.5
0.8
0.6
0.5
0.2
0.5
1.5
0.7
1.5
1.2
0.5
1.3
1.3
1.2
0.9
1.8
1.8
2.1
2.1
2.6
0.4
0.4
2.1
2.1
1.2
1.1
1.5
2.1
1.2
1.3
0.4
1.3
2.1
0.5
1.4
1.7
1.7
0.5
0.5
0.5
1.3
1.3
0.5
1.4
0.5
1.0
1.3
0.6
1.5
0.9
1.7
0.9
1.9
2.3
0.9
0.5
1.0
0.8
0.7
1.5
0.6
0.8
0.7
1.3
1.5
0.7
1.1
1.4
1.9
1.2
1.7
1.5
0.9
1.1
0.7
0.8
0.8
0.3
0.6
1.5
0.7
1.2
1.1
0.6
1.2
1.2
1.1
1.5
1.1
1.1
1.1
1.6
2.5
0.1
0.3
1.7
1.6
1.3
1.2
1.7
1.6
1.3
1.2
0.3
0.3
1.6
0.3
1.2
1.7
1.8
0.3
0.4
0.4
1.5
1.2
0.5
1.3
0.4
0.9
1.2
0.4
1.2
1.1
2.0
1.1
1.7
2.9
0.4
0.4
0.5
0.7
0.5
1.4
0.8
0.7
0.4
1.1
1.2
0.5
1.2
1.2
1.7
1.2
1.6
2.0
0.5
0.8
0.8
0.7
0.7
0.5
0.8
1.2
0.7
1.3
1.2
0.9
1.4
1.3
1.2
2.4
26.4
29.5
30.2
32.9
33.6
3
3
3
3
4.2
4.8
5.6
5.8
EEL
PoEE
LnELn
AAA
AALn
PEE
38.1
39.0
40.2
41.8
42.4
42.5
AEL
AEPo
ALnLn
LnEL
EEP
LnLnLn
LnEPo
AEM
LnEM
AAL
42.9
4
4
4
4
3.0
3.5
4.3
4.9
(
2014) analyzed total fatty acids, while we analyzed only FAs con-
45.4
46.7
tained in TAGs, see below. Yet Table 1 clearly shows that the same
phenomenon took place in both cases, i.e. loss of PUFAs and an
increase in the production of saturated FAs in cultivations at higher
temperatures. This statement is entirely consistent with the results
of PCA analysis of both our data and the findings of Cepák et al.
AAPo
ALnL
LEL
4
4
4
4
7.4
7.8
8.0
8.3
AEP
LnAPo
PoEL
LnLnL
AAM
LnEP
LnPoLn
PoEPo
ALnM
MEL
LnLnM
MEPo
MEM
ALL
49.0
49.1
49.8
50.0
50.2
50.5
(
2014), see Figs. 1S and 2S.
2.2. Identification and production of triacylglycerols
Unlike the analysis and identification of fatty acids, which is the
50.7
50.9
51.0
51.3
51.6
subject of thousands of studies (13,500 according to the Web of
Science), the analysis and identification of TAGs is much less inten-
sively studied (630 studies according to the Web of Science). Out of
this number, papers describing analysis of diastereoisomers and
enantiomers of TAGs can be counted in units. This analysis is not
a trivial task, see e.g. the studies cited in this article. The method
most commonly used for analyzing the molecular species of the
TAGs is liquid chromatography–tandem mass spectrometry (LC–
MS/MS) using soft ionization techniques. Either atmospheric pres-
sure chemical ionization (APCI) or electrospray ionization (ESI) is
most commonly used, non-aqueous reversed-phase high-perfor-
mance liquid chromatography (NARP-HPLC) being the most com-
monly used LC technique. Alternatively, one can also use direct
injection of a mixture of TAGs into mass spectrometer, but this
method cannot identify the regioisomers and certainly not enanti-
omers. Based on our previous experience with identifying regioi-
somers and enantiomers we used here two methods, which
proved to be successful, viz. APCI and chiral chromatography with
MS/MS APCI. NARP-LC/tandem MS allowed us to identify dozens of
molecular species, see Table 2 and Fig. 1.
53.4
53.8
AAP
LnLL
5
5
5
4.3
4.6
5.1
PoAL
LnAP
PEL
PoAPo
LnPoL
LnPLn
AML
PoEP
LnPoPo
MAPo
LnML
MEP
LnMPo
AMM
LnMM
LLPo
55.6
56.2
5
5
5
5
7.4
8.1
8.5
9.0
59.2
59.3
59.6
60.0
60.2
60.5
60.6
60.7
60.8
60.9
61.0
LLL
PoPoL
LnPL
PAL
LPoL
PoAP
PEP
Positive ion APCI mass spectra of TAGs typically exhibited
[
M+H]+ ion and fragment ions of type [M+HꢀRCOOH] ([DAG] ),
+
+
61.2
61.3
61.5
resulting from the loss of an acyl chain from TAG. Monoacyl glyc-
erol ([MAG] ) is equivalent to [RCO+74] ion. The positive ion APCI
mass spectrum (EEA, for abbreviations of FAs see Table 1), i.e. the
+
+
6
6
6
1.6
1.9
2.2
LnPoP
MLL
+
peak at RT 26.4 min shows [M+H] ion at m/z 947.7 and the frag-
+
+
ment ions [M+HꢀRCOOH] at m/z 645.5 [EA] and at m/z 643.5
62.3
62.4
62.8
MAP
PoPoPo
PoML
PoMPo
LnMP
MMPo
+
+
+
[
EE] . Further ions at m/z 361.3 [A+74] and at m/z 359.3 [E+74]
+
or [RCO] at m/z 287.2 [arachidonoyl] and at m/z 285.2 [eicosapen-
taenoyl] were also present. These ions were observed with suffi-
cient intensity to analyze the mass spectrum.
6
6
4.2
4.5
65.6
Further TAGs were identified in a similar manner; the identifi-
cation was often complicated by the coelution of several TAGs,
(
continued on next page)
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Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013

ˇ
T. Rezanka et al. / Phytochemistry xxx (2015) xxx–xxx
4
Table 2 (continued)
and regioisomers are demonstrated on the example of di-eicosa-
pentaenoyl arachidonoins-and di-arachidonoyl-eicosapentaenoins
Figs. 3S–10S, see Supplements). One can use for the purpose the
RT
TAG
ECN
ACN
10 °C
25 °C
32 °C
(
6
6
6
6
6
6
6
6
6
7
7
7
7
7
7
7
8
6.1
MMM
MML
LPPo
PPoPo
LMP
LnPP
PMM
PAP
PoPL
PML
PoMP
PLL
MMP
PPL
PPoP
PMP
PPP
42
42
44
44
44
44
44
44
44
44
44
44
44
46
46
46
48
42:0
46:2
50:3
48:2
48:2
50:3
44:0
52:4
50:3
48:2
46:1
52:4
44:0
50:2
48:1
46:0
48:0
0.5
1.8
0.2
0.5
1.1
0.5
0.0
0.6
0.6
0.1
1.0
0.5
1.3
0.6
0.3
0.1
0.1
1.4
1.6
0.4
0.6
1.3
0.7
0.1
0.6
0.8
0.3
1.3
0.9
1.7
0.7
0.6
0.3
0.1
2.7
1.7
0.5
1.1
1.7
0.9
0.2
0.7
1.0
0.8
1.8
1.5
2.4
1.2
1.2
1.1
0.5
well-known rule (Byrdwell, 2005; Mottram, 2005), which says
6.7
6.9
7.3
7.9
8.1
8.5
9.1
9.5
1.3
1.5
1.6
3.0
5.3
5.6
6.2
1.3
+
that, based on the intensities of ions [M+HꢀRiCOOH] , it is possible
to determine whether a TAG is a 1,2-isomer (2,3-isomer) or a 1,3-
isomer. Conversely, enantiomers have almost identical mass spec-
tra, see Figs. 4S and 5S, so it cannot be determined from the mass
spectra which enantiomer is present and has to be differentiated
on the basis of comparison of retention times of the standards.
To prepare standards of the individual enantiomers of TAGs, we
used an already described method, the principle of which consists
in the hydrolysis of commercially available diacylphosphatidylch-
oline by lipase C and esterification of the free OH group of the cor-
responding diacylglycerol with a fatty acid (Rezanka et al., 2013).
As is evident from Fig. 2, optical purity, i.e. the yield of the desired
enantiomer, always exceeded 95%.
The elution order of the individual enantiomers and regioisom-
ers can partly be predicted, because, as described previously (Nagai
et al., 2011; Rezanka et al., in press), the enantiomer having a more
unsaturated FA at the sn-1 position of the glycerol backbone is
eluted earlier than the other enantiomer. Achiral TAG is eluted as
the last of the three. So, for example, the elution order of di-arach-
idonoyl-sn-eicosapentaenoin is sn-EAA, sn-AAE and sn-AEA. This
sequence however holds only under strictly defined conditions,
i.e. a certain type of stationary and mobile phases, temperature,
etc., see also Nagai et al. (2011), where the mere substitution of
see Table 2 and Fig. 1. For instance, three TAGs, AMM, LnMM, and
LLPo, were coeluted to form the peak at retention time
6
0.6 ± 0.1 min. They, as well as other coeluted peaks, were identi-
+
fied by tandem MS, which clearly identified peaks of the [DAG]
+
+
+
type, i.e. [AM] , at m/z 571.5, [LnM] at m/z 545.5, and [LL] at m/
z 599.5. TAGs often occur as regioisomers, i.e. positional isomers
on the glycerol backbone (Mottram, 2005). The overlapping of
TAG peaks means that overlapping of total ion current (TIC) takes
place, see also Fig. 1. However, if one uses neutral loss scan tandem
mass spectrometry (NLS), see Fig. 2, then one can distinguish and
thus also quantify, e.g. TAGs having nearly identical retention
times, such as, e.g., RT 42.9 min (EEP) and RT 43.0 min (LnLnLn).
3
CH group for Cl group in the stationary phase led to an inverse order
of elution of enantiomers. These findings indicate that, to ensure
correct identification, at least one of the two enantiomers must be
obtained either from commercial sources or by organic synthesis.
This statement does not apply to regioisomers, as they can be iden-
+
+
+
+
+
To quantify ions of the [DAG] type – ([EP] and ([LnLn] ) one
can thus use in fact NLS-E and NLS-Ln, which have different m/z,
see Table 2.
tified based on the abundance of ions of type ([AA] to [AB] ), see
above. Identification of most regioisomers having the same fatty
acyls in the sn-2 and sn-1 (sn-3) positions therefore usually does
not pose a problem.
To identify the enantiomers and diastereoisomers TAGs one
must use standards, since it is known that regioisomers and both
enantiomers have the same m/z values of ions but not the same
abundance of the ions. An achiral TAG, e.g. sn-EAE, differs as a
diastereomer in its mass spectrum from each of the two
enantiomers (sn-EEA and sn-AEE). Mass spectra of enantiomers
Selected ion monitoring (SIM) was used to determine the
appropriate enantiomers and regioisomers, see Fig. 3 acquired for
a mixture of two TAGs, i.e. di-arachidonoyl-eicosapentaenoins
and di-eicosapentaenoyl-arachidonoins obtained from algae cul-
tured at various temperatures (10, 25 and 32 °C).
Fig. 1. RP-LC/APCI-MS chromatogram total ion current (TIC) of the TAGs from the alga T. minutus, cultivated at 10, 25 and 32 °C.
ˇ
Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013

ˇ
5
T. Rezanka et al. / Phytochemistry xxx (2015) xxx–xxx
Fig. 2. Chiral LC of synthetic standards, see Table 3. The prefix ‘‘sn’’ was omitted in order to abbreviate the acronyms in the figure. TAGs containing myristic, arachidonic and
+
+
+
eicosapentaenoic acids as revealed by the neutral loss scan for [M+H] – 228, [M+H] – 304, and [M+H] – 302 Da, respectively, during APCI of synthetic TAGs.
Fig. 3 illustrates an obvious dependence of the change in the
ratios of individual enantiomers and regioisomers on cultivation
temperature. In the case of di-arachidonoyl-eicosapentaenoin it is
clearly seen that the percent proportion of achiral TAG (sn-AEA)
decreases with increasing temperature of cultivation, while Table 2
and Fig. 1, i.e. the results obtained from NARP-LC/APCI-tandem MS
obviously show that the total content of di-arachidonoyl-eicosapen-
taenoin decreases with increasing cultivation temperature. Next
examined was the TAG di-eicosapentaenoyl-arachidonoin. Here
both Table 3 and Fig. 3 shows that the proportion of achiral TAG
The biosynthesis of TAGs involves three acyl transferases
(Coleman and Douglas, 2004), each of which esterifies only one
of the three hydroxyls of glycerol; the resulting combinations are
not random. From the above findings it is clear that the achiral
TAGs are preferably synthesized at low temperatures, implying a
different activity of the enzymes acyl-CoA: sn-glycerol-3-phos-
phate acyltransferase (GPAT), acyl-CoA: lyso-phosphatidic acid
acyltransferase (LPAAT), and acyl-CoA: diacylglycerol acyltransfer-
ase (DGAT) at various temperatures.
This phenomenon is demonstrated using a principal component
analysis, where the first axis scores of the samples are clearly cor-
related with the temperature of cultivation (Fig. 4). PCA analysis of
all 18 measured abundant TAGs is shown in Fig. 11S, even if the
image is not as illustrative as Fig. 4.
Large differences between the proportion of enantiomers and
regioisomers were detected by Nagai et al. (2011) for marine fish
(skipjack tuna, sardine) and mammals (steller sea lion, harp seal).
Basically they are final predators in the food chain that begins with
phytoplankton which is the source of C20 polyunsaturated fatty
acids – arachidonic and eicosapentaenoic acid. A similar phenom-
enon was recorded also in algae grown under both stressful condi-
tions and under different nutritional conditions (Rezanka et al.,
2012), and in strains of the genus Stichococcus obtained from loca-
tions with different temperature conditions (Svalbard, a lake in a
temperate region, thermal springs) (Rezanka et al., in press).
Based on the knowledge about the influence of temperature on
the crystalline structure of TAGs (Craven and Lencki, 2013) it is
clear that TAGs exhibit polymorphism and usually crystallize in
(sn-EAE) has an increasing tendency with temperature increasing
from 10 °C to 25 °C and then it decreases at 32 °C, the sum of asym-
metric (sn-EEA and sn-AEE) to symmetrical sn-EAE having a rising
characteristic, i.e. 0.56:0.74:1.20. Here again there is a sharp reduc-
tion in the proportion of the di-eicosapentaenoyl-arachidonoin,
which is shown in Fig. 1 and in Table 2. It is also confirmed by the
data of Table 1, which clearly point to the loss of both C20 PUFAs
with increasing temperature of cultivation. SIM and tandem MS
were similarly used to identify and further semi-quantify 4 trios of
enantiomers and regioisomers of TAGs (di-eicosapentaenoyl myris-
toin-, di-arachidonoyl-myristoin, di-myristoyl-eicosapentaenoin,
and di-myristoyl-arachidonoin). The results are shown in Table 3.
For TAGs having two myristic acids in the molecule, i.e. di-myr-
istoyl-eicosapentaenoin and di-myristoyl-arachidonoin the con-
tent of achiral TAG, i.e. sn-MEM and sn-MAM remains
substantially constant. In both cases the content of the two pairs
of enantiomers is rapidly increasing with increasing cultivation
temperature. It can therefore be concluded that cultivation tem-
perature has a significant impact not only on the amount of PUFAs
three polymorphic forms, exhibiting with increasing temperature
0
(
(
a phenomenon known for decades) but also on the content of TAG
which has also been reported by Olofsson et al., 2012) and espe-
increasing stability of the sub-cells, i.e.
most stable form.
a
? b ? b, where b is the
cially on the ratio of enantiomers and regioisomers of the relevant
molecular species. Unfortunately, the research in this area has so
far been fully focused on those TAGs that are useful in the food
industry (Craven and Lencki, 2013) and not on the oil bodies in liv-
ing cells of photosynthesizing plants, not even on oleaginous crops
such as sunflower, canola, soya, etc. So far, all research has been
directed toward understanding the adaptation of membranes
However, natural TAGs are always a mixture of different molec-
ular species that differ in chain length and number of double
bond(s), and therefore they are a mixture of polymorphic forms
that can coexist simultaneously. The crystalline structure of TAGs
is important for fluidity of oil bodies in the cells of living organ-
isms, since each crystalline form presents unique properties with
respect to plasticity and solubility. The crystallization and poly-
morphic behavior for achiral, chiral racemic, and chiral-enantio-
pure TAGs is remarkably different (Craven and Lencki, 2013). All
(
membrane fluidity) at low temperatures, completely neglecting
the above-mentioned oil bodies (Morgan-Kiss et al., 2006).
ˇ
Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013

ˇ
6
T. Rezanka et al. / Phytochemistry xxx (2015) xxx–xxx
Fig. 3. Chiral-LC of TAGs (SIM – selected ion monitoring) from T. minutus, cultivated at various temperatures (10, 25 and 32 °C).
Table 3
TAG cannot form the b polymorph because both enantiomers must
be present in the unit cell.
Relative quantities (%) of regioisomers and enantiomers of TAGs from T. minutus.
TAG
10 °C
25 °C
32 °C
The life functions of the cell have to be retained throughout
these transitions. One of the possibilities how to achieve this is
to keep oil bodies in a fluid – non-crystalline state, which could
be achieved by transitions between crystalline forms. Regretfully,
the data on the melting point of crystalline forms of TAG regioi-
somers as well as enantiomers are available for only a few of them.
Despite this, some data have been gathered (Serebrennikov et al.,
AMM
MMA
MAM
EMM
MME
AAM
MEM
MAA
AMA
AAA
EEM
MEE
EAA
0.23
0.13
1.19
0.34
0.22
0.19
1.45
0.10
0.80
1.19
0.35
0.17
0.06
2.22
0.04
0.54
0.48
0.31
1.40
3.92
0.26
0.30
1.09
0.50
0.40
0.21
1.40
0.17
0.64
1.10
0.41
0.42
0.07
1.74
0.04
0.25
0.52
0.38
1.21
1.84
0.33
0.39
0.91
0.67
0.73
0.18
1.52
0.23
0.45
1.19
0.57
0.71
0.08
1.21
0.07
0.17
0.29
0.30
0.49
1.08
1
961), e.g. for SLO and SOL, in which the melting points for individ-
ual
a
and b forms are ꢀ2 to ꢀ4 °C and ꢀ15 to ꢀ17 °C and ꢀ11.5 to
ꢀ13 °C and ꢀ18 to ꢀ19 °C, respectively. Although this holds for
two regioisomers, i.e. compounds having the same FAs and the
same molecular weight, it is surprising that a mere change in the
positions of two FA (O and L) on glycerol backbone shifts the melt-
ing point by as much as 10 °C.
EME
AAE
AEA
EEA
AEE
The composition of TAGs and/or the proportion of individual
regioisomers and enantiomers are thus seen to affect whether
the oil bodies are, at a certain temperature, fluid or crystalline.
We can therefore hypothesize that an alga could regulate their
adjustment to the temperature changes by changing the content
EAE
EEE
TAGs (achiral and chiral, racemic mixture or enantiomers) can, in
of FA and phospholipids in the membranes but also by changing
0
0
principle, adopt
a
and b forms, whereas only achiral and racemic
TAG fluidity, i.e. transition between
oil bodies.
a
? b ? b forms and back in
mixtures of chiral TAGs can form the b polymorph. Enantiopure
ˇ
Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013

ˇ
7
T. Rezanka et al. / Phytochemistry xxx (2015) xxx–xxx
Fig. 4. Principal component analysis (PCA) of TAG composition in the alga T. minutus cultivated at 10 °C, 25 °C and 32 °C. The first two canonical axes are plotted; they
explained 94.5% of the variability. For FA abbreviations, see Table 1.
Our literature search did not reveal any such case and this
observation thus requires further measurements in order to
establish a valid hypothesis about the biosynthesis of TAGs.
of Botany AS CR (http://ccala.butbn.cas.cz/index.php). Laboratory
cultivation units consisted of glass cultivation flasks, which were
placed in a thermostatic bath and continuously illuminated by
white fluorescent light tubes (Osram L 36W/830 Lumilux, Ger-
2
1
many). The incident light intensity was 80 lmol/m /s photosyn-
3
. Conclusion
thetic active radiation (PAR). The light intensity was measured
using a digital lux-meter PU 550 (Metra, Czech Republic) equipped
with a PAR sensor calibrated according to the LI-185 quantum sen-
sor (LI-COR, USA). Cultures were grown at 2ꢁ concentrated ‘‘Z’’
nutrient medium (Staub, 1961) and aerated by the mixture of air
The ratio of regioisomers and enantiomers of TAGs in T. minutus
has been shown to be affected by cultivation temperature. We used
the principal component analysis to detect and illustrate this
effect. This approach can conceivably be used on a broader scale
to identify if and to what degree the production of some compound
of interest is influenced by factors such as cultivation conditions.
and CO
tivation flask was 900 mL. Three different temperatures were used:
0, 25 a 32 °C. The cultures were cultivated 21 days. The biomass
2
(2%, v/v). The volume of the algal suspension in each cul-
1
was harvested by centrifugation, frozen at ꢀ80 °C and lyophilized.
4
. Experimental
4.1. Chemical and standards
4.3. Enzymatic synthesis of 1,2-dieicosapentaenoyl-, 1,2-
diarachidonoyl- and 1,2-dimyristoyl-sn-glycerols
Acetonitrile, 2-propanol, THF (tetrahydrofuran), hexane, dichlo-
romethane, 4-dimethylaminopyridine (DMAP) EDCI (1-ethyl-3-(3-
dimethylaminopropyl) carbodiimide), 1,2-dieicosapentaenoyl-sn-
glycero-3-phosphocholine, arachidonic and cis-5,8,11,14,17-eico-
sapentaenoic acids and phospholipase C from Clostridium perfrin-
gens (Clostridium welchii) Type I, lyophilized powder, 10–50 units/
mg protein were purchased from Sigma–Aldrich (Prague, CR). Tri-
arachidonin (5c,8c,11c,14c) and trieicosapentaenoin (5,8,11,14,
1,2-Dieicosapentaenoyl-sn-glycero-3-phosphocholine,
1,2-
diarachidonoyl-sn-glycero-3-phosphocholine and 1,2-dimyristoyl-
sn-glycero-3-phosphocholine were treated with phospholipase C
(Type I from C. perfringens (C. welchii)) as described by Christie
(1989). In brief, phospholipase C from C. welchii (3 mg) in 0.5 M
tris(hydroxymethyl)methylamine (tris) buffer (pH 7.5; 2 mL), con-
taining 2 mM calcium chloride, was added to phosphatidylcholine
(15 mg) in diethyl ether (2 mL). The mixture was shaken at room
temperature for 3 h and extracted three times with diethyl ether
(4 mL portions). The ether layer was dried over anhydrous sodium
sulfate before being evaporated in a stream of nitrogen at ambient
temperature. Pure 1,2-diacyl-sn-glycerols were obtained by pre-
parative TLC on a layer of silica gel G impregnated with boric acid
1
7-all cis) was purchased from Larodan Fine Chemicals,
Malmö, Sweden. 1,2-Diarachidonoyl-sn-glycero-3-phosphatidyl-
choline and 1,2-dimyristoyl-sn-glycero-3-phosphocholine were
purchased from Avanti Polar Lipids, Inc., AL, USA.
4.2. Cultivation
(
10%, w/w), with hexane–diethyl ether (50:50, v/v) as solvent sys-
T. minutus Lukavsky et Pribyl 2005/1 CCALA 931 was obtained
from Culture Collection of Autotrophic Organisms of the Institute
tem. The appropriate band was located under UV light and was
eluted from the adsorbent with diethyl ether (Christie, 1989).
ˇ
Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013

ˇ
T. Rezanka et al. / Phytochemistry xxx (2015) xxx–xxx
8
The esterification of glycerol derivatives with acids was
described previously (Ziegler and Berger, 1979; Halldorsson et al.,
003). Briefly, a solution of 1,2-diacyl-sn-glycerol (57 mg) and
eicosapentaenoic or arachidonic acids (33 mg) as a free acid in
dichloromethane (1.0 mL) was added to DMAP (6.1 mg, 50 mol)
and EDCI (18.6 mg, 0.12 mmol) in dichloromethane (1.0 mL) under
an inert atmosphere (argon). The resulting solution was stirred at
room temperature for 15 h. The reaction was stopped by passing
the reaction mixture in ether/dichloromethane (10:90) through a
short column packed with silica gel. Solvent removal in vacuo
afforded the pure products as yellowish oil. The yields of the target
sn-EEA and sn-AAE were 45 and 38 mg, respectively. Other TAGs
were prepared similarly, the list of all synthesized triacylglycerols,
see Table 1S.
current 7 lA, sheath gas – high-purity nitrogen, pressure
0.45 MPa, and auxiliary gas (also nitrogen) flow rate 15 ml/min.
Positively charged ions with m/z 200–1000 were scanned with a
scan time of 0.5 s. The whole HPLC flow (0.35 ml/min) was intro-
duced into the APCI source without any splitting. TAGs were sepa-
rated using a gradient solvent program with acetonitrile (ACN) and
2-propanol (iPrOH) as follows: initial ACN/iPrOH (99:1, vol/vol);
linear from 5 min to 120 min ACN/iPrOH 30:70, vol/vol); held until
30 min; the composition was returned to the initial conditions over
10 min. A peak threshold of 0.07% intensity was applied to the
mass spectra. Data acquisition and analyses were performed using
PC with MassLab 2.0 for Windows XP applications/operating
software.
2
l
4.7. TAGs analysis by chiral LC/MS-APCI
4.4. Isolation of TAGs
The LC system used for separation in the chiral mode was the
The extraction procedure was based on the method of Bligh and
Dyer (1959), except that 2-propanol was substituted for methanol,
since 2-propanol does not serve as a substrate for phospholipases
same as that used in the reversed-phase mode. TAGs (ꢂ1 mg/mL
in methanol) were chromatographed on two Astec cyclobond™ I
2000 DMP (3,5-dimethyl-phenyl-carbamate modified b-cyclodex-
(
Kates, 1986). The alcohol–water mixture was cooled and one part
trin) chiral LC columns (5
ies, from Sigma. Mobile phase was a gradient from 90% of A and
0% of B at 0 min to 50% of A and 50% of B during 180 min, and then
lm, 25 cm ꢁ 4.6 mm) connected in ser-
chloroform was added and the lipids from the lyophilized cells of
different cultivations of T. minutus were extracted for 30 min.
Insoluble material was sedimented by centrifugation and the
supernatant was separated into two phases. The aqueous phase
was aspirated off and the chloroform phase was washed three
times with two parts 1 M KCl each. The resulting chloroform phase
was evaporated to dryness under reduced pressure.
1
isocratically for 30 min, where A is hexane and B is hexane-2-pro-
panol (99:2, v/v) mixture (Rezanka and Sigler, 2014). The flow rate,
column temperature, and injection volume were 0.5 mL/min, 25 °C,
and 10 ll, respectively. Other separation conditions were as used
in the RP-LC, see above.
First, total lipid extracts were applied to Sep-Pak Vac Silica car-
tridge 35cc (Waters; with 10 grams of silica normal-phase), and
TAGs were subsequently eluted from the cartridge with 40 mL of
chloroform-methanol (9:1). The eluate was evaporated and the
oil samples were dissolved in an acetonitrile-2-propanol-hexane
mixture (1:1:1, v/v/v), which was injected for HPLC analysis.
Acknowledgements
The research was supported by GACR (Grant Agency of the
Czech Republic) projects P503/11/0215 and P503 14-00227S, by
MEYS (Czech Republic Ministry of Education, Youth and Sports)
project Algatech, CZ.1.05/2.1.00/03.0110, by Institutional Research
Concept RVO61388971 and Biorefinery Research Centre of Compe-
tence (BIORAF, TE 01020080), Technology Agency of the Czech
Republic.
4.5. FAMEs analysis
The TAGs (ꢂ5 mg) were saponified overnight in 10%
KOH-MeOH at room temperature. A fatty acid fraction obtained
from saponification was partitioned between alkali solution (pH
Appendix A. Supplementary data
9
2
) and Et O to remove basic and neutral components. The aqueous
phase, containing fatty acids, was acidified to pH 2 and extracted
with hexane. The fatty acid fraction was methylated using
Supplementary data associated with this article can be found, in
the online version, at http://dx.doi.org/10.1016/j.phytochem.
2 2
CH N . GC–MS of fatty acid methyl ester (FAME) mixture was
2
014.12.013.
done on a Finnigan 1020 B in EI mode. Splitless injection was at
1
6
00 °C, and a fused silica capillary column (Supelcowax 10;
0 m ꢁ 0.25 mm i.d., 0.25 mm film thickness; Supelco, Prague)
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HPLC equipment consisted of a 1090 Win system, PV5 ternary
pump and automatic injector (HP 1090 series, Agilent, USA) and
m par-
two Hichrom columns HIRPB-250AM 250 ꢁ 2.1 mm ID, 5
l
ticle size, in series. This setup provided us with a high-efficiency
column – approximately 26,000 plates/250 mm. A quadrupole
mass spectrometer system Navigator (Finnigan MAT, San Jose,
CA, USA) was used for analysis. The instrument was fitted with
an atmospheric pressure chemical ionization source (vaporizer
temperature 390 °C, capillary heater temperature 260 °C, corona
ˇ
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Please cite this article in press as: Rezanka, T., et al. Temperature dependence of production of structured triacylglycerols in the alga Trachydiscus minutus.
Phytochemistry (2015), http://dx.doi.org/10.1016/j.phytochem.2014.12.013