Biosynthesis and mobilization of arachidonic-acid-rich triacylglycerols in the green microalga Parietochloris incisa
Thesis submitted in partial fulfillment of the requirements for the degree of "DOCTOR OF PHYLOSOPHY"
by
Pushkar Shrestha
Submitted to the Senate of Ben-Gurion University of the Negev
חשון תשס"ו 11/2005
Biosynthesis and mobilization of arachidonic-acid-rich triacylglycerols in the green microalga Parietochloris incisa
Thesis submitted in partial fulfillment of the requirements for the degree of "DOCTOR OF PHYLOSOPHY"
by
Pushkar Shrestha
Submitted to the Senate of Ben-Gurion University of the Negev
Approved by the advisors:
Prof. Zvi Hacohen ______
Prof. Bezalel Kessler ______
Dr. Inna Khozin-Goldberg ______
חשון תשס"ו 11/2005
Beer-Sheva TABLE OF CONTENTS
Acknowledgments I
Summary II
List of figures and tables V
List of abbreviations and symbols IX
1. INTRODUCTION 1
1.1. Polyunsaturated fatty acids 1
1.2. Importance of PUFAs 2
1.3. Occurrence of PUFA-rich TAG 6
1.4. Biosynthesis of TAG 7
1.4.1. De novo synthesis of FA 8
1.4.2. Chloroplastic and extrachloroplastic lipids 10
1.4.3. Membrane desaturases 12
1.4.4. Biosynthesis of C20 PUFA in algae 13
1.4.5. Partitioning of FA to TAG 16
1.4.6. Acyltransferases of TAG biosynthetic pathway 23
1.5. Factors affecting the biosynthesis of TAG 25
1.5.1. Temperature 25
1.5.2. Light 26
1.5.3. Nutrient deprivation 27
1.6. Mutant studies 30
1.7. Role of TAG 31
1.8. Working hypothesis 32
2. MATERIALS AND METHODS 34
2.1. Growth conditions 34 2.2. Growth Parameters 35
2.3. Lipid analysis 35
2.3.1. Lipid extraction 35
2.3.2. Fatty acid analysis 36
2.3.3. Lipid separation 37
2.3.4. Molecular species separation 38
2.3.5. Positional analysis of fatty acids distribution in individual lipids 39
2.4. Nitrogen starvation and recovery experiments 40
2.5. Inhibitor studies 40
2.5.1. Salicylhydroxamic acid (SHAM) 40
2.5.2. Sethoxydim 41
2.6. Characterization of TAG biosynthetic enzymes 41
2.6.1. Grinding and homogenization 41
2.6.2. Cellular fractionation 41
2.6.3. Enzymatic assays 42
2.7. Mutagenesis 44
3. RESULTS 45
3.1. Lipid studies 45
3.1.1. Lipid classes and fatty acid composition 45
3.1.2. Molecular species composition of PC 48
3.1.3. Molecular species composition of PE 50
3.1.4. Molecular species composition of DGTS 52
3.1.5. Molecular species composition of TAG 53
3.2. Role of extrachloroplastic lipids in TAG synthesis 56
3.2.1. Sethoxidim treatment 56
3.2.2. Phosphate starvation 58
3.2.3. P- and N-starvation 63
3.3. Enzymes of TAG biosynthesis 64
3.3.1. Optimization of cell homogenisation 64
3.3.2. Development of a protocol for cellular fractionation 67
3.3.3. DAGAT assay utilizing [1_14C]oleoyl-CoA and non-labeled DAG 71
3.3.4. DAGAT activity in microsomes 74
3.3.4.1. Effect of time and protein 74
3.3.4.2. Substrate specificity 74
3.3.4.3. Solubility and availability of DAG substrate in assay 75
3.3.4.4. Effect of thiol reagents 77
3.3.4.5. Effect of cations 78
3.3.4.6. The origin of labeled DAG and the evidence for the MAGAT activity 79
3.3.5. DAGAT assay in microsomes with [1_14C]DAG and non-labeled oleoyl-CoA 80
3.3.5.1. Protein and time dependence 81
3.3.5.2. Effect of oleoyl-CoA concentrations 83
3.3.5.3. Selectivity of acyl-CoA substrates 84
3.3.5.4. Effect of ethanol 85
3.3.5.5. Effect of thiol-reagent PCMB 85
3.3.5.6. Effect of salts 86
3.3.6. Acyl-CoA independent TAG synthesis from [1_14C]1,2-dioleoyglycerol 88
3.3.6.1. Time and protein dependence 88
3.3.6.2. pH and temperature dependence 89
3.3.6.3. Effect of thiol reagents PCMB, DTT and CuCl2 90
3.3.6.4. Distribution of activity in the cellular fractions 91
3.3.7. DAGAT activity in oil bodies in assay with [1-14C]oleoyl-CoA 92
3.3.8. DAGAT assay in oil bodies with [1-14C]dioleoylglycerol 94
3.3.8.1. Factors affecting DAGAT activity in oil bodies 95
3.3.9. MAGAT activity in microsomes 100
3.3.9.1. Effect of MAG concentration 101
3.3.9.2. Time dependence 101
3.3.9.3. Effect of ethanol 102
3.3.9.4. Effect of dioleoylglycerol addition 105
3.3.9.5. Effect of MgCl2 105
3.3.10. Lipolytic activity in microsomes 107
3.3.10.1. pH and temperature 107
3.3.10.2. Time and protein 110
3.3.10.3. Effects of divalent metals and inhibitors on DAG lipase activity 111
3.3.10.4. Positional specificity 112
3.3.11. Evidence for the activity of TAG lipase 113
3.4. Role of AA-rich TAG 114
3.4.1. Nitrogen starvation and recovery 114
3.4.1.1. Nitrogen starvation 114
3.4.1.2. Recovery from the nitrogen starvation 115
3.4.1.3. Alterations in lipid and fatty acid content and composition 117
3.4.1.4. Molecular species analysis 119
3.4.1.5. Radiolabelling 121
3.4.2. SHAM treatment and recovery 123
3.5. Mutant studies 128
4. DISCUSSION 132
4.1. Lipids involved in the biosynthesis of AA-rich TAG 132
4.2. Role of extrachloroplastic lipids in TAG synthesis 136
4.3. Enzymes of TAG biosynthesis 138
4.4. Role of TAG 150
4.4.1. Nitrogen starvation and recovery 150
4.4.2. Recovery from SHAM treatment 155
4.5. Mutant studies 156
5. REFERENCES 158
6. HEBREW ABSTRACT
ACKNOWLEDGMENTS
I would like to express my deep gratitude to my supervisors Prof. Zvi HaCohen, Prof.
Bezalel Kessler and Dr. Inna Khozin-Goldberg for providing me an excellent opportunity to have this study and for their kind supervision and support to accomplish this dissertation research. I am very thankful to Inna for her sincere guidance in laboratory and in everyday life during the whole tenure in Israel, without which this study wouldn’t have been materialized.
I am grateful to Shosh Didi-Cohen, Ilana Saller, Dorit Levin, Ben Friehoff and Amos
Masika for their crucial help during this study and thanks to all my friends whom I met in
Midreshet Ben Gurion for their kindness.
I also would like to thank my teachers, Prof. Sanu Devi Joshi, Prof. Govinda Prasad
Ghimire, Dr. Micha Guy, Dr. Micha Volokita and friends, Dr. Deepak Khadka and Dr. Roshan
Shrestha, whose goodwill promoted me to commence this endeavor.
I am highly indebted to my parents and aunt, who bestowed me with selfless love and care and continuous encouragement for study since my childhood. Thanks for their blessings.
Finally, my heartfelt thank goes to my wife, Shiru and son, Sakar for their patience, understanding and every support in good and hard moments in Sde Boker. Her support, encouragement and companionship greatly facilitated my journey through this study.
I SUMMARY
The very long chain-polyunsaturated fatty acid (VLC-PUFA), arachidonic acid (AA, 20:4ω6) is essential for brain development and critical biological functions of human health. Over 30% of dry weight of the green microalga Parietochloris incisa are triacylglycerol (TAG) and over 95% of cellular AA are deposited in these TAG. The accumulation of AA-rich TAG brought about a great interest in exploring the mechanism of its biosynthesis and distinctive role in this alga.
Analysis of lipid classes revealed that AA and its precursors are concentrated mostly in the extraplastidic lipids. The molecular species and stereo-specific analyses of fatty acid distribution of phosphatidylcholine (PC) revealed the presence of various C18 PUFAs and AA in the sn-2 and 16:0 in the sn-1 position and indicated that ∆12 and ∆6 desaturations occurred at the sn-2 position. Molecular species of phosphatidylethanolamine (PE), containing AA at the sn-1 position and C20 PUFA at the sn-2 position are likely to be involved in the ∆5 desaturation of
20:3. Diarachidonoyl-PC and -PE were among the major molecular species in both the exponential and stationary phases. The presence of the intermediates, C18:1-3 and C20:3, primarily in the sn-2 position of diacylglycerol (N,N,N)-trimethylhomoserine (DGTS), suggests the role of this position in the ∆12, ∆6 and possibly also in ∆5 desaturations.
Most molecular species of TAG contain 2 or 3 AA moieties. We assume that AA synthesized in PE is exported to PC, DGTS and TAG and partly to the chloroplastic lipids. The
AA/AA molecular species of PC and PE may have a key role in donating AA for TAG by either providing AA moieties to the acyl-CoA pool via the acyltransferases of the Kennedy pathway, or directly as DAG (diacylglycerol). AA derived from the sn-2 position of PE (18:1ω7/AA) can be also incorporated into the sn-1 and sn-3 positions of TAG.
The significance of extraplastidic polar lipids in AA-rich TAG biosynthesis was studied following administration of sethoxidim, an inhibitor of the de novo fatty acids synthesis, and under P-starvation.
II In order to characterize the final and committed step of TAG biosynthesis involving diacylglycerol acyltransferase (DAGAT), a cellular fractionation protocol was developed. This protocol enabled to minimize TAG degradation and ensure the isolation of intact oil bodies.
When membrane fractions and oil bodies were applied in in vitro assays containing labeled substrates, [14C]oleoyl-DAG or [14C]oleoyl-CoA and unlabelled acyl-CoA or DAG, respectively, labeled TAG was formed, indicating the activity of DAGAT. In the both types of DAGAT assays, the activity was drastically reduced in the presence of the thiol-modifying reagent p- chloromercuribenzoic acid (PCMB), suggesting that a serine or a cysteine residue is important for the enzyme activity. In the presence of [1-14C]oleoyl-CoA, DAGAT of P. incisa showed some preference towards the 18:1/18:1 molecular species of DAG over the AA/AA, 18:2/18:2, and 16:0/16:0 molecular species. An acyl-CoA-independent TAG biosynthesis has been also revealed, probably accomplished by DAG:DAG transacylase (DGTA). This activity was pronounced in the presence of 150 mM Mg2+ and had the highest specific activity for TAG formation.
In the DAGAT assay with [14C]oleoyl-CoA, in addition to TAG, PC, PE, DGTS, DAG, monoacylglycerol (MAG), was also labeled, indicating the activity of several acyltransferases that might be directly or indirectly involved in the biosynthesis of TAG. The MAGAT activity was shown when 2-MAG was utilized as the sole acceptor of [14C]oleoyl-CoA.
The lipase activity in the microsomes was deduced by the release of FFA from the labeled
DAG. This activity demonstrated positional selectivity to the acyl composition of DAG. Oleic acid was released from both position of 18:1/18:1 DAG, however, AA was not released from the sn-2 position of 18:0/20:4 DAG and was retained in the form of MAG. These data may also suggest a MAGAT–mediated pathway leading to AA-rich TAG.
Delipidated oil bodies proteins catalyzed the incorporation of [1-14C]oleoyl-CoA into TAG,
Untreated oil bodies showed activity only in assay with [1-14C]1,2 dioleoylglycerol. This activity
III was inhibited by niacin, an inhibitor of DAGAT in mammals, and thiol-modifying reagents. In
14 the presence of 150 mM MgCl2, oil bodies, as microsomes, could utilize [1- C]1,2 dioleoylglycerol as their sole acyl donor for the synthesis of TAG. Distinctly from microsomal enzyme, the activity in oil bodies was inhibited by PCMB, suggesting the presence of a different isoform of DGTA.
Taken together, it can be concluded that oil bodies and membrane fractions of P. incisa can accomplish the final step of TAG assembly in vitro. The source of the activity was not DAGAT exclusively. Probably, several enzymes are assigned to maximize the specific incorporation of
AA into the glycerol backbone of the TAG at the different metabolic steps.
The utilization and importance of AA-rich TAG for the construction of chloroplastic lipids was revealed by a study of the changes in AA distribution in lipids during recovery from nitrogen starvation at 24 and 12 °C. At both temperatures, TAG was mainly consumed to support growth, however, there was a significant increase in the content of AA in the chloroplastic lipids, predominantly, in the eukaryotic molecular species of monogalactosyldiacylglycerol (MGDG) at
24 °C, but much less so at 12 °C. Our data point to the existence of different modes of operation for the construction of chloroplastic lipids that the alga can utilize to support growth under changing environmental conditions. Comparatively slower build-up of chlorophyll and biomass following growth recovery of the salicylhydroxamic acid (SHAM)-treated cells with decreased
AA content, also indicated to the importance of AA-rich TAG for the synthesis of chloroplastic lipids during the growth recovery from unfavorable conditions.
In order to facilitate the understanding of the mechanisms, underlying the biosynthesis of
AA-rich TAG and their role in the organism, mutants using chemical mutagenesis. Lipid analysis of the B105, B107 and B109 mutants showed lower levels of neutral lipids in comparison to the
WT. In the B107 mutant, a significant increase in 18:1ω9 and decrease in 18:3ω6 of PC, PE and
DGTS, suggested a possible inhibition of the ∆12 and ∆6 desaturases.
IV
V
LIST OF FIGURES AND TABLES
Figures
Figure 1. Metabolic pathways of PUFAs in the human and animal body. 3
Figure 2. De novo fatty acid synthesis in higher plants and algae. 9
Figure 3. Biosynthesis of membrane glycerolipids by prokaryotic and eukaryotic
pathways in Arabidopsis leaves. 11
Figure 4. Various pathways of TAG biosynthesis in plants. 17
Figure 5. The Kennedy pathway of TAG biosynthesis. 18
Figure 6. Pathway for unsaturated TAG biosynthesis. 18
Figure 7. DAG-DAG transacylation pathway for TAG biosynthesis. 19
Figure 8. PC-DAG transacylation pathway of TAG biosynthesis. 20
Figure 9. HPLC-ELSD chromatogram of the molecular species of PC in the
logarithmic and stationary cultures of P. incisa. 49
Figure 10. HPLC-ELSD chromatogram of the molecular species of PE in the
logarithmic and stationary cultures of P. incisa. 51
Figure 11. HPLC-ELSD chromatogram of the molecular species of DGTS in the
logarithmic and stationary cultures of P. incisa. 52
Figure 12. HPLC-ELSD chromatogram of the molecular species of TAG in the
logarithmic and stationary cultures of P. incisa. 54
Figure 13. Comparison of chlorophyll, dry weight, total fatty acid and AA contents
in control and P-starved cultures. 59
Figure 14. Effect of pH on the degree of TAG degradation in cell homogenates. 65
Figure 15. Effect of the composition of homogenization buffers on the degradation
of TAG. 66
Figure 16. Outline of the cellular fractionation protocol of P. incisa. 68
V Figure 17. Isolated oil bodies under a light microscope. 68
Figure 18. TLC separation of total lipid extract of oil bodies. 70
Figure 19. Incorporation of radioactivity from [1-14C]oleoyl-CoA into various lipids
in presence of cell-free homogenate, pellet 25,000 x g, pellet 100,000 x g,
supernatant 100,000 x g and oil bodies. 71
Figure 20. Incorporation of radioactivity from [1-14C]oleoyl-CoA into TAG in the
absence and presence of dioleoylglycerol. 74
Figure 21. Substrate specificity of DAGAT towards four DAG species. 75
Figure 22. Effect of detergents and ethanol on the incorporation of radioactivity from
[1-14C]oleoyl-CoA into TAG. 76
Figure 23. The incorporation of [1-14C]oleoyl-CoA into TAG activity by delipidated
microsomes in the absence and presence of dioleoylglycerol. 77
Figure 24. Effect of PCMB on incorporation of [1-14C]oleoyl-CoA into TAG and
free fatty acid production in the presence and absence of ethanol. 78
Figure 25. Effect of different concentrations of MgCl2 on DAGAT activity in the
presence and absence of ethanol. 79
Figure 26. Incorporation of radioactivity from [1_14C]dioleoylglycerol into TAG in
the presence of cell free extract, pellet 25,000 x g, soluble fraction,
pellet 100,000 x g. 80
Figure 27. Effect of protein content on the incorporation of radioactivity from
[1_14C]1,2-dioleoylglycerol into TAG. 82
Figure 28. Time-dependence of the incorporation of radioactivity from
[1_14C]1,2-dioleoylglycerol into TAG. 82
Figure 29. Effect of concentration of oleoyl-CoA on the incorporation of radioactivity
from [1_14C]1,2-dioleoylglycerol into TAG. 83
VI Figure 30. Incorporation of radioactivity from [1_14C]1,2-dioleoylglycerol to TAG
in the presence of different acyl-CoA substrates. 84
Figure 31. Effect of ethanol and PCMB on the incorporation of radioactivity from
[1_14C]1,2-dioleoylglycerol into TAG. 85
Figure 32. Effect of the concentration of MgCl2 on the incorporation of radioactivity
from [1_14C]1,2-dioleoylglycerol into TAG. 86
_14 Figure 33. Effect of MgCl2 on incorporation of radioactivity from [1 C]1,2-
dioleoylglycerol into TAG in the absence and the presence of oleoyl-CoA. 87
Figure 34. Time-dependence of the incorporation of [1_14C]1,2-dioleoylglycerol into
TAG at 150 mM of MgCl2. 88
Figure 35. Protein dependence of the incorporation of [1_14C]1,2-dioleoylglycerol into
TAG at 150 mM of MgCl2. 88
Figure 36. Effect of pH on incorporation of radioactivity from [1_14C]1,2-
dioleoylglycerol into TAG at 150 mM MgCl2. 89
Figure 37. Effect of temperature on incorporation of radioactivity from [1_14C]1,2-
dioleoylglycerol into TAG at 150 mM MgCl2. 89
Figure 38. Effect of oleoyl-CoA, PCMB and DTT on the incorporation of [1_14C]1,2-
dioleoylglycerol into TAG in the presence of 150 mM MgCl2. 90
Figure 39. Incorporation of radioactivity from [1_14C]dioleoylglycerol into TAG in the
presence of 150 mM MgCl2 in cell-free homogenate, pellet 25,000 x g,
supernatant 100,000 x g, pellet 100,000 x g. 91
Figure 40. Effect of concentration of dioleoylglycerol on the incorporation of radioactivity
from [1-14C]oleoyl-CoA into TAG by delipidated proteins of oil bodies. 92
Figure 41. Time-dependence of the incorporation of radioactivity from
[1-14C]oleoyl-CoA into TAG by delipidated proteins of oil bodies. 93
VII Figure 42. Protein dependence of the incorporation of radioactivity from
[1-14C]oleoyl-CoA into TAG by delipidated proteins of oil bodies. 93
Figure 43. Effect of delipidation and fatty acid composition of DAG on the incorporation
of radioactivity from [1-14C]oleoyl-CoA into TAG by oil bodies proteins. 94
Figure 44. Time dependence of the incorporation of radioactivity from
[1-14C]dioleoylglycerol into TAG by oil bodies. 95
Figure 45. Effect of components of the assay mixture on the incorporation of
radioactivity from [1-14C]dioleoylglycerol into TAG by oil bodies. 96
Figure 46. Effect of different chemicals on the incorporation of radioactivity from
[1-14C]dioleoylglycerol into TAG by oil bodies. 96
Figure 47. Effect of PCMB on the incorporation of radioactivity from
[1-14C]dioleoylglycerol into TAG by oil bodies. 97
_14 Figure 48. Effect of addition of MgCl2 and PCMB on the incorporation of [1 C]1,2-
dioleoylglycerol to TAG by oil bodies. 98
Figure 49. Incorporation of radioactivity from [1_14C]1,2-dioleoylglycerol to TAG in
the absence and presence of different acyl-CoAs at 150 mM MgCl2. 98
Figure 50. Effect of pH on the incorporation of [1-14C]dioleoylglycerol into TAG by
oil bodies in the presence of 150 mM MgCl2. 99
Figure 51. Incorporation of radioactivity from [1-14C]oleoyl CoA into DAG in the
absence or the presence of sn-2-monooleoylglycerol. 100
Figure 52. Effect of concentration of sn-2-monooleoylglycerol on the incorporation of
radioactivity from [14C]oleoyl-CoA into DAG. 101
Figure 53. Time-dependence of incorporation of radioactivity from [14C]oleoyl-CoA
into DAG and TAG in the presence or absence of sn-2-monooleoylglycerol. 102
Figure 54. Effect of mode of MOG preparation on the incorporation of [14C]oleoyl CoA
VIII into DAG. 104
Figure 55. Effect of ethanol on the incorporation of radioactivity from [14C]oleoyl CoA
into DAG. 104
Figure 56. Incorporation of radioactivity from [14C]oleoyl-CoA into DAG in the
presence or absence of dioleoylglycerol during the time course. 105
Figure 57. Effect of the shift in MgCl2 concentration on incorporation of radioactivity
from [14C]oleoyl-CoA into TAG in the MAGAT assay, after the shift to
150 mM MgCl2. 106
Figure 58. Effect of pH on the release of radioactivity from [1_14C]1,2-dioleoylglycerol
into [1_14C]oleic acid. 107
Figure 59. Effect of pH on the release of radioactivity from [1_14C]1,2-dioleoylglycerol
into [1_14C]oleic acid. 108
Figure 60. Effect of temperature on the release of radioactivity from [1_14C]1,2-
dioleoylglycerol into [1_14C]oleic acid. 109
Figure 61. Time dependence of the release of radioactivity from [1_14C]1,2-
dioleoylglycerol into [1_14C]oleic acid. 110
Figure 62. Effect of protein content in assay on the release of radioactivity from
[1_14C]1,2-dioleoylglycerol into [1_14C]oleic acid. 110
Figure 63. The release of radioactivity from 1-stearoyl-2-[14C] arachidonyl-glycerol
into sn-2[14C] -MAG . 112
Figure 64. Release of radioactivity from [14C]trioleoylglycerol into FFA, DAG and
MAG showing the activity of TAG-lipase. 113
Figure 65. Changes in total fatty acids and arachidonic acid during recovery from
N-starvation at 24 °C and 12 °C. 114
Figure 66. Changes in chlorophyll and dry weight during recovery from N-starvation
IX at 24 °C and 12 °C. 115
Figure 67. Changes in the volumetric content of arachidonic acid in TAG, MGDG,
DGDG and SQDG during recovery from N-starvation at 24 °C and 12 °C. 118
Figure 68. Redistribution of radioactivity in chloroplastic lipids and TAG of P. incisa
during nitrogen replenishment to the nitrogen starved cells at 24 °C or 12 °C. 122
Figure 69. Redistribution of radioactivity in extraplastidial lipids and TAG of P. incisa
during nitrogen replenishment to the nitrogen starved cells at 24 °C or 12 °C. 123
Figure 70. Changes in the biomass content of TFA and AA during growth recovery in
control and SHAM treated cultures. 125
Figure 71. Changes in the culture contents of TFA and AA during growth recovery of
control and SHAM-pretreated cultures. 126
Figure 72. Changes in the contents of chlorophyll and dry wt in P. incisa cultures
during growth recovery in control and SHAM-pretreated cultures. 127
Figure 73. Changes in the volumetric content of 18:3ω3 during growth recovery in
control and SHAM-pretreated cultures. 127
Figure 74. Proposed pathways of TAG synthesis. 147
Figure 75. Outline of suggested pathways in the biosynthesis of MGDG in P. incisa. 152
X Tables
Table 1. Fatty acid composition of biomass and individual lipids of P. incisa in the
logarithmic and stationary cultures. 47
Table 2. Positional analysis of PC, PE and DGTS in the logarithmic culture
of P. incisa. 48
Table 3. Molecular species composition of PC in the logarithmic and stationary
cultures of P. incisa. 50
Table 4. Molecular species composition of PE in the logarithmic and stationary
cultures of P. incisa. 51
Table 5. Molecular species composition of DGTS in the logarithmic and stationary
cultures of P. incisa. 53
Table 6. Molecular species composition of TAG in the logarithmic and stationary
cultures of P. incisa. 55
Table 7. Effect of sethoxidim on fatty acid composition of nitrogen-starved culture
of P. incisa. 57
Table 8. Effect of sethoxidim on the accumulation of biomass and fatty acids
(µg/mg DW) of nitrogen-starved cultures of P. incisa. 57
Table 9. Effect of phosphate starvation on fatty acid composition and content
in P. incisa. 60
Table 10. Effect of phosphate starvation on distribution and fatty acid composition
of polar lipids in P. incisa. 61
Table 11. Effect of phosphate and nitrogen starvation on fatty acid composition and
content in P. incisa. 63
Table 12. Composition of homogenization buffers tested to control the lipolytic activity
during cell homogenization. 66
XI Table 13. Parameters characterizing various components of oil bodies. 69
Table 14. Distribution of neutral lipid (NL) lipid classes in oil bodies of P. incisa. 69
Table 15. Specific and total activity of [1-14C]oleoyl-CoA incorporation into TAG
in presence of cellular fractions of P. incisa. 72
Table 16. Specific activity of [1-14C]1,2-dioleoylglycerol incorporation into TAG in
the presence of cellular fractions of P. incisa. 81
Table 17. Specific activity of incorporation of radioactivity from [1-14C]1,2-
dioleoylglycerol into TAG in the presence of various cellular fractions
of P. incisa at 150 mM MgCl2. 91
Table 18. Effects of divalent metals and inhibitors on DAG lipase activity. 111
Table 19. Fatty acids composition of major lipid classes of P. incisa following
recovery from N-starvation. 116
Table 20. Changes in the molecular species distribution and content in MGDG of
P. incisa, following recovery from N-starvation. 120
Table 21. Distribution of fatty acids of P. incisa in the presence of SHAM and during
the recovery from SHAM. 124
Table 22. Fatty acid composition and content of selected mutant strains of P. incisa in
comparison to WT. 129
Table 23. Fatty acid composition of neutral and polar lipid fractions of selected
mutants of P. incisa in comparison to WT. 130
Table 24. Major fatty acid composition of selected polar lipid classes of WT and C18
desaturase mutant 107. 131
XII LIST OF ABBREVIATIONS AND SYMBOLS
∆X: A double bond in a fatty acid chain, positioned at carbon X, counted from the carboxyl end
∆XD: desaturase inserting a double bond at carbon X
AA: arachidonic acid
ACCase: acetyl-CoA carboxylase
ACP: acyl carrier protein
ACS: acyl-CoA synthase
ALA: α-linolenic acid
ATP: adenosine triphosphate
BSA: bovine serum albumin
C: carbon
CHES: 2-(N-cyclohexylamino)ethane sulfonic acid
CoA: coenzyme A
CPT: cholinephosphotransferase
DAG: diacylglycerol
DAGAT: diacylglycerol acyltransferase
DGDG: digalactosyldiacylglycerol
DGLA: dihomo-γ-linolenic acid
DGTA: diacylglycerol transacylase
DGTS: diacylglycerol trimethylhomoserine
DHA: docosahexaenoic acid
DIDS: 4,4'-diisothiocyanatostilbene-2,2'-disulfonate
DMSO: dimethyl sulfoxide
DTT: dithreitol
IX DW: dry weight
E: elongase
ECN: equivalent chain number
EDTA: ethylenediamine tetraacetic acid
EGTA: ethylene glycol-bis(β-aminoethylether)-N,N,N',N'-tetraacetic acid
ELSD: evaporative light scattering detector
EPT: ethanolamine phosphotransferase
ER: endoplasmatic reticulum
FAD: fatty acid desaturase (protein)
Fad: fatty acid desaturase (gene)
FAS: fatty acid synthase
FFA: free fatty acid
FID: flame ionization detector
G3P: glycerol-3-phosphate
G3PAT: glycerol-3-phosphate acyltransferase
GC: gas chromatography
GLA: γ-linolenic acid
GPAT: glycerol-3-phosphate acyltransferase
HEPES: 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid
HPLC: high performance liquid chromatography
LA: linoleic acid
LPA: lyso-phosphatidic acid
LPAAT: lyso-phosphatidic acid acyltransferase
LPCAT: lyso-phosphatidylcholine acyltransferase
X Lyso-PC: lyso-phosphatidylcholine
MAGAT: monoacylglycerol acyltransferase
ME: malic enzyme
MGDG: monogalactosyldiacylglycerol
MNNG: 1-methyl-3-nitro-nitrosoguanidine
MOG: monooleoylglycerol
N: nitrogen
NADPH: nicotinamide adenine dinucleotide phosphate
NEM: n-ethylmaleimide
NL: neutral lipids
PA: phosphatidic acid
PAP: phosphatidic acid phosphatase
PC: phosphatidylcholine;
PCMB: p-chloromercuribenzene sulfonic acid
PDAT: phospholipid-diacylglycerol acyltransferase
PE: phosphatidylethanolamine
PEP: phosphoenolpyruvate
PG: phosphatidylglycerol
PI: phosphatidylinositol
PL: polar lipids
PLA: phospholipase A
PLA2: phospholipase A2
PLAT: phospholipid acyltransferase
PLC: phospholipase C
PLD: phospholipase D
XI PMSF: phenylmethylsulphonylfluoride
PUFA: polyunsaturated fatty acids
SH-: sulfhydryl group
SHAM: salicylhydroxamic acid sn-: stereospecific position of fatty acid in the glycerol backbone of a lipid
SQDG: sulphoquinovosyldiacylglycerol
TAG: triacylglycerols
TFA: total fatty acids
TLC: thin-layer chromatography
Tris: hydroxymethylaminoethane
VLC-PUFA: very-long-chain polyunsaturated fatty acid
WT: wild type
ω: position of a double bond in a fatty acid chain, counted from the methyl end
XII 1. Introduction
Arachidonic acid (AA) is a very long chain-polyunsaturated fatty acid (VLC-PUFA) containing
20 carbon atoms and 4 double bonds at positions 5,8,11 and 14; the last double bond is at a distance of 6 carbons from the terminal methyl end of the fatty acid (20:4ω6).
The potentialities of PUFA in therapeutic and nutritional applications have increased the interest in them. AA is a major component of brain cell membranes and the predominant PUFA of human breast milk. In search of a source for AA production, the microalga Parietochloris incisa was found to contain remarkably high amount of triacylglycerols (TAG) that are rich in
AA. The main aim of this work was to study the unique mechanisms that lead to the accumulation of AA-rich TAG in this alga.
1.1 Polyunsaturated fatty acids
PUFAs are long chain fatty acids whose carbon chain has more than one methylene-interrupted double bond per molecule. The higher the unsaturation level of the fatty acid, the lower the melting point. A double bond, either of cis- or trans- configuration, is introduced between two adjacent carbon atoms of the aliphatic chain by the loss of two hydrogen atoms. The cis geometrical configuration produces a rigid kink of about 30 º in the chain whereas the trans forms have nearly the same conformation as that of the saturated chains. In biological systems, most double bonds are in the cis-configuration. PUFAs are divided into ω3 and ω6 families according to the distance of the last double bond from the terminal methyl group of the fatty acid,
3 in the former and 6 in the latter family. The ω6 fatty acids include linoleic acid (LA, 18:2ω6),
γ-linolenic acid (GLA, 18:3ω6), dihomo γ-linolenic acid (DGLA, 20:3ω6) and arachidonic acid
(AA, 20:4ω6), whereas α-linolenic acid (ALA, 18:3ω3), eicosapentaenoic acid (EPA, 20:5ω3), docosahexaenoic acid (DHA, 22:6ω3) are the major ω3-fatty acids.
1 1.2 Importance of PUFAs
PUFAs are one of the important components of the present day biomedical and nutraceutical
(food and nutritional products) world due to their specific therapeutic roles, especially in relation to certain clinical conditions, ranging from infant development to heart diseases.
PUFAs are essential components in higher eukaryotes that confer fluidity, flexibility and selective permeability to cellular membranes. PUFAs affect many cellular and physiological processes in both plants and animals, including cold adaptation and survival (Wada et al. 1990,
Miquel et al. 1993), modulation of ion channels (Meves 1994, Xiao et al. 2001), endocytosis/exocytosis (Schmidt et al. 1999), pollen formation, pathogen defense, chloroplast development in plants (Wallis and Browse 2002), and activities of membrane-associated enzymes that are sensitive to the biophysical properties of lipid membranes (Goldberg and
Zidovetzki 1997, Tsutsumi et al. 1995). In addition, PUFAs also regulate the expression of certain genes, including those coding for fatty acid synthase, nitric oxide synthase, sodium channel proteins and cholesterol-7-α-hydrolase, and thereby affect processes including fatty acid biosynthesis, cancer induction (Jiang et al. 1998) and cholesterol regulation (Clarke and Jump
1996).
All essential fatty acids in human are PUFAs, but not all PUFAs are essential fatty acids.
They are essential fatty acids (linoleic acid, α- and γ- linolenic acids, the major components of most plant lipids) in that they cannot be synthesized in animal tissues. On the other hand, as linoleic acid is almost always present in foods, it tends to be relatively abundant in animal tissues. In turn, these fatty acids are the biosynthetic precursors in animal systems of C20 and
C22 PUFAs, with three to six double bonds, via sequential desaturation and chain-elongation steps. The desaturases in animal tissues can insert a double bond only on the carboxyl side of an existing double bond. The metabolic pathways by which the parent essential fatty acids are
2 converted to very long chain PUFAs (VLC-PUFAs), within the human and animal body are summarized in Fig. 1.
ω6 PUFAs ω3 PUFAs
18:2ω6 18:3ω3 ▼ ∆6-desaturase ▼ 18:3ω6 18:4ω3 ▼ Elongation ▼ 20:3ω6 20:4ω3 ▼ ∆5-desaturase ▼ 20:4ω6 20:5ω3
Figure 1. Metabolic pathways of PUFAs in the human and animal body.
Studies on premature infants suggest that AA and DHA improve infant development, since they are the major acyl components of brain membrane phospholipids (Hansen et al. 1997).
Pre- and post-natal accumulation of DHA and AA is necessary for brain and retina development of infants (Crawford et al. 1997). Dietary VLC-PUFAs affects positively the growth and development of the infant and ameliorates the visual and cognitive functions, particularly in preterm infants. Likewise, VLC-PUFAs improves intestinal repair in severe protein-energy malnutrition; therefore, its qualitative and quantitative dietary supply should be considered (Gil et al. 2003). DHA and AA are transferred directly from mother to infant during the last intrauterine trimester through placenta, and following birth by feeding PUFA-rich human milk
(Agostoni et al 1994). The level of endogenous synthesis of DHA and AA in preterm and term infants is insufficient for optimal retinal and brain formation. Therefore, they require an external supply either by breast-feeding or by infant formula. Addition of AA and DHA to the infant formula has been approved by various health agencies (Boswell et al. 1996). FAO/WHO (Food and Agriculture Organization/World Health Organization) have indicated that the infant formula
3 shall contain: 40 mg DHA/day/kg of infant weight and 60 mg AA /day/ kg of infant weight
(FAO/WHO Nutrition Reviews 1995).
Another important function of VLC-PUFAs in mammals is the provision of precursors for short-lived eicosanoids, like prostaglandins, thromboxanes, and leukotrienes that regulate critical biological functions like fever, inflammation, vasodilation, blood pressure and pain
(Marx 1982, Funk 2001). VLC-PUFAs and their eicosanoids derivatives are fundamental for the maintenance of homeostasis and are linked to serious physiological and pathophysiological syndromes (Horrobin 1997b). Because the production of various classes of these molecules depends in part upon the availability of their PUFAs precursors in membrane phospholipids, modulation of PUFAs is a potential target of pharmaceuticals and nutraceuticals (Colquhoun
2001).
Data suggest that marine ω3-PUFA (e.g., ALA, DHA, EPA) have a beneficial impact in patients at high risk of sudden cardiac death (Christensen 2003), impair platelet aggregation and thromboxane formation, two major events in arterial thrombosis (Dyerberg et al. 1978, Mowat et al. 1997), reduce blood pressure, induce vasodilation and lower cholesterol levels (Horrobin
1997a), inhibit mental stress induced adrenal activation (Delarue et al. 2003). Similarly, ω6-
VLC-PUFA (AA) has shown its positive role in schizophrenia (Yao and Reddy 2002, Horrobin
1996) and in depression (Edwards et al. 1998). AA produced by Phospholipase A2 (PLA2) appears to play a crucial role throughout the generation of downstream-oxygenated products for the intracellular signaling mechanisms that mediate the regulation of parathyroid hormone secretion by parathyroid glands (Canalejo et al. 2003). In rodents, about 5% of brain AA and of
DHA are lost daily by metabolism and are replaced from dietary sources through the plasma demonstrating the active roles of PUFA in signal transduction and other processes (Rapoport
2003).
4 AA is an essential fatty acid, found in liver, brain, glandular organs, and depot fats of animals, and is a constituent of animal phosphatides. But, the discovery of proinflammatory and prothrombotic activity of leukotrienes and prostaglandins derived from AA (thromboxane A) has indicated that the presence of AA beyond certain level is harmful for health. On the other hand, studies have proved the beneficial effects of AA, especially if it can be kept as AA, and if the metabolism can be diverted away from eicosanoids to prostacyclins. Inclusion of ω3 PUFA in the diet has shown to decrease the synthesis of harmful AA-derived eicosanoids (James et al.
2000). Similarly, fish fats containing EPA were shown to impair platelet aggregation and thromboxane formation, two major events in arterial thrombosis. It is due to the fact that EPA replaces arachidonic acid in the platelet membrane phospholipids giving rise to prostaglandins and thromboxanes of 3-series, with very limited pro-aggregatory and vasoconstrictive activity
(Dyerberg et al. 1978). Hence, the ω6 to ω3 ratio of PUFA in the food is very important, and an optimal ratio 4 to 1 in diet is a major issue (Ristic and Ristic 2003). Hence, PUFAs have shown its obvious importance in dietary supplements, food additives, infant formulae and medicinal products such as, for cholesterol reduction, cardiovascular and various health indications.
In lower animals, such as insects and marine invertebrates, eicosanoids mediate metamorphosis, reproduction and host parasite interactions (Gerwick and Bernart 1993). In plants, C18 PUFAs are substrates for the synthesis of jasmonate and other oxylipins that serve as anti-infectives, wound response mediators, hormonal and chemotactic agents in plant defense and development (Wallis and Browse 2002, Feussner and Wasternack 2002). In Arabidopsis, trienoic fatty acids of thylakoid membrane lipids are required for low-temperature recovery from photoinhibition (Vijayan and Browse 2002). In this way, PUFA have an impact on cellular biochemical activities, transport processes, and cell-stimular responses.
5 1.3 Occurrence of PUFA-rich TAG
PUFAs occur throughout animal, plant, algae (Cohen et al. 1995), fungi (Totani and Oba 1988) and bacteria (Yazawa et al. 1988). They are found widely in many lipid compounds such as glycolipids, phospholipids, sphingolipids and lipoproteins in membranes and storage oils. Most of the higher plants do not produce PUFAs longer than C18 since they are devoid of the elongases and desaturases required for the production of C20-VLC-PUFAs. Mammals are unable to synthesize linoleic acid and α-linolenic acid, but they can further metabolize these fatty acids obtained from outer sources to produce LC-PUFA since they have ∆4, ∆5, and ∆6 desaturases.
However, lower plants and some microorganisms can produce both C18 and C20 PUFAs.
The conventional sources for C18 PUFAs (LA, ALA) are the seed plants, while marine fish and certain animal tissues are the source of VLC-PUFAs (AA, EPA, DHA). Although oils derived from fatty fish like herring, mackerel, sardine or salmon contain DHA and EPA, these oils are often inappropriate for human consumption or for inclusion in infant formula for its inconsistent quality, expensive refining process and limited fish stock. AA, widespread in the animal kingdom, can be isolated from lipids extracted from pig adrenal gland or pig liver and sardines as well; however, the yield of AA is only 0.2 % or lower (Ahern 1984), which is very difficult to industrialize.
Therefore, other sources, for example, microbial PUFA producers, are now actively being sought for commercial large-scale production of PUFA-containing oils. Some alternative sources for AA are fungi: Pythium, Mortierella; algae: Porphyridium; mosses: Rhytidialephus,
Brachythecium (Gill and Valivety 1997). Likewise, fungi: Mortierella, Pythium; algae: Monodus,
Navicula, Phaeodactylum, Porphyridium, Nannochloropsis; bacteria: Rhodopseudomonas,
Shewanella contain significant amounts of EPA. Similarly, fungi: Thraustochytrium,
Entomophthora; algae: Cryptocodinium, Gonyaulax; and bacteria: Rhodopseudomonas are potent sources of DHA. Companies like Rhone Pulenc and Martek Bioscience have set up large scale
6 production facilities of AA using Mortierella alpina, which consists of a very high lipid content
(6 g/L of DW) and with high proportions of AA, ranging from 40-60% of total fatty acids
(Yamada et al. 1987).
Regarding algae, many algal species are oleaginous, accumulating oil, mostly as TAG, but algal TAG are generally characterized by the presence of saturated and monounsaturated fatty acids, which are thought to serve as storage material (Henderson et al. 1990). Some other microalgae are able to produce TAG rich in EPA and AA but are not oleaginous (Eichenberger and Gribi 1997, Falk-Peterson et al. 1998, Makewicz et al. 1997). However, a rhodophyte
Porphyridium cruentum (Cohen 1990) was found to be capable of producing AA-rich TAG and its AA content reaching 2.5 % of its dry weight (Cohen et al. 1988, 1995).
Recently, in a search for PUFA-rich microalgae, an alga has been isolated from a snow patch in Japan, which contains more than 20% AA of its dry weight (Cohen et al. 2000). This alga has been identified as the chlorophyte, Parietochloris incisa (Trebouxiophyceae). Over 95% of the AA in the alga is deposited in TAG, constituting 60% of the fatty acids of these lipids. The major molecular species of the TAG is the very rare triarachidonylglycerol.
Lately, EPA and DHA were found to accumulate into TAG in stationary cells of the marine microalgae Nannochloropsis oculata (Eustigmatophyceae), diatom Phaeodactylum tricornutum , Thalassiosira pseudonana (Bacillariophyceae), and the Haptophyte Pavlova lutheri
(Poisson et al. 2002). DHA-containing lipid bodies were produced in the marine alga Isochrysis sp. to yeild up to 16 mg per liter of culture (Liu and Lin 2001).
1.4 Biosynthesis of TAG
Biosynthesis of membrane lipids in leaves and storage lipid, mostly TAG, in seeds have been thoroughly studied in higher plants (Browse and Sommerville 1994). However, little is known about the biosynthesis of algal PUFAs and PUFA-rich TAG. Basically, a TAG molecule is
7 formed by esterification of three fatty acid molecules to a glycerol molecule. Glycerol-3- phosphate (G3P) and acetyl CoA, the precursor of fatty acid synthesis, are obtained through glycolysis. The fatty acids are transferred to G3P sequentially in endoplasmic reticulum (ER) and in oil bodies by a class of specific enzymes, called acyltransferases to produce TAG. The fatty acids may be short-chained or long-chained, saturated, unsaturated or polyunsaturated. The fatty acids are synthesized de novo in chloroplast and desaturated by membrane-bound desaturases in the inner envelop of chloroplasts as well as in the endoplasmic reticulum (ER), being attached to different glyco- and phospholipids. Whereas, elongation of fatty acid chain longer than C18, takes place in the cytoplasm, while it is attached to coenzyme A (CoA). In this way, fatty acids synthesized de novo are converted to VLC-PUFA and subsequently, synthesis of PUFA-rich
TAG occur outside the chloroplast.
1.4.1. De novo synthesis of FA
The de novo synthesis of fatty acids is apparently the same in higher plants and algae, and is similar to that described for animals. In the plant kingdom, the de novo synthesis of C16 and C18 fatty acids takes place almost exclusively in the plastids (Ohlrogge et al. 1991, 1993).
Subsequent elongation steps to produce C20 or longer fatty acids are believed to occur in the ER.
Production of a C18 fatty acid from C2 acetyl-CoA and C3 malonyl-CoA precursors is catalyzed by seven enzymes, requiring acyl carrier protein (ACP) as cofactor (Browse and Somerville
1991, Ohlrogge et al. 1993) (Fig. 2).
Recently, cytosolic carbon sources such as glucose-6-phosphate, PEP, pyruvate and malate are seen as important substrates for plastidial fatty acid synthesis rather than acetate
(Rawsthorne 2002). The enzyme acetyl-CoA carboxylase, which catalyzes the synthesis of malonyl-CoA from acetyl CoA, and CO2 (the first committed step in the pathway) is supposed to be tightly regulated to determine the overall rate of fatty acid synthesis (Ohlrogge and Jaworski
8 1997). The final 2-carbon elongation from 16:0 to 18:0 occurs in plastids and requires 3- ketoacyl-ACP synthase II (Wu et al. 1994, Lightner et al. 1994). Some of the 16:0-ACP is released from the fatty acid synthesis machinery, but most molecules which are elongated to
18:0-ACP, are introduced with a first double bond at the ∆9 position by a stromal stearoyl-ACP desaturase (Lindqvist et al. 1996). The only known soluble desaturase is the plant stearoyl-ACP desaturase and it was cloned from castor (Ricinus communis) and cucumber (Cucumis sativus)
(Shanklin and Somerville 1991).
Figure 2. De novo fatty acid synthesis in higher plants and algae (Hildebrand et al. 2001).
9 Crystallization of plastid ∆9-18:0-ACP desaturase of castor revealed that it contains a di- iron cluster and a narrow channel adjacent to it (Lindqvist et al. 1996). The di-iron cluster catalyses the introduction of the double bond whereas the channel seems to represents the binding pocket for the stearic acid (18:0) portion of the substrate (Cahoon et al. 1997). Further desaturations involve complex lipid substrates such as galactolipids, phospholipids and in certain algae, betaine lipids.
1.4.2. Chloroplastic and extrachloroplastic lipids
16:0-ACP and 18:1-ACP, the major products of the de novo fatty acid synthesis in plastids, are used to make complex glycerolipids of cell membranes and oil bodies. Two acylation reactions, which transfer acyl residues from acyl-ACP (in chloroplast) or from acyl-CoA (in ER) to glycerol-3-phosphate, form the first glycerolipid, phosphatidic acid (PA). PA is converted to diacylglycerol (DAG) by hydrolysis of the phosphate group, to phosphatidylinositol (PI) or to phosphatidylglycerol (PG), while DAG can give rise to phospholipids, such as phosphatidylethanolamine (PE), phosphatidylcholine (PC), to galactolipids, such as monogalactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG), or to the sulfolipid sulfoquinovosyldiacylglycerol (SQDG) by esterification of the terminal –OH to respective polar head groups. Synthesis of complex lipids by modification or exchange of the head groups is also common (Browse and Sommerville 1991) (Fig. 3).
Browse and Sommerville (1994) have suggested that, in higher plants and algae, the 16:0- and 18:1-ACP products of the chloroplastic fatty acid synthesis may be used for the production of chloroplastic lipids by the ‘prokaryotic pathway’ inside the chloroplast, or may be exported to the cytoplasm as free fatty acids, which are later converted into acyl-CoA and incorporated into extrachloroplastic lipids in the ER by an independent set of acyltransferases (eukaryotic pathway) (Fig. 3).
10 In the prokaryotic scheme, PA initially incorporates 18:1 into the sn-1 position and 16:0 to the sn-2 position. PA then, is hydrolyzed to DAG, which ultimately gives rise to chloroplastic galactolipids, sulfolipid and phospholipid PG. These lipids can be constructed of molecular species of varying degrees of unsaturation and containing different combinations fatty acids.
Figure 3. Biosynthesis of membrane glycerolipids by prokaryotic and eukaryotic pathways in
Arabidopsis leaves (Browse and Sommerville 1994).
In the eukaryotic pathway, PA is synthesized by transferring 18:1 or 16:0 to the sn-1 position and 18:1 to the sn-2 position resulting in the 18:1/18:1 and/or 16:0/18:1 molecular species. PA is later transformed into lipids of extrachloroplastic membranes, such as PC, PE and
PI. Phospholipids can be further desaturated, reimported to the chloroplast as 18:2/18:2 DAG and finally produce chloroplastic galactolipids and sulfolipid, which can be further desaturated in the chloroplast (Lemieux et al. 1990, Miquel and Browse 1992, Browse et al. 1993).
11 In some higher plants, including Arabidopsis and spinach, the two pathways contribute equally to chloroplast lipid synthesis. However, in many other higher plants, only PG is produced by the prokaryotic pathway, and the remaining chloroplastic lipids are synthesized completely by the eukaryotic pathway.
1.4.3. Membrane desaturases
The fatty acids produced by de novo synthesis are further desaturated while they are attached to the complex lipids, either by chloroplastic desaturases or by desaturases of the endoplasmic reticulum (ER). In Arabidopsis, the mechanisms and regulation of the desaturases were revealed through the characterization of seven classes of mutants, each one deficient in a specific desaturation step (Browse and Sommerville 1991; Somerville and Browse 1991) (Fig. 3). The chloroplastic desaturases encoded by fad4 and fad5 are highly substrate specific while the products of genes, fad6, fad7 and fad8 have no apparent specificity for the chain length, the position of the fatty acids and the nature of the lipid head group. The gene product of fad4
(FAD4) desaturates the ∆3 position of 16:0 attached to the sn-2 position of PG and FAD5 adds
∆7 double bond in 16:1 of MGDG and DGDG (Browse et al. 1985, Kunst et al. 1989). FAD6 desaturates 16:1/18:1 whereas two isozymes FAD7 and FAD8 are responsible for the desaturation of 16:2/18:2 molecular species of the major chloroplast lipids. The ER desaturases encoded by fad2 and fad3, act on 18:1 and 18:2 of PC, respectively, and probably also on other phospholipids. Thus, in plants, generally, 18:1 produced by de novo synthesis is incorporated into glycerolipids of the chloroplasts and ER, and membrane-bound desaturases in these organelles add a second double bond at ∆12 to form 18:2, and a third double bond at the ∆15 position to form 18:3ω3, which are essential fatty acids as substrates for the synthesis of C20 PUFA
(Somerville and Browse 1996).
12 The plant glycerolipid desaturases not only produce the C18 PUFAs, essential for membrane function in plants, but also provide 18:2 and 18:3ω3 of seed oils that are the main source of these fatty acids in the human diet (Wallis et al. 2002).
1.4.4. Biosynthesis of C20 PUFA in algae
The biosynthesis of PUFAs occurs outside the plastid. The pathways for the synthesis of C20
PUFA, such as AA (20:4ω6) and EPA (20:5ω3) involve alternating fatty acid desaturation and elongation reactions. The membrane-bound desaturase insert double bonds at specific carbon atoms in the fatty acid chain and the fatty acid elongation system elongates the precursors in two- carbon increments (Wallis et al. 2002).
Although the biosynthesis of C18 PUFAs has been studied in detail, very little is known about that of C20 PUFA in algae (Bigogno et al. 2002a). In algae, the pathways for the production of C18 PUFA are similar to that of higher plants (Norman et al. 1985). However, the biosynthesis of C20 PUFA appears to be more complex. Multiple pathways have been reported for the biosynthesis of AA, EPA and DHA, resulting from the differences in the order of elongation and desaturation steps in different organisms (Cohen et al. 1995, Bigogno et al. 2002,
Wallis et al. 2002, Baoxiu et al. 2004, Sayanova and Napier 2004).
Biosynthesis of AA and EPA in the red microalga Porphyridium cruentum proceeds by stepwise desaturation of oleate to 18:3ω6, which is then elongated to 20:3ω6 and further desaturated to AA and EPA (Sequence 1) (Shiran et al.1996, Khozin et al. 1997). In this alga,
AA is synthesized in PC, re-imported to chloroplast and esterified to MGDG, where it is further desaturated to EPA. However, in Euglena gracilis (Euglenophyceae), Nichols and Appleby
(1969) reported the elongation of 18:2ω6 to 20:2ω6 (eicosadienoic acid) precedes the desaturations to 20:3ω6, and to AA (Sequence 2).
13 ∆12D ∆6D E ∆5D ω3D 18:1 → 18:2ω6 → 18:3ω6→ 20:3ω6 → 20:4ω6 → 20:5ω3 [1]
∆12D E ∆8D ∆5D 18:1 → 18:2ω6 → 20:2ω6 → 20:3ω6 → 20:4ω6 [2]
In the diatom, Phaeodactylum tricornutum, EPA is synthesized through four different routes, starting from 18:1ω9, utilizing PC as the lipid carrier for the desaturations (Arao et al.
1994, Arao and Yamada 1994). In one pathway (Sequence 3), 18:1 is successively desaturated via 18:2ω6 and 18:3ω3, to 18:4ω3 (octadecatetraenoic acid), elongated to 20:4ω3
(eicosatetraenoic acid), subsequently re-linked to PC and desaturated to 20:5ω3. In the second pathway (Sequence 4), 18:3ω3 is elongated to 20:3ω3 before being desaturated to 20:4ω3 and
20:5ω3. In the mixed ω6/ω3 pathway, 18:2ω6 is desaturated to 18:3ω6, which is ω3 desaturated to 18:4ω3 and elongated to 20:4ω3 before the final desaturation to 20:5ω3 (Sequence 5).
∆15D ∆6D E ∆5D 18:2ω6 → 18:3ω3 → 18:4ω3 → 20:4ω3 → 20:5ω3 [3]
∆15D E ∆8D ∆5D 18:2ω6 → 18:3ω3 → 20:3ω3 → 20:4ω3 → 20:5ω3 [4]
∆6D ∆6D E ∆5D 18:2ω6 → 18:3ω6 → 18:4ω3 → 20:4ω3 →20:5ω3 [5]
The biosynthesis of another important C20 PUFA, DHA (22:6ω3) may follow two pathways. 22:5ω3, produced by elongation of 20:5ω3 is desaturated to produce 22:6ω3 by ∆4 desaturase in the marine protist Thraustochytrium spp. and the freshwater species Euglena
(Meyer et al. 2003, Qi et al. 2002). In the so-called "Sprecher" pathway in mammals, 24:5ω3, produced by two consecutive C2 elongation cycles in 20:5ω3, undergoes ∆6 desaturation to
14 produce 24:6ω3 and one cycle of C2-shorteninig via β-oxidation in the peroxisome to yield DHA
(Sprecher et al. 1995).
The types of lipids, to which the fatty acyl groups are attached prior to undergoing the desaturations, seem to be species-specific (Bigogno et al. 2002). In the green alga,
Chlamydomonas reinhardtii, the ∆12, ∆15, and ∆6 desaturations on C18 fatty acids occur while they are esterified to the sn-2 position of diacylglycerol (N,N,N)-trimethylhomoserine (DGTS) or
PE (Giroud and Eichenberger 1989). In the cryptomonad, Chroomonas salina, C18 PUFAs are located predominantly in galactolipids, whereas EPA and DHA are found almost exclusively in phospholipids (Vogel and Eichenberger 1992). Schneider and Roessler (1994) suggested that in the eustigmatophyte, Nannochloropsis, PC and PE were the lipid carriers for the desaturations of
C18 and C20 fatty acids, respectively. In the rhodophyte P. cruentum, PC is the major substrate for the ∆12, ∆6, and ∆5 desaturations, whereas the final ∆17 desaturation of 20:4 to 20:5 is predominantly chloroplastic, involving eukaryotic-like (MGDG) and prokaryotic-like (MGDG and DGDG) molecular species (Khozin et al. 1997). In the freshwater eustigmatophyte Monodus subterraneus, PC is mostly involved in the desaturation of C18 fatty acids, whereas PE and
DGTS are involved in the desaturation of C20 PUFAs, leading to the synthesis of EPA (Khozin-
Goldberg 2002b). In P. incisa, PC and DGTS are involved in the ∆12 and, subsequently, the ∆6 desaturations of oleic acid, whereas PE and PC are the major substrate for the ∆5 desaturation of
20:3ω6 to AA (Bigogno et al. 2002a).
Starting from acetyl-CoA, the synthesis of 22:6 requires approximately 30 distinct enzyme activities and nearly 70 reactions, including four repetitive steps of the fatty acid synthesis cycle and the energetically demanding desaturase reactions (Wallis et al. 2002).
Recently, a novel PUFA biosynthetic mechanism was found in marine bacteria, Shewanella sp.,
Moritella marinus and in the marine alga Schizochytrium sp. that does not require the fatty acid desaturase/elongase system, instead uses a polyketide synthase to produce 20:5ω3 and 22:6ω3
15 (Metz et al. 2001). The polyketide synthase contain several identifiable domains, which are homologous to FAS enzymes (Wallis et al. 2002).
Once the PUFA are formed, they can remain linked to the membrane lipids, transferred to other membrane lipids, or accumulated in TAG. In Phaeodactylum tricornutum, EPA is exported from PC to another membranal lipid, MGDG (Arao et al 1994, Arao and Yamada 1994). In M. subterraneus, PE and DGTS transfer EPA to the eukaryotic-like and prokaryotic-like molecular species of MGDG, respectively (Khozin-Goldberg 2002b). AA is transported from PC to TAG in
Porphyridium cruentum (Khozin et al. 1997) whereas, in Chroomonas salina, 18:3ω3 is transferred from DGTS to TAG (Henderson and Mackinlay 1992). In P. incisa, AA is formed in
PE and PC, and mostly transferred to TAG in the stationary phase (Bigogno et al. 2002a). Over
90% of the cellular AA was found to accumulate in TAG during the growth under nitrogen starvation conditions (Khozin-Goldberg et al. 2002a). However, AA was distributed to all the polar lipids, including chloroplastic lipids.
1.4.5. Partitioning of FA to TAG
The mechanism of TAG synthesis in developing oilseeds is less understood than that of membrane lipid synthesis in leaves (Browse and Somerville 1994). A scheme developed to describe the lipid metabolism of developing Arabidopsis seeds (Slack et al. 1985, Stymne and
Stobart 1987, Stymne et al. 1987) is suitable also for many other oilseed species. The major reactions of TAG synthesis in seeds are shown in Fig. 4.
TAG is synthesized via the so-called Kennedy pathway using the acyl-CoA pool, deriving from the de novo synthesis in plastids, through glycerol-3-phosphate (G3P), lyso-phosphatidic acid (Lyso-PA), phosphatidic acid (PA) and diacylglycerol (DAG) (Figs 4 and 5) (Kennedy
1961). This pathway apparently operates in the ER and the TAG accumulates in structures known as oil bodies, which are surrounded by a phospholipid membrane monolayer rather than
16 the usual bilayer. The Kennedy pathway involves 4 enzymatic steps involving 3 acyltransferases and a phosphatase. The first and second acyltransferase reactions transfer fatty acids from acyl-
CoAs to the sn-1 and the sn-2 positions of glycerol to form PA, and phosphatase convert PA to
DAG. These enzymes are common also to membrane lipid synthesis.
Figure 4. Various pathways of TAG biosynthesis in plants (Hildebrand et al. 2001)
The final step of TAG synthesis is catalyzed by the third acyltransferase, diacylglycerol acyltransferase (DAGAT) (Moore 1982), which esterifies a fatty acid at the sn-3 position, and it is the enzyme devoted only to the synthesis of TAG (Fig. 5).
17
Figure 5. The Kennedy pathway of TAG biosynthesis. P - Phosphate, A - Acyl group, GPAT-
Glycerol-3-phosphate acyltransferase, LPAAT - Lysophosphatidic acid acyltransferase, PAP -
Phosphatidic acid phosphatase, DAGAT - Diacylglycerolacyltransferase.
In addition to the Kennedy pathway, several pathways of the biosynthesis of TAG, containing unsaturated fatty acids, have been elucidated in oil seeds. Since the extraplastidial desaturations of fatty acids occurs mainly in PC, the hydrolysis of the phosphocholine headgroup of PC by the reversible enzyme cholinephosphotransferase (CPT) produces DAG, containing unsaturated fatty acids (Slack et al. 1985). The obtained DAG can be converted to TAG by
DAGAT (Fig. 6). Kamisaka et al. (1999) also have shown that PA and PC are converted to DAG and then to TAG in the membrane and lipid body fraction in the fungus Mortierella ramanniana.
A reversible enzyme lyso-PC acyltransferase (LPCAT), esterifying and releasing fatty acids at the sn-2 position of PC was found in the microsomal preparations of developing safflower cotyledons (Stymne and Stobart 1984). LPCAT transfers oleate from oleoyl-CoA to the sn-2 position of PC where it undergoes desaturation to linoleate and finally released to the acyl-CoA pool, enriching it with PUFAs for TAG assembly. LPCAT and CPT work together to increase
C18 PUFAs in TAG of oil seeds (Stymne and Stobart 1987).
Figure 6. Pathway for unsaturated TAG biosynthesis. PCh-Phosphocholine head group,
LPCAT-Lyso-phosphatidylcholine acyltransferase, CPT-Choline phosphotransferase, A-
Acyl group, DAGAT-Diacylglycerol acyltransferase.
18 Recently, Stobart et al. (1997) and Fraser et al. (2000) demonstrated a novel and alternative pathway by which two molecules of DAG disproportionate to produce TAG and monoacylglycerol (MAG) (Fig. 7). This reaction does not require DAGAT and does not utilize acyl-CoA.
Figure 7. DAG-DAG transacylation pathway for TAG biosynthesis. DGTA- Diacylglycerol transacylase.
Waters et al. (2003) have recently discovered that the sn-2 monounsaturated monoacylglycerols (MAG), the product of the transacylation reaction, is recycled to produce
DAG by monoacylglycerol acyltransferase (MAGAT) in developing cotyledons of sunflower.
It has been recently demonstrated that in plants and yeast, an enzyme called phospholipid:diacylglycerol acyltransferase, PDAT, can catalyze the transfer of acyl group from the sn-2 position of major phospholipids to DAG , forming TAG and lyso-phospholipids without using acyl-CoA (Dahlqvist et al. 2000) (Fig. 8). In different plants, PDAT showed different selectivities for acyl groups, which are transferred into TAG. In microsomal preparations from developing castor bean (Ricinus communis) seeds, which accumulate TAG with about 85% of an unusual hydroxy fatty acid, ricinoleate (12-hydroxy-octadeca-9-enoate), PDAT activity showed high specificity for ricinoleoyl groups (Dahlqvist et al., 2000). The activity of yeast PDAT was shown to be dependent on the type of polar head-group of the donor lipid, the acyl group transferred, and the acyl chains of the acceptor molecule DAG (Dahlqvist et al. 2000).
19
Figure 8. PC-DAG transacylation pathway of TAG biosynthesis. PDAT- phospholipid diacylglycerol acyltransferase
Any oleoyl substrate entering PC by the activity of CPT is readily desaturated and the
PUFA products are incorporated into TAG (Stobart et al. 1997). Transacylation, therefore, could account for the further enrichment of preformed TAG with PUFA as observed in vivo (Garces et al. 1994, Dahlqvist et al. 2000).
Although TAG biosynthesis in general has been extensively studied in various organisms, little is known about the biosynthetic pathways for TAG molecular species (Pillai et al. 2002).
Stereospecific analysis of TAG in various organisms has shown that fatty acids are not randomly distributed at the sn-1,2,3 positions (Padley et al. 1994), suggesting mechanisms distributing appropriate fatty acids to each position.
In microalgae, the fatty acid composition of TAG generally reflects that of the polar lipids from which they derive. Comparison of the fatty acid composition and positional distribution in the various molecular species of both polar lipids and TAG could indicate which of these lipids is a likely contributor to TAG biosynthesis. Furthermore, it could indicate whether distinct acyl moieties or DAG are contributed. For example, nitrogen starvation affects an increase in the proportion of AA in TAG of Porphyridium cruentum from 20-31% (of fatty acids). At the same time, the proportion of AA in PC increases from 46 to 61% (Cohen 1992). In Parietochloris incisa however, under N-starvation, the proportion of AA in TAG is as high as 50-60%, in comparison to 25% or less, in the phospholipids. Possibly, the fatty acids of TAG arrive from distinct positions, presumably the sn-2 position of the specific polar lipid.
20 Different pathways of TAG biosynthesis have been found to proceed together at the same time. In mice, inactivation of the DAGAT encoding gene resulted in a lack of TAG synthesis in most of tissues, however serum TAG level and TAG in white adipose tissues were normal, indicating the existence of alternative pathways of TAG synthesis (Buhman et al. 2001).
Different importance and activity of enzymes of TAG synthesis during different growth phases were observed in yeast. In yeast, the PDAT-pathway is the most important for TAG synthesis during active cell division (Oelkers et al. 2002) whereas the DAGAT-pathway is dominant in the stationary growth phase when the cells are storing significant amounts of TAG (Stahl et al.
2004). In oil seeds, producing unusual fatty acids, different enzymes contribute to the final acyl composition of TAG (Dahlqvist et al. 2000).
It is also important to determine the relative contributions of the Kennedy pathway and the cytoplasmic lipids in the construction of PUFA-rich TAG. This can be studied by using inhibitors, which are specific for certain pathways. Study of the molecular species of TAG in the oleaginous fungus, Mortierella ramanniana showed the existence of several pathways for different TAG molecular species synthesis, which exhibited different sensitivities to low temperature and inhibitors of lipid metabolism (Pillai et al. 2002). The use of inhibitors of de novo fatty acid synthesis, such as sodium azide and cerulenin suggested that biosynthetic pathway of triolein (OOO) is different from that of palmitic acid containing molecular species
(OPP), which are more readily synthesized through the de novo pathway.
The time-course of PUFA production and its partition into TAG are different in various marine microalgae. A much higher percentage of total cellular EPA production and partition to
TAG upon the transition to stationary phase, was seen in Nannochloropsis oculata
(Eustigmatophyceae) when nitrate level in the culture fell to a minimum level. However, in
Phaeodactylum tricornutum cell division continued for about 100 h in this condition and TFA increased strongly only after this period (Tonon et al. 2002). Although P. tricornutum produces
21 both EPA and DHA, only EPA (40% of total EPA) was partitioned into TAG, suggesting a substrate specificity of the acyltransferases. Both Thalassiosira pseudonana (Bacillariophyceae) and the haptophyte Pavlova lutheri produce EPA and DHA and partition these to TAGs during the stationary phase of growth (Tonon et al. 2002).
TAG production is a mere extension of the glycerolipid pathway essential for the biosynthesis of membrane lipids. Both processes occur simultaneously in the cells of developing oil-storage tissues and share substrates (DAG, acyl-CoA) and common location, ER. However, plants producing highly saturated or unusual fatty acids (medium and long chains or oxygenated) accumulate these fatty acids in TAG, leaving only traces in membranal phospholipids (Voelker and Kinney, 2001). Mature Arabidopsis seeds expressing a Ricinus hydroxylase, contained 17% hydroxylated fatty acids in their TAG but only about 1% in their seed phospholipid fraction
(Broun and Somerville 1997). The channeling of these deleterious-to-membrane fatty acids to
TAG could be achieved either by a spatial separation of TAG and membrane biosynthesis in ER
(Vogel and Browse 1996, Frentzen 1993) at the stage of DAG utilization by DAGAT and CPT, or there may be a selective and efficient removal of unusual fatty acids from phospholipids by a more efficient editing mechanism (Dahlqvist et al. 2000, Stahl et al. 1995, Stahl et al. 1998).
Plants containing PUFAs in polar lipids and in TAG should have a mechanism of rapid transfer between the two ER domains.
Due to the presence of C20 fatty acids, the acyl-CoA pool in the seed of some plants has a much more complex collection of fatty acyl groups than in leaf mesophyll cells. Membrane lipids are composed mainly of five fatty acids, palmitate (16:0), stearate (18:0), oleate (18:1), lenoleate
(18:2), and linolenate (18:3). However, TAGs are known to accumulate >300 fatty acids/fatty acid derivatives. This is apparently due to a combination of unique fatty acid biosynthetic/modification enzymes in oil accumulating tissues and specificity of acyltransferases directing these unusual fatty acids from the acyl-CoA pool into TAG.
22 1.4.6. Acyltransferases of TAG biosynthetic pathway
Membrane-bound glycerol-3-phosphate acyltransferase (GPAT) initiates the TAG synthesis by transferring the acyl chain from CoA to the sn-1 position of glycerol-3-phosphate, forming lysophosphatidic acid (LPA). This enzyme appears to have a low selectivity for acyl chains
(Frentzen, 1998) and even prefers saturated CoA substrates (Cao and Huang 1987).
Lysophosphatidic acid acyltransferase (LPAAT) catalyzes the transfer of acyl-chain from the
CoA ester to the sn-2 position of LPAAT, creating phosphatidic acid. In plants, this enzyme prefers unsaturated acyl chains (Frentzen, 1998). In safflower, LPAAT preferentially incorporates 18:2 and 18:3 moieties into the sn-2 position of PA (Bafor et al. 1990). In genetically modified Canola, producing short chain 12:0 fatty acid in seeds, the laurate (12:0) was deposited almost exclusively at the sn-1 and the sn-3 positions (75% laurate) of TAG and represented only 5% at the sn-2 (Voelker et al. 1996) indicating that the canola LPAAT strongly discriminated against laurate, but GPAT and DAGAT accommodated the novel substrate.
However, in high saturate producers, the sn-2 position of TAG often selects medium chains preferentially. In coconut oil, the sn-2 position always consists of 80% of the TAG’s laurate
(Padley et al. 1994, Wiberg et al. 1997). Similarly, in Cuphea lanceolata, which is rich in C10:0, the sn-2 position of TAG is composed of 97% C10:0, whereas the sn-1 and the sn-3 positions together are composed of only 76% of this fatty acid (Bafor et al. 1990).
Ultimately, the fatty acids composition of TAG depends upon the composition of the acyl-CoA pool in the cytoplasm. Acyl-CoA synthases in Brassica and castor bean are responsible for the unique composition of the acyl-CoA pools that include very long chain fatty acids in
Brassica and ricinoleic acid in castor bean (Ichihara et al. 1997). In Cuphea, it was shown that the acyl-CoA pool composition influences the selectivity of DAGAT (Bafor and Stymne 1992).
DAGAT, the unique enzyme of TAG synthesis might be an important step in control of
TAG synthesis (Hobbs et al. 1999). DAGAT is a membrane-bound enzyme, which is mainly
23 thought to be located in the ER. However, DAGAT activity has also been reported in oil bodies.
In the oleaginous fungus, Mortierella ramanniana, DAGAT activity in the lipid body fraction, was much higher than that in the membrane fraction, in terms of both total activity and specific activity (Kamisaka and Nakahara 1994). In contrast to seeds, leaves contain most of their
DAGAT activity in the envelope membranes of the chloroplasts (Martin and Wilson 1984). TAG usually accumulates in low amounts in plastids but, under stress conditions, the DAGAT converts DAG, released by degradation of the membrane lipids, into TAG (Browse et al.1988,
Sakaki et al. 1990).
DAGAT shows broad acyl-CoA selectivities in some organisms and it shows some degree of preferential selectivity of certain acyl-CoA in other cases. In vitro assays with oilseeds suggest that microsomal GPAT and DAGAT usually display broad acyl–CoA selectivities and even prefer saturated CoA substrates (Cao and Huang 1987, Frentzen 1998). Preference to saturates (80% saturates at the sn-1 and the sn-3) were shown also in vivo in transgenic canola, when FA production was engineered to medium-chain production (Voelker et al. 1996, Wiberg et al. 2000). In Mortierella ramanniana, palmitoyl-CoA was more readily incorporated over oleoyl-CoA in the TAG by DAGAT (Pillai et al. 1998) whereas GPAT and LPAAT had a strong preference toward oleoyl-CoA as a substrate. In maturing safflower seeds, DAGAT showed no strict selectivity for acyl-CoA (Ichihara et al. 1988). Thus, the acyl-composition at the sn-1 and the sn-3 positions in TAGs depends on the acyl composition of the available substrates rather than acyltransferase specificity (Voelker and Kinney 2001). In contrast, the microsomal DAGAT of Cuphea procumbens showed pronounced selectivities for caproyl (10:0) containing substrates
(Wiberg et al. 1994).
Some seeds contain high amount of unusual fatty acids (medium chain, very long chain, epoxy, hydroxy) in the TAGs but are nearly excluded from membrane lipids. The channeling of these unusual fatty acids into TAG is probably achieved through the selectivity of
24 acyltransferases, including DAGAT (Frentzen 1993). The castor bean enzyme showed selectivities for substrates containing ricinoleate (18:1-OH) (Wiberg et al. 1994).
1.5. Factors affecting the biosynthesis of TAG
The triacylglycerol biosynthesis in algae has been found to be affected by various growth conditions such as light, temperature, nitrogen and carbon availability. The increase in lipid accumulation has been observed under the growth-limiting conditions at the expense of protein and carbohydrate production. The highest levels of TAGs are typically found at the stationary phase when the nutrients are depleted (Iwamoto et al. 1955, Klyachko-Gurvich et al. 1967).
Optimal conditions for TAG production may vary from species to species and may be different from the optimal growth condition.
In oilseed rape, diversion of photosynthate from starch synthesis to the precursors of fatty acid synthesis was observed during the embryonic development (Eastmond and Rawsthrone
2000). Net increase in carbon flux to lipid synthesis, and decrease to starch production, accompanied by changes in the plastids ability to import metabolites via transporter proteins, and an increase in uptake and utilization of pyruvate were seen, while there was a decrease in the uptake of the another metabolite, glucose-6-phosphate and its utilization for fatty acid synthesis.
In Arabidopsis, fatty acids of thylakoid galactolipids are channeled to the TAG synthesis in chloroplast during leaf senescence by DGAT1, which has high sequence similarity with sterol acyltransferase (Kaup et al. 2002).
1.5.1. Temperature
The growth temperature is one of the most important factors for biomass productivity, lipid synthesis and unsaturation of fatty acids. The optimum temperature for maximum lipid production is different in different organisms. The lipid content of microalga Ochromonas danica
25 for instance, increased from 39 to 53% as the temperature was raised from 15 °C to 30 °C
(Aaronson 1973) whereas in Nannochloropsis sp., the FA content was higher at 18 °C than at 32
°C (Sukenik et al. 1989). The optimum temperature for the growth and for the synthesis of lipid may or may not be the same. When the green alga Botryococcus was grown at the supra-optimal temperature, the synthesis of all intracellular lipids, except for triacylglycerides, were considerably inhibited and the content of trienoic fatty acids was significantly lower than at the optimal cultivation temperature (Kalacheva et al. 2002). In the fungus, Pythium irregulare, the maximum growth rate was obtained at 25 °C, while maximum EPA production was obtained at
12 °C (Stinson et al. 1991). Temperature changes the lipid composition in most organisms. The lipid composition was affected by growth temperature in Anacystis nidulans, but not in
Anabaena variabilis (Sato et al. 1979).
It has been well established that the decrease in the growth temperature results in an increase in the degree of fatty acids unsaturation to maintain membrane fluidity at lower temperatures. A temperature shift from 38 °C to 22 °C stimulated the desaturation of C16 and
C18 FAs in DGDG in the cyanobacterium, Anabaena variabilis (Sato and Murata 1980). In
Motierella alpina, total PUFA, EPA and AA production increased by 12, 84.4 and 46.1%, respectively, when the culture temperature was shifted from 20 °C to 12 °C (Jang et al. 2000).
However , PUFA-rich microalgae retain a high level of PUFA even at elevated temperatures.
When P. cruentum was grown under supra-optimal temperatures, the proportion of EPA was decreased whereas that of arachidonic acid was increased (Cohen et al. 1988).
1.5.2. Light
Light intensity and light availability affects not only the growth rate of an alga, but also the pigments, the structure, and the composition of the photosynthetic apparatus. Under high light intensities, a reduction of thylakoid membranes, an increase in starch granules, and a large
26 accumulation of lipid droplets, which contain triacylglycerols (63%) polar lipids, (12%) and carotene (35%) were observed in the unicellular alga Dunaliella bardawil (Rabbani et al. 1998).
However a decrease in the FA content of biomass was reported under high light in
Phaeodactylum tricornutum (Fernandez et al. 2000). Higher light intensities have been reported to increase the PUFA levels in Phaeodactylum tricornutum (Fernandez et al. 2000),
Porphyridium cruentum (Cohen et al. 1988) and in Fucus serratus (Smith and Harwood 1984).
However, opposite effect have been reported in some other algal species, Nannochloropsis sp
(Sukenik et al. 1989), Monodus subterraneus (Cohen 1994), and Isochrysis galbana (Harrison et al. 1990).
Light-dark cycles may also have an effect on fatty acid composition. Sicko-Goad et al.
(1988) showed that in the diatom Cyclotella meneghiniana, the levels of EPA and other PUFAs were lowest in the early part of the light period and highest in the dark. They suggested that EPA accumulation precedes cell division, which takes place at the end of the dark period or at the beginning of the light period.
1.5.3. Nutrient deprivation
The availability of nitrogen during the growth was shown to affect, not only the growth rate, but also the lipid content and composition (Shifrin and Chisholm 1981). TAG accumulation, up to
70% of dry weight was demonstrated in many algae (Shifrin and Chisholm 1981, Roessler,
1990). Imposing nitrogen limitation when light is in excess, results in cessation of growth. Since photosynthetic fixation of carbon continues, the cellular ratio of C/N is thereby increased
(Mayzaud et al. 1989) and energy could be channeled into production of non-nitrogenous reserve materials such as TAG, which serve as a sink for photosynthetically fixed carbon. N-starvation adversely affects the pigments and protein contents of the cells. In the marine diatom,
Phaeodactylum tricornutum, during N-starvation, cell nitrogen and chlorophyll a decreased,
27 mainly as a consequence of continued cell division, keeping the cell volume constant, which led to a decrease in the rate of photosynthesis and respiration, while carbon and lipid increased
(Larson and Rees 1996). N-starvation in Dunaliella tertiolecta (Butcher) was characterized by slow reduction in cell chlorophyll, protein content and chlorophyll/carotenoid ratio, and a decline in photosynthetic capacity and maximum quantum yield of photosynthesis (Fv/Fm) (Young and
Beardall 2003).
A high C/N ratio favors lipid accumulation, which is triggered by nitrogen depletion in the culture (Ratledge 1989). Shifrin and Chisholm (1981) reported a 130-320% increase in oil content under nitrogen-deficient conditions in fifteen chlorophycean strains. Similarly, in the red microalga Porphyridium cruentum, N-starvation stimulated an additional accumulation of TAG
(Cohen 1990). Moreover, under nitrogen-starvation conditions, the amount of TAG increased to over 90% of total acyl lipids and AA constituted up to 60% of the fatty acids of TAG in the freshwater alga Parietochloris incisa (Khozin-Goldberg et al. 2002a). However, during heterotrophic cultivation of the green microalga Chlorella sorokiniana, cellular lipid content was minimal at a C/N ratio of 20 and increased at both higher and lower C/N ratios (Chen and Johns
1991).
In heterotrophic oleaginous organisms, the exhaustion of nitrogen from the culture medium triggers lipid accumulation in the cells, but glucose continues to be assimilated. N- starvation decreases AMP within the cells, which slows or stop the activity of isocitrate dehydrogenase in the mitochondria, resulting in the accumulation of cirtrate. The citrate is cleaved to acetyl-CoA, the substrate for FA synthesis by ATP:citrate lyase, which is found only in oleaginous species in the cytosol. The extent of lipid accumulation is considered to be controlled by the activity of malic enzyme (ME), which acts as the sole source of NADPH for fatty acid synthase (FAS). If the ME is inhibited, or genetically disabled, then lipid accumulation is very low. The ME could be physically attached to FAS as part of the lipogenic metabolon. ME
28 activity correlates closely with lipid accumulation in two filamentous fungi, Mucor circinelloides and Mortierella alpina (Ratledge 2002).
While nutrient depletion often results in an enhancement of triglycerides, in many cases, nitrogen (Ben Amoz et al. 1985) and silicon (Engel et al. 2000), depletion resulted in an increase in short chain saturated fatty acids and a decrease in long chain polyunsaturated fatty acids. In
Chlorella sorokiniana, a high proportion of unsaturated fatty acids were observed in a low C/N ratio (Chen and Johns 1991). In Porphyridium cruentum, the proportion of EPA decreased and that of AA increased in neutral and polar lipids under nitrogen starvation (Cohen et al. 1992), while no difference in the fatty acid composition was observed in the case of Navicula saprophilla (Kyle et al. 1989). In P. incisa, the fatty acid composition demonstrated a sharp increase in the proportion of AA, from 40.1% to 58.9% (of total fatty acids) under N-starvation, in comparison to only 46.2% in the control (Khozin-Goldberg et al. 2002a).
The lipid accumulation can be activated also by the depletion of other nutrients, such as phosphate and silicon. Lipid yield in the diatom Chaetoceros muelleri increased with silicon- depletion, whereas growth rate decreased (Engel et al. 2000). Experiments with the phosphate- deficient pho1 mutant of Arabidopsis showed that phosphate deprivation decreases the amount of phospholipids whereas that of non-phosphorus lipids (sulfolipid and digalactosyldiacylglycerol) is increased; indicating the substitution of phospholipids by non-phosphorous lipids is a general phenomenon in plants (Hartel et al. 1998). Hence, phosphate deprivation is a useful tool to assess the role of phospholipids in TAG synthesis. The accumulation of DGTS and decrease in the amounts of glycerophospholipids in the photosynthetic purple bacterium Rhodobacter sphaeroides and the bacterium Sinorhizobium meliloti, under phosphate-limiting growth conditions indicated that betaine lipids may substitute for phospholipids (Klug and Benning
2001).
29 1.6. Mutant studies
The most commonly used techniques for studying the biosynthesis of lipids in plants and algae are the incorporation of radiolabeled precursors, isolation of mutants, and the use of enzyme inhibitors. Mutants producing higher amount of TAG could be very useful for the production of
AA by P. incisa in a large scale. At the same time, TAG-deficient mutants can be analyzed to determine the steps of TAG synthesis.
An ethyl methanesulfonate-induced mutation in Arabidopsis thaliana impaired the activity of DAGAT, reduced the triacylglycerol biosynthesis and altered the seed fatty acid composition (Katavic et al. 1995). Genetic analyses indicated that the fatty acid phenotype is caused by a semidominant mutation in a single nuclear gene, designated TAG1, located on chromosome 2.
The developing seeds of the Arabidopsis wri1 mutant exhibited decreased accumulation of oil and an increased accumulation of carbohydrates, suggesting that the conversion of carbohydrates into the precursors of fatty acid and triacylglyceride synthesis (pyruvate and acetate) are in some way adversely affected (Focks and Benning 1998). Over-expression of wri1 could potentially result in more hexose sugars being converted to pyruvate and acetate, and ultimately favoring the synthesis of triacylglycerides over starch synthesis.
A mutant of the red alga, P. cruentum, which has impaired growth at sub-optimal temperatures, contained an elevated level of TAGs and reduced level of the 20:5/20:5 eukaryotic- like molecular species of monogalactosyldiacylglycerol (MGDG), suggesting that TAG can contribute to the biosynthesis of eukaryotic-like species of MGDG (Khozin-Goldberg et al.
2000).
30 1.7. Role of TAG
TAG, the main storage lipids of plants, is widely found as a major energy reserve in seeds and fruits. But, the role(s) of TAG other than as the energy reserve, especially that of the PUFA-rich-
TAGs have not been well understood.
When grown in a light-dark regime, Nannochloropsis produced 16:0 and 18:1-rich cytosolic lipid globules during the light period, which were utilized for polar lipid synthesis in the dark (Sukenik and Carmeli 1990). There are some indications of the role of TAG in reproductive functions in certain green algae. In Chlamydomonas reinhardtii, the accumulation of TAG, induced by nitrogen depletion, promotes the transformation of vegetative cells into male and female gamete forms (Martin and Goodenough 1975). Similarly, appearance of dense lipid bodies, made up mostly of TAG was reported to occur in Chlamydomonas moewusii during the fusion of the gametes (Brown et al. 1968). Under stress conditions such as high light intensity or nutrient starvation, cells of the unicellular alga Dunaliella bardawil overproduce β-carotene, which is accumulated in the plastids in newly formed triacylglycerol droplets. When the synthesis of triacylglycerol is blocked, the overproduction of β-carotene is also inhibited. Thus, TAG acts as a plastid-localized sink for the end product of the carotenoid biosynthetic pathway (Rabbani et al. 1998).
Labeled-AA in the TAG of the green alga P. incisa were transferred to polar lipids at low temperatures indicating that the PUFA of TAG can be mobilized for the membranes construction in response to the low-temperature-induced stress (Cohen et al. 2000). It has been suggested that algae that contain long-chain PUFAs and whose natural habitat is characterized by rapid changes in environmental conditions, like temperature, light, particularly PUFA-rich TAG might be involved as a buffer capacity for PUFA to swiftly provide its PUFA to construct membranes and increase the degree of unsaturation of membranal fatty acids during the sudden drops in temperature (Cohen et al. 2000).
31 TAG lipase(s) are required to utilize TAG. TAG is stored in intracellular lipid particles that are also named lipid bodies, lipid droplets, oil bodies, oleosomes, and spherosomes in plants.
The lipid body in seeds has a hydrophobic core, mainly formed of TAG and/or steryl esters
(STE), which is surrounded by a phospholipid monolayer, containing oleosin proteins. The oleosins are assumed to be involved in the mobilization of the neutral lipid core of the particle by serving as a docking and/or activating protein for TAG lipases and STE hydrolases (Athenstaedt and Daum 2003). Alternatively, such proteins were assumed to protect the stored lipids from random degradation (Murphy and Vance 1999).
In plants, many TAG lipases were isolated and characterized, although their subcellular localization has not been well studied. By using antibodies raised against animal lipases, Belguith et al. (2000) detected lipases of rapeseed in different subcellular fractions, isolated by differential centrifugation. During germination of maize kernel, lipases synthesized on free polyribosomes are specifically bound to oil bodies (Huang 1992).
1.8. Working hypothesis
The discovery of unusually high occurrence of the pharmaceutically and nutritionally important
PUFA, AA (up to 60%) in the abundant lipid TAG, constituting more than 30% of DW, in P. incisa, brought about a great interest in exploring the unique mechanism of accumulation of AA- rich TAG in this alga. Understanding the reasons for the accumulation of AA-rich TAG in this alga was also equally interesting and important. This alga was isolated from the slopes of a snow mountain in Japan, where sudden and intense climatic changes (temperature, light) are the environmental characteristics. So it has been assumed that the algae should contain some efficient mechanisms or enzymes, which can synthesize and utilize PUFA-rich TAG during a short span of environmental changes. The alga could have some specific role for abundant TAG in relation to the harsh environmental conditions. The goal of this research was thus to elucidate
32 the lipid intermediates of the biosynthetic pathway(s) of AA-rich TAG and the cellular distribution and characterization of the enzymes that are involved in the unusual targeting of AA into TAG and utilization of TAG. Additionally, determination of the role of AA-rich TAG in the alga was also aimed. These goals were accomplished by addressing the following objectives.
1. Elucidation of the fatty acid and molecular species composition of polar lipid classes
(PC, DGTS, PE), involved in the biosynthesis of AA-rich TAG.
2. Screening and characterization of TAG (or AA)-deficient mutants.
3. Characterization of the cellular distribution, activity, specificity and selectivity of the
diacylglycerol acyltransferase (DAGAT).
4. Elucidation of the role of AA-rich TAG in membrane construction.
5. Characterization of the triacylglycerol acyl hydrolase (lipase).
This research would allow better understanding of the underlying mechanisms of the biosynthesis, accumulation and utilization of AA in the unique system of P. incisa. The results of the research can be useful to increase AA production in this organism. The results can be also useful in searching for the genes encoding enzymes of TAG biosynthesis.
33 2. MATERIALS AND METHODS
The microalgal strain utilized in this work was isolated from a snow sample of Mt. Oyama
(Japan) in the Laboratory of Microalgal Biotechnology, Blaustein Institute for Desert Research.
This microalga has been identified as the chlorophyte, Parietochloris incisa (Trebouxiophyceae)
(Watanabe et al. 1996).
2.1 Growth conditions
Cultures were cultivated indoors in BG-11 nutrient medium in 1 L glass columns under controlled temperature and light conditions. The columns were placed in a temperature regulated water bath at 24 °C and illuminated by cool white fluorescent lights from one side at a light intensity of 170 µmol photon m-2s-1 as previously described (Cohen 1994, Bigogno et al.
2002a,b). The light intensity was measured at the middle and the center of the empty column with a quantum meter (Lamda L1-185). The cultures were provided with a continuous bubbling of air and CO2 mixture (99:1, v/v) from the bottom of the column.
In radiolabeling experiments, inhibitors and mutant studies, cultures were grown in 150 mL Erlenmeyer flasks under air:CO2 (99:1) atmosphere in an incubator-shaker at a speed of 170 rpm, and the illumination measuring to 115 µmol photon m-2s-1 was provided from above
(Khozin and Cohen 1996, Bigogno et al. 2002a). The flasks were closed by cotton or paper plugs, and the plugs were covered by aluminum foil.
For nitrogen-starvation experiments, NaNO3 was omitted from the medium and ferric ammonium citrate was substituted by ferric citrate. Similarly, K2HPO4 was substituted by KNO3 in the phosphate-starvation experiments.
34 2.2 Growth Parameters
Growth of the cultures was estimated on the basis of chlorophyll content and dry weight measurements. Chlorophyll concentration (µg/mL) was measured by extracting the chlorophyll pigments by dimethyl sulfoxide (DMSO). The biomass was collected from 3-5 mL of culture by centrifugation and extracted with a known volume of DMSO at 70 °C for 5 min. The extract was spun for 5 min (3500 rpm) and the absorbance of the supernatant was measured at 666 nm
(Diode array spectrophotometer HP 8452A or UV-visible spectrophotometer Cary 50 Bio
1 cm Varian). The chlorophyll concentration (µg/mL) was calculated using E 1% = 898 (Seely et al.
1972).
The biomass concentration was estimated by dry weight determination. Five mL of cultures were filtered through pre-weighed 25 mm glass fiber paper filters (Whatman GF/C,
Schleicher & Schuell Co.) and dried at 105 °C for 5-12 h until the constant weight.
2.3 Lipid analysis
2.3.1 Lipid extraction
Biomass was collected in 50 mL plastic tubes by centrifugation at 3400 rpm for 5 minutes. The supernatant was discarded by suction; the residue was frozen at -20 °C and lyophilized in a freeze dryer (10 MR-TR, Virtis Co.) for 24-48 h according to the amount of the biomass.
Lipid extraction from freeze-dried samples of biomass were carried out, first, by heating with DMSO (200 µL/ 50 mg biomass), while stirring it continuously with a magnetic stirrer at 70
°C for 5 min in 15 mL glass tubes, closed with caps containing teflon seal and then, by continuous mixing of the biomass with 5 mL methanol at 4 °C for 30 min. The mixture was centrifuged, the supernatant was transferred to a 50 mL teflon tube and the pellet was re- extracted with 1 mL of methanol. The combined extract was mixed with diethyl ether, hexane and water to form a final ratio of 1:1:1:1 (v/v/v/v). The mixture was vortexed vigorously, spun
35 for 5 min at 3500 rpm and the upper phase was collected to a round bottom flask. The water phase was acidified with 10-15 drops of 1 M KCl in 0.2 M H3PO4 to facilitate the extraction of the acidic lipids, PG and SQDG, and to improve phase separation. The water phase was re- extracted twice with the diethyl ether:hexane mixture (1:1, v/v). The organic phases were combined and evaporated to dryness in BUCHI Rotavapor R-114. At every step, the tubes and flasks were filled with argon to protect polyunsaturated fatty acids from oxidation.
2.3.2 Fatty acid analysis
Fatty acid profile and content in the samples were determined as their methyl esters by capillary
GC (Cohen et al. 1992). Transmethylation of fatty acids were carried out by incubation of the freeze-dried cells, or total lipid extracts, or individual lipids (either adsorbed on silica of TLC plates or extracted from it) in dry methanol containing 2% H2SO4 at 70 °C for 1-1.5 h under argon atmosphere and continuous mixing. Heptadecanoic acid (Sigma) in petroleum ether (bp
80-100 °C) was added as an internal standard. The reaction was terminated by addition of water and the methyl esters were extracted by hexane.
Gas chromatographic analysis of fatty acid methyl esters were performed on a
Supecowax 10 (Supleco Inc., Bellefonte, PA) fused silica capillary column (30 m x 0.32 mm) using temperature gradient of 185 °C to 210 °C (5890 Gas Chromatograph, Hewlette Packard).
Initially, temperature was programmed for 185 °C isothermically for 10 min. Then it was increased to a final temperature of 210 °C at a rate of 10 °C/min, and the final temperature was maintained for 5 min at 210 °C before the program cycle finished. Flow of the carrier gas, helium, was maintained 50 mL/min and the pressure of the column head was 80 kPa.
Temperature of the injector was 280 °C and that of the flame ionization detector (FID) was 300
°C. Fatty acid methyl esters were identified by co-chromatography with authentic standards
(Sigma) and by comparison of their equivalent chain length (Ackman 1969). The data shown
36 represent mean values with a range of less than 5% for major peaks (over 10% of fatty acids) and
10 % of minor peaks, of at least two independent samples, each analyzed in duplicate. Total fatty acid numbers were calculated on the basis of the identified FAME peaks. These included over
98% of total fatty acid peaks.
2.3.3 Lipid separation
Total lipid extract was separated into neutral and polar lipids by silica Sep-Pak (Waters) or Bond-
Elute (Varian) chromatography cartridges using chloroform or 0.5% methanol in chloroform to elute neutral lipids, and methanol to elute polar lipids (Cohen et al. 1992).
Polar lipids were separated into individual lipids by two dimensional TLC (Silica Gel 60,
10x10 cm, 0.25 mm thickness, Merck, Darmstadt, Germany) using a solvent system of chloroform:methanol:water (65:25:4, v/v/v) for the first direction, and of chloroform: methanol:
1-ethylpropylamine:conc.ammonia (65:35:0.5:5, v/v/v/v) for the second direction (Khozin et al.
1997). Neutral lipids were resolved with petroleum ether:diethyl ether:acetic acid (70:30:1, v/v/v).
Lipids on TLC plates were visualized by brief exposure to iodine vapors, scraped from the plates and were transmethylated for the fatty acid analysis as previously described, or sequentially extracted with mixtures of chloroform:methanol (2:1, v/v) and (1:1, v/v) for further analysis. Individual lipids were identified by spraying with specific reagents and/or by comparison of Rf values to Rf values provided in literature (Christie 1982) and/or by running commercial lipid standards along with the samples.
The individual lipids were further purified by a silica column of normal phase HPLC
(LiChrospher, Si-60, 5 µM, Merck). DGTS and PE obtained from TLC spots, were purified by employing a linear gradient system of hexane:isopropanol:water (from 40:60:0 to 40:53:7) with
37 flow-rate of 1 mL/min. PC was purified with a mobile system of hexane:isopropanol:water
(40:53:7) with flow-rate of 1 mL/min.
2.3.4 Molecular species separation
Individual lipids were resolved into component molecular species by reverse-phase HPLC utilizing an HPLC system equipped with a reverse phase column (LiChrospher 100, RP-18, 5
µM, Merck) variable-wavelength (UV-visible) detector (Lamda-Max 481, Waters), a mass detector (Evaporative light scattering detector, ELSD II A, Varex). DGTS, PE and PC molecular species were separated by using a mobile systems of methanol:acetonitrile:water, 80:12:8,
60:35:5 and 60:35:5 (v/v/v), respectively. Five µM of ethanolamine was added to the acetonitrile in the case of PE molecular species separation. TAG molecular species were resolved by a gradient system of acetonitrile:(acetonitrile:ethanol:hexane, 40:20:20), which started from a proportion of 70:30 to 0:100 at the end of 90 min run.
To identify the individual molecular species composition, elution of each peak of molecular species as detected at 205 nm (where isolated double bonds absorb) by UV-visible detector was collected in separate tube and its fatty acid composition was analysed by GC. ELSD was utilized to quantify the individual molecular species employing evaporation temperatures of
123 °C for DGTS, and 125 °C for PE, PC and TAG respectively. The proportion of molecular species (% of total) were calculated on the basis of their mass values obtained by HPLC-ELSD.
The proportion of molecular species (% of total) were calculated on the basis of their mass values obtained by HPLC-ELSD. Equivalent carbon number (ECN) of each molecular species were calculated according to an equation:
ECN = TOTAL NUMBER OF CARBON ATOMS IN THE FATTY ACIDS −2 X TOTAL NUMBER OF DOUBLE BONDS
Under isocratic conditions, molecular species that contain fatty acids with longer carbon chain and less desaturation are eluted before those with shorter chains and higher desaturation. The
38 molecular species with the same or close ECN number could not be separated by HPLC.
However, the major and minor molecular species found within a peak of HPLC were estimated by the amount of respective fatty acids in the peak, determined by GC.
2.3.5 Positional analysis of fatty acids distribution in individual lipids
Positional analysis of fatty acids of the molecular species of DGTS was performed according to
Norman and Thompson (1985). DGTS was incubated with Rhizopus lipase (Sigma) in 1 mL of diethyl ether, 0.25 mL of 10 mM sodium phosphate buffer (pH 7.4) containing 0.5 mM CaCl2 for
1 h at 37 °C. The reaction was terminated by evaporation of the diethyl ether under the flow of
N2. Extraction was carried out twice using diethyl ether/petroleum ether (1:1, v/v). Separation of free fatty acids, representing sn-1 position of DGTS, and lyso-DGTS, representing sn-2 position of DGTS, was done by TLC using a solvent system of chloroform:methanol:acetic acid:water
(85:15:10:3.5, v/v/v/v). The fatty acid analysis of TLC bands was carried out by GC, as described above.
Similarly, positional analysis of fatty acids of PC and PE were carried out by incubating lipids with phospholipase A2 (from Crotalus admanteus venom poison, Sigma) in 1 mL diethyl ether, 50 µl of 100 mM boric acid-Borax buffer pH 7.6 for 1 h at 37 °C. The reaction was terminated by evaporation of the diethyl ether, and extraction was carried out by the Bligh-Dyer method (1959). The products were separated by HPLC, utilizing normal phase Si-60 column
(Merck). A solvent system of hexane:isopropanol:water (40:53:7), at a rate of 1.5 mL/min, eluted unesterified fatty acids (representing sn-2 position) immediately after injection and lyso-PC after
30 minutes of injection.
39 2.4 Nitrogen starvation and recovery experiments
Cells growing in the early stationary phase were collected by centrifugation, washed several times with sterile double distilled water, and the pellet was resuspended in the equal volume of nitrogen-free BG-11 medium and cultivated for another 14 days. For the recovery experiments, the nitrogen-starved culture was centrifuged and the pellet was resuspended in the full BG-11 medium with a four-fold dilution. The lipid separation, analysis of fatty acids, separation of molecular species and positional analysis of fatty acids were carried out as described above.
In growth recovery experiments after nitrogen replenishment, after 13 days of N-
14 starvation, the cultures were labeled with the ammonium salt of [1- C] 18:1 (5 µCi, specific activity 52 mCi/mmol, Amersham) for 24 h, washed and diluted four-fold in full medium and chased for 48 hours. Lipids were extracted, separated by TLC and the distribution of radioactivity in the lipid spots was determined by a phosphoimager (BAS 1000, Fujix). TLC plates were exposed to imaging plates for overnight before scanning. Radioactive spots were scrapped and counted with a radioactivity counter (1600 TR-liquid scintillation analyzer,
Packard) using universal liquid scintillating cocktail (Ultima Gold, Packard Bioscience).
2.5 Inhibitor studies
2.5.1 Salicylhydroxamic acid (SHAM)
Stock solutions of 30 mM salicylhydroxamic acid (Sigma-Aldrich) in DMSO was added to 150 mL of stationary phase cultures (grown for 6 days in a glass column) to achieve a final concentration of 300 µM. The cultures were centrifuged; pellets were washed, resuspended in 3 volumes of the medium, and grown for another 6 days to study a recovery from the inhibition.
40 2.5.2 Sethoxydim
The cells harvested from the culture of early stationary phase (6 days) were washed twice with water, resuspended in nitrogen-free BG-11 medium and incubated in the presence of sethoxydim
(Riedel-de Haën) for 3 days. Stock solution of 10 mM sethoxydim in DMSO was administered to the cultures to achieve final concentrations of 50, 100 and 200 µM.
2.6 Characterization of TAG biosynthetic enzymes
2.6.1 Grinding and homogenization
The biomass from 200 mL of 14 days old batch culture, stored at -70 °C in a plastic tube, was frozen in liquid nitrogen and ground into fine powder in a liquid nitrogen-cooled mortar with a pestle. Thirty mL of 50 mM CHES-NaOH buffer (pH 9.0) was used for further grinding and homogenization of the cells. Lypolytic activity discovered in cell-free extracts of the alga was minimized by using this high pH grinding buffer containing 5 mM EDTA, 1 mM EGTA, 1% w/v
BSA, 20 mM KCl, 2 mM MgCl2, 10% v/v glycerol, 0.6 M sucrose, 0.005% w/v Triton X-100.
PMSF (1 mM), DTT (2 mM) and catalase (1000 units/mL of buffer) were added freshly to the buffer before grinding.
2.6.2 Cellular fractionation
The homogenate was centrifuged at 1500 x g for 15 min (Sorvall RC plus 5C, HB4-swinging bucket rotor) to remove cell debris and unbroken cells. The supernatant and the floating layer were collected, mixed by a 10 mL glass pipette and subjected to a discontinuous sucrose gradient centrifugation at 25000 x g for 1 h. For that, 12 mL of the homogenate, 10 mL of 0.4 M and 10 mL of 0.2 M sucrose gradient buffers were placed one above another in a plastic tube for the gradient centrifugation. The sucrose gradient buffers contained 50 mM CHES pH 9.0, 1 mM
EDTA, 20 mM KCl, 1 mM MgCl2, and 1 mM DTT. The floating layer of oil bodies above 0.2 M
41 sucrose was collected with a cut needle of a 1 mL syringe, mixed well in 5 mL of 0.4 M sucrose gradient solution by a 1 mL plastic tip for washing, and subjected again to the gradient centrifugation for 1 h (25,000 x g) by overlaying with 20 mL of 0.2 M sucrose gradient solution.
The floating layer of oil bodies above 0.2 M sucrose was collected, gently homogenized, divided into eppendorf tubes and stored at -70 °C until use. The pellet obtained by 25000 x g centrifugation was homogenized in the storage buffer (pH 9.0), containing 50 mM CHES, 0.33
M sucrose, 1mM DTT and stored at –70 °C until use.
The supernatant obtained after 25000 x g centrifugation was filled in ultracentrifuge tubes and centrifuged at 100,000 x g for 1 h (Beckman-H; SW 41Ti-swinging bucket rotor). The pellet
(100,000 x g, microsomes) was collected, washed by homogenizing it in 50 mM CHES buffer pH 9.0 with 1 mM DTT and centrifuged at 100,000 x g for 1 h again. The pellet obtained after washing was homogenized in the CHES buffer (pH 9.0, 1 mM DTT) and stored at -70 °C until use.
All procedures were carried out at 4 °C. Protein content in cellular fractions was quantified by the method of Bradford (1976). In oil bodies, protein content was measured after delipidation with diethyl ether. For that, the oil bodies were mixed with diethyl ether and sonicated to facilitate the extraction of lipids. The upper organic phase was removed after centrifugation. The process was repeated 2-3 times for complete delipidation. The β-carotene
1 cm content of the oil bodies was determined spectrophotometrically using E 1% absorption coefficient of 2592 at 450 nm in petroleum ether (Britton 1995).
2.6.3 Enzymatic assays
The formation of [1-14C] TAG from 0.4 mM [1-14C] 1,2-oleoylglycerol (20000–30000 DPM) was measured in the presence or absence of 20 µM oleoyl-CoA in the final volume of the assay mixture of 100µL (Hobbs and Hills 2000). In some experiments 1-stearoyl-2-[14C] arachidonyl-
42 14 14 DAG was utilized. [1- C] 1,2-dioleoylglycerol or 1-stearoyl-2-[ C] arachidonyl-DAG were obtained by the action of phospholipase C on their respective PCs (121 mCi/mmol; Amersham
Biosciences, UK) according to Vogel and Browse (1995). Otherwise, 20 µM [1-14C] oleoyl-CoA
(20000 DPM) and unlabeled 0.4 mM 1,2-oleoylglycerol were utilized (Bouvier-Nave et al.
2000). Reactions were initiated by addition of cellular fractions. Assay mixtures containing substrates, 0.125% BSA, 100 mM Tris-HCl (pH 7.5), were incubated at 30 °C for indicated times. MgCl2 (150 mM), 20% glycerol and other chemicals were added when indicated. In selectivity experiments, different DAG and acyl-CoA molecular species were supplied. Stock solutions of stearoyl-, oleoyl-, linoleoyl- and arachidonoyl- CoA (Larodan) were prepared in 100 mM sodium acetate buffer pH 5.0; stock solutions of sn-1,2 stearoyl-, oleoyl-, linoleoyl- and arachidonoyl-glycerol (Nu-Check Prep) were prepared in 0.2 % (w/v) Tween-20.
Similarly, synthesis of DAG from unlabeled 2- or 1- monoacylglycerols was determined by applying 20 µM [1-14C] oleoyl-CoA (20000 DPM) to the reaction mixture (100 µL). The sequential acylation of the lipids of the Kennedy pathway were studied by providing 20 µM [1-
14C] acyl-CoA (20000 DPM) to unlabeled glycerol-3-phosphate in the reaction mixture.
The stock solutions of DAG and MAG in Tween-20 were heated to 70 °C and sonicated warm until transparent solution to break the micelles. Ten microliters DAG or MAG stock solutions were added to the reaction medium, sonicated again for 1 min, and protein was added to start the reaction. Magnetic stirrer maintained the continuous mixing of the reaction medium and a constant temperature was maintained in a water bath. Reactions were terminated by the addition of 0.8 mL ice-cold mixture of chloroform:methanol:acetic acid (50:50:1, v/v/v) and 0.4 mL of ice-cold water and vortexing.
Lipid extraction was repeated thrice and followed by TLC separation of the reaction products. TLC was first run for 3 cm from the origin of 10 cm-long plate in chloroform:methanol:water:acetic acid (73:25:4:2, v/v/v/v) system. After drying the plate in air
43 for 10 min, it was run up to the top in a neutral lipid system (petroleum ether:diethyl ether:acetic acid, 70:30:1, v/v/v) twice. Labeled lipids were detected and estimated by a phosphoimager
(BAS 1000, Fujix) as mentioned earlier. Radioactive spots were scrapped and counted with a radioactivity counter (1600 TR-liquid scintillation analyzer, Packard) using universal liquid scintillating cocktail (Ultima Gold, Packard Bioscience).
Each enzymatic assay was repeated at least twice mostly using different batches of cellular fractions isolated from different batches of algal cells. Otherwise specified, the representative experiment was shown in figures.
2.7 Mutagenesis
Fifty mL of log phase cultures were sonicated in flasks to break the cell clumps. Ten mL of cell suspension, containing mostly single cells, were exposed to a mutagen, 1-methyl-3-nitro- nitrosoguanidine (MNNG, Aldrich) at final concentration of 50 or 100 µg/mL for 1 h in an incubator shaker. The stock solution of MNNG was prepared in DMSO (5 mg/mL) to ease the penetration of the mutagen across the cell tough cell wall of the alga. The cultures were washed with BG-11 medium several times. Finally, the cultures were well sonicated in 10 mL of medium and cell numbers of untreated and treated cultures were counted to further determine the percent of survived cells. The cultures were sequentially diluted to 1000 cells/mL. Fifty or 100 µL of cultures of different dilutions were plated on BG-11 agar plates and grown under fluorescent light at the room temperature. Colonies, which showed poor or better growth, were selected and grown in eppendorfs or glass tubes under fluorescent light at room temperature for a few days.
After the growth was observed, 5-10 µL of the cultures were plated to agar plates containing BG-
11, N-free BG-11 and P-free BG-11 for further screening. Mutant and wild type cultures were also grown in a incubator shaker at 25 °C and/or 15 °C. Lipid and fatty acid content were analyzed as described above.
44 3. Results
3.1. Lipid studies
3.1.1. Lipid classes and fatty acid composition
Aiming to elucidate the role of major extraplastidial lipids (PC, PE, DGTS) as suppliers of building blocks for the biosynthesis of TAG we separated and analyzed these lipids by TLC and
HPLC. Separation of major lipid classes, determination of their content and fatty acid composition, identification of the molecular species and positional analysis of fatty acids in PC,
PE, DGTS were carried out using cells harvested at the logarithmic and stationary phases.
Decreased nutrient availability was shown to affect an enhanced biosynthesis of AA-rich TAG in stationary phase. Previous work conducted in our laboratory was focused on the comparative analysis of molecular species composition of major chloroplastic lipids, MGDG and DGDG under those conditions (Bigogno et al. 2002b).
The amount of total lipids in the biomass increased by 4.5 times in the stationary phase in comparison to the logarithmic phase of growth (Table 1). The proportions of the individual lipid classes changed during growth. Neutral lipids were represented by the predominant TAG and low amounts of DAG and FFA. TAG shared the largest proportion of total lipids in both stages, but especially in the stationary phase (82.1 and 38.8% of total acyl lipids in stationary and logarithmic phases, respectively). Among the polar lipids, the chloroplastic galactolipids MGDG,
DGDG and the sulpholipid SQDG, the major glycolipids of the chloroplastic membranes, were the predominant polar lipids in the log phase (overall 38% of total lipids). However, the total share of chloroplastic lipids decreased sharply to 10% at the stationary phase. Phospholipids constituted a rather smaller portion (13.2%) of total lipids. PC comprised the largest proportion
(6.6%) among the phospholipids. In the stationary phase, except of betaine lipid DGTS and PA, the proportion of all polar lipids decreased sharply, while TAG increased drastically.
45 The fatty acid analysis of the biomass of P. incisa has revealed that AA is the major fatty acid in both the exponential and stationary phases (25.8 and 45.8% of TFA, respectively) (Table
1). The fatty acids 16:0, 18:1, 18:2 and 18:3(ω3 and ω6) appeared in significant proportions at both phases, except for 18:3ω3, which decreased sharply in the stationary phase with the increase of AA.
PC, PE and DGTS, which comprise 78% of extrachloroplastic lipids, were rich in 18:0-3,
20:3ω6 and exceptionally rich in AA. PC and DGTS contained comparatively higher proportions of 16:0, 18:1ω9, 18:2 and 18:3(ω6 and ω3), whereas PE contained higher proportions of 18:1ω7,
20:3 and AA. The proportions of 20:3ω6 and AA were highest in PE among the polar lipids. At the stationary phase, the proportion of 18:2 increased while that of 18:3ω6, 20:3ω6 and AA decreased in keeping with radiolabeling data on turnover of AA precursors in these lipid followed by the transfer of AA to TAG (Bigogno et al. 2002a). PE contained also a high proportion of 18:1ω7, a fatty acid, which is common to bacteria (Asselineau 1966).
The highest proportions of 16:1 fatty acids were found in PG. The 16:1trans ∆3, which is unique to this chloroplast phospholipid, was shown to play a role in the electron transport of PS
II (Douce and Joyard 1996). In the stationary phase, similarly to glycolipids, the proportions of the unsaturated 18:3ω3 and AA decreased in PG, but that of 16:1∆3 increased, suggesting the importance of 16:1trans ∆3 for thylakoid membrane function in this phase.
PA is an important lipid intermediate of the biosynthesis of membrane lipids and the
Kennedy pathway of TAG biosynthesis. This lipid being the product of phospholipase D is also involved in phospholipid turnover and signaling. Because of its fast turnover, it is usually not accumulated in significant proportions. Indeed, PA comprised 1.4 and 1.1% of TFA in the log and stationary phase, respectively. This lipid contained the highest proportion of 18:0 and 18:1ω9 among the polar lipids and higher 18:3ω3 in comparison to 18:3ω6. During the stationary phase, the proportions of 18:2 and AA increased sharply in PA.
46 Table 1. Fatty acid composition of biomass and individual lipids of P. incisa in the logarithmic
(L) and stationary (S) cultures of P. incisa
Lipid Growth TFA** Fatty acid composition (% of total) class condition %DW 16:0 16:1 16:1 16:2 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:0 20:1 20:2 20:3 20:4 20:5 22:0 ω7 ω5 ω6 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 ω3
Biomass L 14.9 4.5 tr 1.3 5.1 1.0 8.5 6.5 15.0 1.6 12.2 0.6 - tr 0.8 25.8 1.7 0.2 S 9.1 0.2 0.2 1.0 0.4 2.8 15.6 3.7 16.1 0.8 1.1 0.2 0.5 0.6 0.8 45.8 0.6 0.2
MGDG L 15.2 3.0 1.5 tr 7.1 23.4 0.3 5.3 0.9 13.2 0.8 33.9 0.2 0.3 tr tr 8.9 1.0 0.3 S 2.7 2.4 1.4 - 29.7 11.2 0.2 3.2 0.5 34.1 0.5 13.8 0.2 tr - - 2.7 --
DGDG L 11.8 29.2 1.4 0.4 2.7 3.9 0.8 8.8 2.2 20.1 1.1 21.4 tr 0.5 - 0.2 7.8 1.4 0.7 S 2.9 15.0 1.7 0.4 11.7 - 0.7 5.3 1.2 43.1 1.0 6.6 0.5 - - 0.3 1tr 0.3 1.0
SQDG L 11.1 62.5 0.3 0.2 - - 0.6 8.6 5.8 12.8 0.2 11.1 - 0.2 - - 0.6 tr 0.4 S 4.4 63.7 0.2 0.2 tr - 0.6 3.3 11.7 17.8 tr 1.3 - - - - 0.3 tr 0.4
PG* L 2.7 36.6 10.3 - 1.3 - 0.9 8.7 2.7 22.4 3.6 5.1 - 0.6 - - 1.4 - - S 0.5 21.2 17.9 - 5.0 - 0.8 8.7 1.6 17.6 0.6 1.8 - 0.3 0.5 - 0.6 - - PC L 6.6 17.1 0.3 0.9 0.2 0.3 3.7 7.9 7.0 18.4 7.4 4.7 0.4 0.2 - 3.1 25.4 1.1 0.7 S 0.6 19.0 0.6 0.8 1.3 - 4.2 7.0 8.8 26.4 2.8 1.3 0.6 - 1.0 1.1 25.0 - -
DGTS L 4.6 32.2 0.4 1.2 0.3 0.3 2.6 6.0 2.8 21.0 8.7 4.6 0.6 0.2 - 1.1 14.9 0.9 1.3 S 4.1 35.9 0.4 1.0 0.2 tr 4.7 4.5 3.4 30.3 2.7 0.6 0.5 - 0.4 0.5 13.9 0.3 0.7
PE L 1.8 6.5 - 1.1 - - 3.2 3.4 18.5 5.7 2.9 1.3 0.4 0.5 - 11.3 41.3 1.7 1.3 S 0.5 10.8 - 1.0 1.1 - 5.2 4.7 20.9 15.1 2.1 1.3 0.6 - 0.8 4.0 31.8 --
PA L 1.4 48.4 1.0 0.2 0.7 1.3 4.5 19.2 3.2 13.6 0.3 5.9 - - - 1.8 - - S 1.1 33.2 0.8 0.2 2.4 tr 1.9 10.4 2.3 37.4 0.4 2.3 - - - 0.3 8.4 - -
PI L 0.7 53.7 tr 0.8 tr - 3.7 15.4 4.5 15.8 1.0 1.4 0.3 0.3 tr tr 2.2 - - S 0.9 57.0 0.3 0.5 tr - 1.2 8.3 3.4 23.8 1.8 0.2 - - 0.3 - 3.1 - - TAG L 38.8 9.9 0.2 0.2 0.2 0.6 2.4 24.0 3.5 12.1 1.2 2.2 0.4 0.8 - 1.1 38.1 2.4 0.6 S 82.1 9.1 0.2 0.2 tr - 2.9 19.0 4.2 14.5 0.7 0.4 0.2 0.3 0.5 0.8 45.0 0.6 -
DAG L 2.3 14.7 0.2 0.5 0.4 1.1 3.3 27.5 4.0 13.8 1.1 2.5 0.4 0.5 - 0.8 27.3 1.3 0.3 S 0.4 25.2 6.4 0.9 - 0.3 8.9 19.4 4.4 12.5 1.9 0.5 0.5 - 0.2 1.1 16.6 - -
FFA L 1.1 25.0 1.0 0.4 0.4 - 4.0 19.5 5.3 12.0 1.2 4.2 0.4 0.3 tr 0.7 22.6 1.4 0.3 S tr 26.4 0.3 0.4 - 1.0 7.3 39.6 6.4 9.3 0.5 0.6 0.9 1.6 0.3 0.9 4.4 0.2 -
*16:1 trans ∆3 constituted for 6.3 and 23.4% in PG at logarithmic and stationary phases, respectively.
** TFA (Total fatty acid) content was 6.6 and 29.4% of dry wt at logarithmic and stationary phases, respectively.
47 3.1.2. Molecular species composition of PC
Positional analysis of polar lipids was conducted in order to investigate the stereospecific acyl distribution in glycerol backbone. Fatty acid composition of sn-1 and sn-2 positions, determined following action of phospholipase A2 on PC and PE and Rhizopus lipase on DGTS (Materials and Methods- 2.3.5), is presented in Table 2. The positional analysis of fatty acids of PC did not reveal strict positional specificity. However, the sn-1 position of PC showed preference to 16:0,
AA and 18:1ω7, comprising 78.7% of fatty acids of the sn-1 position whereas the sn-2 position favored 18:1ω9, 18:2, 18:3(ω6 and ω3) and AA, comprising 75.4% of fatty acids of the sn-2 position (Table 2). It thus follows that the fatty acids in the sn-2 position are likely substrates for the desaturases of C18 fatty acids in the course of the biosynthesis of AA.
Table 2. Positional analysis of PC, PE and DGTS in the logarithmic culture of P. incisa
Lipid Fatty acid composition (% of total fatty acids) 16:0 16:1 16:2 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:3 20:4 20:5 ω7 ω6 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω3 Lyso-PC (sn-1) 33.7 1.0 tr tr 6.0 4.5 12.2 4.5 0.7 0.9 0.7 32.8 1.8 FFA-PC (sn-2) 8.0 4.4 0.2 0.4 2.1 8.3 3.2 30.8 7.3 5.5 2.3 23.5 3.3
Lyso-PE (sn-1) 15.0 1.8 tr tr 6.7 8.2 35.5 3.8 tr tr 5.7 19.6 3.7 FFA-PE (sn-2) 5.9 2.6 1.2 1.6 3.9 7.3 2.9 16.6 1.8 2.1 9.3 42.3 2.4
FFA-DGTS (sn-1) 80.3 0.2 0.6 tr 5.8 2.0 2.2 4.5 0.3 1.4 tr 0.3 tr Lyso-DGTS (sn-2) 4.8 1.9 1.5 tr 1.0 12.1 2.6 40.7 14.9 9.4 1.2 8.5 tr
Out of the eight major peaks of molecular species of PC detected by HPLC (Fig. 9 and
Table 3), five contained molecular species with 18:2 and 18:3 at the sn-2 position, constituting
54.3% and 60.7% of total PC in the log and stationary phases, respectively (Table 3). On the basis of the fatty acid composition of individual peaks and selectivity in the sn-1 position of PC, three groups of molecular species were implied, containing AA, 16:0 or 18:1ω7 at this position.
The AA- group shared the highest proportion (57.6%) followed by the 16:0 - (28.3%) and the
48 18:1ω7 - (14.1%) groups. During the stationary phase, the 16:0- group decreased sharply to
17.1% while the other two groups increased. The proportion of AA/18:2 increased sharply (from
25.4 to 37.5%) at the expense of AA/18:3(ω6 and ω3), which decreased from 6.4 to 1.7%.
Molecular species having the same or close equivalent chain number (ECN) could not be separated by HPLC. ECN describes an elution order of molecular species and it is calculated according to the equation:
ECN = total number of carbon atoms in the fatty acids- 2x(total number of double bonds).
Under isocratic conditions, molecular species that contain fatty acids with longer carbon chain and lower desaturation are eluted before those with shorter chains and higher desaturation.
However, the major and minor molecular species found within a same peak were estimated by
GC analysis of the fatty acids.
2 3 A
1 6 8 5 7 4
3 B 2 Detector response
5 7 8 1 4 6
0 20406080 Time (min)
Figure 9. HPLC-ELSD chromatogram of the molecular species of PC in the logarithmic (A) and stationary (B) cultures of P. incisa.
49
Table 3. Molecular species composition of PC in the logarithmic and stationary cultures of P. incisa
Log phase Stationary phase Peak ECN Molecular species (% of total) (% of total) 3 26 AA/18:2 25.4 37.5 1 24 AA/18:3ω6 (70.3%*), AA/18:3ω3 6.4 1.7 2 24 AA/AA 25.8 25.4 8 30 16:0/18:2 12.7 8.5 4 28 16:0/18:3ω6 (73.3%), 16:0/18:3ω3 3.8 3.3 6 28 16:0/AA 11.8 5.3 7 30 18:1ω7/18:2 (82.3%), 18:1ω9/18:2 6.0 9.7 5 28 18:1ω7/AA (70.6%), 18:1ω9/AA 8.1 8.7
The molecular species are grouped according to the fatty acids at the sn-1 position. *The proportion of the molecular species in the peaks was calculated for the log phase only. Positional analysis of individual molecular species was not carried out. The positional distribution of the fatty acids is based on the positional analysis of total PC (Table 2).
3.1.3. Molecular species composition of PE
Positional analysis of PE showed that the sn-1 position was dominated by 18:1ω7, AA and 16:0, comprising 70.1% of TFA (Table 2). The fatty acids, 18:2, 18:3ω6, 18:3ω3, 20:3ω6 and AA, constituted 72.1% of the fatty acids at the sn-2 position. The sn-2 position selectively incorporated 18:2, 18:3(ω6 and ω3), moreover, much higher proportions of 20:3ω6 and AA were observed in the sn-2 position of PE in comparison to other polar lipids, suggesting this lipid as a major substrate for the ∆5 desaturation. The major molecular species of PE was 18:1ω7/AA
(54.8%), followed by AA/AA (Peaks 7 and 3, respectively, Fig. 10 and Table 4). Several molecular species contained AA in the sn-1 position (AA/18:2, AA/18:3, AA/20:3, AA/AA), and their proportion increased in the stationary phase (from 23.2 to 32.9%), indicating an increase in the biosynthesis of AA within this lipid.
50 7 3 A
4 5 1 2 9 6 8
7 B Detector response
3
1 4 2 5 9 6 8
0 5 10 15 20 25 30 35 40 Time (min)
Figure 10. HPLC-ELSD chromatogram of the molecular species of PE in the logarithmic (A) and stationary (B) cultures of P. incisa.
Table 4. Molecular species composition of PE in the logarithmic and stationary cultures
Log phase Stationary phase Peak ECN Molecular species (% of total) (% of total) 4 26 AA/18:2 3.8 8.2 2 24 AA/18:3 1.7 2.0 5 26 AA/20:3 1.5 1.5 3 24 AA/AA 17.2 22.7 1 22 AA /20:5 1.2 4.2 7* 28 18:1ω7/AA 54.8 48.9 5 26 18:1ω7/20:5 2.6 2.6 6 26 18:3/18:2 tr tr 9 28 20:3/18:2 3.7 1.7 8 28 20:3/20:3 3.5 tr 7* 28 16:0/AA 9.2 8.2
*The proportion of molecular species in the peaks was calculated for the log phase only. Positional analysis of the individual molecular species was not carried out. The positional distribution of the fatty acids is based on the positional analysis of total PE (Table 2). tr- traces
51 3.1.4. Molecular species composition of DGTS
The sn-1 position of DGTS was dominated by 16:0 (80%) (Table 2). The sn-2 position was occupied by C18 fatty acids, mainly 18:1ω9, 18:2, 18:3(ω6 and ω3), and AA. Similarly to PC, the molecular species analysis revealed the presence of different groups of molecular species that contained 16:0, 18:1(ω7 and ω9) or AA at the sn-1 position (Table 5). The major molecular species of DGTS were 16:0/18:2, 16:0/18:3(ω6 and ω3) and 16:0/AA (Peaks 7, 5 and 6, respectively, Fig. 11), in the log phase. In the stationary phase, the proportion of 16:0/18:2 (#7) increased from 55.0 to 68.1% at the expense of 16:0/18:3(ω6 and ω3) (#5), which sharply decreased from 12.5 to 1.9%. However, 16:0/AA, AA/18:2, AA/18:3(ω6 and ω3) did not change significantly. There was some decrease in AA/AA. Since some HPLC peaks contained more than one molecular species due to the same ECN, their proportions were estimated by GC.
7 A
6
5
3 2 1 4
7 B Detector response Detector
6
3 4 2 5 1
020406080100 Time (min)
Figure 11. HPLC-ELSD chromatogram of the molecular species of DGTS in the logarithmic (A) and stationary (B) cultures of P. incisa.
52 Table 5. Molecular species composition of DGTS in the logarithmic and stationary cultures of P. incisa
Log phase Stationary phase Peak ECN Molecular species (% of total) (% of total) 3 26 AA/18:2 5.9 5.6 1 24 AA/18:3ω6 (95%*), AA/18:3ω3, 20:5/20:3 0.2 0.1 (tr) 2 24 AA/AA (82.9%), 3.2 1.6 26 18:3ω6/18:2, 18:3ω3/18:2 7 30 16:0/18:2 (90.9%), 18:1ω7/18:2 (7.6%), 55.0 68.1 18:1ω9/18:2 (3.3%) 5 28 16:0/18:3ω6 (66.9%), 16:0/18:3ω3 (33.1%) 12.5 1.9 6 28 16:0/AA (83.3%), 18:1ω7/AA (6.7%), 21.2 19.1 18:1ω9/AA (5.5%) 4 28 18:2/18:2 (95.7%), AA/20:3 (4.3%) 1.1 3.3
*The proportion of molecular species in the peaks was calculated for the log phase only. Positional analysis of the individual molecular species was not carried out. tr- traces Positional distribution of the fatty acids is based on the positional analysis of total PE (Table 2).
3.1.5. Molecular species composition of TAG
In contrast to most algae, where TAG are made up of mainly saturated and monounsaturated fatty acids, in P. incisa, AA is the major fatty acid of TAG (Table 1). TAG, which is the major lipid of the alga, contained AA (38%), 18:1ω9 (24%), 18:2ω6 (12%) and 16:0 (10%) in the exponential phase. In the stationary phase, the proportion of TAG was doubled and the proportion of AA increased to 45%. Consequently, the share of cellular AA deposited in TAG increased from 57% to 81%. The proportion of 18:1ω9, 18:3 and 16:0 decreased in the stationary phase.
The major molecular species of TAG contained two arachidonyl moieties per molecule
(AA/AA/18:1ω9, AA/AA/18:2 and AA/AA/16:0; Peak 8, 5 and 9, respectively, Fig. 12 and
Table 6), constituting 33.7% of total TAG or even three arachidonyl moieties (AA/AA/AA, peak
4, Fig. 12, Table 6) per molecule, amounting to 6.4% of total TAG. Only minor changes were
53 noted on transfer to the stationary phase. AA/AA/16:0 and AA/AA/18:1ω9 have shown some increase.
8 A
13 5 14 11 10 4
9 16 7 12 1 2 3 6 15 1718
8 B Detector response Detector
11 5 14 10 13
9 4 16 7 12 18 6 15 19 20
020406080 Time (min)
Figure 12. HPLC-ELSD chromatogram of the molecular species of TAG in the logarithmic (A) and stationary (B) cultures of P. incisa.
54 Table 6. Molecular species composition of TAG in the logarithmic and stationary cultures of P. incisa
Log phase Stationary phase Peak ECN Molecular species of TAG (% of total) (% of total) 1 36 AA/AA/18:3ω3 0.6 tr 2 34 AA/AA/20:5 1.0 tr 3 34 20:5/18:3ω6/AA 2.1 tr 4 36 AA/AA/AA 6.4 5.4 5 38 AA/AA/18:2 12.9 9.8 6 38 20:5/AA/18:1ω9 1.0 0.1 7 X1 2.0 1.0 8 40 AA/AA/18:1ω9 18.9 21.7 9 40 AA/AA/16:0 1.9 5.0 10 42 18:1ω9/18:2/AA, 18:1ω7/ 18:2/AA 10.6 12.8 11 42 AA/18:2/16:0, 20:3ω6/x/x 10.4 13.0 12 X2 0.4 1.3 13 44 18:1ω9/18:1ω9/AA 16.3 11.8 14 44 16:0/18:1ω9/AA, 18:0/18:2/AA 11.6 13.2 15 46 18:1ω9/18:1ω9/18:2 1.4 1.1 16 46 16:0/18:1ω9/18:2 1.8 1.0 17 46 16:0/16:0/18:2 tr tr 18 46 AA/18:0/18:1 0.8 1.8 19 48 18:1/18:1/18:1(ω9-major + ω7), tr 0.9 16:0/18:0/18:2 20 48 16:0/18:1/18:1 tr 0.7
X – unidentified molecular species; tr- traces The positional distribution of fatty acids was performed on the basis of the positional analysis of polar lipids, from which TAG is derived.
55 3.2 Role of extraplastidic lipids in TAG synthesis
As a part of the study on the involvement of the extraplastidic polar lipids (PC, PE, DGTS) in the biosynthesis of AA-rich TAG, it was important to assess the relative contribution of the pathways utilizing these lipids and the de novo Kennedy pathway, in supplying building blocks for TAG biosynthesis. We hypothesized that upon inhibition of the de novo synthesis of fatty acids by sethoxidim, an inhibitor of the acetyl-CoA carboxylase (ACCase) (Burton et al. 1991), the steady state level of the products of the Kennedy pathway would decrease and the existing polar lipids would support the biosynthesis of TAG during short-term experiments.
Alternatively, phospholipid –linked pathways can be minimized by growing the alga in phosphate-deprived medium to decrease the pool of phospholipids.
3.2.1 Sethoxidim treatment
The herbicide sethoxidim inhibits the multidomain plastidial acetyl-CoA carboxylase in weeds
(Graminae) by reducing malonyl-CoA supply for the de novo synthesis of fatty acids (Page et al.
1994), indicating a strong control by the ACCase-mediated reaction over fatty acid biosynthesis.
This inhibitor was tested with several species of green algae e.g., Dunaliella bardawil and
Haematococcus pluvialis (Rabbani et al. 1998, Zhekisheva et al. 2005). Severe inhibition of FAS was observed in the range of 10-50 µM sethoxydim. When logarithmically growing cells of P. incisa were transferred to nitrogen-free medium to enhance TAG accumulation, and maintained for 3 d in the presence of 200 µM of sethoxidim, dry weight and total fatty acid content increased only slightly in comparison to time-zero. In contrast, in the inhibitor-free control, dry weight and total fatty acid content increased significantly after 3 d of N-starvation (Table 7). The biomass content of TFA increased by 39.2% in the control, in comparison to only 9% in the presence of sethoxidim, indicating an inhibition of the de novo synthesis of fatty acids. However, under the sethoxidim treatment, the proportion of AA increased from 29.0 at time-zero to 38.1%
56 (43.4% in the control) after 3 d of N-starvation (Table 7). From the fatty acid content data
(µg/mg) (Table 8), it is clear that despite a little change in the TFA content, AA specifically increased in comparison to 16:0 and C18 unsaturated fatty acids, but less than in the N-starved control. The content of 16:0, 18:1ω9 (the products of the de novo synthesis of fatty acids) decreased or did not change much in comparison to time-zero (Table 8). These data may indicate to the contribution of pathways, involving preexisting fatty acids of polar lipids in the production of AA when de novo FAS is inhibited. The same conclusion can be derived from the data related to the volumetric content of fatty acids (data not shown).
Table 7. Effect of sethoxidim (200 µM) on fatty acid composition of nitrogen-starved culture of
P. incisa
Growth Fatty acid composition (% of total fatty acids) Condition 16:0 16:1 16:2 16:3 16:4 18:0 18:1 18:1 18:2 18:3 18:3 20:0 20:1 20:2 20:3 20:4 20:5 Others ω11 ω6 ω3 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 ω3 Zero-Time 16.3 0.8 5.6 2.3 0.9 1.6 7.4 3.6 23.9 1.2 4.7 0.3 0.2 0.2 1.1 29.0 0.8 3.9 3d N-starvation Control 11.9 0.6 2.2 0.9 0.3 1.6 12.9 3.4 16.8 1.1 2.2 0.2 0.4 0.3 0.8 43.4 0.8 2.0 Sethoxydim 13.0 1.9 3.6 1.2 0.7 1.2 10.6 3.4 19.8 1.0 2.7 0.1 0.3 0.4 0.8 38.1 1.0 2.5
Table 8. Effect of sethoxidim (200 µM) on the accumulation of biomass (DW) and fatty acids
(µg/mg DW) of nitrogen-starved cultures of P. incisa
Growth DW Fatty acid content (µg/mg DW) Condition mg/mL TFA 16:0 16:1 16:2 16:3 164 18:0 18:1 18:1 18:2 18:3 18:3 20:0 20:1 20:2 20:3 20:4 20:5 ω11 ω6 ω3 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 ω3 Zero-Time 3.3 112.0 18.2 0.9 6.2 2.5 1.0 1.8 8.3 4.0 26.8 1.3 5.3 0.3 0.2 0.2 1.2 32.4 0.9 3d N-starvation Control 5.0 156.0 18.6 0.9 3.4 1.4 0.5 2.6 20.2 5.3 26.2 1.8 3.4 0.3 0.6 0.5 1.2 67.9 1.3 Sethoxydim 4.3 122.0 15.9 2.3 4.4 1.4 0.8 1.4 12.9 4.2 24.1 1.3 3.3 0.1 0.4 0.4 1.0 46.6 1.3
The sethoxidim concentration used in this work was higher than that previously used for
Dunaliella bardawil and Haematococcus pluvialis. Concentrations lower than 100 µM were much less effective. This could be due to the presence of an adamant cell wall, which prevents the penetration of the inhibitor into the cells; alternatively the enzymes in this alga are less sensitive to sethoxydim.
57 3.2.2 Phosphate starvation
In attempts to highlight the role of extrachloroplastic polar lipids, phospholipids - PE and PC, and the phosphate-devoid betaine lipid - DGTS, as contributors of precursors for the synthesis of
AA-rich TAG, cultures grown on complete BG11 were transferred to phosphate-free BG11 medium. We hypothesized that severe phosphate limitation would result in a decrease of phospholipids to minimum levels, which will allow us evaluate the importance of phospholipids in the biosynthesis of AA-rich TAG and, simultaneously, the role of the non-phosphorous lipid
DGTS could be emphasized.
When a 6 d batch-cultivated culture was subjected to phosphate starvation, a significant decrease in the chlorophyll and somewhat lower biomass production were observed in comparison to control nutrient-complete medium (Fig. 13). Until 8 d, phosphate-starved cultures accumulated more TFA and AA (Fig. 13), but further, the production of TFA and AA was slowed down in the P-starved culture. After 8d, when control cultured reached a late stationary phase, volumetric chlorophyll (Chl), dry weight (DW) and fatty acid contents were not significantly different in two cultures. The P-starved cells were able to produce AA, however, the proportion of AA of total lipids was also not different from control culture, supplied with phosphate (Table 9). However, there were clear increases in proportion of 18:0 and 18:1 until day 6, indicating the impaired pathway of fatty acids through phospholipids. The data obtained reveal that under P-starvation, P. incisa produced less biomass and was not able to quickly accumulate storage lipids and AA, as shown for N-starvation. However, the culture was able to tolerate phosphate starvation for substantial period and still synthesize AA.
58 16 80 14 Dry weight (mg/mL)
60 12 g/mL) µ µ µ µ 10 40 8
6 Chlorophyll ( 20 4
0
1.5
1
Fatty acid (mg/mL) 0.5
0 02468101214 Time (days)
Figure 13. Comparison of chlorophyll (●, ○), dry weight (■, □), total fatty acid (♦, ◊) and AA
(▲, ∆) contents in control (closed symbols) and P-starved cultures (open symbols), respectively.
59 Table 9. Effect of phosphate starvation on fatty acid composition and content in P. incisa
Growth TFA AA Fatty acid composition (% of total fatty acids) Condition 16:0 16:1 16:2 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:2 20:3 20:4 20:5 % DW ω11 ω6 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 ω3 0d (6 d 14.5 4.7 12.1 1.1 1.7 1.4 4.0 15.9 3.2 18.4 1.6 3.4 0.3 1.6 32.6 0.9 stationary)
2d (+P) 11.6 4.5 12.6 2.2 1.9 1.8 3.0 8.1 3.3 18.3 1.3 4.8 0.3 1.0 38.7 1.2 2d (-P) 16.4 6.5 10.6 1.0 1.4 1.1 3.9 13.3 3.4 17.5 1.2 2.6 0.3 1.6 39.5 0.9
4d (+P) 12.3 4.7 13.5 1.1 2.3 1.3 3.6 9.4 3.5 17.8 1.6 3.3 0.3 1.3 37.9 1.1 4d (-P) 17.5 6.7 10.3 0.5 1.2 0.9 4.6 15.4 3.5 17.7 1.2 2.1 0.4 1.5 38.4 0.7
6d (+P) 15.1 6.1 12.1 0.8 2.0 0.9 4.0 11.6 3.5 16.9 1.5 2.3 0.3 1.3 40.2 0.9 6d (-P) 19.8 8.1 9.3 0.5 0.9 0.7 4.5 15.6 3.5 16.8 1.1 1.8 0.5 1.3 41.1 0.7
10d (+P) 21.5 9.4 10.5 0.5 1.3 0.6 3.3 15.2 3.6 15.0 1.2 1.5 0.4 1.2 43.8 0.7 10d (-P) 21.7 9.6 9.2 0.3 0.7 0.5 4.2 15.9 3.8 15.2 0.9 1.3 0.6 1.1 44.4 0.6
14d (+P) 25.4 11.3 10.0 0.5 1.1 0.5 3.0 16.9 3.7 14.3 1.0 1.2 0.5 1.1 45.8 0.7 14d (-P) 29.5 14.3 8.3 0.3 0.5 0.4 3.7 15.6 3.8 13.2 0.8 1.1 0.6 1.0 48.7 0.6
It was further important to analyze changes in distribution and fatty acid composition of major lipid classes. The lipid class analysis of control and P-starved cultures on 14 d showed that the proportion of PC in lipids decreased from 4.8 to 0.5% and PE decreased from 2.3 to 0.9%
(Table 10). The proportion of the non-phosphorous extrachloroplastic lipid DGTS instead increased sharply from 18.6 to 33.5% of total polar lipids. Among the chloroplastic lipids,
MGDG and DGDG decreased, whereas SQDG increased from 24.9 to 35.9%. However, the proportion of TAG was not significantly affected by phosphate starvation, comprising for 88.8 and 89.4% in control and P-starvation, respectively. Fatty acid composition of TAG reflected that of TFA (Table 9).
60 Table 10. Effect of phosphate starvation on distribution and fatty acid composition of polar lipids in P. incisa (cultures were grown in presence or absence of phosphate for 14 days)
Lipid Growth % of Fatty acid composition (% of total fatty acids) class condition PL** 16:0 16:1* 16:2 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:2 20:3 20:4 ω7 ω6 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 MGDG Con 18.0 4.5 1.4 27.8 8.8 0.6 4.2 0.7 33.2 0.6 11.6 0.4 0.2 4.2 -P 12.1 2.3 0.7 23.1 14.2 0.4 2.3 0.6 29.4 0.5 19.3 0.4 0.0 6.1
DGDG Con 18.0 23.7 1.3 7.8 0.6 0.8 5.5 1.6 41.0 0.7 5.5 0.4 0.4 9.8 -P 10.0 13.5 0.8 6.5 1.0 0.7 3.8 1.1 48.9 0.9 9.3 0.6 0.5 11.3
SQDG Con 24.9 62.9 0.1 0.0 0.0 0.8 2.8 11.4 18.1 0.3 1.7 0.3 0.2 1.1 -P 35.9 56.6 0.0 0.0 0.0 0.7 1.7 22.2 15.9 0.1 2.1 0.2 0.0 0.4
PG* Con 3.2 26.6 14.9 5.3 0.2 0.8 8.6 1.1 17.4 1.1 0.4 1.8 0.0 0.9 -P 1.1 14.1 21.9 12.9 2.1 2.5 6.1 1.5 11.9 0.7 1.2 1.5 0.0 0.7
DGTS Con 18.6 51.5 0.5 0.4 0.1 7.0 6.3 4.4 18.8 3.0 0.3 0.5 0.3 4.0 -P 33.5 42.6 0.2 0.0 0.0 7.8 3.1 5.0 24.8 5.7 0.4 0.4 0.1 7.2
PC Con 4.8 41.2 0.2 0.0 0.0 4.0 5.5 12.5 17.4 3.9 0.5 1.0 1.3 11.4 -P 0.5 29.5 1.3 0.0 0.0 7.8 4.6 10.4 21.0 5.7 0.7 2.6 2.0 14.5
PE Con 2.3 9.0 0.5 0.4 0.5 8.1 2.7 36.0 6.2 2.8 0.6 1.3 5.3 26.3 -P 0.9 9.9 1.0 0.0 0.5 9.2 3.4 22.6 9.0 3.4 0.8 2.0 5.0 30.4
PI Con 5.0 62.2 0.2 0.0 0.0 1.4 8.4 2.6 18.5 2.9 0.1 0.9 0.2 2.2 -P 1.9 57.1 0.4 0.0 0.0 2.2 5.3 2.1 22.4 3.0 0.1 0.9 0.4 5.4
PA Con 4.6 44.3 0.4 0.8 0.0 4.1 12.1 3.8 26.0 0.2 1.2 1.3 0.5 4.5 -P 3.8 35.6 0.5 1.0 0.0 4.8 10.2 2.7 34.6 0.3 2.2 1.0 0.5 5.3
* 16:1 trans ∆3 was observed only in PG and constituted for 20.8 and 22.1% in PG of control and P- starved culture, respectively.
** PL – Polar lipids
The fatty acid analysis of polar lipid classes revealed significant changes in their composition as a result of P-starvation (Table 10). Except for PE, all the polar lipid classes showed a decrease in the proportion of 16:0, and except for SQDG, all the polar lipids showed an increase in the proportion of AA. In the chloroplastic lipids, a decrease in proportions of 16:0 and 18:1ω9 and an increase in 18:3ω3 was observed. In MGDG, a decrease in 16:2 and 18:2 was
61 accompanied by increase in 16:3ω3 (from 8.8 to 14.2%) and 18:3ω3 (from 11.6 to 19.3%), respectively, indicating more chloroplastic ω3-desaturation.
In PC, the important phospholipid precursor of TAG biosynthesis, under P-starvation, the proportion of 16:0 decreased from 41.2 to 29.5%, indicating a decrease in the group of molecular species, which contain 16:0 at the sn-1 position (Table 3, section 3.1.2). However, the proportions of 18:0, 18:2, 18:3ω6, 20:2, 20:3ω6 and AA increased in PC, indicating that residual
PC was enriched in the C20/C20 and C18/C20 groups of molecular species. Similarly in PE,
18:1ω7, which resides at the sn-1 position of the predominant molecular species of PE (Table 4, section 3.1.3), decreased from 36.0 to 22.6%, indicating a decrease of this molecular species during P-starvation. However, an increase in the proportion of 18:1ω9, 18:2, 18:3ω6, 20:2, and
AA indicated that molecular species, which contain these fatty acids and involved in the desaturation of C18 and C20 fatty acids, were less affected. We can thus conclude that both residual PE and PC can still serve, at least partially as substrates for desaturations of C18 and
C20 precursors in AA biosynthesis.
In comparison to other phospholipids, the proportion of PA was decreased only slightly
(from 4.6 to 3.8%). In PA, similarly to PC, an increase in 18:2 and AA and a decrease in 16:0 were observed, suggesting that at least part of this phospholipid intermediate might derive from
PC by the activity of phospholipase D, a possible enzymatic step involved in catabolism of phospholipids during P-starvation. The non-phosphorous betaine lipid, DGTS, showed a decrease in the proportion of 16:0 (51.1 to 42.6%) and an increase in the proportion of 18:2
(from 18.8 to 24.8%), 18:3ω6 (from 3.0 to 5.7%) and AA (from 4.0 to 7.2%), indicating an increase in the ∆12 and ∆6 desaturations of C18 fatty acids.
62 3.2.3 P- and N-starvation
When cultures of P. incisa were starved both for phosphate and nitrogen (-P and -N), the proportion of AA of total fatty acids significantly decreased in comparison to the culture starved to N alone (-N) (Table 11). The decrease in AA was accompanied by the increase in 20:3ω6 and corresponding increases in 18:1 and 18:2, indicating the impaired AA biosynthesis. However, this finding indicates that P-deficiency does not significantly affect TAG accumulation under N starvation in this alga, but the resultant TAG contains substantially less AA and more of less unsaturated C18 fatty acids, indicating the importance of phospholipids in biosynthesis of AA.
Table 11. Effect of phosphate and nitrogen starvation on fatty acid composition and content in
P. incisa
Days Conditions TFA AA Fatty acid composition (% of total fatty acids) % DW 16:0 16:1 16:2 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:2 20:3 20:4 20:5 ω11 ω6 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 ω3 0 12.1 3.9 13.1 2.7 1.3 1.9 2.2 13.4 4.2 17.9 1.6 5.9 0.3 1.5 324 0.6 4 -N 23.3 10.7 10.9 0.8 0.4 0.6 2.5 17.4 4.1 11.1 1.2 1.8 0.2 1.4 45.9 0.9 7 -N 27.5 14.0 9.9 0.4 0.3 0.4 2.8 14.0 5.2 10.9 0.9 1.2 0.2 1.2 50.8 1.0 14 -N 30.6 16.5 9.9 0.5 0.3 0.3 2.6 12.1 5.6 10.3 0.8 0.8 0.3 1.1 53.9 1.0
4 -N -P 20.8 7.8 11.7 1.0 0.6 0.8 1.5 21.4 4.5 13.6 1.6 2.2 0.2 2.0 37.6 0.8 7 -N -P 30.9 11.7 10.7 0.4 0.3 0.5 1.5 25.9 4.3 12.8 1.1 1.4 0.2 1.8 38.0 0.7 14 -N -P 39.2 14.7 9.5 0.4 0.2 0.3 1.4 27.4 2.0 13.6 0.8 1.1 0.3 1.3 37.5 0.7
63 3.3 Enzymes of TAG biosynthesis
The unique biosynthesis of AA-rich TAG in P. incisa has prompted us to study the enzymes involved in this process. Although it was proposed that AA is transferred from PE and/or PC to
TAG (Bigogno et al. 2002c), the enzymatic steps involved are not known. The identification of
TAG biosynthetic enzymes in the microalga and characterization of their specificity and selectivity towards acyl substrates would help to understand the unique mechanism of AA channeling to TAG in this alga. Very little is known about the enzymes of TAG biosynthesis in algae. In plants, it is thought that the relative abundance of fatty acids in PL and TAG is determined by the selectivities and specificities of the acyltransferases that control the incorporation of fatty acids into TAG (Voelker and Kinney 2001). The main objective of the present study was to localize, determine and characterize the activity of DAGAT, which is committed to TAG biosynthesis and accomplish the final step of the acylation of DAG to form
TAG. Lipases, which initiate the metabolism and the catabolism of AA-rich TAG in P. incisa, were also detected and characterized. The determination of activity and characterization of these enzymes may be useful for future molecular biological and biotechnological applications. In order to achieve those goals, firstly it was necessary to develop a protocol for the cell homogenization and the cellular fractionation suitable for P. incisa.
3.3.1 Optimization of cell homogenization
Since the cells of P. incisa contain an adamant cell wall, homogenization was tested in a cooled
(4 °C) French press at 10,000 psi in HEPES buffer (pH 7.5). Under these conditions, we discovered high lipolytic activity that was indicated by the production of a large amount of free fatty acids in the lipid extract of the homogenate. Very little oil bodies and TAG were recovered, as observed by light microscopy and TLC analysis of the extract. Optimum conditions for the cell homogenization and the cellular fractionation were further determined in order to diminish
64 the lipolytic activity and to obtain the intact oil bodies. Homogenization of the biomass at pH 9.0 in a French press (10,000 psi) showed a significant decrease of lipolysis, since TAG was preserved in the lipid extracts of the homogenates (Fig. 14).
Figure 14. Effect of pH on the degree of TAG degradation in cell homogenates.
Total lipids were extracted from whole cells a) by the DMSO method (control); b), c), e) by the
Bligh-Dyer method (1959) from homogenates prepared in a French press under 10,000 psi at pH
6.0, 7.5 and 9.0, respectively; and d) from a homogenate prepared in a French press under 5,000 psi at pH 7.5. Lipids were resolved by TLC on silica gel plates in a system composed of hexane:diethyl ether:acetic acid 70:30:1 (v/v/v/v). The plate was charred with 5% H2SO4 in methanol at 180 °C.
A more gentle breakage of cells, aiming to limit organelle rupture and lipid degradation and to preserve the integrity of oil bodies, was achieved by freezing the biomass in liquid nitrogen and grinding to a fine powder in a mortar in CHES buffer (pH 9.0), containing the thiol- protecting reagent DTT, the protease inhibitor PMSF, catalase and glycerol. Moreover, the use of
5 mM EDTA, 1 mM EGTA and 1% (w/v) BSA to decrease lipolytic activity, as recommended by Chapman and Barber (1984), further reduced the degradation of TAG, leaving oil bodies intact (Table 12, Fig. 15).
65 Table 12. Composition of homogenization buffers tested to control the lipolytic activity during cell homogenization
K-Pia (100 mM) K-Pib (100 mM) HEPES (50 mM) CHES (50 mM) pH 7.4 pH 7.4 pH 7.5 pH 9.0 KCl (mM) - 20.0 20.0 20.0 MgCl2 (mM) - 2.0 2.0 2.0 EDTA (mM) 5.0 2.0 2.0 2.0 EGTA (mM) 1.0 - - - BSA (%, w/v) 1.0 0.5 0.5 0.5 Glycerol (v/v) - 10.0 10.0 10.0 Sucrose (mM) 0.25 0.25 0.25 0.25 Triton X-100 (w/v) - 0.005 0.005 0.005 PMSF (mM) - 1.0 1.0 1.0 DTT (mM) - 2.0 2.0 2.0 Catalase (units/mL) - 1000 1000 1000
80 70 60 50 40 30 % of total% of NL 20 10 0 abcd
Figure 15. Effect of the composition of the homogenization buffer on the degradation of TAG.
Results are presented as % of total NL.
Gray, black and white bars represent TAG, FFA and DAG, respectively. Cell homogenization in a) K-Pia buffer (pH 7.4), b) K-Pib buffer (pH 7.4), c) HEPES (pH 7.5), and d) CHES (pH 9.0).
Details of the composition of the buffers are given in the Table 13.
66 Total lipid of the homogenates were extracted by the Bligh-Dyer method (1959) and subjected to TLC in a NL system (see Materials and Methods). The amounts of neutral lipids were quantified by GC on the basis of spiked internal standard (C17:0) as mentioned in
‘Methods’.
3.3.2 Development of a protocol for cellular fractionation
Cellular fractionation was carried out by sequential centrifugation and a floatation discontinuous sucrose gradient. The cell-free homogenate, containing 0.6 M sucrose, was overlaid with 0.4 M and 0.2 M sucrose gradient buffers, and centrifuged for 1 h at 25,000 x g to purify oil bodies during their movement from the bottom to the surface of the gradient (Fig. 16). Oil bodies possess low density due to the accumulation of TAG, thus facilitating their purification by floatation (Huang 1984). The yellow floating layer of oil bodies was collected and purified by a second gradient centrifugation. The supernatant was utilized for the preparation of microsomes at
100,000 x g. Microsomes are a membrane fraction enriched in vesicles of circular membranes of the endoplasmic reticulum which were reported to locate various enzymes of TAG biosynthesis
(Stymne and Stobart 1987). The membrane pellet of 25,000 x g, which might contain different heavier membrane particles, was contaminated with chloroplast membranes indicated by its high chlorophyll content.
67
Figure 16. Outline of the cellular fractionation protocol of P. incisa.
Oil-bodies, observed under light microscopy, were coloured yellow-orange due to the presence of β-carotene, which was identified spectrophotometrically after extraction with diethyl ether (Fig. 17 and Table 13).
Figure 17. Isolated oil bodies under a light microscope (magnification x 400).
68 The acyl lipids of the oil bodies (Table 13) were composed of neutral lipids (96 ±%) 2.3 and polar lipids; β-carotene and proteins were minor components of oil bodies (Table 13). Oil bodies contained high proportion of AA (44 ± 4.5 % of total fatty acids).
Table 13. Parameters characterizing various components of oil bodies (The amounts of total fatty acids, β-carotene and protein was determined in aliquots of oil bodies suspensions (n=6), obtained by the developed protocol)
Parameter Value Ratio (TFA / β-carotene) 114 ± 29 Ratio (TFA / protein) 96 ± 28 NL (% of TFA) 96 ± 2.3 AA (% of TFA) 44 ± 4.5
TAG made up over 92% of the neutral lipid of oil bodies, the rest being DAG and MAG
(Table 14, Fig. 18).
Table 14. Distribution of neutral lipid (NL) lipid classes in oil bodies of P. incisa
Lipid % of total NL
TAG 92.4
FFA <1.0
DAG 6.6
MAG <1.0
69 Figure 18. TLC separation of total lipid extract of oil bodies.
Lipids were resolved in petroleum ether:diethyl ether:acetic
acid (70:30:1, v/v/v) and plates were sprayed with iodine
vapor to reveal unsaturated lipids. As evident from the
picture (left lane), TAG constituted the major lipid of the oil
bodies. Neutral lipid standards (right lane).
Microscopic observations confirming the integrity of oil bodies as well as the high AA and
TAG content in this fraction as determined by GC and TLC, respectively, allowed us to utilize the developed protocol for the cellular fractionation of P. incisa and minimize the possible contaminations of membrane fractions with the products of oil bodies degradation (FFA, oil bodies proteins).
The main purpose of this study was to determine, localize and characterize the activity of
DAGAT in the obtained cellular fractions, mainly in microsomes in oil bodies. Since DAGAT utilizes two substrates, acyl-CoA and DAG, the activity of the enzyme can be assayed by employing either of the radiolabelled substrates (Vogel and Browse 1996, Cases et al. 1998,
Bouvier- Nave et al. 2000) by measurement of their incorporation into TAG.
70 3.3.3 DAGAT assay utilizing [1_14C]oleoyl-CoA and non-labelled DAG
Cellular fractions were incubated in the DAGAT assay medium, containing 20 µM
[1_14C]oleoyl-CoA (20,000 DPM, 10,000 DPM/nmole), 400 µM 1,2 dioleoyl DAG, 0.125%
BSA and 100 mM Tris-HCl (pH 7.5), 100 µL final volume (Bouvier-Nave 2000). The production of labelled TAG was detected by Phosphoimaging (Fig. 19). Initially, we compared the specific activity of [1_14C]oleoyl-CoA incorporation into TAG in different fractions.
Different amounts of cellular fractions in terms of their protein content were used. It is important to note that both the cell-free homogenate and the soluble fraction contained BSA, thus, preventing the accurate determination of protein content and comparison of specific activities.
Figure 19. Incorporation of radioactivity from [1-14C]oleoyl-CoA into various lipids in presence of a) cell-free homogenate (270 µg protein), b) pellet 25,000 x g (30 µg protein), c) pellet
100,000 x g (30 µg protein), d) supernatant 100,000 x g (133 µg protein), e) oil bodies (7.5 µg protein).
71 Despite this obstacle, the higher specific activity of TAG formation was clearly observed in the microsomal fraction (Table 15). The fraction of oil bodies incorporated very little labelled acyl-CoA into TAG. It was difficult to increase the protein content of the oil bodies in the assay due to the limited volume of the assay mixture. The 25,000 x g pellet showed only a tiny amount of TAG synthesis in comparison to microsomes. Due to the presence of a high amount of protein in the soluble fraction, most of the total activity (84.5%) appeared in the soluble fraction; however, the specific activity was low. Microsomes represented 9.4% of the total activity.
Table 15. Specific and total activity of [1-14C]oleoyl-CoA incorporation into TAG in presence of cellular fractions of P. incisa
Specific activity Total activity Cellular fractions (pmol/min/mg protein) (pmol/min/mL culture) Cell-free homogenate 13.1 21.7 Oil bodies 6.5 0.2 Pellet 25,000 x g 18.7 1.1 Pellet 100,000 x g 96.3 2.1 Supernatant 100,000 x g 6.2 19.0
DAGAT has been reported as a membrane-bound protein located in the ER (Stymne and
Stobart 1987, Byers et al. 1999) and in membrane of oil bodies (Kamisaka and Nakahara 1994), originated in turn from ER membranes (Murphy, 2001). Thus, microsomes and oil bodies of P. incisa were utilized in the present work for the characterization of membrane-bound DAGAT.
The presence of activity in the soluble fraction seems to be an important observation, because the activity of soluble DAGAT was also discovered (Triki et al. 2000, Gangar et al. 2001).
However, the appearance of the activity in this fraction could have aroused from several artifacts, as incomplete sedimentation of microsomes, or solubilization of membrane-bound proteins in the presence of free fatty acids, possessing detergent properties. Indeed, very high amount of free fatty acids was produced in the DAGAT assay in the presence of the soluble fraction (Fig. 19).
72 The labelled lipids were resolved by the two-step one dimensional TLC that enabled to follow the radioactive lipid distribution in polar and neutral lipids simultaneously. As follows from Fig. 19, under the DAGAT assay conditions, several other lipids were also labelled in addition to TAG, indicating the activity of other enzymes involved in the acylation and interconversion of labelled lipids. Acyl-CoA is a common substrate for several acyltransferases and might acylate endogenous polar lysolipids, lyso-PA, lyso-PC (PE), lyso-DGTS and as well
DAG and MAG. It is important to note that there was a significant amount of FFA in all the fractions, likely due to the action of acyl-CoA thioesterase on the labelled substrate as frequently observed in DAGAT assays (Triki et al. 2000) and probably also due to the activity of lipases on labelled lipids.
Various cellular fractions were different in their lipid labeling profile. There was a very slight acylation of DAG, DGTS, PA and TAG with the 25,000 x g fraction. Little labeling of
DAG and DGTS was observed in the presence of the soluble fraction, and almost none in PC,
PE.
73 3.3.4 DAGAT activity in microsomes
3.3.4.1 Effect of time and protein
With microsomes, the incorporation of [1-14C]oleoyl-CoA into TAG was linear with time for 30 min (Fig. 20) and the protein content up to 60 µg. In the further experiments, DAGAT was assayed for 20 min with 25- 30 µg microsomal protein. A substantial synthesis of TAG was observed also in the absence of DAG (Fig. 20), indicating the presence of endogenous DAG in the membrane fraction. The endogenous DAG is a cause for the absence of DAG dependence in the DAGAT assays with microsomes and even with detergent-solubilized enzymes (Byers et al.
1999). However, the incorporation into TAG was significantly higher in the presence of 0.4 mM dioleoylglycerol (Fig. 20).
5
4
3
2 C]TAG formed 14 [ (nmol/mg protein) (nmol/mg 1
0
0 20406080 Time (min)
Figure 20. Incorporation of radioactivity from [1-14C]oleoyl-CoA into TAG in the absence (○) and presence (●) of 400 µM dioleoylglycerol.
3.3.4.2 Substrate specificity
The selectivity of DAGAT towards DAG substrates was studied by providing different DAG species to [1-14C]oleoyl-CoA. The dioleoyl molecular species of DAG was found to be the most preferred substrate for the reaction with oleoyl-CoA (Fig. 21).
74 250
200
150 C] TAG formed TAG C]
(pmol/min/mg) 100 14 [ 50
0 abcde
Figure 21. Substrate specificity of DAGAT towards four DAG species. Incorporation of radioactivity from [1-14C]oleoyl-CoA into TAG in a) in the absence of DAG; and in the presence of b) dioleoylglycerol; c) diarachidonylglycerol; d) dipalmitoylglycerol and e) dilinoleoylglycerol.
3.3.4.3 Solubility and availability of DAG substrate in assay
The low solubility of DAG in aqueous medium is known to be a major obstacle in the DAGAT assay. Since DAG forms micelles in an aqueous medium, making it unavailable for the acylation, the DAG stock solution (4 mM) routinely was prepared in 0.2% Tween-20, heated at 70 °C and immediately sonicated in order to break the micelles. The transparent DAG solution was added to the assay medium and it was again sonicated for 1 min before adding cellular fractions.
In attempt to increase the solubility of DAG, detergents CHAPS (0.1%, w/v), Triton X-
100 (0.1%, w/v) and solvent ethanol (10%, v/v) were used. The activity of TAG synthesis was severely inhibited in the presence of detergents (Fig. 22). The addition of ethanol significantly increased the incorporation of label into TAG (Fig. 22, lanes g and h).
75
Figure 22. Effect of detergents and ethanol on the incorporation of radioactivity from [1-
14C]oleoyl-CoA into TAG in the absence or presence of DAG. a) 0.02% Tween-20, b) 0.02% Tween-20 + 400 µM dioleoylglycerol; c) 0.1%, w/v CHAPS, d) 0.1%, w/v CHAPS + 400 µM dioleoylglycerol; e) 0.1%, w/v Triton, f) 0.1%, w/v Triton + 400 µM dioleoylglycerol; g) 10%, v/v ethanol, h) 10%, v/v ethanol + 400 µM dioleoylglycerol.
However, in the presence of ethanol, the dependence of TAG production on the exogenous DAG was not observed (Fig. 22, lanes g and h), likely due to an increase in the solubility and availability of endogenous DAG. The enhanced incorporation of label into TAG in the presence of ethanol can be explained by the better solubility of exogenous DAG that accepted most of the acyl-CoA, making it less available for other acyltransferases. Indeed, the labeling of other lipids (DAG, DGTS, PC) in the assay decreased (Fig. 22).
DAGAT activity in membrane fractions is often independent on exogenous DAG
(Weselake et al. 1999) and DAG dependence can be achieved only after delipidation of membranes with organic solvents in order to remove the endogenous DAG (Valencia-Turcotte and Rodríguez-Sotres 2001). Microsomes of P. incisa were delipidated by acetone and diethyl
76 ether at 4 °C. After diethyl ether treatment, the DAGAT activity of the delipidated proteins was low and was not reconstituted by the application of exogenous DAG. However, after acetone treatment, the DAGAT specific activity even though low, but was enhanced by the addition of
DAG. Differently from DAGAT of maize microsomes, DAGAT of P. incisa did not resist completely to acetone washing (Fig. 23), and the recovered activity was lower in comparison to untreated microsomes.
8
6
4 C] TAG formed C] TAG
14 a b [ 2 a b pmol/min/mg protein
0 Diethyl ether Acetone
Figure 23. The incorporation of [1-14C]oleoyl-CoA into TAG activity by delipidated microsomes in the absence of DAG (a), in the presence of 400 µM dioleoylglycerol (b).
3.3.4.4 Effect of thiol reagents
Thiol- modifying reagents were shown to affect DAGAT in microsomes of rats (Lehner and
Kuksis 1995). The inhibition of the activity of the enzyme with thiol-modifying reagents is presumed to indicate the presence of cysteine or serine residue(s) in the active site of an enzyme.
When the thiol reagent, p-chloromercuribenzene sulfonic acid (PCMB, 1 mM), was used,
DAGAT activity indeed was reduced over 50% in the presence of ethanol; PCMB also considerably inhibited the production of FFA (Fig. 24).
77 120
100
80
60
% of control 40
20 a b c dda b c 0 No EtOH 10% EtOH
Figure 24. Effect of PCMB on [1-14C]oleoyl-CoA into TAG and free fatty acid production in the presence and absence of ethanol.
Incorporation of radioactivity from [1-14C]oleoyl-CoA into TAG in (a) control and (b) in the presence of 1 mM PCMB; (c) released free [1-14C]oleic acid in control and (d) in the presence of
1 mM PCMB. Specific activities of TAG formation in the controls assays were 42.7 and 266.4 pmol/min/mg protein, in the absence and presence of ethanol, respectively.
3.3.4.5 Effect of cations
The bivalent cation Mg2+ was shown to affect DAGAT activity in microsomes (Byers et al.
1999). Commonly, a range of 0 to 3 mM of MgCl2 is utilized in DAGAT assays with plant microsomes (Hobbs et al. 1999, Valencia-Turcotte and Rodríguez-Sotres 2001). However, higher concentrations of MgSO4 and MgCl2 (25 and 10 mM, respectively) have been found to enhance DAGAT activity in microsomes of Brassica napus (Byers et al. 1999). Increase in
MgCl2 concentration above 3 mM showed an inhibition of DAGAT activity both in the presence and the absence of ethanol indicating an inactivation of DAGAT at higher Mg+2 (Fig. 25). The removal of Mg2+ cations by addition of the chelator EDTA increased the activity in the absence
78 of ethanol. It is believed that high concentrations of Mg2+ decrease the solubility of acyl-CoA and cause its aggregation in the medium (Constantinides and Steim 1985).
600
500
400
300
C]TAG formed 200 14 [ pmol/min/mg protein pmol/min/mg 100
0 + EDTA 0 3 10 150 MgCl (mM) 2
Figure 25. Effect of different concentrations of MgCl2 on DAGAT activity in the presence (grey bars) and absence of ethanol (black bars).
3.3.4.6 The origin of labelled DAG and the evidence for the MAGAT activity
We observed a significant production of labelled DAG in the DAGAT assay. Several sources may lead to the formation of DAG, the central intermediate in lipid metabolism. It could originate either from lipolysis of TAG, or derived from the polar lipids, PC, PE, PA, and DGTS.
However, so far, no enzymatic activity has been reported which produces DAG from DGTS.
Yet, the high labeling of DGTS, the major extraplastidic polar lipid in this alga, suggests an activity of the putative lyso-DGTS acyltransferase, which might contribute to further conversion of DGTS to DAG. Alternatively, DAG can be formed via the acylation of endogenous MAG or
DAG-derived MAG by the monoacylglycerol acyltransferase (MAGAT). MAGAT contributes significantly to the biosynthesis of TAG via an acyltransferase pathway in mammalian systems
79 (Cao et al. 2003). This enzyme has been recently discovered in microsomes of higher plants
(Tumaney et al. 2001) and proved to be involved in the synthesis of TAG via sequential transacylation of two molecules of DAG. Therefore, we conducted also experiments to characterize MAGAT activity in microsomes of P. incisa, which will be described in section
3.3.9.
3.3.5 DAGAT assay in microsomes with [1_14C]DAG and non-labelled oleoyl-CoA
As described in section 3.3.3, when labelled oleoyl-CoA was utilized in DAGAT assay several other lipids were labelled along with TAG, demonstrating the activity of different acyltransferases. On the other hand, the use of [1_14C] dioleoylglycerol and non-labelled acyl-
CoA could support the activity of DAGAT more clearly, avoiding undesirable artifacts. Indeed, when the cellular fractions were incubated in the same assay medium, but containing instead radiolabelled DAG and oleoyl-CoA, the incorporation of label into TAG was observed, apparently due to the activity of DAGAT (Fig. 26, Table 16).
Figure 26. Incorporation of radioactivity from
[1_14C]dioleoylglycerol into TAG in the presence of a)
cell free extract, b) pellet 25,000 x g, c) soluble fraction,
d) pellet 100,000 x g, e) control (no added protein).
Assay conditions: 100 µL of medium contained 100 mM
Tris HCl (pH 7.5), 400 µM [1_14C]dioleoylglycerol
(22,900 DPM), 0.02% Tween-20, 20 µM oleoyl-CoA,
0.125% (w/v) BSA, 30 µg protein, 10 min, 30 °C.
80
Among the cellular fractions, the highest specific activity was observed in microsomes.
The decrease in specific activity of DAGAT in microsomal fraction in comparison to cell- free homogenate might be due to inactivation of enzymatic activity in microsomes during fractionation or storage. In addition to TAG, some other lipids, including MAG and PC, were also labelled. MAG could be labelled due to the activity of DAG lipase or DAG:DAG transacylase, whereas labeling of PC could be due to DAG-CPT. The high amount of FFA obtained, indicated the activity of lipases, most probably on the labelled DAG. The activity of
DAGAT in oil bodies was determined separately before and after delipidation by organic solvents, which will be described under a separate heading.
Table 16. Specific activity of [1-14C]1,2-dioleoylglycerol incorporation into TAG in the presence of cellular fractions of P. incisa
Cellular fractions Specific activity (nmol/min/mg protein) Control 0.4 Cell-free homogenate 8.6 Pellet 25,000 x g 1.4 Pellet 100,000 x g 2.0 Supernatant 100,000 x g 0.7 Oil bodies n.d.* *n.d. – non determined
3.3.5.1 Protein and time dependence
The activity of DAGAT in the microsomal fraction was linear up to about 30 µg protein (Fig. 27) and up to 30 min (Fig. 28). The assays were conducted within this time and protein content range.
81
15 30 45 60 µg protein
15
10
(pmol/min)
C] TAG formed 5 14 [
0
0 15304560
Protein (µg) Figure 27. Effect of protein content on the incorporation of radioactivity from [1_14C]1,2- dioleoylglycerol into TAG.
10 15 30 45 60 90 min
20
15
10 C] TAG formed formed TAG C] 4 nmol/mg protein nmol/mg
[1 5
0 0 153045607590 Time (min) Figure 28. Time-dependence of the incorporation of radioactivity from [1_14C]1,2- dioleoylglycerol into TAG.
82 3.3.5.2 Effect of oleoyl-CoA concentrations
We observed a synthesis of TAG even without the application of acyl-CoA, indicating either the presence of endogenous acyl-CoA, or acyl-CoA independent TAG synthesis. However, when oleoyl-CoA was added, a significant enhancement in the TAG synthesis was observed, indicating the activity of DAGAT (Fig. 29). The TAG synthesis reached maximum at oleoyl-
CoA concentration of 30 µM.
200
150
100 % to control% to C] TAG C] formed 14 [ 50
0 010203040 Oleoyl-CoA (µM)
Figure 29. Effect of concentration of oleoyl-CoA on the incorporation of radioactivity from
[1_14C]1,2-dioleoylglycerol into TAG.
The specific activity in the absence of oleoyl-CoA (control) was 350 pmol/min/mg protein.
The results shown are based on the average of three independent experiments.
83 3.3.5.3 Selectivity of acyl-CoA substrates
When four types of acyl-CoAs were compared in assay with [1_14C]1,2-dioleoylglycerol, the higher TAG production was observed in the presence of linoleoyl- and oleoyl-CoA, indicating some selectivity of DAGAT towards the C18 rather than C16 and C20 acyl-CoA substrates (Fig.
30).
0.7
0.6
0.5
0.4
0.3 C] TAG formed 14
[ 0.2 nmol/min/mg protein 0.1
0 No acyl-CoA 20:4-CoA 16:0-CoA 18:1-CoA 18:2-CoA
Figure 30. Incorporation of radioactivity from [1_14C]1,2-dioleoylglycerol to TAG in the presence of different acyl-CoA substrates.
84 3.3.5.4 Effect of ethanol
The addition of ethanol (10%, v/v) drastically increased the incorporation of radioactivity from
[1-14C]1,2-dioleoylglycerol into TAG in the presence of 20 µM oleoyl-CoA (Fig. 31), which is in accord with our results on the enhancement of the DAGAT activity by ethanol in the assay with [1-14C]oleoyl-CoA (Fig. 22).
800
600
400 % of control% of C] TAGC] formed 14 [ 200
c abc a b 0 No EtOH 10% EtOH
Figure 31. Effect of ethanol and PCMB on the incorporation of radioactivity from [1_14C]1,2- dioleoylglycerol into TAG. a) Control (no addition); b) 20 µM oleoyl-CoA; c) 20 µM oleoyl-
CoA + 1 mM PCMB. The specific activity of the control (in the absence of ethanol) was 1.08 nmol/min/mg protein.
3.3.5.5 Effect of thiol-reagent PCMB
The thiol-reagent PCMB inhibited the DAGAT activity in the assay with radio-labelled oleoyl-
CoA (Fig. 25), thus we tested the effect of this reagent also on the incorporation of radioactivity from [1_14C]1,2-dioleoylglycerol into TAG, in the presence of acyl-CoA. PCMB considerably inhibited the enhancement of TAG synthesis caused by the addition of oleoyl-CoA (Fig. 31), both in the absence or presence of ethanol, indicating an inhibition of DAGAT activity.
85 3.3.5.6 Effect of salts
In the course of the characterization of the DAGAT activity in microsomes, an increase in TAG synthesis from [1_14C]1,2-dioleoylglycerol was observed with an increase in the concentration of
_14 2+ MgCl2 (Fig. 32). In contrast, in the assays utilizing [1 C]oleoyl-CoA, increasing the Mg concentration up to 150 mM, inhibited the TAG synthetic activity (Fig. 25), in accordance with a decrease in the solubility of acyl-CoA at high Mg2+ concentration (Constantinides and Steim
1985). The addition of 150 mM KCl did not affect TAG synthesis; however, both 10 mM ZnCl2 and 10 mM FeCl3 completely inhibited the activity (data not shown).
2
1.8
1.6
1.4 C] TAG formed TAGC] 14 [ nmol/min/mg protein nmol/min/mg 1.2
1
0 20406080100 MgCl (mM) 2
Figure 32. Effect of the concentration of MgCl2 on the incorporation of radioactivity from
[1_14C]1,2-dioleoylglycerol into TAG.
Furthermore, at the high Mg2+ concentration (150 mM), TAG synthesis was independent of exogenous acyl-CoA, while in the absence of Mg2+, TAG synthesis was dependent on exogenous acyl-CoA (Fig. 33). These observations indicated the presence of an acyl-CoA independent TAG synthetic activity utilizing DAG in the microsomes. An acyl-CoA independent
86 TAG formation by a transacylation reaction between two molecules of DAG, releasing a molecule of MAG, has been reported in plants (Stobart et al. 1997) (Fig. 7, Chapter-Introduction, section-1.4.5). Further, we tested factors affecting an acyl-CoA independent TAG formation by microsomes.
2.5
2
1.5
1 C] TAG formed formed TAG C] 14 [ nmol/min/mg protein 0.5
0 abcd
_14 Figure 33. Effect of MgCl2 on incorporation of radioactivity from [1 C]1,2-dioleoylglycerol into TAG in the absence and the presence of 20 µM oleoyl-CoA. a) and b), without and with addition of 20 µM oleoyl-CoA in the absence of MgCl2; c) and d), without and with addition of 20 µM oleoyl-CoA in the presence of 150 mM MgCl2.
Figure 33 represents the mean of three replicates.
87 3.3.6 Acyl-CoA independent TAG synthesis from [1_14C]1,2-dioleoyglycerol
3.3.6.1 Time and protein dependence
Acyl-CoA independent TAG synthesis from [1_14C]1,2-dioleoyglycerol was assayed at 150 mM
MgCl2, and was linear within 15 min (Fig. 34) and within protein content up to 40 µg (Fig. 35).
40
30
20 C] TAG formed nmol/mg protein 14 [ 10
0 0 5 10 15 20 25 30 Time (minutes) Figure 34. Time-dependence of the incorporation of [1_14C]1,2-dioleoylglycerol into TAG at 150 mM of MgCl2.
0.15
0.1 nmol/min
C] TAGC] formed 0.05 14 [
0 0 5 10 15 20 25 30 35 40 Protein (µg) Figure 35. Protein dependence of the incorporation of [1_14C]1,2-dioleoylglycerol into TAG at
150 mM of MgCl2.
88 3.3.6.2 pH and temperature dependence pH and temperature optima were observed at 7.5 and 30 °C, respectively (Fig. 36 and Fig. 37, respectively).
120
100
80
60 % of control % of C] TAG C] formed
14 40 [
20
0 456789 pH
Figure 36. Effect of pH on incorporation of radioactivity from [1_14C]1,2-dioleoylglycerol into
TAG at 150 mM MgCl2.
3
2.8
2.6
2.4
C] TAG formed TAG C] 2.2 14 [ nmol/min/mg protein nmol/min/mg 2
1.8
20 25 30 35 40 ο Temperature ( C)
Figure 37. Effect of temperature on incorporation of radioactivity from [1_14C]1,2- dioleoylglycerol into TAG at 150 mM MgCl2.
89 In conclusion, the acyl-CoA independent TAG formation can be assayed under conditions similar to DAGAT, e.g. pH, temperature, components of the assay, but requires high
MgCl2 and does not require acyl-CoA.
3.3.6.3 Effect of thiol reagents, PCMB, DTT and CuCl2
_14 The incorporation of radioactivity from [1 C]1,2-dioleoylglycerol into TAG at 150 mM MgCl2 was not inhibited by PCMB and was enhanced to some degree by the thiol-protecting reagent
DTT (Fig. 38), while it was inhibited by 1 mM PCMB in the absence of MgCl2 (Fig. 31).
However, 1 mM CuCl2 sharply inhibited the activity (data not shown).
Figure 38. Effect of 20 µM oleoyl-CoA (b), 1 mM PCMB (c) and 1 mM DTT (d) on the
_14 incorporation of [1 C]1,2-dioleoylglycerol into TAG in the presence of 150 mM MgCl2; a) control (no acyl-CoA).
90 3.3.6.4 Distribution of activity in the cellular fractions
When the incorporation of radioactivity from [1_14C]1,2-dioleoyglycerol to TAG in the presence of 150 mM MgCl2 was tested in different cellular fractions, highest specific activity was observed in microsomes (Fig. 39, Table 17).
Figure 39. Incorporation of radioactivity from [1_14C]dioleoylglycerol into TAG in the presence of 150 mM MgCl2 in a) cell-free homogenate, b) pellet 25,000 x g, c) supernatant 100,000 x g, d) pellet 100,000 x g.
Table 17. Specific activity of incorporation of radioactivity from [1-14C]1,2-dioleoylglycerol into
TAG in the presence of various cellular fractions of P. incisa at 150 mM MgCl2
Cellular fractions Specific activity (nmol/min/mg protein) Cell-free homogenate 0.39 Pellet 25,000 x g 1.20 Pellet 100,000 x g 3.28 Supernatant 100,000 x g 0.12 Oil bodies n.d.*
In conclusion, evidences concerning the activities of two different types of TAG synthesis (acyl-CoA dependent and independent) from radioactive DAG were obtained in microsomes of P. incisa. Our data may suggest the involvement of both DAGAT and DGTA activities in the biosynthesis of TAG in this alga.
91 3.3.7 DAGAT activity in oil bodies in assay with [1-14C]oleoyl-CoA
Oil bodies failed to incorporate [1-14C] oleoyl-CoA into TAG in the presence or absence of ethanol (Fig. 19, Table 15). However, when oil bodies were delipidated by sequential extractions with diethyl ether to remove the bulk of NL, the residual proteins were able to incorporate labelled acyl-CoA into TAG and the rate of incorporation was dependent on the concentration of
DAG (Fig.40).
200
180
160
140 C] TAG C] formed 14 [
pmol/ min/ mg protein mg min/ pmol/ 120
100
0 0.1 0.2 0.3 0.4 Dioleoylglycerol (mM)
Figure 40. Effect of concentration of dioleoylglycerol on the incorporation of radioactivity from
[1-14C]oleoyl-CoA into TAG by delipidated proteins of oil bodies.
Assay conditions: 20 µM oleoyl-CoA (20,000 DPM), 0.4 mM dioleoylglycerol, 0.02% Tween 20
(w/v), 0.25% BSA (w/v), 100 mM Tris-HCl buffer pH 7.5 in 100 µL; 30 min, 30 °C.
The time and protein dependence of TAG synthesis by delipidated proteins of oil bodies are presented in Fig. 41 and 42. A linear TAG synthesis was observed over a long incubation period of 120 min. TAG synthesis was saturated at protein concentration higher than 3 - 4 µg in
100 µL of assay.
92 20
15
10 C] TAG formed TAG C] nmol/mg protein nmol/mg 14
[ 5
0
0 30 60 90 120 Time (min)
Figure 41. Time dependence of the incorporation of radioactivity from [1-14C]oleoyl-CoA into
TAG by delipidated proteins of oil bodies.
1
0.8
0.6
pmol/min 0.4 C] TAG C] formed 14 [ 0.2
0 024681012 protein (µg)
Figure 42. Protein dependence of the incorporation of radioactivity from [1-14C]oleoyl-CoA into
TAG by delipidated proteins of oil bodies.
Further, we compared the incorporation of [1-14C]oleoyl-CoA into TAG in the presence of two DAG species: dioleoylglycerol and diarachidonoylglycerol (Fig. 43). Between two DAG species, dioleoylglycerol was the better acyl acceptor.
93 100
75
50 % of control C] TAGC] formed 14 [ 25 abc 0
Figure 43. Effect of delipidation and fatty acid composition of DAG on the incorporation of radioactivity from [1-14C]oleoyl-CoA into TAG by oil bodies proteins
(a) dioleoylglycerol with delipidated oil bodies proteins (control);
(b) diarachidonoylglycerol with delipidated oil bodies proteins;
(c) dioleoylglycerol with native oil bodies
3.3.8 DAGAT assay in native oil bodies with [1-14C]dioleoylglycerol
Under assay conditions for DAGAT with [1-14C]dioleoylglycerol, differently from the assay with [1-14C]oleoyl-CoA, native oil bodies were able to catalyze the incorporation of [1-
14C]dioleoylglycerol into TAG in the presence of 20 µM oleoyl-CoA (Fig. 44).
94 300
200 C] TAG formed TAG C] nmol/mg protein 14
[ 100
0
0 5 10 15 20 25 30 Time ( min)
Figure 44. Time dependence of the incorporation of radioactivity from [1-14C]dioleoylglycerol into TAG by oil bodies.
Assay conditions: 20 µM oleoyl-CoA, 0.4 mM dioleoylglycerol (20,000 DPM), 0.02% (w/v)
Tween 20, 0.25% (w/v) BSA, 20% glycerol, 100 mM Tris-HCL buffer pH 7.5 in 100 µL, 2.4 mg protein, 30 °C.
3.3.8.1 Factors affecting DAGAT activity in oil bodies in assay with [1-14C]dioleoylglycerol
While optimizing DAGAT assay for oil bodies, we found out that addition of 0.25% BSA and
20% glycerol enhanced the activity. In the presence of 3 mM MgCl2 (Hobbs and Hills 2000), the activity was reduced. The same was true for 10 mM MgCl2, but one should note the higher activity in the presence of 10 mM in comparison to 3 mM MgCl2 (Fig. 45).
Since the detergent CHAPS was frequently utilized for the DAG preparation for DAGAT assays (Wiberg et al. 1994, Lehner and Kuksis 1995), we tested this detergent. However, replacement of Tween-20 with CHAPS inhibited the activity (Fig. 46).
95 120
100
80
60
% of control % of 40
20 abcde 0
Figure 45. Effect of components of the assay mixture on the incorporation of radioactivity from
[1-14C]dioleoylglycerol into TAG by oil bodies. a) control; b) no BSA; c) no glycerol; d) 3 mM MgCl2; e) 10 mM MgCl2.
Control assay contained 0.25% (w/v) BSA, 20% (v/v) glycerol and no MgCl2. (Specific activity of control was 6.2 nmol/min/mg protein).
120
100
80
60
% of control 40
20 abcde 0
Figure 46. Effect of different chemicals on the incorporation of radioactivity from [1-
14C]dioleoylglycerol into TAG by oil bodies. a) control (as in Fig. 45), b) CHAPS, c) 1 mM DTT, d) 1 mM PCMB and e) 1 mM nicotinic acid
We also tested the effect of several chemicals known to affect DAGAT in various systems. As follows from Fig. 46, nicotinic acid (an inhibitor of DAGAT in mammalian
96 systems) (Ganji et al. 2004), as well as the thiol-modifying reagents, PCMB and DTT, suppressed the activity. The severe concentration-dependent inhibition by PCMB (Fig. 47) suggests the presence of a catalytically important serine or cysteine group (Lehner and Kuksis
1995). The inhibitory effect of DTT, which protects SH-group is difficult to explain.
100
75
50 % ofcontrol C] TAG formed TAG C] 14 [ 25
0 0.0 0.01 0.1 1.0 PCMB (mM)
Figure 47. Effect of PCMB on the incorporation of radioactivity from [1-14C]dioleoylglycerol into TAG by oil bodies.
Similarly to microsomes, increasing MgCl2 concentration to 150 mM drastically enhanced the incorporation of [1-14C] 1,2 dioleoylglycerol into TAG in oil bodies (Fig. 48) but it was completely abolished by 1mM PCMB. At 150 mM MgCl2, oil bodies could utilize [1-
14C]1,2 dioleoylglycerol as a sole acyl donor for the synthesis of TAG and the acyl-CoA independent incorporation of DAG to TAG seems to be the dominant activity under the assay
14 conditions in the presence of 150 mM MgCl2. However, [1- C]1,2 dioleoylglycerol incorporation into TAG was enhanced by the addition of oleoyl- and arachidonoyl-CoAs (Fig.
49).
97
_14 Figure 48. Effect of addition of MgCl2 and PCMB on the incorporation of [1 C]1,2- dioleoylglycerol to TAG by oil bodies. a) control (absence of MgCl2); b) 150 mM MgCl2; c) 150 mM MgCl2 + 1 mM PCMB.
200
150
100 (% to control) to (% C] TAG formation 50 14 [
0 abc
Figure 49. Incorporation of radioactivity from [1_14C]1,2-dioleoylglycerol to TAG in the absence and presence of different acyl-CoAs at 150 mM MgCl2. a) control (no acyl-CoA); b) oleoyl-CoA; c) arachidonoyl-CoA
Based on average of two independent assays. Specific activity of control was 34 nmol/min/mg protein
98 pH optima for the incorporation of [1-14C]1,2 dioleoylglycerol into TAG in the oil bodies was observed at 7.5 similarly to microsomes (Fig. 50).
40
30
20 C] TAG formed 14 [ 10 nmol/min/mg protein
0 456789 pH
Figure 50. Effect of pH on the incorporation of [1-14C]dioleoylglycerol into TAG by oil bodies in the presence of 150 mM MgCl2. Na-Succinate, Na-MES, TRIS-HCL, Na-HEPES and Na-
CHES were utilized in different pH ranges.
99 3.3.9 MAGAT activity in microsomes
As mentioned above (Section 3.3.4.6), the formation of labelled DAG in the DAGAT assay with
[1-14C]oleoyl-CoA suggested MAGAT activity, utilizing MAG, either endogenous or produced via enzymatic conversion of DAG. Indeed, when microsomes were incubated with [1-14C]oleoyl-
CoA and sn-2-monooleoylglycerol (MOG) as acyl acceptor, significantly higher formation of labelled DAG was observed (Fig. 51). The formation of DAG was considerably higher in comparison to the assay conducted in the absence of MOG.
Figure 51. Incorporation of radioactivity from [1-14C]oleoyl CoA into DAG in the absence (lane a) or the presence (lane b) of sn-2-monooleoylglycerol. The brightness of the phosphoimage was adjusted to emphasize the changes in DAG labeling.
Assay conditions: 30 µg microsomal protein were added to a well sonicated mixture of 20 µM
[14C]oleoyl-CoA (21,000 DPM), 200 µM sn-2-monooleoylglycerol, 0.02% Tween-20, 0.125%
BSA and 100 mM Tris-HCl (pH 7.5) in 100 µL for 30 min at 30 °C.
100 3.3.9.1 Effect of MAG concentration
The incorporation of radioactivity from [14C]oleoyl-CoA into DAG was dependent upon the concentration of the sn-2-monooleoylglycerol (Fig. 52). Maximal specific activity was observed at a concentration of 200 µM. There was an activity in the absence of exogenous MAG, which indicated the presence of endogenous MAG in microsomes.
100
90
80 C] DAG formation DAG C] pmol/mg protein pmol/mg 14 [ 70
60 0 100 200 300 400 sn-2-monooleoylglycerol (µM)
Figure 52. Effect of concentration of sn-2-monooleoylglycerol on the incorporation of radioactivity from [14C]oleoyl-CoA into DAG.
3.3.9.2 Time dependence
The MAGAT activity was linear within 15 min (Fig. 53). In the absence of exogenous MAG, the incorporation of [14C]oleoyl-CoA into DAG during the time course increased only slightly, indicating that the concentration of endogenous MAG was not sufficient. Interestingly, the incorporation of [14C]oleoyl-CoA into TAG did not increase when MOG was applied to the assay.
101 10 A 8
6
4 C] DAG formation DAG C] nmol/mg protein
14
[ 2
0
B 8
6
4 C] TAG formation TAG C] nmol/mg protein nmol/mg
14 2
[
0
0 153045607590 Time (min)
Figure 53. Time dependence of incorporation of radioactivity from [14C]oleoyl-CoA into DAG
(A) and TAG (B) in the presence (●) or absence (○) of sn-2-monooleoylglycerol.
3.3.9.3 Effect of ethanol
Further, we compared the mode of administration of MAG into the assay medium. As can be seen from Fig. 54, DAG synthesis was similar when MOG was added as an ethanol solution
(followed by the solvent evaporation) or as a Tween 20-containing solution. However, the addition of ethanol (10%, v/v) into the MAGAT assay reduced the incorporation of [14C]oleoyl-
CoA into DAG, while TAG synthesis was significantly enhanced (Fig. 54 and Fig. 55).
102
bm a b c
Figure 54. Effect of mode of MOG administration on the incorporation of [14C]oleoyl CoA into
DAG. bm) boiled microsomes (non-enzymatic control); a) MOG added as ethanol stock, followed by the solvent evaporation; b) MOG in 0.02% Tween-20; c) MOG in 0.02% Tween-20 + 10% (v/v) ethanol.
100
80
60
40 C] DAG formation DAG C] 14 [ pmol/min/mg protein pmol/min/mg 20
0 abc
Figure 55. Effect of ethanol on the incorporation of radioactivity from [14C]oleoyl CoA into
DAG. Column headings as in Fig. 54.
104 3.3.9.4 Effect of dioleoylglycerol addition
When DAG (0.4 mM) was present in assay, labeling of DAG increased (Fig 56). It is possible that DAG was converted to MAG by a lipase activity or by DAG:DAG transacylase. MAG was further converted to DAG by the activity of MAGAT.
5
4
3
2 C] DAG formed formed C] DAG 14 nmol/mg protein nmol/mg
[1 1
0 0 153045607590 Time (min)
Figure 56. Incorporation of radioactivity from [14C]oleoyl-CoA into DAG in the presence (●) or absence (control, ○) of dioleoylglycerol during the time course.
Assay conditions: 20 µM [14C]oleoyl-CoA (21,000 DPM), 400 µM 1,2-dioleoylglycerol, 0.02%
Tween-20, 0.125% BSA and 100 mM Tris-HCl (pH 7.5) in 100 µL, 30 µg microsomal protein,
30oC.
3.3.9.5 Effect of MgCl2
DAG produced in the MAGAT assay was not further converted to TAG. As shown above
(Section 3.3.6.3), when MgCl2 increased, a transacylation reaction between two DAG molecules via DGTA was favored. Indeed, when MgCl2 (150 mM) was added to the MAGAT assay after
15 min of incubation, enhanced synthesis of TAG from labelled DAG was observed (Fig. 57).
105 These results suggest that DAG produced under the MAGAT assay conditions, was not available for DAGAT, but was probably utilized by DGTA, following the shift in Mg2+ concentration.
Figure 57. Effect of the shift in MgCl2 concentration on incorporation of radioactivity from
[14C]oleoyl-CoA into TAG in the MAGAT assay.
Lane a) control (30 min),
Lane b) addition of 150 mM MgCl2 after 15 min.
106 3.3.10 Lipolytic activity in microsomes
3.3.10.1 pH and temperature
During the study on the conversion of [1_14C]1,2-dioleoylglycerol to TAG, the production of free fatty acids was observed, indicating the presence of a lipase, hydrolyzing DAG and, possibly, also TAG. Since temperature and pH optimum for the lipolysis of DAG were observed at 37 °C and 4.5, respectively (Fig. 58, 59, 60), the assays for the lipolysis of DAG were carried out in a succinate buffer (100 mM, pH 4.5). FFA were the principal product, however, MAG was also observed, indicating that the lipolytic activity can release labelled fatty acids from both positions of [1_14C]1,2-dioleoylglycerol.
Figure 58. Effect of pH on the release of radioactivity from [1_14C]1,2-dioleoylglycerol into
[1_14C]oleic acid. a) pH 4.5 (Succinate buffer), b) pH 7.5 (Tris buffer) and c) pH 9.0 (Ches buffer).
Assay conditions: 400 µM dioleoylglycerol (30,000 DPM ), 0.02% Tween-20, 100 mM pH buffer in 100 µL, 1 h, 30 µg protein.
As follows from the image, DAG lipolysis was minimal at pH 7.5 (conditions for
DAGAT assay) and at pH 9.0 (conditions for cell homogenization), respectively (Fig. 58).
107 25
20
15 C] oleic acid oleic C]
14 10 [1- nmol/min/mg protein nmol/min/mg 5
0 3456789 pH
Figure 59. Effect of pH on the release of radioactivity from [1_14C]1,2-dioleoylglycerol into
[1_14C]oleic acid. Succinate, MES, TRIS, HEPES and CHES buffers were utilized for
different pH ranges.
Lipolysis of DAG was observed at a wide range of temperatures, with 37 °C being optimal (Fig. 60). A significant hydrolysis of DAG occurred even at 55 °C and was not completely abolished at 77 °C.
108 14
12
10
8
6 C] oleic acidC] 14
[1- 4 nmol/min/mg protein nmol/min/mg 2
0 20 30 40 50 60 70 80 Temperature (oC)
Figure 60. Effect of temperature on the release of radioactivity from [1_14C]1,2-dioleoylglycerol into [1_14C]oleic acid.
109 3.3.10.2 Time and protein
A linear time dependence of the lipolysis of DAG was observed during 45 min of incubation and the lipolysis was linear with protein concentration under the conditions of assay (Fig. 61,62).
1400
1200
1000
800
600
400 nmol/mg protein C] oleic acid released acid oleic C] 14 [ 200
0 0 1020304050 Time (min)
Figure 61. Time dependence of the release of radioactivity from [1_14C]1,2-dioleoylglycerol into
[1_14C]oleic acid.
0.6
0.5
0.4
0.3 C]oleic acid C]oleic 14 nmole/min 0.2 [1-
0.1
0
0 5 10 15 20 25 30 Protein (µg) Figure 62. Effect of protein content in assay on the release of radioactivity from [1_14C]1,2-
dioleoylglycerol into [1_14C]oleic acid. Assay conditions: 400 µM dioleoylglycerol (24,000
DPM), 0.2% Tween-20, 100 mM succinate buffer (pH 4.5) in 100 µL, 15 min, 37 °C.
110 3.3.10.3 Effects of divalent metals and inhibitors on DAG lipase activity
Further, we tested a number of compounds known to affect the activity of lipases. The activity was inhibited by several thiol-modifying reagents, PCMB, NEM and iodoacetamide at a concentration of 1 mM and was stimulated by the thiol-protecting chemical DTT, suggesting the importance of SH-group containing amino acids for the enzyme activity (Table 18).
Table 18. Effects of divalent metals and inhibitors on DAG lipase activity.
Chemical Concentration Activity (mM) % of control Control 100 Thiol-reagents PCMB 1 58 NEM 1 75 Iodoacetamide 1 68 DTT 1 122 Cations MgCl2 2.5 124 CaCl2 2.5 80 CuCl2 2.5 50 EDTA 1 95 MgCl2 150 80
DAG-lipase inhibitor RHC 80267 (µM) 125 49 Free amino group reagent DIDS (µM) 100 27
Among divalent salts tested, only Mg2+ was effective in stimulating the activity. This lipase probably does not require Ca2+. The inhibition by Cu2+ is in line with the inhibition by SH- group- modifying reagents. Incubation with DIDS (100 µM), which interacts with free amino groups of basic amino acid residues, also inhibited the release of FFA from DAG, suggesting the potential involvement of a free amino group in the acyl-hydrolyzing reactions.
During studies on lipolytic enzymes in microsomes, we observed, along with FFA, also a release of labelled MAG. MAG can be an intermediate in the hydrolysis of DAG. MAG may be also a product of a specific DAG-lipase. The DAG-lipase is well studied in mammalian systems,
111 since it is involved in the release of AA from DAG, derived from phospholipids, and is an important component in the cell signaling. Thus, we tested the effect of RHC 80267, a specific inhibitor of DAG-lipase on the lipolytic activity in our assays. RHC 80267 (125 µM) inhibited the production of [1_14C]FFA from [1_14C]1,2-dioleoylglycerol up to 50%, indicating the activity of a DAG-lipase in the microsomes (Table 18).
3.3.10.4 Positional specificity
[1_14C]1,2-dioleoylglycerol was mainly hydrolyzed to the free oleic acid, indicating that lipase activity did not differentiate between two sn-positions of DAG when DAG contained C18 fatty acids at both positions. The mammalian sn-1–specific DAG lipase catalyses the hydrolysis of sn-
2-AA- DAG species to 2-arachidonoylglycerol (2-AG), the most abundant endocannabinoid in mammalian tissues (Bisogno et al. 2003). In microsomes of P. incisa, when arachidonoyl- containing 1-stearoyl-2-[14C] arachidonoyl-glycerol was utilized, AA was not released as a FFA, but was preserved in the form of MAG (2-arachidonoylglycerol) (Fig. 63). These data may suggest either the activity of a sn-1 specific DAG-lipase on AA-containing DAG, or lipase selection against VLC-PUFAs, as known for the mammalian pancreatic lipase.
Figure 63. Release of radioactivity from 1-stearoyl-2-
[14C] arachidonoyl-glycerol into sn-2-MAG .
112 3.3.11 Evidence for the activity of TAG lipase
We tested also the ability of lipolytic enzymes to hydrolyse TAG. To obtain radiolabelled TAG, we incubated P. incisa cells with [14C] oleate under nitrogen starvation conditions. When the resulting labelled TAG was extracted and applied to the assay, the production of FFA, DAG and
MAG was observed, indicating the presence of a TAG lipase in the microsomes. Most of the label was found in MAG and FFA followed by DAG. As shown in Fig. 64, MAG, but not FFA, was the most strongly labelled after 1 h of incubation, suggesting that formation of MAG is an important step in the utilization of TAG in the microalga.
Figure 64. Release of radioactivity from
[14C]trioleoylglycerol into FFA, DAG and MAG
showing the activity of TAG-lipase.
113 3.4 Role of AA-rich TAG
3.4.1 Nitrogen starvation and recovery
This study was carried out to investigate the utilization of AA-rich TAG in P. incisa during the onset of a favorable condition of growth.
3.4.1.1 Nitrogen starvation
Cultures of P. incisa were maintained in nitrogen (N)-free medium for 14 d at 25 °C to maximize the cellular content of AA-rich TAG. Fatty acid analysis of the N-starved cultures showed that net synthesis of fatty acids and particularly AA proceeded during starvation, reaching after 14 d, a content of 29.2% (of dry weight) and 14.4%, respectively (Fig. 65). TAG accounted for 86% of total lipids, comprising 25.1% (of dry weight) of the cell biomass. AA was the major fatty acid of TAG constituting 50% of its fatty acid content (Table 19). More than 80% of total cellular AA was accumulated in TAG, during this period.
30 TFA 24 oC TFA 12 oC 25 AA 24 oC AA 12 oC 20
15 Fatty acids (% dw) (% acids Fatty
10
5 0123456 Time (days)
Figure 65. Changes in total fatty acids (!, ") and and arachidonic acid (#, $) during recovery from N-starvation at 24 °C (solid symbols) and 12 °C (hollow symbols).
114 3.4.1.2 Recovery from nitrogen starvation
Recovery from N-starvation was induced by resuspension of the cells in complete nutrient medium. Recovery of growth in terms of increase in chlorophyll and biomass contents of the cultures was followed for six days. Samples for lipid analysis were taken after 2 d at room temperature (24 °C) and after 4 d at low temperature (12 °C). These time points were chosen on the basis of similar increases in dry weight. After a 1d lag, growth was resumed and significant chlorophyll and biomass synthesis commenced at room temperature (Fig. 66). At low temperature, however, chlorophyll accumulation and biomass production were much slower
(Fig. 66).
100 7
o Chl 24 C DW 24 oC o Chl 12 C DW 12 oC 80 6 ) -1 Dry wt. (mgL wt. Dry 60 5
40 4 -1 ) Chlorophyll (mg L (mg Chlorophyll
20 3
0 2 0123456
Time (days)
Figure 66. Changes in chlorophyll (!, ") and dry weight (#, $) during recovery from N- starvation at 24 °C (solid symbols) and 12 °C (hollow symbols).
115 Table 19. Fatty acids composition of major lipid classes of P. incisa following recovery (Rec.) from N-starvation (Starv.)
Temp % TFA Fatty acid composition (% of total) o -1 Lipid Conditions ( C) TFA (µg mL ) 16:0 16:1 16:2 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:3 20:4 ω7 ω6 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 TAG Starv. 86.0 779.2 8.0 0.2 0.4 0.3 3.5 16.1 3.6 13.1 1.0 0.9 1.0 50.0 Rec. 24 77.6 546.6 9.0 0.2 0.3 0.7 3.1 12.6 3.6 13.2 0.9 2.1 0.9 51.1 Rec. 12 78.2 532.4 8.6 0.2 tr 0.8 3.2 11.9 3.7 10.8 1.0 3.6 1.0 52.4
MGDG Starv. 1.8 16.0 3.6 0.3 23.6 14.6 0.3 3.2 0.4 28.9 0.5 20.4 - 2.7 Rec. 24 4.3 30.5 2.1 0.3 4.4 13.3 0.7 6.2 2.6 22.5 1.2 21.9 0.4 22.9 Rec. 12 2.7 18.3 2.1 0.5 1.7 23.2 0.6 2.2 2.2 6.7 1.1 42.0 - 16.1
DGDG Starv. 1.6 14.1 16.7 0.3 6.3 1.2 0.5 5.8 0.9 42.7 1.1 10.5 0.3 11.3 Rec. 24 3.8 26.6 7.1 0.4 1.8 0.9 2.3 8.8 3.9 37.0 1.1 13.9 0.4 20.9 Rec. 12 3.2 21.5 7.3 0.5 2.4 3.0 1.2 2.4 2.9 12.6 1.2 44.5 0.5 20.2
SQDG Starv. 3.2 28.8 62.3 0.2 - - 0.7 2.7 8.5 21.1 0.3 3.3 - 0.4 Rec. 24 4.9 34.8 54.2 0.3 - - 2.5 5.7 9.6 17.4 0.5 7.9 - 1.8 Rec. 12 5.5 37.5 56.8 0.2 - - 1.6 1.2 16.8 4.7 0.4 16.2 - 1.4
DGTS Starv. 2.1 19.1 35.6 1.5 tr 0.2 5.5 3.7 3.8 22.1 6.2 0.6 0.6 13.1 Rec. 24 2.2 15.7 36.2 1.2 - - 6.9 1.9 3.3 22.7 2.8 1.3 0.3 17.4 Rec. 12 3.4 23.4 43.8 1.1 - 0.2 7.5 1.0 5.7 13.1 4.3 2.3 0.7 16.4
PE Starv. 0.5 4.8 12.0 0.4 - - 7.3 1.5 32.9 5.5 2.6 0.2 4.6 27.2 Rec. 24 0.7 5.1 14.9 - - - 9.8 1.1 38.7 5.1 0.8 0.2 3.1 25.6 Rec. 12 0.8 5.6 18.1 0.3 - - 10.0 1.1 34.3 3.5 0.9 0.4 4.3 24.4
PC Starv. 0.9 8.5 23.6 1.3 tr 0.2 6.5 5.0 15.7 16.8 4.9 0.3 1.5 18.6 Rec. 24 1.9 13.1 18.7 0.8 0.2 - 6.0 3.4 11.6 18.9 2.8 1.0 1.1 31.8 Rec. 12 1.9 13.3 33.0 0.7 - - 6.8 2.3 14.8 9.9 3.4 2.3 1.5 21.9
DAG Starv. 1.1 9.9 15.6 1.1 - 0.5 8.4 30.9 6.3 12.0 1.1 0.3 0.5 20.3 Rec. 24 1.0 6.9 21.4 - - - 9.3 25.7 5.3 12.2 0.8 0.8 - 24.5 Rec. 12 0.8 5.6 17.5 0.5 - 0.6 8.8 22.1 5.4 11.9 1.1 1.3 0.6 27.0
FFA Starv. 0.8 6.8 22.2 2.5 - 1.0 11.1 25.0 6.2 12.3 3.6 0.5 3.1 3.8 Rec. 24 0.5 3.4 19.9 - - - 11.1 15.6 3.5 10.8 1.3 0.5 2.0 31.1 Rec. 12 0.4 2.5 27.3 1.4 - - 14.8 15.3 3.6 6.3 5.9 1.2 6.2 12.9
Cultures were resuspended in full medium and grown at 24 oC (2 d) or 12 oC (4d). tr-traces.
116 3.4.1.3. Alterations in lipid and fatty acid content and composition
After 4 days of recovery the total fatty acid content decreased by 33.5 and 50.4% at 24 °C and 12
°C, respectively (Fig. 65). The decrease in AA content was lower, amounting to only 28.3 and
45.7%, respectively. At both temperatures, recovery was accompanied by a decrease in the proportion of TAG (from 86 to 78% of total lipids) and by an increase in the relative proportions of the chloroplastic lipids, MGDG (from 1.8 to 4.3 and 2.7%, respectively), DGDG (from 1.6 to
3.8 and 3.2%, respectively) and SQDG (from 3.2 to 4.9 and 5.5%, respectively) (Table 19).
The volumetric content of TAG decreased from 779 to 547 and 533 µg mL-1. At 24 °C the net buildup of DGDG and especially MGDG was more intense than at 12 °C (Table 19).
MGDG increased from 16.0 to 30.5 and 18.3 µg mL-1, DGDG from 14.1 to 26.6 and 21.5 µg mL-1 and SQDG from 28.8 to 34.8 and 37.5 µg mL-1 at 24 °C and 12 °C, respectively (Fig. 67).
During the recovery, there was also a decrease in the content of DAG, from 9.9 to 6.9 and 5.6 µg mL-1, and of FFA, from 6.8 to 3.4 and 2.5 µg mL-1, respectively.
During recovery at 12 °C, but much less so at 24 °C, the proportion of 18:3ω3 in the three major chloroplastic lipids and of 16:3ω3 (in MGDG and DGDG), increased at the expense of their respective ω6 precursors, 18:2 and 16:2 (Table 19). The proportion of AA in MGDG increased sharply from 2.7 to 22.9 and 16.1% and in DGDG from 11.3 to 20.9 and 20.2%, at 24
°C and 12 °C, respectively. The content (% dw) of these lipids also increased, especially at 24
°C. Consequently, in MGDG, AA increased from 0.4 to 7.6 and 1.7 µg mL-1 and in DGDG from
1.3 to 7.9 and 2.7 µg mL-1, at 24 °C and 12 °C, respectively. The proportion of AA increased also in the extraplastidial polar lipids, PC and DGTS. The proportion of AA in TAG didn’t change but the content of AA in TAG decreased from 285 to 139 and 203 µg mL-1, respectively.
These findings suggest a transfer of AA from TAG to polar lipids at 24 °C, but much less so at
12 °C (Table 19, Fig. 67). There was also a sharp increase in the proportion of AA in FFA (from
117 3.8 to 31.1 and 12.9%, respectively), indicating an enhanced activity of TAG lipase. At 12 °C, but not at 24 °C, the proportion of 18:2 in the extrachloroplastic lipids DGTS, and PC decreased from 22.1 and 16.8% to 13.1 and 9.9%, respectively. Lower decreases were noted in the proportion of 18:1. Correspondingly, the proportion of 16:0 increased. These findings indicate
DGTS and PC as likely sources of acyl moieties that can be remodeled into chloroplastic lipids.
8 300 8 300 o 24 oC 12 C 7 7 240 240 6 6 TAG ( TAG
g/mL) 5 5 µ µ µ µ 180 180 µ µ TAG µ µ
4 4 g/mL) MGDG 120 120 3 3 DGDG
Polar lipids ( SQDG 2 2 60 60 1 1
0 0 0 0 024024
Time (days) Time (days)
Figure 67. Changes in the volumetric content of arachidonic acid in TAG ("), MGDG (#),
DGDG (!) and SQDG (!) during recovery from N-starvation at 24 °C (left panel) and 12 °C
(right panel).
118 3.4.1.4 Molecular species analysis
In order to further elucidate the transfer of AA from TAG to chloroplastic lipids, we have analyzed the molecular species composition of MGDG, which was found to be the major lipid sink of AA during recovery. Under nitrogen starvation, the molecular species of MGDG were mostly of the 18/16 type (Table 20), constituting about 84% of total MGDG. During recovery at
24 °C, these molecular species were diluted by the production of 18/18, 20/18 and 20/20 molecular species, whose content increased from 1 to 4.7, 1.4 to 11.8 and 0.2 to 2.8 µg mL-1, respectively. Within the 18/16 group there was a decrease in the content of those containing ω6 fatty acids (16:2 and 18:2), in favor of the fully desaturated species, 18:3ω3/16:3ω3. The share of the latter increased from 28.8 to 68.5% of total 18/16 molecular species, indicating an intensive
ω3 desaturation. At 12 °C too, the content of the 18/16 species decreased (from 13.4 to 10.0 µg mL-1), whereas that of the 18/18, and especially the 20/18 and the 20/20 increased, but much less than at 24 oC (from a total of 2.6 to 8.2 µg mL-1). Molecular species containing ω3 fatty acids
(16:3 and 18:3) dominated at the expense of their less unsaturated ω6 precursors. In each of the first 3 groups, the share of 18:3ω3-containing molecular species constituted 96.7, 83.3 and
70.6% of their respective groups.
119 Table 20. Changes in the molecular species distribution (% of TFA) and content (µg mL-1) in
MGDG of P incisa, following recovery (Rec.) (2 d at 24 °C, or 4 d at 12 °C) from N-starvation
Molecular Molecular species distribution and content species N-starvation Rec. 24 oC Rec. 12 oC 14 d 2 d 4 d % of µg mL-1 % of µg mL-1 % of µg mL-1 TFA TFA TFA 18:2/16:2 31.3 5.0 2.6 0.8 tr tr 18:3/16:2 16.0 2.6 4.9 1.5 1.2 0.2
18:2/16:3ω3 12.5 2.0 4.3 1.3 0.6 tr 18:3ω3/16:3ω3 24.2 3.9 25.4 7.7 53.1 9.7 Total 18/16 83.9 13.4 37.1 11.3 54.9 10.0 18:1/18:2 tr tr 3.8 1.2 tr tr 18:2/18:2 2.0 0.3 6.4 1.9 tr tr
18:1/18:3ω3 0.8 tr tr tr tr tr 18:2/18:3ω3 2.1 0.3 2.8 0.8 1.6 0.3 18:3ω3/18:3ω3 1.3 0.2 2.4 0.7 8.0 1.5
Total 18/18 6.1 1.0 15.3 4.7 9.6 1.7 20:4/18:1 2.4 0.4 10.1 3.1 1.8 0.3 20:4/18:2 4.4 0.7 21.2 6.5 6.7 1.2 20:4/18:3ω3 2.1 0.3 7.2 2.2 20.2 3.7 Total 20/18 8.9 1.4 38.5 11.8 28.6 5.2
20:4/20:4 1.0 0.2 9.0 2.8 6.9 1.3
Positional analysis was not performed. TFA-total fatty acids; tr-traces; Unless otherwise indicated, PUFA are of the ω6 family.
120 3.4.1.5 Radiolabelling
A radiolabelling study was carried out to monitor the transfer of AA from TAG to the chloroplastic lipids following recovery. Cells of P. incisa were labeled with [1-14C]18:1 for 24 h under N-starvation conditions. During this period most of the label was converted to [1-14C]20:4 and deposited in TAG (data not shown). The redistribution of label was followed during the recovery period at room and low temperatures. The label of TAG continuously decreased and was partially transferred to the chloroplastic lipids, MGDG, DGDG and SQDG during recovery at 24 °C, but much less so at 12 °C (Fig. 68). At 12 °C, the decrease of label of TAG was observed only in the first 4 h and the transfer to chloroplastic lipids was much lower. Decreases in label were noted also in DGTS (at both temperatures) and in PC (only at 12 °C) (Fig. 69). An early transitory rise was noted in the label of AA in PC and even more in DGTS. At 12 °C, it was less pronounced, mostly in DGTS. The relatively high label of DGTS and its subsequent decrease may indicate this lipid together with PC as the primary intermediates for both TAG and
MGDG.
121 8 90 8 o 90 MGDG 24 oC TAG 12 C DGDG 7 85 7 TAG 85 SQDG 6 6 80 80 5 5 75 75 MGDG 4 4 DGDG 70 70 3 3 SQDG 65 65 2 2
60 1 60
Radioactivity (% of total decompositions) total of (% Radioactivity 1 Radioactivity (% of total decompositions) 0 55 0 55 0 8 16 24 32 40 48 0 8 16 24 32 40 48 Time (hours after nitrogen replenishment) Time (hours after nitrogen replenishment)
Figure 68. Redistribution of radioactivity in chloroplastic lipids (left scale) and TAG (right scale) of P. incisa during nitrogen replenishment to the nitrogen starved cells at 24 °C (left panel) or 12
°C (right panel).
Labeling with [1-14C]18:1 (24 hours) was carried out on P. incisa cells maintained on N-free medium for 14 d. Changes in label distribution was followed after nitrogen replenishment;
TAG ("), MGDG (#), DGDG (!) and SQDG (!), respectively.
122
8 8 24 oC o DGTS 12 C DGTS 7 PC 7 PC PE PE 6 6
5 5
4 4
3 3
2 2
1 1 Radioactivity (% of total decompositions) of total (% Radioactivity 0 0 0 8 16 24 32 40 48 0 8 16 24 32 40 48 Hours after nitrogen replenishment Hours after nitrogen replenishment
Figure 69. Redistribution of radioactivity in extraplastidial lipids (left scale) and TAG (right scale) of P. incisa during nitrogen replenishment to the nitrogen starved cells at 24 °C (left panel) or 12 °C (right panel).
Labeling with [1-14C]18:1 (24 hours) was carried out on P. incisa cells maintained on N-free medium for 14 d. Changes in label distribution was followed after nitrogen replenishment;
DGTS (▼), PC ($), PE (").
3.4.2 SHAM treatment and recovery
The aim of this part of the research was to investigate whether cells of P. incisa with lower AA content would recover differently from cells with higher AA content, upon transferring of a stationary culture onto fresh nutrient medium. To obtain cells with lower AA content, the 6 d batch culture was split into two parts: the first part was kept under the same conditions, the second part was exposed to 300 µM salicylhydroxamic acid (SHAM) for another 5 d. The latter culture showed higher accumulation of 18:1ω9 and 18:2 and decrease in 18:3ω6 and AA, indicating an inhibition of the ∆12 and ∆6 desaturases, as shown previously (Bigogno et al.
123 2002b). After another 5 d, the proportion of 18:1ω9 was twice that of control (32.9 and 15.5% of total fatty acids, respectively) while that of AA was about half of the control (21.7 and 37.8%, respectively) (Table 21).
Table 21. Distribution of fatty acids of P. incisa in the presence of SHAM and during the recovery from SHAM
Growth TFA AA Fatty acid composition (% of total fatty acids) Condition % % 16:0 16:1 16:2 16:3 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:0 20:1 20:2 20:3 20:4 20:5 DW DW ω7 ω6 ω6 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 ω3 6d Stationary 10.6 2.7 13.9 0.6 1.3 0.4 0.2 3.1 22.0 3.0 18.2 2.3 4.6 0.5 0.6 0.3 1.1 26.0 1.1 Further 5d treatment Control 13.7 5.2 12.2 0.4 1.0 0.2 1.1 2.6 15.5 3.6 17.3 1.1 3.1 0.3 0.5 0.5 0.9 37.8 1.4 SHAM 17.5 3.8 10.9 0.3 0.7 0.2 0.9 2.4 32.9 2.5 20.2 0.7 2.9 0.3 0.6 0.8 0.5 21.7 0.8 Recovery 2d Con 12.5 5.2 12.1 0.6 0.7 0.2 1.5 2.3 10.9 4.1 15.5 1.1 4.8 0.3 0.4 0.5 0.7 41.9 1.7 2d SHAM 16.4 4.9 10.9 0.3 0.5 0.3 1.3 2.2 22.8 3.2 18.5 1.6 4.5 0.3 0.6 0.7 0.7 30.1 1.1
6d Con 8.8 3.4 14.1 1.1 1.5 0.1 2.3 1.3 6.6 4.8 15.7 1.2 8.1 0.2 0.3 0.4 0.7 38.8 2.2 6d SHAM 11.1 4.3 12.7 0.7 1.1 0.2 1.7 1.4 9.8 5.0 16.2 1.3 6.5 0.2 0.3 0.5 0.7 38.9 1.9
Prior to the recovery, the contents of AA in control and treatment were 5.2 and 3.8% of
DW, respectively. However, the total fatty acid content was higher in the SHAM-treated culture,
13.7 and 17.5 % of DW, in control and treatment, respectively. The biomass volumetric content increased from 2.5 to 3.1 and 2.7 mg/mL, in control and treatment, respectively.
Recovery from the inhibitor was obtained by washing the cells three times and transferring into the complete inhibitor-free medium (after a 3-fold dilution). The control culture was subjected to a similar treatment. After 6 d recovery, the proportion of TFA (% of DW) decreased in both control and SHAM-pretreated cultures similarly by 35.8 and 36.6%, respectively (Table 21, Fig. 70) while chlorophyll and dry weight increased (Fig. 72). The increase in dry wt and chlorophyll content of SHAM-pretreated culture was slower than in control. During recovery, AA-rich TAG could export AA to chloroplastic lipids (Bigogno et al.
2002b; section 3.4.2). In keeping, the control culture, which has higher amount of AA in its cells,
124 mostly in TAG, showed a shorter lag and a sharper increase of chorophyll (from 9.5 to 48.8
µg/mL in 6 d) while the low AA-containing, SHAM-pretreated cells showed a longer lag and
lower increase in chlorophyll during recovery (from 9.0 to 31 µg/mL in 6 d) (Fig. 72).
18 8
16 7
14 DW) of (% AA 6 12
10 5
8 Total fatty acids (% of DW) 4 6
0246 Time after recovery (days)
Figure 70. Changes in the biomass content of TFA (○, ●) and AA (∆, ▲) during growth recovery in control (hollow symbols) and SHAM (solid symbols) treated cultures, respectively.
After 6d under similar decreases in TFA (% of DW), major monounsaturated 18:1ω9
decreased by 72.7 and 81.1% in the control and SHAM-pretreated culture respectively, while the
content of AA in the biomass decreased in the control by only 36.1% (Table 21). Surprisingly, in
the SHAM-pretreated cells, AA increased continuously for the first 4 d crossing the level of AA
in control after 4 d (Table 21, Fig. 70). This finding suggests the reactivation of the ∆12 and ∆15
desaturases, resulting in an increase in the proportion of AA after 6 d from 21.7 to 38.9% in
SHAM-treated cells, but not in the control (Table 21).
During 2 days of recovery, the volumetric content of TFA did not change while that of
AA increased (Fig. 71). After 2 d, both TFA and AA synthesis increased, but faster in the
SHAM-pretreated culture. After 6 d, the volumetric content of AA in SHAM-pretreated culture
125 increased sharply since the onset of recovery by 185%, in comparison to only 67% in the control
(by 10.1 and 2.4 µg/mL, respectively). TFA increased almost similarly, by 59 and 63% in
SHAM-pretreated culture and control, respectively, indicating a particularly active synthesis of
AA during recovery from SHAM (Fig. 71). The fast recovery of AA synthesis was not likely the
consequence of impaired or slower synthesis of main chloroplastic polyunsaturated fatty acids.
Indeed, the SHAM-pretreated culture featured levels of 18:3ω3 similar to that of control during
recovery (Fig. 73).
250 140
120
200 AA ( 100 µ µ µ µ g/mL) g/mL) µ µ µ µ 150 80 TFA ( 60 100 40
0123456 Time after recovery (days)
Figure 71. Changes in the culture contents of TFA (○, ●) and AA (∆, ▲) during growth recovery of control (hollow symbols) and SHAM-pretreated (solid symbols) cultures, respectively.
126 50 5 Dry wt (mg/mL culture) wt (mg/mL Dry 40 4 g/mL) µ µ µ µ 30 3 20 2 Chlorophyll ( 10
1 0 0246 Time after recovery (days)
Figure 72. Changes in the contents of chlorophyll (○, ●) and dry wt (∆, ▲) in P. incisa cultures
during growth recovery in control (hollow symbols) and SHAM-pretreated (solid symbols)
cultures, respectively.
20
16
12 g/mL ) g/mL µ µ µ µ ( -linolenic acid α α α α 8
4 0123456 Time after recovery (days)
Figure 73. Changes in the volumetric content of 18:3ω3 (○, ●) during growth recovery in control
(hollow symbols) and SHAM-pretreated (solid symbols) cultures, respectively.
127 3.5 Mutant studies
Logarithmically growing cells of P. incisa were treated with 100 µg mL-1 1-methyl-3-nitro- nitrosoguanidine (NNG) solution for 1 h. NNG stock solution was prepared in DMSO in order to attain a better penetration of mutagen through the adamant cell wall of the microalga and a stronger mutagenic effect. The survived cells were grown on solidified BG11 medium. Overall
300 colonies were tested during this study for their fatty acid composition and content.
Additional screening of these cultures was carried out by plating 5 µl of individual mutant cell suspension as spots onto three agar plates prepared with BG11, nitrogen-free BG11 and phosphate-free BG11, respectively. We expected that colonies in which biosynthesis of TAG were impaired would show a poor growth on nitrogen-free medium, in contrast to overproducing mutants that would show a better growth. We also incubated Petri dishes, containing the same replicates at 4 °C to check their growth at low temperature, which may be affected by the PUFA content of the cells. During screening, we also compared TAG in the cells visually under a fluorescence microscope following staining with fluorescent dye Nile Red. The selected colonies were further subcultured on agar plates to obtain sufficient biomass, before growing in small bottles or in 10 mL tubes. The biomass obtained from the stationary culture or from the growth in nitrogen-free medium was collected and fatty acid analysis was carried out.
The screening by fatty acid analysis of the selected 300 colonies did not produce any AA- deficient mutant. Several mutants showed, however, variations in the fatty acid distribution and lipid content. Either lower or higher proportion of AA and total lipid in comparison to the wild type were observed among the mutant strains. Those showing the largest deviations from the wild type were selected for further analysis. Alternatively, strains that demonstrated the lower or higher AA and total lipids contents than average value (drawn among them) were also selected.
The selected inocula were grown in flasks with dilution to obtain few generations of cells, which were transferred to nitrogen-starved condition for maximum lipid accumulation. The
128 fatty acid distribution and lipid content were determined as mentioned above. The mutants producing lower level of AA were grown also at lower temperature to compare their growth rate with wild type. Among mutants (Table 22) B33, 50, 105, 114 and NB5 contained high proportions of AA in TFA, however, still lower than in WT. B83, 107, 109, contained lower proportions of AA as compare to other mutants and to WT.
Table 22. Fatty acid composition and content of selected mutant strains of P. incisa in comparison to WT
Fatty acid composition (% of total fatty acids) Strain AA TFA 16:0 16:1 16:1 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:3 20:4 20:5 % DW % DW ω11 ω7 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω3
WT 6.9 0.5 0.3 0.2 1.8 10.3 5.8 9.3 0.6 0.8 1.4 61.7 0.0 24.5 39.6 B33 11.8 0.5 0.4 0.4 1.6 9.0 5.3 9.5 0.7 1.1 1.3 57.6 0.7 14.2 24.7 B50 10.6 0.6 0.4 0.2 2.4 8.0 5.4 11.8 0.9 0.8 1.2 57.0 0.3 20.9 36.7 B83 9.5 0.6 0.3 0.3 1.2 13.1 7.0 11.7 0.8 0.9 1.0 53.1 0.3 15.5 29.2 B105 8.5 0.5 0.3 0.3 2.2 11.3 4.7 9.7 0.6 0.7 1.0 59.4 0.4 26.7 45.0 B107 9.8 0.5 0.4 0.3 2.0 22.1* - 14.9 0.4 0.6 0.6 47.7 0.3 13.0 27.2 B109 11.0 0.8 0.3 0.8 2.2 11.5 5.5 12.1 - 1.0 1.4 53.4 - 7.7 14.3 B114 10.0 0.5 0.3 0.4 1.9 8.9 6.9 9.7 0.6 1.0 1.5 57.8 0.4 17.5 30.3 NB5 8.8 0.4 0.2 0.3 1.8 10.9 5.7 9.6 0.5 0.8 1.5 59.3 0.2 23.7 39.9 * 18:1 isomers did not resolve
A higher biomass content of TFA was observed in B105 in comparison to WT (45.0 and
39.6 % DW, respectively). There was not a significant change in the fatty acid composition in this mutant, indicating an enhanced TAG biosynthesis. The fatty acid analysis of B107 and B109 suggested an inhibition in the synthesis of AA and accumulation of TFA as shown by their lower percentages of biomass in comparison to WT (Table 22). Substantial increases in 18:1 and 18:2 were observed in those mutants, indicating a mutation in ∆6 desaturase. The high AA-containing
B105 and low AA-containing B107 and B109 were chosen for their lipid class analysis along with the WT.
Lower levels of neutral lipids were observed in mutants B105, B107 and B109 than in
WT (Table 23). In lipid proportions, the lowest level of NL and highest level of PL were
129 observed in B107. The proportions of AA of fatty acids were also lowest in NL and PL in this mutant among the three mutants studied and WT, which may indicate some problem in the transfer of acyl moieties from PL to NL.
Table 23. Fatty acid composition of neutral and polar lipid fractions of selected mutants in comparison to WT
Lipid Fatty acid composition (% of total fatty acids) Total 16:0 16:1 16:1 16:3 18:0 18:1 18:1 18:2 18:3 18:3 20:2 20:3 20:4 Lipid NL/PL ω11 ω7 ω3 ω9 ω7 ω6 ω6 ω3 ω6 ω6 ω6 % NL-WT 9.8 0.7 - - 1.7 10.9 5.1 9.3 0.5 0.2 tr 0.7 60.1 94.7 17.7 NL-B105 8.5 0.6 - - 2.1 10.2 4.7 8.6 0.6 0.2 tr 0.7 63.4 91.5 10.7 NL-B107 8.9 0.2 - tr 1.7 21.9 0.0 14.2 0.5 0.5 tr 0.4 50.9 85.4 5.8 NL-B109 9.1 0.3 - 0.9 2.2 11.4 5.6 10.0 0.5 0.6 - 0.5 58.2 92.9 13.1
PL-WT 26.2 1.2 4.3 3.6 1.1 2.2 7.4 24.0 2.7 4.0 0.4 1.0 18.1 5.3 PL-B105 26.1 1.2 5.2 3.2 1.4 2.7 6.3 27.0 2.9 3.8 0.7 0.9 16.2 8.5 PL-B107 25.0 1.8 4.0 2.6 0.8 6.6 7.0 30.4 0.9 3.5 0.5 0.4 14.3 14.6 PL-B109 27.3 1.1 2.9 4.2 1.6 3.8 12.1 18.7 2.2 4.4 1.0 0.4 17.5 7.1
Indeed, when polar lipid classes of B107 were analyzed (Table 24), an increase in the content of phosphatidylcholine (PC), phosphatidylethanolamine (PE) and DGTS, were observed.
These lipids are involved in the desaturation of C18 and C20 fatty acids to produce AA. Among three, PC was almost doubled in comparison to that in WT. 18:3ω6 was almost absent in PC, PE and DGTS, while18:1ω9 was clearly increased in all three, indicating the inhibition of ∆12 and
∆6 desaturases. When ∆12 and ∆6 desaturases were inhibited in P. incisa by the application of the inhibitor SHAM (Bigogno et al. 2002b), AA synthesis was inhibited and 18:1 increased, but the TFA content was not inhibited. In B107, the inhibition of desaturation of C18 fatty acid together with the decrease in TFA content, may indicate the inhibition of enzymes, which release acyl moieties from these lipids to the acyl/FFA pool or produce DAG (phospholipases or lyso- phospholipid acyltransferase) to enable AA and TAG production.
130 Table 24. Major fatty acid composition of selected polar lipid classes of WT and C18 desaturase mutant 107
Fatty acids Strain Lipid Fatty acid (% of total) (µg mg-1) 16:0 18:0 18:1 18:2 18:3 20:3 20:4 AA 18:2 TFA
ω9+ω7 ω6 ω6 ω6 ω6 WT DGTS 51.7 3.7 10.9 18.2 2.7 0.5 7.8 8.8 20.6 113.4 107 49.4 3.2 18.2 23.8 0.6 - 3.9 4.6 28.2 118.3 WT PC 59.6 1.7 9.4 21.8 5.3 - 2.2 0.9 8.8 40.4 107 50.4 0.6 23.8 21.0 - 1.4 1.4 1.0 15.2 72.1 WT PE 23.1 4.6 26.3 5.9 1.2 4.8 28.1 9.9 2.1 35.1 107 18.9 4.0 31.3 10.9 - 2.3 27.0 11.5 4.6 42.5
131 4. Discussion
4.1 Lipids involved in the biosynthesis of AA-rich TAG
In the stationary phase P. incisa accumulates AA-rich TAG as a result of nutrient limitation. We studied the changes in lipid distribution, their molecular species profiles and fatty acid composition following the growth of the batch culture. The decrease in the content of major chloroplastic lipids, MGDG, DGDG, SQDG and PG is in line with the decrease in chlorophyll content during the stationary phase. The major extraplastidial phospholipids, PC, PE, PI and PA, except for the betaine lipid, DGTS, also decreased. These data allow suggesting a role for the phosphate-devoid lipid DGTS in the maintenance of the membrane structure of starved cells.
The presence of higher proportions of 18:2ω6 and its down stream ω6 products 18:3ω6,
20:3ω6 and AA, in the extraplastidic lipids PC, PE and DGTS indicated to the role of this lipids in the desaturation of C18 and C20 fatty acids in the synthetic pathway leading to AA (Bigogno et al. 2002a). Among the polar lipids, the proportions of 20:3ω6 and AA were highest in PE, and the proportions of 18:2ω6 and 18:3ω6 were the lowest, indicating this lipid as a preferred substrate for the ∆5 desaturase (Table 1). The lower proportion of 20:3ω6 and higher proportion of 18:2ω6 and 18:3ω6 in PC and DGTS suggests these lipids as the substrates for the ∆12 and
∆6 desaturations. At the stationary phase, when TAG synthesis is enhanced, the sharp increase in
18:2 of PE was accompanied by a decrease in 20:3 and AA, which is likely due to a higher turnover of AA into TAG. In fact, at the stationary phase, there was a decrease in the proportion of the ω6 intermediates, 18:3ω6 and 20:3ω6, in PC, PE and DGTS, but not in 18:2. This may indicate the ∆6 desaturation of 18:2 to 18:3ω6 as the rate-limiting step in the biosynthesis of AA.
In several algae, PC, PE and DGTS are the substrates for the desaturases in the biosynthesis of PUFAs. In Chlamydomonas reinhardtii, which is devoid of PC, desaturation of
C18:1-4 takes place at the sn-2 position of DGTS and PE (Giroud and Eichenberger 1988). In
Ochromonas danica, ∆12 desaturation takes place on DGTS while ∆15 and ∆6 desaturases act
132 preferentially on PE (Vogel and Eichenberger 1992). Different substrate specificity for the desaturation of fatty acids of different chain length was reported in Nannochloropsis, where desaturations at ∆12 and ∆6 positions of C18 take place in PC while ∆5 and ∆17 desaturations of
C20 occur in PE (Schneider and Roessler 1994). In Monodus subterraneus, PC was suggested as the substrate of choice for C18 desaturations, whereas PE and DGTS as that of C20 (Khozin-
Goldberg et al. 2002b). Differently, in the red microalga P. cruentum, desaturations of C18 and
C20 fatty acids to produce AA, occur in PC, whereas further desaturation of AA at the ∆17 position to produce EPA, takes place in the chloroplastic lipid, MGDG (Khozin et al. 1997).
Previous radiolabeling studies, carried out to determine the lipid intermediates involved in the biosynthesis of AA in P. incisa, have shown that the polar lipids, PE, PC and DGTS are involved in the fatty acid desaturations and the biosynthesis of AA-rich TAG (Bigogno et al.
2002a). However, detailed analysis of the molecular species composition and positional analyses of these lipids were not conducted.
The molecular species and positional analyses of PE, PC, DGTS (Tables 2-5) carried out in the present work, have provided a better insight into the role of these lipids in the biosynthesis of TAG in P. incisa at the molecular lipid level. Specific pattern of fatty acid positioning and the presence of some specific molecular species, which play a key role in TAG synthesis, were presumed. Comparison of molecular species structures of these lipids and TAG has led to the determination of the role of some particular molecular species or acyl moieties at certain positions of the lipid in the biosynthesis of TAG.
The presence of high proportion of 18:2- and 18:3ω6-containing molecular species in PC indicated PC as a substrate for the ∆12 and ∆6 desaturases (Table 3). The presence of various
C18 PUFAs in the sn-2 position of PC revealed that those desaturations occur specifically at the sn-2 position. A substantial increase in AA/18:2 at the expense of AA/18:3(ω6 and ω3) molecular species, in the stationary phase, may indicate a faster turnover of 18:3 from this
133 species for the sequential elongation and ∆5 desaturation in PE. AA was located at both positions of PC. The AA in the sn-1 position is likely released from PE, where most AA is produced, into the acyl-CoA pool. The acylation of the sn-1 position could occur at the level of G3PAT or by exchange with another acyl moiety residing at the sn-1 position of PC. Another group of PC molecular species contain 16:0 or 18:1ω7 at the sn-1. Possibly, the latter type of molecular species might serve as a structural component of the lipid, or contribute sn-2 fatty acid for the acylation of DAG molecule by the activity of PDAT to produce TAG. Diarachidonoyl-PC was one the major molecular species in both the exponential and stationary phases.
The absence of intermediate fatty acids of C18 and C20 desaturation (C18:1-3, 20:3) in the most abundant molecular species of PE containing 18:1ω7 in the sn-1 position revealed that these are less likely to be involved in these desaturations of fatty acids, instead they accumulate
AA (Table 4). Thus, just a small proportion (about 30%) of the molecular species of PE, i.e., those containing AA at the sn-1 position and C20 at the sn-2 position are likely to be involved in the desaturation. Diarachidonoyl-PE was the dominant molecular species in this group, its share slightly increased in the stationary phase.
The presence of the AA intermediates C18:1-3 and C20:3-4 primarily in the sn-2 position of DGTS (Table 2) indicated the role of this position of DGTS in the ∆12, ∆6 desaturations and possibly also in ∆5 desaturation. Indeed, the major group of molecular species contained 16:0 at the sn-1 position and an ω6 PUFA at the sn-2 position (Table 5). The sharp increase in the proportion of 16:0/18:2 in the stationary phase at the expense of 16:0/18:3 (ω3 and ω6 indicates the fast turnover of 18:3 from this species. Similarly, in Chlamydomonas reinhardtii it was shown that the ∆12, ∆15, and ∆6 desaturations of C18 fatty acids occur while residing in the sn-2 position of DGTS or PE (Giroud and Eichenberger 1989).
Indeed, in P. incisa, radiolabeling studies showed that C18 desaturations occur mainly within PC and DGTS, and the product, 18:3ω6, is elongated to 20:3ω6 and re-esterified mainly
134 to PE, where it is further ∆5 desaturated to produce AA (Bigogno et al. 2002a). However, in
Monodus subterraneus, labeling with linoleic acid showed that PC was the first polar lipid to be labeled, followed by PE and DGTS (Khozin-Goldberg et al. 2002b). The presence of similar molecular species in PC and PE of P. incisa allow speculating an interconversion between these phospholipids. Indeed, PE and PC may be interrelated by methylation of PE (Hubscher et al.
1959) or by ethanolamine exchange with a pre-existing PC head group (Miura and Kanfer 1976,
Shin and Moore 1990). In the radiolabeling study of P. incisa, more labeled AA was shown to appear in PC than in PE, suggesting a rapid methylation of AA-PE to AA-PC (Bigogno et al.
2002a).
Other than the rare triarachidonylglycerol, most molecular species of TAG contain at least two arachidonyl moieties (Fig. 12, Table 6). Since one of the major molecular species of both PC and PE contain two arachidonyl moieties, the corresponding TAG molecular species, may be obtained by acylation of the sn-3 position of the corresponding DAG, derived by the activity of DAGAT on these phospholipids. The relationship between the fatty acid composition of PC and TAG was demonstrated by inhibitor experiments. Following treatment with SAN
9785, TAG decreased significantly whereas AA synthesis was not affected (Bigogno et al.
2002c). The level of AA in PC and TAG increased sharply (from 21 to 42.5% and from 40 to
58%, respectively), indicating PC as the primary source of AA for TAG. However, the level of
AA in PE was not affected, suggesting that PC, rather than PE, is the source of AA for TAG.
When another inhibitor, salicylhydroxamic acid (SHAM), was applied, the proportion of 18:1 in
PC, DGTS and TAG increased sharply while the proportion of 18:2 and further down stream products up to AA were sharply decreased, indicating PC and DGTS as precursors for the ∆12 desaturation. TAG synthesis was not affected by SHAM and the substantial increase of 18:1 in
PC (14 to 57%) and TAG (28 to 66%) further supported a connection between these lipids
135 (Bigogno et al. 2002a,c). Here again, although the proportion of 18:1 increased in PE, 18:2 did not decrease, revealing it as a less likely substrate for ∆12 desaturation.
Thus, we assume that AA synthesized in PE is exported to PC, DGTS and TAG and partly to the chloroplastic lipids. The AA/AA molecular species of PC and PE may have a major role in donating AA for TAG by either providing AA moieties to the free fatty acid or acyl-CoA pool or directly as DAG. The TAG assembly can be completed using DAGAT for the esterification of AA/AA- and AA/18:1-3- DAG, derived from phospholipids by the action of
DG-cholinephosphotransferase (CPT), DG-ethanolaminephosphotransferase (EPT) or by phospholipases C and D. The absence of 18:1ω7 in the sn-1 position of TAG suggests that the molecular species of PE and PC, which has 18:1ω7 at the sn-1 position, contribute acyl moieties from the sn-2 position, mainly AA, to the acyl-CoA pool, rather than being converted into DAG and then to TAG. AA thus derived from the sn-2 position of the major molecular species of PE
(18:1ω7/AA) can be incorporated into the sn-1 and sn-3 positions of TAG via the acyltransferases of the Kennedy pathway (G3PAT and DAGAT, respectively) (Stymne and
Stobart 1987). The role of alternative DAG-utilizing enzymes for the synthesis of TAG should also be considered.
4.2 Role of extrachloroplastic lipids in TAG synthesis
Following 3d of N-starvation in the presence of the de novo lipid synthesis inhibitor sethoxidim
(200 µM), the volumetric content of TFA in the culture increased insignificantly. The inhibition of the de novo synthesis affected differently the production of C18 fatty acids and AA. Despite lower volumetric accumulation of AA in the presence of inhibitor, AA was the only fatty acid, whose percentage significantly increased (Table 7, 8), indicating a contribution of extrachloroplastic lipids to provide AA by desaturations and elongation of preexisting fatty acids.
136 Under P-starvation, P. incisa survived and produced TAG by utilizing its phospholipids and probably other endogenous phosphate resources. However, under P-starvation, P. incisa produced less biomass and TFA than in control cultures, and is not able to overproduce storage lipids as shown under N-starvation. Under N-starvation, the lack in phosphate affected significantly the proportion of AA, indicating importance of phospholipids as substrates for desaturases in the pathway of AA biosynthesis.
The proportion of all phospholipids decreased during P-starvation, whereas the proportion of the extrachloroplastic DGTS and the chloroplastic SQDG, of polar lipids increased sharply (Table 10). In higher plants, a novel biochemical mechanism aiding in phosphate conservation has been recently revealed. The decrease in the phospholipid content is accompanied with an increase in the proportions of the non-phosphorous membrane lipids,
DGDG and SQDG (Hartel et al. 1998, Essigmann et al. 1998). Several genes, necessary for the syntheses of extraplastidial DGDG are activated by P-deprivation (Awai et al. 2001, Kelly and
Dormann 2002). In the photosynthetic DGTS-containing bacteria Rhodobacter sphaeroides and
Sinorhizobium meliloti, P-deprivation resulted in the accumulation of different P-devoid lipids
(SQDG, DGTS and an ornithine lipid) (Benning et al. 1993, Benning et al. 1995, Geiger et al.
1999). The replacement of the acidic phospholipids PG by SQDG was observed in chloroplast membranes of Arabidopsis thaliana (Essigmann et al. 1998, Hartel et al. 1998). In P-deprived
Arabidopsis thaliana, accumulation of the extraplastidic DGDG that resembles PC in fatty acid composition was observed (Hartel et al. 2000). It is evident that P. incisa is not replacing PC/PE by DGDG, but utilizing another strategy, similarly to photosynthetic bacteria, increasing the involvement of DGTS to support ∆6 and ∆5 desaturations during the shortage of phospholipids.
The increase in unsaturation of MGDG and DGDG, found in P. incisa under P- starvation, may reflect an attempt of the cells to adapt residual chloroplastic membranes to unchanged light conditions. An increase in the proportion of AA in MGDG and DGDG under P-
137 starvation in P. incisa appears to evolve from import of AA-containing DAG moieties or CoA- esters derived from PC and PE. Jouhet et al. (2003) reported that in the early stage of P- starvation in cell cultures of Acer sp., a decrease in PE and PG was accompanied by a transient increase of PC and accumulation of DAG with fatty acid composition similar to that of PC, mediated by the activity of phospholipase C. It was suggested that PC-derived molecular species of DAG were selected by galactolipid-synthesizing enzymes (Jouhet et al. 2003). In P. incisa, similar changes in the fatty acid composition of PC and PA suggest an activity of Phospholipase
D, which converts PC to PA. Taking together, there could be a conversion of PE and PC to PA and finally to SQDG or DGTS via DAG.
At the level of individual lipids, it seems that, during P- starvation, the lower proportion of 16:0 in PC is a result of a decrease in the group of molecular species that contain 16:0 at the sn-1 position, while those containing AA at the sn-1 position are still utilized as substrates for the desaturations and contributed AA for TAG synthesis. Although the proportion of PE decreased, desaturation of C18 and C20 appeared to be continued. It appears that there is a decrease in the molecular species of PE (Table 4, section 3.1.3), containing 18:1ω7 at the sn-1 position, which do not participate in desaturations, since they lack intermediate fatty acids of C18 and C20.
However, molecular species containing AA at the sn-1 position were intact. The decrease in
18:1ω7 of PE may be related to the increase of 18:1ω7 in SQDG. Similarly, the decrease in 16:0 may lead to an increase in the molecular species of DGTS, which contain 18:1ω7 or AA at the sn-1 position.
4.3 Enzymes of TAG biosynthesis
Following the elucidation of the biosynthesis of AA in P. incisa (Bigogno et al. 2002a) and the determination of the lipid and molecular species intermediates involved (Section 3.1), it was important to study the enzymes, which may participate in the channeling of AA to TAG,
138 attempting to understand the mechanisms responsible for the deposition of AA in TAG.
The enzymatic steps involved in the biosynthesis of TAG has been well studied in mammals (Bell and Coleman 1983, Mayorek et al. 1989, Lehner and Kuksis 1995), oleaginous yeasts (Oelkers et al. 2002), the oleaginous fungus Mortierella (Pillai et al. 1998) and in the seeds of oil plants (Cao and Huang 1986, 1987, Harwood 1996, Hobbs et al. 1999, Voelker and
Kinney 2001). However, very little is known about the enzymes involved in the biosynthesis of
TAG in algae. TAG biosynthesis occurs mainly in the ER by membrane-bounded enzymes.
Thus, owing to the membrane-bound nature of the enzymes and the hydrophobicity of many lipid substrates and products, the study of lipid metabolism in ER is very difficult. It is also complicated by the fact that many different processes might occur simultaneously in isolated microsomes, making the interpretation of the data difficult (Voelker and Kinney 2001).
To enable studies of activities of enzymes in microsomes (ER-enriched membrane fraction) and oil bodies in P. incisa, optimum conditions for the cell homogenization and fractionation has been determined. The endogenous lipolytic activity was reduced by utilizing a buffer with pH 9.0, containing Mg2+- and Ca2+- chelators, EDTA, EGTA as well as BSA
(Chapman and Barber 1984), which enabled us to obtain intact oil bodies, thus minimizing the contaminations of membrane fractions with the products of oil bodies degradation (FFA, oil bodies proteins).
When DAGAT activity was tested in cellular fractions utilizing [1_14C]acyl-CoA and unlabeled DAG, labeled TAG was produced. Among the cellular fractions, the microsomes showed the highest specific activity. Unexpectedly, a high percentage of total activity was observed in the supernatant 100,000 x g. High proportion of total activity of DAGAT was also recovered in the cellular fraction, that did not precipitate at 100,000 x g, in sunflower oilseeds
(Triki et al. 2000) and in the oleaginous yeast, Rhodotorula glutinis (Gangar et al. 2001).
Nevertheless, in sunflower seeds, this activity was lost after some days, whereas that of
139 microsomes remained for several months. Thus, microsomal preparation was used in the
DAGAT study (Triki et al. 2000). The determination of the cytosolic 10S multienzyme complex, which contains enzymes of TAG synthesis, including DAGAT suggests that the cytosol is one of the sites for triacylglycerol biosynthesis in the oleaginous yeast (Gangar et al. 2001). In P. incisa, however, the presence of the DAGAT activity in the supernatant 100,000 x g could be an artefact. During the homogenization of P. incisa cells, FFA having detergent properties are released and can solubilize DAGAT from membranes. In several cases, the activity of DAGAT has been reported also in the oil bodies. A high level of DAGAT activity was observed in the lipid body fraction of Mortierella ramanniana var. angulispora (Kamisaka and Nakahara 1994) and in the delipidated proteins of oil bodies from developing and mature maize embryos
(Valencia-Turcotte and Rodríguez-Sotres 2001).
However, the majority of the studies with plants located the activity of DAGAT to the microsomal membranes, since the synthesis of TAG occurs in the ER. In developing and germinating seeds of oilseed plants, TAG accumulation and DAGAT activity have been shown to be associated with the ER (high-speed microsomal fraction) (Cao and Huang 1986, Stobart et al. 1986, Stymne and Stobart 1987, Frentzen 1993, Settlage et al. 1995, Lacey and Hills 1996).
The biochemical properties of microsomal DAGAT have been examined in a number of plant systems (Frentzen 1993), including developing seeds (Cao and Huang 1987, Bernerth and
Frentzen 1990, Vogel and Browse 1996) and embryo cultures of Brassica napus seeds (Taylor et al. 1991, 1992, Weselake et al. 1991, Little et al. 1994). In general, studies with developing seeds indicate that DAGAT activity increases rapidly during the active phase of oil accumulation and then decreases markedly as seed lipid content reaches a plateau (Tzen et al. 1993, Weselake et al. 1993).
Taking into account the literature data, a high specific activity in microsomes and low specific activity distributed in the large volume of the supernatant 100,000 x g, to avoid the
140 possibility of an artefact, we concentrated our study on the DAGAT activity in microsomes. In microsomes of P. incisa, the time-dependent incorporation of radioactivity from [1_14C]oleoyl-
CoA into TAG and from [1_14C]dioleoylglycerol into TAG has demonstrated the activity of
DAGAT. The acyltransferase activity was observed without addition of the second substrate,
DAG, but was significantly enhanced by the addition of DAG (Fig. 20).
In many studies with plant microsomes, the activity of DAGAT was shown to be independent of the exogenous DAG, thus complicating studies on substrate (DAG) selectivities in assays with labelled acyl-CoA. For example, in microsomes from cell-suspension cultures of oilseed rape, DAGAT activity was independent of exogenous sn-1,2-dioleoylglycerol in the presence of 3 mM Mg2+, but 100 µM (bulk concentration) sn-1,2-dioleoylglycerol caused a 1.3- fold increase in activity when 25 mM Mg2+ was present (Byers et al. 1999). Little et al. (1994) reported that even the microsomal DAGAT of cell-suspension cultures of oilseed rape, partially purified by solubilization with detergent, was still not dependent on exogenous DAG. The independence of the DAGAT towards exogenous DAG could be due to the presence of a higher concentration of endogenous DAG in the microsomes, or due to unavailability of the DAG to the enzyme.
The formation of DAG micelles and the low solubility of DAG in the aqueous medium is the major obstacle in the DAGAT assay. However, in the present work, the preparation of DAG solution in 0.2% Tween-20 (Cao and Huang 1986, Jako et al. 2001, Triki et al. 2000), followed by heating at 70 °C and vigorous sonication to break the micelles before the addition to the assay medium (Wiberg et al. 1994), has proven to be effective in enhancing DAGAT activity. Another approach that was utilized in the present work to increase DAG solubility and availability for acylation, was the addition of an organic solvent, ethanol (10%, v/v). The stability of DAGAT in the presence of various organic solvents was also observed by Valencia-Turcotte and Rodríguez-
Sotres (2001). The enhanced TAG synthesis in the presence of ethanol might indicate a better
141 solubility of DAG and its availability for the enzyme, or an enhancement of the DAGAT activity. However, the independence of TAG synthesis of exogenous DAG in the presence of ethanol is likely due to the better solubility and adequate availability of endogenous DAG.
Indeed, the labelling of other lipids in the assay decreased with the addition of ethanol, suggested that DAG accepted most of the acyl-CoA, making it less available for other acyltransferases.
In microsomes of P. incisa, the presence of activity in the absence of exogenous DAG, although in a lower scale, also indicates the existence of endogenous DAG in the microsomes.
DAG has been reported to be the most abundant metabolite of the Kennedy pathway in developing seeds of Brassica napus (Perry and Harwood 1993 a,b). Valencia-Turcotte and
Rodríguez-Sotres (2001) attempted to extract the endogenous DAG from maize microsomes with the organic solvents, acetone, benzene, diethylether, etc. Interestingly, the DAGAT activity tolerated sequential acetone washings as shown by the restoration of the activity by the addition of DAG that to a level as high as that obtained to the unwashed freeze-dried microsomes.
However, in P. incisa, the activity recovered after delipidation of microsomes was very low in comparison to untreated microsomes.
In the presence of [1-14C]oleoyl-CoA, DAGAT of P. incisa showed some preference towards the 18:1/18:1 molecular species of DAG over the AA/AA, 18:2/18:2, and 16:0/16:0 molecular species. Possibly, DAGAT prefers acyl-CoA and DAG with the identical acyl residues. Arachidonoyl-CoA, the most prevalent acyl-CoA in the cellular pool under conditions of TAG accumulation, thus DAGAT might preferentially select DAG with one or two arachidonoyl moieties, as most of the TAG molecular species contains two or three arachidonoyl moieties. For example, the microsomal DAGAT of Cuphea procumbens (a high C10:0 oilseed) preferentially selects 10:0-containing substrates over oleoyl-containing substrates (Wiberg et al.
2000). However, in several conventional or transgenic oilseeds, microsomal DAGAT displays broad acyl-CoA selectivities and even preferred saturated CoA substrates (Cao and Huang 1987,
142 Frentzen 1998). Transgenic canola, which is high in medium chain fatty acids, accumulated up to
80% saturates at the sn-1 and the sn-3 positions of TAG, suggesting a dependence of the acyl composition at the sn-1 and the sn-3 positions on the acyl composition of the available substrates rather than acyl transferase specificity (Voelker and Kinney 2001). In P. incisa, when the synthesis of AA was inhibited and oleic acid was accumulated by the effect of an inhibitor
(SHAM), acyl composition of TAG was changed to oleic-rich species (Bigogno et al. 2002c).
This data may indicate an absence of strict specificity of acyltransferases for the acyl- composition of the substrates.
An inhibition of the enzyme activity by thiol-modifying reagents is regarded as an indicator of the presence of sulfhydryl (SH-) group in the amino acid residues of the active site, and accordingly, the presence of cysteine or serine in the vicinity of active site. In microsomes of P. incisa, the significant inhibition of the activity of DAGAT in the presence of the sulfhydryl reagent, PCMB, indicates the importance of cysteine or serine residues for the activity of
DAGAT, which is in line with the findings of the inhibition of the in vitro DAGAT activity in rat microsomes with thiol-modifying reagents (Lehner and Kuksis 1995).
The bivalent cation Mg2+ was shown to affect DAGAT activity in many cases. Generally, a range of 0 to 3 mM of MgCl2 is utilized with plant microsomes (Valencia-Turcotte and
Rodríguez-Sotres 2001, Hobbs et al. 1999). However, magnesium salts, MgSO4 and MgCl2 (25 mM), were found to enhance DAGAT activity in microsomes of Brassica napus (Byers et al.
1999). In the present study, an increase in the MgCl2 concentration above 3 mM resulted in an inhibition of DAGAT, in agreement with the decrease in the solubility of acyl-CoA at high concentrations of Mg2+, causing its aggregation in the medium (Constantinides and Steim 1985).
To avoid the possibility of artifacts arising due to the labeling and interconversion of several other lipids during the assay conditions, labeled DAG and non-labeled acyl-CoA were also utilized to assess the DAGAT activity. The time-dependent incorporation of label from
143 DAG into TAG was dependent on the acyl-CoA concentration. This type of assay allows testing the selectivity of the enzyme towards the acyl-CoA substrate. When different types of acyl-CoAs were utilized as acyl donors for the acylation of [1-14C]dioleoylglycerol, the activity was preferentially selective to C18 acyl-CoA substrates. Arachidonoyl-CoA was less utilized. The selectivity of the enzyme to acyl-CoA substrate may be dependent upon the type of acyl composition of DAG. The enhancement of TAG synthetic activity by ethanol and inhibition by
PCMB in assays with labeled DAG, has been similar to that in assays with labeled acyl-CoA, suggesting the same type of activity observed in both types of DAGAT assays.
On the other hand, a sharp enhancement of the incorporation of label from [1-
14C]dioleoylglycerol into TAG at the higher Mg2+ concentration (Fig. 32), together with an independence of the activity on added acyl-CoA (Fig. 33), is likely due to another mechanism of
TAG synthesis, which does not require acyl-CoA. Since acyl CoA has been reported to be less soluble at high Mg+2 and the application of acyl CoA did not change the activity, it is less likely that acyl-CoA dependent TAG synthesis occur. Instead, it allows suggesting the activity of another enzyme of TAG synthesis, which also utilizes DAG as substrate, but does not utilize acyl-CoA. The acyl-CoA independent incorporation of radiolabeled DAG into TAG allows to suggest the activity of a DAG:DAG transacylase. Stobart et al. (1997) has reported an acyl-CoA independent TAG synthesis by a transacylation reaction between two molecules of DAG, releasing a molecule of MAG and catalyzed by the enzyme DGTA (Fig. 7, Section 1.4.5). In contrast to DAGAT, this activity in microsomes is inhibited by ethanol, but not by PCMB. This suggests that in contrast to DAGAT, DGTA does not contain thiol groups (or SH- group containing cystein or serine residues) in the active site of the enzyme. DGTA is likely to require
Mg2+ as co-factor for its enzymatic activity.
The cellular fractionation of P. incisa produces a thick floating layer of oil bodies, which are bright yellow in color due to the presence of β-carotene. Oil bodies are rich in NL and also
144 contain some PL and proteins. In addition to microsomes, biosynthetic activity leading to the production of TAG was also shown in oil bodies (Christiansen 1978, Kamisaka and Nakahara
1994, Valencia-Turcotte and Rodríguez-Sotres 2001, Sorger and Daum 2002). In the oleaginous fungus, Mortierella ramanniana var. angulispora, the highest activity of DAGAT was observed in oil bodies (Kamisaka and Nakahara 1994).
In P. incisa, the activity of DAGAT was rather minor in native oil bodies, when [1-14C] oleoyl-CoA was applied. However, oil bodies proteins, delipidated with diethyl ether, had catalyzed the incorporation of labeled acyl-CoA into TAG, the rate of the incorporation was dependent on the presence of DAG in the assay medium. The DAGAT activity was substantially increased also in oil bodies of maize following delipidation (Valencia-Turcotte and Rodríguez-
Sotres 2001). In P. incisa, DAGAT has shown a preferential selectivity to the dioleoylglycerol substrate over the diarachidonoylglycerol, in the presence of [1-14C] oleoyl-CoA. When [1-
14C]dioleoylglycerol was applied as a labeled substrate, even native oil bodies showed a DAGAT activity as shown by the time-dependent incorporation of [1-14C]dioleoylglycerol into TAG (Fig.
41). Apparently, labeled DAG was more available than acyl-CoA to the DAGAT of the oil bodies. Inhibition of the activity by nicotinic acid (an inhibitor of DAGAT in mammalian system, Ganji et al. 2004) further ascertained the activity of DAGAT in the alga.
The thiol-modifying reagents PCMB and DTT suppressed the activity. The severe inhibition of the activity by the thiol-reagent, PCMB allows us to suggest the presence of catalytically important serine or cysteine residues in the active site of the enzyme (Lehner and
Kuksis 1995).
Similarly to microsomes, increasing MgCl2 concentration to 150 mM highly enhanced
14 the incorporation of [1- C]1,2 dioleoylglycerol into TAG in oil bodies. At 150 mM MgCl2, oil bodies could utilize [1-14C]1,2 dioleoylglycerol as their sole acyl donor for the synthesis of TAG, revealing the presence of a DAG:DAG transacylase (DGTA) activity as in microsomes.
145 However, in contrast to microsomes, TAG synthesis from [1-14C]1,2 dioleoyl-DAG in oil bodies was enhanced by the addition of oleoyl- and arachidonoyl-CoAs and was inhibited by the thiol- specific reagent PCMB. This indicates to the presence of DGTA in oil bodies, which is sensitive to PCMB, indicating that free sulfhydryl groups of cysteine residues were involved in the acyl transfer reactions as in DAGAT of microsomes. It appears that there are two forms of DGTA;
PCMB sensitive in oil bodies and PCMB insensitive in microsomes.
In the DAGAT assay with labeled oleoyl-CoA using microsomes, in addition to TAG, several other lipids were also labeled, indicating the activity of several enzymes involved in the lipid metabolism of this alga, directly or indirectly involved in the biosynthesis of TAG (Fig. 19,
74). Thus, our results indicate the activity of various active acyltransferases, utilizing endogenous substrates in the microsomes of P. incisa as shown in Fig 74. For example, PC and
PE could be labeled by acylation of endogenous lysophospholipids and further converted to
DAG by reverse action of DG-CPT/EPT- phosphotransferase. A similar mechanism might be proposed for the labeling of the betaine lipid DGTS, however, neither lyso-DGTS acyltransferase nor an enzyme, involved in DGTS – DAG conversion, were characterized so far.
The particularly high labeling of DAG, the immediate precursor of TAG and a central intermediate in lipid metabolism, may result from the acylation of endogenous MAG by
MAGAT. Degradation of labeled TAG by a lipase or of phospholipids by phospholipase C and
D can also bring about the formation of labeled DAG (Fig. 74). PA phosphatase or a reverse action of DAGAT can also contribute to DAG formation. MAG could be produced by lipolysis of DAG or as a result of transacylation of two molecules of DAG (DGTA). It is likely that these enzymes, may work in some sort of coordination, to channel AA into TAG (Fig. 74).
146 LPL Observed in our assays G3P Observed in our inhibitor studies PLA
PLAT Not studied in the present work PC, PE LPA PLD PLC
DG-CPT/EPT PDAT LPAAT G3PAT DGTA PAP PA DAG DAGAT TAG Lipase ? Lipase DGTS MAGAT MAG
Figure 74. Proposed pathways of TAG biosynthesis in P. incisa. G3PAT- Glycerol-3-phosphate acyltransferase, LPAAT- Lysophosphatidic acid acyltransferase, PAP- Phosphatidic acid phosphatase, PLAT- Phospholipid acyltransferase, PLA- Phospholipase A, PLC- Phospholipase
C, PLD- Phospholipase D, DG-CPT- Diacylglycerol cholinephosphotransferase, DG-EPT-
Diacylglycerol ethanolaminephosphotransferase, MAGAT- Monoacylglycerol acyltransferase,
PDAT- Phospholipid-diacylglycerol acyltransferase, DGTA- Diacylglycerol transacylase (the thicker arrow represents a stronger activity), DAGAT- Diacylglycerol acyltransferase.
Several evidences were obtained to support the MAGAT activity in the microsomes of P. incisa. The formation of labeled DAG was observed in the DAGAT assay with [14C]oleoyl-CoA, possibly due to the acylation of the endogenous MAG. More directly, when 2-MAG was utilized as the sole acceptor of labeled oleoyl-CoA, the increase in labeling of DAG was linear with time
147 (Fig. 53). When ethanol was present in the DAGAT assay, an increase of oleoyl-CoA incorporation into TAG was concomitant to a decrease in DAG labeling. This could also indicate a faster utilization of DAG formed by MAGAT for further acylation by DAGAT. However, other experiments have shown that ethanol inhibited the activity of MAGAT (Fig. 54, 55) thus probably only DAGAT activity was responsible for the incorporation of label into TAG.
To avoid the interference of endogenous substrates, DAG and acyl-CoA, and several other enzymes utilizing the same substrates in DAGAT assay, purification of the enzyme or obtaining the enzyme by the expression of the DAGAT gene in a heterologous system can be suggested for future studies.
Another observation in the DAGAT assay was that the labeling of DAG increased when unlabeled DAG was provided (Fig. 56). This was likely due to partial lipolysis of unlabeled
DAG, releasing MAG, which in turn was reacylated with labeled acyl-CoA (MAGAT).
However, we cannot exclude a lipolysis of labeled TAG. In microsomes of developing peanut cotyledons, labeled DAG was obtained by MAGAT activity, rather than by hydrolysis of TAG or PC (Tumaney et al. 2001). This was the first plant MAGAT that was further purified to homogeneity and characterized.
Along with activities of acyltransferases, the lipase activity was deduced in the microsomes by the release of FFA from the labeled DAG substrate. The pH, temperature, time and protein dependence for the activity has been determined. The pH optimum at 4.5 suggested a presence of acidic lipase in P. incisa. The thiol-modifying reagents, PCMB, NEM and iodoacetamide, inhibited the activity while the thiol-protecting reagent DTT enhanced the activity, indicating the importance of amino acids with SH-group for the enzyme activity. The presence of a catalytically essential -SH group at the active site is well-known property of lipases. We also tested the inhibitor RHC, which specifically inhibits the DAG-lipase in mammalian system (Caroll and Severson 1992, Sutherland and Amin 1982). The lipolysis of
148 DAG was inhibited, however, up to 50% only, despite a range of concentrations of the inhibitor.
We infer that about 50% of the activity seen in the assay, can be related to a non-specific lipase, able to hydrolyze, for example, also TAG. Indeed, labeled TAG was hydrolyzed in an assay with microsomes (data not shown).
The lipolytic activity demonstrated fatty acid and positional selectivity. When 1,2 [14C] dioleoylglycerol was utilized, oleate was released from both positions. Whereas, the lipolysis of
1-oleoyl-2-[14C] arachidonoylglycerol, preferentially released oleoyl moieties from the sn-1 position, preserving the sn-2 arachidonoyl moiety intact in MAG. We interpret these data as an evidence for the existence of a pathway by which AA originated from the DAG intermediate is preserved in MAG rather being released to the FFA pool. This MAG can thus be sequentially acylated by MAGAT and DAGAT to enrich TAG with AA.
Taking together, several enzymes, directly or indirectly, may play a role in the biosynthesis of AA-rich TAG in the alga. Probably, they are assigned to maximize the specific incorporation of AA into the glycerol backbone of the TAG at the different metabolic steps. For example, PC and PE may produce AA-rich DAG, by the reverse action of phosphotransferases
CPT or EPT or by the action of phospholipase C and D. DAG is produced in the Kennedy pathway via lyso-phosphatidic acid acyltransferase (LPAAT) and phosphatidic acid phosphatase
(PAP). DAG in turn may form MAG by lipolysis or by the activity of DGTA. MAG can again be acylated to produce DAG, possibly with AA moieties from acyl-CoA pool. TAG containing shorter and less desaturated fatty acids, may also be hydrolyzed by a lipase to produce DAG. The
DAG, either by the activity of DAGAT or DGTA, can be transformed to TAG. In addition to the acylation of lipid substrates by transfer of acyl moieties from one lipid to another, the acylation of those are, mostly, contributed by the acyl-CoA pool of the cytosol, which collects different acyl-CoA obtained from the extrachloroplastic lipids as well as de novo lipids formed in the chloroplasts.
149 4.4 Role of TAG
4.4.1 Nitrogen starvation and recovery
The unique accumulation of AA-rich TAG in oil bodies, especially under N-starvation lead to hypothesis that in P. incisa TAG may have an additional role other than being a storage of carbon and energy (Bigogno et al. 2002c; Khozin-Goldberg et al. 2002a). While most microalgae reside in large water bodies where temperature and nutritional changes are relatively slow, quite a few microalgae grow in ecological niches that may be subject to rapid short-term fluctuations e.g., temperature, salinity or nitrogen availability. Indeed, P. incisa was isolated from an alpine environment characterized by sudden changes in environmental conditions
(Bigogno et al. 2002b). Adaptation to these changes may require significant and rapid alterations in the fatty acid and molecular species composition of chloroplast membrane lipids. However, under such conditions, the de novo synthesis of PUFA would be too slow. We have thus hypothesized that in algae growing in such habitats, PUFA-rich TAG can be metabolically active serving as a buffering capacity for PUFA, providing specific acyl groups, in order to enable rapid adaptation of the membranes.
When N-starved cells of P. incisa are transferred to optimal growth conditions, resumption of photosynthesis requires swift construction of the various photosynthetic membrane components, among them the chloroplastic lipids. This requirement could be crucial since the window of opportunity for exponential growth could be rather narrow. During recovery the AA content decreased but somewhat less than the fatty acid content (Fig. 65). There was a significant reduction in the content of TAG and a smaller increase in that of chloroplastic lipids
(Fig.67). The simultaneous decrease of AA-rich TAG and increase of AA in chloroplastic lipids
(Table 19, Fig. 67) strongly supports our hypothesis. Similarly, when the growth temperature of an exponentially growing culture of P. incisa labeled with [1-14C]arachidonic acid was suddenly dropped to 4 °C, the label in TAG, which was mostly associated with AA, was turned over to
150 polar lipids (Bigogno et al. 2002c). Likewise, it was found that AA- rich TAG of P. cruentum contributed AA moieties for the production of eukaryotic-like molecular species of MGDG, containing EPA at the sn-1 and sn-2 positions of the glycerol skeleton following N- replenishment (Khozin-Goldberg et al. 2000). Also in higher plants, lipid bodies were recently shown to be metabolically active in seeds and other plant organs (Murphy, 2000). Furthermore,
Stobart et al. (1997) produced evidence that supports a transacylation mechanism that can account for TAG turnover in microsomal membranes of developing safflower seeds.
Under optimal conditions the major PUFA of MGDG in P. incisa, as in many other green algae, are 16:3ω3 and 18:3ω3. However, it contains also 8.9% AA (of total fatty acids) (Table
1). Under N-starvation the share of AA decreased to 2.4% (Table 19) and the ω3 fatty acids were mostly replaced by their ω6 precursors, 16:2 and 18:2. Attenuation of ω3 desaturation and accumulation of the ω6 precursors, under conditions that do not support optimal growth, is a well documented phenomenon in both higher plants (Somerville and Browse 1996) and algae
(Klyachko-Gurvich et al. 1997, Routaboul et al. 2000). Similarly, in P. cruentum, under suboptimal growth conditions, chloroplastic lipids, predominantly MGDG, accumulate AA
(20:4ω6) rather than EPA (20:5ω3). The increase in membrane unsaturation and consequently of its fluidity, as a means of cold adaptation, is a well-studied phenomenon in both lower and higher plants (Murata and Wada, 1995). Indeed, upon recovery at 12 °C, the need for high level of unsaturation in MGDG is provided by an intensive ω3 desaturation of the C16- and C18- containing molecular species (Table 20).
Our findings indicate that in P. incisa there are two pathways leading to the production of molecular species of chloroplastic lipids (Fig. 75). The first pathway is the prokaryotic pathway that in similarity to 16:3-plants such as Arabidopsis thaliana (Browse and Somerville 1991) and several green algae, e.g., Chlorella (Thompson 1996, Sato et al. 2003) and Chlamydomonas
151 (Giroud et al. 1988), gives rise to the 18/16 type molecular species of MGDG (and presumably other chloroplastic lipids) in P. incisa. The second pathway is the eukaryotic pathway, also
TAG
DGTS PC
DAG
Cytoplasm Chloroplast
18:1 18:1 20:4 16:1 18:2 18:1 Gal Gal Gal
ω6D
18:2 18:2 20:4 16:2 18:2 18:2 Gal Gal Gal
ω3D
18:3ω3 18:3ω3 20:4 20:4 16:3ω3 18:3ω3 18:3ω3 20:4 Gal Gal Gal Gal
Prokaryotic Eukaryotic
Figure 75. Outline of suggested pathways in the biosynthesis of MGDG in P. incisa. Dashed
arrow- Rapid deployment pathway. Gal- Galactose moiety.
common in higher plants and green algae. In these organisms, this pathway most likely import
DAG from extrachloroplastic lipids and provides membranal lipids with C18-containing molecular species that can be further desaturated by the chloroplastic ω3 desaturase (Browse and
152 Somerville 1991, Sato et al. 2003). In P. incisa, 3 types of molecular species, 18/18, 20/18 and
20/20 are produced via this pathway. These molecular species appears to derive from DGTS and
PC but also in TAG (Figs. 68, 69).
We hypothesize that P. incisa have three modes of operation with respect to the production of chloroplastic lipids. When environmental conditions do not support growth, e.g., in the stationary phase or under N-starvation, the prokaryotic pathway predominates, comprising over 70% of total MGDG. Upon recovery from nitrogen deficiency that is compounded by a low temperature shift, maximal growth cannot be immediately supported, as the organism has to cope with both recovery and low temperature. Under such circumstances, the eukaryotic pathway is invoked, importing acyl (mostly 18:1, 18:2 and AA) moieties from DGTS and PC (presumably as DAG). This mode is also contributing under exponential conditions. The third mode is utilized when there is a sudden requirement for enhanced desaturation of chloroplastic lipids under conditions that support exponential growth, e.g., upon recovery from N-starvation at room temperature or when an exponentially growing culture is undergoing a drastic temperature down shift (Bigogno et al. 2002b). In such cases, the acyl content available from DGTS and PC would not suffice and TAG are rapidly deployed to enhance the flux in the eukaryotic pathway, releasing AA-rich acyl moieties that can be exported into the chloroplast. We presume that AA is imported as DAG that is released from a phospholipid or TAG (Fig. 75). A TAG lipase is likely the key enzymatic activity involved in the release of AA from TAG as evidenced by the increase in AA in FFA. Indeed, the presence of a highly active TAG lipase was detected in cell-free homogenates and microsomes of P. incisa (section 3.3.1 and 3.3.11).
In higher plants the ability to change the ratio of eukaryotic to prokaryotic molecular species, under different environmental conditions, is rather limited. However, in the red alga, P. cruentum, the share of eukaryotic-like molecular species of MGDG increases from 42% (of total acyl MGDG) at 30 °C to 58% at 20 °C (Adlerstein et al. 1997). Indeed, this alga was isolated
153 from wet saline soil, another ecological niche characterized by rapid fluctuations in environmental conditions. Recently, Falcone et al. (2004) have shown that in higher plants too, the ratio of eukaryotic to prokaryotic molecular species is affected by temperature.
The data we have shown suggest that upon transfer to growth conditions at low temperatures, the major adaptation processes used by cells of P. incisa are the construction of new molecular species, produced via the eukaryotic pathway and the enhancement of ω3 desaturation of preexisting, as well as newly formed, molecular species of MGDG (and apparently also of
DGDG). At 24 °C however, ω3 desaturation is lesser than at 12 °C. Instead, the eukaryotic mechanism is significantly intensified. The share of molecular species produced in the eukaryotic pathway increased from 16.0% to 62.8% of total MGDG (Table 20). However, the reason why two diverse types of PUFA, 16:3 and 18:3 of the ω3 family, and AA of the ω6 family, are preferentially produced under different conditions is still unclear. Possibly, the role of the AA-containing molecular species is to provide increased chain length, as well as enhanced desaturation.
While the ratio of the different groups of molecular species was significantly different between 12 °C and 24 °C, the ratio of the eukaryotic molecular species of the 18/18, 20/18 and
20/20 types was only slightly different, being 24:61:14 at 24 °C and 21:63:15 at 12 °C, respectively. This finding indicates that the rapid deployment of acyl groups does not change the ratio of these groups in the sink although under N-starvation, the AA content of TAG is 50% in comparison to only 13.1 and 18.6% in DGTS and PC, respectively. Indeed, the enrichment of the latter lipids with 16:0 during recovery at 12 °C suggests that only certain molecular species are selected for export.
Plant desaturases are classified according to their specificities. Several desaturases introduce a double bond at a certain distance from the carboxylic end, e.g., ∆6 or ∆9, while other desaturate at a position that is relative to the methyl end of the fatty acid, e.g., ω3. There is a
154 general agreement in the literature that the ultimate chloroplastic desaturase is an ω3, rather than a ∆15, desaturase, since it is able to desaturate both 16:2 and 18:2 (Browse and Somerville
1991). Surprisingly, it appears that in P. incisa, this enzyme can desaturate both fatty acids, but not 20:4ω6 to 20:5ω3. In P. cruentum, the chloroplastic ω3 desaturase can desaturate AA to
EPA, however the 18:2/16:0 molecular species of MGDG is not further desaturated (Khozin et al. 1997). In contrast, Sakuradani et al. (2005b) recently showed that the ω3 desaturase of the
AA-producing fungus Mortierella alpina1S-4 can desaturate both C18 and C20 PUFA. This is, to the best of our knowledge, the first reported case of an ω type enzyme, i.e., an enzyme that
“counts” from the methyl end of the fatty acid chain that is specific both to the carbon atom position and to the chain length.
4.4.2 Recovery from SHAM treatment
Comparatively slower build-up of chlorophyll and dry weight following growth recovery in low-
AA, SHAM-treated cultures indicated to the importance of AA-rich TAG for chloroplast development. The similar pattern of increase in 18:3ω3 during growth recovery in control and
SHAM-treated cultures indicated that the lag in chlorophyll synthesis in SHAM-treated culture could be due to the lower level of AA, but not to insufficient production of 18:3 ω3. One may suggest that the slower recovery was due to presence of residual amounts of the inhibitor.
However, the synthesis of AA under these conditions suggests otherwise.
The SHAM-treated, low AA-containing cells, showed a rapid synthesis of AA during recovery, possibly because of the availability of accumulated 18:1ω9 precursor in extraplastidial lipids and TAG. The volumetric content of TFA did not change during first 2 d of recovery in
SHAM-treated culture, but the content of 18:1ω9 was reduced, simultaneously with the increase in AA content.
155 During 1-2 d of recovery, while the de novo synthesis of fatty acids was not yet sufficient, AA content was increasing much faster in SHAM treated culture, indicating the recycling of existing C18 precursors, located in polar lipids and TAG, for the synthesis of AA
(Table 21). Likewise, in sunflower (Helianthus annuus L.) when the developing seeds were transferred to low temperature, the total amount of oleate found in TAG decreased as that of linoleate increased, while the contents of total lipids and TAG remained unchanged suggesting that oleate from TAG were used for desaturation (Garces et al. 1994). Hence, it can be proposed that upon sudden change in nutrient availability in P. incisa, C18 FA are released from TAG, desaturated, elongated into AA, and deposited in either TAG or membrane lipids. Similarly to the recovery from N-starvation, we can propose that AA supply for chloroplastic lipids may be prerequisite for growth recovery from unfavorable conditions.
4.5. Mutant studies
This part of the research was carried out to generate mutants, which are disrupted or impaired in the biosynthesis of AA and TAG. Fatty acid and lipid analysis of such mutants will aid understanding the mechanism underlying the biosynthesis of AA-rich TAG and also their role in the organism. For example, mutant of Nannocloropsis devoid of EPA was obtained by random selection of mutant colonies, obtained by NNG treatment (Schneider et al. 1995). The preliminary selection out of thousands of NNG-treated colonies was carried out visually by selecting poor- grown and better-grown colonies. Further screening of selected colonies was carried out by plating replicates onto three agar plates prepared with BG11, nitrogen-free BG11 and phosphate-free BG11, respectively, to follow their growth and performance.
The screening of 300 selected colonies by fatty acid analysis showed the absence of AA- deficient mutants. Many of the colonies thus selected were not mutants with the altered fatty acid composition and content. However, variations in the fatty acid distribution and lipid content
156 indicated the production of lipid-biosynthesis mutants. For example, the mutants B50, 83, 107,
109, 114, NB5 contained very low proportions of AA, while B33 and 105 contained higher proportions of AA.
The mutant B105, which contained a higher proportion of TFA in comparison to WT, but no significant change in the fatty acid composition, indicated an enhanced TAG biosynthesis. In contrast, mutants B107 and B109, which contained a lower proportion of AA and TFA as shown by their lower percentages of biomass in comparison to WT indicated an inhibition of AA and
TAG synthesis. However, significant increase in 18:1 and 18:2 in those mutants indicated mutations in ∆12 or ∆6 desaturases. In AA-producing fungus, Mortierella alpina, mutants deficient in ∆12 or ∆6 desaturases were characterized by elevated levels of these fatty acids
(Sakuradani et al. 2005a). Lipid type analysis of B105, B107 and B109 showed lower levels of neutral lipids in comparison to WT. In B107, the presence of the lowest level of NL and highest level of PL among the three mutants studied, might indicate some problem in the transfer of acyl moieties from PL to NL.
Indeed, analysis of polar lipid classes of B107 showed an increase in the content of PC,
PE and DGTS, which are involved in the desaturation of C18 and C20 fatty acids. 18:3ω6 was almost absent from PC, PE and DGTS, while18:1 ω9 was significantly increased in all three polar lipids, supporting the inhibition of ∆12 and ∆6 desaturases. When ∆12 and ∆6 desaturases were inhibited in P. incisa by the administration of the inhibitor SHAM, AA synthesis was inhibited and 18:1 increased, but not the TFA content (Bigogno et al. 2002c). In B107, the inhibition of desaturation of C18 fatty acid together with the decrease in TFA indicated the inhibition of enzymes, which release acyl moieties from these lipids to the acyl/FFA pool or produce DAG (phospholipases of lyso-phospholipid acyltransferase) to enable AA and TAG production.
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184
תקציר החומצה הארכידונית (AA), חומצה שומנית רב-בלתי-רוויה (PUFA), היא בעלת חשיבות רבה להתפתחות תאי מוח בתינוקות. האצה Parietochloris incisa מכילה כמויות גדולות ביותר של AA, עד 21% מהמשקל היבש. רוב ה-AA נמצא בטריגליצרידים. הבהרה של המסלולים המעורבים בביוסינתיזה של טריגליצרידים אלה היא על-כן בעלת חשיבות מרובה. בעבודה זו נעשה מאמץ לזיהוי האנזימים והליפידים המעורבים בסינתיזה ברמת molecular species. ה-PUFA 20:3 ו-AA נמצאו בעיקר בליפיד PE ואילו 18:2 ו18:3- בליפידים PC ו-DGTS. נתונים אלה מצביעים על מעורבות PE בדסטורציה מסוג 5∆ ומעורבות PC ו-DGTS בשלבים הקודמים בעמדות 12∆ ו6-∆ . אנליזה של molecular species הראתה כי הדסטורציות 12∆ ו6-∆ מתרחשות בעמדה sn-2 של השלד הגליצרידי. לעומת זאת, דסטורציה 5∆ ב-PE מתרחשת גם ב- sn-1 וגם בsn-2- . רוב ה-molecular species של TAG מכילים 2 או 3 קבוצות AA. על בסיס הרכב ה- molecular species של PE ניתן לשער מסלול בו PE משחרר דיאציל גליצרול (DAG) המכיל קבוצה אחת או שתיים של AA וע"י פעולת האנזים DAGAT הופך ל-TAG. על-מנת לאפיין את פעילות DAGAT פותח פרוטוקול ובוצעה פרקציונציה של התאים. יצירה in vivo של TAG נמדדה תוך שימוש בסובסטרטים רדיואקטיביים 18:1-DAG או 18:1-CoA. פעילות DAGAT נמצאה בפרקציות ממברנליות ובחלבונים של גופיפי שמן (oil bodies). בנוסף, נמצאה פעילות של אנזים אחר, DGTA, היוצר TAG משתי מולקולות של DAG ע"י העברה של קבוצות אציל. על בסיס הממצאים בעבודה זו הוצעה סכמה לביוסינתיזה של AA באצה. כמו-כן, הוצעה היפותיזה הטוענת כי התכולה הגבוהה של AA ב-TAG באצה מסייעת להתמודדות עם שינויים בתנאי הסביבה. באצות של מקווי מים גדולים שינויים בתנאים כגון טמפרטורה, מתרחשים בהדרגה. עם ירידת הטמפרטורה, עולה בהדרגה דרגת האי - ריוויון של החומצות השומניות בממברנה על-מנת לשמור על פלואידיות גבוהה. לעומת זאת, באצות שמוצאן בסביבה אלפינית השינויים מתרחשים תוך דקות ועל - מנת לספק לממברנה במהירות דרגת אי-ריוויון נאותה, יש צורך באגירה של PUFA בגופיפי שמן. על-מנת לאשש היפותיזה זו, נבדקו שינויים בהרכב חומצות שומן בליפידים השונים ב-oC 24 ו- oC 12, לאחר התאוששות מהרעבה לחנקן. בשני המקרים רוב ה-TAG נצרך והיה מעבר של AA לליפידים כלורופלסטיים, בעיקר MGDG. על בסיס ממצאים אלה ואחרים, הוצע כי בניית הליפידים הכלורופלסטיים מתרחשים ב3- אופנים. כאשר תנאי הסביבה אינם תומכים בגידול, המסלול העיקרי הוא פרוקריוטי שבו נוצרים molecular species מסוג 18/16. כאשר מתרחש שינוי בתנאי הסביבה והגידול מתאפשר, המסלול האאוקריוטי גובר ובו נוצרים molecular species מסוג 18/18 או 20/18. כאשר מתאפשר גידול מהיר ביותר, מופעל המסלול השלישי המאופיין על-ידי יצירת molecular species מסוג 20/20 ב-MGDG.
ביוסינתיזה ושינוע של טריאצילגליצרולים עתירי חומצה ארכידונית באצה הירוקית פריאטוכלוריס אינציסה
מחקר לשם מילוי חלקי של הדרישות לקבלת תואר "דוקטור לפילוסופיה"
מאת
פושקאר שרסטה
הוגש לסינאט אוניברסיטת בן - גוריון בנגב
חשון תשס"ו נובמבר 2005
באר-שבע
ביוסינתיזה ושינוע של טריאצילגליצרולים עתירי חומצה ארכידונית באצה הירוקית פריאטוכלוריס אינציסה
מחקר לשם מילוי חלקי של הדרישות לקבלת תואר "דוקטור לפילוסופיה"
מאת
פושקאר שרסטה
אישור המנחים
פרופ' צבי הכהן ______
פרופ' בצלאל קסלר ______
ד"ר אינה חוזין-גולדברג ______
אישור דיקן בית הספר ללימודי מחקר מתקדמים ______
חשון תשס"ו נובמבר 2005
באר-שבע