FUNCTIONS AND MECHANISMS OF CHLOROPLAST LIPID REMODELING IN PLANTS
By
Kun Wang

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Biochemistry and Molecular Biology – Doctor of Philosophy
2017

ABSTRACT
FUNCTIONS AND MECHANISMS OF CHLOROPLAST LIPID REMODELING IN PLANTS
By
Kun Wang
Chloroplasts in plants and algae are organelles that carry out a number of important
functions including photosynthesis, fatty acid biosynthesis and oxylipin mediated plant immune
responses. Chloroplast membranes with a unique lipid composition are crucial for the functions of
chloroplasts. Chloroplast membranes are composed of a specialized glycerolipid matrix with
predominately galactolipids, mono- and digalactosyldiacylglycerol (MGDG and DGDG,
respectively),

and

two

anionic

lipids,

phosphatidylglycerol

(PG)

and

sulfoquinovosyldiacylglycerol. The lipid composition of the chloroplast membranes is finely tuned
in response to stresses or during various developmental stages to maintain plant fitness.
SENSITIVE TO FREEZING 2 (SFR2) is a galactolipid : galactolipid galactosyltransferase
that is highly conserved in land plants. In response to freezing, SFR2 is activated in freezing
tolerant plants like Arabidopsis and remodels MGDG to form a class of oligogalactolipids that
stabilize the bilayer structures. Phylogenetically, SFR2 is highly conserved in land plants including
those that are freezing sensitive. Therefore, I hypothesized that SFR2 in freezing sensitive plants
may be involved in plant resilience to other abiotic stresses, like salt or drought stresses because
similar to freezing, salt and drought stress cause cell dehydration. In Chapter 2, freezing sensitive
tomato was used as a model to test this hypothesis. I generated the SlSFR2 RNAi lines and
observed their hypersensitivity to salt and drought responses. Decreased tolerance was correlated
with compromised production of oligogalactolipids in the SlSFR2 RNAi lines, suggesting SFR2
plays a role in salt and drought tolerance in freezing sensitive plants.

Maintenance of the chloroplast membranes requires a finely tuned lipid anabolic and
catabolic machinery. Contrary to the chloroplast lipid biosynthetic enzymes, a large number of the
predicted lipid-degrading enzymes in the chloroplasts have unknown functions. In Chapter 3, I
identified over 50 chloroplast localized lipases in silico. Using a reverse genetic approach, I first
focused on the characterization of an Arabidopsis thylakoid membrane-associated lipase,
PLASTID LIPASE1 (PLIP1). PLIP1 is a phospholipase A1. In vivo, PLIP1 hydrolyzes
polyunsaturated acyl groups from a unique chloroplast-specific PG that contains 16:1∆3trans at the
sn-2 glyceryl position. PLIP1 is predominately expressed in seeds, and the plip1 mutant seeds
contain less oil while the PLIP1 overexpression seeds contain more compared to the wild type.
Pulse-chase labeling assays suggest that the acyl groups released by PLIP1 are exported from the
chloroplasts, reincorporated into phosphatidylcholine before ultimately entering seed
triacylglycerol. Therefore, 16:1∆3trans uniquely labels a biochemical active PG pool in developing
Arabidopsis embryos, which is subject to PLIP1 mediated acyl export and seed oil biosynthesis.
The Arabidopsis genome encodes two putative PLIP1 paralogs which are designated PLIP2
and 3. PLIP2 and PLIP3 are also localized in the chloroplasts, but with different subplastid
locations. PLIP2 possesses similar biochemical properties in vitro to PLIP1, but in vivo studies
suggest that PLIP2 prefers MGDG as substrate, while PLIP3 prefers PG. Overexpression of PLIP2
or PLIP3 severely stunts plant growth and leads to accumulation of oxylipins. Genetically blocking
jasmonate signaling restored the growth of the overexpression plants. The expression of PLIP2
and PLIP3, but not PLIP1 is induced by abscisic acid (ABA), and plip1,2,3 triple mutants exhibit
compromised jasmonate biosynthesis in response to ABA. Therefore, in Chapter 4, I am proposing
that PLIP2 and PLIP3 are involved in linking ABA responses with jasmonate production.

ACKNOWLEDGEMENTS

First and foremost, I would like to thank my advisor and mentor Christoph Benning. He
has been supportive since the day I joined the lab. I appreciate all his contribution of time, ideas
and funding to make my Ph.D. experience productive and stimulating. He has taught me,
consciously and unconsciously, what a great scientist should be. He is a very patient mentor.
Despite my lack of experience in plant biology and biochemistry training when I joined the lab, he
saw my potential and accepted me. I still remember vividly that he took time to teach me plant
physiology himself every Friday afternoon in my third year to prepare me for my tomato project.
After I was a bit mature, he allowed me more freedom to pursue the directions that I am interested
in and trusted me to independently design projects. He always encouraged me to get out of my
comfort zone, and to challenge and improve myself, an attitude surely will benefit me for my whole
life.
I would also like to thank the members of my guidance committee, Dr. Sheng Yang He,
Dr. Thomas Sharkey, Dr. Dean DellaPenna and Dr. R. Michael Garavito for their thoughtprovoking discussions, insightful suggestions as well as valuable career advices. They have been
part of my inspiration in many ways. My Ph.D. experience would not be so profitable without their
expertise, encouragement and support. Several other professors have also helped me kindly on
experimental equipment and materials. Dr. Gregg A. Howe generally provided suggestions on my
projects and jasmonates related materials. Dr. Shin-Han Shiu kindly spent time teaching me the
bioinformatic knowledge for my projects. Dr. Giovanni Stefano and Dr. Melinda Frame gave
helpful advice on confocal microscopy.

iv

I also want to thank all the Benning lab members, past and present, for their companionship.
All of them have been very nice to me and provided enormous help and precious suggestions on
my research. I thank Yang Yang, Wei Ma, Chia-Hong Tsai and Xiaobo Li for their helpful
discussions and entertainment; Dr. Barb Sears, Anna Hurlock and Anastasiya Lavell for critically
reading my manuscripts and giving constructive feedbacks among many other things. Special
mention goes to Dr. Rebecca Roston for patiently teaching me the fundamental bench skills during
the first three years of my Ph.D. and Hope Hersh for her assistantship in experiments for the past
four years.
I was initially recruited by the Biochemistry and Molecular Biology Department and joined
the DOE-Plant Research Laboratory program in 2015. I would like to acknowledge the BMB
secretary Jessica Lawrence, PRL staff Eunice Hoeve, Igor Houwat for their help and support
through the years, which made my Ph.D. life smooth. I am also hugely appreciative to Jim Klug
and Cody Keilen for organizing and maintaining the plant chambers and greenhouses.
More broadly, I thank MSU for providing the wonderful academic and life environment.
Besides the world-class scientific facility and faculty, I am also entertained tremendously by the
college sports. The Spartan Will spirit will be certainly carried on wherever I go.
Finally, my gratitude goes to friends and family. My time at MSU was made enjoyable and
enriched in a large part due to the time spent with many friends. I am grateful to my parents and
my wife for may years of love, support and understanding. My wife and I have been having a longterm relationship for over six years. I owe her many thanks to her unconditional love,
understanding and support regardless the decision I made in my career and life.

v

TABLE OF CONTENTS

LIST OF TABLES ......................................................................................................................... ⅸ
LIST OF FIGURES ........................................................................................................................ ⅹ
KEY TO ABBREVIATIONS ...................................................................................................... ⅻi
CHAPTER 1 Introduction............................................................................................................... 1
Galactolipid Synthesis and Remodeling in Response to Stress .................................................. 2
Lipases in Lipid Catabolism ....................................................................................................... 5
Oxylipin Biosynthesis and Signaling ......................................................................................... 7
Overview .................................................................................................................................. 10
APPENDIX ................................................................................................................................... 11
REFERENCES.............................................................................................................................. 15
CHAPTER 2 SENSITIVE TO FREEZING2 Aids in Resilience to Salt and Drought in FreezingSensitive Tomato .......................................................................................................................... 23
ABSTRACT .............................................................................................................................. 24
INTRODUCTION ..................................................................................................................... 25
MATERIALS AND METHODS .............................................................................................. 28
Plant Materials and Growth Conditions............................................................................ 28
Rnai Plasmid Constructions and Tomato Transformation ................................................ 28
Quantitative Real-Time PCR ............................................................................................ 29
Lipid Analysis ................................................................................................................... 29
Protein Production ............................................................................................................ 29
Stress Assays ..................................................................................................................... 30
Chlorophyll Measurements ............................................................................................... 31
Protein Measurement ........................................................................................................ 31
RESULTS.................................................................................................................................. 33
Salt and Drought Treatment Similarly Affect Arabidopsis Wild Type and sfr2-3 Mutant
........................................................................................................................................... 33
Identification of an SFR2 Orthologue of Solanum lycopersicum ..................................... 34
SlSFR2 Has Higher Specific Activity Than AtSFR2 ........................................................ 35
Generation of SlSFR2 RNAi Lines in Tomato ................................................................. 36
SlSFR2 RNAi Transgenic Lines Show Increased Salt and Drought Sensitivity .............. 37
Chlorophyll and Protein Content of SlSFR2 RNAi Transgenic Lines .............................. 38
SFR2 is Activated in Older Leaves in Response to Salt and Drought Treatment ............ 39
DISCUSSION ........................................................................................................................... 40
AtSFR2 and SlSFR2 are Genuine Orthologues ................................................................. 40
Oligogalactolipid Formation by SlSFR2 Protects Against Salt and Drought Stress......... 41
Lipid Remodeling by SFR2 Protects Against Different Abiotic Stresses in Different
Plants ................................................................................................................................. 43
ACKNOWLEDGEMENT ........................................................................................................ 46

vi

APPENDIX ................................................................................................................................... 47
REFERENCES.............................................................................................................................. 67
CHAPTER 3 A Plastid Phosphatidylglycerol Lipase Contributes to the Export of Acyl Groups
from Plastids for Seed Oil Biosynthesis ....................................................................................... 71
ABSTRACT .............................................................................................................................. 72
INTRODUCTION ..................................................................................................................... 73
MATERIALS AND METHODS .............................................................................................. 76
Plant Material and Growth Conditions ............................................................................. 76
Quantitative Reverse-Transcription PCR ......................................................................... 76
Confocal Laser Scanning Microscopy .............................................................................. 77
Protein Extraction and Immunoblot Analysis ................................................................... 77
Recombinant Protein and Antiserum Production ............................................................. 78
Chloroplast Import Assay ................................................................................................. 79
PLIP1 Lipase Assay .......................................................................................................... 79
Lipid Analysis ................................................................................................................... 80
Pulse-Chase Labeling........................................................................................................ 81
Observation of Embryo Morphology ................................................................................ 82
Hormone Extraction and Arabidopside measurement ...................................................... 82
Phylogenetic analysis ........................................................................................................ 83
Accession Numbers .......................................................................................................... 84
RESULTS.................................................................................................................................. 85
PLIP1 is a Chloroplast Thylakoid-Associated Protein ..................................................... 85
PLIP1 is a Phospholipase A1 with a Preference for Unsaturated Acyl Groups ................. 87
18:3/16:1Δ3t-PG is the Preferred Substrate of PLIP1 In Vivo ........................................... 89
Overexpression of PLIP1 Accelerates Recycling of Plastid PG Acyls Groups and Their
Transfer to PC ................................................................................................................... 92
PLIP1 Contributes a Small Fraction of Precursors for TAG Biosynthesis During
Embryogenesis .................................................................................................................. 94
Overexpression of PLIP1 Increases PG Recycling and TAG Biosynthesis During
Embryogenesis .................................................................................................................. 98
DISCUSSION ......................................................................................................................... 100
PLIP1 Prefers PG as Natural Substrate ........................................................................... 100
The Location of PLIP1 Limits the Availability of Likely Substrates In Vivo ................ 101
PLIP1 Contributes to TAG Biosynthesis in Developing Embryos ................................. 102
PLIP1 Enables Channeling of Acyl Groups from Plastid 16:1Δ3t-PG to TAG at the ER 103
PLIP1 Takes Part in Acyl Group Export from Chloroplasts .......................................... 106
ACKNOWLEDGEMENTS .................................................................................................... 108
APPENDIX ................................................................................................................................. 109
REFERENCES............................................................................................................................ 140
CHAPTER 4 Two Abscisic Acid Responsive Plastid Lipase Genes Involved in Jasmonic Acid
Biosynthesis in Arabidopsis thaliana .......................................................................................... 147
ABSTRACT ............................................................................................................................ 148
INTRODUCTION ................................................................................................................... 149
MATERIALS AND METHODS ............................................................................................ 152
vii

Plant Material and Growth Conditions ........................................................................... 152
Phylogenetic Analysis ..................................................................................................... 152
Confocal Laser Scanning Microscopy ............................................................................ 153
Chloroplast Import Assay ............................................................................................... 153
Recombinant Protein Production .................................................................................... 153
PLIP2 Lipase Assay ........................................................................................................ 154
Lipid Analysis ................................................................................................................. 154
Pulse-Chase Labeling...................................................................................................... 155
Hormone Measurements and Oxylipin Profiling ............................................................ 155
Accession Numbers ........................................................................................................ 156
RESULTS................................................................................................................................ 157
PLIP2 and PLIP3 are Putative Paralogs of PLIP1 in Arabidopsis.................................. 157
PLIP2 and 3 are Located in the Chloroplast ................................................................... 157
PLIP2 is a Phospholipase A1 with Ser428 as the Catalytic Site ........................................ 159
PLIP2 Uses MGDG as Primary Substrate, While PLIP3 Uses PG ................................ 160
The Growth and Lipid Phenotypes are Not Correlated in PLIP-OX Plants ................... 162
Oxylipin Production and Signaling in PLIP2- and 3-OX Plants .................................... 164
PLIP2 and PLIP3 Contribute to Oxylipin Production in response to ABA .................... 165
PLIP 1,2,3 Triple Mutants are Hypersensitive to ABA .................................................. 168
DISCUSSION ......................................................................................................................... 169
PLIP2 and PLIP3 Share Biochemical Properties with PLIP1 ........................................ 169
PLIP2 and PLIP3 are Involved in JA Biosynthesis ........................................................ 170
PLIP2 and PLIP3 Link ABA Signaling with JA Biosynthesis ....................................... 171
ACKNOWLEDGEMENTS .................................................................................................... 175
APPENDIX ................................................................................................................................. 176
REFERENCES............................................................................................................................ 200
CHAPTER 5 Future perspectives ............................................................................................... 207
SFR2 in Engineering Stress Tolerant Plants ........................................................................... 208
PLIP1 in Seed Oil Biosynthesis .............................................................................................. 210
PLIP2 and 3 in the Interaction Between ABA and JA ............................................................ 212
REFERENCES............................................................................................................................ 214

viii

LIST OF TABLES

Table 2. 1. Primers used in this study ........................................................................................... 48
Table 3. 1. Primers used in this study ......................................................................................... 110
Table 3. 2. Leaf acyl group contents in different genotypes. ...................................................... 112
Table 3. 3. Sequences used in the PLIP1 and FAD4 phylogenetic analyses. ............................. 113
Table 4. 1. Primer sequences used in this study ......................................................................... 177

ix

LIST OF FIGURES

Figure 1. 1. Proposed model of SFR2-dependent galactolipid remodeling during Freezing........ 12
Figure 1. 2. Ester bond specificity of the phospholipases. ............................................................ 13
Figure 1. 3. Jasmonic acid biosynthesis and signaling in the context of cellular compartments.. 14
Figure 2. 1. SFR2 is not involved in protection against salt or drought stress in Arabidopsis. .... 49
Figure 2. 2. SFR2 is not involved in protection against salt stress in Arabidopsis. ...................... 50
Figure 2. 3. SFR2 is not involved in protection against drought in Arabidopsis .......................... 51
Figure 2. 4. Protein sequence alignment between AtSFR2 and SlSFR2. ...................................... 52
Figure 2. 5. SlSFR2 is a galactolipid : galactolipid galactosyltransferase (GGGT) ..................... 53
Figure 2. 6. Activity of SlSFR2 in vitro. ....................................................................................... 54
Figure 2. 7. Generation the SlSFR2-RNAi lines. .......................................................................... 55
Figure 2. 8. Genotyping of wild type (WT) and SlSFR2-RNAi lines. .......................................... 56
Figure 2. 9. Activation of SlSFR2 in additional SlSFR2-RNAi lines. ......................................... 57
Figure 2. 10. SlSFR2-RNAi lines are sensitive to salt and drought. ............................................. 58
Figure 2. 11. Effects of salt treatment on two-week-old tomato plants. ....................................... 59
Figure 2. 12. Effect of drought treatment on WT and SlSFR2-RNAi plants. ............................... 60
Figure 2. 13. Chlorophyll in the leaves of untreated and stressed three-week-old tomato plants. 61
Figure 2. 14. Chlorophyll in the leaves of untreated and salt-treated two-week-old tomato plants
....................................................................................................................................................... 62
Figure 2. 15. Total protein in leaves from untreated and stress treated three-week-old tomato
plants. ............................................................................................................................................ 63
Figure 2. 16. SFR2 activation in three-week-old tomato plants subjected to salt or drought stress.
....................................................................................................................................................... 64
Figure 2. 17. SFR2 activation in two-week-old tomato plants subjected to salt stress. ................ 65
Figure 2. 18. SlSFR2 transcript levels during salt treatment. ........................................................ 66

x

Figure 3. 1. Subcellular localization of PLIP1 in Arabidopsis. .................................................. 115
Figure 3. 2. Recombinant PLIP1 has lipase activity ................................................................... 117
Figure 3. 3. Alignment of PLIP1 with class 3 lipase protein sequences in the NCBI conserved
domain database (CDD). ............................................................................................................. 118
Figure 3. 4. Time course of PLIP1 lipase activity with phosphatidylcholine (PC) as substrate. 119
Figure 3. 5. Specificity of PLIP1 in vitro activity ....................................................................... 120
Figure 3. 6. Analysis of polar lipids in the leaves of 4-week-old, soil-grown PLIP1-OX and
empty vector (EV) control plants. ............................................................................................... 122
Figure 3. 7. Growth and lipid phenotypes of PLIP1-overexpression plants. .............................. 123
Figure 3. 8. Expression of jasmonic acid-responsive genes. ...................................................... 124
Figure 3. 9. Quantification of plant hormones and Arabidopsides in leaves of the wild type (WT)
and PLIP-OX1. ........................................................................................................................... 125
Figure 3. 10. Phenotypes of PLIP1-OX and coi1 plants............................................................. 126
Figure 3. 11. PLIP1 precursor–product relationships in PLIP1-overexpression plants.............. 127
Figure 3. 12. Effect of sn-2 acyl 16:1Δ3t on PLIP1 function. ....................................................... 129
Figure 3. 13. Lipid analysis of offspring from a cross between fad3-2 and PLIP1-OX1. .......... 130
Figure 3. 14. Effect of PLIP1 on seed oil biosynthesis and germination. ................................... 131
Figure 3. 15. Genotyping, phenotypic and lipid analysis of the plip1 mutants. ......................... 132
Figure 3. 16. Hormone quantification in plip1 and PLIP1-OX1 dry seeds. ............................... 133
Figure 3. 17. Growth phenotypes of mature plip1 mutant and PLIP1-OX plants. ..................... 134
Figure 3. 18. Relative composition of total acyl groups in the wild type (WT), fad3-2, and fad32, PLIP1-OX1 dry seeds. ............................................................................................................ 135
Figure 3. 19. Analysis of fad4-2 and fad4-3 plants..................................................................... 136
Figure 3. 20. Phylogenetic trees of PLIP1 and FAD4 in land plants. ......................................... 137
Figure 3. 21. Phenotypic analysis of PLIP1-OX1 embryos........................................................ 138
Figure 3. 22. Hypothesis for the role of PLIP1 in triacylglycerol biosynthesis. ......................... 139
Figure 4. 1. Time course of PLIP2 lipase activity with phosphatidylcholine (PC) as substrate. 179
xi

Figure 4. 2. Phylogenetic analysis of the top 20 PLIP1 similar sequences in Arabidopsis. ....... 180
Figure 4. 3. Subcellular localization of PLIP2 and PLIP3 in Arabidopsis. ................................ 181
Figure 4. 4. Alignment of PLIP2 (A) and PLIP3 (B) with class 3 lipase protein sequences
available in the NCBI conserved domain database (CDD)......................................................... 182
Figure 4. 5. Production of the recombinant PLIP2 proteins and its lipase activity in vitro. ....... 183
Figure 4. 6. In vitro lipase activity of PLIP2. ............................................................................. 184
Figure 4. 7. The steady-state and dynamic lipid phenotypes of the PLIP2-OX plants. .............. 185
Figure 4. 8. Analysis of polar lipids from the 4-week-old soil-grown PLIP2-OX and empty
vector (EV) control plant leaves ................................................................................................. 186
Figure 4. 9. Analysis of polar lipids from 4-week-old soil-grown PLIP3-OX and empty vector
(EV) control plant leaves ............................................................................................................ 187
Figure 4. 10. In vivo pulse-chase acetate labeling of polar lipids in the empty vector (EV) control
and the PLIP3-OX plants. ........................................................................................................... 188
Figure 4. 11. Morphology of the PLIP-OX plants. ..................................................................... 189
Figure 4. 12. Phytohormone quantification and comparative oxylipin profiling in the WT and the
PLIP2-OX plants. ....................................................................................................................... 190
Figure 4. 13. Phytohormone quantification and oxylipin profiling in the wild-type (WT) and the
PLIP3-OX plants. ....................................................................................................................... 191
Figure 4. 14. Rescue of growth in PLIP2-OX coi1 homozygous plants. ................................... 192
Figure 4. 15. Phenotypes of PLIP3-OX coi1 plants. .................................................................. 193
Figure 4. 16. Transcriptional induction of PLIP2 and PLIP3 as shown in the eFP database ..... 194
Figure 4. 17. Induction of PLIP transcription by ABA. .............................................................. 195
Figure 4. 18. Growth and the lipid phenotype of the plip single and triple mutants. ................. 196
Figure 4. 19. Genotyping of the plip-CRISPR line ..................................................................... 198
Figure 4. 20. The plip-tri plants are hypersensitive to ABA. ..................................................... 199

xii

KEY TO ABBREVIATIONS

ABA – abscisic acid
ABI – ABA INSENSITIVE
ACP – Acyl Carrier Protein
AOC – ALLENE OXIDE CYCLASE
AOS – ALLENE OXIDE SYNTHASE
COI1 – CORONATIN INSENSITIVE 1
DAG – diacylglycerol
DGAT – DAG-acyltransferases
DAD1 – DEFECTIVE IN ANTHER DEHISCENCE1
DGD1 – DGDG Synthase
DGDG – digalactosyldiacylglycerol
ER – endoplasmic reticulum
FA – fatty acid
FAD4 – FATTY ACID DESATURASE4
FAP – FATTY ACID BINDING PROTEINS
FAX1 – FATTY ACID EXPORT1
JA – jasmonic acid
JA-Ile – JA- isoleucine
JAR1 – JASMONATE RESISTENT 1
JAZ – JA ZIM-domain transcriptional repressors
LHCb1 – Light Harvesting Complex b1
LOX – lipoxygenase
xiii

MGD1 – MGDG Synthase
MGDG – monoalactosyldiacylglycerol
OPDA – 12-oxo phytodienoic acid
PA – phosphatidic acid
PC – phosphatidylcholine
PDAT – phospholipid: DAG acyltransferases
PE – phosphatidylethanolamine
PG – phosphatidylglycerol
PGD1 – Plastid Galactolipid Degradation1
PI – phosphatidylinositol
PLA1 – phospholipase A1
PLA2 – phospholipase A2
PLC – phospholipase C
PLD – phospholipase D
PLIP1 – PLASTID LIPASE1
SDP1 – SUGAR DEPENDENT1
SFR2 – SENSITIVE TO FREEZING 2
SQDG – sulfoquinovosyldiacylglycerol
TAG – triacylglycerol
TGD1 – TRIGALACTOSYLDIACYLGLYCEROL1
TGDG – trigalactosyldiacylglycerol
TLC – thin-layer chromatography

xiv

CHAPTER 1
Introduction

1

In plants and algae, chloroplasts are organelles that carry out a number of crucial functions,
including photosynthesis, fatty acid biosynthesis and plant immune responses. Chloroplast
membranes with a unique lipid composition are essential for the proper functions of chloroplasts.
In Arabidopsis, chloroplasts lipids are synthesized by two pathways (Benning, 2009; Hurlock et
al., 2014). De novo synthesized fatty acids (FAs) can either directly enter the plastid pathway or
are exported to the endoplasmic reticulum (ER) and assembled into lipids there. Lipid precursors
synthesized in the ER can also return to the plastid and contribute to chloroplast lipid biosynthesis
(Wang and Benning, 2012; Hurlock et al., 2014). To date, many of the genes and the respective
enzymes needed for chloroplast lipid biosynthesis have been characterized in Arabidopsis. Certain
chloroplast lipid biosynthetic genes are induced, or the corresponding enzymes are activated only
in response to specific stresses. They function to remodel the chloroplast membrane composition
affecting its biophysical properties to cope with unfavorable conditions.
Besides lipid biosynthesis, maintenance of the chloroplast membranes also requires a finely
tuned lipid catabolic machinery. A number of lipid degrading enzymes are predicted to be localized
in the chloroplast with unknown physiological functions. It has been proposed that lipases are
crucial for various aspects of chloroplast physiology, e.g., membrane turnover, lipid recycling and
signaling (Wang, 2002, 2004; Scherer et al., 2010; Richmond and Smith, 2011).

Galactolipid Synthesis and Remodeling in Response to Stress
The galactolipids monogalactosyldiacylglycerol (MGDG) and digalactosyldiacylglycerol
(DGDG) are the most abundant lipids in plant leaf tissues and they comprise up to ~50% and ~25%
of total chloroplast lipids, respectively (Block et al., 1983). During normal growth and
development, MGDG and DGDG are exclusively restricted to the chloroplasts. However, DGDG

2

was also found in the extraplastidic membranes following phosphate deprivation (Härtel et al.,
2000).
Under normal conditions, MGDG synthesis is catalyzed by MGDG synthase (MGD1) with
UDP-Gal and diacylglycerol (DAG) as substrates (Awai et al., 2001), while DGDG is synthesized
by DGDG synthase (DGD1) using UDP-Gal as the head group donor to further galactosylate
MGDG (Dörmann et al., 1995; Dörmann et al., 1999). Phosphate is essential for the synthesis of
biological macromolecules including DNA and phospholipids. Highly regulated transcriptional
and post-transcriptional mechanisms are evolved in plants in response to phosphate limiting
conditions (Rubio et al., 2001; Aung et al., 2006; Chiou et al., 2006). Phosphate liberation from
phospholipids and replacement of phospholipids by galactolipids is one of the mechanisms that
enables plants to cope with phosphate deficiency (Dörmann and Benning, 2002). Two genes
encoding paralogs of MGD1 (MGD2 and MGD3) and a single gene encoding a DGD1 paralog
(DGD2) are transcriptionally induced during phosphate deprivation (Awai et al., 2001; Kelly and
Dörmann, 2002). Presence of the MGD1 and DGD1 paralogs enables the maintenance of
galactolipid levels under phosphate deprivation and allows the export of DGDG to the
extraplastidic membranes, even though the export mechanisms remain enigmatic thus far
(Moellering and Benning, 2011).
Another pathway for galactolipid biosynthesis and remodeling is the SFR2 catalyzed
oligogalactolipid biosynthesis during freezing. The sfr2 mutant was discovered in 1996 by Gareth
Warren and his colleagues by screening of freezing sensitive mutants in Arabidopsis thaliana. As
one of the seven identified sensitive to freezing mutants, sfr2 distinguishes itself by displaying a
strong and severe phenotype on the whole plant but maintaining an intact plasma membrane under
freezing conditions (Warren et al., 1996). Later, transmission electron microscopy showed that the

3

chloroplast membranes rather than the plasma membrane of the sfr2 mutant are disrupted. The
chloroplast membranes are fused with other organelle membranes after freezing treatment, which
leads to cell disruption upon rewarming (Fourrier et al., 2008). By mapping, SFR2 was identified
as At3g06510, which encodes a beta glycosidase classified as a family I glycosyl hydrolase
belonging to a distinct lineage (Thorlby et al., 2004; Xu et al., 2004). Despite the conserved
catalytic site suggesting it to be a glycosyl hydrolases, SFR2 possesses no hydrolase activity, but
functions as a transferase (Roston et al., 2014). Structural modeling and site-directed mutagenesis
of SFR2 revealed three unique regions that might evolutionally separate SFR2 from hydrolases: a
loop insertion, the C-terminal peptide, and a hydrophobic patch adjacent to the catalytic site
(Roston et al., 2014). The transcript level of SFR2 and the level of the encoded protein remain
constant under all conditions examined including freezing. Instead, SFR2 activity is directly turned
on by decreased cytosolic pH and increased cytosolic Mg2+ concentration changes concomitant to
the freezing (Roston et al., 2014; Barnes et al., 2016). As shown in Figure 1.1, when activated by
freezing, this outer-envelope membrane bound enzyme processively transfers the galactosyl
residues from MGDG to a second galactolipid acceptor, which gives rise to the formation of
oligogalactolipid and DAG (Moellering et al., 2010; Roston et al., 2014). Depending on the
galactosyl acceptor, the galactolipid product can be DGDG or higher order oligogalactolipids like
trigalactosyldiacylglycerol (TGDG) or tetragalactosyldiacylglycerol (TeDG). However, distinct
from the DGDG produced by the UDP-Gal-dependent DGD1/DGD2 lipid galactosyltransferases,
which is in βα conformation with respect to the glycosidic linkage (Dörmann et al., 1999; Kelly
and Dörmann, 2002), the DGDG produced by SFR2 is in all-β conformation (ββDGDG) (Xu et
al., 2003). The other product of SFR2 activity, DAG is converted to triacylglycerol (TAG) and
sequestered into lipid droplets. Based on these observations, it is proposed that conversion of

4

MGDG into oligogalactolipids and DAG replaces non-bilayer forming lipids with bilayer
stabilizing ones. This process also adjusts chloroplast envelope membranes as the chloroplast
shrinks in response to cellular dehydration accompanying ice formation in the apoplast during
freezing. Presence of the oligogalactosyl residues as the head group also increases the hydration
of the chloroplast envelope membranes. As a result, the chloroplast membranes are stabilized and
do not fuse with other membranes during freezing conditions.
SFR2-like genes are present in the plant kingdom over a wide phylogenic distribution based
on the observation of sequenced genomes and expressed tag sequences (EST), including species
both resistant and sensitive to freezing (Fourrier et al., 2008). Introduction of SFR2 cDNAs from
freezing intolerant plant species, such as soy bean (Glycine max) and rice (Oryza sativa) can also
rescue the freezing phenotype in the Arabidopsis sfr2 mutant. Therefore, it is likely that SFR2
activity has other physiological roles in plants besides avoiding freezing damage. Freezingtriggered dehydration and cell shrinkage is shared with some other stresses, primarily drought and
high salinity. In fact, oligogalactolipid accumulation was observed in desiccated leaves of
Arabidopsis under room temperature and recently, lipid analysis of the resurrection plant
Craterostigma plantagineum revealed that MGDG is converted to DGDG and DAG during the
desiccation stage (Gasulla et al., 2013). Even though the responsible enzyme was not confirmed
to be SFR2, it implies that MGDG conversion to oligogalactolipids might be a common strategy
for plants to deal with dehydration.

Lipases in Lipid Catabolism
Lipases are enzymes that belong to a subclass of the esterases and catalyze the hydrolysis
of lipids. They are the key enzymes for lipid turnover and play important roles in a wide range of
5

biological processes in most, if not all, living organisms (Wang, 2004; Scherer et al., 2010;
Richmond and Smith, 2011). Due to their distinct catalytic properties even in organic solvents,
lipases also constitute an important group of biocatalysts for biotechnological applications (Jaeger
and Eggert, 2002). Based on their substrate preference, lipases from plants are broadly classified
into TAG lipases, phospholipases, glycolipases, sulfolipases and monoacylglycerol lipases. They
are more commonly classified based on the preference on the glyceryl positions at which the
hydrolysis reaction happens. For example, phospholipases can be classified into four major types
based on the cleavage sites, phospholipase D (PLD), phospholipase C (PLC), phospholipase A1
(PLA1), and phospholipase A2 (PLA2) (Figure 1.2). PLD releases the polar head group and
produces phosphatidic acid while PLC cleaves the phosphodiester bond at the glyceryl sn-3
position and produces the phosphorylated head group and DAG. PLAs release acyl groups from
the glyceryl moiety at the sn-1 or sn-2 positions (Wang et al., 2012).
The Arabidopsis genome encodes about 300 proteins that are annotated as lipases, but most
of them have not been biochemically verified and have unknown physiological functions (LiBeisson et al., 2013; Troncoso-Ponce et al., 2013; Kelly and Feussner, 2016). A group of well
characterized lipases named patatin-related PLAs are encoded by 10 genes in Arabidopsis. These
lipases are structurally related to patatins, which are potato tuber proteins with acyl-hydrolyzing
activities (Scherer et al., 2010). Members of this groups use diverse polar lipids as substrates and
are involved in an array of physiological functions, including responses to auxin, pathogens and
abiotic stresses (Scherer et al., 2010). One of the members named SUGAR DEPENDENT1 (SDP1)
is a lipase that is capable of releasing acyl groups from both TAG and DAG. During seedling
development, SDP1 is required for degrading seed storage oil to provide carbon skeletons and
energy for the post-germinative growth (Eastmond, 2006). Another recently identified and

6

characterized lipase is Plastid Galactolipid Degradation1 (PGD1) in Chlamydomonas reinhardtii
(Li et al., 2012). Expression of PGD1 is induced under nitrogen deprivation and the protein
functions as an MGDG lipase. It almost exclusively hydrolyzes newly synthesized acyl groups that
have not yet been desaturated from MGDG, and the acyl chains are later exported to the ER and
incorporated into TAG. The PGD1 deficient mutant exhibits reduced TAG accumulation,
decreased cell viability and altered MGDG and TAG fatty acid (FA) composition following
nitrogen depletion.
A number of lipases are predicted to be located in the chloroplast, but most of them have
not been characterized so far and have unknown physiological functions. Conceivably, these
lipases are required for the membrane maintenance, turnover and oxylipin biosynthesis in the
chloroplasts. One of the characterized plastid lipases is DEFECTIVE IN ANTHER
DEHISCENCE1 (DAD1) which is a PLA1 that is primarily present in the reproductive tissues,
where it initiates JA biosynthesis critical for pollen maturation, anther dehiscence and flower
opening in Arabidopsis (Ishiguro et al., 2001). In leaf tissues, DONGLE encodes a DAD1homologous lipase contributing to wounding-induced JA production (Hyun et al., 2008).

Oxylipin Biosynthesis and Signaling
Lipids are frequently subject to oxidation through enzymatic and non-enzymatic processes
(Mosblech et al., 2009). Oxylipins comprise a diverse class of lipophilic metabolites derived from
the oxidation of polyunsaturated FAs, including FA hydroperoxides, hydroxy-, oxo-, or keto-FAs
(Grechkin, 1998). Oxylipins are constitutively present at low levels in seed plants and their levels
increase in response to a variety of responses (Mueller, 2004). The enzymatic process is primarily
mediated by lipoxygenases (LOX) and some of their oxylipin products, e.g., jasmonate, are

7

essential for plant defense and reproduction (Porta and Rocha-Sosa, 2002). In parallel, oxidative
stress and the formation of reactive oxygen species can also lead to chemically mediated lipid
oxidation independent of lipoxygenases. Even though non-enzymatic lipid oxidation is generally
regarded as deleterious, lipid peroxidation derived reactive electrophile species (RES) have
emerged as signaling molecules that are capable of reprogramming plant transcriptomes through
class II TGA transcription factors (Farmer and Mueller, 2013).
The enzymatic lipid oxidation process is initiated by the LOXs to produce lipid
hydroperoxide using polyunsaturated FAs as substrates. The most common substrates in plants are
linolenic (18:3; carbon number : double bonds number) and linoleic acids (18:2) (Grechkin, 1998),
expects the debate that FAs esterified to the glycerol backbone could also be used as substrates
(Feussner et al., 2001; Stelmach et al., 2001). Their products, 9- and 13-hydroperoxy FAs, can
then enter an array of pathways leading to various products (Mosblech et al., 2009). One of the
best characterized pathway is the ALLENE OXIDE SYNTHASE pathway which leads to the
synthesis of oxylipins, including 12-oxo phytodienoic acid (OPDA), jasmonic acid (JA) and JA
derivatives, e.g., the bioactive JA-isoleucine (JA-Ile) (Turner et al., 2002; Wasternack and Hause,
2013).
JA production and signaling are primarily induced by wounding damage caused by
herbivorous insects or necrotrophic microbes (Creelman et al., 1992; Howe and Schilmiller, 2002;
Trusov et al., 2006). ~80% of wounding and insect attach related genes are under the control of
JA in Arabidopsis leaves (Reymond et al., 2004). During plant growth and development, JA is
essential for stamen development, sizes of petioles and petals, as well as lignin biosynthesis (Feys
et al., 1994; McConn and Browse, 1996; Ellis et al., 2002; Buseman et al., 2006; Denness et al.,
2011). Upon wounding damage or herbivory attack, JA production and activation of the JA
8

signaling represses plant growth by changing the plant transcriptome and interfering with the
gibberellin signaling cascade (Yang et al., 2012). This was generally explained by the concept of
growth-defense tradeoff (Huot et al., 2014; Zust et al., 2015). Recently, JA signaling mediated
growth-defense tradeoff was genetically uncoupled in the jazQ phyB mutant which is deficient in
five JA ZIM-domain transcriptional repressors (JAZs) and the photoreceptor phyB (Campos et al.,
2016). In addition, synergetic or antagonistic interactions between JA and other phytohormones
have also emerged. Classically, salicylic acid signaling antagonizes JA signaling for resistance
against biotrophic and necrotrophic pathogens, respectively, while ethylene works synergistically
with JA in activating the defense against the necrotrophic pathogens (Glazebrook, 2005). JA
biosynthesis is also stimulated by applied ABA or during ABA-inducing abiotic stress conditions
(Creelman and Mullet, 1995; Adie et al., 2007; Avramova, 2017), but the underlying mechanisms
are not well understood. During abiotic stress, rising ABA levels induce JA synthesis, which
synergistically optimizes the plant’s abiotic defense by altering the plant transcriptome, facilitating
stomata closure, inhibiting cell division, as well as epigenetically modifying defense genes during
dehydration responses (Murata et al., 2015; Riemann et al., 2015; Liu et al., 2016; Valenzuela et
al., 2016). A basic-helix-loop-helix transcription factor MYC2 plays roles in both JA perception
and ABA signaling, and has been proposed as one of the factors linking ABA and JA signaling
(Kazan and Manners, 2013).
As shown in Figure 1.3, JA biosynthesis starts in the plastids. Released poly-unsaturated
FAs are oxidized by LOXs and ALLENE OXIDE SYNTHASE (AOS). The unstable allene oxides
are subsequently cyclized by ALLENE OXIDE CYCLASE (AOC), leading to the production of
12-oxo phytodienoic acid (OPDA) and dinor-OPDA (Wasternack and Hause, 2013; Heitz et al.,
2016). OPDA and dinor-OPDA are exported from the chloroplasts through unknown mechanisms,

9

and subsequently enter peroxisomes where they are subject to reduction and β-oxidation before
the completion of JA synthesis (Wasternack and Hause, 2013; Heitz et al., 2016). Among many of
the possible subsequent JA modifications, the conjugate JA-Ile produced by JASMONATE
RESISTENT 1 (JAR1) is the active hormone molecule for JA signaling (Staswick and Tiryaki,
2004; Fonseca et al., 2009).
JA perception and signaling happens in the nucleus. Similar to other phytohormones, JA
perception and signaling also fit the ‘relief of repression model’ (Santner and Estelle, 2009). In the
absence of JA-Ile, key mediators including MYC transcription factors are repressed by JAZs in
the resting state (Zhang et al., 2015). In the presence of JA-Ile, the F-box subunit of an SCF-type
ubiquitin E3 ligase, CORONATIN INSENSITIVE 1 (COI1), forms the JA receptor complex
together with JA-Ile and JAZs (Katsir et al., 2008). This subsequently leads to the ubiquitination
mediated degradation of JAZs, liberating MYCs from the transcriptional repression and initiating
the downstream signaling (Thines et al., 2007; Zhang et al., 2015).

Overview
Many aspects of the function and mechanisms of chloroplast lipid remodeling remain to be
determined. The results in Chapter 2 expands the physiological functions of SFR2 to freezing
sensitive plants. Chapter 3 explores chloroplast lipid catabolism and describes a novel lipase,
PLASTID LIPASE 1 (PLIP1), that connects lipid catabolism with seed oil biosynthesis. In Chapter
4, the focus is on two paralogs of PLIP1, PLIP2 and PLIP3, that release FAs which are
subsequently converted to oxylipins. A new hypothesis is provided suggesting that these lipases
link ABA signaling with oxylipin biosynthesis. In Chapter 5, future perspectives are discussed
and possible experiments are proposed for each chapter.

10

APPENDIX

11

Figure 1. 1. Proposed model of SFR2-dependent galactolipid remodeling during Freezing.
SFR2 is activated in the wild-type Arabidopsis plant during freezing. It removes the MGDG head
group and transfers it to another MGDG (or DGDG) for the production of DGDG (or TGDG). The
other product, DAG, is converted to TAG. This transgalactosylation activity stabilizes the
membrane bilayer. Acyl chains and glycerol bonds are represented as green sticks, while galactosyl
head groups are represented as yellow circles. DAG, diacylglycerol; DGDG,
digalactosyldiacylglcyerol; MGDG, monogalactosyldiacylglycerol; TAG, triacylglycerol; TGDG,
trigalactosyldiacylglycerol. oEM, outer envelope membrane; iEM, inner envelop membrane.

12

Figure 1. 2. Ester bond specificity of the phospholipases.
Phospholipase A1 (PLA1) and phospholipase A2 (PLA2) hydrolyze the ester bond from the sn-1
(1) and sn-2 (2) glyceryl positions, respectively. Phospholipase C (PLC) and phospholipase D
(PLD) catalyze the hydrolysis of the phosphodiester bond adjacent or distal to the sn-3 glyceryl
bond, respectively. X represents the derivatives for the phosphoryl-head group, such as choline,
glycerol, ethanolamine, etc.

13

Figure 1. 3. Jasmonic acid biosynthesis and signaling in the context of cellular compartments.
Jasmonic acid (JA) biosynthesis starts in the plastids, where the OPDA and dinor-OPDA are
synthesized and exported from the plastids to peroxisomes before the completion of JA synthesis.
JA is converted to the bioactive form, JA-Ile, in the cytosol. JA-Ile triggers the degradation of the
JAZ proteins together with COI1, and the released TFs initiate gene expression in the nucleus.
Enzymes are shown with black text in colored circles, while the metabolites are shown in blue
text. AOC, ALLENE OXIDE CYCLASE; AOS, ALLENE OXIDE SYNTHASE; JAZ, Jasmonate
ZIM domain (JAZ) transcriptional repressors; JAR, JASMONATE RESISTENT1; LOX,
lipoxygenases; OPDA, 12-oxo Phytodienoic Acid OPR3; OPR3, OXOPHYTODIENOIC ACID
REDUCTAXSE3;
SCF-COI1,
SKP1-CUL1-F-box
(SCF)CORONATINE
INSENSITIVE1(COI1); TFs, transcription factors.

14

REFERENCES

15

REFERENCES

Adie, B.A., Perez-Perez, J., Perez-Perez, M.M., Godoy, M., Sanchez-Serrano, J.J., Schmelz,
E.A., and Solano, R. (2007). ABA is an essential signal for plant resistance to pathogens
affecting JA biosynthesis and the activation of defenses in Arabidopsis. Plant Cell 19,
1665-1681.
Aung, K., Lin, S.I., Wu, C.C., Huang, Y.T., Su, C.L., and Chiou, T.J. (2006). pho2, a
phosphate overaccumulator, is caused by a nonsense mutation in a microRNA399 target
gene. Plant Physiol 141, 1000-1011.
Avramova, Z. (2017). The jasmonic acid-signalling and abscisic acid-signalling pathways cross
talk during one, but not repeated, dehydration stress: a non-specific 'panicky' or a
meaningful response? Plant Cell Environ 40, 1704-1710.
Awai, K., Marechal, E., Block, M.A., Brun, D., Masuda, T., Shimada, H., Takamiya, K.,
Ohta, H., and Joyard, J. (2001). Two types of MGDG synthase genes, found widely in
both 16:3 and 18:3 plants, differentially mediate galactolipid syntheses in photosynthetic
and nonphotosynthetic tissues in Arabidopsis thaliana. Proc Natl Acad Sci U S A 98,
10960-10965.
Barnes, A.C., Benning, C., and Roston, R. (2016). Chloroplast membrane remodeling during
freezing stress is accompanied by cytoplasmic acidification activating SENSITIVE TO
FREEZING 2. Plant Physiol 171, 2140-2149.
Benning, C. (2009). Mechanisms of lipid transport involved in organelle biogenesis in plant
cells. Annu Rev Cell Dev Biol 25, 71-91.
Block, M.A., Dorne, A.J., Joyard, J., and Douce, R. (1983). Preparation and characterization
of membrane fractions enriched in outer and inner envelope membranes from spinach
chloroplasts. II. Biochemical characterization. J Biol Chem 258, 13281-13286.
Buseman, C.M., Tamura, P., Sparks, A.A., Baughman, E.J., Maatta, S., Zhao, J., Roth,
M.R., Esch, S.W., Shah, J., Williams, T.D., and Welti, R. (2006). Wounding
stimulates the accumulation of glycerolipids containing oxophytodienoic acid and dinoroxophytodienoic acid in Arabidopsis leaves. Plant Physiol 142, 28-39.
Campos, M.L., Yoshida, Y., Major, I.T., de Oliveira Ferreira, D., Weraduwage, S.M.,
Froehlich, J.E., Johnson, B.F., Kramer, D.M., Jander, G., Sharkey, T.D., and Howe,
G.A. (2016). Rewiring of jasmonate and phytochrome B signalling uncouples plant
growth-defense tradeoffs. Nat Commun 7, 12570.
Chiou, T.J., Aung, K., Lin, S.I., Wu, C.C., Chiang, S.F., and Su, C.L. (2006). Regulation of
phosphate homeostasis by MicroRNA in Arabidopsis. Plant Cell 18, 412-421.
16

Creelman, R.A., and Mullet, J.E. (1995). Jasmonic acid distribution and action in plants:
regulation during development and response to biotic and abiotic stress. Proc Natl Acad
Sci U S A 92, 4114-4119.
Creelman, R.A., Tierney, M.L., and Mullet, J.E. (1992). Jasmonic acid/methyl jasmonate
accumulate in wounded soybean hypocotyls and modulate wound gene expression. Proc
Natl Acad Sci U S A 89, 4938-4941.
Denness, L., McKenna, J.F., Segonzac, C., Wormit, A., Madhou, P., Bennett, M.,
Mansfield, J., Zipfel, C., and Hamann, T. (2011). Cell wall damage-induced lignin
biosynthesis is regulated by a reactive oxygen species- and jasmonic acid-dependent
process in Arabidopsis. Plant Physiol 156, 1364-1374.
Dörmann, P., and Benning, C. (2002). Galactolipids rule in seed plants. Trends Plant Sci 7,
112-118.
Dörmann, P., Balbo, I., and Benning, C. (1999). Arabidopsis galactolipid biosynthesis and
lipid trafficking mediated by DGD1. Science 284, 2181-2184.
Dörmann, P., Hoffmann-Benning, S., Balbo, I., and Benning, C. (1995). Isolation and
characterization of an Arabidopsis mutant deficient in the thylakoid lipid digalactosyl
diacylglycerol. Plant Cell 7, 1801-1810.
Eastmond, P.J. (2006). SUGAR-DEPENDENT1 encodes a patatin domain triacylglycerol
lipase that initiates storage oil breakdown in germinating Arabidopsis seeds. Plant Cell
18, 665-675.
Ellis, C., Karafyllidis, I., Wasternack, C., and Turner, J.G. (2002). The Arabidopsis mutant
cev1 links cell wall signaling to jasmonate and ethylene responses. Plant Cell 14, 15571566.
Farmer, E.E., and Mueller, M.J. (2013). ROS-mediated lipid peroxidation and RES-activated
signaling. Annu Rev Plant Biol 64, 429-450.
Feussner, I., Kuhn, H., and Wasternack, C. (2001). Lipoxygenase-dependent degradation of
storage lipids. Trends Plant Sci 6, 268-273.
Feys, B., Benedetti, C.E., Penfold, C.N., and Turner, J.G. (1994). Arabidopsis mutants
selected for resistance to the phytotoxin coronatine are male sterile, insensitive to methyl
jasmonate, and resistant to a bacterial pathogen. Plant Cell 6, 751-759.
Fonseca, S., Chini, A., Hamberg, M., Adie, B., Porzel, A., Kramell, R., Miersch, O.,
Wasternack, C., and Solano, R. (2009). (+)-7-iso-Jasmonoyl-L-isoleucine is the
endogenous bioactive jasmonate. Nat Chem Biol 5, 344-350.

17

Fourrier, N., Bedard, J., Lopez-Juez, E., Barbrook, A., Bowyer, J., Jarvis, P., Warren, G.,
and Thorlby, G. (2008). A role for SENSITIVE TO FREEZING2 in protecting
chloroplasts against freeze-induced damage in Arabidopsis. Plant J 55, 734-745.
Gasulla, F., Vom Dorp, K., Dombrink, I., Zahringer, U., Gisch, N., Dörmann, P., and
Bartels, D. (2013). The role of lipid metabolism in the acquisition of desiccation
tolerance in Craterostigma plantagineum: a comparative approach. Plant J 75, 726-741.
Glazebrook, J. (2005). Contrasting mechanisms of defense against biotrophic and necrotrophic
pathogens. Annu Rev Phytopathol 43, 205-227.
Grechkin, A. (1998). Recent developments in biochemistry of the plant lipoxygenase pathway.
Prog Lipid Res 37, 317-352.
Härtel, H., Dörmann, P., and Benning, C. (2000). DGD1-independent biosynthesis of
extraplastidic galactolipids after phosphate deprivation in Arabidopsis. Proc Natl Acad
Sci U S A 97, 10649-10654.
Heitz, T., Smirnova, E., Widemann, E., Aubert, Y., Pinot, F., and Menard, R. (2016). The
rise and fall of jasmonate biological activities. Subcell Biochem 86, 405-426.
Howe, G.A., and Schilmiller, A.L. (2002). Oxylipin metabolism in response to stress. Curr
Opin Plant Biol 5, 230-236.
Huot, B., Yao, J., Montgomery, B.L., and He, S.Y. (2014). Growth-defense tradeoffs in plants:
a balancing act to optimize fitness. Molecular plant 7, 1267-1287.
Hurlock, A.K., Roston, R.L., Wang, K., and Benning, C. (2014). Lipid trafficking in plant
cells. Traffic 15, 915-932.
Hyun, Y., Choi, S., Hwang, H.J., Yu, J., Nam, S.J., Ko, J., Park, J.Y., Seo, Y.S., Kim, E.Y.,
Ryu, S.B., Kim, W.T., Lee, Y.H., Kang, H., and Lee, I. (2008). Cooperation and
functional diversification of two closely related galactolipase genes for jasmonate
biosynthesis. Dev Cell 14, 183-192.
Ishiguro, S., Kawai-Oda, A., Ueda, J., Nishida, I., and Okada, K. (2001). The DEFECTIVE
IN ANTHER DEHISCIENCE gene encodes a novel phospholipase A1 catalyzing the
initial step of jasmonic acid biosynthesis, which synchronizes pollen maturation, anther
dehiscence, and flower opening in Arabidopsis. Plant Cell 13, 2191-2209.
Jaeger, K.E., and Eggert, T. (2002). Lipases for biotechnology. Curr Opin Biotechnol 13, 390397.
Katsir, L., Schilmiller, A.L., Staswick, P.E., He, S.Y., and Howe, G.A. (2008). COI1 is a
critical component of a receptor for jasmonate and the bacterial virulence factor
coronatine. Proc Natl Acad Sci U S A 105, 7100-7105.
18

Kazan, K., and Manners, J.M. (2013). MYC2: the master in action. Molecular plant 6, 686703.
Kelly, A.A., and Dörmann, P. (2002). DGD2, an arabidopsis gene encoding a UDP-galactosedependent digalactosyldiacylglycerol synthase is expressed during growth under
phosphate-limiting conditions. J Biol Chem 277, 1166-1173.
Kelly, A.A., and Feussner, I. (2016). Oil is on the agenda: Lipid turnover in higher plants.
Biochim Biophys Acta 1861, 1253-1268.
Li-Beisson, Y., Shorrosh, B., Beisson, F., Andersson, M.X., Arondel, V., Bates, P.D., Baud,
S., Bird, D., Debono, A., Durrett, T.P., Franke, R.B., Graham, I.A., Katayama, K.,
Kelly, A.A., Larson, T., Markham, J.E., Miquel, M., Molina, I., Nishida, I.,
Rowland, O., Samuels, L., Schmid, K.M., Wada, H., Welti, R., Xu, C., Zallot, R.,
and Ohlrogge, J. (2013). Acyl-lipid metabolism. Arabidopsis Book 11, e0161.
Li, X., Moellering, E.R., Liu, B., Johnny, C., Fedewa, M., Sears, B.B., Kuo, M.H., and
Benning, C. (2012). A Galactoglycerolipid Lipase Is Required for Triacylglycerol
Accumulation and Survival Following Nitrogen Deprivation in Chlamydomonas
reinhardtii. Plant Cell 11, 4670-4686.
Liu, N., Staswick, P.E., and Avramova, Z. (2016). Memory responses of jasmonic acidassociated Arabidopsis genes to a repeated dehydration stress. Plant Cell Environ 39,
2515-2529.
McConn, M., and Browse, J. (1996). The critical requirement for linolenic acid is pollen
development, not photosynthesis, in an arabidopsis mutant. Plant Cell 8, 403-416.
Moellering, E.R., and Benning, C. (2011). Galactoglycerolipid metabolism under stress: a time
for remodeling. Trends Plant Sci 16, 98-107.
Moellering, E.R., Muthan, B., and Benning, C. (2010). Freezing tolerance in plants requires
lipid remodeling at the outer chloroplast membrane. Science 330, 226-228.
Mosblech, A., Feussner, I., and Heilmann, I. (2009). Oxylipins: structurally diverse
metabolites from fatty acid oxidation. Plant Physiol Biochem 47, 511-517.
Mueller, M.J. (2004). Archetype signals in plants: the phytoprostanes. Curr Opin Plant Biol 7,
441-448.
Murata, Y., Mori, I.C., and Munemasa, S. (2015). Diverse stomatal signaling and the signal
integration mechanism. Annu Rev Plant Biol 66, 369-392.
Porta, H., and Rocha-Sosa, M. (2002). Plant lipoxygenases. physiological and molecular
features. Plant Physiol 130, 15-21.

19

Reymond, P., Bodenhausen, N., Van Poecke, R.M., Krishnamurthy, V., Dicke, M., and
Farmer, E.E. (2004). A conserved transcript pattern in response to a specialist and a
generalist herbivore. Plant Cell 16, 3132-3147.
Richmond, G.S., and Smith, T.K. (2011). Phospholipases A1. Int J Mol Sci 12, 588-612.
Riemann, M., Dhakarey, R., Hazman, M., Miro, B., Kohli, A., and Nick, P. (2015).
Exploring jasmonates in the hormonal network of drought and salinity responses.
Front Plant Sci 6, 1077.
Roston, R.L., Wang, K., Kuhn, L.A., and Benning, C. (2014). Structural determinants
allowing transferase activity in SENSITIVE TO FREEZING 2, classified as a family I
glycosyl hydrolase. J Biol Chem 289, 26089-26106.
Rubio, V., Linhares, F., Solano, R., Martin, A.C., Iglesias, J., Leyva, A., and Paz-Ares, J.
(2001). A conserved MYB transcription factor involved in phosphate starvation signaling
both in vascular plants and in unicellular algae. Genes Dev 15, 2122-2133.
Santner, A., and Estelle, M. (2009). Recent advances and emerging trends in plant hormone
signalling. Nature 459, 1071-1078.
Scherer, G.F., Ryu, S.B., Wang, X., Matos, A.R., and Heitz, T. (2010). Patatin-related
phospholipase A: nomenclature, subfamilies and functions in plants. Trends Plant Sci 15,
693-700.
Staswick, P.E., and Tiryaki, I. (2004). The oxylipin signal jasmonic acid is activated by an
enzyme that conjugates it to isoleucine in Arabidopsis. Plant Cell 16, 2117-2127.
Stelmach, B.A., Muller, A., Hennig, P., Gebhardt, S., Schubert-Zsilavecz, M., and Weiler,
E.W. (2001). A novel class of oxylipins, sn1-O-(12-oxophytodienoyl)-sn2-O(hexadecatrienoyl)-monogalactosyl Diglyceride, from Arabidopsis thaliana. J Biol Chem
276, 12832-12838.
Thines, B., Katsir, L., Melotto, M., Niu, Y., Mandaokar, A., Liu, G., Nomura, K., He, S.Y.,
Howe, G.A., and Browse, J. (2007). JAZ repressor proteins are targets of the
SCF(COI1) complex during jasmonate signalling. Nature 448, 661-665.
Thorlby, G., Fourrier, N., and Warren, G. (2004). The SENSITIVE TO FREEZING2 gene ,
required for freezing tolerance in Arabidopsis thaliana , encodes a β-glucosidase 16,
2192-2203.
Troncoso-Ponce, M.A., Cao, X., Yang, Z., and Ohlrogge, J.B. (2013). Lipid turnover during
senescence. Plant Sci 205-206, 13-19.

20

Trusov, Y., Rookes, J.E., Chakravorty, D., Armour, D., Schenk, P.M., and Botella, J.R.
(2006). Heterotrimeric G proteins facilitate Arabidopsis resistance to necrotrophic
pathogens and are involved in jasmonate signaling. Plant Physiol 140, 210-220.
Turner, J.G., Ellis, C., and Devoto, A. (2002). The jasmonate signal pathway. Plant Cell 14
Suppl, S153-164.
Valenzuela, C.E., Acevedo-Acevedo, O., Miranda, G.S., Vergara-Barros, P., Holuigue, L.,
Figueroa, C.R., and Figueroa, P.M. (2016). Salt stress response triggers activation of
the jasmonate signaling pathway leading to inhibition of cell elongation in Arabidopsis
primary root. J Exp Bot 67, 4209-4220.
Wang, G., Ryu, S., and Wang, X. (2012). Plant phospholipases: an overview. Methods Mol
Biol 861, 123-137.
Wang, X. (2002). Phospholipase D in hormonal and stress signaling. Curr Opin Plant Biol 5,
408-414.
Wang, X. (2004). Lipid signaling. Curr Opin Plant Biol 7, 329-336.
Wang, Z., and Benning, C. (2012). Chloroplast lipid synthesis and lipid trafficking through ERplastid membrane contact sites. Biochem Soc Trans 40, 457-463.
Warren, G., McKown, R., Marin, a.L., and Teutonico, R. (1996). Isolation of mutations
affecting the development of freezing tolerance in Arabidopsis thaliana (L.) Heynh. Plant
Physiol 111, 1011-1019.
Wasternack, C., and Hause, B. (2013). Jasmonates: biosynthesis, perception, signal
transduction and action in plant stress response, growth and development. An update to
the 2007 review in Annals of Botany. Ann Bot 111, 1021-1058.
Xu, C., Fan, J., Riekhof, W., Froehlich, J.E., and Benning, C. (2003). A permease-like
protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J 22, 23702379.
Xu, Z., Escamilla-Trevino, L., Zeng, L., Lalgondar, M., Bevan, D., Winkel, B., Mohamed,
A., Cheng, C.L., Shih, M.C., Poulton, J., and Esen, A. (2004). Functional genomic
analysis of Arabidopsis thaliana glycoside hydrolase family 1. Plant Mol Biol 55, 343367.
Yang, D.L., Yao, J., Mei, C.S., Tong, X.H., Zeng, L.J., Li, Q., Xiao, L.T., Sun, T.P., Li, J.,
Deng, X.W., Lee, C.M., Thomashow, M.F., Yang, Y., He, Z., and He, S.Y. (2012).
Plant hormone jasmonate prioritizes defense over growth by interfering with gibberellin
signaling cascade. Proc Natl Acad Sci U S A 109, E1192-1200.

21

Zhang, F., Yao, J., Ke, J., Zhang, L., Lam, V.Q., Xin, X.F., Zhou, X.E., Chen, J., Brunzelle,
J., Griffin, P.R., Zhou, M., Xu, H.E., Melcher, K., and He, S.Y. (2015). Structural
basis of JAZ repression of MYC transcription factors in jasmonate signalling. Nature
525, 269-273.
Zust, T., Rasmann, S., and Agrawal, A.A. (2015). Growth-defense tradeoffs for two major
anti-herbivore traits of the common milkweed Asclepias syriaca. Oikos 124, 1404-1415.

22

CHAPTER 2
SENSITIVE TO FREEZING2 Aids in Resilience to Salt and Drought
in Freezing-Sensitive Tomato

The work presented in this chapter has been published in Plant Physiology
(www.plantphysiol.org) with the copyright belonging to American Society of Plant Biologists:
Wang, K., Hersh, H.L., and Benning, C. (2016). SENSITIVE TO FREEZING2 Aids in
Resilience to Salt and Drought in Freezing-Sensitive Tomato. Plant Physiology 172, 1432-1442.

23

ABSTRACT
SENSITIVE TO FREEZING2 (SFR2) is crucial for protecting chloroplast membranes
following freezing in Arabidopsis (Arabidopsis thaliana). It has been shown that SFR2 homologs
are present in all land plants, including freezing-sensitive species, raising the question of SFR2
function beyond freezing tolerance. Similar to freezing, salt and drought can cause dehydration.
Thus, it is hypothesized that in freezing-sensitive plants SFR2 may play roles in their resilience to
salt or drought. To test this hypothesis, SlSFR2 RNAi lines were generated in the cold / freezingsensitive species tomato (Solanum lycopersicum [M82 cv]). Hypersensitivity to salt and drought
of SlSFR2-RNAi lines was observed. Higher tolerance of wild-type tomatoes was correlated with
the production of trigalactosyldiacylglycerol, a product of SFR2 activity. Tomato SFR2 in vitro
activity is Mg2+-dependent and its optimal pH is 7.5, similar to that of Arabidopsis SFR2, but the
specific activity of tomato SFR2 in vitro is almost double that of Arabidopsis SFR2. When salt and
drought stress were applied to Arabidopsis, no conditions could be identified at which SFR2 was
induced prior to irreversibly impacting plant growth, suggesting that SFR2 protects Arabidopsis
primarily against freezing. Discovery of tomato SFR2 function in drought and salt resilience
provides further insights into general membrane lipid remodeling-based stress tolerance
mechanisms and together with protection against freezing in freezing-resistant plants such as
Arabidopsis, it adds lipid remodeling as a possible target for the engineering of abiotic stressresilient crops.

24

INTRODUCTION
As one of the major environmental plant stresses, freezing greatly afflicts the production
of crops and limits the geographic distribution of naturally occur- ring plants (Pearce, 2001).
Adaptations to freezing have been studied at the physiological, metabolic, molecular, and genetic
levels (Andrews, 1996; Browse and Xin, 2001; Griffith and Yaish, 2004). The Arabidopsis
(Arabidopsis thaliana) mutant sensitive to freezing2 (sfr2) was discovered as a freezing-sensitive
plant during a forward genetic mutant screen (Warren et al., 1996). The SFR2 protein is
associated with the outer envelope membrane of chloroplasts and is crucial for maintaining
chloroplast membrane integrity after exposure to freezing temperatures (Fourrier et al., 2008;
Roston et al., 2014). Despite the fact that this protein was originally classified as a family I
glycosyl-hydrolase and originally described as glucosidase (Thorlby et al., 2004), SFR2 has
galactosyltransferase activity (Moellering et al., 2010; Roston et al., 2014). Once activated under
freezing conditions, SFR2 processively transfers galactosyl residues from the most abundant
chloroplast membrane lipid monogalactosyldiacylglycerol (MGDG) to a second galactolipid
acceptor, forming oligogalactolipids and diacylglycerol (DAG). The DAG is then converted to
triacylglycerol (Moellering et al., 2010). This mechanism of remodeling chloroplast membrane
lipids upon freezing helps adjusting envelope membranes by removing extra polar lipids as the
organelle shrinks due to apoplastic ice formation and cellular dehydration. Membranes are
stabilized by increasing the ratio of bilayer-forming to nonbilayer-forming lipids. In addition,
through increased abundance of polar oligo- galactolipid head groups, the hydration of the
envelope membrane is increased during freezing-related dehydration (Moellering et al., 2010).
Phylogenetic analysis shows that SFR2 orthologs are ubiquitous in the genomes of land
plants, including freezing-sensitive species. Ectopic expression of SFR2 orthologs from freezing-

25

sensitive soybean (Glycine max) and rice (Oryza sativa) has been shown to revert the sfr2
phenotype in Arabidopsis (Fourrier et al., 2008). This finding suggests that SFR2 orthologs are
functionally interchangeable regardless of their origin from freezing-tolerant or -sensitive species.
Therefore, we hypothesized that SFR2 must play physiological roles beyond freezing resilience in
plants that never encounter freezing temperatures in their natural habitats. Given the sessile nature
of all plants, encountering unfavorable growth conditions even in lush tropical environments is at
times inevitable. Unfavorable conditions might include periods of high temperature and drought,
or increased salt concentrations in soils following coastal flooding. Salt and drought stress are
conceptually closely related to freezing, as all are accompanied by severe dehydration of plant
cells (Andrews, 1996; Verslues et al., 2006). Drought stress directly decreases the water potential
in the apoplast and causes water deficiency within the symplast. Increased salt exposure can change
the cellular ion and solute homeostasis, which directly limits water availability for other cellular
processes such as biochemical reactions or membrane hydration. Upon freezing, cellular
dehydration is caused by extracellular ice formation, which acts as a nucleation site for water
drawn out of the cell (Andrews, 1996). Cellular dehydration in plants usually is accompanied by
the shrinkage of the symplast and organelles, bringing different cellular membranes into close
proximity, which can lead to the fusion of membranes along with membrane bilayer structure
disruption. To alleviate cellular dehydration, plants have evolved common signaling pathways,
transcriptional responses, and metabolic adjustments shared in response to freezing, salt, and
drought exposure (Zhu, 2002; Mahajan and Tuteja, 2005; Krasensky and Jonak, 2012), further
suggesting that these abiotic stresses elicit similar protective mechanisms in plants.
As outlined above, SFR2-mediated remodeling of membrane lipids was shown to protect
chloroplast membrane structure during dehydration caused by freezing. The same may be true for

26

cellular dehydration as the result of salt or drought stress. In fact, in the resurrection plant
Craterostigma plantagineum MGDG is increasingly converted to digalactosyldiacylglycerol
(DGDG) and DAG during desiccation (Gasulla et al., 2013). Even though the responsible enzyme
has not yet been identified, this observation suggests that MGDG conversion to higher order
galactolipids might be a general strategy for plants coping with dehydration. Recently, SFR2 was
shown to interact in guard cells with Open Stomata1, an SnRK2-type protein kinase involved in
mediating ABA responses in Arabidopsis (Waadt et al., 2015). It remains to be shown whether
this interaction is physiologically relevant or whether SFR2 may be involved in remodeling of
chloroplast membranes in guard cells in response to dehydration. Here, tomato (Solanum
lycopersicum) was chosen as a representative cold-sensitive plant to explore possible roles of SFR2
in the protection of plants against dehydration-inducing abiotic stresses other than freezing. In
addition, we tested whether SFR2 could protect Arabidopsis against other abiotic stresses than
freezing.

27

MATERIALS AND METHODS
Plant Materials and Growth Conditions
Tomato cultivar M82 seeds were generously provided by Dr. Cornelius Barry, Michigan
State University. Seed were germinated by being plated on a moisturized filter paper for a week,
then seedlings were transferred to soil and kept in a growth chamber with the condition set as 200300 μE m-2s-1 with a 16 h light/8 h dark and 24/20C (day/night). Plants were watered with tomato
nutrient formula water, specifically, 12.2 g 15-30-15 (N-P-K, %weight) fertilizer, 0.13 g Fe iron
and 0.05 g calcium nitrate in one gallon deionized water.
Arabidopsis (Arabidopsis thaliana) Columbia-2 and sfr2-3 seeds (Moellering et al., 2010a)
were sterilized and plated on Murashige and Skoog medium (Murashige and Skoog, 1962)
containing 1% (w/v) Sucrose. Plants were grown under 100 µE m−2s−1 with a 16-h-light / 8-h-dark
cycle and 22°C.

Rnai Plasmid Constructions and Tomato Transformation
Total RNA was extracted from tomato M82 leaves using the Plant RNeasy RNA extraction
kit (Qiagen); 600 ng RNA was reverse transcribed using SuperScript® III Reverse Transcriptase
(life technology) and oligo(dT)18 following the manufacturer’s protocols. The SlSFR2 RNAi
construct was built by first amplifying the 509-bp (121-629bp) cDNA of SlSFR2 Using primers as
shown in Table 2.1. PCR products were purified by QIAquick PCR Purification Kit (Qiagen) and
then inserted into the pENRT/D vector according to the pENTR/D-TOPO Cloning kit instruction
(Life Technologies). The 509-bp SlSFR2 fragment was finally recombined into pHELLSGATE12
vector by the LR reaction using Gateway LR Clonase II Enzyme Mix (Life technologies). Tomato
transformation was done essentially as described (McCormick, 1991).

28

Quantitative Real-Time PCR
Total RNA from the 4th leaves of 8-week old tomatoes was isolated and reverse transcribed
as described above. qRT-PCR was performed using the SYBR Green PCR Core Reagents mix
(Life Technologies) based on the manufacturer's instructions. Table 2.1 lists the primers used. The
2-ΔΔCt calculation was used to determine the relative mRNA levels.

Lipid Analysis
Lipids were extracted from approximately 50 mg 4-week-old tomato 2nd and 4th leaves as
previously described (Moellering et al., 2010b).

Polar lipids were analyzed on activated

ammonium sulfate-impregnated silica gel TLC plates (Si250; Mallinckrodt Baker, Phillipsburg,
NJ, USA, which are no longer available, and which were later substituted by DC-Fertigplatten SIL
G-25; MACHEREY-NAGEL, Germany) using a solvent system consisting of chloroform:
methanol: acetic acid: water (85:20:10:4, v/v/v/v). Lipids were visualized by brief exposure to
iodine vapor or staining with α-naphthol to detect glycolipids.

Protein Production
An SlSFR2 yeast expression construct was built by amplifying the full SlSFR2 coding
sequence from a cDNA, which was inserted into the pYES2.1 vector according to the
manufacturer’s instructions (Invitrogen). The recombinant protein contained a C-terminal 6XHis
tag. Arabidopsis AtSFR2 and cucumber CsMGD1 constructs are described in (Roston et al., 2014).
Protein production was essentially done as described in (Roston et al., 2014). Microsomes were
harvested from yeast cells post induction as described in (Dahlqvist et al., 2000) and stored at -80
℃ until use. Protein extraction was done according to (Kushnirov, 2000) and protein production

29

was analyzed by SDS-PAGE and immunoblotting as described in (Roston et al., 2014). SFR2
assays were performed as described in (Roston et al., 2014), except that the MgCl2 concentration
was elevated to 8 mM.

Stress Assays
For Arabidopsis, after 7 days of growth in regular medium seedling were transferred to MS
plates with extra concentrations of salts for salt treatment, or of PEG-8000 to generate different
water potentials for drought treatment. For the drought assay, the detailed recipe for preparing
PEG-infused plates is descried in (van der Weele et al., 2000). Plants were grown for 3 weeks
before ion leakage or lipid analysis.
Tomato seeds were first germinated on moisturized filter paper for 1 week under standard
conditions as described above, and then transferred to soil and grown for 2 weeks before
application. For the salt assay, three-week-old plants were acclimated by daily watering with
nutrient solution, to which 50 mM NaCl was added, for 5 days and then followed by increasing
NaCl to 100 mM for a week and 150 mM for another 2 days. Plant and leaf images were taken at
the end of the salt treatment. Plants were watered daily with normal nutrient solution for one week
after the end of the salt treatment to ensure they resume normal growth.
For the drought assay, three-week-old tomato plants were not watered for 3 days to
acclimate the plants followed by watering with nutrient solution for 4 days. The then the threeweek-old plants were not watered for another 9 days, followed by daily watering. Lipids were
extracted and analyzed at the end of the drought treatment. Plant and leaf images were taken 2
days after resuming watering.

30

Water potential and relative water content measurements were conducted before the
rewatering phase. Water potential was measured with a pressure chamber (PMS instrument
company, Model 600) following the equipment manual.

Water content measurement was

performed according to (Barrs and Weatherley, 1962).
The electrolyte assays were performed as described (Gilmour et al., 1988). Briefly, 3 to 5
leaf blades were immersed in 3 ml of deionized water and the samples were gently agitated for 3
hours. Conductivity was measured using a conductivity meter (YSI model 35). One hundred
percent leakage was defined by placing the leaves in -80°C for 10 hours and then agitating them
for 3 hours before conductivity of the solution was measured. Electrolyte leakage was expressed
as a percentage of the final conductivity.

Chlorophyll Measurements
Chlorophyll was quantified as described (Lichtenthaler and Wellburn, 1983). In brief, Leaf
pigments were extracted in 80% (v/v) acetone from punched discs and measured
spectrophotometrically on an Uvikon 930 spectrophotometer (Kontron instruments, Germany).
Chlorophyll a, chlorophyll b and total carotenoids were calculated from leaf absorbance at 663nm
and 646nm with the following equations: Chlorophyll a (mg/l) = 12.21A663 - 2.81A646; Chlorophyll
b (mg/l) = 20.13A646 - 5.03A663; Total chlorophyll (mg/l) = chlorophyll a + chlorophyll b.

Protein Measurement
Leaf disks were ground with liquid nitrogen to which 100 µl extraction buffer (0.1M Tris
HCl, pH 6.8; 1% SDS; 15% glycerol; 5% (v/v) β-mercaptoethanol) was added, followed by
vigorous mixing. Samples were boiled at 100℃ for 10 minutes followed with centrifugation at

31

13,000 x g for 5 mins and the proteins were quantified using a Bradford assay (BioRad) following
the manufacturer’s instructions.

32

RESULTS
Salt and Drought Treatment Similarly Affect Arabidopsis Wild Type and sfr2-3 Mutant
SFR2 has been shown to be essential for freezing tolerance in Arabidopsis. In principle, if
SFR2 provides protection against cellular dehydration in general, one might expect that it also
protects Arabidopsis against salt and drought stress. To test this hypothesis, we investigated
whether salt or drought tolerance is compromised in the Arabidopsis sfr2-3 mutant, which carries
an sfr2 loss-of-function allele (Moellering et al., 2010a). For the salt stress assay, wild-type (WT,
Col-2) and mutant sfr2-3 seedlings were grown for one week on regular MS medium prior to the
transfer to MS plates containing different concentrations of NaCl (Figure 2.1). Within three weeks
of treatment, all plants were increasingly stunted with increasing NaCl concentrations (Figure 2.1).
No significant difference in growth was observed between the wild type and sfr2-3 (Figure 2.1).
Ion leakage of the leaf tissues also increased in parallel with NaCl concentrations with no
difference observed for the wild type and sfr2-3 (Figure 2.1). To check whether SFR2 was
activated during the treatment, lipids were analyzed by thin-layer chromatography (TLC) (Figure
2.1). To activate SFR2 in vivo, leaves were treated by MgCl2 infiltration (Moellering et al., 2010b)
and as a result WT leaves contained TGDG, a specific product of SFR2, while sfr2-3 did not
(Figure 2.1). No TGDG was detected in all salt treated plant leaves, suggesting that SFR2 was not
activated under the conditions tested (Figure 2.1). Root bending assays in response to NaCl
treatment and direct salt treatment of soil-grown plants showed consistently no differences
between the wild type and the sfr2-3 mutant (Figure 2.2).
For the drought assay, WT and sfr2-3 plants were grown on control MS plates (Ψ=-2.5
MPa) for a week before transfer to other plates with lower water potentials (Figure 2.1). When
grown on MS medium with Ψ as low as -7.5MPa, both wild type and sfr2-3 were dead after 2

33

weeks. After three weeks of growth on MS medium with Ψ=-5 MPa, plants were alive but growth
was pronouncedly compromised (Figure 2.1). No differences were observed between WT and sfr23 (Figure 2.1). In addition, no TGDG accumulation was observed suggesting that SFR2 was not
activated during the drought treatment (Figure 2.1). Similar results were also observed for soilgrown plants as shown in Figure 2.3.
Apparently, for Arabidopsis no condition of salt or drought treatment could be identified
that would lead to activation of SFR2 before the plants were irreversibly compromised. One
interpretation is that salt and drought stress affect essential cellular processes other than chloroplast
membrane stability and if those become compromised in Arabidopsis prior to the membranes, the
hypothesis stated above cannot be tested in Arabidopsis. Therefore, we shifted the focus of this
study to the freezing-sensitive tomato, in which we were able to study the function of SFR2 in
other abiotic stresses than freezing.

Identification of an SFR2 Orthologue of Solanum lycopersicum
To identify the SFR2 orthologue of tomato (M82), a BLASTp search of the tomato genome
protein database (http://solgenomics.net/) was performed using the Arabidopsis SFR2 sequence
(AtSFR2) as the query (Fernandez-Pozo et al., 2015). Tomato gene Solyc01g058140.2.1 encoded
the top scoring protein fragment with a reported E-value of zero and the respective protein was
designated SlSFR2. The two aligned protein sequences showed 59% identity overall and 72%
similarity (Figure 2.4). To corroborate that SlSFR2 has galactolipid : galactolipid
galactosyltransferase activity, the respective cDNA was expressed in a yeast strain that also
expresses a Cucumis sativus MGDG synthase-encoding cDNA (CsMGD1) (Shimojima et al.,
1997) ensuring the production of MGDG, which is the substrate for SFR2. Arabidopsis and tomato

34

SFR2 cDNAs were tested in parallel. SFR2 protein production in yeast was confirmed by
immunoblot using an anti-His antibody against a c-terminal 6xHIS-TAG included with both
constructs (Figure 2.5) and the lipids were extracted from induced cultures. When LacZ, AtSFR2
or SlSFR2 were expressed in yeast lacking MGDG, no galactolipids were detected by TLC (Figure
2.5). When CsMGD1was expressed alone, only MGDG was present. Co-expression of SlSFR2 or
AtSFR2 with CsMGD1 led to the formation of the oligogalactolipids DGDG, tri- (TGDG) and
tetragalactosyldiacylglycerol (TeGDG). Therefore, SlSFR2 encodes an SFR2 protein with
galactolipid : galactolipid galactosyl transferase activity.

SlSFR2 Has Higher Specific Activity Than AtSFR2
It seemed possible that the specific activity and activation of SFR2 orthologs from freezinginsensitive and -sensitive plants differ. The activity of AtSFR2 is Mg2+ and pH-dependent
(Moellering et al., 2010b; Roston et al., 2014; Barnes et al., 2016) and we tested whether this was
also the case for SlSFR2. To estimate SlSFR2 and AtSFR2 specific activities, microsomes were
prepared from the respective yeast cultures and MGDG-based galactosyl transfer activity was
assayed in the presence of 8 mM MgCl2 at pH 7.5 (Moellering et al., 2010b) using approximately
equal amounts of SFR2 proteins as estimated by immunoblotting (Figure 2.6). A LacZ expressing
line was included as a background control. Under these conditions, SlSFR2 activity was nearly
twice that of AtSFR2 (Figure 2.6) and given that approximately equal amounts of protein were
used, its specific activity must be higher than that of the Arabidopsis enzyme. Using different
concentrations of MgCl2 ranging from 0 to 12 mM (Figure 2.6), both enzymes showed similarly
strong Mg2+ dependency of their activity. When the pH-dependency of SlSFR2 was tested using a
series of buffers with varying pH, it was apparent that the optimal pH for SlSFR2 is approximately

35

7.5 (Figure 2.6), which agrees with that observed for AtSFR2 (Roston et al., 2014). Thus, aside
from an observed higher specific activity of SlSFR2, both orthologues behaved essentially the
same under the conditions tested.

Generation of SlSFR2 RNAi Lines in Tomato
To explore physiological functions of SlSFR2 in cold/freezing-sensitive tomato, an
SlSFR2-RNAi construct was introduced into tomato M82 (Figure 2.7). In total, 23 transgenic lines
were generated and the presence of the SlSFR2-RNAi construct was confirmed by PCR using
genomic DNA (Figure 2.8). To examine the RNAi effect, SlSFR2 transcript abundance of the
transgenic lines was quantified by quantitative real-time PCR (Figure 2.7) and the majority of lines
showed an 80% reduction in transcript abundance.
To estimate the maximal in vivo SlSFR2 activity in the RNAi transgenic lines, we
incubated detached tomato leaves in a 0.5 M MgCl2 aqueous solution for 6 hours, which fully
activates the enzyme, or water only (mock) followed by lipid extraction and TLC of the polar lipids
(Fig. 4; Supplemental Fig. S5 for additional lines). Arabidopsis and tomato WT produced TGDG
in the MgCl2-treated leaves only. However, most of the 13 lines with low SlSFR2 RNA abundance
showed a decreased ability to produce TGDG following MgCl2 treatment (Figure 2.7; Figure 2.7).
Based on the extent of SlSFR2 transcript levels, TGDG reduction and number of seeds recovered
from individual transgenic plants, the two transgenic lines 2 and 29 (L2 and L29) were selected
for further study. As shown in Figure 2.7, L2 and L29 produced very little TGDG or TeGDG in
response to MgCl2 treatment showing the effect of RNAi suppression of SlSFR2.

36

SlSFR2 RNAi Transgenic Lines Show Increased Salt and Drought Sensitivity
When grown under normal conditions, SlSFR2 RNAi transgenic lines L2 and L29 had no
obvious growth defects at all ages tested (Figure 2.10, 2.11 and 2.12). To detect responses to
cellular dehydration, SlSFR2 RNAi lines and M82 WT plants were subjected to salt and drought
stress conditions. To apply salt stress, two- (Figure 2.11) or three-week-old plants (Figure 2.10)
were watered over a period of time with an aqueous nutrient solution containing an increasing
amount of NaCl to allow slow acclimatization. Plants of two different ages were included in the
analysis to observe the robustness of the response during at least two different developmental
stages. Precautions were taken that at the end of the treatment both, SlSFR2 RNAi lines and M82
WT plants, were able to recover and resume normal growth after rewatering with nutrient solution,
from which NaCl was omitted. At the end of the treatment with NaCl, SlSFR2 RNAi lines and
M82 WT plants showed a visible reduction in growth compared to nutrient solution only-watered
plants (Figure 2.10 and 2.11). Fresh weight measured for the two-week-old L2 and L29 lines was
significantly reduced (p<0.01; t-test) compared to the WT plants (Figure 2.11). The three-weekold L2 and L29 plants showed increased chlorosis and decreased size on older leaves, e.g. the
second leaf which is the second oldest true leaf (Figure 2.10). On the two-week-old plants the
cotyledons were most chlorotic (Figure 2.11). On the three-week-old plants the second and the
fourth true leaves were compared as representatives of older and younger leaves, respectively
(Figure 2.10). While the fourth leaves were slightly chlorotic on all plants but did not show much
difference between WT and SlSFR2 RNAi transgenic lines (Figure 2.10), the second leaves of L2
and L29 were more chlorotic and had visibly reduced leaf sizes (Figure 2.10). Hence in general,
the older leaves on either plant showed the most severe response to salt treatment, which could be
described as accelerated senescence.

37

To apply drought stress, water was withheld from three-week-old plants for 3 days
followed by watering for two days to acclimatize the plants to drought. This acclimatization
treatment was followed by withholding water for 9 days followed by rewatering for 2 days. Before
rewatering, water potential and relative water content of the WT plants were measured as indicated
in Figure 2.12. Even though SlSFR2 RNAi lines were not affected at the level of the whole plant
with regard to fresh weight or plant height, the older leaves as represented by the second leaves
were smaller and had less fresh weight at the end of the drought period (p<0.01; t-test) (Figure
2.12). After two days of rewatering, most severe chlorosis of older leaves was observed (Figure
2.10), similar as was seen after salt treatment (Figure 2.10). It should be noted that both, WT and
SlSFR2 RNAi transgenic lines were able to recover from this drought treatment following
rewatering.

Chlorophyll and Protein Content of SlSFR2 RNAi Transgenic Lines
Leaf protein and chlorophyll content are sensitive markers for abiotic stresses, including
salt and drought. Thus, chlorophyll contents were quantified on both, the 4th and 2nd leaves of
three-week-old-plants (Figure 2.13) or the cotyledons and 2nd leaves of two-week-old plants
(Figure 2.14) undergoing stress treatment to corroborate the visual phenotypes. Consistent with
the observed chlorosis of three-week-old plants, a decrease in chlorophyll was only observed for
the 2nd leaves during salt (Figure 2.13) and drought stress (Figure 2.13) or the cotyledons of twoweek-old plants following salt treatment (Figure 2.14). Similarly, measuring the total leaf protein
content of three-week-old-plants (Figure 2.15), the 2nd but not the 4th leaves of the transgenic lines
showed a decrease in protein content in response to both stress treatments.

38

SFR2 is Activated in Older Leaves in Response to Salt and Drought Treatment
To assess a direct correlation of the observed phenotypes of SlSFR2 RNAi transgenic lines
with SFR2 function, lipids were extracted and analyzed for oligogalactolipids as a measure of
SFR2 activation. The 2nd and 4th leaves of three-week-old (Figure 2.16), and the cotyledon and 2nd
leaf of two-week-old plants (Figure 2.17) of WT and SlSFR2 RNAi transgenic lines treated as
described above were assayed. Unstressed WT and SlSFR2 RNAi transgenic lines did not produce
detectable amounts of TGDG or TeGDG (Figure 2.16 and 2.17). However, following salt treatment
TGDG was detected in the 2nd leaves in three-week-old (Figure 2.16), and cotyledons of twoweek-old (Figure 2.17) WT plants, but not in the SlSFR2 RNAi transgenic lines L2 and L29.
Younger leaves on both sets of plants did not show an induction of SlSFR2 in WT or transgenic
lines (Figure 2.16 and 2.17). It should be noted that similar to AtSFR2, the SlSFR2 transcript level
did not change during the salt treatment (Figure 2.18), suggesting that the tomato SFR2 protein is
present at all times and directly activated.
Lipid analysis following drought stress was done for three-week-old plants (Figure 2.16).
Lipids were extracted from the 2nd and 4th leaves directly following the 9 day drought treatment
before the recovery phase to avoid the turnover of potential oligogalactolipids during recovery.
Similar to the salt treatment, no TGDG was detected in the 2nd leaves of the SlSFR2 RNAi
transgenic lines L2 and L29, but in the 2nd leaves of the WT plants (Figure 2.16). Therefore, the
growth and chlorosis phenotypes observed for older leaves of the SlSFR2 RNAi transgenic lines
above coincided with the absence of SFR2 activity following salt and drought treatment.

39

DISCUSSION
AtSFR2 and SlSFR2 are Genuine Orthologues
The current study was based on the premise that all plant genomes characterized harbor a
presumed SFR2 orthologue, including plants that do never encounter freezing in their natural
environments. Functionality of different predicted orthologues including those from freezingsensitive plants was indirectly shown by heterologously restoring the freezing tolerance of the
Arabidopsis sfr2 mutant (Fourrier et al., 2008). At that time the biochemical activity of AtSFR2
was not accurately known, but later shown to be the processive transfer of galactosyl residues from
MGDG to a galactolipid acceptor to form oligogalactolipids and DAG (Moellering et al., 2010b;
Roston et al., 2014). Here we conclusively determined that the recombinant SFR2 orthologue from
tomato, SlSFR2, when produced in a suitable yeast host containing the MGDG substrate has
galactolipid : galactolipid galactosyltransferase activity comparable to its Arabidopsis orthologue
AtSFR2 (Figure 2.6). Further biochemical comparison of the two recombinant proteins in yeastderived microsomal fractions indicated that both enzyme respond qualitatively similarly to
changes in pH and MgCl2 concentrations in the buffer (Figure 2.6). Thus, the two enzymes from a
freezing-tolerant and a -sensitive species appear to function fundamentally in similar ways at the
biochemical level. However, it should be noted that SlSFR2 exhibited about 2 fold higher
galactolipid : galactolipid galactosyltransferase activity than AtSFR2 (Figure 2.6).
AtSFR2 was shown to contain multiple sequence regions that are important for its
galactosyl transferase activity (Roston et al., 2014). The SlSFR2 sequence is only 72% similar to
that of AtSFR2 (Figure 2.4), allowing room for sufficient diversity to explain these subtle
differences. Hence, surveying a larger range of SFR2 orthologues from diverse plants for their
specific SFR2 activity might yield enzymes with different activation profiles or extent of activities

40

that might be better suited for future efforts to engineer tolerance to cellular dehydration based on
SFR2 for increasingly stress tolerant crops. For example, overexpression of a cDNA encoding a
tightly regulated SFR2 enzyme may not lead to crop protection, while a constitutively activated
SFR2 orthologue or one with higher sensitivity to MgCl2 might be more suitable for engineering
purposes.
In Arabidopsis, the AtSFR2 transcript and protein levels are kept constant in the tissues and
under all conditions tested (Thorlby et al., 2004). Similarly, we observed a constant abundance of
SlSFR2 transcripts at all instances tested in tomato (Figure 2.18), suggesting that as in Arabidopsis
activation of SFR2 occurs post-transcriptional/translational.
Aside from the direct determination of the activity of SlSFR2, reducing the abundance of
SlSFR2 transcripts in SlSFR2 RNAi lines decreased the formation of the oligogalactolipids TGDG
and TeGDG in response to MgCl2 (Figure 2.7), providing further corroboration that SlSFR2 is
involved in the formation of these oligogalactolipids in vivo. It should be noted that 0.5 M MgCl2
used in this experiment constitutes a much higher concentration than found under physiological
conditions, which is usually 2-10 mM (Shaul, 2002). We used this concentration of MgCl2 here as
a simple method to induce the enzyme in vivo and estimate the maximal enzyme activity.

Oligogalactolipid Formation by SlSFR2 Protects Against Salt and Drought Stress
Having established in multiple ways that SlSFR2 is a genuine orthologue of AtSFR2 with a similar
activity profile, we can pursue the question whether SlSFR2 protects tomato, a generally
cold/freezing sensitive plant, against other forms of cellular dehydration induced by osmotic stress,
either high salt or drought. A set of SlSFR2 RNAi lines with up to 80% reduction in mRNA
abundance was generated and analyzed to answer this question. Exposing different WT and

41

transgenic plants at different ages to salt and drought stress using generally accepted protocols
(Conroy et al., 1988; Shalata and Tal, 1998; Umezawa et al., 2000), we were able to show that
older leaves on the SlSFR2 RNAi plants remain smaller and senesce earlier than corresponding
leaves on WT plants (Figure 2.10, 2.11 and 2.12). Most importantly, this phenotype strictly
correlated with the inability to generate oligogalacotlipids in those leaves in the transgenic lines
(Figure 2.16 and 2.17). Hence we conclude that the ability to produce oligogalcotlipids as a result
of induction of SlSFR2 activity under salt and drought stress allows WT tomato plants to continue
to produce biomass under these adverse conditions.
The fact that older leaves are most severely affected by different abiotic stress regimes
while young leaves stay relatively protected has been previously observed. For example, the
drought-tolerant plant Senecio medley-woodii responds to drought stresses with a gradual
dehydration of leaves progressing from the oldest to the young leaves until the older leaves were
eventually shed while the younger leaves were still be able to maintain or even increase
transpiration (Donatz and Eller, 1993). In Arabidopsis during drought stress, enzymes protecting
against oxidative stress including catalase, peroxidase, superoxide dismutase and glutathione
reductase accumulate in mature but not in young leaves, suggesting indirectly that older leaves
may be more susceptible to drought-induced oxidative damage (Jung, 2004). Even though the
concept that water translocation from older to younger leaves during dehydration stress may still
be debatable, it is well known that younger photosynthetically active tissues are preferably
protected (Munns and Tester, 2008; Chaves et al., 2009). In rice, young leaves are protected from
salt stress as NaCl accumulates preferably in the older leaves (Wang et al., 2012). Similarly,
younger leaves are more effective than older in adjusting the osmotic potential by increasing solute
concentrations, which helps maintain water status and turgor pressure during dehydration

42

conditions (Hajlaoui et al., 2010). It is possible that the salt stress phenotype of the older tomato
leaves observed in our hands is caused by a combination of NaCl toxicity and cellular dehydration.
However, dehydration must be at least partially responsible, because a similar phenotype was
observed during drought stress alone.

Lipid Remodeling by SFR2 Protects Against Different Abiotic Stresses in Different Plants
SFR2 has been shown to play a crucial role in freezing tolerance in Arabidopsis, but is not
activated during salt or drought stress, at least prior to the occurrence of irreversible damage
(Figure 2.1, 2.2 and 2.3). While Arabidopsis is freezing tolerant, it is much more sensitive to salt
and drought stress and effects of salt or drought become lethal before the SFR2-based lipid
remodeling system is activated. However, in freezing-intolerant tomato, SFR2 is activated during
salt and drought stress before the plants are irreversibly damaged and, hence, is able to protect old
leaves from stress-induced senescence (Figure 2.10 and 2.16).
What are the common principles by which SFR2 potentially protects against cellular
dehydration in different plants? Given that SFR2 mRNA levels are not regulated in different plants,
the enzyme itself must be activated by post-transcriptional mechanisms. We recently showed that
SFR2 in Arabidopsis is activated upon freezing by a decrease in cytosolic pH and an increase in
the concentration of MgCl2 (Barnes et al., 2016). Typically, chloroplasts contain up to 10 mM
MgCl2, a higher concentration than found in the cytosol (Shaul, 2002). The SFR2 protein is likely
ubiquitously present in the outer envelope membranes of plant chloroplasts as has been directly
shown for Arabidopsis (Fourrier et al., 2008). It should be noted that we tested the available
antibody raised against AtSFR2, but failed to detect cross reactivity with SlSFR2. In addition, we
were unable to assay SlSFR2 in isolated tomato chloroplasts, perhaps due to interference with

43

secondary metabolites present in the tomato extracts. However, as in vivo activation in tomato
shows (Figure 2.16 and 2.17), SlSFR2 has access to its substrate MGDG, which is exclusively
present in the plastid. Hence we assume that SlSFR2 is likely also present in the outer envelopes
of chloroplasts as shown for Arabidopsis. Therefore, a simple hypothesis for the protective
function of SlSFR2 during drought and salt stress causing severe cellular dehydration is that
SlSFR2 acts as a first line of defense being present at all times in the chloroplast envelopes. It is
activated when the chloroplast membranes are directly disrupted during dehydration allowing
leakage of MgCl2 from the chloroplast. Alternatively, the MgCl2 concentration increases due to
loss of water from the cytosol during dehydration. Once activated by the cytosolic increase in
MgCl2, the removal of the non-bilayer forming lipid MGDG and the synthesis of higher order
oligogalactolipids stabilize the chloroplast membranes similarly as proposed for AtSFR2 function
during the response to freezing in Arabidopsis (Moellering et al., 2010b; Moellering and Benning,
2011). Therefore, SFR2 constitutes a first line of defense providing a common molecular
membrane lipid remodeling mechanism against cellular dehydration accompanying different
abiotic stresses such as drought, increased salt, or freezing, thereby explaining its presence in all
plants currently tested.
However, other mechanisms than disruption of the chloroplast membrane raising the
cytosolic MgCl2 concentration and lowering the pH to activate SFR2 cannot be excluded. It is
possible that MgCl2 or protons released from vacuoles are sensed by SFR2 upon salt or drought
stress in tomato, and MgCl2 and protons could enter the cytosol through specific channels that may
be differently regulated in different plants in response to different abiotic stresses. Therefore,
understanding the mechanisms of SFR2 activation in different plants in response to different

44

abiotic stresses may provide new plant-specific avenues to engineer crops to be more tolerant to
specific abiotic stresses.

45

ACKNOWLEDGEMENT
This work was done with the assistance of Hope Lynn Hersh, who helped with the plant
maintenance and contributed Figure 2.18. We thank Dr. Cornelius Barry (Michigan State
University) for generously providing tomatoM82 seeds, Ms. Kathleen Imre and Dr. Rob Last
(Michigan State University) for the tomato transformation protocol, Dr.Wayne Loescher
(Michigan State University) for advice on salt and drought assays and providing the pressure
chamber, Dr. Sarah J. Gilmour (Michigan State University) for providing the conductivity meter,
and Dr. Rebecca Roston (University of Nebraska) for scientific discussions.

46

APPENDIX

47

Primer name

Sequence 5’ – 3’

Purpose

Sl_SFR2 RNAi F

CACCCGCTCTCCCATAGACGACTC

For RNAi
constructs

Sl_SFR2 RNAi R

GGCATGATCCTTGACCAATC

For RNAi
constructs

Sl_SFR2 F

AGCTCCAAACTGCTGCTAAA

For qPCR

Sl_SFR2 R

CTCATCAAGTCCTCCTGCATAC

For qPCR

Sl_TIP41-like F

GGTTCCTATTGCTGCGTT

qPCR
reference gene

Sl_TIP41-like R

CGAAGACAAGGCCTGAAA

qPCR
reference gene

Sl_PDF2 F

GCCAATCAAGCACTCCAG

qPCR
reference gene

Sl_PDF2 R

ACCTTTCCCCTTTTGCTTCT

qPCR
reference gene

pHELLSGATE12-FW

GAAGGAGAAAAACTAGAAATTTA
CC

For genotyping

Table 2. 1. Primers used in this study.

48

Figure 2. 1. SFR2 is not involved in protection against salt or drought stress in Arabidopsis.
(A) and (D) Images of plants grown on MS agar plates with, (A), different concentrations of NaCl
or with, (D), different water potentials as indicated. One-week-old normal MS-grown seedlings
were transferred to salt or drought mimicking plates and grown for 3 weeks before the pictures
were taken.
(B) and (E) Ion leakage measurements of the, (B), salt or, (E), drought-treated plant leaves (n=4,
± SD).
(C) and (F) Thin-layer chromatography of the lipids from, (C), salt-stressed leaves or, (D),
drought-stressed leaves. MgCl2 treated WT samples served as positive controls for SFR2
activation. Lipids were stained for sugar head groups. DGDG, digalactosyldiacylglycerol; MGDG,
monogalactosyldiacylglycerol; TGDG, trigalactosyldiacylglycerol.

49

Figure 2. 2. SFR2 is not involved in protection against salt stress in Arabidopsis.
(A) and (B) Root bending assay with salt treated plants. 1-week-old vertically grown WT and sfr23 seedlings were transferred to MS agar plates with different concentrations of NaCl and grown
on plates 180° inverted for 2 weeks. (A), representative images of the 3-week-old plants. (B),
bended roots were isolated and measured (n=6-12, ± SD).
(C) and (D) Salt stress assay with soil-grown plants. (C), plant image of untreated and salt treated
plants. To avoid possible growth differences for different pots, WT and sfr2-3 were grown together
in the same pots. Two-week old plants were watered with Arabidopsis nutrient solution with extra
50 mM NaCl for one week, 100 mM NaCl for one week, 150 mM NaCl for four days and 200 mM
NaCl for five days. Pictures were taken at the end of the treatment. (D), TLC analysis of lipids
extracted from leaves of untreated or salt treated plants. MgCl2 treated WT samples served as
positive controls for SFR2 activation. Lipids were stained for sugar head groups. DGDG,
digalactosyldiacylglycerol;
MGDG,
monogalactosyldiacylglycerol;
TGDG,
trigalactosyldiacylglycerol.

50

Figure 2. 3. SFR2 is not involved in protection against drought in Arabidopsis.
(A) Plant image of untreated and drought treated soil-grown plants. WT and sfr2-3 were grown in
the same pots. Water was withheld from two-week-old plants and pictures were taken after three
weeks of drought treatment.
(B) Ion leakage measurement of the untreated or drought- treated plant leaves (n=5-8, ± SD). C,
TLC analysis of lipids extracted from leaves of untreated or drought stressed plants as indicated.
MgCl2-treated WT samples served as positive controls for SFR2 activation. Lipids were stained
for sugar head groups. For lipid abbreviations, refer to the legend of Supplemental Figure S1.

51

AtSFR2
SlSFR2

1 EEGKFFFGLATAPAHAEDDLDDAWLQFAKETPCSAEEAE----------1 GEKGFFFGLATAPAHVEDRLDDAWLQFAKNTESHEIQQPQTADAIMGSAT

AtSFR2
SlSFR2

40 --------------AADKKARRKKVKLAVGAITKGLAKNTHGKEDKNAAD
51 GDGGSQQALLPQREATKTIKRKKSLKIAIEAQIRGFEKYIEVEE--LTPT

AtSFR2
SlSFR2

76 KPPSKNVAAWHNAPHAEDRLKFWSDPDKEVKLAKDTGVTVFRMGVDWSRI
99 EQCPHNVAAWHNVPHPEERLRFWSDPDIELKLAKNTGVQVFRMGVDWSRI

AtSFR2
SlSFR2

126 MPVEPTKGIKEAVNYEAVEHYKWILKKVRSNGMKVMLTLFHHSLPPWAAD
149 MPEEPLGGLKETVNFAALERYKWIINRVRSYGMKVMLTLFHHSLPPWAGE

AtSFR2
SlSFR2

176 YGGWKMEKTVDYFMDFTRIVVDSMYDLVDSWVTFNEPHIFTMLTYMCGSW
199 YGGWKLEKTVDYFMEFTRLIVDSVADIVDYWVTFNEPHVFCMLTYCAGAW

AtSFR2
SlSFR2

226 PGNNPDFLEIATSTLPMGVFHRALHWMAVAHSKAYDYIHGKISLKKPLVG
249 PGGNPDMLEVATSALPTGVFNQTMNWIAIAHTKAYDYIHEKSKPASAIVG

AtSFR2
SlSFR2

276 VAHHVSFMRPYGLFDIGAVTISNSLTIFPYIDSICEKLDFIGINYYGQVR
299 VAHHVSFMRPYGLFDVAAVSVANSMTLFPFLDCISDKMDYIGINYYGQ--

AtSFR2
SlSFR2

326 ELQVKIAIRSQILINNIAFSRISMLESDSRNQEAVCGAGLKLVETDEYSE
347 --------------------------------EVICGAGLKLVETDEYSE

AtSFR2
SlSFR2

376 SGRGVYPDGLYRVLLMFHERYKHLKVPFIVTENGVSDETDVIRRPYLIEH
365 SGRGVYPDGLFRVLLQFDERYKHLNLPFIITENGVSDGTDLIRQPYLLEH

AtSFR2
SlSFR2

426 LLALYAAMLKGVPVLGYIFWTISDNWEWADGYGPKFGLVAVDRSHDLART
415 LLATYAAMMMGVRVLGYLFWTISDNWEWADGYGPKFGLVAVDRANDLARI

AtSFR2
SlSFR2

476 LRQSYHLFSKIVKSGKVTRKDRSLAWNELQKAAKAGKLRPFYRGVDNHNL
465 PRPSYNLFSKVAESGKITREDREQVWGELQTAAKEGKRRPFYRSVNKYGL

AtSFR2
SlSFR2

526 MYADGLDKPQWRPFVDRDWRFGHYQMDGLQDPLSRVARTLLIWPLIMKKR
515 MYAGGLDEPIWRPYIKRDWRFGHYEMEGLQDPLSRLARYLLH-PLSFKQK

AtSFR2
SlSFR2

576 IRKVKIKHTDDAGLVLHPALASPFD
564 AQTQ----RESDQLTLEPLSANI--

Figure 2. 4. Protein sequence alignment between AtSFR2 and SlSFR2.
Amino acid sequences were aligned by Clustal Omega (http://www.ebi.ac.uk/Tools/msa/clustalo/)
and
consensus
sequences
were
highlighted
using
BOXSHADE
3.21
(http://www.ch.embnet.org/software/BOX_form.html). Identical residues were highlighted black
and similar residues were highlighted grey. Identity between two sequences is 59% (407/679);
positives are 72% (493/679) and gaps are 9% (67/679).
52

Figure 2. 5. SlSFR2 is a galactolipid : galactolipid galactosyltransferase (GGGT).
(A) Immunoblot of the proteins extracted from yeast that are transformed with constructs as
indicated. LacZ, AtSFR2, and SlSFR2 constructs incorporate a His-tag at the C-terminus of the
protein, but not the CsMGD1 construct.
(B) Thin-layer chromatography of the lipids extracted from yeast strains as indicated.
Galactolipids were selectively stained by α-naphthol. Strains solely producing LacZ, AtSFR2, or
SlSFR2 cannot produce galactolipids (left three lanes). Expression of CsMGD1 produces MGDG
(the fourth lane) and co-expressing CsMGD1 with AtSFR2 or SlSFR2 results in the production of
MGDG and DGDG, TGDG and tetragalactosyldiacylglycerol (TeGDG) (fifth and sixth lanes).
For lipid abbreviations, refer to the legend of Figure 2.1.

53

Figure 2. 6. Activity of SlSFR2 in vitro.
(A) Immunoblot of yeast microsomes containing approximately equal amounts of recombinant
His-tagged LacZ and SFR2 proteins. AtSFR2 and SlSFR2 microsomes with these comparable
protein amounts were used for the assays.
(B) Activity comparison between AtSFR2 and SlSFR2 in a buffer with pH 7.5 and 8 mM MgCl2.
Activity is shown as the molar ratio between the product DGDG and the substrate MGDG (n=3,
±SD).
(C) Magnesium dependency of SFR2 activity. Assay buffers contained MgCl 2 concentrations
ranging from 0 to 12 mM (n=3-4, ±SD). D, pH dependency of SlSFR2 activity. Assays buffers had
a range of pHs from 6.72 to 8.26 (n=3, ±SD).

54

Figure 2. 7. Generation the SlSFR2-RNAi lines.
(A) Simplified diagram of the SlSFR2-RNAi construct. RB, right border; p35S, Cauliflower
mosaic virus (CaMV) 35S promoter; Intron, pyruvate dehydrogenase kinase (PDK) intron; Tnos,
NOS-terminator; LB, left border.
(B) Quantitative RT-PCR probing SlSFR2 mRNA abundance in wild type (SlWT) and SlSFR2RNAi lines. C, thin-layer chromatography of the lipids from Arabidopsis WT plants (AtWT),
tomato WT plants (SlWT) or select transgenic lines (SlL2 and SlL29). The leaves were incubated
in either water or 0.5 M MgCl2 aqueous solution for 6 hours before lipid extraction. For lipid
abbreviations, refer to the legend of Figure 2.1.

55

Figure 2. 8. Genotyping of wild type (WT) and SlSFR2-RNAi lines.
Genomic DNA was extracted from 8-week old plant leaves and used as template to generate PCR
fragments containing parts of the pHELLSGATE12 vector and SlSFR2 sequences using primers 1
and 9 as shown in Supplemental Table 1. Amplicon size is expected to be 583 bp. M, 1Kb plus
DNA ladder (Cat. No. 10787-018, Invitrogen); NC, negative PCR control without genomic DNA.

56

Figure 2. 9. Activation of SlSFR2 in additional SlSFR2-RNAi lines.
Detached leaves were incubated in 0.5 M MgCl2 aqueous solution for 6 h. Tomato wild-type leaf
was incubated with water (Mock) (A) and wild-type samples (WT) incubated in 0.5 M MgCl2
aqueous solution for 6 h (B) were included for comparison. Lipids were extracted at the end of the
treatment and separated on TLC plates. For lipid abbreviations, refer to the legend of Figure 2.2.

57

Figure 2. 10. SlSFR2-RNAi lines are sensitive to salt and drought.
(A) Whole plant images of wild-type (WT) and SlSFR2-RNAi lines L2 and L29 grown under
normal condition or (B), with salt treated or (C), drought treated. The plants were three-week-old
at the beginning of the treatment and images were taken at five weeks of age. In the panels on the
right leaf appearances of the representative fourth (4th) and second (2nd) leaves are shown.

58

Figure 2. 11. Effects of salt treatment on two-week-old tomato plants.
Wild-type tomato M82 (WT) and 2 SlSFR2-RNAi lines L2 and L29 were daily watered with
nutrient solution to which 50 mM NaCl was added for 1 day, 100 mM for 2 days for acclimation,
followed by watering with 150 mM for 9 days. The plants were two-week-old at the beginning of
the treatment and images were taken at 26 days of age.
(A) Plant and leaf phenotype of untreated plants.
(B) Plant and leaf phenotype of salt-treated plants.
(C) Fresh weight quantification of the above ground parts. Statistics: n=4 for untreated controls,
±SD; n=9-12 for salt treated plants, ±SD. * Indicates p<0.01, t-test. The size bars equal 1 cm.

59

Figure 2. 12. Effect of drought treatment on WT and SlSFR2-RNAi plants.
Water was withheld three days from three-week-old plants for acclimation. Watering was resumed
for four days, followed by withholding of water for nine days.
(A) Water potential measurement of the untreated control and drought-treated WT plants at the
end of the treatment. n=10 for untreated plants; n=8 for drought-treated plants, ± SD.
(B) Relative water content measurement of the 2nd and 4th leaves of the untreated and drought
treated WT plants (n=3-5, ± SD).
(C) Measurement of the plant height and internode length of untreated controls and drought treated
plants. n=4 for untreated plants; n=7-8 for drought treated plants, ± SD.
(D) Fresh weight measurement of the above ground parts of the whole plant of drought-treated
WT and SlSFR2 RNAi lines L2 and L29 (n=8, ± SD).
(E) Fresh weight measurement of the 2nd and 4th leaves of the drought-treated WT or SlSFR2 RNAi
lines L2 and L29. Statistics: n=4-5, ± SD. *indicates p<0.01, t-test.

60

Figure 2. 13. Chlorophyll in the leaves of untreated and stressed three-week-old tomato
plants.
(A) Total chlorophyll and chlorophyll a, b content in untreated or (B) Salt-treated and (C), of
drought-treated wild-type (WT) and transgenic L2 and L29 lines plants as shown in Figure 2.10.
Statistics: n = 3, ±SD.* indicates p<0.01, t-test.

61

Figure 2. 14. Chlorophyll in the leaves of untreated and salt-treated two-week-old tomato
plants.
(A) Total chlorophyll and chlorophyll a, b content in untreated, and (B), salt-treated wild-type
(WT) and transgenic L2 and L29 lines plants as shown in Figure 2.11. Statistics: n = 4, ±SD. *
indicates p<0.01, t-test.

62

Figure 2. 15. Total protein in leaves from untreated and stress treated three-week-old
tomato plants.
(A) Total protein in untreated, (B), salt-treated and (C), of drought-treated wild-type (WT) and
transgenic L2 and L29 lines plants as shown in Figure 2.10. Statistics: n = 3-5, ±SD. * indicates
p<0.01, t-test.

63

Figure 2. 16. SFR2 activation in three-week-old tomato plants subjected to salt or drought
stress.
(A) Lipids were extracted from the 2nd and 4th leaves of untreated wild-type (WT) and SlSFR2RNAiplants (L2, L9) and analyzed by thin-layer chromatography.
(B) Lipids were extracted from the 2nd leaves and (C), from the 4th leaves of salt and droughttreated plants as indicated. Lipids were stained for sugar head groups. Extracts of MgCl2-treated
wild-type leaves (WT, MgCl2; SFR2 induced) were included (A, C) to detect the diagnostic
trigalactosyldiacylglycerol
(TGDG)
following
SFR2
induction.
DGDG,
digalactosyldiacylglycerol;
MGDG,
monogalactosyldiacylglycerol;
SQDG,
sulfoquinovosyldiacylglycerol. A representative result is shown.
64

Figure 2. 17. SFR2 activation in two-week-old tomato plants subjected to salt stress.
(A) Lipids were extracted from cotyledons and 2nd leaves of untreated wild-type (WT) and SlSFR2RNAi plants (L2, L9) and analyzed by thin-layer chromatography.
(B) Lipids were extracted from the cotyledons and (C), from the 2nd leaves of salt-treated plants
as indicated. Lipids were stained for sugar head groups. Extracts of water-treated (WT, H2O, mock)
or MgCl2-treated wild-type leaves (WT, MgCl2; SFR2 induced) were included (A, C) to detect the
diagnostic trigalactosyldiacylglycerol (TGDG) following SFR2 induction. DGDG,
digalactosyldiacylglycerol;
MGDG,
monogalactosyldiacylglycerol;
SQDG,
sulfoquinovosyldiacylglycerol. A representative result is shown.
65

Figure 2. 18. SlSFR2 transcript levels during salt treatment.
Three-week-old tomato plants were treated with nutrient solution to which 100 mM NaCl was
added. RNA was extracted from 2nd and 4th leaves at different time points, 0, 2, 6 hours (h), 1, 3
and 5 days (d). mRNA abundance was normalized to mRNA abundance at 0 hour in the 4th leaf.

66

REFERENCES

67

REFERENCES

Barnes, A.C., Benning, C., and Roston, R. (2016). Chloroplast membrane remodeling during
freezing stress is accompanied by cytoplasmic acidification activating SENSITIVE TO
FREEZING 2. Plant Physiol, 171, 2140-2149.
Barrs, H.D., and Weatherley, P.E. (1962). A Re-Examination of the Relative Turgidity
Technique for Estimating Water Deficits in Leaves. Aust J Biol Sci 15, 413-428.
Chaves, M.M., Flexas, J., and Pinheiro, C. (2009). Photosynthesis under drought and salt
stress: regulation mechanisms from whole plant to cell. Ann Bot 103, 551-560.
Conroy, J.P., Virgona, J.M., Smillie, R.M., and Barlow, E.W. (1988). Influence of Drought
Acclimation and CO(2) Enrichment on Osmotic Adjustment and Chlorophyll a
Fluorescence of Sunflower during Drought. Plant Physiol 86, 1108-1115.
Dahlqvist, A., Stahl, U., Lenman, M., Banas, A., Lee, M., Sandager, L., Ronne, H., and
Stymne, S. (2000). Phospholipid:diacylglycerol acyltransferase: an enzyme that catalyzes
the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Natl
Acad Sci U S A 97, 6487-6492.
Donatz, M., and Eller, B.M. (1993). Plant water status and water translocation in the drought
deciduous CAM-succulent Senecio-Medley-Woodii. J Plant Physiol 141, 750-756.
Fernandez-Pozo, N., Menda, N., Edwards, J.D., Saha, S., Tecle, I.Y., Strickler, S.R.,
Bombarely, A., Fisher-York, T., Pujar, A., Foerster, H., Yan, A., and Mueller, L.A.
(2015). The Sol Genomics Network (SGN)--from genotype to phenotype to breeding.
Nucleic Acids Res43, D1036-1041.
Fourrier, N., Bedard, J., Lopez-Juez, E., Barbrook, A., Bowyer, J., Jarvis, P., Warren, G.,
and Thorlby, G. (2008). A role for SENSITIVE TO FREEZING2 in protecting
chloroplasts against freeze-induced damage in Arabidopsis. Plant J 55, 734-745.
Gilmour, S.J., Hajela, R.K., and Thomashow, M.F. (1988). Cold Acclimation in Arabidopsis
thaliana. Plant Physiol 87, 745-750.
Hajlaoui, H., El Ayeb, N., Garrec, J.P., and Denden, M. (2010). Differential effects of salt
stress on osmotic adjustment and solutes allocation on the basis of root and leaf tissue
senescence of two silage maize (Zea mays L.) varieties. Ind Crop Prod 31, 122-130.
Jung, S.Y. (2004). Variation in antioxidant metabolism of young and mature leaves of
Arabidopsis thaliana subjected to drought. Plant Sci 166, 459-466.
Kushnirov, V.V. (2000). Rapid and reliable protein extraction from yeast. Yeast 16, 857-860.
68

Lichtenthaler, H.K., and Wellburn, A.R. (1983). Determinations of total carotenoids and
chlorophylls a and b of leaf extracts in different solvents. Biochem Soc Trans 11, 591592.
McCormick, S. (1991). Transformation of Tomato with Agrobacterium tumefaciens.
In K Lindsey, ed, Plant Tissue Culture Manual. Kluwer Academic, Dordrecht, The
Netherlands, pp 1–9
Moellering, E.R., and Benning, C. (2011). Galactoglycerolipid metabolism under stress: a time
for remodeling. Trends Plant Sci 16, 98-107.
Moellering, E.R., Muthan, B., and Benning, C. (2010). Freezing tolerance in plants requires
lipid remodeling at the outer chloroplast membrane. Science 330, 226-228.
Munns, R., and Tester, M. (2008). Mechanisms of salinity tolerance. Annu Rev Plant Biol 59,
651-681.
Murashige, T., and Skoog, F. (1962). A Revised Medium for Rapid Growth and Bio Assays ith
Tobacco Tissue Cultures. Physiol Plant 15, 473-497.
Roston, R.L., Wang, K., Kuhn, L.A., and Benning, C. (2014). Structural determinants
allowing transferase activity in SENSITIVE TO FREEZING 2, classified as a family I
glycosyl hydrolase. J Biol Chem 289, 26089-26106.
Shalata, A., and Tal, M. (1998). The effect of salt stress on lipid peroxidation and antioxidants
in the leaf of the cultivated tomato and its wild salt-tolerant relative Lycopersicon
pennellii. Physiol Plant 104, 169-174.
Shaul, O. (2002). Magnesium transport and function in plants: the tip of the iceberg. Biometals
15, 309-323.
Shimojima, M., Ohta, H., Iwamatsu, A., Masuda, T., Shioi, Y., and Takamiya, K. (1997).
Cloning of the gene for monogalactosyldiacylglycerol synthase and its evolutionary
origin. Proc Natl Acad Sci USA 94, 333-337.
Thorlby, G., Fourrier, N., and Warren, G. (2004). The SENSITIVE TO FREEZING2 gene ,
required for freezing tolerance in Arabidopsis thaliana , encodes a beta-glucosidase.
Plant Cell 16, 2192-2203.
Umezawa, T., Shimizu, K., Kato, M., and Ueda, T. (2000). Enhancement of salt tolerance in
soybean with NaCl pretreatment. Physiol Plant 110, 59-63.
van der Weele, C.M., Spollen, W.G., Sharp, R.E., and Baskin, T.I. (2000). Growth of
Arabidopsis thaliana seedlings under water deficit studied by control of water potential in
nutrient-agar media. J Exp Bot 51, 1555-1562.

69

Wang, H., Zhang, M., Guo, R., Shi, D., Liu, B., Lin, X., and Yang, C. (2012). Effects of salt
stress on ion balance and nitrogen metabolism of old and young leaves in rice (Oryza
sativa L.). BMC Plant Biol12, 194.

70

CHAPTER 3
A Plastid Phosphatidylglycerol Lipase Contributes to the Export of Acyl Groups from
Plastids for Seed Oil Biosynthesis

The work presented in this chapter has been published in The Plant Cell (www.plantcell.org)
with copyright belonging to American Society of Plant Biologists:
Wang, K., Froehlich, J.E., Zienkiewicz, A., Hersh, H.L., and Benning, C. (2017). A Plastid
Phosphatidylglycerol Lipase Contributes to the Export of Acyl Groups from Plastids for Seed Oil
Biosynthesis. Plant Cell 29, 1678-1696.

71

ABSTRACT
The lipid composition of thylakoid membranes inside chloroplasts is conserved from leaves
to developing embryos. A finely tuned lipid assembly machinery is required to build these
membranes during Arabidopsis thaliana development. Contrary to thylakoid lipid biosynthetic
enzymes, the functions of most predicted chloroplast lipid-degrading enzymes remain to be
elucidated. Here, we explore the biochemistry and physiological function of an Arabidopsis
thylakoid membrane-associated lipase, PLASTID LIPASE1 (PLIP1). PLIP1 is a phospholipase
A1. In vivo, PLIP1 hydrolyzes polyunsaturated acyl groups from a unique chloroplast-specific
phosphatidylglycerol that contains 16:1Δ3trans as its second acyl group. Thus far, a specific function
of this 16:1Δ3trans-containing phosphatidylglycerol in chloroplasts has remained elusive. The PLIP1
gene is highly expressed in seeds, and plip1 mutant seeds contain less oil and exhibit delayed
germination compared to wild type. Acyl groups released by PLIP1 are exported from the
chloroplast, reincorporated into phosphatidylcholine, and ultimately enter seed triacylglycerol.
Thus, 16:1Δ3trans uniquely labels a small but biochemically active plastid phosphatidylglycerol pool
in developing Arabidopsis embryos, which is subject to PLIP1 activity, thereby contributing a
small fraction of the polyunsaturated fatty acids present in seed oil. We propose that acyl exchange
involving thylakoid lipids functions in acyl export from plastids and seed oil biosynthesis.

72

INTRODUCTION
Life on the Earth is driven by photosynthesis, which converts light energy into chemical
energy and provides us with food and industrial feedstock. Photosynthesis takes place in
chloroplast thylakoids, where protein complexes containing chlorophyll, carotenoids and other
cofactors are embedded into photosynthetic membranes (Kobayashi et al., 2016). Photosynthetic
membranes are composed of a specialized glycerolipid matrix with four major polar lipids (in order
of abundance): mono- and digalactosyldiacylglycerol, (MGDG and DGDG, respectively),
phosphatidylglycerol (PG) and the sulfolipid sulfoquinovosyldiacylglycerol (SQDG) (Andersson
and Dörmann, 2009; Benning, 2010; Boudière et al., 2014). To date, many of the genes and the
respective enzymes needed for photosynthetic membrane lipid biosynthesis have been
characterized in Arabidopsis thaliana. However, we are only beginning to explore the dynamic
changes in photosynthetic membranes under various developmental and environmental conditions
and the underlying mechanisms of plastid lipid turnover and its regulation.
Lipid turnover requires lipases, which are enzymes that hydrolyze ester bonds of
glycerolipids (Troncoso-Ponce et al., 2013; Kelly and Feussner, 2016). Lipases are involved in a
large number of cell biological processes, from maintaining lipid homeostasis to lipid signaling
(Wang, 2004; Scherer et al., 2010; Richmond and Smith, 2011). Phospholipases can be classified
into four major types based on their lipid substrate cleavage sites: phospholipase D (PLD),
phospholipase C (PLC), phospholipase A1 (PLA1), and phospholipase A2 (PLA2). PLD releases
the polar head group and produces phosphatidic acid, while PLC cleaves the phosphodiester bond
at the glyceryl sn-3 position and produces the phosphorylated head group and diacylglycerol. PLA1
and PLA2 release acyl groups from the glyceryl moiety at the sn-1 and sn-2 positions, respectively
(Wang et al., 2012). The Arabidopsis genome encodes approximately 300 proteins that are

73

annotated as lipases, but most of them have not been biochemically verified or have unknown
physiological functions (Li-Beisson et al., 2013; Troncoso-Ponce et al., 2013; Kelly and Feussner,
2016). Some of the characterized chloroplast-located lipases have intriguing physiological
functions. For example, DEFECTIVE IN ANTHER DEHISCENCE1 (DAD1) (Ishiguro et al.,
2001) is a chloroplast-located PLA1 that catalyzes the initial step of jasmonic acid (JA) production,
which is necessary for proper pollen development and biotic stress resistance. Despite their
potentially important functions in membrane maintenance and signaling, the bulk of chloroplastlocalized lipases remain uncharacterized.
In land plants, fatty acid (FA) biosynthesis occurs in plastids. In Arabidopsis, two pathways
are responsible for glycerolipid biosynthesis (Benning, 2009; Hurlock et al., 2014). De novo
synthesized FAs either directly enter the prokaryotic pathway in plastids or are exported to the
endoplasmic reticulum (ER) to be assembled into glycerolipids by the eukaryotic pathway. In
developing embryos, the bulk of synthesized FAs is incorporated into triacylglycerol (TAG),
which serves as the primary energy repository to fuel seed germination. Oleic acid (18:1; carbon :
double bonds) is the FA predominantly exported from chloroplasts. Exported FAs are activated to
acyl-CoAs and initially incorporated into phosphatidylcholine (PC), which is present in the outer
envelope membrane of chloroplasts and in the ER, before reentering the cytosolic acyl-CoA pool
by a process referred to as acyl-editing (Bates et al., 2007). Acyl-editing allows 18:1 to be further
desaturated into polyunsaturated acyl groups attached to PC before reentering the acyl-CoA pool
for incorporation of FAs into other lipids, including TAG. In fact, acyl-editing is one of the
reported mechanisms for directing polyunsaturated FAs into TAG during embryogenesis through
the action of diacylglycerol acyltransferase (Katavic et al., 1995; Routaboul et al., 1999), in parallel
with direct head group exchange between PC and diacylglycerol (DAG) (Bates et al., 2012) or

74

acyl transfer from PC to DAG by phospholipid : diacylglycerol acyl transferase (Dahlqvist et al.,
2000). Whether lipids other than PC are subject to acyl exchange remains to be determined.
Here, we report on the function of PLASTID LIPASE1 (PLIP1). This gene encodes a thylakoidassociated PLA1 that specifically releases polyunsaturated sn-1 acyl groups from 16:1Δ3t-PG
(16:1Δ3t, 16 carbons with one trans double bond following carbon 3 counting from the carboxyl
end, which is always at the sn-2 glyceryl position), a chloroplast-specific lipid. The unusual trans
double bond is introduced by FA DESATURASE4 (FAD4), which has diverged during evolution
from other desaturases introducing cis double bonds (Gao et al., 2009). In developing seeds, the
polyunsaturated acyl groups released by PLIP1 from 16:1Δ3t-PG in the chloroplast are incorporated
into PC before being incorporated into TAG. PLIP1 initiates a mechanism leading to FA export
from the chloroplast and the incorporation of FAs derived from the thylakoid membrane lipid pool
into seed TAG. In addition, 16:1Δ3t-PG serves as a specific substrate for PLIP1 in vivo, providing
insights into the specialized function of this unusual chloroplast lipid.

75

MATERIALS AND METHODS
Plant Material and Growth Conditions
Experiments were performed with Arabidopsis thaliana ecotype Columbia (Col-0). Seeds
of T-DNA insertion lines SALK_102149 (plip1-1) and SALK_147687 (plip1-2) were obtained
from the Arabidopsis Biological Resource Center, Ohio State University. Lines overexpressing
PLIP1 (or PLIP1S422A) were generated by subcloning the coding sequence of PLIP1 or PLIP1S422A
(see below for their origin) into pEarleyGate 101 (YFP at the C-terminus) (Earley et al., 2006),
followed by introducing constructs into Col-0 plants by Agrobacterium tumefaciens-mediated
floral dip (Clough and Bent, 1998). Transformed seeds were initially screened for resistance to
Basta, followed by confirmation by RT-PCR. Primers used for genotyping of T-DNA insertion
lines or for RT-PCR analysis of overexpression lines are given in Supplemental Table 1.
Arabidopsis seeds were vernalized at 4 °C in the dark for two days before being sown on soil and
grown under 100 μmole m−2s−1 (Sylvania FO/741/ECO) in a 16 h light (22 °C) and 8 h dark (20
°C) cycle. Alternatively, sterilized and vernalized seeds were sown onto Phytoagar plates
containing 1 × Murashige and Skoog (MS) growth medium (Murashige and Skoog, 1962) and 1%
sucrose under 100 μmole m−2s−1 in the same light/dark cycle at 22 °C (Wang et al., 2016).

Quantitative Reverse-Transcription PCR
Total RNA was isolated from leaves of 4-week-old Arabidopsis plants grown on soil as
previously described (Wang et al., 2016) using an RNeasy Plant Mini kit (Qiagen). Total RNA
(600 ng) was used to synthesize complementary DNA using SuperScript III Reverse Transcriptase
(Invitrogen). qRT-PCR was performed using SYBR Green PCR Core Reagents mix (Life
Technologies) based on the manufacturer’s instructions. The 2-ΔΔCt calculation was used to

76

determine the relative mRNA levels. Table 3.1 lists the primers used. Reference primers were as
previously described (Robinson and Bolhuis, 2001). For tissue-specific expression analysis, 100
mg dry seeds, five seedlings, roots harvested from five plants, 10 fully opened flowers or 10
siliques harvested nine days after flowering were pooled as one biological repeat.

Confocal Laser Scanning Microscopy
Imaging of YFP fusions was performed on leaves of 4-week-old Arabidopsis grown on soil
using an Olympus FluoView 1000 confocal laser scanning microscope (Olympus) with excitation
at 514 nm and emissions at 600 nm. Chlorophyll autofluorescence was visualized using excitation
at 633 nm and emission at 700 nm. Images were merged and pseudocolored using Olympus
FluoView 1000 confocal microscope software (Olympus).

Protein Extraction and Immunoblot Analysis
Intact chloroplasts were isolated from 4-week-old Arabidopsis plants grown on MS
medium essentially according to (Aronsson and Jarvis, 2002; Roston et al., 2011), followed by
sub-fractionation into stroma and thylakoid according to (Keegstra and Yousif, 1986; Roston et
al., 2012) with minor modifications. In brief, isolated intact chloroplasts were pelleted and ruptured
by resuspension in hypertonic solution (0.6 M sucrose in TE buffer) and the suspension was
homogenized with a Dounce tissue homogenizer. After incubation on ice for 10 min, bulk
thylakoid fractions were harvested by three 1500 × g 5-min centrifugations at 4 °C. Supernatants
were subjected to another 100,000 × g 2-h centrifugation at 4 °C to remove envelope membranes,
and the final supernatants were harvested as the stroma fraction. Total protein from each fraction
was extracted using a Plant Total Protein Extraction Kit (Sigma) according to the manufacturer’s

77

instructions, and protein was quantified using the Bio-Rad Bradford assay. Appropriate amounts
of extracted organellar or total cellular protein were separated by SDS-PAGE (4-20% gradient,
Bio-Rad), transferred to polyvinylidene fluoride membranes (Bio-Rad) and subjected to
immunoblot analysis using primary antisera in 1:1000 to 1:5000 dilutions in TBST buffer (137
mM NaCL; 20 mM Tris base pH 7.5; 0.5% Tween-20). Secondary anti-rabbit or anti-chicken IgG
antibodies were diluted 1:10,000. Positive immunoreactions were visualized using the Horseradish
Peroxidase reaction with SuperSignal West Dura Extended Duration Substrate (Thermo
Scientific), and the chemiluminescent signal was captured using the ChemiDocTM MP imaging
system (Bio-Rad) according to the manufacturer’s instructions.

Recombinant Protein and Antiserum Production
The PLIP1 sequence was amplified from Arabidopsis wild-type cDNA (see above under
RT-PCR procedure) and inserted into pGEM-T-EASY plasmid (Promega). It was then subcloned
into the pET41a plasmid through BamHI and XhoI restriction sites. The PLIP1S422A point mutation
construct was generated with a Q5 Site-Directed Mutagenesis Kit (New England Biolabs).
Constructs were confirmed by sequencing. Final pET41a-PLIP1 and pET41a-PLIP1S422A
constructs were transformed into BL21 (DE3) E. coli strains for protein production. Cultures
grown in LB medium (containing 0.1% glucose) were inoculated with fresh E. coli colonies and
grown to log phase (OD260 0.8) at 37 °C. Protein production was then induced by adding isopropylβ-D-thiogalactopyranoside (IPTG) to a final concentration of 0.2 mM, and the culture was
transferred to 14 °C. Cells were harvested after 3 h of induction. Cultures were harvested and
sonicated to lyse cells. Supernatant was collected after centrifugation at 10,000 × g for 30 min, and
subjected to another 1-h centrifugation at 100,000 × g to remove the majority of membrane bound

78

PLIP1. The supernatant was collected and used to extract and purify PLIP1 recombinant proteins
using a Ni-NTA column as described (Lu and Benning, 2009), except with a modified washing
buffer (50 mM Tris HCl, pH 7.5; 600 mM NaCl; 40 mM imidazole). Purified protein was
concentrated using an Amicon Ultra-15 Centrifugal Filter (Millipore, UFC901024) and recovered
with 1 × PBS buffer. The protein was quantified using the Bio-Rad Bradford assay, aliquoted, and
stored at -20 °C in 30% glycerol.
Recombinant PLIP1S422A was produced in E. coli and purified with a Ni-NTA column as
described above. Purified protein was separated by SDS-PAGE and the corresponding band of
PLIP1S422A was isolated. Protein was recovered by immersing the gel bands into 1 × PBS buffer at
4 °C overnight with gentle agitation. Recovered proteins were concentrated with an Amicon Ultra15 Centrifugal Filter to a final purity above 98%. Antisera were raised in rabbits by Cocalico
Biologicals, Inc. using their standard protocol.

Chloroplast Import Assay
The N-terminal 6×His tag and TEV cleavage site of pET41a-PLIP1 were removed using a
Q5 Site-Directed Mutagenesis Kit (New England Biolabs) and the construct was confirmed by
sequencing prior to use for import assays. The FtsH8 gene was used as a control. Isolation of pea
chloroplasts, import assays and post-import trypsin treatment were done as previously described
(Xu et al., 2005).

PLIP1 Lipase Assay
Commercial lipid substrates were purchased from Avanti Polar Lipids Inc.. For each PLIP1
lipase reaction, 60 µg lipids were used. The organic solvent was removed under an N2 stream, and

79

the lipids were resuspended in 300 µL reaction buffer (0.1 M PBS, pH 7.4; 4.2 mM Anzergent 312 (Anatrace)) and dispersed by sonication for 3 × 10 s on ice (Misonix; Sonicator 3000 with
microprobe; power setting 1.5). Then, 0.5 µg protein in 20 µL 1 × PBS with 30% glycerol was
added to each reaction. The mixture was sonicated again for 10 s with the same parameters
mentioned above and incubated at ambient temperature (~22 °C) for 1.5 h or as indicated for time
courses. The linear range of the reaction was determined by plotting the substrate degradation
percentage along a time course as indicated in Supplemental Figure 2. The reaction was stopped
by lipid extraction, followed by lipid analysis with TLC and gas chromatography as described
below.
To prepare tobacco PG substrates, total lipids were isolated from 4-week-old plant leaves
and resolved by polar TLC. The PG bands were isolated and lipids were recovered from silica
powder by extraction with chloroform-methanol (1:1 by volume).

Lipid Analysis
Lipid extraction, TLC of polar and neutral lipids, transesterification, and gas
chromatography were done as described in (Wang and Benning, 2011). Polar lipids were analyzed
on activated ammonium sulfate-impregnated silica gel TLC plates (TLC Silica gel 60; EMD
Chemical, Germany) using a solvent system consisting of acetone, toluene and water (91:30:7-7.5
by volume). The amount of water was adjusted according to ambient humidity (in general, 7 for
summer; 7.5 for winter). This solvent system was also used for separation of lyso-lipids derived
from MGDG and PG during in vitro lipase assays. For TAG quantification, lipids were resolved
by TLC on DC-Fertigplatten SIL G-2 (MACHEREY-NAGEL, Germany) using a solvent system
consisting of petroleum ether, ether and acetic acid (80:20:1 by volume). For total FA analysis of

80

dry seeds, 3 h transesterification was conducted directly on a number of seeds as specified. Lipids
were visualized on TLC plates by brief exposure to iodine vapor. To separate lyso-lipids from PC,
PE or PI, a solvent system consisting of chloroform, methanol, glacial acetic acid and water
(65:35:8:5 by volume) was used. To separate lyso-PS from PS, the running solvent consisted of
chloroform, methanol and ammonium hydroxide (28-30% NH3 in water) (65:25:5 by volume). To
separate lyso-lipids from DGDG and SQDG the running solvent contained chloroform, methanol,
glacial acetic acid and water (85:20:10:4 by volume).

Pulse-Chase Labeling
For the leaf labeling experiments, detached leaves from 4-week-old soil-grown plants were
incubated in non-radioactive medium (25mM MES-KOH, pH 5.7; 0.01% Triton X-100) in the
light (25W incandescent bulb, ~40 μmole m−2s−1) at ambient temperature for 1 h. Radiolabeling
was initiated by adding sodium [14C]-acetate (specific activity 100 mCi/mmol in ethanol;
American Radiolabeled Chemicals, Inc.) to the medium to provide 1 µCi/mL, followed by a 1-h
incubation with gentle agitation. The leaves were then washed twice in non-radioactive medium
prior to incubation in non-radioactive medium for another 48 h. At various time points after
application of the label, samples were harvested and the metabolism was halted by immediate lipid
extraction. Lipids were extracted and separated by TLC as described above, and radioactivity in
each lipid fraction was analyzed using a scintillation counter (MicroBeta Trilux, Perkin Elmer)
with 3 ml of scintillator solution (4a20, Research Products International Corporation) for 1 min
per sample, or using phosphorimaging (FBCS 810, Fisher Biotech) with quantification by Quantity
One (V 4.6.6).
Embryo labeling experiments were done as described (Bates et al., 2012). Briefly, the

81

newly opened flowers of 4-week-old soil-grown plants were tagged, and nine days later, siliques
were harvested for embryo isolation. For each time point, a 100 µL volume of embryos was
collected from approximately 50 siliques and pre-incubated in non-radioactive buffer (5 mM MES,
pH 5.8; 0.5% sucrose; 0.5 × MS) in the light (~40 μmole m−2s−1) for 20 min with gentle agitation
at room temperature. Labeling was initiated by removing the old medium and replacing it with the
same medium containing 5 µCi sodium [14C] acetate. Pulse labeling lasted for one hour, followed
by washing and replacing with non-radioactive medium to start the chase. Samples were collected
at the indicated time points, and the reaction was quenched by immediate lipid extraction as
described above.

Observation of Embryo Morphology
Siliques were harvested nine days after flowering and subsequently cleared with a clearing
solution (chloral hydrate : glycerol : water = 8 :2 :1) according to (Herr Jr, 1993). Developing
embryos were dissected from siliques after clearance and observed under a Nikon C2 microscope.

Hormone Extraction and Arabidopside measurement
Hormone extraction and quantification was done according to (Zeng et al., 2011) with
minor modifications. Briefly, hormones were extracted from 100 mg (fresh weight) leaf tissue or
50 mg dry seeds with an extraction buffer composed of methanol∶water (80∶20 v/v) containing
0.1% formic acid, 0.1 g/L butylated hydroxytoluene (BHT) and 100 nM abscisic acid (ABA)-d6
as an internal standard. Hormones were analyzed by LC/MS/MS as described (Zeng et al., 2011).
Standard curve samples were prepared by serial dilution of stock ABA, IAA, SA, JA, JA-Ile, and
OPDA to reach final concentrations from 1 nM to 1000 nM of each hormone with a fixed

82

concentration of ABA-d6 of 100 nM. Peak integration and calibration curve processing was
performed using Masslynx software (version 4.1).
Arabidopsides were extracted following the hormone extraction procedure without adding
internal standards. Samples were analyzed using a Waters Xevo G2-XS Q-TOF mass spectrometer
interfaced with a Waters Acquity binary solvent manager and Waters 2777c autosampler. 10 µl
samples were injected onto an Acquity UPLC CSH C18 column (100 x 2.1 mm, Waters Corp)
held at 40 ˚C. A gradient of solvent A (acetonitrile/water (60:40) containing 10 mM ammonium
formate and 0.1% formic acid) and solvent B (Isoproponal / acetonitrile (90:10) containing 10 mM
ammonium formate and 0.1% formic acid) was applied in a 10-minute program with a mobile
phase flow rate of 0.3 ml/minute. The elution was performed as follows: hold for 1 min after
injection at 95% A / 5% B, followed by a seven-minute linear ramp to 0% A / 100% B, hold for
one minute at 100% B, then return to 95% A / 5% B and equilibrate for one minute. LC separated
analytes were ionized by negative ion mode electrospray ionization and mass spectra were
acquired using an MSe method in continuum mode over m/z 50 to 1500 to provide data under nonfragmenting and fragmenting conditions (collision energy ramp from 15-60 V). Peak areas from
extracted ion chromatograms for OPDA/OPDA-MGDG ([M-H]-, m/z 847.5), OPDA/OPDADGDG ([M-H]-, m/z 1009.57), OPDA/dnOPDA-MGDG ([M-H]-, m/z 819.5) were obtained using
Masslynx (version 4.1) software. Identification was based on accurate mass and OPDA or
dnOPDA fragment ions (m/z values are 291.19 and 263.16, respectively). The peaks of the
Arabidopsides were normalized to that of 18:3/18:3-DGDG ([M-H]-, m/z 981.61).

Phylogenetic analysis
The presumed orthologues land plant sequences of Arabidopsis PLIP1 and FAD4 were

83

obtained by BLAST analysis against the NCBI database with default parameters. Only one
sequence per plant with the highest alignment score and over 40% sequence similarity with the
Arabidopsis sequence was considered a presumed orthologue. The amino acid alignment was
created using MUSCLE with default settings in MEGA (version 7.0.21) software. The Maximum
Likelihood phylogenetic trees were built using MEGA (version 7.0.21), and the bootstrap
percentages were based on 1000 replicates.
The correlation coefficiency score was obtained essentially according to (Pazos et al.,
2005). The 18S rRNA sequences were obtained from the SILVA rRNA database (Quast et al.,
2013). The tree built with 18S rRNA sequences was subtracted as background from both the FAD4
and PLIP1 trees before calculating the Pearson’s correlation coefficient. The accession numbers
of the PLIP1, FAD4 orthologous sequences and the 18S rRNA sequences are listed in
Supplemental Table 1.

Accession Numbers
Sequence data from this article can be found in the Arabidopsis TAIR database
(https://www.arabidopsis.org/) under the following accession numbers: At3g61680 for PLIP1,
At2g29980 for FAD3, At4g27030 for FAD4, At5g42020 for BIP2, At1g06430 for FTSH8.

84

RESULTS
PLIP1 is a Chloroplast Thylakoid-Associated Protein
The Arabidopsis genome encodes approximately 300 putative lipases (Li-Beisson et al.,
2013; Troncoso-Ponce et al., 2013; Kelly and Feussner, 2016), among which 46 were included in
the Chloroplast 2010 Project, which was aimed at assigning functions to nearly all plastid-localized
proteins (Lu et al., 2008; Ajjawi et al., 2010). We hypothesized that some of these putative
chloroplast lipases may play roles in the maintenance of photosynthetic membranes and perhaps
have specialized roles in tissues with high demands on lipid metabolism, such as developing seeds
that accumulate TAG. We initially focused on one of the predicted chloroplast lipase genes,
At3g61680, which encodes a protein with a conserved Lipase 3 domain and a strongly predicted
transit peptide, which we subsequently named PLIP1. With its Lipase 3 domain, which is a
signature for triacylglycerol lipases, this Arabidopsis protein shares similarities with a bona fide
lipase of Chlamydomonas, PLASTID GALACTOGLYCEROLIPID DEGRADATION1 (PGD1),
involved in the turnover of chloroplast MGDG, leading to the export of acyl groups and their
incorporation into TAG following nutrient deprivation (Li et al., 2012), although the two proteins
do not share sequence similarity outside the Lipase 3 domain and are not orthologues.
To experimentally verify the subcellular location of PLIP1, the PLIP1 coding sequence
derived from an Arabidopsis wild-type (Col-0) cDNA was spliced at its 3’-end (creating a Cterminal fusion) to the open reading frame of yellow fluorescent protein (YFP). When the PLIP1YFP construct was stably expressed in the wild type under the control of the cauliflower mosaic
virus (CaMV) 35S promoter, the YFP and chlorophyll signals overlapped (Figure 3.1 A). Although
transgenic lines used in this experiment had constitutive expression of PLIP1-YFP, ubiquitous
YFP signals were not detected. Instead, only about 10 to 15% of mesophyll chloroplasts showed

85

YFP signals, and the cells observed in the PLIP1-YFP transgenic plants were slightly smaller than
wild type.
To corroborate the suborganellar location of PLIP1, we isolated intact chloroplasts from 4week-old wild-type seedlings and fractionated them into thylakoid membranes and stroma.
Immunoblot analysis of PLIP1 showed increasing signal intensity from whole plant tissue to intact
chloroplasts and thylakoids, consistent with an association of PLIP1 with the thylakoids (Figure
3.1B). Fractionation quality was controlled for by including marker proteins for each fraction. The
thylakoid protein Light Harvesting Complex b1 (LHCb1) showed a similar intensity pattern to that
of PLIP1. To exclude the possibility of contamination with ER-associated proteins, the ER-specific
marker BiP2 was also included (Figure 3.1 B). The fractionation of the stroma-specific Rubisco
large subunit and the thylakoid-specific light-harvesting chlorophyll a/b-binding protein (LHCP)
were visible on a Coomassie Brilliant Blue-stained SDS-PAGE gel (Figure 3.1 C).
The PLIP1 gene is predicted to encode a 71,735 D protein for which a molecular or
biochemical function had not been experimentally determined. Based on the ARAMEMNON
database, PLIP1 is not predicted to contain any transmembrane domains (Schwacke et al., 2003).
Therefore, it seemed likely that PLIP1 is a peripheral thylakoid membrane protein. To learn more
about its suborganellar location and processing en route, we translated PLIP1 cDNA in vitro in the
presence of labeled methionine. During the import of the translation product into isolated pea
(Pisum sativum) chloroplasts, the PLIP1 precursor was processed into a smaller intermediate
protein that was present in both the stroma and chloroplast membrane fractions. In addition, the
trypsin-resistance of PLIP1 suggested that it is located inside the chloroplast (Figure 3.1 D).
Interestingly, the intermediate PLIP1 form was further processed into a smaller mature protein,
which was mainly associated with the stroma fraction. We used thylakoid lumen-localized FtsH8

86

as a control for proper fractionation and import (Rodrigues et al., 2011), which was processed and
imported into the thylakoids with a pattern different from that of PLIP1, suggesting that PLIP1 is
probably not imported into the thylakoid lumen but instead attaches to the outside leaflet of
thylakoid membranes and can be released into the stroma with additional processing. Summing up
all localization data, PLIP1 is likely a thylakoid membrane-associated protein.

PLIP1 is a Phospholipase A1 With a Preference for Unsaturated Acyl Groups
PLIP1 is annotated as a TAG lipase in TAIR (https://www.arabidopsis.org/). To verify its
activity and determine its enzymatic properties, we tried to develop an in vitro lipase assay with
recombinant PLIP1 purified from E. coli. However, expression of the recombinant 6×His-PLIP1
construct was very low, as detected by immunoblotting against the His tag. Analysis of transgenic
E. coli-derived lipid extracts by thin-layer chromatography (TLC) showed that, when PLIP1 was
expressed, PG and phosphatidylethanolamine (PE), the two major polar lipids of E. coli, were
degraded, leading to the accumulation of free FAs (Figure 3.2 A). This observation suggested that
PLIP1 is a lipase that releases acyl groups from PG and PE.
Lipases belong to a group of serine esterases with a lipase signature motif, an Asp-His-Ser
triad, with some exceptions having only a Ser-Asp dyad, and in all cases Ser serves as the active
site residue, participating in the reaction mechanism (Brady et al., 1990; Winkler et al., 1990;
Richmond and Smith, 2011; Kelly and Feussner, 2016). Alignment of the PLIP1 protein sequence
with those of classic lipases using NCBI’s conserved domain database (Marchler-Bauer et al.,
2015) identified two potential catalytic residues, Asp483 and Ser422, but the conserved His site was
not present (Figure 3.3). Replacing these two residues with Ala, respectively, abolished PLIP1
lipase activity, and the respective mutant proteins were abundantly produced in E. coli, as PG and

87

PE were not degraded (Figure 3.2 B). Taken together, these data suggest that PLIP1 is a lipase
with Ser422 as the catalytic residue. Taking advantage of the enhanced production of nonfunctional
PLIP1S422A in E. coli, we purified the mutant protein and raised an antibody in rabbits to
specifically detect PLIP1.
To develop an in vitro lipase assay, recombinant PLIP1 and PLIP1S422A were produced in
E. coli and affinity-purified from the soluble fraction (Figure 3.2 C). We chose Anzergent 3-12 as
the solubilizing detergent from a series of other reagents, because of its high compatibility with
PLIP1 enzyme activity and because it was not co-chromatographing with native plant membrane
lipids during subsequent TLC analysis. In the final optimized system (Figure 3.4), in vitro lipase
activity of PLIP1 was observed on a wide range of substrates. As an example (shown in Figure
3.2D), PC was provided to PLIP1 and PLIP1S422A. At the end of the reaction, lipids were extracted
and separated by TLC. Lipase activity based on the production of lyso-PC was only observed with
PLIP1, but not with PLIP1S422A (Figure 3.2 D).
PLIP1 in vitro activity with PC and other glycerolipids as substrates always resulted in
lyso-lipid products, implying that PLIP1 can only hydrolyze one of the two acyl-glyceryl ester
bonds in glycerolipids. To investigate which glyceryl position PLIP1 prefers, two PCs with
reversed acyl compositions were offered to PLIP1. At the end of the reaction, lipids were extracted,
lyso-PC was isolated by TLC, and FA methyl esters derived from the lyso-lipid were analyzed by
gas-liquid chromatography (Figure 3.5A). For PC with a composition of 18:1Δ9/16:0 (sn-1/sn-2),
18:1Δ9 was selectively released and 16:0 was retained in the lyso-product. The result was reversed
with PC containing 16:0/18:1Δ9 as the lyso-product contained 18:1Δ9. Therefore, PLIP1 is a lipase
that prefers the sn-1 glyceryl position of the respective glycerolipid. Furthermore, to determine a
possible acyl group preference of PLIP1 at the sn-1 glyceryl position, PLIP1 was offered different

88

combinations of pure PC molecules carrying the same acyl groups at the sn-2 but acyl groups with
different degrees of saturation at the sn-1 position (Figure 3.5 B). Comparing 18:0/18:1 with
18:1/18:1, PLIP1 enzyme activity was approximately twice as high for 18:1/18:1-PC, and when
comparing 18:0/18:2 with 18:2/18:2, its activity was elevated nearly four-fold for 18:2/18:2-PC.
Therefore, PLIP1 is a phospholipase A1 with a preference for more unsaturated acyl groups.
To survey PLIP1 substrate preference in vitro, we offered the majority of the plant
glycerolipids, including galactoglycerolipids, phospholipids, as well as TAG (Figure 3.5 C). High
enzyme activity was detected for all tested phospholipids and MGDG. Given the plastid location
of PLIP1, possible native substrates were limited to PG and MGDG. The low activities detected
for SQDG, DGDG and TAG suggested that these are not likely substrates of PLIP1 (Figure 3.5
C). Based on these results, despite its conserved Lipase 3 domain, PLIP1 is apparently not a TAG
lipase.

18:3/16:1Δ3t-PG is the Preferred Substrate of PLIP1 In Vivo
The in vitro assays in combination with its established chloroplast location narrowed down
possible native PLIP1 substrates to MGDG and PG (Figure 3.5 C). However, given the complexity
of native plant acyl compositions, this limited survey based on an in vitro lipase assay alone could
only provide a first approximation of the likely PLIP1-preferred substrate in vivo. To assess PLIP1
activity in its native biological context, we took advantage of the Arabidopsis transgenic lines used
for PLIP1 localization described above. In total, 30 independent PLIP1-YFP (PLIP1-OX) and 14
PLIP1S422A -YFP (PLIP1S422A-OX) overexpression lines were generated and three PLIP1-OX lines
were selected as representatives. As shown in Figure 4A, compared to wild type, the PLIP1-OX
lines had smaller rosettes and fewer leaves, which were slightly pale yellow, whereas PLIP1S422A-

89

OX plants were indistinguishable from wild-type and empty vector control plants. On a fresh
weight basis, the total leaf acyl group content of the smaller PLIP1-OX plants was not reduced
(Table 3.2). A comparison of the relative abundance of polar lipids (Figure 3.6 A) and the acyl
group composition of individual polar lipids of EV control plants and two PLIP1-OX lines is
shown in Figure 3.7B, C and Supplemental Figure 3.5B-E. In PLIP1-OX lines, the levels of lipids
associated with chloroplasts (MGDG, PG and DGDG) decreased, while the levels of lipids mostly
associated with the ER (PC, PE and PI) increased (Figure 3.6 A), suggesting a reduced ratio of
plastid-to-extraplastidic membranes in the PLIP1-OX lines. Acyl group analysis of individual
membrane lipids showed the greatest changes for PG (Figure 3.7 B). Specifically, the ratio of 16:0
to 16:1Δ3t was increased in the PLIP1-OX lines. For 18-carbon acyl groups, which are primarily
present at the sn-1 position of plastid PG, polyunsaturated 18:3 levels decreased with a concurrent
increase in the relative abundance of 18:1 and 18:2. Based on these changes in the molecular
composition of PG, we hypothesize that 18:3/16:1Δ3t-PG is a preferred substrate of PLIP1 in its
native environment.
MGDG, the most abundant lipid in chloroplasts, also showed a subtly reduced ratio of 16:3
to 18:3 in PLIP1-OX lines (Figure 3.6 B). For ER lipids, a decrease in 18:2 and an increase in 18:1
were observed for PC (Figure 4.7 C), as well as for PE and PI (Figure 3.6 E). Since PLIP1 is
located in the chloroplast and is spatially separated from ER lipids, observation of this ER lipid
alteration suggested that turnover of chloroplast lipids can affect the synthesis of ER lipids,
assuming that lipid precursors are transported from the chloroplast to the ER. The other two
photosynthetic membrane lipids, DGDG and SQDG, showed very minor changes in their
molecular compositions in the PLIP1-OX lines (Figure 3.6 C and D), which is consistent with the
low activity of PLIP1 on DGDG and SQDG in vitro (Figure 3.5 C).

90

The altered morphology of the PLIP1-OX lines raised a concern that the lipid phenotype might be
a secondary phenotype due to the plants’ abnormal development and growth. Therefore, we
investigated whether growth and lipid phenotypes could be separated. Besides stunted growth, the
three PLIP1-OX lines had shorter leaf petioles than wild type and accumulated anthocyanin under
normal growth conditions (Figure 3.7 A), phenotypes reminiscent of plants with activated JA
signaling (Yan et al., 2007; Zhang and Turner, 2008; Campos et al., 2016). Given that PLIP1
releases polyunsaturated FAs including 18:3, a precursor of JA and other oxylipins, we
hypothesized that the reduced growth of PLIP1-OX plants is due to the production of oxylipins
and the activation of JA signaling. Quantitative PCR analysis showed that the expression of JAresponsive marker genes was constitutively increased in PLIP1-OX lines compared to wild type
(Figure 3.8). Targeted hormone analysis of mesophyll tissues showed that the levels of 12-oxo
Phytodienoic acid (OPDA) and OPDA-containing glycerolipids called Arabidopsides were
strongly elevated in the PLIP1-OX mesophyll tissues (Figure 3.9). JA and to some extent OPDA
signaling require the coreceptor CORONATINE INSENSITIVE1 (COI1) (Xie et al., 1998; Ribot
et al., 2008). To corroborate that the growth phenotype is secondary and caused by the activation
of JA signaling pathways, we crossed PLIP1-OX lines with a coi1 mutant to block COI1dependent OPDA and JA signaling. As a result, the growth phenotype was largely (but not fully)
restored in the offspring of this cross as validated by analysis of variance, while the lipid phenotype
remained identical to that of the PLIP1-OX lines (Figure 3.10). Thus, we were able to at least
partially separate the growth and lipid phenotypes, suggesting that the changes in lipid composition
observed represent the primary phenotype resulting from PLIP1 overproduction in the PLIP-OX
lines and are not the result of abnormal growth.

91

Overexpression of PLIP1 Accelerates Recycling of Plastid PG Acyls Groups and Their
Transfer to PC
The analysis of the PLIP1-OX lines described above depicts the lipid composition at
steady-state. However, lipid metabolism is a dynamic process, and pulse-chase labeling is an
effective way of probing the dynamics of lipid metabolism and movement of acyl groups through
different lipid pools and between organelles (Xu et al., 2008; Li et al., 2012). Therefore, we
employed pulse-chase labeling of membrane lipids using [14C]-acetate, which can be readily
incorporated into acyl groups in plastids by the FA synthase complex. The pulse phase of the
experiment (shown in Figure 3.11 A) indicated that MGDG, PG and PC contained the majority of
the label in empty vector (EV) control leaves, with PG accounting for approximately 15% of the
label after 1 hour. However, in PLIP1-OX1, incorporation of label into PG accounted for nearly
70% of total label at the end of the pulse phase. This result suggests that incorporation of de novo
synthesized acyl groups into PG is greatly accelerated in PLIP1-OX lines. During the chase phase
(Figure 3.11 B), PG rapidly lost most of the label (within a day), and the label concomitantly
increased in PC and to a smaller extent in PE in the PLIP1-OX1 line. The EV control line showed
less drastic changes in labeling during the chase phase. The rapid increase and subsequent loss of
PG label in PLIP1-OX1 during the pulse and chase phases, respectively, suggested a rapid acyl
exchange, preferably on PG, in these lines and supported our hypothesis that PG is the preferred
PLIP1 substrate in its native environment.
The most notable acyl group change observed in the PLIP1-OX lines was the increased
16:0-to-16:1Δ3t ratio in PG (Figure 3.7 B). However, 16-carbon FAs only exist at the sn-2 position
of plastid PG. This implies that acyl groups at the glycerol sn-2 position affect PLIP1-catalyzed
hydrolysis at the glyceryl sn-1 position of PG. To test this possibility, we provided purified

92

recombinant PLIP1 with a set of commercial PCs with 16:0 at the glyceryl sn-1 position, but 18carbon acyl groups of different saturation levels at the glyceryl sn-2 position. The highest enzyme
activity was observed for PC with 18:2 at the sn-2 position, followed by 18:1, with the lowest
activity observed for 18:0 (Figure 3.11 C), suggesting that unsaturated sn-2 acyl groups enhance
PLIP1 activity. Therefore, it follows that 16:1Δ3t should be favored over 16:0 at the sn-2 position
of PG. To test this hypothesis, plant-derived PG composed of species containing 16:0 or 16:1Δ3t at
the glyceryl sn-2 position was extracted from tobacco (Nicotiana benthamiana) leaves and offered
to PLIP1 in vitro. Total PG was degraded, while lyso-PG was produced over time (Supplemental
Figure 6A). The fraction of 16:1Δ3t in lyso-PG increased over time, while 16:0 decreased (Figure
3.11 D), indicating that 16:1Δ3t-PG is preferred by PLIP1 under these conditions when native PG
substrate is offered. The opposite pattern between 16:0 and 16:1 Δ3t was observed in retained PG
(Figure 3.12 B). Therefore, we concluded that 18:3/16:1Δ3t-PG is the preferred native substrate of
PLIP1 based on data gathered from the above-described combination of in vitro and in vivo
experiments.
Another interesting observation during the chase phase of the labeling experiment was the
sequential labeling of PG and PC, which points towards a precursor-product relationship between
these two lipids. This observation was also consistent with the decreased plastid-to-extraplastidic
lipid ratio observed during steady-state lipid analysis of the PLIP1-OX lines (Figure 3.6 A).
Therefore, we hypothesized that 18:3 released from 18:3/16:1Δ3t-PG was exported from the plastid
and incorporated into PC. PC is known for its intermediate role in acyl editing involving
desaturation of PC-acyl groups (18:1 to 18:2 and 18:3) followed by acyl exchange (Bates et al.,
2007). We reasoned that the reduced 18:2 content of PC in the PLIP1-OX lines might be due to
the increased competition of incorporation of plastid-derived 18:3 into PC via the activity of ER

93

desaturases FAD2 and FAD3, generating 18:3 from 18:1 and 18:2 bound to PC. To test this
hypothesis, we crossed a PLIP1-OX1 plant to a fad3-2 mutant plant (Figure 3.11 E and 3.13),
which is deficient in the desaturation of 18:2 to 18:3 on ER lipids (PC, PI and PE). We expected
that PLIP1 overexpression might rescue the fad3-2 defect. As reported (Browse et al., 1993), the
fad3-2 mutant had a reduced 18:3 content in ER lipids. The overexpression of PLIP1 in the fad32 mutant background partially reversed this phenotype by increasing the 18:3 content in ER lipids
PC, PI, and PE (Figure 3.11 E and 3.13 F). These 18:3 acyl groups must have been derived from
the chloroplast, where the FAD7/8 desaturases (Li-Beisson et al., 2013) catalyze the lipid-linked
desaturation of acyl groups from 18:2 to 18:3. This increase in 18:3 in PC is paralleled by a
decrease in 18:3/16:1Δ3t-PG (Figure 3.13 E). Taken together, these data support our hypothesis that
in the PLIP1-OX lines, 18:3 increasingly moves from plastid 18:3/16:1Δ3t-PG to PC, which
interferes with the desaturation of acyl groups on PC by ER desaturases FAD2 and 3 and the PCbased acyl editing process.

PLIP1 Contributes a Small Fraction of Precursors for TAG Biosynthesis During
Embryogenesis
The analysis of PLIP1, thus far, has focused on its biochemical function in vitro and in
vivo using overexpression lines. However, these experiments can shed only limited light on the
physiological functions of PLIP1. Querying the native tissue-specific and developmental
expression of PLIP1, the highest gene expression was detected primarily in seeds and reproductive
tissues, including flowers and siliques (Figure 3.14 A). Since PLIP1 encodes a lipase and is highly
expressed in seeds, we postulated that it might play a role in seed lipid metabolism, which is
dominated by the biosynthesis of TAG. In fact, towards the end of seed development, over 90%

94

of total acyl groups are stored in TAG (Li et al., 2006; Li-Beisson et al., 2013). To explore the
physiological function of PLIP1 during embryogenesis, two independent T-DNA insertion lines
were obtained (Alonso et al., 2003). The T-DNA allele corresponding to SALK_102149 was
designated as plip1-1, and the second, corresponding to SALK_147687, was designated as plip12. The T-DNA insertions were in the 3’ and 5’ UTRs, respectively (Figure 3.15 A). Quantitative
RT-PCR analysis indicated that both lines carry leaky alleles (Figure 3.15 B, numbers on panel).
Under normal growth conditions, the two plip1 mutant alleles were physiologically
indistinguishable from the wild-type plants (WT) (Figure 3.15 B). Lipid analysis also showed no
changes in vegetative tissues (Figure 3.15 C-J). However, in dry seeds, where PLIP1 mRNA is
highly abundant, the insertion lines showed an approximate 10% reduction in total seed acyl group
content indicative of a decrease in TAG, while the overexpression lines had a 40-50% increased
seed acyl group content (Figure 3.14 B). Seed weight changes were consistent with the changes in
seed TAG levels; the plip1 mutants had smaller seeds than wild type, while the seeds of the
overexpression lines were larger (Figure 3.14 C). Compromised germination was also observed
for the seeds of the two plip1 mutants (Figure 3.14 D), a typical phenotype related to the reduction
in the major seed carbon reserve, TAG. Surprisingly, seeds from the overexpression lines also had
slower germination rates than wild type, despite their elevated TAG content (Figure 3.14 D). To
investigate whether this is caused by altered seed development due to altered plant hormone levels,
we analyzed the major hormones typically present in dry seeds. As shown in Supplemental Figure
9, the hormone levels of the plip1 mutant seeds were not significantly different from those of the
WT seeds, whereas the PLIP1-OX1 seeds had elevated OPDA and JA-Ile levels (Figure 3.16),
both of which are effective germination inhibitors (Dave et al., 2011; Dave and Graham, 2012).
Arabidopsides were not detected in seeds of any genotypes. It must be noted that mature PLIP1-

95

OX lines had shorter and bushier inflorescences than wild type (Figure 3.17 A), and that seed yield
was reduced by approximately 60% for the PLIP1-OX lines (Figure 3.17 B). Thus, overall oil yield
was not elevated in the PLIP1-OX plants. Nevertheless, the phenotype of the plip1 reduced
function mutant suggested that PLIP1 might play a role in TAG biosynthesis during
embryogenesis.
To gain more information on how the plastid-located PLIP1 contributes to TAG
biosynthesis during embryogenesis, we analyzed TAG acyl groups in dry seeds. In the insertional
mutants, especially in the slightly stronger plip1-2 allele, 18:3 levels increased relative to 18:1
(Figure 3.14 E). For the two PLIP1-OX lines, 18:2 levels decreased, while 18:3 and 18:1 levels
increased, a pattern that resembled the leaf PC acyl group profile in the PLIP1-OX lines (Figure
3.7 C). This suggested that increased TAG may be derived from increased flux of acyl groups
through PC in the overexpression lines. Based on the lipid analysis in vegetative tissues of the
PLIP1-OX lines (Figure 3.7), we hypothesized that PLIP1 in developing embryos contributes to
TAG biosynthesis by catalyzing the turnover of PG, increasing the flux of acyl groups into PC and
ultimately TAG. To corroborate this hypothesis, we analyzed PG and PC content and composition
in developing seeds isolated from siliques seven days after flowering. Total PG content was
approximately 1.6% of total lipids in both WT and plip1-2, while developing seeds of PLIP1-OX1
contained approximately 50% less PG (Figure 3.14 F, inserted panel). With regard to acyl
composition, relative levels of 16:1Δ3t and 18:3 decreased in the PLIP1-OX1 line, while they
increased in plip1-2, supporting a role for PG as the substrate of PLIP1 in developing seeds. PC
accounts for approximately 10% of the total lipids at this stage of developing seeds (Figure 3.14
G, inserted panel), and the acyl compositions of PC in different genotypes resembled that of TAG

96

in dry seeds (Figure 3.14 G), implying that the altered TAG acyl compositions in PLIP1-OX and
plip1 mutant seeds at least partially reflect the metabolism of PC.
It has been proposed that acyl editing of PC, head group exchange, and PDAT activity
represent the mechanisms for directing polyunsaturated FAs from PC into TAG during seed
development (Dahlqvist et al., 2000; Bates et al., 2012). In the fad3-2 mutant, approximately 15%
of 18:3 was retained in total seed FAs (Figure 3.18), suggesting that the proposed PLIP1-based
mechanism is a relatively small but significant contributor to the incorporation of 18:3 into seed
TAG compared to the PC-based FAD2/3 desaturation pathway. In fact, consistent with the
observation in leaf tissues (Figure 3.11 E), overexpressing PLIP1 in the fad3-2 background
restored the 18:3 level to 36% (Figure 3.18).
As discussed above, 16:1Δ3t-PG is likely the preferred substrate of PLIP1 and 16:1Δ3t only
exists at the sn-2 position of plastid PG. FAD4 is the enzyme in Arabidopsis that specifically
introduces trans-double bonds into the 16:0 acyl chain of PG (Gao et al., 2009). Transcript analysis
of FAD4 during seed development showed that this gene is upregulated during embryogenesis, in
a time course that correlates well with oil biosynthesis during embryogenesis (Figure 3.19 A). It
should be noted that the transcript levels of FAD4 are on the same order of magnitude in leaf and
seed tissues (Winter et al., 2007) (Figure 3.19 A), but the 16:1Δ3t-PG level varies drastically
between these two tissues.
Our hypothesis that PLIP1 contributes to embryonic TAG biosynthesis predicts that
removal of 16:1Δ3t-PG should result in a similar seed phenotype to that observed for the plip1
mutants. To test this hypothesis, we characterized two FAD4 loss-of-function lines, fad4-2 and
fad4-3 (Gao et al., 2009). As shown in Figure 8B, 16:1Δ3t was not detected in PG in either fad4-2
or fad4-3 leaf tissues. Similar to the plip1 mutants, fad4-2 and fad4-3 showed a nearly 10%

97

reduction in total seed acyl group content (Figure 3.19 C), reduced seed weight, but no altered seed
yield (Figure 3.19 D). The fad4-2 and fad4-3 mutants also had altered seed acyl group profiles,
specifically reduced 18:2 and increased 18:3 content (Figure 3.19 E), similar to the changes in
plip1-2 seed acyl group composition (Figure 3.14 E). Taken together, these observations led us to
conclude that acyl groups in plastid 16:1Δ3t-PG contribute to TAG biosynthesis during
embryogenesis and that this requires PLIP1 activity.
Alignment of the predicted PLIP1 sequences found in land plants (Table 3.3 for species
and accession numbers) followed by phylogenetic analysis shows that PLIP1 proteins are
conserved in land plants (Figure 3.20), except for the orchid family, for which genomes of two
species are available in NCBI (Phalenopsis equestris and Dendrobium catentum. Interestingly,
presumed FAD4 homologs are also ubiquitous among land plants, except for the two
representatives of the orchid family (Figure 3.20). Calculation of the linear correlation coefficient
between the PLIP1 and FAD4 phylogenetic trees after extracting the 18S rRNA background from
26 sequenced land plant species (Pazos et al., 2005) resulted in a score of 0.971, suggesting that
these two proteins are coevolving. This supports our hypothesis that PLIP1 and FAD4 function in
the same pathway.

Overexpression of PLIP1 Increases PG Recycling and TAG Biosynthesis During
Embryogenesis
To determine whether increased turnover of plastid PG is responsible for increased TAG
biosynthesis during embryogenesis in the PLIP1-OX lines, we harvested siliques at nine days after
flowering from WT and PLIP1-OX1 plants and isolated the developing seeds. Seeds at this
developmental stage have robust lipid metabolism (Le et al., 2010; Bates et al., 2012). However,

98

siliques of the same age collected from PLIP1-OX1 plants were shorter than those from WT
(Figure 3.21 A), which raised the concern that embryos from PLIP1-OX1 and WT might be at
different developmental stages. However, upon closer examination, WT and PLIP1-OX1 had
nearly mature embryos with fully developed cotyledons and radicals (Figure 3.21 B), indicating
they were at a similar developmental stage and likely metabolically comparable. Therefore, we
performed [14C]-acetate pulse-chase labeling on the isolated developing seeds. Pulse time points
were collected after 20 and 60 minutes and are shown before time 0 of the chase start on the Xaxis, followed by three chase time points; a representative example of multiple repeats is shown
(Figure 3.21 C). Compared to PC and TAG, plastid lipids PG and MGDG were not highly labeled
during embryogenesis, likely due to their small pool size; therefore, an expanded view for PG is
shown in the lower graph of Figure 9C. PLIP1-OX1 had higher incorporation of label into PG and
increased turnover during the chase phase (Figure 3.21 C), as was observed for the equivalent
experiment done on leaves. The altered labeling patterns for PG and MGDG resembled those
observed in the leaf labeling assays (Figure 3.11 A and B). The most strongly labeled lipids were
TAG and PC, reflecting their end-product status (TAG) or large pool size (PC) in developing seeds.
However, the much smaller PG pool (mostly in the chloroplast as 16:1Δ3t-PG) seemed to be more
metabolically active in PLIP1-OX1 than in WT. Incorporation of label into PG during the pulse
under the conditions tested was faster than could be captured by the earliest sampling time points.
The rate and extent of incorporation into TAG was increased in the PLIP1-OX1 line, consistent
with increased total acyl group content in these seeds, while the PC pool was similarly labeled in
the WT and overexpression lines.

99

DISCUSSION
PLIP1 Prefers PG as Natural Substrate
Recombinant lipases are notoriously difficult to produce and to study in vitro. PLIP1 is no
exception, as its production in E. coli led to membrane degradation, which confirmed its general
lipase activity, but made PLIP1 challenging to purify (Figure 3.2). Furthermore, while recombinant
PLIP1 was specifically acting on the glyceryl sn-1 position, it is active on a range of polar lipids
found in plants and bacteria in vitro (Figure 3.5 C). Notably, it did not act on TAG and had very
little activity on the two glycolipids found in the chloroplast, DGDG and sulfolipid, but showed
high activity towards PG and MGDG. Because it was impractical to test the full spectrum of all
possible combinations of glycerolipid molecular species occurring in chloroplasts in vitro, we
explored the in vivo specificity of PLIP1 by overproducing the protein in chloroplasts, which we
determined was the location of PLIP1 in plant cells using multiple independent approaches (Figure
1). This allowed us to test PLIP1 lipase activity in a quasi-native environment with the caveat that
PLIP1 is normally not abundant in leaf chloroplasts and is more likely active in the chloroplasts
of developing embryos based on the gene’s expression profile and the reduced-function phenotype
visible in seeds (Figure 3.14). However, leaf chloroplasts are much more readily accessible for
assays than embryo chloroplasts, and we assumed that findings on PLIP1 activity would be
transferable between the two tissues, which we ultimately confirmed. Based on the in vivo analysis
of PLIP1, 18:3/16:1Δ3t-PG emerged as the most likely in vivo substrate for PLIP1, which was
corroborated in vitro using a native, leaf-isolated molecular species mixture of PG (Figure 3.11
D). It should be noted that in the overexpression lines, effects on MGDG and PC were observed
in addition to PG (Figure 3.6 B and Figure 3.7 B and C). This could be directly due to the activity
of PLIP1 on MGDG or caused by secondary effects related to acyl exchange and acyl transfer in

100

the case of PC, which is not in the thylakoid membranes and should not be directly accessible to
PLIP1.

The Location of PLIP1 Limits the Availability of Likely Substrates In Vivo
To explain the observed substrate preference of PLIP1 in vivo, one might invoke the
presence of factors in its native environment that are simply not present in vitro. Another, more
likely explanation might be the limited accessibility of certain lipid molecular species to PLIP1
due to its specific membrane location, assuming that specific membrane leaflets or lateral
subdomains have a specific lipid composition. We have some clarity now on the location of PLIP1,
its likely peripheral association with thylakoid membranes based on fractionation, chloroplast
import, and protease protection experiments, and its dual processing (Figure 3.1). Fractionation
showed that PLIP1 is associated with thylakoid membranes, while import and protease protection
assays were consistent with three possible suborganellar locations for PLIP1: stroma, thylakoid or
the stroma leaflet of the inner envelope membrane (Figure 3.1). PLIP1 is not predicted to contain
transmembrane domains, but it must be a peripheral membrane protein to gain access to its
substrate. Most likely, PLIP1 is a peripheral thylakoid protein, but it also has the ability to be
transiently free in the stroma or even has access to the inner envelope membrane. Double
processing of PLIP1, as observed, can be interpreted as first generating an intermediate during
protein import into the stroma, while the second processing possibly releases the majority of the
mature protein from the thylakoid fraction into the soluble stroma fraction. A conserved twinarginine motif is generally required for importing proteins into thylakoids (Robinson and Bolhuis,
2001; Goosens and van Dijl, 2016), but PLIP1 is missing a canonical motif, although it contains
two sets of twin-arginine in its transit peptide usually part of such a motif. Therefore, PLIP1 may

101

peripherally attach to the thylakoid membrane but is likely prevented from being further imported
into the thylakoid lumen.
For the likely substrate of PLIP1, 18:3/16:1Δ3t-PG, we only know that it is exclusively
present in chloroplasts, where the FAD4 desaturase required for its synthesis is located as well
(Gao et al., 2009). However, the presence of 18:3/16:1Δ3t-PG in a specific suborganellar
membrane, leaflet, or lateral membrane domain is not known. All we can conclude based on our
localization of PLIP1 is that 18:3/16:1Δ3t-PG must be present in the stroma leaflet of the thylakoid
or envelope membranes to be accessible to PLIP1.

PLIP1 Contributes to TAG Biosynthesis in Developing Embryos
PG is required for proper embryo development. The development of embryos in a pgp1
pgp2 double mutant affected in PG biosynthesis in chloroplasts, mitochondria and the endoplasmic
reticulum is delayed, and maturing seeds shrink during desiccation, resulting subsequently in
compromised germination (Tanoue et al., 2014). Originally, it was proposed that the chloroplastspecific molecular lipid species, 16:1Δ3t-PG, is critical for the function of the photosynthetic
membrane, but its complete replacement with 16:0-PG in the Arabidopsis fad4 mutant had only
mild effects on leaf photosynthesis (Browse et al., 1985; McCourt et al., 1985). What was possibly
neglected before was that the expression level of FAD4 is on the same order of magnitude in leaf
and developing seeds (Figure 3.19 A), but the level of 16:1Δ3t-PG in developing seeds was much
lower than that in leaves. The discovery that PLIP1 preferentially metabolizes 16:1Δ3t-PG during
seed development potentially explains the small 16:1Δ3t-PG pool, and points towards roles for this
lipid in seeds. In fact, decreased PLIP1 expression in T-DNA insertional lines reduced seed TAG
content without affecting growth or seed hormone levels (Figure 3.14 B), corroborating a possible

102

contribution of PLIP1 to seed TAG biosynthesis. The plip1 low-TAG seed phenotype is
recapitulated in the fad4 mutant (Figure 3.19 C), and further phylogenetic analysis suggests
coevolution of PLIP1 and FAD4 in land plants. These findings support a role for PLIP1 in seed
TAG biosynthesis and provide a possible function for this unusual lipid molecular species.
It should be noted that PLIP1-OX plants produced seeds with higher TAG content, but
40% fewer seeds, compared to wild type (Figure 3.17). Overall, seed oil production per plant was
reduced in line with the reduced growth of the PLIP1-OX plants, likely due to the activation of
oxylipin-induced defense pathways (Figure 3.9 and 3.10). During the labeling experiment with
developing seeds, saturating substrate levels were provided (Figure 3.21 C). Therefore, higher
carbon incorporation into TAG in PLIP1-OX seeds reflects an increased capacity for TAG
biosynthesis in individual embryos, in spite of the reduced plant growth and the decreased overall
seed yield of the plants (Figure 3.7 A and Figure 3.17 B).

PLIP1 Enables Channeling of Acyl Groups from Plastid 16:1Δ3t-PG to TAG at the ER
How can PLIP1, a lipase, be a component of a mechanism directing FAs synthesized in the
plastid into TAG lipid droplets in the cytosol during embryogenesis? A large body of evidence
suggests that PC is a critical precursor for TAG biosynthesis in developing seeds. As shown in
Figure 3.22, several pathways, including acyl exchange (Figure 3.22, reactions 1) and desaturation
of acyl groups on PC by FAD2/3 followed by transfer of 18:3 from the acyl-CoA pool to DAG
(Figure 10, reaction 4), head group exchange generating DAG with 18:3 acyl groups (Figure 3.22,
reaction 3), and transfer of acyl groups from PC to DAG by PDAT (Figure 3.22, reaction 5)
contribute to the incorporation of polyunsaturated FAs into TAG during seed development
(Dahlqvist et al., 2000; Bates et al., 2012; Li-Beisson et al., 2013). However, even in the rod1

103

lpcat1 lpcat2 triple mutant carrying the strongest known alleles at each locus, which in
combination should completely disrupt acyl editing and head group exchange, the capacity of
seeds to produce 18:3-containing TAGs is only reduced by half (Bates et al., 2012). Therefore,
other compensatory mechanisms likely exist for incorporating polyunsaturated FAs into ER lipids
and TAGs that could involve PLIP1 or PDAT. We propose that PLIP1 is part of a chloroplast acyl
editing mechanism augmenting the supply of polyunsaturated FAs to the cytosolic acyl-CoA and
PC pools, as depicted in Figure 3.22, reaction 2. This is particularly apparent whenever PLIP1 is
highly abundant, as the turnover of plastid PG accelerates (Figure 3.11 and Figure 3.21). Acyl
group flux is increased from PG to PC in PLIP1 overexpression lines, and these acyl groups
eventually end up in TAGs, which is evident from the pulse-chase labeling experiments (Figure
3.11 and 3.21). Partial restoration of the low 18:3-PC lipid phenotype of the fad3-2 mutant by
overexpression of PLIP1 in mesophyll tissues and seeds (Figure 3.11 E and 3.18) supports a role
for PLIP1 in plastid acyl export. The resemblance of acyl compositions of PC and TAG in seeds
is indicative of the precursor-product relationship between these two lipids (Figure 3.14), but the
detailed mechanism needs further investigation. In vegetative and seed tissues, overexpression of
PLIP1 results in a marked decrease in 18:2 and increase of 18:1 in PC, a phenotype reminiscent
of the fad2 mutant. This suggests that the elevated 18:3 flux from plastids out competes the
incorporation of unsaturated fatty acids by the FAD2/3 pathway. Consistent with this assumption,
in the plip1 mutant, which lacks the PLIP1-dependent plastid acyl export pathway, one would
expect that the ER-based FAD2/3 mechanism for the incorporation of unsaturated FAs into PC
and TAG to be more active. In fact, the decreased 18:1 and increased 18:3 contents in PC and total
acyl groups (reflecting mostly TAG), as observed in the plip1 mutant seeds (Figure 3.14 E and G),
corroborate this prediction. The FAD2/3 desaturation pathway counteracts the reduced 18:3 export

104

in the plip1 mutants but does not affect loss of plastid-derived acyl group flux, leading to the
observed decrease in seed TAG. The reversibly compensatory activity of the PLIP1-mediated
plastid export of 18:3 acyl groups and the ER-based FAD2/3 desaturation pathway explains the
seemingly paradoxical observation that 18:3 contents are high in both the plip1 mutants and the
PLIP1 overexpression lines.
It should be noted that nearly 85% of the seed TAG 18:3 is dependent on FAD3, and the
contribution of PLIP1 to seed 18:3 reflects the finding that the PLIP1-mediated pathway
contributes about 10% of the acyl groups in TAG in seeds, even though overexpression of PLIP1
is able to significantly increase 18:3 content in the fad3-2 seeds (Figure 3.18). This may seem like
a minor contribution. However, even the ablation of the presumed key enzyme of TAG
biosynthesis, DGAT1, leads to only a 20-40% reduction in seed oil content (Katavic et al., 1995;
Routaboul et al., 1999). Furthermore, in a PDAT-loss of function mutant affecting a pathway
parallel to DGAT (Figure 3.22, reactions 5 and 4, respectively), seeds show no changes in oil
content nor composition (Mhaske et al., 2005), even though a double mutant deficient in these two
pathways exhibits up to 80% reduction in seed oil content and a loss in seed viability. These results
indicate that other parallel mechanisms exist to account for the remaining small fraction of TAG.
The regulatory network governing the different TAG biosynthesis pathways is complex, allowing
for redundancy and compensatory mechanisms. The PLIP1-mediated pathway described here
could represent one of the alternative mechanisms for TAG biosynthesis. At the present time, the
lack of a PLIP1 loss-of-function mutant prevents us from more accurately quantifying the
contribution of a PLIP1-dependent pathway of polyunsaturated FA incorporation into TAG under
different conditions or in different loss-of-function mutants. Despite repeated attempts, we have
not yet succeeded in isolating a homozygous PLIP1 loss-of-function mutant using targeted gene

105

disruption methods. Thus, we cannot rule out that PLIP1 has additional essential functions in
embryo metabolism that still need to be discovered.

PLIP1 Takes Part in Acyl Group Export from Chloroplasts
The hypothesis outlined above (Figure 3.22) also implies that PLIP1 activity leads to acyl
export from the plastid. Assuming that PLIP1 acts at the stroma surface of the thylakoids or the
inner envelope membrane, additional chloroplast proteins are likely necessary to direct acyl groups
from chloroplast PG into TAGs. Recently, chalcone isomerase-like chloroplast proteins were
shown to be FATTY ACID BINDING PROTEINS (FAP), which may be associated with
chloroplast fatty acid export (Ngaki et al., 2012). The expression pattern of FAPs resembles that
of PLIP1, and all FAPs are located in the stroma of chloroplasts. One FAP, FAP1, shows high
proclivity for binding 18:3. Therefore, it seems possible that 18:3 released by PLIP1 from
18:3/16:1Δ3t-PG is bound by FAP, thereby sequestering it to avoid cytotoxicity of free FAs or to
mediate FA transfer to the chloroplast envelope membrane. Another protein possibly involved is
FATTY ACID EXPORT1 (FAX1), a likely acyl group or FA transporter of the inner envelope
membrane of plastids (Li et al., 2015). In the fax1 mutant, of the four thylakoid lipids, PG levels
are increased the most, especially 18:3/16:1Δ3t-PG, and the levels of PC are reduced. Also, TAG
biosynthesis, especially its polyunsaturated FA content, correlates with the presence of FAX1 in
reproductive tissues. Hence, it seems possible that PLIP1, FAP and FAX1 work together to channel
a small fraction of plastid synthesized acyl groups through 16:1Δ3t-PG into PC outside the plastid
and ultimately into TAG during seed development.
Movement of de novo synthesized lipid groups through the chloroplast membrane lipid
pool has been observed in Chlamydomonas, in which PGD1 is a lipase specific for newly

106

synthesized 18:1/18:1-MGDG, while 18:3/16:4 MGDG is resistant to its activity (Li et al., 2012).
In this case, PGD1 expression is induced following N-deprivation and participates in the
channeling of acyl groups into TAG biosynthesis under these conditions. Although plants
synthesize TAG in vegetative tissues under stress (Moellering et al., 2010), they generally produce
bulk TAG in developing embryos. PLIP1 is too distantly related to PGD1 to be an orthologue, and
Chlamydomonas does not contain PC, while Arabidopsis lacks 18:3/16:4-MGDG. However, both
lipases point towards a common theme, the need for channeling of plastid-synthesized acyl groups
through the chloroplast lipid pool prior to incorporation into extraplastidic TAGs. The specific
substrate selectivity of these two lipases also partially explains the existence of unusual molecular
species of chloroplast lipids, 18:3/16:4 MGDG in Chlamydomonas and 18:3/16:1Δ3t-PG in
Arabidopsis and most other plants and algae. It seems likely that unusual acyl groups tag specific
molecular species for specific purposes. In the case of 18:3/16:4 MGDG in Chlamydomonas, it is
tagged as a structural thylakoid membrane lipid resistant to PGD1, while in the case of
18:3/16:1Δ3t-PG in Arabidopsis, it is the preferred substrate for PLIP1, leading to 18:3 acyl export,
rather than having a specific function related to photosynthetic light capture and conversion as
previously assumed. Chlamydomonas also contains 16:1Δ3t-PG, and genomes of plants and algae
encode many more potential plastid-targeted lipases. Therefore, it is likely that acyl hydrolysis
catalyzed by specific plastid lipases and their respective native substrates is a common process for
maintaining photosynthetic membrane lipid homeostasis while enabling the exchange and export
of acyl groups for the synthesis of extraplastidic lipids or as precursors for retrograde signaling
molecules, a possibility that still begs to be explored.

107

ACKNOWLEDGEMENTS
This work was done in collaboration with several colleagues. John E. Froehlich
conducted the chloroplast import assay in Figure 3.1. Agnieszka Zienkiewicz assisted with the
confocal microscopy experiment in Figure 3.1. I would also like to thank Dr. Danny Schnell,
Michigan State University, for providing antibodies against LHCb1, Dr. Gregg Howe for
providing the coi1-30 seeds, Dr. Dan Jones for providing the ABA-d6 internal standard and Dr.
Anthony Schilmiller for assisting with plant hormone and Arabidopside measurement. We would
also like to thank Dr. Rebecca Roston, University of Nebraska, Lincoln, and Dr. Barb Sears,
Michigan State University for their advice on conducting the experiments or critically reading
the manuscript. This work was supported by the Division of Chemical Sciences, Geosciences
and Biosciences, Office of Basic Energy Sciences of the United States Department of Energy
Grants DE-FG02-98ER20305 and DE-FG02-91ER20021, the US Department of Energy-Great
Lakes Bioenergy Research Center Cooperative Agreement DE-FC02-07ER64494, and by
AgBioResearch, Michigan State University.

108

APPENDIX

109

Primer name

Sequence 5’ – 3’

PEG_PLIP1 F

CACCATGGCGTTTAATACGGCTAT
G

For pEarlyGate 101/4

PEG101_PLIP1
R

GACACGTGTCATGATCTCCTCGG

For pEarlyGate 101

TCGGATCCATGGCGTTTAATACGG
CTATG
GACTCGAGTTAGACACGTGTCATG
PLIP1_XhoI R
ATCTCC
ATGTATATCTCCTTCTAAAGTAAA
pET41a_His del F
CAAA
pET41a_His del
ATGGCGTTTAATACGGCTATG
R
AGTCTCTCATTAATAGTGAATTTG
PLIP1_S422A F
ATGCTTATC
PLIP1_S422A R
GCCTCCAAGAGCATGACCCGTG
GAGCCTTTTCGTGTAATTATCCTGA
PLIP1_D483A F
CCA
PLIP1_D483A R GTGGGACGATAGCTCTATGCATCA
PLIP1_qPCR F
AGTTCTATAATCCCAAGTCCGA
PLIP1_qPCR R
CTCCTTATCTCAAGCAGCCT
TIP-41_like
GTGAAAACTGTTGGAGAGAAGCA
qPCR F
A
TIP-41_like
TCAACTGGATACCCTTTCGCA
qPCR R
PDF2 qPCR F
TAACGTGGCCAAAATGATGC
PDF2 qPCR R
GTTCTCCACAACCGCTTGGT
BamHI_PLIP1 F

LBb1.3

ATTTTGCCGATTTCGGAAC

plip1-1 LP
plip1-1 RP
plip1-2 LP
plip1-2 RP
fad3-2 genotype F
fad3-2 genotype
R
VSP1 F
VSP1 R
LOX2 F
LOX2 R
VSP2 F
VSP2 R

AGATTCTAGCGGAGCTTGGTC
GCCTCTTCAAACCAAATCTCC
TTATTACCGGAGCGACAACAC
TCCAATAACGGTTAAGCAACG
GTCACGATGAGAAGTTGCCTTGG
CAATGTCGTGATGAATGTTGTTAA
AGAAT
AGAGCTGGAGCTGGTGTTGTTAAG
GAACACCCATTCCGGTAACACCAT
GGGCGTACTGGTCGTGGTTA
TCCCGAGTTCCAAGAGGTTTT
GCAAGTGGTGTACAAGTCAAAGGT
ATCCTCAACCAAATCAGCCCATTG

Table 3. 1. Primers used in this study.
110

Purpose

For the pET41a to produce
PLIP1

pET41a construct for
chloroplast import

S422A point mutation

D483A point mutation
For qPCR analysis of PLIP1
expression
qPCR reference gene

qPCR reference gene
For genotyping of plip1
mutants
For genotyping of plip1-1
For genotyping of plip1-2
For genotyping of fad3-2
For qPCR analysis of VSP1
(At5g24780) expression
For qPCR analysis of LOX2
(At3g45140) expression
For qPCR analysis of VSP2
(At5g24780) expression

Table 3. 1. (cont’d)
JAZ1 F
JAZ1 R
JAZ10.1-3 F

GCACCGCTAATAGCTTAGCCAAGA
TCTGTCAATGGTGTTGGGGAGGAT
AGAGCTGGAGCTGGTGTTGTTAAG

JAZ10.1-3 R

GAACACCCATTCCGGTAACACCAT

CYP94C1 F

TCTCACTCTTTCACTTACTCTTCC

CYP94C1 R

CGTTTGATGGATTCGCTGTG

111

For qPCR analysis of JAZ1
(At1g19180) expression
For qPCR analysis of
JAZ10.1, 10.2 and 10.3
(At5g13220) expression
For qPCR analysis of
CYP94C1 (AT2G27690)
expression

Genotype

Acyl Groups (µg/mg FW)

Col-0

3.48 ± 0.067

plip1-1

3.51 ± 0.076

plip1-2

3.51 ± 0.063

EV control

3.55 ± 0.055

PLIP1S422A-OX1

3.53 ± 0.055

PLIP1S422A-OX2

3.59 ± 0.049

PLIP1-OX1

3.56 ± 0.062

PLIP1-OX2

3.54 ± 0.063

PLIP1-OX3

3.56 ± 0.067

Table 3. 2. Leaf acyl group contents in different genotypes.
Acyl group contents were determined by measuring total leaf fatty acid methyl esters. Plants
were grown on soil for 4 weeks. Four independent samples were averaged, and the SD is
indicated. FW, fresh weight.

112

Organism

PLIP1

FAD4

Arabidopsis thaliana

NP_191727.2

NP_194433.1

Arabidopsis lyrata

XP_002876633.1 XP_002869582.1

ADBK01000671

Eutrema salsugineum

XP_006402456.1 XP_006413113.1

AHIU01024615

Brassica rapa

XP_009138759.1 XP_009109956.1

AENI01003060

Manihot esculenta

OAY49032.1

AB233568

Populus trichocarpa

XP_002301404.2 XP_006370685.1

AARH02003126

Cucumis sativus

KGN44530.1

XP_004152938.1

ACHR02000003

Glycine max

XP_014627545.1 XP_003551889.1

ACUP02007833

Citrus sinensis

KDO66297.1

AJPS01002884

Solanum lycopersicum

XP_004231212.1 XP_004244133.1

AB971540

Zea mays

NP_001183891.2 XP_008662704.1

AC150267

Brachypodium distachyon

XP_010233458.2 XP_003571398.1

ADDN02000002

Selaginella moellendorffii

XP_002982750.1

XP_002962129.1

ADFJ01002076

Gossypium raimondii

XP_012437750.1 XP_012460522.1

ALYE01001348

Elaeis guineensis

XP_010913778.1 XP_010908683.1

ASJS01014373

Oryza sativa

XP_015630701.1 XP_015648482.1

AF069218

Ananas comosus

XP_020098248.1 XP_020103087.1

AP014632

Physcomitrella patens

XP_001766893.1 XP_001752601.1

AB251495

Capsella rubella

XP_006292392.1 XP_006285104.1

KR029093

Capsella grandiflora

XP_006292392.1 XP_006285104.1

KR029092

OAY28524.1

XP_006452281.1

18S rRNA
AC004708

Table 3. 3. Sequences used in the PLIP1 and FAD4 phylogenetic analyses.
For PLIP1 and FAD4 protein sequences, accession numbers in NCBI GenBank are listed for the
corresponding organism. For the 18S rRNA sequences, accession numbers in the SILVA rRNA
database are listed.

113

Table 3. 3. (cont’d)
Ricinus communis

EEF39083.1

Theobroma cacao

XP_017970999.1 EOY12432.1

AF207040

Malus domestica

XP_017185317.1 XP_008385990.1

ACYM01009980

Camelina sativa

XP_010413364.1 XP_010433413.1

GABL01042315

Sorghum bicolor

KXG32911.1

XP_002443967.1

ABXC02001831

Eucalyptus grandis

XP_010053810.1 XP_010060414.1

AUSX01001791

114

XP_002522972.1

AASG02002472

Figure 3. 1. Subcellular localization of PLIP1 in Arabidopsis.
(A) Subcellular localization of PLIP1-YFP in leaf mesophyll cells of 3-week-old Arabidopsis Col0 transformed with PLIP1-YFP driven by the 35S promoter or empty vector (EV) control using
confocal laser scanning microscopy. Chlorophyll autofluorescence is shown in red, and YFP
fluorescence is shown in green. Overlay of chlorophyll and YFP is shown as well (Merge).
Representative images from one experiment are presented. Scale bars: 30 µm.
(B) PLIP1 enrichment in chloroplast fractions analyzed by immunoblotting. Intact and
subfractionated chloroplasts were prepared using 4-week-old Arabidopsis (Col-0) plants grown on
MS medium. Equal amounts of protein of leaf tissues from the whole plant (wp), intact chloroplasts
(chl), thylakoid (thy) and stroma (str) were separated by SDS-PAGE or further subjected to
immunoblotting analysis using an antibody against PLIP1S422A, a non-functional mutant of PLIP1.
Immunoblotting was used to detect marker proteins BiP2 (endoplasmic reticulum) and LHCb1
(thylakoid). For protein loading, 12 µg per fraction were loaded for PLIP1; 2 µg per fraction for
BiP2 and LHCb1.
115

Figure 3. 1. (cont’d)
(C) SDS-PAGE Coomassie Brilliant Blue staining was used to detect Rubisco large subunit
(stroma) and light-harvesting chlorophyll a/b-binding protein (LHCP) (thylakoid), which were
used as markers. Numbers indicate protein molecular mass in kDa. For protein loading, 12 µg per
fraction were loaded.
(D) Chloroplast import experiments with labeled PLIP1 and control protein FtsH8. Chloroplasts
were treated with (+) or without (−) trypsin. Total chloroplast membranes (P) or soluble (S)
fractions were analyzed by SDS-PAGE followed by fluorography. TP, translation products; p,
precursor; i, intermediate; m, mature form; MW, molecular weight markers.

116

Figure 3. 2. Recombinant PLIP1 has lipase activity.
(A) Thin layer chromatographic analysis of polar (left) and neutral (right) lipids in E. coli
containing a 6×His-PLIP1 expression construct or empty vector control (EV) at 6 h following
induction. FFA, free fatty acid; O, origin of sample loading; PE, phosphatidylethanolamine; PG,
phosphatidylglycerol. TLC plates were stained with iodine vapor.
(B) Mutation of the PLIP1 active site motif. Lipid extracts of E. coli cultures 6 h after induction
expressing 6×His-PLIP1 or two point mutation alleles, 6×His -PLIP1S422A or 6×His-PLIP1D483A,
or containing an empty vector control (EV) were analyzed by thin layer chromatography to detect
FFA products (top panel). Protein extracts were analyzed for protein production using an antibody
against the 6×His tag.
(C) SDS-PAGE analysis of purified PLIP1 and PLIP1S422A proteins. Loading was 5 µg per lane
for both samples. SDS-PAGE separated proteins were stained by Coomassie Brilliant Blue (left)
or detected by immunoblotting with an antibody raised against PLIP1S422A. Numbers indicate
protein molecular mass in kDa. 6×His-PLIP1 and 6×His-PLIP1S422A are indicated by the arrow.
(D) Thin-layer chromatogram of products of a representative in vitro lipase reaction using PC with
wild type (PLIP1 + PC) and the mutant enzyme (PLIP1S422A + PC). Substrate without enzyme
(Buffer + PC) and enzyme without substrate (PLIP1) were included as controls. PC,
phosphatidylcholine. O, origin of sample loading. Lipids were visualized by iodine vapor staining.

117

Figure 3. 3. Alignment of PLIP1 with class 3 lipase protein sequences in the NCBI
conserved domain database (CDD).
The alignment results were obtained by BLAST analysis of the PLIP1 protein sequence against
the CDD database using CD-search (https://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi) with
default settings. Reference protein names are represented by either Protein Data Bank ID (e.g.,
1TIA and 3TGL) or NCBI GI number. The conserved catalytic triads are highlighted in yellow.

118

Figure 3. 4. Time course of PLIP1 lipase activity with phosphatidylcholine (PC) as
substrate.
(A) Results of thin layer chromatography of substrate PC and lyso-PC product at different time
points. O, origin of sample loading. 120 µg substrate 18:1/18:1-PC was provided to 1 µg
recombinant PLIP1 and the lipase assay was stopped at the indicated time points. Lipids were
subsequently extracted and analyzed by TLC and stained with iodine vapor.
(B) Fraction of PC degradation at different time points, as indicated in (A). Fatty acid methyl esters
of acyl groups of both PC and lyso-PC at each time point were analyzed by liquid gas
chromatography. The fraction of PC degradation is calculated as 2 (molarity of lyso-PC acyl
groups) / (2 (molarity of lyso-PC acyl groups) + (molarity of PC acyl groups)) *100.

119

Figure 3. 5. Specificity of PLIP1 in vitro activity.
(A) Gas-liquid chromatograms of methyl esters derived from commercial PC substrates or lysoPC fractions from PLIP1 lipase reactions with different PC substrates. 15:0 was used as an internal
standard.
(B) PLIP1 lipase activity on commercial PC substrates (carbon number: double bond number; sn1/sn-2) with different degrees of saturation of the sn-1 acyl groups. PC containing 18:0/18:1 and
18:1/18:1 and PC containing 18:0/18:2 and 18:2/18:2 were compared, respectively. n=4, ± SD.
Student’s t-test was applied (**indicates p<0.01).
(C) PLIP1 activity on different substrates. For each PLIP1 lipase reaction, 60 µg lipids and 0.5 µg
protein were used. The reactions were incubated at ambient temperature (~22 °C) for 1.5 h, which
120

Figure 3. 5. (cont’d)
was still during the linear portion of the reaction time course for PC (Supplemental Figure 2).
PLIP1S422A was included as a negative control and is shown in the top panel. All lipids contained
two oleic acids (18:1), except MGDG, DGDG, and SQDG, which were isolated from plants, and
PI, which was isolated from bovine liver. n=3-4 for each substrate, ± SD. DGDG,
digalactosyldiacylglycerol; MGDG, monogalactosyldiacylglycerol; PA, phosphatidic acid; PC,
phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PI,
phosphatidylinositol; PS, phosphatidylserine; SQDG, sulfoquinovosyldiacylglycerol; TAG,
triacylglycerol.

121

Figure 3. 6. Analysis of polar lipids in the leaves of 4-week-old, soil-grown PLIP1-OX and
empty vector (EV) control plants.
(A) Relative abundance of major polar lipid classes in PLIP1-OX and EV control lines.
(B-E) Relative acyl group composition of MGDG, DGDG, SQDG, PI and PE supplemental to data
shown in Figure 4B and C. Leaf samples harvested from one plant were analyzed as one biological
repeat; n=3, ± SD. Student’s t-test was applied comparing the EV control with each PLIP1-OX
line (* indicates p<0.05; ** p<0.01). DGDG, digalactosyldiacylglycerol; MGDG,
monogalactosyldiacylglycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG,
phosphatidylglycerol; PI, phosphatidylinositol; SQDG, sulfoquinovosyldiacylglycerol. Acyl
groups with a molar percentage less than 0.5% were omitted.

122

Figure 3. 7. Growth and lipid phenotypes of PLIP1-overexpression plants.
(A) Growth of 4-week-old soil-grown plants. Arabidopsis wild-type plant (WT), one empty vector
control line, two PLIP1S422A-OX and three PLIP1-OX overexpression lines are shown. Scale bar:
5 cm.
(B) and (C) Relative acyl composition of phosphatidylglycerol (PG) and phosphatidylcholine (PC)
in PLIP1-OX and empty vector (EV) control lines. Mature leaf lipids were extracted and analyzed
from the plants shown in (A). Leaf samples harvested from one plant were taken as one biological
repeat; n=4, ± SD. Student’s t-test was applied to compare the EV control with each PLIP1-OX
line, respectively (* indicates p<0.05; ** indicates p<0.01).

123

Figure 3. 8. Expression of jasmonic acid-responsive genes.
The expression levels of the indicated genes were determined by quantitative RT-PCR and
expressed relative to UBQ10 gene expression. Analysis of JAZ10 includes JAZ10.1, 10.2 and 10.3,
but not the 10.4 splicing isoform. RNA was extracted from the leaves of 3-week-old soil-grown
plants. Leaves harvested from one plant were analyzed as one biological repeat; n=3, ± SD.
Student’s t-test was applied comparing the EV control with each PLIP1-OX line (** indicates
p<0.01).

124

Figure 3. 9. Quantification of plant hormones and Arabidopsides in leaves of the wild type
(WT) and PLIP-OX1.
(A) Plants were grown on soil for three weeks before the leaves were harvested for hormone
extraction and quantification by LC-MS/MS. Leaves harvested from one plant were analyzed as
one biological repeat; n=3, ± SD. Student’s t-test was applied (**indicates p<0.01).
(B) Relative amounts of the major Arabidopsides in wild-type (WT) and PLIP-OX plant leaves.
The indicated Arabidopside level was normalized to the level of 18:3/18:3digalactosyldiacylglycerol, which did not change in WT or PLIP-OX1 plants. Leaves harvested
from one plant were analyzed as one biological repeat; n=3-4, ± SD. Student’s t-test was applied
(**indicates p<0.01).
125

Figure 3. 10. Phenotypes of PLIP1-OX and coi1 plants.
(A) Image of the 4-week-old plants grown on soil and the rosette diameter of the indicated plants.
n=6, ± SD, one-way ANOVA with post-hoc Turkey HSD test was applied. Rosette diameters
indicated by different letters (a, b and c) are significantly different (p<0.01).
(B) Relative acyl group compositions of leaf phosphatidylglycerol (PG) and phosphatidylcholine
(PC) in the indicated plants in (A). Leaf samples harvested from one plant were treated as one
biological repeat; n=3, ± SD. Student’s t-test was applied to compare WT with each of the
remaining genotypes (** indicates p<0.01).

126

Figure 3. 11. PLIP1 precursor–product relationships in PLIP1-overexpression plants.
(A) and (B) In vivo pulse-chase acetate labeling of lipids in wild-type and PLIP1-OX1 plants. The
length of the [14C]-acetate labeling pulse was 60 min (A), after which medium was replaced with
non-labeled free acetate to initiate the chase with a duration of three days (B). The fractions of
label in all polar lipids are given as percentages of total incorporation of label in polar lipids.
127

Figure 3. 11. (cont’d)
Experiments were repeated three times with similar results, and one representative result is shown.
MGDG, monogalactosyldiacylglycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine;
PG, phosphatidylglycerol.
(C) Activity of purified recombinant PLIP1 on PC with different sn-2 acyl groups. PC containing
16:0/18:0, 16:0/18:1, and 16:0/18:2 were used as substrates. n=4, ± SD. Student’s t-test was
applied (** indicates p<0.01).
(D) PLIP1 enzyme activity preference for molecular species of phosphatidylglycerol isolated from
tobacco leaves. Acyl groups of lyso-phosphatidylglycerol are shown as molar percentages of total
acyl groups at any given time point. Experiments were repeated three times with similar results
and data from one representative experiment are shown.
(E) Relative acyl composition of PC in wild-type (WT), fad3-2 and fad3-2, PLIP1-OX1 plants.
Leaf lipids were extracted and isolated by TLC and fatty acid methyl esters derived from the lipids
were analyzed by GC. Mature leaf samples harvested from one plant were taken as one biological
repeat; n=4, ± SD. Student’s t-test was applied (** indicates p<0.01).

128

Figure 3. 12. Effect of sn-2 acyl 16:1Δ3t on PLIP1 function.
(A) Change in the molar percentage of native phosphatidylglycerol (PG) isolated from 4-week-old
tobacco leaves and lyso-PG in a time course of in vitro PLIP1 lipase reaction.
(B) PLIP1 enzyme activity preference for molecular species of PG isolated from tobacco leaves.
Acyl groups of PG are shown as molar percentages of total acyl groups at any given time point.
These data are supplemental to the acyl group analysis of lyso-PG shown in Figure 6D.
Experiments were repeated three times with similar results, and data from one representative
experiment are shown.

129

Figure 3. 13. Lipid analysis of offspring from a cross between fad3-2 and PLIP1-OX1.
(A) Sequencing of the FAD3 locus from the fad3-2 mutant plant revealed a point mutation of
G875A, which leads to a G292E mutation in the FAD3 protein.
(B) Genotyping of fad3-2 and PLIP1-OX1 plants. PCR products amplified from plant genomic
DNA were subjected to EcoRI digestion. The mutated fad3-2 fragment was susceptible to
digestion. Hetero, heterozygous; Homo, homozygous plants.
(C-F) Relative abundance of major polar lipid classes and relative acyl composition of other
individual glycerolipids supplemental to Figure 6E. n=3-4 for each substrate, ± SD. Student’s ttest was applied (** indicates p<0.01). For lipid abbreviations, refer to the legend of Figure 3.6.
Acyl groups with a molar percentage less than 0.5% were omitted.

130

Figure 3. 14. Effect of PLIP1 on seed oil biosynthesis and germination.
(A) PLIP1 transcript levels in different tissues or developmental stages determined by quantitative
PCR. Expression levels were normalized to those lowest in 4-week-old leaf tissues and shown as
relative fold changes. n=3 for each tissue, ± SD.
(B) Total acyl group content in dry seeds of wild type (WT), plip1-1, plip1-2, PLIP1-OX1 and
PLIP1-OX2. 30 seeds were analyzed in bulk for each repeat; n=5, ± SD.
(C) Weight of the seeds shown in (B). 200 seeds were analyzed for each repeat; n=4-7, ± SD.
(D) Germination of WT, plip1-1, plip1-2, PLIP1-OX1 and PLIP1-OX2 seeds. The fraction of
seeds showing radical emergence was determined 40 h after stratified seeds were sown on MS
medium. 100 seeds were used for each repeat, n=3, ± SD.
(E) Relative acyl group composition of dry seeds shown in (B). Acyl groups with a molar
percentage less than 0.5% were omitted. 30 seeds were analyzed in bulk for each repeat; n=5, ±
SD.
(F) and (G) content and compositional analysis of phosphatidylglycerol (PG) and
phosphatidylcholine (PC) in developing seeds isolated from WT, plip1-1 and PLIP1-OX1 siliques
harvested seven days after flowering. Molar percentages of PG and PC in lipids are shown in the
inserted figures. 100 µL equivalent volume of developing seeds was used for each repeat. n=3, ±
SD. Where appropriate, Student’s t-test was applied to compare WT values with those each other
line, respectively (* indicates p<0.05; ** indicates p<0.01).
131

Figure 3. 15. Genotyping, phenotypic and lipid analysis of the plip1 mutants.
(A) Genotyping of the plip1 mutants. Top panel, positions of T-DNA insertions of plip1-1 and
plip1-2 in the PLIP1 gene. Gray box, untranslated regions; black box, exons; black lines, introns.
Bottom panel, genotyping of the homozygous plip1 alleles.
(B) Growth phenotypes of 4-week-old wild-type (WT) plants and two plip1 mutants grown on soil
(scale bar: 5 cm). Numbers indicate the relative PLIP1 transcript levels in 2-week-old seedlings of
wild-type (WT), plip1-1 and plip1-2 plants grown on MS medium. Leaves harvested from one
plant were analyzed as one biological repeat; n=3-4, ± SD.
(C-J) Relative abundance of major polar lipid classes and relative acyl group composition of the
major glycerolipids from leaves of 4-week-old soil-grown wild-type (WT), plip1-1 and plip1-2
plants. Leaves harvested from one plant were analyzed as one biological repeat; n=3-4, ± SD.
DGDG,
digalactosyldiacylglycerol;
MGDG,
monogalactosyldiacylglycerol;
PC,
phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PS,
phosphatidylserine; SQDG, sulfoquinovosyldiacylglycerol; TAG, triacylglycerol.

132

Figure 3. 16. Hormone quantification in plip1 and PLIP1-OX1 dry seeds.
100 mg dry seeds of each genotype were used for hormone extraction as one biological repeat;
n=4, ± SD. Student’s t-test was applied to compare the WT with every other genotype (** indicates
p<0.01).

133

Figure 3. 17. Growth phenotypes of mature plip1 mutant and PLIP1-OX plants.
(A) Growth phenotypes of 8-week-old wild type (WT), two plip1 mutants, empty vector control
(EV), and two PLIP1-OX plants.
(B) Seed yield measurement from plants in (A). Seeds harvested from four plants grown in the
same pot were used in each repeat; n=3, ± SD. Student’s t-test was applied to compare the WT
with every other genotype (** indicates p<0.01).

134

Figure 3. 18. Relative composition of total acyl groups in the wild type (WT), fad3-2, and
fad3-2, PLIP1-OX1 dry seeds.
30 seeds per replicate, n=6 ± SD. Student’s t-test was applied to compare the WT with every other
genotype (** indicates p<0.01).

135

Figure 3.19. Analysis of fad4-2 and fad4-3 plants.
(A) Absolute expression levels of FAD4 in leaves and developing seeds at different stages. Data
were extracted from the Arabidopsis eFB browser. Average values from triplicates are shown, and
the error bars represent SD.
(B) Leaf phosphatidylglycerol (PG) acyl group composition of 3-week-old MS medium-grown
wild type (WT), fad4-2 and fad4-3 plants. Leaves harvested from one plant were pooled as one
biological repeat; n=3-4, ± SD. n.d., not detected.
(C) Total acyl group content in dry seeds of WT, fad4-2 and fad4-3. 30 seeds were used for each
repeat; n=5, ± SD.
(D) Seed weight and yield measurements of WT, fad4-2 and fad4-3 plants. For seed weight
measurement, 200 seeds were used for each repeat, n=3, ± SD. For seed yield measurement, seeds
harvested from 4 plants grown in the same pot were used as one repeat, n=3, ± SD.
(E) Relative acyl group composition of seed lipid extracts from of WT, fad4-2 and fad4-3. 30 seeds
were used for each repeat, n=5, ± SD. Acyl groups with a molar percentage less than 0.5% were
omitted. Where appropriate, Student’s t-test was to compare the WT and each fad4 mutant (*
indicates p<0.05; ** p<0.01).
136

Figure 3. 20. Phylogenetic trees of PLIP1 and FAD4 in land plants.
The phylogenetic trees were built using the Maximum Likelihood method with PLIP1 or FAD4
homologous sequences from 26 land plant species with sequenced genomes. All sequence
accession numbers are shown in Supplemental Table 2. Bootstrap values (based on 1000
replicates) are indicated at the tree nodes. The scale measures evolutionary distances in
substitutions per amino acid.

137

Figure 3. 21. Phenotypic analysis of PLIP1-OX1 embryos.
(A) Morphology of the wild-type (WT) and PLIP1-OX1 siliques nine days after flowering. Scale
bar: 0.5 cm. The numbers indicate the length of siliques. n=9-12, ± SD. Student’s t-test was applied
(** indicates p<0.01).
(B) Differential interference contrast images of embryos isolated from siliques of WT and PLIP1OX1 plants. Scale bars: 50 µm. Representative images are shown.
(C) Pulse-chase labeling of developing embryos isolated from siliques of wild-type (WT) and
PLIP1-OX1 plants. 100 µL equivalent volume of developing seeds were used for each time point.
The first two time points represent the labeling pulse. Embryos were transferred to unlabeled
medium after one hour. Values represent the fraction of label in select individual lipids compared
to label in total lipids. The top panels show four lipids, as indicated. The lower panels show PG
and MGDG again, but on an expanded scale. Experiments were repeated three times on PLIP1OX1 seeds and one time on PLIP1-OX2 seeds, with similar results. One representative result is
shown.
MGDG,
monogalactosyldiacylglycerol;
PC,
phosphatidylcholine;
PG,
phosphatidylglycerol; TAG, triacylglycerol.
138

Figure 3. 22. Hypothesis for the role of PLIP1 in triacylglycerol biosynthesis.
The left panel depicts the wild type (WT), the middle panel the PLIP1-overexpression lines, and
the right panel the plip1 mutant. The thickness of the arrows indicates the relative fluxes in the
three different lines. Reactions or sets of reactions are numbered as follows: 1. In WT (left panel)
acyl exchange on phosphatidylcholine (PC) involving desaturation of acyl groups by FAD2/3
provides one mechanism to introduce polyunsaturated fatty acids (FAs) into phosphatidylcholine
(PC). 2. A parallel mechanism to introduce PUFAs into PC involves PLIP1. In the chloroplast,
PLIP1 hydrolyzes 18:3/16:1Δ3t-phosphatidylglycerol (PG) and other 16:1Δ3t-phosphatidylglycerol
species (not shown here) at the sn-1 glyceryl position and releases 18:3 (carbon: double bonds) or
other acyl groups at the sn-1 position. The released acyl group is exported to the Endoplasmic
Reticulum and incorporated into the acyl-CoA pool and PC. 3. A head group exchange mechanism
leads to diacylglycerol (DAG) formation from PC containing PUFAs. 4. Triacylglycerol (TAG),
which accumulates in lipid droplets (LDs), is formed by the action of DAG-acyltransferases, which
can introduce additional acyl groups into DAG from the acyl-CoA pool. 5. Phospholipid-DAG
acyltransferase provides an additional route for the incorporation of PUFAs from PC into TAG. 6.
DAG can also be formed by de novo assembly through the Kennedy pathway, which, however, is
thought to play a minor role in the synthesis of TAGs in seeds. In the chloroplast, biosynthesis of
PG and monogalactosyldiacylglycerol (MGDG) share the precursor phosphatidic acid (PA), with
more PA being shuttled to MGDG biosynthesis in the wild type. In PLIP1-OX lines (middle
panel), both PG biosynthesis and degradation are accelerated, resulting in increased export of 18:3
and other acyl groups and their direct incorporation into PC (reactions 2). Direct incorporation of
18:3 competes with polyunsaturated FA formation by the acyl-editing pathway of PC involving
FAD2/3 (reactions 1), but leads to increased flux of 18:3 into the end product TAG. As a result of
increased PG turnover in chloroplasts of PLIP1-OX lines, PA is preferably shuttled into PG
biosynthesis, which subsequently reduces its availability for MGDG assembly in the plastid visible
in changes in the MGDG acyl composition. In the plip1 mutant (right panel), the PLIP1-dependent
pathway is deficient, resulting in decreased TAG biosynthesis. Without the competing effect of
PLIP1 on the acyl exchange reactions and FAD2/3, more 18:1 is converted to 18:3, explaining the
altered acyl composition of TAG and other extraplastidic lipids.
139

REFERENCES

140

REFERENCES

Ajjawi, I., Lu, Y., Savage, L.J., Bell, S.M., and Last, R.L. (2010). Large-scale reverse genetics
in Arabidopsis: case studies from the Chloroplast 2010 Project. Plant Physiol 152, 529540.
Alonso, J.M., Stepanova, A.N., Leisse, T.J., Kim, C.J., Chen, H., Shinn, P., Stevenson, D.K.,
Zimmerman, J., Barajas, P., Cheuk, R., Gadrinab, C., Heller, C., Jeske, A.,
Koesema, E., Meyers, C.C., Parker, H., Prednis, L., Ansari, Y., Choy, N., Deen, H.,
Geralt, M., Hazari, N., Hom, E., Karnes, M., Mulholland, C., Ndubaku, R., Schmidt,
I., Guzman, P., Aguilar-Henonin, L., Schmid, M., Weigel, D., Carter, D.E.,
Marchand, T., Risseeuw, E., Brogden, D., Zeko, A., Crosby, W.L., Berry, C.C., and
Ecker, J.R. (2003). Genome-wide insertional mutagenesis of Arabidopsis thaliana.
Science 301, 653-657.
Andersson, M.X., and Dörmann, P. (2009). Chloroplast Membrane Lipid Biosynthesis and
Transport. In The Chloroplast, A.S. Sandelius and H. Aronsson, eds (Berlin, Heidelberg:
Springer Berlin Heidelberg), pp. 125-158.
Aronsson, H., and Jarvis, P. (2002). A simple method for isolating import-competent
Arabidopsis chloroplasts. FEBS Lett 529, 215-220.
Bates, P.D., Ohlrogge, J.B., and Pollard, M. (2007). Incorporation of newly synthesized fatty
acids into cytosolic glycerolipids in pea leaves occurs via acyl editing. J Biol Chem 282,
31206-31216.
Bates, P.D., Fatihi, A., Snapp, A.R., Carlsson, A.S., Browse, J., and Lu, C. (2012). Acyl
editing and headgroup exchange are the major mechanisms that direct polyunsaturated
fatty acid flux into triacylglycerols. Plant Physiol 160, 1530-1539.
Benning, C. (2009). Mechanisms of lipid transport involved in organelle biogenesis in plant
cells. Annu Rev Cell Dev Biol 25, 71-91.
Benning, C. (2010). The Anionic Chloroplast Membrane Lipids: Phosphatidylglycerol and
Sulfoquinovosyldiacylglycerol. In The Chloroplast: Biochemistry, Molecular Biology
and Bioengineering., C.A. Rebeiz, C. Benning, H. Bohnert, H. Daniell, B. Green, K.
Hoober, H. Lichtenthaler, A. Portis, and B. Tripathy, eds (Netherlands: Springer), pp.
171-184.
Boudière, L., Michaud, M., Petroutsos, D., Rébeillé, F., Falconet, D., Bastien, O., Roy, S.,
Finazzi, G., Rolland, N., Jouhet, J., Block, M.A., and Maréchal, E. (2014).
Glycerolipids in photosynthesis: Composition, synthesis and trafficking. Biochim
Biophys Acta, Bioenerg 1837, 470-480.

141

Brady, L., Brzozowski, A.M., Derewenda, Z.S., Dodson, E., Dodson, G., Tolley, S.,
Turkenburg, J.P., Christiansen, L., Huge-Jensen, B., Norskov, L., Thim, L., and
Menge, U. (1990). A serine protease triad forms the catalytic centre of a triacylglycerol
lipase. Nature 343, 767-770.
Browse, J., McCourt, P., and Somerville, C.R. (1985). A mutant of Arabidopsis lacking a
chloroplast-specific lipid. Science 227, 763-765.
Browse, J., McConn, M., James, D., Jr., and Miquel, M. (1993). Mutants of Arabidopsis
deficient in the synthesis of alpha-linolenate. Biochemical and genetic characterization of
the endoplasmic reticulum linoleoyl desaturase. J Biol Chem 268, 16345-16351.
Campos, M.L., Yoshida, Y., Major, I.T., de Oliveira Ferreira, D., Weraduwage, S.M.,
Froehlich, J.E., Johnson, B.F., Kramer, D.M., Jander, G., Sharkey, T.D., and Howe,
G.A. (2016). Rewiring of jasmonate and phytochrome B signalling uncouples plant
growth-defense tradeoffs. Nat Commun 7, 12570.
Clough, S.J., and Bent, a.F. (1998). Floral dip: a simplified method for Agrobacteriummediated transformation of Arabidopsis thaliana. Plant J 16, 735-743.
Dahlqvist, A., Stahl, U., Lenman, M., Banas, A., Lee, M., Sandager, L., Ronne, H., and
Stymne, S. (2000). Phospholipid:diacylglycerol acyltransferase: an enzyme that catalyzes
the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Natl
Acad Sci U S A 97, 6487-6492.
Dave, A., and Graham, I.A. (2012). Oxylipin Signaling: A Distinct Role for the Jasmonic Acid
Precursor cis-(+)-12-Oxo-Phytodienoic Acid (cis-OPDA). Front Plant Sci 3, 42.
Dave, A., Hernandez, M.L., He, Z., Andriotis, V.M., Vaistij, F.E., Larson, T.R., and
Graham, I.A. (2011). 12-oxo-phytodienoic acid accumulation during seed development
represses seed germination in Arabidopsis. Plant Cell 23, 583-599.
Earley, K.W., Haag, J.R., Pontes, O., Opper, K., Juehne, T., Song, K., and Pikaard, C.S.
(2006). Gateway-compatible vectors for plant functional genomics and proteomics. Plant
J 45, 616-629.
Gao, J., Ajjawi, I., Manoli, A., Sawin, A., Xu, C., Froehlich, J.E., Last, R.L., and Benning,
C. (2009). FATTY ACID DESATURASE4 of Arabidopsis encodes a protein distinct
from characterized fatty acid desaturases. Plant J 60, 832-839.
Goosens, V.J., and van Dijl, J.M. (2016). Twin-Arginine Protein Translocation. Curr Top
Microbiol Immunol 404, 69-94.
Herr Jr, J. (1993). Clearing techniques for the study of vascular plant tissues in whole structures
and thick sections. In Tested Studies for Laboratory Teaching, C.A. Goldman, P.L.
Hauta, M.A. O'Donnell, S.E. Andrews, and R. van der Heiden, eds (Toronto, Canada:
Association for Biology Laboratory Education), pp. 63–84.
142

Hurlock, A.K., Roston, R.L., Wang, K., and Benning, C. (2014). Lipid trafficking in plant
cells. Traffic 15, 915-932.
Ishiguro, S., Kawai-Oda, A., Ueda, J., Nishida, I., and Okada, K. (2001). The DEFECTIVE
IN ANTHER DEHISCIENCE gene encodes a novel phospholipase A1 catalyzing the
initial step of jasmonic acid biosynthesis, which synchronizes pollen maturation, anther
dehiscence, and flower opening in Arabidopsis. Plant Cell 13, 2191-2209.
Katavic, V., Reed, D.W., Taylor, D.C., Giblin, E.M., Barton, D.L., Zou, J., Mackenzie, S.L.,
Covello, P.S., and Kunst, L. (1995). Alteration of seed fatty acid composition by an
ethyl methanesulfonate-induced mutation in Arabidopsis thaliana affecting diacylglycerol
acyltransferase activity. Plant physiology 108, 399-409.
Keegstra, K., and Yousif, A.E. (1986). Isolation and Characterization of Chloroplast Envelope
Membranes. Method Enzymol 118, 316-325.
Kelly, A.A., and Feussner, I. (2016). Oil is on the agenda: Lipid turnover in higher plants.
Biochim Biophys Acta 1861, 1253–1268.
Kobayashi, K., Endo, K., and Wada, H. (2016). Roles of lipids in photosynthesis. Subcell
Biochem 86, 21-49.
Le, B.H., Cheng, C., Bui, A.Q., Wagmaister, J.A., Henry, K.F., Pelletier, J., Kwong, L.,
Belmonte, M., Kirkbride, R., Horvath, S., Drews, G.N., Fischer, R.L., Okamuro,
J.K., Harada, J.J., and Goldberg, R.B. (2010). Global analysis of gene activity during
Arabidopsis seed development and identification of seed-specific transcription factors.
Proc Natl Acad Sci U S A 107, 8063-8070.
Li-Beisson, Y., Shorrosh, B., Beisson, F., Andersson, M.X., Arondel, V., Bates, P.D., Baud,
S., Bird, D., Debono, A., Durrett, T.P., Franke, R.B., Graham, I.A., Katayama, K.,
Kelly, A.A., Larson, T., Markham, J.E., Miquel, M., Molina, I., Nishida, I.,
Rowland, O., Samuels, L., Schmid, K.M., Wada, H., Welti, R., Xu, C., Zallot, R.,
and Ohlrogge, J. (2013). Acyl-lipid metabolism. Arabidopsis Book 11, e0161.
Li, N., Gugel, I.L., Giavalisco, P., Zeisler, V., Schreiber, L., Soll, J., and Philippar, K.
(2015). FAX1, a novel membrane protein mediating plastid fatty acid export. PLoS Biol
13, e1002053.
Li, X., Moellering, E.R., Liu, B., Johnny, C., Fedewa, M., Sears, B.B., Kuo, M.H., and
Benning, C. (2012). A Galactoglycerolipid Lipase Is Required for Triacylglycerol
Accumulation and Survival Following Nitrogen Deprivation in Chlamydomonas
reinhardtii. Plant Cell 24, 4670–4686.
Li, Y., Beisson, F., Pollard, M., and Ohlrogge, J. (2006). Oil content of Arabidopsis seeds: the
influence of seed anatomy, light and plant-to-plant variation. Phytochemistry 67, 904915.
143

Lu, B., and Benning, C. (2009). A 25-amino acid sequence of the Arabidopsis TGD2 protein is
sufficient for specific binding of phosphatidic acid. J Biol Chem 284, 17420-17427.
Lu, Y., Savage, L.J., Ajjawi, I., Imre, K.M., Yoder, D.W., Benning, C., DellaPenna, D.,
Ohlrogge, J.B., Osteryoung, K.W., Weber, A.P., Wilkerson, C.G., and Last, R.L.
(2008). New connections across pathways and cellular processes: industrialized mutant
screening reveals novel associations between diverse phenotypes in Arabidopsis. Plant
Physiol 146, 1482-1500.
Marchler-Bauer, A., Derbyshire, M.K., Gonzales, N.R., Lu, S., Chitsaz, F., Geer, L.Y.,
Geer, R.C., He, J., Gwadz, M., Hurwitz, D.I., Lanczycki, C.J., Lu, F., Marchler,
G.H., Song, J.S., Thanki, N., Wang, Z., Yamashita, R.A., Zhang, D., Zheng, C., and
Bryant, S.H. (2015). CDD: NCBI's conserved domain database. Nucleic Acids Res 43,
D222-226.
McCourt, P., Browse, J., Watson, J., Arntzen, C.J., and Somerville, C.R. (1985). Analysis of
photosynthetic antenna function in a mutant of Arabidopsis thaliana (L.) lacking transhexadecenoic acid. Plant Physiol 78, 853-858.
Mhaske, V., Beldjilali, K., Ohlrogge, J., and Pollard, M. (2005). Isolation and characterization
of an Arabidopsis thaliana knockout line for phospholipid: diacylglycerol transacylase
gene (At5g13640). Plant Physiol Biochem 43, 413-417.
Moellering, E.R., Muthan, B., and Benning, C. (2010). Freezing tolerance in plants requires
lipid remodeling at the outer chloroplast membrane. Science 330, 226-228.
Murashige, T., and Skoog, F. (1962). A revised medium for rapid growth and bio assays with
tobacco tissue cultures. Physiol Plant 15, 473-497.
Ngaki, M.N., Louie, G.V., Philippe, R.N., Manning, G., Pojer, F., Bowman, M.E., Li, L.,
Larsen, E., Wurtele, E.S., and Noel, J.P. (2012). Evolution of the chalcone-isomerase
fold from fatty-acid binding to stereospecific catalysis. Nature 485, 530-533.
Pazos, F., Ranea, J.A., Juan, D., and Sternberg, M.J. (2005). Assessing protein co-evolution
in the context of the tree of life assists in the prediction of the interactome. J Mol Biol
352, 1002-1015.
Quast, C., Pruesse, E., Yilmaz, P., Gerken, J., Schweer, T., Yarza, P., Peplies, J., and
Glockner, F.O. (2013). The SILVA ribosomal RNA gene database project: improved
data processing and web-based tools. Nucleic Acids Res 41, D590-596.
Ribot, C., Zimmerli, C., Farmer, E.E., Reymond, P., and Poirier, Y. (2008). Induction of the
Arabidopsis PHO1;H10 gene by 12-oxo-phytodienoic acid but not jasmonic acid via a
CORONATINE INSENSITIVE1-dependent pathway. Plant Physiol 147, 696-706.
Richmond, G.S., and Smith, T.K. (2011). Phospholipases A1. Int J Mol Sci 12, 588-612.
144

Robinson, C., and Bolhuis, A. (2001). Protein targeting by the twin-arginine translocation
pathway. Nat Rev Mol Cell Biol 2, 350-356.
Rodrigues, R.A., Silva-Filho, M.C., and Cline, K. (2011). FtsH2 and FtsH5: two homologous
subunits use different integration mechanisms leading to the same thylakoid multimeric
complex. Plant J 65, 600-609.
Roston, R., Gao, J., Xu, C., and Benning, C. (2011). Arabidopsis chloroplast lipid transport
protein TGD2 disrupts membranes and is part of a large complex. Plant J 66, 759-769.
Roston, R.L., Gao, J., Murcha, M.W., Whelan, J., and Benning, C. (2012). TGD1, -2, and -3
proteins involved in lipid trafficking form ATP-binding cassette (ABC) transporter with
multiple substrate-binding proteins. J Biol Chem 287, 21406-21415.
Routaboul, J.M., Benning, C., Bechtold, N., Caboche, M., and Lepiniec, L. (1999). The
TAG1 locus of Arabidopsis encodes for a diacylglycerol acyltransferase. Plant Physiol
Biochem 37, 831-840.
Scherer, G.F., Ryu, S.B., Wang, X., Matos, A.R., and Heitz, T. (2010). Patatin-related
phospholipase A: nomenclature, subfamilies and functions in plants. Trends Plant Sci 15,
693-700.
Schwacke, R., Schneider, A., van der Graaff, E., Fischer, K., Catoni, E., Desimone, M.,
Frommer, W.B., Flugge, U.I., and Kunze, R. (2003). ARAMEMNON, a novel
database for Arabidopsis integral membrane proteins. Plant Physiol 131, 16-26.
Tanoue, R., Kobayashi, M., Katayama, K., Nagata, N., and Wada, H. (2014).
Phosphatidylglycerol biosynthesis is required for the development of embryos and
normal membrane structures of chloroplasts and mitochondria in Arabidopsis. Febs Lett
588, 1680-1685.
Troncoso-Ponce, M.A., Cao, X., Yang, Z., and Ohlrogge, J.B. (2013). Lipid turnover during
senescence. Plant Sci 205-206, 13-19.
Wang, G., Ryu, S., and Wang, X. (2012). Plant phospholipases: an overview. Methods Mol
Biol 861, 123-137.
Wang, K., Hersh, H.L., and Benning, C. (2016). SENSITIVE TO FREEZING2 Aids in
Resilience to Salt and Drought in Freezing-Sensitive Tomato. Plant Physiol 172, 14321442.
Wang, X. (2004). Lipid signaling. Curr Opin Plant Biol 7, 329-336.
Wang, Z., and Benning, C. (2011). Arabidopsis thaliana polar glycerolipid profiling by thin
layer chromatography (TLC) coupled with gas-liquid chromatography (GLC). J Vis Exp
49, 2518
145

Winkler, F.K., D'Arcy, A., and Hunziker, W. (1990). Structure of human pancreatic lipase.
Nature 343, 771-774.
Winter, D., Vinegar, B., Nahal, H., Ammar, R., Wilson, G.V., and Provart, N.J. (2007). An
"Electronic Fluorescent Pictograph" browser for exploring and analyzing large-scale
biological data sets. PloS One 2, e718.
Xie, D.X., Feys, B.F., James, S., Nieto-Rostro, M., and Turner, J.G. (1998). COI1: an
Arabidopsis gene required for jasmonate-regulated defense and fertility. Science 280,
1091-1094.
Xu, C., Fan, J., Cornish, A.J., and Benning, C. (2008). Lipid trafficking between the
endoplasmic reticulum and the plastid in Arabidopsis requires the extraplastidic TGD4
protein. Plant Cell 20, 2190-2204.
Xu, C., Fan, J., Froehlich, J.E., Awai, K., and Benning, C. (2005). Mutation of the TGD1
chloroplast envelope protein affects phosphatidate metabolism in Arabidopsis. Plant Cell
17, 3094-3110.
Yan, Y., Stolz, S., Chetelat, A., Reymond, P., Pagni, M., Dubugnon, L., and Farmer, E.E.
(2007). A downstream mediator in the growth repression limb of the jasmonate pathway.
Plant Cell 19, 2470-2483.
Zeng, W., Brutus, A., Kremer, J.M., Withers, J.C., Gao, X., Jones, A.D., and He, S.Y.
(2011). A genetic screen reveals Arabidopsis stomatal and/or apoplastic defenses against
Pseudomonas syringae pv. tomato DC3000. PLoS Pathog 7, e1002291.
Zhang, Y., and Turner, J.G. (2008). Wound-induced endogenous jasmonates stunt plant
growth by inhibiting mitosis. PloS One 3, e3699.

146

CHAPTER 4
Two Abscisic Acid Responsive Plastid Lipase Genes Involved in Jasmonic Acid
Biosynthesis in Arabidopsis thaliana

147

ABSTRACT
Chloroplast membranes with their unique lipid composition are crucial for photosynthesis.
Maintenance of the chloroplast membranes requires a finely tuned lipid anabolic and catabolic
machinery. Despite the presence of a large number of predicted lipid-degrading enzymes in the
chloroplasts, their biological functions remain largely unknown. Recently, we described PLASTID
LIPASE 1 (PLIP1) as a plastid phospholipase A1 that contributes to seed oil biosynthesis. The
Arabidopsis genome encodes two putative PLIP1 paralogs which we designated PLIP2 and PLIP3.
PLIP2 and PLIP3 are also located in the chloroplasts, but likely with different locations in the
plastid. PLIP2 possesses similar biochemical properties in vitro to PLIP1, but in vivo studies
suggest that PLIP2 prefers monogalactosyldiacylglycerol as substrate, while PLIP3 prefers
phosphatidylglycerol. Overexpression of PLIP2 or PLIP3 severely reduces plant growth and leads
to accumulation of oxylipins. Genetically blocking jasmonate perception restored the growth of
the overexpression plants. The expression of PLIP2 and PLIP3, but not PLIP1 is induced by
abscisic acid (ABA), and plip1,2,3 triple mutants exhibit compromised oxylipin biosynthesis in
response to ABA. We propose that PLIP2 and PLIP3 provide a mechanistic link between ABA
mediated abiotic stress responses and oxylipin signaling.

148

INTRODUCTION
Chloroplasts in plants and algae are organelles that carry out a number of crucial biological
functions including photosynthesis. They also participate in plant immune responses through the
production of defense signaling compounds such as oxylipins and ultimately jasmonic acid (JA).
Membranes with a unique lipid composition are required for the proper functioning of the
chloroplasts.

Chloroplast

membranes

contain

predominately

galactolipids,

monogalactosyldiacylglycerol (MGDG) and digalactosyldiacylglycerol (DGDG), as well as two
anionic lipids phosphatidylglycerol (PG), and sulfoquinovosyldiacylglycerol (SQDG) (Andersson
and Dörmann, 2009; Benning, 2010; Boudière et al., 2014). In seed plants, lipid assembly begins
in the plastids where fatty acid (FA) synthesis occurs. In Arabidopsis, two pathways are
responsible for plant lipid assembly (Benning, 2009; Hurlock et al., 2014). De novo synthesized
FAs can either directly enter the plastid (prokaryotic) pathway or are exported to the endoplasmic
reticulum (ER) and assembled in to lipids there (eukaryotic pathway). Lipid precursors synthesized
in the ER can also return to the plastid and contribute to chloroplast lipid biosynthesis (Wang and
Benning, 2012; Hurlock et al., 2014). Lipids synthesized by the two pathways can be distinguished
by their glyceryl sn-2-bound acyl groups due to the specificity of the respective acyltransferases
(Frentzen et al., 1983; Frentzen, 1986). Plastid pathway-derived galactolipids contain 16-carbon
acyl groups at the sn-2 glyceryl position, while ER-derived lipids contain 18-carbon acyl groups
in this position.
To date, many of the genes and enzymes responsible for chloroplast lipid biosynthesis have
been identified, yet still little is known about the role of enzymes involved in chloroplast lipid
degradation. Turnover of glycolipids requires lipases to release the acyl or the head groups from
the three different positions of the glyceryl backbone (Troncoso-Ponce et al., 2013; Kelly and

149

Feussner, 2016). Based on their preferred hydrolysis position, phospholipases are classified into
three groups (Wang et al., 2012): Phospholipase C (PLC) and phospholipase D (PLD) cleave the
phosphodiester bonds adjacent and distal to the sn-3 glyceryl position, respectively.
Phospholipases A (PLA) release FAs from glyceryl sn-1 or sn-2 positions; PLA1 hydrolyses at the
sn-1 position while PLA2 at sn-2. Free FAs released by PLAs from the membrane lipids can be
subsequently oxygenated by either enzymatic reactions or chemical oxidation (Blee, 1998;
Mueller, 2004; Mosblech et al., 2009). In plants, enzyme-mediated FA oxidation predominantly
works on polyunsaturated FAs, e.g., linolenic acids (18:3; number of carbons : number of double
bonds in the acyl chain) through the lipoxygenase pathway (Porta and Rocha-Sosa, 2002).
Lipoxygenases initiate the synthesis of FA hydroperoxides. Their subsequent conversion occurs
through an array of alternative pathways (Porta and Rocha-Sosa, 2002; Mosblech et al., 2009).
Structurally and functionally diverse oxylipins are produced during different developmental
stages, in different tissues as well as in response to different environmental cues (Gobel et al.,
2002; Porta and Rocha-Sosa, 2002; Andersson et al., 2006). The best characterized lipid oxidation
pathway is the allene oxide synthase pathway which leads to the synthesis of oxylipins, including
12-oxo phytodienoic acid (OPDA), JA and the conjugates between JA and amino acids, e.g., the
bioactive JA-isoleucine (JA-Ile) (Turner et al., 2002; Wasternack and Hause, 2013). JA production
and signaling are primarily induced by wounding damage caused by herbivore insects or
necrotrophic microbes (Creelman et al., 1992; Trusov et al., 2006). Developmentally, JA is
essential for stamen development, size of petioles and petals, as well as lignin biosynthesis (Feys
et al., 1994; McConn and Browse, 1996; Ellis et al., 2002; Buseman et al., 2006; Denness et al.,
2011). DEFECTIVE IN ANTHER DEHISCENCE1 (DAD1) is a PLA1 that is primarily present in
the reproductive tissues, where it initiates JA biosynthesis critical for pollen maturation, anther

150

dehiscence and flower opening in Arabidopsis (Ishiguro et al., 2001). In leaf tissues, DONGLE
encodes a DAD1-homologous lipase contributing to wounding-induced JA production (Hyun et
al., 2008). In addition, synergetic or antagonistic interactions between JA and other
phytohormones including salicylic acid, ethylene and ABA have emerged (Thomma et al., 1998;
Song et al., 2014; Suzuki, 2016), suggesting that the interaction between different hormones is an
important strategy for plants to fine-tune their responses to environmental cues.
It is known that JA biosynthesis is stimulated by applied ABA or during ABA-inducing
abiotic stress conditions (Creelman and Mullet, 1995; Adie et al., 2007; Avramova, 2017), but the
underlying mechanisms are not well understood. During abiotic stress, rising ABA levels induce
JA synthesis, which synergistically optimizes plant abiotic defenses by altering the plant
transcriptome, facilitating the closure of stomata, inhibiting cell division, as well as epigenetically
modifying defense genes during dehydration responses (Murata et al., 2015; Riemann et al., 2015;
Liu et al., 2016; Valenzuela et al., 2016). A basic-helix-loop-helix transcription factor MYC2 plays
roles in JA perception and ABA signaling, and has been proposed as one of the factors linking
ABA and JA signaling (Kazan and Manners, 2013).
In Arabidopsis, over 50 lipases are predicted to be localized in the chloroplasts with
unverified enzymatic activities and unknown biological functions (Ajjawi et al., 2010). Recently,
we discovered a thylakoid associated lipase named PLIP1, which is a PLA1 that releases
polyunsaturated FAs from chloroplast PG and exports the FAs to the ER for seed oil biosynthesis
(Wang et al., 2017). BLAST searches and phylogenetic analyses identified two putative PLIP1
paralogs in the Arabidopsis genome, which we designated PLIP2 and PLIP3. Both PLIP2 and
PLIP3 are located in the chloroplasts. Their characterization and possible role in JA biosynthesis
in response to increased ABA levels following abiotic stress is explored below.

151

MATERIALS AND METHODS
Plant Material and Growth Conditions
All experiments were performed in the Arabidopsis thaliana Col-0 ecotype. The T-DNA
insertion lines SALK_1234548 (plip2-1), SALK_134525 (plip2-2), SALK_006633 (plip3-1) and
SALK_115539 (plip3-2) were obtained from the Arabidopsis Biological Resource Center, Ohio
State University. For the overexpression lines, the coding sequences of PLIP2 or PLIP3 were
inserted into the pENTRTM/TEV/D-TOPO® vector (Life Sciences; Catalog number: K240020)
before recombining them into the pEarleyGate 101 vector (YFP at the C-terminus) (Earley et al.,
2006) with the Gateway LR Clonase Kit (Thermo Fisher Scientific; Catalog number: 11791-020).
The final constructs were introduced into Col-0 wild-type plants by Agobacterium tumefaciensmediated floral dip (Clough and Bent, 1998). Transformed seeds were initially screened for
resistance to Basta, followed by confirmation by RT-PCR. Primers used for genotyping of T-DNA
insertion lines or for RT-PCR analysis of overexpression lines are given in Table 4.1. The growth
condition for the Murashige and Skoog (MS) medium grown and the soil-grown Arabidopsis
plants are the same as previously reported (Wang et al., 2017).

Phylogenetic Analysis
The top 20 PLIP1 putative paralog amino acid sequences were obtained by comparing the
AtPLIP1 protein sequence against the Arabidopsis proteome using the BLASTp program in TAIR
BLAST 2.2.8 (http://www.arabidopsis.org/Blast/index.jsp) with the default parameters. The amino
acid alignment was created using the MUSCLE program with default settings employing MEGA
(version 7.0.21) software. The Maximum Likelihood phylogenetic trees were built using MEGA
(version 7.0.21) and the bootstrap percentages were based on 1000 replicates.

152

Confocal Laser Scanning Microscopy
The confocal images were taken with an Nikon A1Rsi confocal microscope on leaves of
3-week-old Arabidopsis grown on MS agar plates. YFP fusion proteins were detected with the
configuration A1CYFR using a 514 nm laser for excitation and a 530-600 nm band pass filter for
fluorescence emission. For the chlorophyll autofluorescence, a 647 nm laser was used for
excitation and a 660 nm long pass filter for emission detection. Images were merged and
pseudcolored using the NIS-Elements AR software (version 4.30.01 64-bit).

Chloroplast Import Assay
Isolation of pea chloroplasts, import assays and post-import trypsin treatment were done
essentially as previously described (Xu et al., 2005). The constructs used for production of labeled
proteins were pET41a-PLIP2 and pET41a-PLIP2. N-terminal 6×His tag and TEV cleavage sites
were removed using a Q5 Site-Directed Mutagenesis Kit (New England Biolabs).

Recombinant Protein Production
The PLIP2 coding sequence was amplified from the Arabidopsis wild-type cDNA and
inserted into the pGEM-T-Easy plasmid (Promega). It was then inserted into the pET41a plasmid
at the BamHI and XhoI restriction sites. The PLIP2S428A point mutation construct was generated
with a Q5 Site-Directed Mutagenesis Kit (New England Biolabs). Final pET41a-PLIP2 and
pET41a-PLIP2S428A constructs were introduced into BL21 (DE3) E. coli strains for protein
production. Cultures grown in LB medium (containing 0.1% glucose) were inoculated with fresh
E. coli colonies and grown to log phase (OD600 0.8) at 37 °C. Protein production was then induced
by adding isopropyl-β-D-thiogalactopyranoside (IPTG) to the final concentration of 0.2 mM, and

153

the culture was transferred to 14 °C. Cells were harvested after 1.5 h of induction. Cultures were
harvested and sonicated to lyse cells. Supernatant was collected after centrifugation at 10,000 × g
for 30 min, and subjected to another 1-h centrifugation at 100,000 × g to remove the supernatants.
The pellet fraction was then resuspended with PBS buffer to give rise to microsomes. Microsomes
harvested from the cultures expressing PLIP2, PLIP2S428A or the empty vector were used for the
following lipase assays.

PLIP2 Lipase Assay
The lipase assay was performed essentially according to Wang et al. 2017. Briefly,
microsomes harvested from 1 L induced cultures producing PLIP2 were resuspended in 1 mL PBS
buffer. For each lipase assay, 100 µg dried lipids were first resuspended in 300 µL reaction buffer
(0.1 M PBS, pH 7.4; 4.2 mM Anzergent 3-12 (Anatrace)) and dispersed by sonication. Then, 50
µL microsomes were added to each reaction followed by another 10 s sonication to mix. The
reaction was incubated at ambient temperature (~22 °C) for 1 h or as indicated for time courses
(Figure 4.1). The reaction was stopped by lipid extraction, followed by lipid separation by TLC
and quantification by gas chromatography as described below.

Lipid Analysis
Lipid extraction, TLC of polar and neutral lipids, transesterification, and gas
chromatography were done as described in Wang and Benning 2011. For polar lipids, activated
ammonium sulfate-impregnated silica gel TLC plates (TLC Silica gel 60; EMD Chemical,
Germany) were used for lipid separation with a solvent consisting of acetone, toluene and water
(91:30:7.5 by volume). Lipids were visualized by brief exposure to iodine vapor on TLC plates.

154

Acyl groups of the isolated lipids were then converted to methyl esters, which were subsequently
quantified by a gas chromatography.

Pulse-Chase Labeling
The pulse-chase labeling experiments on 4-week-old soil-grown plant leaves were
performed essentially according to Wang et al. 2017.

Hormone Measurements and Oxylipin Profiling
Hormone extraction and measurements were done based on Wang et al., 2017. 12OH-JA
and 12OH-JA-Ile were added as standards. Their transitions from deprotonated molecules to
characteristic product ions were monitored (12OH-JA, m/z 225.09 > 58.90; 12OH-JA-Ile, m/z
338.23 > 130.09), and the collision energies and source cone potentials were optimized for each
compound with the QuanOptimize software before integration into the LC-MS/MS method.
For oxylipin profiling, oxidized lipids were extracted and analyzed using a Waters Xevo
G2-XS Q-TOF mass spectrometer according to Wang et al. 2017. The metabolite chromatograms
from the WT and the PLIP-OX leaves samples were normalized and aligned using the Progenesis
QI software (version 2.2) to identify peaks accumulated in the PLIP-OX, but not the WT samples.
Identification of the Arabidopside metabolites was based on their accurate masses and the
signature OPDA or dn-OPDA fragment ions (m/z values are 291.19 and 263.16, respectively). The
peak areas of the Arabidopsides were normalized to that of the 18:3/18:3-DGDG ([M-H]-, m/z
981.61).

155

Accession Numbers
Sequence data from this article can be found in the Arabidopsis TAIR database
(https://www.arabidopsis.org/) under the following accession numbers: At3g61680 for PLIP1,
At1g02660 for PLIP2, At3g62590 for PLIP3, At5g24780 for VSP1, At3g45140 for LOX2,
At1g32640 for MYC2, At1g19180 for JAZ1, At5g13220 for JAZ10, AT2G27690 for CYP94C1.

156

RESULTS
PLIP2 and PLIP3 are Putative Paralogs of PLIP1 in Arabidopsis
PLIP1 was included in the Chloroplast 2010 project studying chloroplast targeted proteins
(Ajjawi et al., 2010) and later characterized as a phospholipase A1 enzyme leading to acyl export
from the chloroplasts and contributing to seed oil biosynthesis (Wang et al., 2017). Comparing the
PLIP1 protein sequence against the Arabidopsis proteome using BLASTp with default settings in
TAIR (http://www.arabidopsis.org) identified two putative paralogs of PLIP1 in Arabidopsis
encoded by AT1G02660 and AT3G62590, which were designated PLIP2 and PLIP3, respectively.
Phylogenetic analysis of the top 20 Arabidopsis proteins similar to PLIP1 (Figure 4.2) showed that
PLIP2 and 3 closely clustered with PLIP1, but were distinct from other predicted α / β hydrolases.
However, PLIP2 and 3 peptide sequences only share 41% and 37% identity with the PLIP1
peptide, respectively (57% and 51% similarity, respectively). PLIP2 and 3 are more closely related
to each other sharing 51% identity and 65% similarity.

PLIP2 and 3 are Located in the Chloroplast
PLIP1 is imported into the chloroplast with an N-terminal transit peptide (Wang et al.,
2017). PLIP2 and 3 were also predicted to be located in the chloroplast using different subcellular
location prediction programs, e.g., TargetP (Emanuelsson et al., 2000) or PredSL (Petsalaki et al.,
2006). To experimentally verify this prediction, the coding sequences for PLIP2 or 3 derived from
an Arabidopsis wild-type (Col-0) cDNA were spliced to the open reading frame of YFP at its 5’end (creating a C-terminal fusion of YFP to the PLIPs). When the PLIP-YFP constructs were
stably expressed in wild type under the control of the CaMV 35S promoter, the YFP signals were
observed specifically overlapping with the chlorophyll signals using confocal microscopy (Figure

157

4.3 A). Interestingly, the chloroplast sizes were smaller in plants overexpressing either PLIP2 or
3 than those expressing the empty vector (EV) control. This phenotype is probably corresponding
to the reduced plant size when overexpressing PLIP cDNAs described below.
The PLIP2 and 3 genes are predicted to encode 78,346 D and 73, 044 D proteins,
respectively. Based on the ARAMEMNON database (Schwacke et al., 2003), PLIP2 and 3 have
similar predicted topologies, both containing four transmembrane domains with a very short transit
peptide at the N-terminus. A chloroplast import assay was performed to confirm the plastid
location of PLIP2 and 3. PLIP2 and 3 cDNAs were translated in vitro in the presence of labeled
methionine. The translation products were then tested for import into isolated pea (Pisum sativum)
chloroplasts. As shown in Figure 4.3 B, both PLIP2 and 3 were imported into chloroplasts. During
the import, the precursor protein of PLIP2 was processed to a smaller protein, while PLIP3 did not
show detectable processing possibly due to the small size of the transit peptide. The PLIP2 signal
was present in both the pellet and supernatant fractions. Trypsin is a protease capable of penetrating
the chloroplast outer, but not the inner envelope membrane, respectively. With trypsin treatment,
only the mature PLIP2 protein was retained in the smaller pellet fraction, while the soluble fraction
was not affected (Figure 4.3 B). This suggests that PLIP2 is probably ubiquitously present in the
chloroplast envelope membranes, stroma and thylakoids. For PLIP3, the signal was predominantly
associated with the membrane fractions in the pellet and the signal intensity partially faded with
trypsin treatment, indicating that PLIP3 is a membrane protein embedded in the chloroplast
envelope membranes and thylakoids. Taken together, PLIP2 and 3 are likely chloroplast located
proteins, but have distinct subplastid locations.

158

PLIP2 is a Phospholipase A1 with Ser428 as the Catalytic Site
Both PLIP2 and 3 are annotated as triacylglycerol lipases in TAIR and contain conserved
Lipase 3 domains similar to PLIP1. Lipases are characterized by a signature catalytic triad
comprised of Ser-Asp-His, and reduced occasionally to a dyad (Ser-Asp). Sequence alignment of
PLIP2 and 3 with classic lipases using NCBI’s conserved domain database (Marchler-Bauer et al.,
2015) identified a conserved triad in PLIP2, while only a dyad was identified in PLIP3 (Figure
4.4). To verify whether PLIP2 and 3 possess any lipase activity, recombinant PLIP2 or 3 constructs
were introduced into E. coli or Saccharomyces cerevisiae and protein production was investigated
by immunoblotting against the 6 × His tag included in the N-terminus. Using different variations
of the protocol, we were unable to observe production of the recombinant PLIP3 protein neither
in E. coli nor in yeast. However, production of the recombinant PLIP2 protein and a mutant protein
in which the putative catalytic residue Ser428 was changed to Ala (PLIP2S422A) were successful in
E. coli using a protocol with a brief induction period and growth under low temperature (Figure
4.5 A). Phosphatidylcholine (PC) was then provided to the microsome fractions isolated from E.
coli cultures expressing PLIP2 or PLIP2S422A. At the end of the reaction, lipids were extracted and
analyzed on a thin layer chromatography (TLC) plate (Figure 4.5 B). Degradation of PC and
production of lyso-PC were only observed with PLIP2, but not with PLIP2S422A, suggesting that
PLIP2 is a lipase with a conserved Ser428 as its catalytic residue.
Detection of lyso-lipids in the lipase assay implied that PLIP2 can only hydrolyze one of
the two acyl-glyceryl ester bonds in PC. To determine the positional specificity of PLIP2, two
defined molecular species of PC molecules with inverted acyl compositions were offered to PLIP2.
At the end of the reaction, lyso-PC was isolated by TLC, and FA methyl esters derived from the
lyso-lipid were analyzed by gas-chromatography (Figure 4.6 A). For PC with acyl groups

159

composed of 18:1∆9/16:0 (sn-1/ sn-2) as substrates, 18:1∆9 was selectively released while 16:0 was
retained. The opposite pattern was observed for lyso-PC produced from 16:0/18:1∆9-PC. These
data suggested that PLIP2 preferentially hydrolyzes the sn-1 position and, therefore, is a
phospholipase A1. To further investigate a possible acyl group preference of PLIP2 at the sn-1
glyceryl position, PLIP2 was offered different combinations of defined PC molecules carrying the
same acyl groups at the sn-2 but acyl groups with different degrees of saturation at the sn-1 position
(Figure 4.6 B). Comparing 18:0/18:1 with 18:1/18:1 PC, PLIP2 showed slightly higher activity
with 18:1/18:1 PC, while no statistically significant difference was detected between18:0/18:2 and
18:2/18:2. This result implied that, at least in vitro, PLIP2 does not have a strong preference for
the degree of saturation of acyl groups and is capable of releasing both saturated and unsaturated
acyl groups from the sn-1 position.

PLIP2 Uses MGDG as Primary Substrate, While PLIP3 Uses PG
Plant glycerolipid lipases such as PLIP1 tend to be promiscuous with regard to the
glycerolipid head groups in vitro, and a facile approach to investigate their substrate specificity
and to identify their native substrates is to overproduce them in the plant, thereby providing them
with a native lipid environment (Wang et al., 2017). Accordingly, we then took advantage of the
PLIP2 and 3 overexpressing plants that were used for the localization study above. We generated
over 20 independent overexpression lines for each of PLIP2 and PLIP3 constructs. Three lines for
each construct were chosen for further studies based on the presence of the recombinant protein as
detected by confocal microscopy. It should be noted that despite the use of the strong constitutive
promoters, location of PLIP2-YFP and PLIP3-YFP were strictly limited to the chloroplasts in the
overexpression lines. We performed lipidomic analysis on the relative abundance of polar lipids

160

and the acyl composition of individual polar lipids in one empty vector (EV) control line and two
PLIP2 overexpression (PLIP2-OX) lines (Figure 4.7 A and Figure 4.8), as well as two PLIP3-OX
lines (Figure 4.9). The lipid phenotype of PLIP2-OX and PLIP3-OX plants was very similar to
that of the previously reported PLIP1-OX plants (Wang et al., 2017). Specifically, two chloroplast
lipids (MGDG and PG) and ER lipids (PI, PE and PC) showed statistically significant changes in
their acyl compositions. MGDG exhibited primarily decreased 16:3 to 18:3 ratios, and PG showed
decreased 16:1 to 16:0 ratios in all PLIP2-OX and PLIP3-OX plants. 16:3 acyl groups are
predominantly present at the sn-2 glyceryl position of plastid pathway-derived MGDG. These
changes in acyl ratios suggest that PLIP2 and 3 might have a substrate preference for plastid
pathway-derived MGDG and 16:1-PG.
The steady-state compositional changes observed for the chloroplast lipids MGDG and PG
led us to investigate their synthesis and turnover in vivo with pulse-chase lipid labeling analysis in
both the PLIP2-OX and PLIP3-OX plants. For this purpose, [14C]-acetate was used to label the de
novo synthesized FAs in detached leaves. In the PLIP2-OX leaves, MGDG was labeled
approximately 20% higher during the pulse phase (Figure 4.7 B), suggesting that FA incorporation
into MGDG is accelerated. During the chase phase (Figure 4.7 C), the label in MGDG was lost
within the first 10 hours and labeled MGDG eventually reached a level comparable to that in the
EV control plants. The accelerated turnover of the MGDG pool in the PLIP2-OX leaves suggested
that MGDG is likely the favored native substrate of PLIP2 in vivo. In contrast, for the PLIP3-OX
plants (Figure 4.10), pulse-chase labeling data showed that the labeling of the MGDG pool was
not altered. Instead, PG was labeled almost twice as high during the pulse phase in the PLIP3-OX
plants, and the label in PG was rapidly lost during the chase phase, a pattern that resembled that

161

previously observed for the PLIP1-OX plants (Wang et al., 2017). These data suggested that PLIP3
likely uses PG as its preferred substrate as was also concluded for PLIP1 (Wang et al., 2017).

The Growth and Lipid Phenotypes are Not Correlated in PLIP-OX Plants
One of the striking phenotypes of the PLIP2 and 3 overexpression plants was that their
vegetative growth was severely repressed under normal growth condition (Figure 4.11 A). This
growth phenotype was similar to, but more severe than that observed for the PLIP1-OX plants
(Figure 4.11 A). Despite the fact that overexpression of PLIPs significantly altered plant lipid
composition, it seemed unlikely that these structural lipid changes were the underlying primary
cause for the growth phenotype, because the lipid phenotypes were not correlated with the growth
phenotype. To illustrate this fact, acyl group ratios of the presumed PLIP-preferred substrates
MGDG (18:3/16:3) and PG (16:0/16:1) were chosen as representative indicators (Figure 4.11 A
bottom panel). Based on these indicators, PLIP1-OX plants exhibited the most severe lipid
changes, but showed the mildest growth reduction among all the PLIP-OX plants. On the contrary,
the PLIP2-OX plants exhibited severely reduced vegetative growth, but only showed mild lipid
changes for both MGDG and PG. This inconsistency between the lipid and growth phenotypes
was also visible in the pulse-chase labeling data (Figure 4.7 and 4.10). This result implied that
PLIP2 and 3 might be involved in physiological functions different from PLIP1.
The reduced vegetative growth of PLIP2-OX and PLIP3-OX plants was evident even when
they grew on nutrient rich agar plates, and the PLIP2-OX plants are shown in Figure 6B as an
example. Besides the reduced rosette sizes, these plants had shorter leaf petioles than the wild-type
control plants. Also, a red pigment, likely anthocyanin, accumulated the center of the plants under
normal growth conditions (Figure 4.11 B). The elevated level of the presumed anthocyanin was

162

particularly apparent during lipid extraction and phase partitioning as the red pigment stained the
upper aqueous phase (Figure 4.11 C). Shortened petioles and accumulation of anthocyanin are
typical phenotypes of plants with activated JA signaling or plants that overproduce JA, e.g., the
jazQ (Campos et al., 2016) and the dgl-D mutants (Hyun et al., 2008), respectively. JA is an 18:3FA derived phytohormone, which upon accumulation poises the plant transcriptome from growth
to defense leading, among others, to reduced vegetative growth (Acosta and Farmer, 2010; Huot
et al., 2014). In vitro characterization of PLIP2 determined its PLA1 activity and capability of
releasing polyunsaturated FAs (Figure 4.6). Also, PLIP2’s presumed native substrate MGDG and
PLIP3’s presumed native substrate PG contain predominately 18:3 at the sn-1 position. Therefore,
we hypothesized that the stunted vegetative growth might be caused by the overproduction of JA
and activation of the JA signaling pathway in the PLIP-OX lines.
To test whether JA signaling is relevant to the observed growth inhibition, the expression
of JA-responsive marker genes was examined by quantitative PCR (Figure 4.11 D). As a result,
all the analyzed JA maker genes were constitutively induced in the PLIP-OX plants, including
genes involved in JA biosynthesis (LOX2), JA signal transduction (MYC2), downstream responses
(VSP1), negative regulators of JA signaling (JAZ1, JAZ10.1-3), as well as JA catabolism
(CYP94C1). In addition, the relative gene expression levels appeared correlated with the level of
growth reduction. For example, the smallest PLIP2-OX plants showed the highest induction of the
JA responsive genes, while PLIP1-OX plants only exhibited a mild induction consistent with its
mild growth reduction (Figure 4.11 D). These data indicated that the JA mediated defense response
is fully activated in the PLIP-OX plants.

163

Oxylipin Production and Signaling in PLIP2- and 3-OX Plants
We hypothesized that the constitutively activated defense response in the PLIP2-OX and
PLIP3-OX plants might be a consequence of increased levels of oxylipins, which could be
produced from the FAs released by PLIP2 and 3. To investigate whether the levels of oxylipins
were changed in the PLIP2-OX and PLIP3-OX plants, a targeted metabolite analysis was
conducted to quantify the major phytohormone levels in leaf tissues (Figure 4.12 A). Most of the
oxylipins were maintained at very low basal levels in the WT plants under the chosen growth
condition (Figure 4.12 A). However, the PLIP2-OX plants showed increased levels of JA, OPDA,
Methyl-JA, and JA-Ile. Also, two JA catabolites, 12OH-JA and 12OH- JA-Ile, accumulated to
levels that were even higher than their active forms, JA and JA-Ile, respectively (Figure 4.12 A).
This was consistent with the constitutive induction of JA catabolism genes in the PLIP2-OX plants
(Figure 4.11 D), e.g., CYP94C1 which encodes a hydroxylase involved in JA-Ile turnover (Heitz
et al., 2012). Similar results were also observed in the PLIP3-OX plants (Figure 4.13 A).
We also tested whether other forms of oxylipins than those mentioned above accumulate
in the overexpression lines. Oxylipins include classes of highly diverse and complicated lipid
molecules, in which a typical fatty acid is replaced with an oxidized fatty acid (Gobel and Feussner,
2009; Mosblech et al., 2009). Oxylipins other than JA start to be recognized as important signaling
molecules contributing to various aspects of plant physiology (Porta and Rocha-Sosa, 2002). To
more comprehensively observe changes in oxylipin metabolism in the PLIP2-OX and PLIP3-OX
plants, a non-targeted metabolite profiling was focused on extracted oxidized complex lipids using
liquid chromatography-QTOF-MS. Through comparative metabolic profiling, a few metabolites
were highly accumulated in the PLIP2-OX plants but not in the WT (Figure 4.12 B). By
investigation of their accurate mass and their molecular fragmentation patterns, these metabolites

164

were tentatively identified as Arabidopsides (Figure 4.12 C). Arabidopsides are structurally very
similar to galactolipids except they contain OPDA or dinor-OPDA instead of acyl groups at the
sn-1 and sn-2 glyceryl positions. Arabidopsides do not usually accumulate in plants grown under
normal condition unless plants are wounded. However, in the PLIP2-OX plants, the levels of
Arabidopsides A, B and D were highly elevated. Arabidopsides have been proposed as storage
pools for OPDA, and this is possibly a compensatory mechanism for fine-tuning free OPDA levels
in the cells. Similarly, accumulation of these compounds was also detected in the PLIP3-OX plants
(Figure 4.13 B).
The targeted phytohormone measurement and the non-targeted metabolite profiling
strongly indicated that JA metabolism and consequently signaling were affected in the PLIP2-OX
and PLIP3-OX plants. To directly test whether activated JA signaling specifically caused the
reduced vegetative growth, one each of the PLIP2-OX and PLIP3-OX lines was crossed with a
mutant defective in CORONATINE INSENSITIVE1 (coi1), carrying an knock-out mutant of the
JA receptor. As a result, the growth inhibition of the PLIP-OX plants was completely reversed in
PLIP-OX coi1 homozygous offspring (Figure 4.14 A and Figure 4.15 A), while the lipid
phenotypes remained the same as in the PLIP-OX plants (Figure 4.14 B and Figure 4.15 B). Thus,
using this genetic approach, we were able to fully separate the primary lipid phenotype from the
growth phenotype, confirming that the stunted growth phenotypes were caused by constitutive JA
signaling due to oxylipin accumulation in the PLIP2-OX and PLIP3-OX plants.

PLIP2 and PLIP3 Contribute to Oxylipin Production in response to ABA
The data presented thus far have only indirectly provided clues towards the biological
functions of PLIP2 and 3. Taking into consideration the severely reduced vegetative growth of the

165

PLIP2-OX and PLIP3-OX plants it seemed plausible that PLIP2 and 3 are possibly only induced
under certain conditions or in specific tissues requiring increases in oxylipin levels. Based on data
available at the Arabidopsis eFP browser (http://bar.utoronto.ca/efp/cgi-bin/efpWeb.cgi) (Winter
et al., 2007), an online Arabidopsis transcriptomic database, PLIP2 and 3 were specifically induced
by ABA treatment as well as under ABA-mediated abiotic stress conditions (Figure 4.16).
Interestingly, PLIP2 and 3 are responsive to different abiotic stressors. Expression of PLIP2 seems
to be specifically induced by cold, while PLIP3 is induced by osmotic and drought stress (Figure
4.16 B and D). To experimental verify ABA induction of PLIP2 and 3, two-week-old WT
seedlings were transferred to MS agar plates containing ABA or the equivalent ethanol solvent as
the Mock treatment (Figure 4.17 A). The expression of PLIP2 and 3, but not of PLIP1, was
elevated within the first two-hours of exposure to ABA, and the response gradually attenuated
during the following 48 hours, confirming that PLIP2 and 3 are transcriptionally induced by ABA
supplementation.
Characterization of the PLIP2-OX and PLIP3-OX plants suggested that the activity the
two proteins gives rise to increased oxylipins levels and signaling. Under natural biological
conditions, the expression of these genes is induced by ABA. In fact, a body of evidence shows
that JA-responsive genes are also induced by ABA or conditions that lead to increased ABA levels,
although mechanistic details of this relationship remain elusive (Adie et al., 2007; Valenzuela et
al., 2016). To test the hypothesis that ABA induction of JA responsive genes is mediated by PLIP
gene activation, we constructed plip triple mutant lines that are deficient in all three PLIP lipases.
The triple mutant allowed us to address their potential functional redundancy. Two independent
T-DNA insertion lines were first obtained for both PLIP2 and 3 (Alonso et al., 2003). These TDNA insertion lines were physiologically indistinguishable from WT under normal growth

166

conditions (Figure 4.18 A), and their leaf lipid profile was indistinguishable from wild type as
well. We further constructed the double mutant between plip2-2 and plip3-1 without showing any
physiological phenotype under normal growth conditions. Since the PLIP3 and PLIP1 genes are
closely linked on the same chromosome, we obtained the PLIP triple mutant using two different
approaches, either with artificial microRNA to reduce the PLIP1 expression, or with the CRISPRCas9 technology to knock out PLIP1 in the plip2-2/3-1 double mutant background. We obtained
14 independent microRNA knock-down lines in total, and the final two representative lines, pliptri1 and plip-tri2 had no expression of PLIP2 and 3, but ~60% reduced expression of PLIP1
(Figure 4.18 B). These plants were also physically nearly identical to the WT plants under normal
growth conditions, and did not exhibit any lipid phenotype (Figure 4.18 C to I). For the CRISPRCas9 approach, we only obtained one homozygous line (plip-CRP) so far. Sequencing results
showed that the plip-CRP plants have a single nucleotide insertion after the 223rd nucleotide in
the PLIP1 coding sequence, causing a frameshift (Figure 4.19).
To test the hypothesis whether PLIPs contribute to oxylipin production in response to
ABA, plants were first grown on filter papers placed on top of the normal MS medium plates for
two weeks. Following transfer to MS agar plates containing 10 µM ABA, quantification by LCMS showed that oxylipins including OPDA, JA and JA-Ile were significantly elevated after 24
hours treatment (Figure 4.17 B to D) in the wild type, but their accumulation was abolished in the
two plip-tri lines and one plip-CRP line (Figure 4.17 B to D), confirming that PLIPs are involved
in ABA induced JA production.

167

PLIP 1,2,3 Triple Mutants are Hypersensitive to ABA
Aside from their inability to produce oxylipins in response to ABA application, plip-tri
mutant germination and seedling establishment were hypersensitive to ABA. With ABA in the
medium, the germination of the plip-tri seeds was severely reduced, although the seeds were able
to germinate within two days without exogenous ABA (Figure 4.20 A). ABA hypersensitivity was
also observed during seedling establishment for the plip-tri plants as they showed reduced growth
and paler rosettes compared with the wild-type seedlings when grown on ABA-containing MS
medium for two weeks (Figure 4.20 B). As established above, PLIP2 and 3 hydrolyze chloroplast
membrane lipids providing 18:3 FA precursors for JA production when induced by ABA.
Furthermore, decreased endogenous ABA levels were observed in the PLIP2-OX and PLIP3-OX
plants which have elevated levels of oxylipins (Figure 4.12 A). To explain these observations, we
hypothesize that JA antagonizes ABA anabolism or signaling, providing a feedback regulatory
loop for the cooperative action of ABA and JA under certain conditions. To investigate how JA
might affect ABA metabolism in WT and PLIP-compromised plants, we grew the plants on regular
MS agar plates for two weeks, and subsequently transferred the plants to Methyl-JA containing
plates. In a time-course assay from 1 to 48 h, endogenous ABA contents in different genotypes
were quantified (Figure 4.12 C). ABA levels generally decreased in all the genotypes, suggesting
JA might repress ABA metabolism. Furthermore, endogenous ABA levels were also found to be
slightly lower in the PLIP compromised plants compared to the WT plants.

168

DISCUSSION
PLIP2 and PLIP3 Share Biochemical Properties with PLIP1
PLIP2 and PLIP3 were discovered based on their sequence similarity to PLIP1. They are
more closely related to PLIP1 compared to other α / β hydrolases in the Arabidopsis genome
(Figure 4.2). All three contain a conserved Lipase 3 domain. Even though we were unable to
produce PLIP3 in in E. coli nor yeast, the PLIP2 recombinant protein was obtained (Figure 4.5).
Further biochemical characterization of PLIP2 showed that it is also a Phospholipase A1 (Figure
4.6). Lipases tend to be promiscuous with regard to offered lipid substrates in vitro. By ectopically
overexpressing the respective lipase cDNAs in the plant, the recombinant lipases under
investigation gain access to their native lipid substrates in a quasi-native environment causing
changes in the lipid composition indicative of their respective substrate preference in vivo.
Lipidomic analysis of the PLIP2-OX and PLIP3-OX plants showed very similar steady-state lipid
changes (Figure 4.7 A, 4.8 and 4.9) compared to that of the previously reported PLIP1-OX plants
(Wang et al., 2017), corroborating their similarity in their enzymatic activities. Especially, the
correlative changes for chloroplast lipids (MGDG and PG) with ER lipids (PI, PE and PC) suggest
that all three PLIPs can efficiently release polyunsaturated FAs that are subsequently exported
from chloroplasts to the ER.
Further Pulse-Chase labeling analysis suggested that different PLIP-OX plants affect
different lipid pools in vivo (Figure 4.7 and 4.10). In the PLIP2-OX plants, labeling and turnover
of the MGDG pool was accelerated. On the other hand, the PG pool was labeled and turned over
at an accelerated rate in the PLIP3-OX plants. These data imply that PLIP2 and PLIP3
preferentially access different substrates in vivo, PLIP2 the galactolipid MGDG and PLIP3 the
phospholipid PG. Biosynthesis of both the plastid assembled MGDG and PG uses phosphatidic

169

acid (PA) as the common precursor. Accelerated degradation of either PG or MGDG pools requires
that more plastid-derived PA, the precursor for both lipids, is diverted to other lipid pools,
eventually indirectly leading to a similar ER lipid phenotype during steady state. Acyl
compositional analysis of MGDG showed reduced 16:3, a signature FA of the plastid-assembled
MGDG in the PLIP2-OX plants, corroborating the hypothesis that prokaryotic MGDG is likely
the native substrate of PLIP2. Although PLIP2 and likely PLIP3 can act on MGDG and PG based
on the available in vitro data for PLIP 1 and 2, their distinct subchloroplastid location might give
s them preferential access to one or the other lipid explaining the observed apparent lipid
preferences in vivo.

PLIP2 and PLIP3 are Involved in JA Biosynthesis
Overexpressing anyone of the three PLIP cDNAs causes a growth defect (Figure 6). This
growth phenotype, however, does not correlate with the steady changes in lipid composition and
hence, simply degradation of the membrane lipid composition due to increased lipase activity
cannot explain this phenotype. Morphologically, the PLIP2-OX and PLIP3-OX plants resemble
plants with activated defense responses (Figure 4.11). Indeed, JA-responsive marker genes were
constitutively induced in the PLIP-OX plants, and their induction levels matched the severity of
the growth reduction (Figure 4.11). Oxylipins including Arabidopsides accumulated in the PLIP2OX and PLIP3-OX plants, (Figure 4.12). Arabidopsides were previously observed at very high
levels in vegetative tissues after wounding or during the pathogen-induced hypersensitive response
(Andersson et al., 2006; Buseman et al., 2006). Even though some of the Arabidopside species
have been shown to directly participate in plant defense (Kourtchenko et al., 2007), they are
generally thought to provide a temporary, inert storage pool for OPDA (Mosblech et al., 2009).

170

The biosynthesis of Arabidopsides is still a matter of debate, but in the PLIP2-OX and PLIP3-OX
plants, Arabidopsides possibly serve to sequester the levels of free OPDA as a result of increased
18;3 release by the PLIPs. After PLIP2 or PLIP3 release the polyunsaturated FAs, free OPDA
readily accumulates in the plastids possibly by oxidation of 18:3. OPDA is structurally a fatty acid,
and may be detrimental to the cell in high quantities simply due its detergent properties.
Presumably, a fraction of the free OPDA is converted to OPDA-Acyl Carrier Protein (ACP), which
serves as substrate for the plastid acyl transferases that either assemble membrane lipids de novo
or catalyze acyl exchange on existing membrane lipids giving rise to the observed Arabidopsides.
This mechanism can also likely provide feedback regulation for the fine tuning of JA biosynthesis
and signaling along with JAZ repressors and JA catabolism (Figure 4.11 and 4.12).
Finally, to examine whether the growth repression phenotype was specifically caused by
JA metabolism and signaling, and no other factors, the PLIP2-OX and PLIP3-OX plants were
crossed with a coi1 mutant allele, which abolishes JA perception in plants (Figure 4.14).
Homozygous coi1 offspring in the PLIP-OX backgrounds reversed their stunted growth without
affecting the lipid phenotype. Taken together, these data led to our hypothesis that PLIP2 and
PLIP3 might be involved in initiating JA metabolism under conditions when they are
overproduced, for example, following activation of their respective genes in response to
environmental cues.

PLIP2 and PLIP3 Link ABA Signaling with JA Biosynthesis
As the three PLIPs possess similar biochemically characteristics, the question arises how
they exert their distinct physiological roles. One possible explanation is their different subplastidic
locations. PLIP1 is specifically associated with the thylakoid membranes (Wang et al., 2017).

171

Based on our current analysis, PLIP3 is a membrane protein associated with both the envelope and
thylakoid membranes, while PLIP2 is found in the soluble and membrane fractions, and seems to
be ubiquitously present in envelope membranes, stroma and thylakoids (Figure 4.3). Possibly,
PLIP2 and PLIP3 are enriched in microenvironments that are in close proximity to other enzymes
required for oxylipin synthesis, like lipoxygenase, Allene Oxide Synthase and Allene Oxide
Cyclase to readily convert free FAs to oxylipins.
Another possible factor that could lead to distinct physiological functions might be the
differing transcriptional regulation of their respective genes. PLIP1 is specifically expressed
during seed developmental stages, and has much lower basal expression levels in the leaf tissues
compared to PLIP2 and PLIP3. In addition, the expression of PLIP2 and PLIP3 in leaf tissues is
ABA responsive, but that of PLIP1 is not (Figure 4.16 and 4.17). Moreover, despite the fact that
both PLIP2 and 3 can be induced by applied ABA, public transcriptomic data show that they are
induced by different abiotic stresses in vivo (Figure 4.16), implying that each of the two genes
contain their own unique cis-elements in their promoters that can be recognized by different
transcription factors sensing different stressors.
Having roles in both ABA responses and JA biosynthesis, PLIP2 and PLIP3 we
hypothesize that they can mediate ABA induction of JA biosynthesis. Given the relatedness of
their biochemical properties and subtle phenotypes of the individual mutants, triple PLIP1, 2, 3
mutants were generated. Two approaches, artificial microRNA and CRISPR-Cas9, were
successful in overcoming the the close genetic linkage between PLIP2 and PLIP3 (Figure 4.18
and 4.19). All plip triple mutant lines exhibited compromised oxylipin biosynthesis in response to
applied ABA (Figure 4.17), corroborating their roles in mediating ABA responsiveness of JA
biosynthesis. JA production is most commonly induced by wounding damage due to herbivore

172

insects or necrotrophic microbes (Creelman et al., 1992; Trusov et al., 2006). Furthermore, a
growing body of evidence suggests that JA biosynthesis is also stimulated by applying ABA to
plants or during abiotic stress conditions that lead to ABA accumulation (Creelman and Mullet,
1995; Adie et al., 2007; Avramova, 2017). The transcription factor MYC2 has been proposed as a
potential component in linking ABA and JA mediated physiological responses given its
involvement in both JA perception and ABA signaling (Kazan and Manners, 2013). So far, it is
still unknown how or even whether MYC2 contributes to ABA induced JA production. In the
future, it would be interesting to investigate whether MYC2 is directly involved in the
transcriptional induction of PLIP2 or PLIP3, to determine whether MYC2 and PLIPs work in the
same pathway. It is also important to study each PLIP gene’s unique timing and its range of
responsiveness to different environmental cues. In nature, plants continually encounter different
adverse environmental conditions. Possibly, the PLIP genes evolved allowing plants to cope with
different abiotic / biotic stresses and integrate their protective responses for example during
coinciding drought conditions and insect outbreaks (Suzuki, 2016).
Aside of their inability to induce JA biosynthesis in response to ABA application, the plip
triple mutants were also hypersensitive to ABA during germination and early seedling
development (Figure 4.20). Application of JA treatment to the WT and the plip triple mutant
seedlings led to a trend of gradually decreasing ABA levels in all the genotypes overtime (Figure
4.20). Decreased ABA content was also observed in the PLIP2-OX plants (Figure 4.12).
Therefore, one possible explanation for the ABA hypersensitivity phenotype might be that JA
negatively regulates ABA biosynthesis. However, other possibilities cannot be precluded at this
point. For example, besides ABA metabolism, it is important to investigate whether PLIP2 and
PLIP3 are directly or indirectly involved in ABA signaling. It is possible that other oxylipin

173

products as a result of PLIP2 and PLIP3 activation serve as negative regulators of ABA signaling,
a possibility that still needs to be further explored.

174

ACKNOWLEDGEMENTS
This work was done in collaboration with several collaborators. John E. Froehlich
conducted the chloroplast import assay in Figure 4.3. Hope L. Hersh generated the plip-CRISPR
lines in Figure 4.19. I would also like to thank Dr. Ronghui Pan and Dr. Jianping Hu, Michigan
State University for providing the artificial microRNA constructs; Dr. Sheng Yang He, Michigan
State University and Dr. Jian-Kang Zhu, Purdue University for providing the CRISPR-Cas9
constructs and instructions; Dr. Anthony Schilmiller, Michigan State University for assisting with
the plant hormone measurement and oxylipin profiling assays. We would also like to thank Dr.
Koichi Sugimoto, Michigan State University for generally sharing the standards of JA-Ile, 12OHJA and 12OH-JA-Ile; Dr. Jian Yao in Sheng Yang He lab for providing the coi1-30 mutant seeds,
Michigan State University for helpful discussions and suggestions.

175

APPENDIX

176

Primer
PEG_PLIP2 F
PEG_PLIP2 R

Sequence 5’ – 3’

pET41a_His del R
PLIP2_S428A F
PLIP2_S428A R
PLIP2_qPCR F
PLIP2_qPCR R
PLIP3_qPCR F
PLIP3_qPCR R
TIP-41_like F
TIP-41_like R
UBQ10 qPCR F
UBQ10 qPCR R

CACCATGGACAGTTTGTGTTTGAA T
TCAACGGAAACTAAAGACGTTGA
CACCATGGAGGGTGTTTTCTTAAAA
AT
TCACTTGAAGAAACTAAAATGTGCT
TG
TCGGATCCATGGACAGTTTGTGTTT
GAATAG
GACTCGAGTCAACGGAAACTAAAG
ACGTT
TCGGATCCATGGACAGTTTGTGTTT
GAATAG
GAGTCGACTCACTTGAAGAAACTA
AAATGTGC
GCCATGGACAGTTTGTGTTTGAATA
G
ACGGAAACTAAAGACGTTGAAAGC
GCCATGGAGGGTGTTTTCTTAAAAA
T
CTTGAAGAAACTAAAATGTGCTTGT
GA
ATGTATATCTCCTTCTAAAGTAAAC
AAA
ATGGCGTTTAATACGGCTATG
ACCGGTCATGCATTAGGTGGA
GAAACGGAATTTAGCGCTGGT
AGCGACAGATCGGCTTATGG
GCGATGCTCCCTTTTTGCTC
CAGATGTTTGCCAAATTGTCGT
TCCTCGCCATTGTTGTTCTC
GTGAAAACTGTTGGAGAGAAGCAA
TCAACTGGATACCCTTTCGCA
TCCGGATCAGCAGAGGCTTA
TCAGAACTCTCCACCTCAAG

LBb1.3

ATTTTGCCGATTTCGGAAC

plip2-1 LP
plip2-1 RP
plip2-2 LP
plip2-2 RP

CACAATCCACTTCGTCTCCTC
CAACAGTAGAAGACAAGGCGC
AAATCGCTAGCTATGCTGCAG
TCCAATAACGGTTAAGCAACG

PEG_PLIP3 F
PEG_PLIP3 R
BamHI_PLIP2 F
PLIP2_XhoI R
BamHI_PLIP3 F
PLIP3_SalI R
PLIP2_pYes2.1 F
PLIP2_pYes2.1 R
PLIP3_pYes2.1 F
PLIP3_pYes2.1 R
pET41a_His del F

Table 4. 1. Primer sequences used in this study.

177

Purpose
For pEarlyGate 101
For pEarlyGate 101
For pEarlyGate 101
For pEarlyGate 101
For the pET41a to produce
PLIP2 in E. coli

For the pET41a to produce
PLIP3 in E. coli
For the pYes2.1 to produce
PLIP2 in yeast
For the pYes2.1 to produce
PLIP3 in yeast
pET41a construct for
chloroplast import
S428A point mutation
For qPCR analysis of PLIP2
expression
For qPCR analysis of PLIP3
expression
qPCR reference gene
qPCR reference gene
For genotyping of plip2 and 3
mutants
For genotyping of plip2-1
For genotyping of plip2-2

Table 4. 1. (cont’ed)
AAAGGAAGATTTGATCGGACG
plip3-1 LP
plip3-1 RP
plip3-2 LP
plip3-2 RP

JAZ1 F
JAZ1 R
JAZ10.1-3 F

CAACAGTAGAAGACAAGGCGC
TTACCAGAAGTTCATGCCCAC
AAATCTCTTGCATTTCCTCGC
gaTCTTAAAACGTCATTTGCCGGctctc
ttttgtattcca
agCCGGCAAATGACGTTTTAAGAtcaa
agagaatcaatga
agCCAGCAAATGACGATTTAAGTtcac
aggtcgtgatatg
gaACTTAAATCGTCATTTGCTGGctaca
tatatattccta
AGAGCTGGAGCTGGTGTTGTTAAG
GAACACCCATTCCGGTAACACCAT
GGGCGTACTGGTCGTGGTTA
TCCCGAGTTCCAAGAGGTTTT
AGAAACTCCAAATCAAGAACCAGC
TC
CCGGTTTAATCGAAGAACACGAAG
AC
GCACCGCTAATAGCTTAGCCAAGA
TCTGTCAATGGTGTTGGGGAGGAT
AGAGCTGGAGCTGGTGTTGTTAAG

JAZ10.1-3 R

GAACACCCATTCCGGTAACACCAT

CYP94C1 F

TCTCACTCTTTCACTTACTCTTCC

CYP94C1 R

CGTTTGATGGATTCGCTGTG

coi1-30 F
coi1-30 R
pROK2-LB3
PLIP1 CPR F
PLIP1 CPR R
PLIP1 CPR_geo F
PLIP1 CPR_geo R

atggaggatcctgatatcaag
AGTCACTGACAATCATCCGTCG
TTGGGTGATGGTTCACGTAG
gattGATGAAAACGAGTCGATCTG
aaacCAGATCGACTCGTTTTCATC
ATCGAGTTCCGACAATCACTTGT
GACGTGTCCGTTTCAAATAGCAAC

PLIP1-miRNA I
PLIP1-miRNA II
PLIP1-miRNA III
PLIP1-miRNA IV
VSP1 F
VSP1 R
LOX2 F
LOX2 R
MYC2 F
MYC2 R

178

For genotyping of plip3-1

For genotyping of plip3-2

For knocking down PLIP1
with artificial microRNA

For qPCR analysis of VSP1
(At5g24780) expression
For qPCR analysis of LOX2
(At3g45140) expression
For qPCR analysis of MYC2
(At1g32640) expression
For qPCR analysis of JAZ1
(At1g19180) expression
For qPCR analysis of
JAZ10.1, 10.2 and 10.3
(At5g13220) expression
For qPCR analysis of
CYP94C1 (AT2G27690)
expression
For genotyping the coi1-30
locus
For knock out PLIP1 with
CRISPR-Cas9
For genotyping of the
CRISPR-Cas9 lines

Figure 4. 1. Time course of PLIP2 lipase activity with phosphatidylcholine (PC) as substrate.
As substrate, 120 µg 18:1/18:1-PC was provided to microsomal recombinant PLIP2 and the lipase
assay was stopped at times indicated. Lipids were subsequently extracted and analyzed by TLC.
Remaining PC and produced lyso-PC were isolated and their acyl contents were quantified as
methyl esters by liquid gas chromatography. Fractions of PC degradation at different times is
calculated as 2 (molarity of lyso-PC acyl groups) / (2 (molarity of lyso-PC acyl groups) + (molarity
of PC acyl groups)) *100.

179

Figure 4. 2. Phylogenetic analysis of the top 20 PLIP1 similar sequences in Arabidopsis.
The phylogenetic tree was built using the Maximum Likelihood method with PLIP1and the top 20
Arabidopsis similar protein sequences identified from the BLASTp search. Sequences alignments
are shown in Supplemental File 1. Bootstrap values (based on 1000 replicates) are indicated at the
tree nodes. The scale measures evolutionary distances in substitution per amino acid.

180

Figure 4. 3. Subcellular localization of PLIP2 and PLIP3 in Arabidopsis.
(A) Subcellular localization of YFP tagged PLIP2 and PLIP3 in leaf mesophyll cells of 3-weekold Arabidopsis Col-0 transformed with PLIP2-YFP or PLIP2-YFP driven by the 35S promoter,
or empty vector (EV) control using confocal laser scanning microscopy. Chlorophyll
autofluorescence is shown in red, and YFP fluorescence is shown in yellow. Overlay of
chlorophyll and YFP are shown as well (Merge). Representative images from one experiment are
presented. Scale bars: 5 µm.
(B) Chloroplast import experiments with radiolabeled PLIP2 and PLIP3. Chloroplasts were treated
with (+) or without (-) trypsin. Total chloroplast membranes (P) or soluble (S) fractions were
analyzed by SDS-PAGE followed by fluorography. TP, translation products; p, precursor; m,
mature form; MW, molecular weight markers.

181

Figure 4. 4. Alignment of PLIP2 (A) and PLIP3 (B) with class 3 lipase protein sequences
available in the NCBI conserved domain database (CDD).
The alignment result was produced by blasting the PLIP2 or PLIP3 protein sequence against the
CDD database using CD-search (https://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi) with
default settings. Reference protein names are represented as either Protein Data Bank ID (e.g.,
1TIA and 3TGL) or as NCBI GI number. The conserved catalytic triads are highlighted in yellow,
and the non-conserved in grey.
182

Figure 4. 5. Production of the recombinant PLIP2 proteins and its lipase activity in vitro.
(A) Detection of PLIP2 and PLIP2S428A protein by immunoblotting against the N-terminus 6 × His
tag (top panel) and Coomassie Brilliant Blue (CBB) staining (bottom panel) in the lysed E. coli
cultures with or without isopropyl β-D-1-thiogalactopyranoside induction. The E. coli strain
carrying the empty vector (EV) was included as a negative control.
(B) Thin-layer chromatogram of products of a representative in vitro lipase reaction using PC with
E. coli microsomes carrying the wild-type (PLIP2 + PC), or the mutant enzyme (PLIP2S428A + PC).
Substrate without enzyme (PC) or with E. coli microsome carrying the EV control (EV + PC) were
included as controls. PC, phosphatidylcholine. O, origin of sample loading. Lipids were visualized
by iodine vapor staining.

183

Figure 4. 6. In vitro lipase activity of PLIP2.
(A) Gas-liquid chromatograms of methyl esters derived from defined PC substrates or lyso-PC
fractions from PLIP2 lipase reactions with different PC substrates. 15:0 was used as an internal
standard.
(B) PLIP2 lipase activity on defined PC substrates (carbon number : double bond number; sn-1/sn2) with different degree of saturation of the sn-1 acyl groups. PC containing 18:0/18:1 and
18:1/18:1 and PC containing 18:0/18:2 and 18:2/18:2 were compared, respectively. n = 4, ± SD.
Student’s t-test was applied (**indicates p<0.01; n.s., not significant).

184

Figure 4. 7. The steady-state and dynamic lipid phenotypes of the PLIP2-OX plants.
(A) Relative acyl composition of monogalactosyldiacylglycerol (MGDG) in PLIP2-OX and empty
vector (EV) control lines. Leaf samples harvested from one plant were pooled as one biological
repeat; n = 4, ± SD. Student’s t-test was applied to compare the empty control (EV) control plants
with each of the two PLIP2-OX plants (**indicates p<0.01).
(B) and (C) In vivo pulse-chase acetate labeling of lipids in the EV control and the PLIP2-OX1
plants. The [14C]-acetate labeling pulse lasted for 60 min. The fractions of label in all polar lipids
were calculated as percentages of total incorporation of label in polar lipids (B). After the pulse,
[14C]-acetate medium was replaced with non-labeled free acetate to initiate the chase with a
duration of three days (C). The fractions of label in all polar lipids is given during time course.
Experiments were repeated three times with similar results and one representative result is shown.
n = 3, ± SD. MGDG, monogalactosyldiacylglycerol; PC, phosphatidylcholine; PE,
phosphatidylethanolamine; PG, phosphatidylglycerol.

185

Figure 4. 8. Analysis of polar lipids from the 4-week-old soil-grown PLIP2-OX and empty
vector (EV) control plant leaves.
(A) Relative abundance of major polar lipid classes in PLIP2-OX and EV control lines.
(B-F) Relative acyl group composition of PG, DGDG, SQDG, PI, PE and PC supplemental to data
shown in Figure 5A. n = 4, ± SD. Student’s t-test was applied (** indicates p<0.01). DGDG,
digalactosyldiacylglycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG,
phosphatidylglycerol; PI, phosphatidylinositol; SQDG, sulfoquinovosyldiacylglycerol. Acyl
groups with a molar percentage less than 0.5% were omitted.

186

Figure 4. 9. Analysis of polar lipids from 4-week-old soil-grown PLIP3-OX and empty
vector (EV) control plant leaves.
(A) Relative abundance of major polar lipid classes in PLIP3-OX and EV control lines.
(B-G) Relative acyl group composition of MGDG, PG, DGDG, SQDG, PI, PE and PC. n = 4, ±
SD. Student’s t-test was applied (** indicates p<0.01). DGDG, digalactosyldiacylglycerol;
MGDG, monogalactosyldiacylglycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine;
PG, phosphatidylglycerol; PI, phosphatidylinositol; SQDG, sulfoquinovosyldiacylglycerol. Acyl
groups with a molar percentage less than 0.5% were omitted.
187

Figure 4. 10. In vivo pulse-chase acetate labeling of polar lipids in the empty vector (EV)
control and the PLIP3-OX plants.
(A) Radioactivity in the lipids at 40 min of the pulse phase. The [14C]-acetate labeling pulse lasted
for 60 min. Detached leaves of 4-week-old plants were used. The fractions of label in individual
polar lipid were calculated as percentages of total incorporation of label in polar lipids.
(B) Chase of the polar lipids in the EV control and the PLIP3-OX plants. After the pulse, [14C]acetate medium was replaced with non-labeled free acetate to initiate the chase with a duration of
24 hours. The fractions of label in all polar lipids were determined during a time course.
Experiments were repeated three times with similar results and one representative result is shown.
n = 3, ± SD. MGDG, monogalactosyldiacylglycerol; PC, phosphatidylcholine; PE,
phosphatidylethanolamine; PG, phosphatidylglycerol.

188

Figure 4. 11. Morphology of the PLIP-OX plants.
(A) Growth of 4-week-old soil-grown plants. One empty vector (EV) control line, two lines of
PLIP1-OX, PLIP2-OX and PLIP3-OX plants are shown. The relative mol % fraction of PG and
MGDG in each line are indicated. n = 4, ± SD.
(B) Morphology of the 24-day-old EV and PLIP2-OX1 plants grown on MS agar medium. Inserts
show representative individual plant for details.
(C) Phase separation of the lipid extracts from the EV and PLIP2-OX plants in (B). Three
independent extracts are shown for each genotype. The upper aqueous phases accumulating
anthocyanin are indicated by an arrow. The image was taken with the plastic reaction tubes inserted
into transparent glass tubes on a rack.
(D) Expression of JA responsive genes in the EV and the PLIP-OX plants. Expression levels of
indicated genes were determined by quantitative RT-PCR and normalized to the levels in the EV
plants as fold changes. Analysis of JAZ10 includes JAZ10.1, 10.2 and 10.3, but not 10.4 splicing
forms; n = 3, ± SD. Student’s t-test was applied to compare the level in the EV plants and every
other genotype (**indicates p<0.01).

189

Figure 4. 12. Phytohormone quantification and comparative oxylipin profiling in the WT
and the PLIP2-OX plants.
(A) Plants were grown on soil for three weeks before the leaves were harvested for hormone
extraction and quantification by LC-MS/MS. The two y-axes apply to compounds with different
order of magnitudes in quantity on the left and right panels, respectively. n = 4, ± SD. Student’s ttest was applied to compare the level in the EV plants and to each other PLIP2-OX line
(**indicates p<0.01).
(B) Partial metabolite profiling chromatographs from three-week-old wild-type (WT) and PLIP2OX1 plant leaves. Numbers indicate the two representative metabolite peaks that are elevated in
the PLIP2-OX1 plants compared to WT. 1 represents Arabidopside A, 2 Arabidopside B.
(C) Relative amount of the three major oxylipins accumulating in the PLIP2-OX1 plant identified
by comparative metabolite profiling in (B). The indicated compound levels were normalized to the
18:3/18:3 digalactosyldiacuylglycerol which remains unchanged in WT or the PLIP2-OX plants.
n = 4, ± SD. Student’s t-test was applied (**indicates p<0.01).

190

Figure 4. 13. Phytohormone quantification and oxylipin profiling in the wild-type (WT)
and the PLIP3-OX plants.
(A) Plants were grown on soil for three weeks before the leaves were harvested for compound
extraction and quantification by LC-MS/MS. n = 4, ± SD. Student’s t-test was applied to compare
the levels in the EV plants and each of the PLIP3-OX lines (**indicates p<0.01).
(B) Relative amounts of the three major classes of Arabidopsides in the WT and the PLIP2-OX1
plant leaves identified by comparative metabolite profiling. The indicated compound levels were
normalized to the 18:3/18:3 digalactosyldiacuylglycerol which remains unchanged in WT or the
PLIP3-OX plants. n = 4, ± SD. Student’s t-test was applied (**indicates p<0.01).

191

Figure 4. 14. Rescue of growth in PLIP2-OX coi1 homozygous plants.
(A) Image of 4-week-old plants grown on soil and the rosette diameter of the indicated plants
below the corresponding images. n = 6, ± SD. one-way ANOVA with post-hoc Turkey HSD test
was applied. Rosette diameters indicated by different letters are significantly different (p<0.01).
(B) Relative acyl group compositions of leaf monogalactosyldiacylglycerol in the indicated plants
in (A). Leaves harvested from one plant were pooled as one biological repeat. n = 3, ± SD.
Student’s t-test was applied to compare the wild type (WT) with each of the remaining genotypes
(**indicates p<0.01).

192

Figure 4. 15. Phenotypes of PLIP3-OX coi1 plants.
(A) Image of the 4-week-old plants grown on soil and the rosette diameter of the indicated plants.
n = 6, ± SD. one-way ANOVA with post-hoc Turkey HSD test was applied. Rosette diameters
indicated by different letters are significantly different (p<0.01).
(B) Relative acyl group compositions of leaf phosphatidylglycerol (PG) and phosphatidylcholine
(PC) in the indicated plants in (A). Leaves harvested from one plant were pooled as one biological
repeat. n = 3, ± SD. Student’s t-test was applied to compare the wild type (WT) with each of the
remaining genotypes (**indicates p<0.01).

193

Figure 4. 16. Transcriptional induction of PLIP2 and PLIP3 as shown in the eFP database.
(A) and (B) Absolute expression values of PLIP2 in response to ABA treatment (A) or under cold
stresses (B).
(C) and (D) Absolute expression values of PLIP3 in response to ABA treatment (C) or under salt
and osmotic stresses (B).
Data were extracted from the Arabidopsis eFB browser
(http://bar.utoronto.ca/efp/cgi-bin/efpWeb.cgi).

194

Figure 4. 17. Induction of PLIP transcription by ABA.
(A) Expression of PLIP genes in response to 7µM ABA or equivalent ethanol solution (Mock)
treatment in 2-week-old seedlings determined by quantitative RT-PCR during a time course as
indicated.
(B) to (D) Oxylipin contents in the wild-type (WT), two plip-tri lines and one plip-CRP line before
and after 24-hour 7µM ABA additon. n = 3, ± SD. Student’s t-test was applied to compare (WT)
with each of the remaining genotypes (**indicates p<0.01).

195

Figure 4. 18. Growth and the lipid phenotype of the plip single and triple mutants.

196

Figure 4. 18. (cont’d)
(A) Morphology of the 4-week-old soil-grown wild type (WT) and two T-DNA insertional alleles
of plip2 and plip3, respectively. The T-DNA insertional line SALK123458 was designated as
plip2-1, SALK134251C as plip2-2, SALK006633 as plip3-1, SALK 006633 as plip3-2.
(B) Morphology of the 5-week-old soil-grown empty vector (EV) control and two independent
plip-tri mutant lines and the expression of PLIPs in these plants. The plip-tri lines were created by
reducing PLIP1 expression by artificial microRNA approach in the plip2 plip3 double mutant. The
expression levels of PLIPs were normalized to those in the EV control (set to 1).
(C) Relative abundance of major polar lipid classes in EV control and the plip-tri lines.
(D-I) Relative acyl group composition of PG, MGDG, SQDG, DGDG, PC, PI and PE. n = 3, ±
SD. Student’s t-test was applied (** indicates p<0.01). For lipid abbreviations, refer to the legend
of Figure 4.9.

197

Figure 4. 19. Genotyping of the plip-CRISPR line.
(A) Digestion patterns of the PLIP1 amplicon from the empty vector (EV) control and the plipCRP genomic DNAs. The PLIP1 amplicon was digested (+) with Dpn II or not (-). The wild type
PLIP1 amplicon (178 bp) is expected to be cut into two fragments (93bp and 85bp).
(B) Sequencing of the PLIP1 locus from the plip-CRP plants revealed a single nucleotide insertion
between the 223rd and 224th nucleotide of the coding sequence, leading to frameshift of the PLIP1
sequence. The inserted nucleotide is highlighted in yellow, and the PAM motif is highlighted in
blue.

198

Figure 4. 20. The plip-tri plants are hypersensitive to ABA.
(A) Germination rates of the wild-type and plip-tri seeds with or without 2 µM ABA addition on
phytoagar plates. Observation of 100 seeds in one plate was treated as one biological repeat. n =
4, ± SD. Student’s t-test was applied to compare (WT) with each of the remaining genotypes
(**indicates p<0.01).
(B) Morphology of the 2-week-old WT and plip-tri seedlings grown on MS agar plates with 0.5µM
ABA or the equivalent ethanol solvent (Mock). Scale bars: 0.5 cm.
(C) ABA content in the 2-week-old WT, plip-tri and plip-CRP seedlings during a 48-h time course
follwoing the addition of 10 µM Methyl-JA to the MS agar medium. 3-5 seedlings were pooled as
one biological repeat. n = 3, ± SD. Student’s t-test was applied to compare (WT) with each of the
remaining genotypes (**indicates p<0.01).

199

REFERENCES

200

REFERENCES

Acosta, I.F., and Farmer, E.E. (2010). Jasmonates. Arabidopsis book 8, e0129-e0129.
Adie, B.A., Perez-Perez, J., Perez-Perez, M.M., Godoy, M., Sanchez-Serrano, J.J., Schmelz,
E.A., and Solano, R. (2007). ABA is an essential signal for plant resistance to pathogens
affecting JA biosynthesis and the activation of defenses in Arabidopsis. Plant Cell 19,
1665-1681.
Ajjawi, I., Lu, Y., Savage, L.J., Bell, S.M., and Last, R.L. (2010). Large-scale reverse genetics
in Arabidopsis: case studies from the Chloroplast 2010 Project. Plant Physiol 152, 529540.
Alonso, J.M., Stepanova, A.N., Leisse, T.J., Kim, C.J., Chen, H., Shinn, P., Stevenson, D.K.,
Zimmerman, J., Barajas, P., Cheuk, R., Gadrinab, C., Heller, C., Jeske, A.,
Koesema, E., Meyers, C.C., Parker, H., Prednis, L., Ansari, Y., Choy, N., Deen, H.,
Geralt, M., Hazari, N., Hom, E., Karnes, M., Mulholland, C., Ndubaku, R., Schmidt,
I., Guzman, P., Aguilar-Henonin, L., Schmid, M., Weigel, D., Carter, D.E.,
Marchand, T., Risseeuw, E., Brogden, D., Zeko, A., Crosby, W.L., Berry, C.C., and
Ecker, J.R. (2003). Genome-wide insertional mutagenesis of Arabidopsis thaliana.
Science 301, 653-657.
Andersson, M.X., and Dörmann, P. (2009). Chloroplast Membrane Lipid Biosynthesis and
Transport. In The Chloroplast, A.S. Sandelius and H. Aronsson, eds (Berlin, Heidelberg:
Springer Berlin Heidelberg), pp. 125-158.
Andersson, M.X., Hamberg, M., Kourtchenko, O., Brunnstrom, A., McPhail, K.L.,
Gerwick, W.H., Gobel, C., Feussner, I., and Ellerstrom, M. (2006). Oxylipin profiling
of the hypersensitive response in Arabidopsis thaliana. Formation of a novel oxophytodienoic acid-containing galactolipid, arabidopside E. J Biol Chem 281, 3152831537.
Avramova, Z. (2017). The jasmonic acid-signalling and abscisic acid-signalling pathways cross
talk during one, but not repeated, dehydration stress: a non-specific 'panicky' or a
meaningful response? Plant Cell Environ 40, 1704-1710.
Benning, C. (2009). Mechanisms of lipid transport involved in organelle biogenesis in plant
cells. Annu Rev Cell Dev Biol 25, 71-91.
Benning, C. (2010). The Anionic Chloroplast Membrane Lipids: Phosphatidylglycerol and
Sulfoquinovosyldiacylglycerol. In The Chloroplast: Biochemistry, Molecular Biology
and Bioengineering., C.A. Rebeiz, C. Benning, H. Bohnert, H. Daniell, B. Green, K.
Hoober, H. Lichtenthaler, A. Portis, and B. Tripathy, eds (Netherlands: Springer), pp.
171-184.
201

Blee, E. (1998). Phytooxylipins and plant defense reactions. Prog Lipid Res 37, 33-72.
Boudière, L., Michaud, M., Petroutsos, D., Rébeillé, F., Falconet, D., Bastien, O., Roy, S.,
Finazzi, G., Rolland, N., Jouhet, J., Block, M.A., and Maréchal, E. (2014).
Glycerolipids in photosynthesis: Composition, synthesis and trafficking. Biochim
Biophys Acta 1837, 470-480.
Buseman, C.M., Tamura, P., Sparks, A.A., Baughman, E.J., Maatta, S., Zhao, J., Roth,
M.R., Esch, S.W., Shah, J., Williams, T.D., and Welti, R. (2006). Wounding
stimulates the accumulation of glycerolipids containing oxophytodienoic acid and dinoroxophytodienoic acid in Arabidopsis leaves. Plant Physiol 142, 28-39.
Campos, M.L., Yoshida, Y., Major, I.T., de Oliveira Ferreira, D., Weraduwage, S.M.,
Froehlich, J.E., Johnson, B.F., Kramer, D.M., Jander, G., Sharkey, T.D., and Howe,
G.A. (2016). Rewiring of jasmonate and phytochrome B signalling uncouples plant
growth-defense tradeoffs. Nat Commun 7, 12570.
Clough, S.J., and Bent, a.F. (1998). Floral dip: a simplified method for Agrobacteriummediated transformation of Arabidopsis thaliana. Plant J 16, 735-743.
Creelman, R.A., and Mullet, J.E. (1995). Jasmonic acid distribution and action in plants:
regulation during development and response to biotic and abiotic stress. Proc Natl Acad
Sci U S A 92, 4114-4119.
Creelman, R.A., Tierney, M.L., and Mullet, J.E. (1992). Jasmonic acid/methyl jasmonate
accumulate in wounded soybean hypocotyls and modulate wound gene expression. Proc
Natl Acad Sci U S A 89, 4938-4941.
Denness, L., McKenna, J.F., Segonzac, C., Wormit, A., Madhou, P., Bennett, M.,
Mansfield, J., Zipfel, C., and Hamann, T. (2011). Cell wall damage-induced lignin
biosynthesis is regulated by a reactive oxygen species- and jasmonic acid-dependent
process in Arabidopsis. Plant Physiol 156, 1364-1374.
Earley, K.W., Haag, J.R., Pontes, O., Opper, K., Juehne, T., Song, K., and Pikaard, C.S.
(2006). Gateway-compatible vectors for plant functional genomics and proteomics. Plant
J 45, 616-629.
Ellis, C., Karafyllidis, I., Wasternack, C., and Turner, J.G. (2002). The Arabidopsis mutant
cev1 links cell wall signaling to jasmonate and ethylene responses. Plant Cell 14, 15571566.
Emanuelsson, O., Nielsen, H., Brunak, S., and von Heijne, G. (2000). Predicting subcellular
localization of proteins based on their N-terminal amino acid sequence. J Mol Biol 300,
1005-1016.

202

Feys, B., Benedetti, C.E., Penfold, C.N., and Turner, J.G. (1994). Arabidopsis mutants
selected for resistance to the phytotoxin coronatine are male sterile, insensitive to methyl
jasmonate, and resistant to a bacterial pathogen. Plant Cell 6, 751-759.
Frentzen, M. (1986). Biosynthesis and Desaturation of the Different Diacylglycerol Moieties in
Higher Plants. J Plant Physiol 124, 193-209.
Frentzen, M., Heinz, E., McKeon, T.A., and Stumpf, P.K. (1983). Specificities and
selectivities of glycerol-3-phosphate acyltransferase and monoacylglycerol-3-phosphate
acyltransferase from pea and spinach chloroplasts. Eur J Biochem 129, 629-636.
Gobel, C., and Feussner, I. (2009). Methods for the analysis of oxylipins in plants.
Phytochemistry 70, 1485-1503.
Gobel, C., Feussner, I., Hamberg, M., and Rosahl, S. (2002). Oxylipin profiling in pathogeninfected potato leaves. Biochim Biophys Acta 1584, 55-64.
Heitz, T., Widemann, E., Lugan, R., Miesch, L., Ullmann, P., Desaubry, L., Holder, E.,
Grausem, B., Kandel, S., Miesch, M., Werck-Reichhart, D., and Pinot, F. (2012).
Cytochromes P450 CYP94C1 and CYP94B3 catalyze two successive oxidation steps of
plant hormone Jasmonoyl-isoleucine for catabolic turnover. J Biol Chem 287, 6296-6306.
Huot, B., Yao, J., Montgomery, B.L., and He, S.Y. (2014). Growth-defense tradeoffs in plants:
a balancing act to optimize fitness. Mol Plant 7, 1267-1287.
Hurlock, A.K., Roston, R.L., Wang, K., and Benning, C. (2014). Lipid trafficking in plant
cells. Traffic 15, 915-932.
Hyun, Y., Choi, S., Hwang, H.J., Yu, J., Nam, S.J., Ko, J., Park, J.Y., Seo, Y.S., Kim, E.Y.,
Ryu, S.B., Kim, W.T., Lee, Y.H., Kang, H., and Lee, I. (2008). Cooperation and
functional diversification of two closely related galactolipase genes for jasmonate
biosynthesis. Dev Cell 14, 183-192.
Ishiguro, S., Kawai-Oda, A., Ueda, J., Nishida, I., and Okada, K. (2001). The DEFECTIVE
IN ANTHER DEHISCIENCE gene encodes a novel phospholipase A1 catalyzing the
initial step of jasmonic acid biosynthesis, which synchronizes pollen maturation, anther
dehiscence, and flower opening in Arabidopsis. Plant Cell 13, 2191-2209.
Kazan, K., and Manners, J.M. (2013). MYC2: the master in action. Mol Plant 6, 686-703.
Kelly, A.A., and Feussner, I. (2016). Oil is on the agenda: Lipid turnover in higher plants.
Biochim Biophys Acta 1861, 1253-1268.
Kourtchenko, O., Andersson, M.X., Hamberg, M., Brunnstrom, A., Gobel, C., McPhail,
K.L., Gerwick, W.H., Feussner, I., and Ellerstrom, M. (2007). Oxo-phytodienoic acid-

203

containing galactolipids in Arabidopsis: jasmonate signaling dependence. Plant Physiol
145, 1658-1669.
Liu, N., Staswick, P.E., and Avramova, Z. (2016). Memory responses of jasmonic acidassociated Arabidopsis genes to a repeated dehydration stress. Plant Cell Environ 39,
2515-2529.
Marchler-Bauer, A., Derbyshire, M.K., Gonzales, N.R., Lu, S., Chitsaz, F., Geer, L.Y.,
Geer, R.C., He, J., Gwadz, M., Hurwitz, D.I., Lanczycki, C.J., Lu, F., Marchler,
G.H., Song, J.S., Thanki, N., Wang, Z., Yamashita, R.A., Zhang, D., Zheng, C., and
Bryant, S.H. (2015). CDD: NCBI's conserved domain database. Nucleic Acids Res 43,
D222-226.
McConn, M., and Browse, J. (1996). The critical requirement for linolenic acid is pollen
development, not photosynthesis, in an arabidopsis mutant. Plant Cell 8, 403-416.
Mosblech, A., Feussner, I., and Heilmann, I. (2009). Oxylipins: structurally diverse
metabolites from fatty acid oxidation. Plant Physiol Biochem 47, 511-517.
Mueller, M.J. (2004). Archetype signals in plants: the phytoprostanes. Curr Opin Plant Biol 7,
441-448.
Murata, Y., Mori, I.C., and Munemasa, S. (2015). Diverse stomatal signaling and the signal
integration mechanism. Annu Rev Plant Biol 66, 369-392.
Petsalaki, E.I., Bagos, P.G., Litou, Z.I., and Hamodrakas, S.J. (2006). PredSL: a tool for the
N-terminal sequence-based prediction of protein subcellular localization. Genomics
Proteomics Bioinformatics 4, 48-55.
Porta, H., and Rocha-Sosa, M. (2002). Plant lipoxygenases. Physiological and molecular
features. Plant Physiol 130, 15-21.
Riemann, M., Dhakarey, R., Hazman, M., Miro, B., Kohli, A., and Nick, P. (2015).
Exploring jasmonates in the hormonal network of drought and salinity responses. Front
Plant Sci 6, 1077.
Schwacke, R., Schneider, A., van der Graaff, E., Fischer, K., Catoni, E., Desimone, M.,
Frommer, W.B., Flugge, U.I., and Kunze, R. (2003). ARAMEMNON, a novel
database for Arabidopsis integral membrane proteins. Plant Physiol 131, 16-26.
Song, S., Huang, H., Gao, H., Wang, J., Wu, D., Liu, X., Yang, S., Zhai, Q., Li, C., Qi, T.,
and Xie, D. (2014). Interaction between MYC2 and ETHYLENE INSENSITIVE3
modulates antagonism between jasmonate and ethylene signaling in Arabidopsis. Plant
Cell 26, 263-279.

204

Suzuki, N. (2016). Hormone signaling pathways under stress combinations. Plant Signal Behav
11, e1247139-e1247143.
Thomma, B.P., Eggermont, K., Penninckx, I.A., Mauch-Mani, B., Vogelsang, R., Cammue,
B.P., and Broekaert, W.F. (1998). Separate jasmonate-dependent and salicylatedependent defense-response pathways in Arabidopsis are essential for resistance to
distinct microbial pathogens. Proc Natl Acad Sci U S A 95, 15107-15111.
Troncoso-Ponce, M.A., Cao, X., Yang, Z., and Ohlrogge, J.B. (2013). Lipid turnover during
senescence. Plant Sci 205-206, 13-19.
Trusov, Y., Rookes, J.E., Chakravorty, D., Armour, D., Schenk, P.M., and Botella, J.R.
(2006). Heterotrimeric G proteins facilitate Arabidopsis resistance to necrotrophic
pathogens and are involved in jasmonate signaling. Plant Physiol 140, 210-220.
Turner, J.G., Ellis, C., and Devoto, A. (2002). The jasmonate signal pathway. Plant Cell 14
Suppl, S153-164.
Valenzuela, C.E., Acevedo-Acevedo, O., Miranda, G.S., Vergara-Barros, P., Holuigue, L.,
Figueroa, C.R., and Figueroa, P.M. (2016). Salt stress response triggers activation of
the jasmonate signaling pathway leading to inhibition of cell elongation in Arabidopsis
primary root. J Exp Bot 67, 4209-4220.
Wang, G., Ryu, S., and Wang, X. (2012). Plant phospholipases: an overview. Methods Mol
Biol 861, 123-137.
Wang, K., Froehlich, J.E., Zienkiewicz, A., Hersh, H.L., and Benning, C. (2017). A plastid
phosphatidylglycerol lipase contributes to the export of acyl groups from plastids for seed
oil biosynthesis. Plant Cell 29, 1678-1696.
Wang, Z., and Benning, C. (2011). Arabidopsis thaliana polar glycerolipid profiling by thin
layer chromatography (TLC) coupled with gas-liquid chromatography (GLC). J Vis Exp
49, 2518.
Wang, Z., and Benning, C. (2012). Chloroplast lipid synthesis and lipid trafficking through ERplastid membrane contact sites. Biochem Soc Trans 40, 457-463.
Wasternack, C., and Hause, B. (2013). Jasmonates: biosynthesis, perception, signal
transduction and action in plant stress response, growth and development. An update to
the 2007 review in Annals of Botany. Ann Bot 111, 1021-1058.
Winter, D., Vinegar, B., Nahal, H., Ammar, R., Wilson, G.V., and Provart, N.J. (2007). An
"Electronic Fluorescent Pictograph" browser for exploring and analyzing large-scale
biological data sets. PloS One 2, e718.

205

Xu, C., Fan, J., Froehlich, J.E., Awai, K., and Benning, C. (2005). Mutation of the TGD1
chloroplast envelope protein affects phosphatidate metabolism in Arabidopsis. Plant Cell
17, 3094-3110.

206

CHAPTER 5
Future perspectives

207

Cell membranes in biological systems primarily serve as barriers for cells and the internal
organelles, providing compartments to make cells work at high efficiency. The membrane
composition with regard to the lipids and membrane associated proteins dynamically changes in a
process referred to as membrane remodeling. In sessile plants, membrane remodeling is not only
important for multiple stages of growth, but also a vital strategy to adapt to the changing
environment including freezing, nutrient scarcity, etc. Chloroplast membranes are essential for
chloroplast functions including photosynthesis. Functional and mechanistic understanding of
chloroplast membrane homeostasis under different conditions is crucial for elucidating basic
chloroplast functions and engineering of more resilient and productive crops.

SFR2 in Engineering Stress Tolerant Plants
SFR2 mediated lipid remodeling was originally discovered as a freezing tolerance
mechanism in Arabidopsis thaliana (Warren et al., 1996; Thorlby et al., 2004; Moellering et al.,
2010). Conservation of SFR2 among all land plants, regardless of their freezing sensitivity, led to
the discovery of its function in salt and drought tolerance in Solanum lycopersicum (Wang et al.,
2016). Thus, functional comparison of the homologous SFR2s from different plants provides a
valuable approach for the molecular investigation and engineering of SFR2.
SlSFR2 and AtSFR2 share high sequence identity (59%) and similarity (72%), but the in
vitro studies suggest that SlSFR2 activity is about twice that of AtSFR2 under similarly optimized
conditions (Wang et al., 2016). This implies that the divergent regions of these two sequences
likely affect the SFR2’s catalytic activity. A domain swap experiment can possibly be performed
between these two proteins to systematically understand the functions of the divergent regions.
For example, a similar approach was conducted between the Arabidopsis and Brachypodium

208

TRIGALACTOSYLDIACYLGLYCEROL1 (TGD1) proteins for identification of a specific
divergent loop that is responsible for the oligogalactolipid production in the tgd mutants in
Arabidopsis (Yang et al., 2017). Even though the crystal structure of SFR2 is not available yet,
structure homology modeling of AtSFR2 identifies three regions that distinguish SFR2 from
glycosyl hydrolases (Roston et al., 2014). Comparison between the sequence fragments identified
from these two approaches might further narrow down the effective regions. Alternatively, more
SFR2 homologous sequences can be included to study the correlation between the SFR2 enzyme
activities and their corresponding organisms’ native tolerance to environmental stresses. For
example, Eutrema salsugineum is a member of the Brassicaceae family and phylogenetically
closely related to Arabidopsis (Bressan et al., 2001; Inan et al., 2004). As an extremophile plant
with a wider range of salinity and drought tolerance than Arabidopsis, Eutrema could provide a
facile tool to explore how the unique sequence regions of EsSFR2 might contribute to its superb
tolerance. With more SFR2s from different organisms being characterized in vitro, specific regions
or residues could be uncovered and engineered to achieve higher efficiency and sensitivity. The
activation mechanism of SFR2 in response to abiotic stresses is not fully understood yet thus far.
Possibly, this approach can also help elucidate regions that are involved in SFR2 activation.
Arabidopsis sfr2 mutant have damaged chloroplast membranes but possesses intact plasma
membranes during freezing, which suggests that chloroplasts are more susceptible to freezinginduced membrane damage and that SFR2 serves as the primary line of defense protecting
membranes against these stresses, or that other mechanisms act on plasma membranes not
involving SFR2. SFR2 is located on the chloroplast outer-envelope membrane facing the cytosol.
Presumably, the outer envelope membrane lipids will be remodeled providing protection during
dehydration. To increase stress tolerance, provide additional insights into the activation of the

209

enzyme, and to further explore the protective function of oligogalactolipids, one potential approach
might be to ectopically relocate SFR2 to other subplastid membranes, e.g., the inner envelope and
thylakoid membranes.
Activation of SFR2 is based on a post-translational mechanism by directly sensing Mg2+
and low pH (Roston et al., 2014; Barnes et al., 2016). In vitro, Mg2+ alone is able to activate SFR2
(Roston et al., 2014; Wang et al., 2016). In vivo, the majority of Mg2+ is sequestered in the vacuole
and the cytosol Mg2+ level is determined by a class of Mg2+ / H+ antiporters on the vacuole
membrane (Shaul, 2002). To expand the impact of SFR2 on plant physiology, genes for certain
vacuolar Mg2+ / H+ exchangers, e.g., AtMHX (Shaul et al., 1999) could be engineered to express
under the control of an ABA responsive promoter. Following drought or salt stresses, more Mg2+
will be pumped into the cytosol to activate SFR2 providing the chloroplasts with a more sensitive
protection mechanism.
Taken together, these approaches and ideas discussed above can potentially be combined
in order to generate a crop plant more tolerant of abiotic stresses.

PLIP1 in Seed Oil Biosynthesis
The PLIP1 and FAD4 pathway contributes to seed oil biosynthesis by releasing fatty acids
(FAs) from plastid PG, and exporting FAs to the ER phosphatidylcholine (PC) pool before finally
entering seed oil biosynthesis. The underlying mechanism of how the acyl groups enter TAG via
PC is not clear. Multiple pathways are in principle available to channel the FAs from PC to TAG.
One pathway is to be the head group exchange mechanism to convert PC to diacylglycerol (DAG)
(Lu et al., 2009). DAG is then acylated to TAG by the action of DAG-acyltransferases (DGAT)
with additional acyl groups from the acyl-CoA pool (Lung and Weselake, 2006). Another pathway

210

is direct incorporation of polyunsaturated FAs from PC into TAG through the phospholipid: DAG
acyltransferases (PDAT) (Dahlqvist et al., 2000). To determine which pathway might work along
with the PLIP1-FAD4 mediated export of FAs from plastids, double mutant lines between plip1
and mutants of key enzymes in different pathways, e.g., rod1, dgat1 and pdat1 can be constructed
by crossing and the seed oil contents can be examined for understanding how PLIP1 fits with the
previously elucidated seed oil biosynthesis pathways. DGAT1 and PDAT1 pathways work
redundantly during seed development and oil biosynthesis, and remarkably, the dgd1 pdat1 double
mutant deficient for the main DGAT1 and PDAT is not viable (Zhang et al., 2009). Conceivably,
because the PLP1 pathway only contributes approximately 10% to seed TAG biosynthesis, the
seeds from the progenies of plip1 crossed with the individual pathway mutants should be viable.
Labeling experiments of seeds with different genotypes can also be conducted for further anlaysis
of the contribution of the different mechanisms to TAG biosynthesis in seeds.
Another interesting question that arises from the study of PLIP1 is how the hydrolyzed
acyls groups are exported. As discussed in Chapter 3, FA exporter 1 (FAX1) and fatty acid-binding
proteins (FAPs) are possible components in FA export in the PLIP1 overexpression plants (Ngaki
et al., 2012; Li et al., 2015). Combining mutants deficient in FAX1 or FAPs with the PLIP1-OX
plants should be a straightforward approach for analyzing the relationship between PLIP1 and
other components. Alternatively, the unique growth reduction phenotype of the PLIP1-OX plants
can be taken advantage of in a forward genetics screening experiment. Ectopical overexpression
of PLIP1 leads to overaccumulation of oxylipins and activation of JA signaling, which causes the
stunted growth in the PLIP1-OX plants (Wang et al., 2017). Screening of repressor mutants that
reverse the growth reduction can conceivably identify novel players in fatty acid and oxylipin
transport, as well as the missing components of JA signaling and its cellular transport.

211

PLIP2 and 3 in the Interaction Between ABA and JA
PLIP2 and 3 have been identified as putative paralogs of PLIP1. They exhibit similar
biochemical activities as PLIP1, but likely play different physiological roles in Arabidopsis. PLIP1
contributes to seed oil biosynthesis, while the plip2 and plip3 seeds show no detectable effects on
either the seed oil content or acyl compositions. The seeds of the plip triple mutant are
indistinguishable from those of the plip1 seeds. Furthermore, the expression of PLIP2 and 3, but
not PLIP1 is induced by ABA. Interestingly, based on the public transcriptomic database, PLIP2
and PLIP3 are induced by different abiotic conditions. Therefore, it is possible that they are
involved in mitigating different abiotic stresses in vivo. It might be important to evaluate the
physiological responses of the individual plip mutants under different conditions to gain a more
comprehensive understanding of their physiological functions. The phenomics platform built by
the Kramer lab can be of help in detecting subtle physiological phenotypes of the mutant plants in
response to dynamic conditions in a more sensitive manner (Davis et al., 2016). This can also help
understanding the real functions of the PLIP-mediated interaction between ABA and JA signaling.
Another interesting observation for the plip triple mutants is their hypersensitivity to
applied ABA. The original hypothesis was that JA signaling antagonizes ABA biosynthesis.
Decreased levels of ABA were indeed observed in both wild-type and plip triple mutant seedings
with JA treatment. However, this is unlikely to be the only reason causing the hypersensitive
phenotype, because the ABA reduction caused by JA treatment is only minor and requires a longtime scale. Therefore, it seems likely that inactivation of PLIPs also affects ABA signaling and
that PLIPs are negative regulators for ABA signaling. To further investigate this issue, it would be
interesting to create higher order mutants that are also deficient in key components in ABA
signaling, e.g., the protein kinases and ABA INSENSITIVE (ABI) transcription factors, in the

212

background of the plip triple mutant (Finkelstein, 2013). This would help understand which ABA
signaling step(s) PLIP takes part in.
To sum up, plant membrane lipid remodeling is a highly regulated dynamic process. This
is still comparatively underexplored topic in plant lipid field. Further studies promise to uncover
more novel mechanisms of lipid remodeling and their relationships with other biological processes,
e.g., lipid signaling in abiotic stress tolerance, etc.

213

REFERENCES

214

REFERENCES

Barnes, A.C., Benning, C., and Roston, R. (2016). Chloroplast membrane remodeling during
freezing stress is accompanied by cytoplasmic acidification activating SENSITIVE TO
FREEZING 2. Plant Physiol 171, 2140-2149.
Bressan, R.A., Zhang, C., Zhang, H., Hasegawa, P.M., Bohnert, H.J., and Zhu, J.-k. (2001).
Learning from the Arabidopsis Experience . The Next Gene Search Paradigm 127, 13541360.
Dahlqvist, A., Stahl, U., Lenman, M., Banas, A., Lee, M., Sandager, L., Ronne, H., and
Stymne, S. (2000). Phospholipid:diacylglycerol acyltransferase: an enzyme that catalyzes
the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Natl
Acad Sci U S A 97, 6487-6492.
Davis, G.A., Kanazawa, A., Schottler, M.A., Kohzuma, K., Froehlich, J.E., Rutherford,
A.W., Satoh-Cruz, M., Minhas, D., Tietz, S., Dhingra, A., and Kramer, D.M. (2016).
Limitations to photosynthesis by proton motive force-induced photosystem II
photodamage. Elife 5.
Finkelstein, R. (2013). Abscisic Acid synthesis and response. Arabidopsis Book 11, e0166.
Inan, G., Zhang, Q., Li, P., and Wang, Z. (2004). Salt cress. A halophyte and cryophyte
Arabidopsis relative model system and its applicability to molecular genetic analyses of
growth and development of extremophiles. Plant Physiol 135, 1718-1737.
Li, N., Gugel, I.L., Giavalisco, P., Zeisler, V., Schreiber, L., Soll, J., and Philippar, K.
(2015). FAX1, a novel membrane protein mediating plastid fatty acid export. PLoS Biol
13, e1002053.
Lu, C., Xin, Z., Ren, Z., Miquel, M., and Browse, J. (2009). An enzyme regulating
triacylglycerol composition is encoded by the ROD1 gene of Arabidopsis. Proc Natl
Acad Sci U S A 106, 18837-18842.
Lung, S.C., and Weselake, R.J. (2006). Diacylglycerol acyltransferase: a key mediator of plant
triacylglycerol synthesis. Lipids 41, 1073-1088.
Moellering, E.R., Muthan, B., and Benning, C. (2010). Freezing tolerance in plants requires
lipid remodeling at the outer chloroplast membrane. Science 330, 226-228.
Ngaki, M.N., Louie, G.V., Philippe, R.N., Manning, G., Pojer, F., Bowman, M.E., Li, L.,
Larsen, E., Wurtele, E.S., and Noel, J.P. (2012). Evolution of the chalcone-isomerase
fold from fatty-acid binding to stereospecific catalysis. Nature 485, 530-533.

215

Roston, R.L., Wang, K., Kuhn, L.A., and Benning, C. (2014). Structural determinants
allowing transferase activity in SENSITIVE TO FREEZING 2, classified as a family I
glycosyl hydrolase. J Biol Chem 289, 26089-26106.
Shaul, O. (2002). Magnesium transport and function in plants: the tip of the iceberg. Biometals
15, 309-323.
Shaul, O., Hilgemann, D.W., de-Almeida-Engler, J., Van Montagu, M., Inze, D., and Galili,
G. (1999). Cloning and characterization of a novel Mg2+/H+ exchanger. EMBO J 18,
3973-3980.
Thorlby, G., Fourrier, N., and Warren, G. (2004). The SENSITIVE TO FREEZING2 gene ,
required for freezing tolerance in Arabidopsis thaliana , Encodes a beta-glucosidase 16,
2192-2203.
Wang, K., Hersh, H.L., and Benning, C. (2016). SENSITIVE TO FREEZING2 Aids in
Resilience to Salt and Drought in Freezing-Sensitive Tomato. Plant Physiol 172, 14321442.
Wang, K., Froehlich, J.E., Zienkiewicz, A., Hersh, H.L., and Benning, C. (2017). A plastid
phosphatidylglycerol lipase contributes to the export of acyl groups from plastids for seed
oil biosynthesis. Plant Cell 29, 1678-1696.
Warren, G., McKown, R., Marin, a.L., and Teutonico, R. (1996). Isolation of mutations
affecting the development of freezing tolerance in Arabidopsis thaliana (L.) Heynh. Plant
Physiol 111, 1011-1019.
Yang, Y., Zienkiewicz, A., Lavell, A., and Benning, C. (2017). Coevolution of domain
interactions in the chloroplast tgd1, 2, 3 lipid transfer complex specific to Brassicaceae
and Poaceae plants. Plant Cell 29, 1500-1515.
Zhang, M., Fan, J., Taylor, D.C., and Ohlrogge, J.B. (2009). DGAT1 and PDAT1
acyltransferases have overlapping functions in Arabidopsis triacylglycerol biosynthesis
and are essential for normal pollen and seed development. Plant Cell 21, 3885-3901.

216