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REGULATION OF LIPID METABOLISM IN RESPONSE TO
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REGULATION OF LIPID METABOLISM IN RESPONSE TO ENVIRONMENTAL
STRESS IN PLANTS AND ALGAE
By
Eric R. Moellering
A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Biochemistry and Molecular Biology
2010
ABSTRACT
REGULATION OF LIPID METABOLISM IN RESPONSE TO ENVIRONMENTAL
STRESS IN PLANTS AND ALGAE
By
Eric R. Moellering
Glycerolipids are the major building blocks of biological membranes which
provide semi-permeable boundaries for plant cells and the organelles within. Insight into
the molecular and genetic factors required for lipid biosynthesis is crucial for
understanding membrane processes as well as organelle biogenesis. The chloroplasts of
eukaryotic plant cells are comprised of three membrane systems: the inner and outer
envelope membranes, and thylakoids. The lipid composition is distinct in chloroplast
membranes in that phosphorous—free galactoglycerolipids are the predominant class. The
central genes and enzymes in galactolipid biosynthesis have been identified in
Arabidopsis thaliana through combined molecular and genetic approaches. While a clear
overall picture of galactolipid biosynthesis in plants has emerged, additional regulatory
and metabolic factors remain to be discovered.
A mutant suppressor screen was conducted in the Arabidopsis dgd] mutant
deficient in the major biosynthetic pathway for the production of
digalactosyldiacylglycerol. One mutant, dgdl suppressor 1 (dgsl) was identified and was
found to cause a mutation in a mitochondrial protein of unknown function, DGS 1.
Chapter 2 of this dissertation focuses on a detailed characterization of DGS 1. The dgsI-I
mutation is semi-dominant and suppresses dng through the activation of (an) alternative
biosynthetic pathway(s), likely through increased levels of reactive oxygen species
present in dgsl—I . Importantly, analysis of a null allele, dgsI-Z, clearly revealed that
effects on chloroplast localized galactolipid metabolism were only found in dgsI-I
backgrounds, indicating that the wild-type DGS] protein is not directly involved in
regulating galactolipid metabolism.
However, dgs] -1 and other suppressors of dng were found to activate a poorly
understood and distinct pathway for galactolipid biosynthesis via the enzyme
galactolipidzgalactolipid galactosyltransferase (GGGT). Through a reverse genetics
approach, the gene encoding GGGT was identified and found to be encoded by a
previously described gene Sensitive T o Freezing 2 (SFR2). SFR2 encodes a glycosyl
hydrolase family 1 protein required for freezing tolerance, and the work described in
chapter 3 shows that SFR2 is sufficient for GGGT activity when expressed in
heterologous form. The role for GGGT in fi'eezing tolerance is as a galactolipid
remodeling enzyme that alters ratios of non-bilayer to bilayer forming lipids in the
chloroplast envelope, which is hypothesized to stabilize this membrane during freeze
induced dehydration.
In a third project, the biosynthesis of triacylglycerols in the green alga
Chlamydomonas reinhardtii during nitrogen deprived growth was investigated. TAGS
accumulate during N-deprivation, and lipid droplets were isolated from cells under these
conditions. Identification of the associated proteins (259 in total) revealed the existence
of one predominant protein designated major lipid droplet protein (MLDP). MLDP is a
protein with no known functional domains and its knockdown through RNA interference
revealed a potential role for MLDP in lipid droplet morphology. In general, molecular
factors involved in green alga are less characterized than their plant counterparts, and this
work contributes initial insight into the cell biology of lipid droplets in green algal cells.
ACKNOWLEDGEMENTS
As I write this—the last section of my dissertation—thoughts and memories of all who
have helped me along the way become overpowering. The past five years I have spent as
a graduate student at Michigan State University have been the most challenging, and at
the same time, most rewarding of my life. However, I must step back prior to my days at
Michigan State, and acknowledge Dr. Raymond Chollet, now W.W. Marshall Family
University Professor Emeritus at the University of Nebraska-Lincoln. My work as an
undergraduate researcher in the Chollet lab solidified my passion for scientific inquiry,
and provided an ideal combination of tutelage and challenge that few receive at this stage
in their scientific career.
I joined Dr. Christoph Benning’s lab at Michigan State in May, 2006, and while
the time seems to have flown right by, it has truly been the most enjoyable of my life.
From the beginning, I feel that Christoph showed trust in my abilities, and allowed me a
great deal of freedom in finding my own way—all-the-while providing guidance to keep
me on track. This is a mark of a truly great professor, and Christoph has been a mentor
and friend to me, both professionally and personally, in too many ways to account for
here. The work enviromnent he has created in his lab is a fertile ground for discovery,
which one can only hope to find in the next step of their career, or one day replicate.
With this in mind, I must acknowledge the Benning lab colleagues whom I’ve had the
privilege to work with over the past five years. Numbering more than 30 over my stay at
Michigan State, I’ll keep this general—mainly for fear of omission: you have all made
work a joy to come to every day. The insight, debate, collaboration, jokes, quirks,
iv
frustration, and collective problem solving that we’ve all shared has been a blessing that
is only rarely found at “work”.
More broadly, I must acknowledge the institutions within Michigan State
University that provided integral support during my graduate work. Firstly, the
Department of Biochemistry and Molecular Biology, in which my degree is based, has
offered nothing but premiere education to a graduate student. The Department of Energy
Plant Research Laboratory—across the street from my BMB home base—is a truly
unique feature of Michigan State (and a one of a kind worldwide institution), and I have
been profoundly impacted as a graduate student in the PRL program. The above-affiliated
faculty and staff, and the facilities maintained within the College of Natural Sciences
have provided me with great opportunity and no restrictions in my research at Michigan
State. In addition, the members of my guidance committee, Drs. Beronda Montgomery,
Gregg Howe, Shelagh Ferguson-Miller, and John Ohlrogge have provided much
appreciated support in my research. Funding provided by grants to Dr. Benning from the
Department of Energy and the Air Force Office of Scientific Research were crucial in my
doctoral research.
I also want to thank all the fiiends I have gained while in Michigan. My time as a
graduate student has been greatly enriched by spending our little free time together.
Lastly, but certainly not least, I must express great love for my family—my parents and
two brothers, for their love and support.
TABLE OF CONTENTS
List of Tables ..................................................................................................................... ix
List of Figures ...................................................................................................................... x
CHAPTER 1
Regulation of chloroplast glycerolipid biosynthesis in response to environmental stress...l
Introduction .............................................................................................................. 2
A general overview fatty acid biosynthesis and glycerolipid assembly pathways in
plants ........................................................................................................................ 3
Galactolipid biosynthesis during normal and nutrient limited growth
conditions. . .............................................................................................................. 7
Galactolipid biosynthesis in non-photosynthetic tissues ....................................... 10
Galactolipid remodeling and turnover during various environmental stresses ...... 12
Production of cycIo-oxylipin-galactolipids during plant wounding and pathogen
stresses ................................................................................................................... 13
Other routes for galactolipid biosynthesis in plants ............................................... l6
Biosynthesis of the chloroplast anionic glycerolipids SQDG and PG, during
normal and stressed growth conditions .................................................................. l9
Perspectives ............................................................................................................ 22
References .............................................................................................................. 23
CHAPTER 2
Functional characterization of the DGS] protein in Arabidopsis thaliana ........................ 31
Abstract .................................................................................................................. 32
Introduction ............................................................................................................ 34
Materials and Methods ........................................................................................... 37
Plant materials and growth conditions ....................................................... 37
Bioinformatic analysis of DGS] ................................................................ 37
Generation of the DGS] antibody and irnmunoblot analysis .................... 38
Isolation of mitochondria and chloroplasts ................................................ 39
RNA analysis ............................................................................................. 40
Lipid analysis ............................................................................................. 40
Results .................................................................................................................... 41
Introduction of dgs1-2 does not suppress the dng growth phenotype ..... 4l
DGSl is a mitochondrial membrane protein ............................................. 4]
DGS] is present in a high molecular weight complex ............................... 46
Phosphate deprivation and dgsI-l allelism have additive effects on lipid
metabolism ................................................................................................. 48
Alternative oxidase protein levels are reduced in the dgsI-I mutant allele
.................................................................................................................... 53
The dgsl -1 mutant shows phenotypes of an AOXla-deficient mutant ..... 59
Discussion .............................................................................................................. 61
References .............................................................................................................. 64
vi
CHAPTER 3
Identification of the gene encoding galactolipid2galactolipid galactosyltransferase and its
role in freezing tolerance .................................................................................................. 69
Abstract .................................................................................................................. 70
Introduction ............................................................................................................ 71
Materials and Methods ........................................................................................... 72
Plant growth conditions ............................................................................. 73
Isolation of sfrZ T-DNA mutants and complementation of sfr2-3 with
SFR2 cDNA ............................................................................................... 73
Lipid analysis ............................................................................................. 74
Heterologous expression of SFR2 in yeast ................................................ 75
Protein quantification and irnmunoblotting ............................................... 76
In vitro GGGT assay .................................................................................. 76
Results .................................................................................................................... 77
Development of a TLC-based screen to identify Arabidopsis mutants
defective in GGGT activity ........................................................................ 77
T-DNA mutants in the SFR2 gene are defective in oligogalactolipid
accumulation .............................................................................................. 79
The sfr2 mutants show defects in lipid remodeling during freezing stress
.................................................................................................................... 79
Effect of various salts and osmolytes on oligogalactolipid and TAG
accumulation following vacuum infiltration .............................................. 85
Genetic complementation of sfr2-3 by transgenic over-expression of the
SFR2 cDNA ............................................................................................... 91
Heterologous expression of the SFR2 protein in yeast and analysis of in
vitro GGGT activity ................................................................................... 91
Discussion .............................................................................................................. 95
A hypothesis for the role of GGGT in stabilizing chloroplast envelopes
during freeze-induced cellular dehydration ............................................. 100
Evolution of glycosyl hydrolase family enzymes towards catalysis of
transglycosidation: a proposed mechanism for GGGT activity ............... 103
Is the protective role of GGGT specific to freeze-induced dehydration?
.................................................................................................................. 104
References ............................................................................................................ 108
CHAPTER 4
Literature Review: Molecular genetics of lipid metabolism in the green alga
Chlamydomonas reinhardlii ............................................................................................ l 14
Abstract ................................................................................................................ 115
Abbreviations ....................................................................................................... l 16
Introduction .......................................................................................................... 1 17
Discussion ............................................................................................................ 118
Fatty acid synthesis and incorporation into glycerolipids ........................ l 18
Chloroplast membrane lipids ................................................................... 122
Extrachloroplastic membrane lipid metabolism ...................................... 127
vii
Fatty acid desaturation ............................................................................. 137
Distinctions between ER-to-chloroplast lipid trafficking in plants and
algae ......................................................................................................... l 36
Neutral glycerolipid metabolism .............................................................. 138
Perspectives .......................................................................................................... 141
References ............................................................................................................ 141
CHAPTER 5
Analysis of triacylglycerol accumulation in the green alga Chlamydomonas reinhardtii
during nitrogen deprivation: Identification and characterization of an algal specific lipid
droplet protein MLDP ...................................................................................................... 154
Abstract ................................................................................................................ 155
Introduction .......................................................................................................... l 56
Materials and Methods ......................................................................................... 158
Strains and growth conditions .................................................................. 158
Isolation of lipid droplets from N-limited C. reinhardrii ......................... 158
Mass spectrometric identification of lipid droplet proteins ..................... 159
Fatty acid and lipid analysis ..................................................................... 160
Construction of vectors and transformation of C. reinhardrii ................. 16]
Quantitative Real-time RT-PCR .............................................................. 162
Fluorescent, confocal, and electron microscopy ...................................... 163
Bioinformatics .......................................................................................... 164
Results .................................................................................................................. 165
TAG accumulation following shift to N-limiting medium ...................... 165
Properties of lipid droplets ....................................................................... 168
Proteins identified in the lipid droplet fraction ........................................ 173
MLDP suppression affects lipid droplet size ........................................... 182
MLDP has orthologs only in green algae ................................................ 187
Discussion ............................................................................................................ 190
References ............................................................................................................ 1 95
CHAPTER 6
Conclusions and perspectives .......................................................................................... 199
DGSl is not directly involved in the regulation of galactolipid biosynthesis in
plants. ................................................................................................................... 201
GGGT is encoded by the SFR2 gene in Arabidopsis and galactolipid remodeling
in the chloroplast envelope is essential for freezing tolerance. ........................... 203
Triacylglycerol accumulation in Chlamydomonas reinhardtii: a role for MLDP in
lipid droplet biogenisis? ...................................................................................... 207
References ............................................................................................................ 209
viii
LIST OF TABLES
Table 5.1. Total fatty acid content of whole cells compared to fatty acids bound in
triacylglycerols isolated from cells which were grown in medium containing 10 mM
NH;+ (+N), or cells switched to 0 mM NH4+ and grown for one day (-N) ..................... 167
Table 5.2. Fatty acid content of whole organelles (eyespot and lipid droplet) compared to
fatty acids bound in triacylglycerols isolated from these organelles ............................... 172
Table 5.3. Complete list of proteins identified in the Chlamydomonas lipid droplet
fraction ............................................................................................................................. 175
LIST OF FIGURES
Images in this dissertation are presented in color.
Figure 1.1. Glycerolipid assembly and trafficking in chloroplast membrane lipid
biosynthesis .......................................................................................................................... 4
Figure 1.2 Chloroplast glycerolipid and fatty acid structures .............................................. 5
Figure 1.3 The four major chloroplast localized glycerolipid biosynthetic pathways ......... 8
Figure 1.4. C yclo—oxylipid-galactolipids (cGLs) produced during environmental stresses
in Arabidopsis .................................................................................................................... 14
Figure 1.5. The GGGT galactolipid remodeling reaction .................................................. 17
Figure 2.1. Comparison of the dgs1-2 T-DNA null allele and the dgsI-l point mutant
gain of function allele in wild type and dgdl backgrounds ............................................... 42
Figure 2.2. Sequence analysis of DGS] ............................................................................. 43
Figure 2.3. Localization of DGSl in mitochondria ........................................................... 45
Figure 2.4. Blue Native PAGE analysis of the DGS] complex ......................................... 47
Figure 2.5. Mitochondria of dgdl dgsl -1 homozygous mutant plants do not accumulate
galactoglycerolipids ........................................................................................................... 49
Figure 2.6. Galactoglycerolipid content in the dgdl single homozygous mutant and the
dng dgsI-I and dgd] dgsI-Z double homozygous mutants under normal and phosphate
deprived growth conditions ................................................................................................ 51
Figure 2.7. Galactolipid Fatty acid profiles ...................................................................... 54
Figure 2.8. Alternative oxidase is reduced in dgsl-I mutant mitochondria ...................... 56
Figure 2.9. Analysis of AOXIa transcript levels in seedlings treated with Antimycin A...
............................................................................................................................................ 58
Figure 2.10. The dgsI -1 but not the dgs1-2 mutant displays a root growth phenotype ..... 60
Figure 3.1. Activation of GGGT activity in leaf tissue by vacuum infiltration of MgClz
............................................................................................................................................ 78
Figure 3.2. Genotypic confirmation of the sfr2 T-DNA mutant alleles used in this
study. ................................................................................................................................ 80
Figure 3.3. Accumulation of oligogalactolipids in galactolipid biosynthetic mutants ...... 82
Figure 3.4. Freeze-induced galactolipid remodeling is not observed in sfr2 mutants ....... 83
Figure 3.5. Freeze-induced triacylglycerol accumulation .................................................. 86
Figure 3.6. ESl-MS/MS of triacylglycerols isolated from freeze-treated wild-type plants
............................................................................................................................................ 88
Figure 3.7. Salt and osmolyte treatments that activate TGDG and TAG accumulation in
wild-type leaves ................................................................................................................. 89
Figure 3.8. Complementation of the sfr2-3 T-DNA mutant with SF R2 cDNA ................. 92
Figure 3.9. Heterologous expression of SFR2 in Saccharomyces cerevisae ..................... 94
Figure 3.10. Recombinant SFR2 in vitro activity from transgenic yeast ........................... 96
Figure 3.11. lH-NMR spectra of galactolipids from plant leaves and isolated from the in
vitro GGGT assay .............................................................................................................. 98
Figure 3.12. Model of SF R2 activity and the effect of freezing on the chloroplast outer
envelope membrane (oEv) and adjacent cell membranes (CM) in the sfi'2 mutant and
wildtype (SFR2) ............................................................................................................... 102
Figure 3.13. A proposed reaction mechanism for GGGT ................................................ 105
Figure 3.14. Phylogenetic relationships of SFR2 orthologs ............................................ 107
Figure 4.1. Overview of glycerolipid biosynthesis in Chlamydomonas .......................... 120
Figure 4.2. Structures of PC and DGTS .......................................................................... 128
Figure 4.3. Overview of acyl-chain desaturation in (’hlamydomonas ............................. 132
Figure 5.1. Accumulation of triacylglycerols in lipid droplets during N-limitation ........ 166
Figure 5.2. Ultrastructure of cells during N-limitation .................................................... 169
Figure 5.3. Characteristics of isolated lipid droplets ....................................................... 170
Figure 5.4. Distribution of lipid droplet associated proteins into functional groups ....... 174
Figure 5.5. MLDP mRNA is induced following nitrogen deprivation ............................ 184
xi
Figure 5.6. Generation of MLDP RNAi-mediated knockdown strains ........................... 185
Figure 5.7. MLDP RNAi knockdown lines exhibit a lipid droplet morphological
phenotype ......................................................................................................................... 186
Figure 5.8. Multiple sequence alignment of MLDP with other green algal orthologs and
hydropathicity plots of MLDP and other known lipid droplet binding proteins ............. 188
xii
Chapter 1
Regulation of chloroplast glycerolipid biosynthesis in response to environmental
stress
Introduction
Membranes provide boundaries for plant cells and the subcellular organelles
within. Additionally, membranes are important sites for energy and signal transduction.
Glycerolipids are major building blocks of biological membranes and membrane
biogenesis requires exquisite regulation at the level of biosynthesis and extensive
trafficking of glycerolipids. This results in subcellular membranes with distinct lipid
compositions and homeostatic control of this composition is important for their
specialized functions within the cell. Chloroplast membranes are an excellent example of
this phenomenon, where non-phosphorous glycolipids are the predominant species. The
chloroplast lipid biosynthetic pathways require enzymes that are localized at both the
inner and outer envelope membranes of the chloroplast, and moreover, require trafficking
of precursor lipid substrates from within the plastid, as well as glycerolipid backbones
that are assembled in the endoplasmic reticulum. During normal growth and
development chloroplast lipid composition remains remarkably constant in order to
maintain an optimal environment for the thylakoid-localized photosynthetic machinery.
However, under a number of environmental stress conditions, plants adapt by altering
chloroplast lipid composition. This chapter will outline the current state of understanding
of chloroplast lipid biosynthesis, with a focus on the factors required for modulating lipid
responses in this dynamic organelle in response to environmental stress.
A general overview fatty acid biosynthesis and glycerolipid assembly pathways in
plants.
De novo fatty acid biosynthesis takes place in the plastid in plant cells, resulting
in the production of 1620- and 18:1 (carbons : double bonds) acyl chains linked to acyl
carrier proteins (ACPs) (Ohlrogge and Browse, 1995, Ohlrogge et al., 1979). These fatty
acids can be directly incorporated into glycerolipids in the plastid through the stepwise
activity of acyl ACP dependent glycerol-3-phosphate acyltransferase (GPAT) and
lysophosphatidic acid acyltransferase (LPAAT) enzymes to produce phosphatidic acid
(PA) (Figure 1.1) (Ohlrogge and Browse, 1995). PA is used directly for the biosynthesis
of phosphatidyl glycerol (PG), which is the only major phospholipid present in thylakoids
(Ohlrogge and Browse, 1995, Xu et al., 2002, Xu eta]., 2006). PA can be further
converted to diacylglycerol (DAG) by a PA phosphatase, providing a substrate for the
biosynthesis of the chloroplast glycoglycerolipids mono- and di-galactosyldiacylglycerol
(MGDG and DGDG, respectively) and sulfoquinovosyldiacylglycerol (SQDG) as firrther
discussed below (Figure 1.2).
Alternatively, fatty acids can be exported after their liberation fi'om ACP to the
endoplasmic reticulum (ER) where they are converted to acyl CoA thioesters and
assembled in to glycerolipids by direct incorporation into the phosphatidylcholine (PC)
pool by acyl CoAzlysoPC acyltransferase or other enzymes (Bates er al., 2007), such as a
set of ER-localized GPAT and LPAAT enzymes (Ohlrogge and Browse, 1995). The ER
lipid biosynthetic pathway provides lipid precursors for the biosynthesis of extraplastidic
membrane lipids including phosphatidylethanolamine (PE) and phosphatidylcholine
(PC). Irnportantly, glycerolipid backbones assembled in the ER
Cytoplasm/ER
cap. E
> L-PAtr' > PA_"* DAG '
Acyl-CoA s r PC at ‘ I
l j“‘~+ PC El
l .5 K,» DGDG
Fi=A gimme °E"
i G3P [2+ L-PA PA DAG I '5"
IZl/v> l\ I\ I
III Acyl-ACP PG 6318006 TE] IE]
-J v v
Thyfakoids
_ Plastic!
Figure 1.1. Glycerolipid assembly and trafficking in chloroplast membrane lipid
biosynthesis. Acyl ACPs generated by the fatty acid synthase complex (FAS, step 1) are
directly utilized in the chlorOplast by GPAT and LPAAT enzymes (steps 2, 3) to produce
PA which is used in the biosynthesis of chloroplast PG (step 7, multiple enzymes). Fatty
acids hydrolyzed fiom Acyl ACPs in the plastid are exported to the cytosol and esterified
in acyl CoAs, which serve as substrates for extraplastidic glycerolipid assembly through
ER—localized GPAT and LPAAT (steps 9, 10) or through acyl editing of the PC pool
(Step 13, multiple enzymes likely). Biosynthesis of extraplastidic phospholipids includes
the major lipid, PC (via steps 11, 12), which provides the precursor backbone for
glycerolipid equivalents that are imported from the ER into the chloroplast (step 14) and
enters the chloroplast DAG pool which is also contributed to by the plastid glycerolipid
assembly pathway via the dephosphorylation of PA by phosphatidic acid phosphatase
(step 4). This DAG pool is used for the biosynthesis of the major galactolipids MGDG
and DGDG (steps 5 and 6, respectively) and the sulfolipid SQDG (step 8, multiple
enzymes). The glycolipids and PG are trafficked to the thylakoid (step 15) through
mechanisms which are incompletely understood. Abbreviations: DAG, diacylglycerol;
DGDG, digalactosyldiacylglycerol; FFA, free fatty acids; G3P, glycerol-3-phosphate;
IBM, inner envelope membrane; L-PA, lysophosphatidic acid; MGDG,
monogalactosyldiacylglycerol; OEM, outer envelope membrane; PA, phosphatidic acid;
PC, phosphatidylcholine; PG, phosphatidylglycerol; SQDG, sulfquinovosyldiacylgycerol.
4
Figure 1.2. Chloroplast glycerolipid and fatty acid structures. (A) Structures for the
galactolipids MGDG and DGDG, the phospholipids PC and PG, and the sulfolipid
SQDG; abbreviations are as in Figure 1.]. Each glycerolipid has a distinct fatty acid
composition esterified to the sn-1 and sn-2 position of the glycerolipid backbone; and the
most common fatty acid species for A. thaliana (several others have been observed) is
indicated by the R1 (for sn-I) and R2 (for sn-2) acyl group species indicated as letters A-F
in parentheses. The identities of A-F are shown in (B). (B) Common fatty acid acyl
groups of glycerolipids. The common names and abbreviation for the corresponding fatty
acids are indicated as is the R-group identifier used in (A). In the drawn structures, the R-
group linkage site is circled, and numbers above carbonzcarbon double bonds indicate the
position counting from the carboxyl end.
HO OH
0 \/L O- o
H DGDG W 0 0’K
P, R2 (A,B)
HO 0 g/
0
HO OH MGDG (DE/R1 (A'E)
O O CH2303' SQDG Q
o m am «a
% H0 R2 (A E)
OH
0
Trivial name R-group
and abbreviation identifier Structure
palmitic acid (16:0)
AIS-trans hexadecenoic
acid (16:1A3!)
oleic acid (18:1)
Iinoleic acid (18:2)
a-Iinolenic acid (18:3)
‘nrnUOm>
palmitolenic acid (16:3)
pathway are trafficked to the chloroplast, feeding into the DAG pool used for the
synthesis of the above-mentioned chloroplast glycolipids (Figure 1.1). Evidence for this
bifurcation in glycerolipid assembly came first from substrate radiotracer studies
(Roughan et al., 1980, Roughan and Slack, 1982). The “two pathway” hypothesis was
later confirmed, when mutants in the plastidial GPAT and the ER and plastidial
desaturases were identified in Arabidopsis (Browse and Somerville, 199], Kunst er al.,
1988). Due to differences in substrate specificities of the plastid and ER-localized lipid
assembly enzymes, glycerolipids derived from either pathway can be distinguished by the
length of the acyl chain esterified to the sn-2 position—where plastid derived lipids
contain 16-carbon acyl chains as opposed to the 18-carbon chains found in ER-derived
lipids. The relative contributions of the plastid and ER-localized glycerolipid assembly
pathways in chloroplast glycoglycerolipid biosynthesis varies among species—where in
spinach and Arabidopsis for example, the plastidial pathway is significant; conu'astingly
in many monocots and in pea, the ER pathway is the sole contributor (Mongrand el al.,
1998). Though not a focus of this chapter, factors required for ER-to-chloroplast lipid
trafficking have recently been identified (Benning, 2009).
Galactolipid biosynthesis in leaves during normal and nutrient-limited growth
conditions.
The galactolipids MGDG and DGDG are the predominant glycerolipid species in
chloroplasts (Dormann and Benning, 2002). The bulk of galactolipid biosynthesis is
carried out by two UDP-galactose (UDP-Gal) dependent enzymes, MGDl and DGD]
(see Figure 1.3) (Benning and Ohta, 2005). MGD] is localized to the inner chloroplast
UTP PPi so3
G1P¥> UDP-GIG UDP-SQ
UDP
Pi
PA DAG SQDG
crp
\ 0034/5 / UDP-Gal
PPi/
i \ UDP
COP-DAG i
G3P \ MGDG
K UDP-Gal
CMP/ \
v UDP
PGP y
DGDG
/
V
PG
Figure 1.3. The four major chloroplast localized glycerolipid biosynthetic pathways.
Substrates, enzymes and steps required for the biosynthesis of the four major chloroplast
glycerolipids MGDG, DGDG, SQDG and PG. Enzymes are indicated in round-edged
boxes, and full names for the abbreviations are: CDS, CDP-DAG synthase; DGD],
DGDG synthase; MGDl, MGDG synthase; PAP, PA phosphatase; PGP], PGP synthase;
PGPase, PGP phosphatase; SQDl, UDP-SQ synthase; SQD2, sulfolipid synthase; UGP3,
UDP-Glc pyr0phosphorylase 3. Abbreviations for molecules are as in Figure 1.1, or:
CDP-DAG, citidinediphosphate—diacylglycerol; CTP, citidinetriphosphate; GlP, glucose-
l-phosphate; Pi, phosphate; PPi, pyrophosphate; 803', sulfite; UDP, uridinediphosphate,
UDP-Gal, UDP-galactose; UDP-Glc, UDP—glucose; UDP-SQ, UDP-sulfoquinovose;
UTP, uridinetriphosphate.
envelope (Awai et al., 2006) where it converts DAG into MGDG. This MGDG can
either be directly transported to the thylakoid membranes, or to the outer chloroplast
envelope, where DGDl acts to convert MGDG to DGDG (Froehlich er al., 2001). A
series of studies on Arabidopsis mutants in this pathway have revealed the importance of
galactolipid biosynthesis during normal plant growth and development A leaky allele of
mng was shown to have defects in chloroplast biogenesis (Jarvis e101,, 2000), and a
more recently described null allele was found to be embryo or seedling—lethal (Kobayashi
et al., 2007). A null allele for dng identified through forward genetics showed a severe
dwarf growth phenotype, but ~10 % of wild-type DGDG levels remained (Diirmann er
al., 1995, Dr'irmann er al., 1999). This latter observation, along with DGDG
accumulation in the dgdl mutant during phosphate deprivation (Hartel er al. , 2000),
indicated the presence of additional pathways for DGDG biosynthesis.
Indeed, a reverse genetics approach identified genes encoding a paralog of DGDI,
DGD2 (Kelly and Dormann, 2002). After initial identification of the MGDG synthetase
purified from cucumber (Shimojima er al., 1997), its ortholog, MGDI, and two paralogs
of MGDI, MGD2,3 were also identified in the Arabidopsis genome (Awai er al., 2001).
MGD2,3 and DGD2 were shown to be coordinately regulated at the transcriptional level,
and highly expressed under phosphate-limiting growth conditions. The MGD2,3 and
DGD2 proteins are localized to the chloroplast outer envelope, and comprise an
alternative pathway for galactolipid biosynthesis that produces DGDG that is trafficked
to extraplastidic membranes (e. g. the tonoplast, mitochondria, and plasma membrane)
during phosphate deprivation stress in a mechanism that conserves phosphate through the
replacement of phospholipids (Hartel et al., 2000, Jouhet et al., 2004, Jouhet er al., 2007,
Tjellstrom et al., 2010). In a detailed study of plasma membranes in phosphate limited
oat (Avena satrva), replacement of phospholipids with DGDG was found to be restricted
to the cytosolic leaflet (Tjellstrom et al., 2010).
More recently, nitrogen (N) or magnesium deprivation was shown to alter
galactolipid composition in Arabidopsis leaves, where MGDG was found to decrease,
with lesser increases in both DGDG and phospholipids (Gaude et al., 2007). Analysis of
transcript levels for the above-described galactolipid biosynthetic genes revealed
increased expression of both DGDI and DGD2, but little or no change in the MGD1/2/3
genes. Fatty acid phytyl esters and triacylglycerols were found to accumulate in response
to N-deprivation, and more detailed analysis revealed an increase in the amount of 16:3
(carbons : double bonds) fatty acid esterified to phytol. As 16:3 is predominately
esterified to MGDG in Arabidopsis during normal growth (Browse 2:01., 1989), these
results suggested that during N-deprivation, MGDG is being turned over and the fatty
acids are converted in part to fatty acid phytyl esters (FAPEs) (Gaude e101,, 2007).
However, the identities of potential lipase and/or transferase enzymes involved in the
formation of FAPEs and their role during N-limitation in plants remain unknown.
Galactolipid biosynthesis in non-photosynthetic tissues.
Not surprisingly, the role of the phosphate-regulated MGD2/3-DGD2 dependent
DGDG biosynthesis pathway is not restricted to photosynthetic tissues. As in leaves,
MGD2/3 and DGD2 are activated in root tissue in Arabidopsis during phosphate
deprivation (Andersson er al., 2003, Benning and Ohta, 2005, Kelly and Dormann, 2002,
Kobayashi et al., 2004, Kobayashi et al., 2006, Lai et al., 2007). Additionally, DGDG
10
has been found to accumulate in phosphate-deprived roots from a number of plant species
in addition to Arabidopsis, including Phaseolus vulgaris (bean) (Hartel er a1. , 2000,
Russo er al., 2007). DGDG also accumulates in peribacteroid membranes (PBMs) in the
legume species soybean and Lotus; the FEM is of plant origin, but surrounds the so-
called “symbiosome” organelle containing nitrogen fixing bacterial species within the
root nodule cells (Gaude er al., 2004). The DGDl/2 orthologs in soybean were shown to
exhibit nodule-enhanced expression (compared to roots), and in situ hybridization
indicated the DGDl/Z mRN As to be enriched in the infected cells of the nodule (Gaude
er a1. , 2004).
A role for galactolipid biosynthesis during pollen tube growth has also emerged
(Kobayashi er al., 2004, Nakamura et al., 2009a). Transgenic Arabidopsis lines
containing promoterzGUS fusion constructs for MGD1,2,3 indicated the expression of all
three genes in anthers, whereas only MGD2 and MGD3 were expressed during pollen
tube growth (Kobayashi etal., 2004). Analyses of lipid content in Lilium Iongzflorum
revealed accumulation of MGDG and DGDG afier pollen tube elongation (Nakamura et
al., 2009a). Given the rapid biosynthesis of membranes required for pollen tube
elongation, a role for the alternative galactolipid biosynthetic pathway in providing
extraplastidic DGDG for this process was hypothesized. However, whether or not
mutants in the alternative galactolipid biosynthetic pathway are defective in pollen tube
growth is currently not known, but major defects are unlikely as null mutants in these
genes have been studied.
11
Galactolipid remodeling and turnover during various environmental stresses.
Although most insight has been gained on the synthesis of galactolipids in plants,
both chilling (Kaniuga and Gemel, 1984) and drought (N avari-lzzo and Rascio, I999,
Torres-Franklin er al., 2007) stress conditions and senescence (Kaup et al., 2002) are
associated with catabolism and/or remodeling of these predominant chloroplast lipids. In
response to low temperature, chilling resistant plants generally show increased fatty acid
desaturation levels (Iba, 2002), but these stress-induced changes are not specific to
galactolipids or chloroplast glycerolipids in general, and as such are not further discussed
here. In response to severe water deficit, the MGDGzDGDG ratio decreases markedly in
Arabidopsis leaves, and transcript levels of MGDI were found to decrease during severe
drought, while DGDI mRNA levels did not (Gigon et al., 2004). Decreases in the
MGDGzDGDG ratio during water deficit and desiccation have been observed in a
number of different plant species (Navari-lzzo and Rascio, 1999), suggesting a conserved
but incompletely understood mechanism for chloroplast lipid remodeling during this
stress. A role for galactolipid remodeling during acquired thermotolerance was revealed
in Arabidopsis, when isolated thennosensitive mutants were mapped and identified as
weak alleles of DGDI (dng-Z and dgd1-3) (Chen et al., 2006). The dng-Z and -3
alleles showed only ~40-50 % decreases in steady state DGDG levels and no apparent
growth defects, and upon exposure to 38° C, did not show the increased DGDG content
and thus altered MGDGzDGDG ratio observed in wild-type plants during heat stress
(Chen et al., 2006). Similar changes have also been observed during exposure of plants
to excess copper or cadmium (Chaffai et al., 2007, Jemal et al., 2000). Whether or not
additional enzymes are involved in stress-induced galactolipid changes remains unclear.
12
In some cases, the changes in lipid composition may be due to changes in transcriptional
or metabolic regulation of the known galactolipid biosynthetic pathways, or due to
increased oxidation-dependent turnover rates that are an indirect consequence of some
stresses (e.g. excess copper).
Recently, a candidate a-galactosidase for the conversion of DGDl/Z derived
DGDG to MGDG has been identified in 0ryza sativa (rice) (Lee et al., 2009, Lee el al.,
2004). The encoding gene, OsAkaGal, produced a functional protein when expressed in
E. coli, and a null allele in rice showed a delayed senescence phenotype (Lee et al.,
2009). However, lipid composition was not reported for wild-type plants or the osakagal
mutant and the in viva role for this enzyme in galactolipid metabolism remains to be
established.
Production of cyclo—oxylipin-galactolipids during plant wounding and pathogen
stresses.
MGDG and DGDG are rich in tri-enoic fatty acids (16:3 and 18:3 in Arabidopsis),
and during stresses such as mechanical wounding (Buseman el al., 2006) or in response
to induced accumulation of the Pseudomonas syn’ngae avirulence protein, Achme , in
planta (Andersson et al., 2006), these galactolipid esterified fatty acids are converted to
oxylipins, resulting in the increased formation of what are collectively referred to as
cyclo-oxylipin-galactolipids (cGLs, also referred to as Arabidopsides A-G, see Figure
1.4) (Bottcher and Pollmann, 2009). cGL formation currently appears to be restricted to
Arabidopsis and other related Brassicaceae (Bottcher and Pollmann, 2009). The esterified
oxylipin species produced are lZ-oxo-phytodienoic acid (OPDA) or dinor- (dn)-OPDA
l3
A B HO R3
anPDA OPDA 0 R2
H 0
HO
0” MGDG
R1
HO
| | O
H
HO 0
0.. $ ; R2
0
° DGDG
c R‘
. MGDG or
cGL s I R R R
pec es DGDG 1 2 3
MGDG—O M OPDA 16:3/18:3 -OH
ArabidopsideA M OPDA OPDA -0H
Arabidopside B M OPDA anPDA ‘0“
Arabidopside c o OPDA anPDA '3:
Arabidopside o o OPDA OPDA N/A
Arabidopside E M OPDA anPDA OPDA
Arabidopside F M 18:3 OPDA .OH
Arabidopside G M OPDA OPDA OPDA
Figure 1.4. Cyclo—oxylipin-galactolipids (cGLs) produced during environmental
stresses in Arabidopsis. (A) structures of (dinor)oxo-phydodienoic acids). (B)
Galactolipids MGDG and DGDG with possible sites of (dn)OPDA esterification
indicated by R], R2, and R3. (C) A legend to the diverse cGL species identified in
Arabidopsis, including MGDG-0 and Arabidopsides A-G. The derived galactolipid is
indicated (M or D for MGDG and DGDG, respectively) and the positions of fatty acids or
their oxidized (dn)OPDA derivatives are indicated for depictied in R1, R2, and R3 (B).
14
from 18:3 and 16:3 fatty acids respectively by the same set of enzymes: A lipoxygenase
(LOX) catalyzes the formation of hydroperoxy- fatty acids that are further converted to
anPDA and OPDA by the stepwise activities of allene oxide synthase (A08) and allene
oxide cyclase (AOC) (Howe and Schilmiller, 2002, Schaller and Stintzi, 2009).
OPDA and anPDA serve as precursors for the biosynthesis of jasmonates, a
class of plant hormones with diverse fimctions in the regulation of abiotic and pathogen
stress responses as well as growth and development (Browse, 2009, Howe and
Schilmiller, 2002, Schaller and Stintzi, 2009). Intriguingly, the levels of OPDAs
esterified in cGLs is far higher than the corresponding free acids (e.g. ~100 fold in
Aerme expressing plants (Andersson et al., 2006)), leading to the hypothesis that
cGLs serve as a stored form of OPDAs that adds an additional level of regulating
substrate levels feeding into the jasmonate biosynthetic pathway (Buseman et al., 2006).
While evidence for direct biosynthesis of cGLs on galactolipid bound fatty acids is
evident in the species formed, the identification of two new cGLs, Arabidopsides E and G
(Figure l.4C), clearly indicate that further modification via transacylation is involved in
their biosynthesis. Arabidopsides E and G are tri-oxylipin containing MGDG derivatives
that, in addition to the OPDAs esterified to the sn-1 and -2 positions of the glycerol
backbone found in other cGLs, contain a third OPDA esterified to the 6’ carbon of the
galactosyl moiety. Arabidopside E (containing anPDA esterified to the sn-2 position of
the glycerol backbone) was shown to inhibit grth of P. syringae, suggesting that some
cGLs may play a direct role in pathogen defense (Andersson et al., 2006). While the
acyltransferase involved in the biosynthesis of Arabidopsides E and G remains unknown,
15
it was hypothesized to be the same or similar to the unknown enzyme responsible for the
synthesis of acyl-MGDG first identified over 50 years ago (Heinz, l967a, Heinz, l967b).
Other routes for galactolipid biosynthesis in plants.
A third and distinct pathway for galactolipid biosynthesis occurs through the
activity of a galactolipidzgalactolipid galactosyltransferase (GGGT) enzyme (Figure 1.5)
(Benning and Ohta, 2005). GGGT was first described at the biochemical level over 40
years ago (van Besouw and Winterrnans, 1978), but its encoding gene(s) and role in
plants has remained enigmatic (Benning and Ohta, 2005, Kelly er al., 2003). Unlike the
above-described MGDl/2/3 and DGDl/2 galactolipid synthases, GGGT is UDP-Gal
independent (Heemskerk er al., 1983, Heemskerk et al., 1990, van Besouw and
Wintermans, 1978), and instead uses two MGDG molecules as substrate to catalyze the
formation of DGDG and DAG via transgalactosylation (Figure l.5A). The DGDG
produced by GGGT is in the all-B conformation with respect to the glycosidic linkages
(BBDGDG), which is distinct from the BaDGDG produced by DGDl/2 (Figure l.5B~C).
Furthermore, through processive activity, GGGT catalyzes the formation of the
oligogalactolipids tri- and tetra-galactosyldiacylglycerol (TGDG and TeDG, respectively)
(van Besouw and Winterrnans, 1978). Due to the high activity of GGGT and lack of a
UDP—Gal dependent effect on DGDG formation from in vitro biochemical studies in
different species, GGGT was hypothesized to be the major contributor to DGDG
biosynthesis in vivo (Heemskerk er al., 1983, Heemskerk et al., 1990, van Besouw and
Wintermans, 1978).
16
0R2
Figure 1.5. The GGGT galactolipid remodeling reaction. (A) Three catalytic cycles of
processive GGGT activity are shown, with the circled number indicating the order of
reactions. The galactolipid products, BBDGDG, BBBTGDG, and BBflBTeDG are produced
along with three DAG molecules. Other GGGT catalyzed transferases reactions are
possible (e.g. transfer of a galactosyl moiety between two BBDGDG molecules). The
difference between the structures of BnDGDG produced by DGDl/2 biosynthetic
pathways (B) and BBDGDG produced by GGGT (C) are shown. The B or (1 letters for the
galactolipid species indicate the anomeric configuration of the l-6 glycosidic linkages
present in each molecule, beginning with the one most distal to the glycerol backbone
moiety. For simplicity, the hydroxyl groups of the galactosyl head group moieties are
drawn as sticks, and the galactosyl protons attached to the l’ C anomeric centers are
indicated as H1’ and H3”.
As described above, molecular genetics revealed a major role played by DGDl
(Dormann e101,, 1999), and further analysis of the dgd] dgd2 double mutant revealed it
to be devoid of detectable levels of DGDG (Kelly etal., 2003), whereas GGGT activity
in isolated chloroplasts of dgdl dgd2 remained indistinguishable from wild-type. Indeed,
GGGT was found to be activated due to osmotic shock arising from certain steps of intact
chloroplast or envelope membrane isolation (Dome er al., 1982, Dome etal., 1985),
suggesting a possible role for GGGT in response to water deficit. GGGT was also found
to be activated in a series of studies on the effect of ozone (03) treatment on chloroplast
lipid metabolism (Sakaki er al., l990a, Sakaki er al., 199%, Sakaki et al., l990c). O3
fumigated spinach leaves accumulated oligogalactolipids as well as triacyl glycerol
(TAG) (Sakaki et al., 199%, Sakaki er al., l990c), and molecular analysis revealed that
DAG derived from GGGT activity was being utilized as a substrate for TAG
biosynthesis. The significance of these galactolipid remodeling events during ozone
stress remained unclear. In vitro studies of GGGT enzyme activity indicated its
activation by divalent cations (e.g. Mg”, Ca”, and Ba2+) as well as free fatty acids added
to the assay (Heemskerk er al., 1983, Sakaki er al., l990a). Most recently GGGT has
been found to be constitutively activated in vivo in the tgd ER-to—chloroplast lipid
trafficking mutants (Awai et al., 2006, Benning, 2009, Lu et al., 2007, Xu et al., 2003,
Xu et al., 2008). Originally isolated as genetic suppressors of dgdl (Xu er al., 2003), the
rgd1/2/3/4 mutants accumulate oligogalactolipids in shoot tissues, although the
molecular/biochemical etiology behind the activation of GGGT in these mutants is
currently unknown. During research conducted in the preparation of this dissertation, the
gene encoding GGGT was identified in Arabidopsis and is further described in Chapter 3.
18
TGDG and/or TeDG are constitutively produced in certain tissues of certain plant species
including rice bran, Adzuki bean, potato tubers, and pumpkin fruit (reviewed in (Holzl
and D'o'rmann, 2007, Kelly and Dormann, 2004). However, the analysis of the anomeric
configuration of the glycosidic linkagese for these constitutively produced
oligogalactolipids (Fujino and Miyazawa, 1979, Galliard, 1969, Kojima et al., 1990)
indicates biosynthetic pathways that are likely to be independent of the all B-
configuration oligogalactolipids produced by GGGT in Arabidopsis (Xu et a]. , 2003)—
possibly involving multiple enzymes. To date, no enzyme known to produce
oligogalactolipids in plants has been characterized at the molecular/genetic level.
Biosynthesis of the chloroplast anionic glycerolipids SQDG and PG during normal
and stressed growth conditions.
Although the head group moieties of the sulfolipid, SQDG, and phospholipid, PG,
may appear to share little structural similarity, these lipids have similar roles in biological
membranes due to the negative charge that they carry (see Figure 1.2). These anionic
lipids are important for maintaining a negative charge density that is important for the
proper function of the photosynthetic machinery (Wada and Murata, 2007).
SQDG biosynthesis begins with the production of UDP-SQ by SQDl (in
Arabidopsis) using UDP-Glc and sulfite as substrates (Figure 1.3). SQDG is then formed
by SQD2, which transfers the sulfoquinovose moiety to DAG. This pathway was first
elucidated in studies on Rhodobacter sphaeroides (Benning and Somerville, 1992a,
Benning and Somerville, 1992b), and through analysis of Rhodobacrer mutants, SQDG
biosynthesis was shown to be of conditional importance during phosphate deprivation
l9
(Benning er al., 1993). An Arabidopsis sqd2 mutant lacking SQDG showed a similar
conditional importance of sulfolipid in higher plants, where no growth defects were
observed until squ was presented with phosphate-deprived conditions (Yu et al., 2002).
The SQD1/2 genes show up-regulation during phosphate deprivation, and under these
conditions, the SQDG levels increase with a concomitant decrease in PG, suggesting a
homeostatic mechanism to maintain a negative charge balance under varying
environmental conditions (Yu et al., 2002). The conditional importance of SQDG may
not be universal among organisms which produce this lipid: a null mutant in
Synechocystis PCC 6803 squ gene (an ortholog of Arabidopsis SQDI) is a sulfolipid
auxotroph (Aoki et a1. , 2004).
Co-expression of Arabidopsis SQDI and SQD2 genes heterologously in E. coli
indicated that the products of these two genes are sufficient for the production of SQDG
in plants (Y 11 et al., 2002). Recently, a third gene required for SQDG biosynthesis in
planta has been identified Utilizing transcriptome co-expression data in Arabidopsis,
Okazaki and colleagues identified a candidate UDP-Glc pyr0phosphorylase encoding
gene (UGP3) that showed strong correlation of expression with known sulfolipid
biosynthetic genes (Okazaki er al., 2009). Null mutants in UGP3 lacked SQDG, and a
UGP-GFP fusion protein was targeted to the chloroplast (Okazaki er al., 2009), providing
convincing evidence that UGP3 is a stroma-localized enzyme required for the synthesis
of the UDP-Glc substrate for SQDI.
PG is the only major phospholipid synthesized in the chloroplast, and as alluded
to above, its biosynthetic pathway is distinct from that of the chloroplast glycolipids. As
seen in Figure 1.3, PG biosynthesis begins with the production of CDP-DAG by the CD8
20
enzyme. Recent molecular/genetic studies in Arabidopsis have identified 5 candidate
CDS encoding genes, and revealed that the CDS4/5 isogenes encode plastidial isozymes
with overlapping functions in PG biosynthesis (Haselier er al., 2010). While no
phenotypes were observed for cds4 or cds5 single mutants, isolation of the corresponding
cds4 cds5 double mutant revealed severe growth phenotypes including non-viability on
soil, and pale yellow leaves when grown on agar medium supplemented with sucrose,
while showing a ~60 % decrease in total leaf PG when compared to wild-type (Haselier
er al., 2010). CDP-DAG produced in the chloroplast is further converted to
phosphatidylglycerolphosphate by PGP] (Figure 1.3) (Wada and Murata, 2007). The
py] mutants in Arabidopsis show similar PG deficiencies and phenotypes as described
for the cds4 cds5 double mutant (Babiychuk etal., 2003, Hagio er al., 2002, Xu et al.,
2002). The last step in chloroplast PG biosynthesis is the dephosphorylation of PGP,
which is catalyzed by a PGP phosphatase that remains unidentified in plants at the
molecular/genetic level. The gene encoding PGP phosphatase have remained unknown in
eukaryotes until only recently, when a mitochondrial protein required for cardiolipin
biosynthesis in Saccharomyces cerevisae, Gep4, was shown to function as a PGP
phosphatase (Osman et al., 2010). Intriguingly, a distant chloroplast targeted (predicted)
ortholog of Gep4 is encoded in the Arabidopsis genome (by the gene At3G58830)—but
its possible role in PG biosynthesis is not known. Modulation of PG levels during
phosphate-limiting stress occurs (Yu et al., 2002), but it is clear from the PG deficient
mutants that PG cannot be functionally replaced by SQDG in plants, where as the
opposite appears to be the case during normal growth and development.
21
Perspectives
Plants have evolved myriad responses to cope with changes in their environment,
and the modulation of lipid metabolism during stress has emerged as a prominent factor
for a number of different responses. While this chapter focused on changes in chloroplast
lipid biosynthesis, responses of extraplastidic lipid metabolism during stress at least equal
that within the chloroplast (Jouhet et al., 2007, Li et al., 2008, Welti etal., 2002).
Indeed, these processes are not mutually exclusive as there is dynamic exchange of lipids
between the chloroplast and extraplastidic membranes through extensive lipid remodeling
and trafficking pathways during normal and stressed growth conditions (Benning, 2009,
Cnrz-Ramirez et al., 2006, Gaude etal., 2008, Li et al., 2006, Nakamura e101,, 2009b).
Since the release of the Arabidopsis genome nearly] 0 years ago (The Arabidopsis
Genome Initiative, 2001), great progress has been made in understanding lipid
biosynthetic pathways in plants (Wallis and Browse, 2010). However, several questions
remain. Among these are: what are the regulatory and metabolic pathways required for
lipid responses to environmental stress? Undoubtedly, the increasingly powerful “omics”
approaches are going to play an expanding role in elucidating the many unknown facets
in plant lipid metabolism (Okazaki et al., 2009). Additionally, the expansion of molecular
tools in other model plants, e. g. Brachypodium (International Brachypodium Initiative,
2010, Pacurar et al., 2008), will aid in the discovery of both conservation and diversity in
lipid metabolism among different plant species.
22
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30
Chapter 2
Functional characterization of DGSI in Arabidopsis thaliana1
I This work has been published in: (i) Xu, C., Moellering, E.R., Fan, J ., and C. Benning.
(2008) Mutation of a mitochondrial outer membrane protein affects chloroplast lipid
biosynthesis. Plant J. 542163-175, and (ii) Moellering, ER. and C. Benning. (2010)
Phosphate regulation of lipid biosynthesis in Arabidopsis is independent of the
mitochondrial outer membrane DGSl complex. Plant Physiol 152: 1951-1959.
Experiments for the data shown in Figure 2.3, panels A-B, were conducted by
Changcheng Xu, as was the production of the antigenic polypeptide used in generating
the DGSl antibody.
31
Abstract
Galactolipid biosynthesis in plants is carried out by UDP-galactose utilizing enzymes that
are localized to the chloroplast envelopes. In Arabidopsis, the bulk of
digalactosyldiacylglycerol (DGDG) is produced by DGDI during normal grth
conditions. A paralog of DGDl , DGD2, is induced during phosphate deprivation in a
mechanism that replaces phospholipids with DGDG in extraplastidic membranes. In
order to identify the largely unknown factors that regulate the DGDI-independent
pathway, and possibly lipid homeostasis in general, a screen for suppressors of the
galactolipid deficient dgd] mutant was conducted. One mutant, dgsI-l (dgdl suppressor
1-1), was identified with partially restored DGDG content. Initial analyses revealed the
DGSI protein to be present in the mitochondria, and that the dgsI-l suppressor
phenotype was semi-dominant. Moreover, dgsI-I plants were found to accumulate H202,
which when applied to dgd] plants, was sufficient to activate dng-independent DGDG
biosynthesis. Here, continued analysis of the gain-of-function dgsl -1 mutant was
accompanied by analysis of a null T-DNA allele, dgsl-Z. The DGSI protein was found
to be part of a large protein complex present in the outer mitochondrial membrane. The
dgsl-I allele causes the loss of the mitochondrial alternative oxidase (AOX) protein
which could possibly explain the accumulation of H202 in this background. The loss-of-
function dgsl-2 allele had no observable effect on plant growth, AOX, or lipid
composition. Moreover, accumulation of DGDG in phosphate-deprived dgdl dgsI -1
double mutants indicates that the dgsI -1-dependent DGDG accumulation is additive to
the phosphate responsive pathway. These results indicate that DGSI is not directly
32
involved in the regulation of lipid metabolism in chloroplasts and mitochondria, and
leaves the molecular function of this protein unresolved.
33
Introduction
In plant cells, the membranes that provide boundaries for the various subcellular
organelles are quite distinct in their glycerolipid composition (Jouhet etal., 2007). This
reflects not only the different lipid biosynthetic pathways which contribute to organelle
biogenesis, but also the unique functions that are carried out by these subcellular
compartments. This is clearly seen in the chloroplast thylakoids that provide a matrix for
the photosynthetic machinery, where the non-phosphorous galactolipids mono- and di-
galactosyldiacylglycerol (MGDG and DGDG, respectively) are the predominant
glycerolipid species (Dormann and Benning, 2002). Under normal growth conditions in
Arabidopsis, MGDG and DGDG are restricted to the chloroplast membranes, and are
primarily synthesized by two UDP-galactose (UDP-Gal) dependent
galactosyltransferases, MGDI and DGD1(Benning and Ohta, 2005). MGDI is present
on the inner chloroplast envelope and converts diacyl glycerol (DAG) derived from both
the plastidic and ER glycerolipid biosynthetic pathways to MGDG (Benning et al., 2006,
Kobayashi et al., 2007, Shirnojima et al., 1997). MGDG is then trafficked to the
thylakoid, or the outer chloroplast envelope, where it can be further galactosylated by
DGDI (Dormann et al., 1999).
The combined activities of the galactolipid biosynthetic and lipid trafficking
pathways give rise, in part, to sub-organellar membranes with defined lipid compositions
during normal growth. However, during environmental stresses, such as phosphate
deprivation, plants adjust their lipid composition in order to survive. The Arabidopsis
null-mutant, dgdl, shows a 90% decrease in DGDG levels and severely stunted growth
(Dormann et al., 1995, Dormann eta]., 1999). However, dgd] plants grown under
34
phosphate-limiting conditions accumulate DGDG to ~60% of the levels found in wild
type plants (Hartel et al., 2000)—indicating that a DGDl-independent DGDG
biosynthetic route exists in plants. In this regard, a set of functional MGD] and DGD]
paralogs, MGD2/3 and DGD2 respectively, have been characterized at the molecular
level and were shown to be coordinately regulated at the transcript level, with strong
induction under Pi-limiting growth conditions (Awai et al., 2001 , Kelly et al., 2003, Kelly
and Dormann, 2002). MGD2/3 and DGD2 have been localized to the chloroplast outer
envelope membrane, and have been proposed to contribute to the biosynthesis of DGDG
destined for extraplastidic membranes (Andersson et al., 2005, Kelly et al., 2003).
Another class of galactolipid biosynthetic enzyme has been found in plants—the
galactolipid:galactolipid galactosyltransferases (GGGTs), which through processive
activity can give rise to oligogalactolipids. While the gene(s) encoding GGGT(s) have
not been identified, GGGT activity was observed in isolated chloroplasts of the
Arabidopsis dgdl dgd2 double null-mutant (Kelly et al., 2003). Regardless,
oligogalactolipids are undetectable in whole-leaf lipid extracts from wild-type
Arabidopsis (Xu et al., 2003), and DGDG was not detected in dng dgd2 double null-
mutant plants (Kelly et a1. , 2003), suggesting that GGGT(s) contribute minimally to
overall galactolipid biosynthesis under normal growth conditions.
While much progress has been made towards the identification and
characterization of genes involved in galactolipid biosynthesis, the regulatory factors that
govern these pathways in plants remain unknown. Toward this end, a suppressor screen
in a dgd] EMS mutagenized population recovered a mutant, dgsl-l (dgdl suppressor ]-
1). The dgsl-I mutant shows constitutive activation of alternative DGDG biosynthesis,
35
with DGDG accumulating to ~60 % of the level found in wild-type leaves. The dgsl -1
mutation has been mapped to a nuclear gene encoding a ~69 kDa protein of unknown
frmction with two predicted trans-membrane spanning domains (hereafier referred to as
DGSI), where it causes a G-to-A mutation resulting in a substitution of Asp457 to Asn
(D457N) in the deduced protein sequence. Initial studies by Changcheng Xu showed that
the over-expressed DGSI-C-terminal-GF P fusion protein was associated with the E"
mitochondria. Interestingly, over expression of the dgsl-I-GFP cDNA resulted in a
severe dominant negative growth defect accompanied by early leaf senescence, while
over-expression of the DGSI-GFP cDNA did not (Xu et al., 2008). These results
suggested that DGSI may be present in a protein complex, as dominant negative
phenotypes are often associated with the poisoning of a protein complex with a mutant
protein (Herskowitz, 1987, Leavitt et al., 1999). When hydrogen peroxide (H202, a
reactive oxygen species [ROS]) levels in leaves were determined, the dgsI -1 -GFP
transgenic lines showed a two-to-three fold elevation over wild-type levels. Likewise, in
homozygous dgsl-I plants, H202 levels were moderately elevated in leaves and roots.
Subsequent experiments revealed that treatment of dgdl plants with H202 resulted in an
increase in the levels of MGD2/3 transcripts as well as the accumulation of DGDG.
These results provided a potential link between the elevated H202 in dgsl-I and the
phenotypic suppression of dgdl that this allele causes. To further investigate the possible
role of DGSI in regulating galactolipid biosynthesis, a comparative study of the dgsl-I
allele with a newly identified null T-DNA allele, dgsI-Z was initiated.
36
Materials and Methods
Plant materials and growth conditions. Wild-type Arabidopsis plants (Col-2), and the
dgsI-I and dgdl dgsl -1 mutants are as previously described (Xu et al., 2003, Xu et al.,
2008). The dgsl-Z T-DNA insertion line (Sessions et a]. , 2002), SAIL_391_F040, was
obtained from the Arabidopsis Biological Resource Center at Ohio State University. This
line was selfed and tested for homozygosity (lack of segregation of the T-DNA marker).
Homozygous dgdl dgsI-Z double mutant plants were obtained from the F2 seeds of a
cross between dgdl and dgsl-Z homozygous mutant plants. Genotyping for dgdl
homozygosity was performed by PCR with a dCAPS marker (Xu et al., 2003).
Homozygosity for dgsI-Z was confirmed by the absence of a PCR amplicon when using
the forward and reverse primers 5’-GAGGTGAGTGAACTAGAACATTCCATGAG-3’
and 5'-ACTCAACATAAAGCTAGAAAGCCCATTG-3’, respectively, which span the
T-DNA insertion, and the presence of a PCR amplicon when the above reverse primer is
used with a primer complementary to the left border of the T-DNA insertion (Alonso et
al., 2003). In general, surface sterilized seeds were grown on agar solidified MS medium
or soil as previously described (Xu et al., 2003).
Bioinformatic analysis of DGSI. Full-length, deduced amino acid sequences were
aligned with ClustalX (version 1.81), and generation of the bootstrapped phylogentic tree
was performed using the PHYLIP software package as previously described (Mamedov
et al., 2005). For transmembrane domain prediction the TMHMM program was
employed (Krogh et al., 2001).
37
Generation of the DGSI antibody and immunoblot analysis. A central portion of the
DGSI cDNA (GenBank accession no. AK118777; bp 1029—1503) encoding the amino
acids between the two predicted transmembrane-spanning domains was isolated by PCR
using primers 5'-ACTGGATCCTCTATATGGCTACTA-3' and 5'-
CATGGTACCTCATAATGCAGCCAGAAT-3'. The resulting fragment was digested
with BamHI and KpnI and inserted into pQE30 (Qiagen). The expression strains were
routinely cultured overnight at 37°C in LB ampicillin (100 pg ml'l), inoculated at 1/250
dilution in LB ampicillin (100 pg mf‘) and cultured at 37°C. The protein was induced
with 0.4 mM isopropyl-B-D-thiogalactopyranoside at an optical density at 600 nm (ODGOO)
of approximately 0.4. At 0D600 0.7—0.8, cultures were cooled on ice and cells were
harvested by centrifugation at 5000 g. Cells from 250 ml of culture were resuspended in
1 ml of wash buffer (50 mM Tris pH 7.5, 600 mM NaCl, 20 mM irnidazole) and lysed by
sonication. The protein was purified from cell Iysate using an Ni-NTA agarose column
(Qiagen) according to the manufacturer's instructions. Fractions with the highest protein
content were combined, and the protein concentration was determined by the Bradford
method (Bradford, 1976) using BSA as a standard. Rabbit polyclonal antiserum against
denatured protein in acrylamide was raised by Cocalico Biologicals (Reamstown, PA,
USA). The antibody was affinity-purified as previously described (Ermolova et al., 2003)
using an affinity matrix consisting of 75 ug of the antigenic recombinant DGSI
polypeptide fragment bound to a PVDF membrane.
38
Isolation of mitochondria and chloroplasts. Mitochondria were isolated from leaf
tissue of three-week-old, soil grown A rabidapsis plants and purified using Percoll
gradients according to Keech et al. (2005), while crude cauliflower mitochondrial
preparations were performed as previously described (Douce et al., 1972). Intact
chloroplasts were isolated from Arabidopsis leaves as previously described (Bessoule et
al., 1995). For SDS-PAGE/immunoblot analyses, denatured crude leaf extracts were
prepared as previously described (Martinez-Garcia et al., 1999), while protein fractions
from purified organelles were denatured in a standard SDS-PAGE buffer (Laemmli,
1970). Protein concentration was quantified using the RCDC protein assay kit from
Biorad (Hercules, CA, USA) with BSA as a standard. For Blue Native PAGE,
solubilized proteins from cauliflower or Arabidopsis mitochondria were prepared using
Triton X-100, dodecylmaltopyranoside, or Tween-20 as described in (Eubel et al., 2003)
at 0.2, 0.33, and 1.0 mg detergent per mg protein (all at 25 mg protein ml"). The
supernatant fractions after a 16,000 x g centrifirgation were removed and mixed with
Coomassie Brilliant Blue R-250, and 50 ill from each sample was separated by Blue
Native PAGE (Eubel et al., 2003, Schagger et al., 1994). lrnmunoblotting was performed
on PVDF membranes using the CPS-1 Chemiluminescent peroxidase substrate detection
system from Sigma (St. Louis, MO, USA). Monoclonal antibodies against a conserved
epitope in plant alternative oxidases (AOX mAb) (Finnegan et al., 1999) were obtained
from stock at the Department of Energy Plant Research Laboratory, Michigan State
University, left by the late Lee McIntosh. Monoclonal antibodies against the outer
mitochondrial membrane VDAC protein were provided by Tom Elthon (University of
Nebraska-Lincoln, USA). Polyclonal antibodies against the pea chloroplast outer
39
envelope membrane protein TOC 75 were kindly provided by John Froehlich (Michigan
State University, USA) (Tranel et al., 1995).
RNA Analysis. Total RNA from liquid N2-frozen leaf tissue of two-week-old seedlings
was isolated with the QIAGEN RNeasy Plant Mini-kit (QIAGEN, Germantown, MD)
according to the manufacturer’s instructions and separated by electr0phoresis through a
formaldehyde-denaturing agarose gel. RNAs were then transferred to a nylon filter
which was processed for RNA blotting using ULTRAhyb hybridization buffer (Ambion,
Austin, TX, USA). The probe used to specifically detect AOX 1 a mRNA was generated
by PCR with the following primers: forward, 5’-TTTGTTCTTCGACGGTCCGTAC-3’;
reverse, S’CGCGATTCCTFTATCTCCCT‘TG-T, and labeled with ”P using the
Megaprime DNA Labeling system (GE Healthcare, USA). Total RNAs were stained with
methylene blue as previously described (Wilkinson et al. , 1991).
Lipid Analysis. Lipids were extracted from leaves (Dormann et al., 1995) on an equal
fresh weight basis and separated by thin-layer chromatography on activated ammonium
sulfate-irnpregnated silica gel TLC plates (Si250, Mallinckrodt Baker, NJ, USA) with a
solvent system of acetone/toluene/water (91 :30:7.5, by vol.). Gas chromatography based
quantification of fatty acid methyl esters (FAMEs) was performed as previously
described (Xu et al., 2005). MGDG and DGDG bands were scraped into individual vials,
while the remaining polar lipids were collectively scraped into a single vial for FAME
derivatization (designated as the remainder of polar lipids). Lipids were extracted fi’om
mitochondria as described above, but loaded on TLC plates on an equal total
40
mitochondrial protein basis. For visualization of total lipids separated by TLC, dried
plates were placed in a chamber containing 12 vapors; for visualization of
glycoglycerolipids, plates were stained with a-naphthol (Benning et al., 1995).
Results
Introduction of dgsI-2 does not suppress the dgdl growth phenotype. To further
characterize the function of DGS I , homozygous plants were generated for a potential null
T-DNA allele (SAIL_391_FO40 (Sessions et al., 2002)) inserted into the third exon of
DGSI (AT5G12290) and confirmed by PCR (Figure 2.1A). This T-DNA allele,
designated dgsl-2, results in an insertion of the left border of the T-DNA plasmid at a
codon corresponding to the 159th amino acid—which is N-temrinal to the D457N
mutation caused by dgsl -1 (Figure 2.1B). Analysis of 30-d-old plants revealed no
obvious growth defects in the dgsl-l or -2 mutants compared to wild-type, where as
these alleles had distinct phenotypes when observed as double mutants in a dgdl
background: dgsl -1 suppressed the growth defect of dgdl, whereas dgsI-2 had no effect
(Figure 2.1C). This result is consistent with the hypothesis that dgsl -1 is a gain-of
function allele, whereas dgsI-Z is a loss-of-firnction allele. Moreover, the lack of a clear
phenotype for dgsI-2 in a wild-type background indicates that DGSI is not essential for
normal growth and development.
DGSI is a mitochondrial membrane protein. DGSI encodes a 602 amino acid, ~69
kDa protein of unknown function. DGSI is predicted to contain two transmembrane-
spanning domains when analyzed with the TMHMM 2.0 program (Keech et al., 2005)
(Figure 2. I B), and a BLAST search (Altschul et al., 1997) revealed several DGSI
41
A RP+LP LB1+LP B
M 1 2 1 2 dgs1-2 (T-DNA) 3%217111
V59
Lo
TM1 TM2
Figure 2.1. Comparison of the dgsI-Z T-DNA null allele and the dgsI-I point mutant
gain of function allele in wild type and dgdl backgrounds. (A) Genotyping of wild-
type (1) and dgsI-Z homozygous (2) plants using the PCR primers RP, LP and LB]
described under MATERIALS AND METHODS. (B) Diagram showing the relative
positions of the dgsl -1 and dgsl-Z mutations in the DGSI protein. The relative positions
of two predicted transmembrane spanning domains are also indicated (TMI and TM2).
(C) Growth phenotypes of 30-day-old plants sown directly on soil. The respective
genotypes are indicated along the bottom.
42
Athaliana pIHrrFs E. DQILRANEI AI
A.oryzae PPVGT R DI ID-LLKSQEL Gr
e.glabramPIKNIVT S IDKMLKSQQL GI
c. albicansPIKYLIT S IDKLLKSQQL AM
c.grobosumPIVGMK D ID-LLKSQEL or
admoideum PLVGAVS D IDSLLKSQEL GF
QNL K a DQILKANEI AI
E‘ftfiifl’ Qurllr D IDKLLQANEL QL
Ow," Rim. T D IDKLLQSQQL GI
' . KGLSS K DQILRANEI AI
O'ta‘mPLTt-JNL D DQLNRANRL SL
P-"O'dumprrx'car N IDSILKSQEL GE‘
P-"lChOCfifPapIRrav N IDRLLKSQEL AT
S-cerewaaeprasar N DEVLEGNDL KI
s.pombe PVKSLI s IDQILKSQEL GI
Y. llpolytica IKNQ G SLLIQLQK IDKMLKSQQL Gv
C. albicans
E. gossypii
- - S. cen'visae
7. cruzi Y. Irpolyttca +
+ C. glabrata
+
0 S. pombe
9 a
+ A. oryzae
a. discoidium 2 p nommm
+ N. crassa
+ C. globosum
P. tn'chocarpa + +
A. thaliana
0.1 O. sativa O. taun'
Figure 2.2. Sequence analysis of DGSI. (A) Partial sequence alignment of DGSI
orthologs showing the region of highest identity. Conserved residues are shaded in black
and the asterisk indicated the Asp residue mutated to Asn in dgsl (D457N). (B) Unrooted
tree showing the relatedness of predicted DGSI orthologs in plants and fungi. Boot
strapping values of >950 are marked by +. Those between 500 and 900 are marked with a
solid circle.
43
orthologs in plant, algal, and fungal species—as well as other organisms such as
Dictyostelium discaideum and Ttypanasoma cruzi—but not in animals or prokaryotes
(Figure 2.2). Intriguingly, the Asp457 residue that is mutated in the dgsl-1 allele
(D457N) is entirely conserved among the DGSI orthologs in a region of high local
similarity (Figure 2.2A).
The DGSI protein is predicted to be targeted to the mitochondrion or chloroplast
according to the Arabidopsis subcellular database (SUBA) (Heazlewood et al., 2007 ),
and was identified in the Arabidopsis mitochondrial proteome (Heazlewood et al., 2004).
The fluorescence of a C-terrninally GFP-tagged DGSI fusion protein was associated with
organelles consistent with the appearance of mitochondria, but not chloroplasts (Figure
2.3A). The localization of the DGSI-GFP fusion protein was further confirmed by
overlap of the corresponding GFP fluorescence with Mitotracker Red fluorescence in root
hairs of the DGSI-GFP transgenic plants (Figure 2.3B). Irnunodetection of DGSI with a
specific polyclonal antibody generated in this study revealed that the protein could only
be detected in isolated Arabidopsis mitochondria, but not crude leaf, or isolated intact
chloroplast preparations (Figure 2.3C). The lack of a signal in the mitochondrial fraction
when using an antibody against the outer chloroplast envelope protein TOC75 ruled out
chloroplast envelope contamination. Together, these results suggest an exclusive
localization of DGSI to mitochondria.
Recent proteomic studies on mitochondrial membrane fractions from yeast and
Neurospora identified homologs of DGSI in outer mitochondrial membrane preparations
(Schmitt et al., 2006, Zahedi et al., 2006). To determine the sub-mitochondrial membrane
association of DGS 1 , mitochondria from cauliflower (Brassica oleracea) florets were
g,
-
kDa
180-
115- g ..
82 , 2 § ..
64- 24‘2” ‘:
DGSt-GFP Mitotracker Merged 3, a
Red 49- El. ...... CBB
w- . "U“
C K 37- . ..
(b \ .\°
0 ‘0 \ ,.
V O ‘1‘ ‘ fl
$\°\\°\ 254:-afi.‘
v‘ DGS] 19' ‘
15- !
._. —- -
C" AOX
— TOC7S
Figure 2.3. Localization of DGSI in mitochondria. (A) Confocal microscopy image of
Arabidopsis wild type stably expressing a DGSI-GFP cDNA. The images for chlorophyll
fluorescence (red) indicative for chloroplasts, and for GFP indicative for the presence of
the DGS 1 -GFP fusion protein (green), were overlaid. The scale bar equals 8 pm. (B)
Confocal microscopy images of a single root hair of Arabidopsis wild type stably
expressing a DGS] -GFP cDNA; the GFP and Mitotracker Red fluorescence images are
shown separately, as well as the merged image as indicated. The scale bars equal 10 pm.
(C) Irnmunodetection of DGSI in isolated Arabidopsis mitochondria. Proteins extracted
from wild-type Arabidopsis leaf (Wt leaf), intact chloroplasts (Int Chl) and intact
mitochondria (Int Mito) were separated by SDS—PAGE (50, 80 and 15 pg, respectively),
and immunoblotted with DGSI , AOX (a mitochondrial inner membrane protein) and
TOC 75 (a chloroplast outer envelope protein) antibodies as indicated.
(D) Association of DGSI with sub-mitochondrial fractions from cauliflower.
Immunoblots with lanes containing total protein (Tot Prot), intact mitochondria (Int
Mito), outer mitochondrial membrane (OMM) and inner mitochondrial membrane (IMM)
are shown. Three identical blots were probed with antibodies against DGS 1 , VDAC (an
outer membrane protein) and AOX (an inner membrane protein). Equal amounts of
protein were loaded as indicated by staining a gel run in parallel with Coomassie Brilliant
Blue (CBB), where apparent molecular weights of an included protein marker set are
indicated (kDa).
45
isolated and inner and outer membrane fractions were prepared. The Arabidopsis DGS 1 -
specific antibody detected a protein of the correct size (Figure 2.3D) as well as a lower
molecular weight protein that was not further investigated. As a marker for the outer
membrane, the voltage-dependent anion channel (VDAC, porin) was used; alternative
oxidase (AOX) was included as an inner membrane marker. DGSI co-purified with the
outer mitochondrial membrane marker VDAC (Figure 2.3D), and was clearly not
enriched in the inner mitochondrial membrane fraction as was observed for AOX. These
results are consistent with the presence of DGSI orthologs that have been localized to the
outer mitochondrial membranes of fungi, but co-localization to the inner membrane
cannot entirely be ruled out.
DGSI is present in a high molecular weight complex. One of the simplest
mechanisms—and a common underlying factor—that gives rise to dominant negative
phenotypes is the. poisoning of a protein complex by a mutated component protein
(V eitia, 2007). To test whether complex poisoning was a possible mechanism that
underlies the dgsl-1 phenotype, we explored whether the DGSI protein is present in a
higher M, complex. Initially, mitochondria isolated from cauliflower florets were used as
a facile source that provides high quantity without contamination of chloroplast
membranes. Following solubilization of mitochondrial proteins using increasing amounts
of Triton X-100 and separation of complexes by blue-native PAGE, a high Mr complex
migrating between the 440 and 669 kD markers (approximately 500 kD) containing a
peptide reacting with anti-Arabidopsis DGSI antibody serum is present (Figure 2.4A).
This complex is stable up to 1 mg Triton-X100 per mg protein, the highest concentration
tested. As a control, complex IV was monitored using a monoclonal antibody that
46
Figure 2.4. Blue Native PAGE analysis of the DGS] complex. (A) Cauliflower
mitochondrial proteins solubilized with varying amounts of Triton X-100 (0.2, 0.33, and
1 mg Triton X-100 per mg mitochondrial protein in lanes 1, 2 and 3, respectively) and
separated by blue native PAGE. The Coonrassie Brilliant Blue stained proteins (CBB)
and corresponding immunoblots with the DGSI antibody (DGSI) or the COXII antibody
(COXII) are shown. The relative positions of protein standards are indicated
(thyroglobulin at 669 kD, ferritin at 440 kD, and catalase at 200 kD). (B) Blue Native
PAGE of cauliflower mitochondrial proteins solublized with different detergents Triton
X-100 (lane 1), Dodecylmaltopyranoside (lane 2), or Tween 20 (lane 3), followed by
immunodetection of DGSI. (C) Blue Native PAGE separation of Triton X-100
solubilized crude cauliflower mitochondria (lane 1), and from isolated Arabidopsis
mitochondria from wild-type (lane 2) and homozygous dgsl-1 plants (lane 3).
47
recognizes subunit II (COXII), and was present in a high M, complex (approximately 300
kD) presumably representing complex Vla of the respiratory electron transport chain as
previously described (Eubel et al., 2003). Contrary to the DGSI complex, this complex
was not stable at 1 mg Triton-X100 per mg protein, where only the previously described
(Eubel et al., 2003) complex M was detected (Figure 2.4A). The DGSI immunogenic
complex was not affected by using other detergents, dodecylmaltoside or Tween 20, for
solubilization (Figure 2.48). Isolated leaf mitochondria from Arabidopsis also showed the
presence of a DGS l-immunogenic complex similar in apparent M, to the cauliflower
DGSI complex (Figure 2.4C). The Arabidopsis DGSI complex was indistinguishable in
wild-type and dgsl-1 mitochondria, indicating that the DGS 1 04573; protein produced in
dgsl -1is not defective in complex formation. These data clearly indicate that DGSI is
present in a multimeric complex, but do not provide any evidence as to whether DGSI is
interacting with other proteins (heteromultimer), or forms a homomultimeric quaternary
structure.
Phosphate deprivation and dgsl-1 allelism have additive effects on lipid metabolism.
Phosphate deprivation has been reported to lead to accumulation of DGDG in
mitochondria of plant cells (Jouhet et al., 2004). As DGSI is a mitochondria-localized
protein (Figure 2.3), potential changes in mitochondrial lipid composition in dgsl-l
plants were investigated. As shown in Figure 2.5 no differences in lipid composition were
observed between mitochondria isolated from wild-type, dgdl , or dgdl dgs1-1 mutant
leaves from plants grown under normal conditions. Irnportantly, no DGDG was found in
the dgdl dgsl-1 mitochondria, indicating that the DGDG accumulating in these mutants
is associated with non-mitochondrial membranes. As such, these results suggest that the
48
A 3‘
A ,t .3
39
K N N N
$9 efil a? ~§K 89 $9
r— whole leaf --1 ,— mitochondria—I
MGDG '
PG/DPG ' "
DGDG —
SQDG
Pl/PS
PE
PC
MGDG O - .-
DGDG ’
SQDG
TGDG ........................
Figure 2.5. Mitochondria of dgdl dgsl-I homozygous mutant plants do not
accumulate galactoglycerolipids. Lipid extracts were prepared from leaves (on an equal
fresh weight basis) and isolated mitochondria (on an equal mg protein basis) separated by
TLC and stained with iodine (A), and after de-staining, with a-naphthol (B). Lipids:
DGDG, digalactosyldiacyl glycerol; DPG, diphosphatidylglycerol (cardiolipin); MGDG,
monogalactosyldiacyl glycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine;
PG, phosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; SQDG,
sulfoquinovosyldiacylglycerol; TGDG; trigalactosyldiacyl glycerol.
49
processes of DGDG accumulation due the presence of the dgsl-I allele and during
phosphate-limiting growth conditions may be distinct.
To test whether DGSI is required for DGDI -independent DGDG biosynthesis
following phosphate deprivation, galactolipid content was determined in the dgdl single
mutant and the dgdl dgsl-1 and dgdl dgsl-2 homozygous double mutants under normal
growth conditions (Figure 2.6A) and under conditions of phosphate deprivation (Figure
2.6B). The dgdl dgsl-2 double mutants were indistinguishable from the dgdl mutant with
regard to DGDG accumulation under phosphate deprivation, suggesting that DGSI is not
required for phosphate-dependent activation of the DGDI-independent pathway. The
accumulation of DGDG in the dgdl dgsl-1 homozygous double mutant under phosphate
deprivation was higher than in dgdl, indicating that dgsl-I is additive. This result
provides further evidence that phosphate deprivation and introduction of the dgsl-I
mutation act independently and represent distinct mechanisms of activating the DGD l -
independent pathway(s) of galactolipid biosynthesis.
In addition to the paralog of DGD], DGD2, which is known to be induced
following phosphate deprivation (Kelly and Dormann, 2002), a distinct processive
galactolipid:galactolipid galactosyltransferase (GGGT) discovered in preparations of
plastids (van Besouw and Wintermans, 1978) but not yet defined at the molecular level
might be activated in dgsl-1 as well. This appears to be the case, as analysis of leaf lipid
crude extracts by TLC revealed the accumulation of the oligogalactolipid
trigalactosyldiacyl glycerol (TGDG) in the dgdl dgsl-l double mutant following
phosphate deprivation (Figure 2.6 C). TGDG is a product of the above-mentioned GGGT
50
Figure 2.6. Galactoglycerolipid content in the dgdl single homozygous mutant and
the dgdl dgsl-1 and dng dgsl-2 double homozygous mutants under normal and
phosphate-deprived growth conditions. Monogalactosyldiacylglycerol (black bars) and
digalactosyldiacylglycerol (grey bars) relative amounts present in leaves of 10—day-old
plants grown on MS agar plates (A), and (B) 10 d old plants transferred to MS agar plates
lacking phosphate for an additional 10 days. Relative amounts are expressed on a % mol
basis of polar glycerolipid and the average and standard deviation of three independent
measurements is shown. (C) A TLC plate stained with u-naphthol to reveal the separated
galactoglycerolipids. Plants were grown in the presence (+) or absence of phosphate (-) as
described for (A) and (B). DGDG, digalactosyldiacylglycerol; MGDG,
monogalactosyldiacylglycerol; TGDG, trigalactosyldiacylglycerol.
51
A
dgd1
dgs 1-2 dgs1-1
dgd1
dgd1
dgd1
dgs1-2 dgs1-1
dgd1
dgd1
000000 00000
54321 54321
0
$225 8:... :28 gag 8.2. 3.8
B
C
G
D
G
D
McDG"
52
and is also found in the tgd mutants defective in endoplasmic reticulum-to-plastid lipid
trafficking (Xu et al., 2003). TGDG also accumulates in ozone-treated spinach (Spinacea
aleracea) leaves (Sakaki et al., 1990). This may be related to the ROS accumulation in
dgsl—I mutant plants (Xu et al., 2008), which would also be produced following ozone
treatment.
The fatty acid profile of MGDG and DGDG in the three different mutant lines
showed no difference between dgd1 and the dgdl dgsl-2 homozygous double mutant line
under phosphate replete and depleted growth conditions (Figure 2.7). The dgd1 dgsl-1
homozygous double mutant showed slight differences from dgd1, with a slight increase in
16:3 (carbons: position of double bonds) and decrease in 18:3 fatty acids in MGDG and
lowered relative amounts of 16:0 and increased 18:3 fatty acids in DGDG independent of
the growth conditions.
Alternative oxidase protein levels are reduced in the dgsl-1 mutant allele. In order to
determine if defects in mitochondrial protein composition are present in the dgsl-I
mutant plants, the protein levels of DGSI , alternative oxidase (AOX), and COX II
subunit of complex IV in isolated mitochondria were investigated. Mitochondria were
prepared from leaves of the wild-type, the dgsl-2, dgd1, dgsl-1, and the dgd1 dgsl-1
homozygous single and double mutants, respectively, and protein levels were compared
using immunoblotting. As shown in Figure 2.8, the dgsl-2 T-DNA allele results in the
ablation of the full length DGSI protein, confirming that this is a null or loss-of-function
allele as indicated above. Moreover, the presence of the dgsl-I allele does not lead to a
53
Figure 2.7. Galactolipid Fatty acid profiles. Fatty acid profiles of mono- (MGDG; A,
B) and digalactosyldiacylglycerol (DGDG; C, D) in the dgd1 single homozygous mutant
and the dgd1 dgsl-1 and dgd1 dgsl -2 double homozygous mutants under normal (A, C)
and phosphate-linrited (B, D) growth conditions. The data are presented on a % mol
basis and represent the average and standard deviation from three independent
experiments. Fatty acids are designated with carbon numbers : number of double bonds.
54
C70
Fatty Acid (mol %)
o a a a a a a
Fatty Acid (mol %) U
A N or b 0'! O) N
O O O O O O O
O
Idgdt
, I dgd1
ndgdt dgs1-1
f
‘ 16:0
16:1
' Idgdt
u dgd1 dgs1-2
adgd1 dgst-t
dgs1-2
MGDG (+Pi)
16:2 16:3118207182‘1'182 18:3
MGDG (-Pi)
18:3
16:0 16:1 16:2 16:3 18:0 18:1 18:2
‘ Idgdt
‘ Edgd1 dgs1-2 _
Idgdt dgst-t DGDG (+Pr)
16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3
' Idgdt
adgdt dgs1-2 .
ldgdt dgs1-1 DGDG (-PI)
A a I A 7 I
A
A
16:3 18:0
18:1 18:2 18:3
55
*“um-
__ one
“m “'"‘ ""5 ""‘W Stain
. I. ~ C
.04.». _ H pow m “_
Figure 2.8. Alternative oxidase is reduced in dgsl-1 mutant mitochondria.
Immunoblots of proteins from isolated mitochondria prepared from leaves of wild type
(WT), dgsl-1, dgsl-2 and dgd1 single homozygous mutants, and the dgd1 dgsl-1 double
homozygous mutant are shown. The blots were probed with affinity purified DGSI
antibody (DGSI), an alternative oxidase antibody (AOX), or a cytochrome c oxidase
subunit II antibody (COX 11). Equal amounts of protein were loaded as shown in the
Coomassie Brilliant Blue (CBB) stained gel at the bottom.
56
decrease in DGS I-immunogenic protein; if anything the protein level might actually be
slightly increased in dgsl-1.
An AOX monoclonal antibody that detects all major forms of this protein
(Finnegan et al., 1999) was initially included as a potential loading control for the
experiment. However, it became apparent that AOX levels are strongly reduced in the
dgsl-l gain-of-function mutant but unaffected in dgsl-2 (Figure 2.8). The observed loss
of the AOX protein was not due to a general deterioration of mitochondrial complexes as
COX II was unaffected in any of the mutant backgrounds (Figure 2.8). Moreover, a
corresponding Coomassie Brilliant Blue-stained gel (Figure 2.8, bottom section) did not
show any major differences in the mutant lines, suggesting that there is no large-scale
aberration in mitochondrial protein patterns in the mutant backgrounds analyzed.
Potential defects in the expression of the AOX1a gene of Arabidopsis were
examined. AOXIa is the dominantly expressed AOX encoding gene in Arabidopsis
seedling tissue. To be able to readily detect AOXla transcript levels in seedling shoot
tissue samples, we treated the seedlings with Antimycin A, which specifically inhibits
respiratory electron transport at complex III and leads to a transient induction of A 0X 1a
expression (Zarkovic et al. , 2005). As shown in Figure 2.9, the presence of neither of the
two dgsl mutant alleles affects induction of A 0X1 a mRNA expression. As such, the
reduction of AOX protein levels observed in isolated mitochondria from dgsl-1 mutant
plants must be due to a posttranscriptional mechanism. Moreover, the wild-type DGSI
protein is not involved in regulating AOX protein or transcript levels, at least under the
conditions tested, because the dgsl-2 loss-of-function allele had no effect. The defect in
57
WT dgs1-2
0246810122402468101224
Btu-"911.0 nuns-cu on1a
lam—‘I— an... .— I an..." — - mun—Ar-
mRNA
—--— . . ,u-u- .- - .. .u . .
it...biOv ,. ore-O...’""—.‘“
l
lbaeaclitocoouuoo.4eoilbaour D’.‘."
:f' " -t.:.-.s,.
Lvr...a.a~ ... _.‘._.,.-.-. x ‘_=!=-A‘Ilu" " F~.\er-.I-~ ‘~ -.4 . 3""! l-a . a." — I ' am .n' '0'" "' 1" W '-' Ill
WT dgs1-1
0246810122402468101224
unit-Diva Hil-IfiiiHonta
m-r-a can“ a... - .H. Witt-n
.ygeauuuuHNu—uflnuu
......auaea-«IOIanIQOOueOduso rRNA
‘ -' Iaaoursla..a.... -:
rarer-
Figure 2.9. Analysis of AOXIa transcript levels in seedlings treated with Antimycin
A. Two week-old seedlings of the wild type (WT) and the dgsl-l and dgsl-2
homozygous mutants were sprayed with 25 pM Antimycin A and samples of aerial tissue
were taken at the indicated times (0-24 h). Total RNA (2.5 pg) from these samples were
isolated, processed for RNA blotting and specifically probed with for A OXla mRNA
Methylene Blue stained total RNA (rRNA) is shown as a loading control.
58
AOX protein in dgs] -1 is thus an additional phenotype specific to this gain-of-function
allele as was observed for the above-mentioned lipid phenotypes.
The dgsl-I mutant shows phenotypes of an AOXla-deficient mutant. Arabidopsis
Mutants deficient in AOX generally have subtle biochemical and morphological
phenotypes under normal that become more apparent only when exposed to multiple
environmental stress conditions (Giraud et al., 2008, Strodtkotter et al., 2009, Umbach et
al., 2005). One easily observable phenotype is reduced root growth of AOXla-deficient
lines (Giraud et al., 2008). Comparing root length of seedlings between the different
Arabidopsis lines used in this study, only lines carrying the dgsl-1 allele have shorter '
roots when compared to the respective wild-type or dgdl mutant backgrounds (Figure
2.10). Thus, the dgsl-I mutant plants, which have reduced protein AOX protein levels
phenocopy the reduced root growth observed in AOXla-deficient T-DNA lines. While
this suggests that some of the phenotypes observed in dgsl-I might arise from the AOX
protein defect, there are differences in phenotypes observed between these two mutant
classes: dgsl-l plants accumulate hydrogen peroxide (Xu et al., 2008), which was
reported not to be the case for the AOXla-deficient lines (Giraud et al., 2008).
59
Root length (mm)
_a N (A) «b 0'1 0’
O O O O O O
O
a
WT dgs1-2 dgs1-1 dgd1 dgd1; dgd1:
dgs1-2 dgst-t
Figure 2.10. The dgsl-I but not the dgsl-2 mutant displays a root growth phenotype.
Seedlings were grown for 10 d on vertically placed MS agar plates supplemented with l
% sucrose, and root length was measured. The data represent the average and standard
deviation from 5 individual plants.
60
Discussion
This study was initiated to determine whether the DGSI protein plays a direct role in the
regulation of DGD] -independent DGDG biosynthesis. Initial studies focused only on the
gain-of-function dgsl-1 allele (Xu et al., 2008) which causes severe growth defects upon
stable over-expression. The DGSI protein was implicated as a potential factor involved
in the regulation of alternative DGDG biosynthesis due to the partial suppression of
DGDG deficiency observed in the dgdl dgsl-1 double mutant, which is similar to the
effect of phosphate-deprivation induced DGDG accumulation (Hartel et al., 2000, Kelly
and Dormann, 2002). Given the increased H202 and reduction in AOX protein levels in
dgsl-1 (Figure 2.8), and that the DGDI -indepdendent pathway is induced by exogenous
H202 application (Xu et al., 2008), one could conceive a simple hypothesis for the
possible function of DGSI in phosphate-mediated regulation of galactoglycerolipid
biosynthesis. Phosphate deficiency is known to lead to a slowing of ATP synthesis in
mitochondria (Theodorou and Plaxton, 1993), causing a backup of the electron transport
chain as the two processes are tightly coupled. The resulting over-reduction of the
respiratory electron transport chain leads to ROS formation that can be compensated for
by AOX activity, which can directly transport electrons from ubiquinone to oxygen. If
DGSI regulated AOX activity or abundance in response to phosphate deprivation, a
mechanism by which ROS could serve as a signal for the activation of the DGDl-
independent pathway can be envisioned. However, two observations from this study
disagree with this hypothesis: (i)dgs1-2 mutants completely lack DGSI but are
indistinguishable from wild-type in their response to phosphate deprivation with regard to
galactolipid metabolism at the level tested, and (ii) DGDG accumulation due to the
61
presence of the dgsl -1 allele and phosphate deprivation are additive and therefore involve
two independent mechanisms. Based on these observations it can be concluded that
DGSI is not directly involved in the low phosphate induction of DGDG biosynthesis by
the DGDl-indepdendent pathway.
Regardless, this work has revealed a clearer understanding of how the dgsl-l
allele may cause the activation of DGDI-independent DGDG accumulation. As
previously observed, this pathway is induced by ROS, which is present in elevated levels
in dgsl -1plants (Xu et al., 2008). In both dgsl-1 and H202-treated wild-type plants,
transcripts are elevated for the MGDG synthase encoding paralogs of MGD] : M602 and
MGD3 (Xu et a]. , 2008). The accumulation of TGDG in phosphate-deprived dgd1 dgs]-
! double mutants indicates that GGGT, which remains to be identified at the
molecular/genetic level, is also contributing to DGDG biosynthesis in dgsl-1. The role
of GGGT in lipid biosynthesis is poorly understood, but is known not to contribute
significantly to DGDG biosynthesis under normal conditions (Kelly et al., 2003). GGGT
is activated in the tgd lipid trafficking mutants in Arabidopsis (Xu et al., 2003, Xu et al.,
2005) and in ozone-treated spinach (Sakaki et al., 1990). The conditional activation of
GGGT in dgsl-1 further demonstrates the importance of identifying the GGGT encoding
gene(s), before a more complete picture of galactolipid biosynthesis can emerge.
The reduction of AOX protein in dgsl-1 mutants seems a plausible cause of the
observed accumulation of hydrogen peroxide in these plants. Indeed, a role for AOX in
the alleviation of mitochondrial ROS has been well documented (Fiorani et al., 2005,
Maxwell et al., 1999, Umbach et al., 2005). This conclusion appears to be in conflict
with a recent report that AOX l a-deficient mutants do not accumulate hydrogen peroxide
62
(Giraud et al., 2008). However, a more recent report suggests that one of the other (five in
total) AOX encoding paralogs in Arabidopsis, AOX 1d, is induced in the AOXla mutant
background and may partially compensate for AOXla deficiency (Strodtkotter et al.,
2009). Thus, at this time the question of whether the reduction of AOX protein observed
in dgsl-1 is the underlying cause of hydrogen peroxide accumulation in this mutant line
remains. How the dgsl-l mutant protein affects AOX protein levels also remains
unresolved, but based on our AOX l a transcript analysis the effect must be
posttranscriptional. Because DGSI is present in the outer membrane (Figure 2.3), while
AOX is associated with the inner membrane, direct interactions between the two seem
unlikely. It is also not likely that the DGSI wild-type protein is required for the
processing or import of AOX protein into the mitochondria, because AOX protein levels
are not affected in dgsl-2 plants. Moreover, the presence of DGSI orthologs in
organisms that lack AOX (e.g., Saccharamyces cerevisiae) also suggest that DGSI is not
necessarily involved in the regulation of AOX. Thus, the effect of the dgsl-I allele on
AOX protein levels may not be directly related to the function of the DGSI wild-type
protein.
This work has revealed that the DGSI protein is not directly involved in the
regulation of DGDl-independent galactolipid biosynthesis during phosphate deprivation,
as originally hypothesized. The reduction in AOX protein also seems to be an indirect
effect of the dgsl -l mutation, and if not directly involved in either of these processes,
what is the molecular function of DGS l ? This question has remained difficult to answer
because analysis of dgsl-2 has lead to no discemable phenotypes under the stress
conditions tested. The biological function at the organismal level of many Arabidopsis
63
proteins including well-studied examples, such as AOX, is not clear as well. Many plant
proteins are likely to be of conditional importance, and help sessile plants to cope with a
number of environmental stresses. Indeed, the conditions under which the loss of the
DGSI protein becomes critical have not yet been identified. While the phenotypes
associated with the gain-of-function dgsl-1 mutant may be more fascinating to the lipid
researcher, a rigorous analysis of the dgs1-2 loss-of-function mutant under gauntlet of
conditions, or detailed biochemical study of the DGSI protein complex and potential
interacting partners, will be required to provide insight into the molecular function of
DGSI.
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68
Chapter 3
Identification of the gene encoding galactolipid:galactolipid galactosyltransferase
and its role in freezing tolerance
2 The data represented in figures 3.2 and 3.8 are derived from experiments conducted by
Bagyalakshmi Muthan.
69
Abstract
UDP-galactose independent biosynthesis of di-, tri-, and tetra-galactoglycerolipids
(DGDG, TGDG, and TeDG, respectively) has long been known to occur in plants. This
distinct pathway of galactolipid biosynthesis is carried out on the chloroplast outer
envelope by galactolipid:galactolipid galactosyltransferase (GGGT). GGGT transfers a
galactosyl moiety from one MGDG molecule to another, forming DGDG and
diacylglycerol, and through processive activity produces oligogalactolipids (TGDG and
TeDG). Prior to this study, the gene(s) encoding GGGT were not known, and the role of
GGGT in lipid metabolism remained enigmatic. Here, the GGGT encoding gene is
identified as SENSITIVE T0 FREEZING 2 (SFR2), a gene of unknown function, which
was previously identified when sfiz mutants showed freezing sensitivity. In viva and in
vitro studies indicate the enzyme is activated by stresses including freezing, dehydration,
and hyperosrnotic treatments. T-DNA mutants in SFR2 cannot produce oligogalactolipids
and exhibit sensitivity to freezing, suggesting a role for GGGT in stabilizing the outer
envelope during this stress. The SFR2 protein shows GGGT activity when expressed in
yeast. Together these results indicate that GGGT is activated under sub-zero
temperatures and remodels galactolipids in the chloroplast outer envelope—possibly by
direct mechanosensation of physical changes that occur in the membrane during freezing.
70
Introduction
The biosynthesis of glycerolipids has been intensely studied, not only because
these molecules are important building blocks for biomembranes that provide the
boundaries of the cell and organelles, but also because membranes are the site of signal
and energy transduction processes that often require distinct lipid compositions for proper
function. Molecular genetics has been a powerful tool in elucidating the factors required
for glycerolipid biosynthesis in plants, where classical biochemical approaches to purify
what are often integral membrane enzymes has been of more limited success. One
example of this is the identification of the enzyme and encoding gene, DGD],
responsible for the bulk of DGDG biosynthesis in Arabidopsis. The dgd1 null mutant
exhibits a severe growth defect, and has a ~90 % decrease in DGDG levels in comparison
to wild-type (Dormann et al., 1995, Dorrnann et al., 1999). Analysis of DGDl revealed it
to be a UDP-galactose (UDP-Gal) dependent enzyme which acts on
monogalactosyldiacyl glycerol (MGDG) as a lipid substrate. These findings were in
seeming conflict with earlier biochemical studies, which suggested that further
galactosylation of MGDG to form DGDG was carried out in a UDP-Gal independent
manner by the unidentified enzyme galactolipid:galactolipid galactosyltransferase
(GGGT) (Heemskerk et al., 1990, van Besouw and Wintermans, I978). GGGT catalyzes
the transfer of a galactosyl moity from one MGDG molecule to another, forming
diacylglycerol (DAG) and DGDG. Through processive activity, DGDG can be further
galactosylated, producing oligogalactolipids (TGDG and TeDG) (Benning and Ohta,
2005, Heemskerk et al., 1983, van Besouw and Wintermans, 1978).
71
More recent studies in Arabidopsis revealed a DGD] paralog, DGDZ, which
comprised an alternative pathway for DGDG biosynthesis that is activated at the
transcript level upon phosphate deprivation. Intriguingly, the dgdl dgd2 double mutants
had no detectable levels of DGDG, but chloroplasts isolated from these mutants were not
defective in radiometrically measured GGGT activity (Kelly et al., 2003). These results
clearly indicated that a distinct gene is encoding GGGT, and that GGGT does not play a
significant role in DGDG biosynthesis under normal growth conditions. The role GGGT
plays in galactolipid metabolism has remained unresolved, but other studies have
indicated that GGGT is activated in response to ozone treatment in spinach leaves
(Sakaki et al., 199%, Sakaki et al., 1990c), and also in the lipid trafficking defective tgd
mutants in Arabidopsis (Awai et al., 2006, Lu et al., 2007, Xu et al., 2003, Xu et al.,
2008), indicating that GGGT may be involved in stress responses.
This study was initiated to identify the GGGT encoding gene using a facile
mutant screen in Arabidopsis, where infiltration of plant leaves with MgCl2 results in the
accumulation of oligogalactolipids. Initial studies focused on finding mutants defective
in oligogalactolipid accumulation in an ethane methyl sulfonate (EMS) mutagenized
population of Arabidopsis. However, it was an alternative approach screening T-DNA
lines of candidate genes—selected on the basis of the localization of the encoded protein
and its potential to catalyze the GGGT reaction (glycosyltransferases and glycosyl
hydrolases)——which lead to a candidate GGGT encoding gene.
72
Materials and methods
Plant growth conditions. Plants were grown on soil or agar solidified Murashige and
Skoog (MS) medium Murashige and Skoog, 1962) containing 1 % sucrose as previously
described (Xu et al., 2005) on a 16 h light] 8 h dark photoperiod. Cold acclimation was
achieved by placing the plants in a cold room at 7 °C at a photosynthetic photon flux
density of 75 pmol rn'2 sec’2 with a 12 h 1i ght/ 12 h dark photoperiod. Freezing conditions
were imposed by incubating plants at -2 °C (24 and 12 h total for MS-agar and soil grown
plants, respectively) with ice nucleation after 3 h (Li et al., 2008). This was followed by
lowering the temperature at 1 °C h’1 to -6 °C and incubation for 24 h. For post freezing
recovery, plants were returned to the above-mentioned cold acclimation conditions for 24
h, and then returned to normal growth conditions.
Isolation of sfr2 T-DNA mutants and complementation of sfi-2-3 with SFR2 cDNA.
The sfi2-3 and sfr2-4 T-DNA lines (SALK_106253 and SALK_112965, respectively)
were obtained from the Arabidopsis Biological Resource Center (Ohio State University,
Columbus, OH). Homozygous T-DNA insertion mutants were identified by PCR-based
genotyping using the following primers: sfr2-3 F P, 5’-GAGCTTGATGT'I‘CTGGAACT-
3’; sfr2-3 RP 5’-TTGAACTTTGAGCTGTCG-3’; sfr2-4 FP, 5’-
CACCAAGATTGT'TGATGCATG-3’; sfi2-3 RP, 5’-
TTAGTCCAGCACCACACACTG-3’. Amplification with these primers indicates the
presence of wild-type genomic DNA at the respective allele; amplification with the sfr2-3
or sfr2-4 RP primers and the T-DNA left border primer LBb1.3, 5’-
ATTTTGCCGATTTCGGAAC-3 ’, indicates the presence of the respective T-DNA
insertion. To genetically complement sfi'2-3, the SFR2 open reading fiame was amplified
73
from cDNA by PCR using gene specific primers 5’-
AAAAAGCAGGCTTCATGGAAT'TATTCGCATTGT'T-3 ’ and 5 ’-
AGAAAGCTGGGTCCTAGTCAAAGGGTGAGGCTA-3 ’. Adapter primers 5 ’-
GGGGACAAGTTTGTACAAAAAAGCAGGCT-3’ and
5’GGGGACCACTTTGTACAAGAAAGCTGGGT-3’ were used for secondary PCR to
introduce Gateway (Invitrogen, Carlsbad, CA) cloning sequences. The resulting PCR
product was inserted into the vector pDONR zeo (Invitrogen) and confirmed by
sequencing. After confirming the sequence, the insert was sub-cloned into the Gateway
binary vector pMDC32 (Curtis and Grossniklaus, 2003). The resulting plasmid DNA was
used to transform the sfi'2-3 T-DNA mutant by Agrobacterium floral dip method (Clough
and Bent, 1998). Seeds were collected from 6 different transformed plants and T1
transgenic plants were selected in the presence of hygromycin B (20pg/ml) on MS
medium containing 1% sucrose. The independent T. transgenic plants were genotyped
using the following PCR primers: P2, 5’-TI‘GAACTTTGAGCTGTCG-3’, and LBb1.3 to
confirm the presence of sfr2-3 T-DNA and P1, 5’-
CACCATGGAATTATTCGCATTGTTAA-3’, and P3, 5’-
TAGAGGGTGGCTTATCAGCTGC-3’, to simultaneously confirm the absence of wild-
type genomic DNA at the sfi2-3 allele and presence of transgenic SFR2 cDNA. After
confirming the genotype, the leaves from independent T1 transgenic plants were
phenotyped for oligogalactolipid accumulation as described below.
Lipid analysis. Lipids were extracted from fresh or liquid N2-frozen MS-agar-grown
plant tissue as previously described (Dormann et al., 1995). Lipids were separated by
thin-layer chromatography (TLC) and visualized with H2SO4 by charring for total lipid
74
staining, or glycolipid staining with u-naphthol (Benning et al., 1995). Quantification of
lipids by gas chromatographic (GC) analysis of derived fatty acid methyl esters was
performed as previously described (Lu et al., 2008). Electrospray ionization mass
spectrometry (ESl-MS/MS) was used as previously described (Bates et al., 2009) to
analyze triacylglycerol (TAG) molecular species isolated according to (Xu et al., 2005).
For 1H NMR analyses, 0.5 — 1.0 mg of DGDG was isolated from Arabidopsis leaf tissue
or a scaled-up in vitro GGGT reaction (see below) by TLC. Lipids were eluted into
CHC13:methanol (2:1 by ml), and were subsequently dried under N2 gas stream. Lipids
were re-dissolved in CDC13:CD3OD:C2D500D (10:10:1 by vol.) and analyzed on a
Varian-SOO spectrometer at 500 MHz as previously described (Xu et al., 2003). Chemical
shifts were measured in relation to residual CHCl3, and data was analyzed using Mnova
software (Mestralab Research, Escondido, CA).
Heterologous expression of SFR2 in yeast. The Open reading frame of SFR2 was
amplified by RT-PCR using Phusion polymerase (New England Biolabs, Beverly, MA)
with the primers 5’-CACCATGGAATTATTCGCATTGTTAA-3’ (SFR2-F) and 5’-
TCAATQATQATQATQATGATQGCCGTCAAAGGGTGAGGCTAAAG-3’ (SFR2-
R); the underlined sequence indicates the coding sequence for a 6 x His-tag added
directly to the C-tenninus of the SFR2 protein. The amplified product was cloned into
the pYESZ. IN 5 His TOPO vector (Invitrogen) to create the vector pYESZ. 1 -SF R2,
which was verified by sequencing. Saccharamyces cerevisae (Strain SCY62; genotype:
MA Ta ADE2 (Sandager et al., 2002)) was transformed with pYESZ. l -SFR2 or the
corresponding negative control (pYESZ). Expression of SFR2 was induced by
inoculating a culture of SC minimal medium containing 2% (w/v) galactose, and 0.5 %
75
(w/v) raffinose to 0D,,00 0.4, followed by continued growth for 8 h at 28 °C. Cells were
pelleted and frozen in liquid N2 and stored at -80 °C. Membranes were isolated from
these cells according to (Dahlqvist et al., 2000) and stored at -80 °C.
Protein quantification and immunoblotting. Membrane protein content was assayed
using the RCDC kit (Biorad, Hercules, CA). For immunoblotting, samples were
denatured according to (Martinez-Garcia et al., 1999), and separated on 12 % SDS-
PAGE gels. Proteins were transferred to PVDF membranes and the recombinant C -
terminally His6-tagged SFR2 protein was detected with anti His(C-term) monoclonal
antibody (R930-25, Invitrogen) using the CPS chemiluminescent detection kit (Sigrna-
Aldrich, St. Louis, MO).
In vitro GGGT assay. The assay was modified from (Heemskerk et al., 1983) as
follows: monogalactosyldiacylglycerol (MGDG) from Larodan (Malmo, Sweden) was
dried under a N2 gas stream (64 nmol per reaction), and overlayed with 150p] per
reaction assay buffer (10 mM Tricine, adjusted to pH 7.2 with Tris, 10 mM MgCl2, 0.426
mM sodium deoxycholate), which results in a final MGDszeoxycholate molar ratio of
1:2 . The reaction components were dispersed by sonication 3 x 10 sec with a Sonicator
3000 (Misonix, F armingdale, NY) equipped with a 1.6 mm diameter microtip at power
setting 1.0. The reaction was initiated by the addition of 15 pg protein equivalents of
isolated membranes (above) followed by sonication 1 x 10 sec as above. Reactions were
stopped by the addition of 600 pl methanol:CHCl3 (1 :2 by vol.). After phase separation
by addition of 600 pl 1 M KCl, 0.2 M H3PO4, and re-extraction with chloroform, lipids
were separated by TLC as above. For radiometric assays, [U-3H-Gal]-MGDG,
specifically labeled on the headgroup was biosynthesized using intact spinach
76
chloroplasts as previously described (Heemskerk et al., 1983). The specific activity of
[U-3H-Gal]-MGDG was determined by GC and scintillation counting, and assays were
performed exactly as above with 1.2 x 104 DPM [U-3H-Gal]-MGDG.
Results
Development of a TLC-based screen to identify Arabidopsis mutants defective in
GGGT activity. Since the diagnostic products of GGGT activity, TGDG and TeDG, are
not constitutively produced in wild-type plants, a treatment which activates this enzyme
in viva was sought out. Earlier biochemical studies suggested increasing amounts of
MgCl2 activate GGGT activity in chloroplast envelope preps (Heemskerk et al., 1983,
Sakaki et al., 1990a). A series of MgCl2 solutions of increasing concentration (0-400
mM) were used to treat Arabidopsis leaves by submergence, followed by vacuum
infiltration and 16-24 h of continued incubation under light. As seen in Figure 3.1,
treatment with M gCl2 solutions above 100 mM resulted in the accumulation of
oligogalactolipids, and 400 mM MgC 12 was used for routine screening of an EMS
mutagenized Arabidopsis population. In ~1500 individual plants analyzed in a
collaborative work with Bagyalakshmi Muthan, a lab technician, and an undergraduate
student, Jason Sherbarth, one putative mutant defective in oligogalactolipid accumulation
was recovered. To our misfortune, this mutant failed to develop and make any seeds.
However, before a second isolate of this putative mutant could be recovered, an
alternative approach using the same treatment to reverse-genetically screen Arabidopsis
T-DNA lines yielded a candidate GGGT encoding gene, as further described below.
77
[M9051 (mM)
0 25 100 200 400
MGDG
DGDG
'TGDG
TeDG
Figure 3.1. Activation of GGGT activity in Ieaftissue by vacuum infiltration of
MgClz. Galactolipids stained with a-naphthol after separation by TLC. Wild-type leaf
samples were treated by vacuum infiltration in solutions containing the indicated
concentration of MgCl2, followed by 24 h incubation.
78
T-DN A mutants in the SFR2 gene are defective in oligogalactolipid accumulation.
Lines for a reverse-genetics approach to identify the GGGT encoding gene were selected
through two criteria: (i) the encoded protein should be present in the available chloroplast
envelope proteomics data sets (Brautigam et al., 2008, F erro et al., 2002, Ferro et al.,
2003, Froehlich et al., 2003)—particularly in the outer envelope where GGGT activity is
unambiguously localized (Dome et al., 1982), and (ii), the encoded protein should be a
member of an enzyme family involved in glycosyl transfer (either glycosyl hydrolases or
glycosyltransferases) (Heinz, 1996). For the outer envelope localized glycosyl hydrolase
family 1 (GH-l) encoding gene, SENSITIVE T 0 FREEZING 2 (SFR2), two homozygous
T-DNA lines, sfr2-3 and sfr2-4, were confirmed by PCR (Figure 3.2). When screened
using the MgCl2 infiltration assay, neither of the sfr2 mutants were able to accumulate
oligogalactolipids, whereas the dgdl and dgd2 mutants defective in UDP-Gal dependent
DGDG biosynthesis were similar to wild-type (Figure 3.3).
The sfr2 mutants show defects in lipid remodeling during freezing stress. SFR2 was
previously identified by other researchers in Arabidopsis, when null mutations in this
gene were found to result in freezing sensitivity (Thorlby et al., 2004); however, the
substrates and products of the encoded glycosyl hydrolase enzyme remained unknown, as
did the precise mechanism through which SFR2 activity contributed to freezing tolerance
(Fourrier et al., 2008). As seen in figure 3.4, the sfr2 mutant alleles exhibited freezing
sensitivity similar to the previously reported alleles (McKown et al., 1996, Thorlby et al.,
1999, Thorlby et al., 2004, Warren et al., 1996). Importantly, freeze-treated wild-type
79
Figure 3.2. Genotypic confirmation of the sfr2 T-DNA mutant alleles used in this
study. (A) Schematic diagram of the SFR2 Gene. Exon (boxes) and intron (lines) regions
are indicated as well as the open reading frame (black bkgd.) and 5’- and 3’-UTRs (white
bkgd). The positions of the T-DNA insertions of the sfr2-3 and -4 alleles are indicated,
as well as other mutant alleles used in other studies (sfr2-I and sfr2-2, first described by
Thorlby et al., 2004 as sfiZ-I and sfi2-i2, respectively). PCR based genotyping to
confirm the homozygosity of sfr2-3 (B) and sfr2-4 (C) alleles. PCR products were
separated on 1% agarose gels after amplification with gene allele specific primers (FP
and RP), or a T-DNA left border and complimentary gene allele specific primer (LB and
RP).
80
1Kb
B WT sfr2-3 WT sfr2-3
sfr2-3 FP/RP sfr2-3 LBIRP
0 WT sfr2-4 wr sfr2-4
sfr2-4 FP/RP sfr2-4 LBIRP
81
no treatment M9012 Treatment
r eff is" 30" 30" a" s e6“ at is” a" c8 ’”
MGDG
_' TGDG
I. A . > - .-
quiz-L . 3g. ,
. ‘ .. F. . bay—:1! - " 13.-
4 - I I ~ ' V ‘ ' _‘.
i U ',l‘5,§72~{~3 31", 3; [”3331 ' as: .. ‘5 . ~-‘. .
... L: >_.. :5 .- 11‘ -:. - - ‘.' ' ,- .
- - .- -7 flag 3 -;~-.-. :44 26,-! _'.~
, ‘j'L. ,1 s -4__ ', ,__-- -_ ' <
‘3 .JJ‘
Figure 3.3 Accumulation of oligogalactolipids in galactolipid biosynthetic mutants.
Lipids were separated by TLC and stained with a-naphtbol after extraction from fresh
leaves, or leaves submerged in 400 mM MgC12, vacuum infiltrated for 5 min, and
incubated for 16 h. Gene names and corresponding gene loci from The Arabidopsis
Information Resource (http://www.arabidopsis.org) for the included mutants are: dgd1,
Digalactosyldiacylglycerol synthase I (AT3611670); dgd2, Digalactosyldiacylglycerol
synthase 2 (AT4G00550); s 2-3 and sfi'2-4, Sensitive T a Freezing 2 (ATBGO6510);
tgd] -I , T rigalactasyldiacylgl ycerol I (AT 1G19800).
82
Figure 3.4. Freeze-induced galatolipid remodeling is not observed in sin mutants.
(A) Appearance of wild-type (C012) and sfr2 mutants at 5 d post freezing treatment. (B)
Thin layer chromatogram of lipid extracts stained for glycolipids with a-naphthol reagent
from cold-acclimated (CA) or freeze-treated (FT) wild-type (WT) and sfr2 plants. (C)
Changes in galactolipid levels in plants treated as in (B). ‘P < 0.05 or "P<0.01 vs. WT
CA levels for 3 biological repeats.
83
B WT sfr2-3 sfr2-4
C mol % polar lipids
2.01.51.o 0.5 0 0 1‘0 20 so 49 so
A
leaves accumulated the oligogalactolipids TGDG and TeDG with a concomitant decrease
in MGDG, while the sfr2 mutants did not (Figure 3.4B, C).
Accumulation of triacylglycerols has been reported to coincide with the activation
of GGGT in both the above-mentioned tgd mutants and in ozone-treated spinach (Awai et
al., 2006, Lu et al., 2007, Sakaki et al., 199%, Sakaki et al., 1990c, Xu et al., 2005, Xu et
al., 2008). Indeed, TAG was found to accumulate to ~6% of total fatty acids as a result
of freezing treatment when compared to cold acclimated wild-type plants (Figure 3.5A,
B). The sfr2 mutants showed defect in TAG accumulation and a large decrease in 16:3
(carbonzdouble bonds) acyl groups esterified to TAG in freeze-treated plants compared to
wild type (Figure 3.5C). The presence of 16:3 containing TAG species in wild-type
leaves was confirmed by mass spectrometry (Figure 3.6). The 16:3 acyl group is
predominately found esterified to MGDG in Arabidopsis (Browse et al., 1989), indicating
that DAG produced by GGGT activity is, in part, frnther acylated to TAG.
Effect of various salts and osmolytes on oligogalactolipid and TAG accumulation
following vacuum infiltration. To test whether there was any specificity for Mng in
activating GGGT by vacuum infiltration, a variety of other salts were tested for their
ability to result in the accumulation of TGDG (Figure 3.7A) or TAG (Figure 3.73). All
salts tested (MgSO4, CaCl2, BaCl2, NaCl, and NH4Cl—all at 0.4 M) activated TGDG and
TAG accumulation. These results indicated that neither the identity nor the ionization
potential of the cation or anion in the salt were specifically required. To test whether the
high ionic strength present in the various salts tested, in general, was required for TGDG
and TAG accumulation, various concentrations of the non-ionic osmolytes glycerol and
85
Figure 3.5. Freeze-induced Triacylglycerol accumulation. (A) Thin layer
chromatogram of neutral lipids visualized by H2SO4 and charting fi'om wild-type (WT)
and sfr2 plants treated as in Figure 3.4. (B) Percent of total fatty acids esterified to TAG
in plants treated as in (A); the average and standard deviation of at least three biological
repeats are shown. (C) Fatty acid profile of TAGs in plants nested as in (B) shown on a
mol % basis of the average and standard deviation of at least three biological repeats; "P
< 0.01 or """"P<10'3 vs. WT FT levels. The labels along the X-axis indicate the fatty acid
species identified (number of carbonsznumber of double bonds).
86
A
WT
sfr2-3 sfr2-4
CA
FTFTFT
TAG-
FFAs-
DAG-
B
8 .
(I) .
:9 7
s 6‘
E 5 '
a 4-
(D 3 .
<
P 2 I
s 1 ,
0 .
WT
C CA
60' nwr CA
50‘ IWI'FT
esfi2-3 FT
40‘ nsfi2-4 FT
101
Fatty acid profile (mol %)
00
9
O
. u 'H H
16:0 16:1 16:3 18:0 18:1 18:2 18:3
wr sfr2-3 sfr2-4
FT FT FT
87
>
52 x 541x
1001 ‘ '
8
C
(B
D
C
3 A
it e".
0
>
'5
g
i 1111
0 1 ._, Ill vllr lllerJn ‘ illi llllllll. ‘ lllllu. ' Y. . Y 1 In #I. 7 ' 1.1 .‘l T I
800 820 840 860 880 900 920 940 960 980 1000
B mlz
100. / lM+NH} NH}18:3C Rcoo'l Daughter ions of 863
l / {mun} NH}16 3c RCOO]
Relative Abundance
(°/o)
0-11 . --
540 550 560 570 580 590 600 610 620 630 640 650 660 670 680 690
C mlz
[M+NH} NH}18:2C Rcoo'] Daughter ions of 865
100' \[Mi-NH} NH}18:3C Flood]
32 ~ [M+NH} NH}16:30 R000]
8
fl
3
.0
2‘.
g
.9
0
a:
540 550 660 570 580 590 600 610 620 630 640 650 660 670 680 690
mlz
Figure 3.6. ESI-MS/MS of triacylglycerols isolated from freeze-treated wild-type
plants. (A) ESI-MS spectra of TAGs isolated from fi'eeze treated wild type leaves. The
numbers above the brackets indicate the number of carbons present in the three acyl
chains; x denotes the variation in degree of unsaturation of the acyl chains. ESI-MS/MS
spectra of the daughter ions of parent ions 863 m/z (B) and 865 (C). Deduced identities
of the daughter ions are indicated in brackets.
88
Figure 3.7. Salt and osmolyte treatments that activate TDGD and TAG
accumulation in wild-type leaves. a-naphthol stained galactolipids (A, C) and
triacylglycerols visualized by H2804 charring (B, D) after separation by TLC from leaves
treated by vacuum infiltration with water (H20) or the indicated salt solutions at 0.4 M
(A, B), or by the osmolytes glycerol and sorbitol (C, D). Vacuum infiltration was for 5
min, followed by 16 h incubation before lipid extraction; 4 M-V indicates that vacuum
infiltration was not used, and the tissue was only submerged in the given osmolyte at 4
molar. In (C) and (D), the numbers indicate the molarity of the osmolyte, and water and
MgCl2 controls as treated in (A) were included.
89
>.
2
V
0
Sorbitol
20410204
0204102040v'0
sorbitol were employed in the vacuum infiltration assay. As seen in Figure 3.7C-D, use of
solutions at a concentration 2 1.0 M of the osmolyte were sufficient to cause TGDG and
TAG accumulation, and at 4.0 M, vacuum infiltration was unnecessary for this effect.
These results indicated high extracellular osmotic pressure is the underlying factor in the
activation of GGGT in planta by this method.
Genetic complementation of sfr2-3 by transgenic over-expression of the SFR2
cDNA. To confirm that the T-DNA insertion in sfi2—3 is causing the oligogalactolipid
accumulation defect described above (Figures 3.3 and 3.4), the SF R2 cDNA was cloned
from wild-type tissue, and inserted into the constitutive expression vector pMDC32
(Figure 3.8) as described in “Methods”. The sfr2-3 mutant plants were transformed with
this plasmid, and the resulting T1 progeny for seven individual transformants were found
to accumulate oligogalactolipids when tested with the MgCl2 infiltration assay (Figure
3.8B-C). These results clearly indicated that the presence of transgenic, wild-type SF R2
cDNA was sufficient to complement the TDGD accumulation defect phenotype of sfi'2-3.
Heterologous expression of the SFR2 protein in yeast and analysis of in vitro GGGT
activity. The SFR2 protein was produced in yeast by expression of the SFR2 cDNA and
localized to the isolated membrane fraction by immunoblotting SDS-PAGE separated
protein preparations with an antibody that recognizes a c-terminal 6 x Histidine tag added
to SFR2 during cloning (Figure 3.9A). The protein:fatty acid ratios (ug nmol") of
membranes isolated from empty vector and SF R2-expressing strains were similar (Figure
3.98). Incubation of SFR2-containing membranes with deoxycholate dispersed MGDG
91
Figure 3.8. Complementation of the sfi-2-3 T-DNA mutant with SFR2 cDNA. (A)
Schematic diagrams of the A. thaliana chromosome 3 (AtChr3) structure of the 5’ end of
the SFR2 encoding gene and the SFR2 cDNA expressing transgenic insertion derived
from the plasmid pMDC32 +SFR2. Positions of primers used in genotyping (B) are
indicated. (B) Genotyping of C012, sfr2-3 and sfr2-3 :: SFR2 complementing lines with
primers that specifically detect the sfr2-3 T-DNA insert (LBb and P2), and primers which
detect the presence of a wild-type genomic copy of SFR2 (638 bp), or the presence of
transgenic SF R2 cDNA (402 bp). (C) Phenotypic complementation of sfi'2-3
oligogalactolipid accumulation defect in the sfi2-3::SFR2 lines. leaves from plants were
vacuum infiltrated with 400 MgCl2 and incubated for 16 h, followed by separation of
extracted lipids by TLC and visualization of galactolipids (MGDG, DGDG, TGDG and
TeDG) by staining with a-naphthol. Only the 7 complementing lines genotyped in (B),
analyzed on two separate TLC plates are shown, along with wild-type and sfi2-3
controls.
92
//L“ 2x 358
ms 577 +SFR2
B .5 sfr2-3 :: SFR2
db ԤL 1 2 3 4 5 6 7
LB
+
P2
0.. P1
(I'll-DII'IIIH-Illliilllas
(3 PER” meez
m + - --~ .....
l-I‘ .' -*0 a . ‘17
MGDG "'7’” “‘wrfm
1:11:— .‘ ,. ,1.
DGDG1"1!H!5'!E!!!P .p-I-IinIUIQp»
TGDG " i ,
TBGDG ref . w- m
“P 1 2 3 ‘1’ 4 5 6 7
E % sfr2-3 :: SFR2 E % sfr2-3 :: SFR2
93
A A N N
c or 'o '01
l l l I
pg protein nmol'1 fatty acid
0
'ur
O
I
EV +SFR2
Figure 3.9. Heterologous expression of SFR2 in Saccharamyces cerevisae. (A) Anti
His (C-term) antibody immunoblot (upper panel) and Coomassie Brilliant Blue stained
gel (lower panel) of SDS-PAGE separated membrane protein preparations (25 pg
protein) for empty vector control (EV memb.) and SFR2 expressing (+SFR2 memb.)
yeast strains; soluble proteins fiom the SFR2 expressing strain (+SF R2 sol.) were also
included. The positions of a molecular weight standard are indicated in kDa. (B) Analysis
isolated membrane fractions for protein content (ug) normalized on a per nmol total fatty
acid basis for the empty vector and SFR2 expressing strains as described in (A).
94
resulted in the production of DGDG, TGDG, and TeGDG at l h incubation, whereas
empty vector control membranes did not (Figure 3.10A). In a time course using [U-3H-
Gal]-MGDG, SFR2-containing membranes converted ~60% of labeled MGDG into
DGDG and oligogalactolipid products in 3 h. This conversion was markedly reduced by
the addition of EDTA equivalent to Mg2+ ions present (10 mM) (Figure 3.10B-C) as was
previously observed in chloroplast envelopes (Heemskerk et al., 1983). The fatty acids
16:3 and 18:3, specific to the plant derived MGDG added to the assay, were present in
the DAG pool only in the SF R2 protein-containing reactions—indicating DAG is being
produced from MGDG by SFR2 in vitro (Figure 3.10D).
Proton-NMR spectra of DGDG isolated from a scaled-up reaction showed the
glycosidic linkages to be all in the fl-nomeric configuration (flflDGDG), which is distinct
from the flaDGDG synthesized by the UDP-Gal dependent DGDI/2 enzymes found in
wild-type leaves during normal grth (Benning and Ohta, 2005, Xu et al. , 2003) (Figure
3.11). The retention of configuration from fiMGDG to fiflDGDG by SFR2 is consistent
with all other GH-l enzymes, which are classified as retaining fl-glycosidases with a
broad range of substrate specificities (Hill and Reilly, 2008).
Discussion
In this work, a longstanding unknown enzyme in chloroplast galactolipid
metabolism, the galactolipid:galactolipid galactosyltransferase (GGGT) has been
identified, and is encoded by the previously identified gene of unknown function, SFR2
(Thorlby et al., 2004). Clearly, GGGT is not essential for normal growth and
development, but rather for sub-zero temperature lipid remodeling of the chloroplast
95
Figure 3.10. Recombinant SFR2 in vitro activity from transgenic yeast. (A) Thin
layer chromatogram of lipid extracts stained for galactolipids with a-naphthol reagent
from in vitro GGGT activity assays using deoxycholate/MGDG dispersions incubated
with membrane fractions from empty vector control (EV) or SFR2 expression strain
(SFR2). The in vitro reactions were stopped after 1 h of incubation. Lipids from the
reactions were extracted, chromatographed and compared to lipids from a freeze-treated
leaf sample (FT Leaf). (B) In vitro GGGT activity using labeled [U-3H-Gal]-MGDG. The
data are presented as the percent of 3 H label found in DGDG, TGDG and TeDG over
time. Empty vector and SFR2 were assayed as in (A), and a SFR2 membrane protein
assay containing EDTA at a concentration equimolar to Mg2+ present (10 mM) is also
shown. Data are representative of two measurements, and the standard deviation is
shown if it exceeds symbol size. (C) Percent of [3H] label found in different galactolipid
species in the SFR2 assay shown in B. (D) Fatty acid composition of diacylglycerol in
empty vector control (EV) and SF R2 reactions at 3 b. Data are presented on 3 mol%
basis as the average and standard deviation of three biological repeats.
96
C100
% label in lipid species
% [3H]MGDG converted m
-o- MGDG D
*DGDG
*TGDG/TeDG
30 60 90 120150180
time (min)
97
80‘
60-
$ 0"
O O
a a
O
a
DAG fatty acids (mol%)
8 0)
_a
C
I
O
a
+SFR2
*SFRZ + EDTA
q—EV
30 60 90 120150180
time (min)
I EV
a SFR2
16:016:1 16:318201821 18:3
fatty acid species
Figure 3.11. IHNMR spectra of galactolipids from plant leaves and isolated from
the in vitro GGGT assay. MGDG (A) from the commercial supplier Laurodan, DGDG
(B) isolated from wild-type (C012) leaves, and (C) DGDG isolated from a scaled-up in
vitro GGGT reaction.
98
H1’
H1a 1b
11 1 11
LL __ ,‘~ 1 ‘
am“ 1’ __ :w~- J“ M -2 All” I &l
4] 4b ' 45 44 43 . 4h . 4H 40 3b ' as
41 1 4b 4b ' 4h 43 ' 42 ' 4H ' 4o . 33 . 33
ppm
99
envelope during freezing stress. Through consideration of (i) what is currently known
about plant cell responses to freezing, (ii) the work conducted by others on the sfr2
mutants in Arabidopsis prior to the functional elucidation of SFR2, and (iii) the work
presented above, a model for the role of GGGT in freezing tolerance can be envisioned.
A hypothesis for the role of GGGT in stabilizing chloroplast envelopes during
freeze-induced cellular dehydration. In order to understand how GGGT contributes to
freezing tolerance in plants, one must first consider the wealth of physiological,
biochemical, and molecular/ genetic insight that has been established in this intensely
studied field. Freezing damage is primarily visited upon the cell membranes and is
attributed to severe cellular dehydration, which is initiated by extracellular ice formation
that results in decreased water potential across the plasma membrane (Steponkus, 1984,
Thomashow, 1999). Under these conditions the formation of non-lamellar lipid phases,
such as the inverted hexagonal II (H11) phase can occur and, thus, stabilization of lamellar
membrane systems within the dehydrated plant cell is a key determining factor in the
survival of freezing (Steponkus, 1984, Thomashow, I999, Uemura et al., 1995). To
survive mild freezing, plants like Arabidopsis require pre-exposure to low, non-lethal
temperatures in a process known as cold acclimation (Hirayarna and Shinozaki, 2010,
Thomashow, 1999, Uemura et al., 1995).
Cold acclimation involves transcriptional (Guy et al., 1985, Kilian et al., 2007,
Zeller et al., 2009) and metabolic (Cook et al., 2004, Kaplan et al., 2004) changes which
result in multiple mechanisms protecting against freezing damage—including increases
in intracellular solutes and the accumulation of cryoprotective metabolites (Uemura et al.,
100
2003) and proteins (Steponkus et al., 1998). The phenotype of Arabidopsis plants
carrying mutations in the SENSITIVE T 0 FREEZING 2 (SF R2) gene, however, clearly
indicates a distinct mechanism required for freezing tolerance (Fourrier et al., 2008,
McKown et al., 1996, Thorlby et al., 2004, Warren et a]. , I996). The sfrZ mutants show
extensive intracellular damage following freezing recovery, with rupture of both
chloroplasts and tonoplasts—likely through fusion of destabilized membranes in these
organelles. The SFR2 protein is constitutively present, and sfr2 mutants are not affected
in the regulation of known cold-acclimation processes.
With cellular membrane damage in mind, alterations in lipid composition during
cold acclimation, such as increased fatty acid unsaturation and phospholipid content,
have long been correlated with enhanced freezing tolerance, but are not yet distinguished
from adaptations that arise from growth at low above-freezing temperatures rather than
freezing stress per se (Nishida and Murata, 1996, Steponkus, 1984, Thomashow, 1999).
More recent lipid profiling studies in Arabidopsis have revealed that more drastic lipid
compositional changes occur during freezing, including a decrease in the chloroplast-
specific lipid monogalactosyldiacylglycerol (MGDG) (Du et al., 2010, Li et al. , 2008,
Welti et al., 2002).
Our hypothesis for the requirement of SFR2-dependent galactolipid remodeling in
freezing tolerance centers on the prevention of non-lamellar Hn phase formation brought
about by dehydration (Figure 3.12). During dehydration, Hn phase is formed at the
interface of apposed bilayers, and believed to initiate at the chloroplast envelope during
freezing (Steponkus, 1984, Steponkus et al., 1998, Thomashow, 1999). This results in
fusion between bilayers (Siegel and Epand, 1997), particularly when membranes are
101
sfr2 SFR2
,. ,“t r. trtrt rutl'Hl
the CM
' l)--t..l rlrlll'l/lllllt I'll ..
r "
3DAG —> STAG
J
MGDG ‘4 teens
MGDG —> TGDG
ZMGDG —" DGDG
All I ;‘
)kéxl‘oI/I.
.. 1.1)-71”,“ I'llf mm 1
H "‘ll"-l'lll.t"l‘ troll/.74?! luVrrlirrllr-ali-YI‘ OEV
Figure 3.12. Model of SFR2 activity and the effect of freezing on the chloroplast
outer envelope membrane (eEv) and adjacent cell membranes (CM) in the sfi'z
mutant and the wild type (SFR2). Lipid bilayers and hexagonal 11 phase (113;)
formation leading to membrane fusions in sfr2 following freezing are indicated. SFR2 is
shown associated with the outer leaflet of the outer chloroplast envelope membrane.
Three cycles of the reaction are indicated consuming 4 moles of MGDG and producing 1
mole of TeGDG and 3 moles of TAG. This assumes that every mole of DAG produced is
converted to TAG which may not be the case as some of the DAG might be converted to
phosphatidic acid.
102
enriched in glycerolipid species with relatively small head groups (e. g. MGDG and
phosphatidylethanolamine) as these show a higher propensity for transition to H1] phase.
Previously, sfr2 mutants showed extensive chloroplast and tonoplast rupture in leaves
during freezing recovery, which was proposed to arise from fusion of destabilized
membranes (Fourrier et al., 2008). This work shows that SFR2 partially converts MGDG
to DGDG and oligogalactolipids (GGGT activity) which are not prone to form Hn phases.
This is analogous to the UDP-Glc-dependent modulation of mono- to di-glucolipid ratios
observed in Acheloplasma laidlawii under different abiotic stress conditions (W ieslander
er al., 1986). One distinction is that DAG is produced by SFR2, and is further
metabolized to TAG, and possibly other lipid species, as accumulation of DAG, which
can form non-lamellar phases, must be prevented. Moreover, the action of SFR2 provides
a mechanism by which membrane lipids and excess membrane are removed to
accormnodate a shrinking organelle following freezing. Under the conditions of fieeze-
induced dehydration, a response initiated by transcriptional activation may not be
practical for reasons including the time required for transcription, translation, protein
folding and lastly, localization, and also that these processes may be defective under this
stress. As such, the constitutive presence of inactive GGGT on the chloroplast envelope,
which could be directly activated by sensing physical changes in the membrane during
freezing, may offer a more successful strategy in mitigating membrane damage.
Evolution of glycosyl hydrolase family enzymes towards catalysis of
transglycosidation: a proposed mechanism for GGGT activity. Many GH-l family
enzymes are known to catalyze moderate levels of transglycosidase activity under certain
103
in vitro conditions, and examples for which transglycosylation predominates over
hydrolysis are known in other GH families (e. g. GH-70 glucansucrases) (Sinnott, 1990).
SFR2, which was found to be most divergent among 253 GH-l family proteins by
phylogenetic analysis (Marques et al., 2003), has evolved to catalyze MGDG
transgalactosidation (GGGT activity) by exclusion of water as an acceptor—which could
be attributed to several intrinsic features of SFR2—including the exclusory binding of
acceptor galactolipids, and/or the close apposition of the catalytic domain to the
membrane surface. As seen in figure 3.13, the proposed GGGT mechanism is a modified
version of the classical Koshland double displacement mechanism (Koshland, 1953),
where the enzyme-linked galactoside intermediate from donor MGDG is transferred to
MGDG (or DGDG/TGDG through processive activity) rather than water as an acceptor.
As such, SFR2 may provide insight into improving the transglycosidase activity of other
GH-l enzymes to produce oligosaccharides of medical or industrial importance (Perugino
et al., 2004).
Is the protective role of GGGT specific to freeze-induced dehydration? Orthologs of
SFR2 are found in all land plants with completed genomes (Fourrier et al., 2008).
Intriguingly, SFR2 is also conserved in species with no tolerance to freezing (e.g. tomato,
maize and rice) (Figure 3.14), suggesting SFR2 function is not restricted to freezing
protection (Fourrier er al., 2008). There is a large overlap in the mechanisms required for
freezing tolerance and dehydration due to water deficit or high salinity (Hirayama and
Shinozaki, 2010, Thomashow, 1999), and SFR2 orthologs in these species may act on
membrane stabilization during other abiotic stresses which cause cellular dehydration.
104
Figure 3.13. A proposed reaction mechanism for GGGT. The reaction is initiated
MGDG substrate binding, followed by nucleophilic attack by an acidic (Glu) residue
(step 1), which is assisted by another Glu residue acting as an acid catalyst in forming the
first transition state (step 2). Cleavage of the glycosidic bond results in the release of the
aglycon moiety (RIOH, diacylglycerol [DAG]), and the formation of a glycosyl enzyme
intermediate (step 3). The glycosyl donor substrate (R20H, MGDG) binds and is
deprotonated by the Glu residue, now acting as a base catalyst, followed by attack of the
enzyme-linked glycoside to form the second transition state (step 4). The enzyme linked
glycosidic bond is cleaved, resulting in the transglycosidation of bound MGDG to form
BBDGDG (step 5). The BBDGDG product is released, or' through processive activity, can
be further transgalactosylated (via steps 1-5) forming TGDG and subsequently TeDG.
The steps are delineated by numbered grey boxes.
105
0-05 r——-P. patens 1
100
P. patens 2
P. tn'chocarpa
100
100 R. comums
~ V. vinifera
64
S. chopersicum
G. max
2. mays
.l 68
O. sativa (lndica)
100
79
———- B. distach yon
A. thaliana
(SFR2)
P. teada
Figure 3.14. Phylogenetic relationships of SFR2 orthologs. Bootstraping values (1000
replicates) are shown at the nodes as percentages. The Genbank accession numbers for
the amino acid sequences are: Arabidopsis thaliana (SFR2), CAD36513; Brachypodium
distachyon, (ACF 22735); Glycine max, (CAJ 87636); Oryza sativa-Indica, (EAY81793);
Physcometrilla palens l, (XP_001759357); P. patens 2, (XP_001756957); Pinus teada,
(CAJ 87639); Populus trichocarpa, (XP_002316058); Ricinus communis,
(XPOOZS 19275); Solanum chopersicum, (CAJ87637); Vitis vinifera, (C8122845); Zea
mays, (NP_001105868).
107
Infiltration of Arabidopsis leaves with any osmotically active compound tested induced
GGGT activity (Figures 3.1 and 3.7). While the sensitivity of sfl2 mutants to severe
drought remains to be tested in detail, a number of physiological studies in crop species
(e.g., wheat and maize) and dessication tolerant resurrection plants (e. g., Ramonda
serbica), have found decreases in MGDGzDGDG ratios and the accumulation of TAG in
response to water deficit treatments (Navari-Izzo and Rascio, 1999). Whether SFR2
orthologs are participating in dehydration-induced lipid remodeling events in these
species is currently unknown. Regardless, it is emerging that modulation of membrane
lipid composition in response to cellular dehydration is a critical factor in the survival of
this stress. Future study of SFR2 may also provide general insight as to how organisms
outside the plant kingdom, including prokaryotes, yeasts, and some animals have evolved
tolerance to severe dehydration—and even desiccation—which was likely an important
factor in the adaptive transition of aquatic species to life on land.
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Chapter 4
Molecular genetics of lipid metabolism in the model green alga Chlarnydomonas
reinharda'r’
3 Parts of this work were published in: Moellering, E. R., Miller, R., and C. Benning.
"Molecular Genetics of Lipid Metabolism in the Model Green Alga C hlamydomonas
reinhardtii" Lipids in Photosynthesis: Essential and Regulatory Functions. H. Wada and
N. Murata, eds, Springer Netherlands-Dordrecht, 2009. Pp. 139-155.
114
Abstract
Lipid biosynthesis in algae has gained renewed attention in recent years, as algal lipids
are increasingly seen as promising biofuel and nutraceutical feedstocks. In contrast to the
molecular-genetic insight in seed plant glycerolipid metabolism, the genetic factors that
govern lipid biosynthesis in the diverse group of species collectively referred to as algae
are less well known. While the unicellular green alga Chlamydomonas reinhardtii is
unlikely to be developed for commercial production, it is currently the best genetic and
genomic model for microalgal lipid research. This chapter focuses on the current
knowledge of lipid metabolism in this alga, as well as the accumulation of neutral
glycerolipids (e.g., triacylglycerols [TAGs]) in different algal species during
environmental stress. Distinctions between seed plant and Chlarnydomonas lipid
metabolism are also discussed. For example, Chlarnydomonas lacks phosphatidylcholine
and has in its place the betaine lipid diacylglyceryl-N, N, N,-trimethylhomoserine. This
has important implications for lipid trafficking and lipid modification—including the
accumulation of TAGS, an increase of which is a major goal in improving algae as a
biofuel feedstock.
115
Abbreviations
ACP
CDP-DAG
DAG
DGDG
DGTS
ER
FAS
PC
PG
PE
Pl
PA
PS
PUFA
MGDG
RNAi
SQDG
TAG
acyl carrier protein
CDP-diacylglycerol
diacylglycerol
digalactosyldiacylglycerol
diacylglyceryl-N, N, N,-trimethyl homoserine
endoplasmic reticulum
fatty acid synthase
phosphatidylcholine
phosphatidylglycerol
phosphatidylethanolamine
phosphatidylinositol
phosphatidic acid
phosphatidylserine
polyunsaturated fatty acid
monogalactosyldiacylglycerol
RNA interference
sulfoquinovosyldiacylglycerol
triacylglycerol
116
Introduction
Glycerolipid biosynthesis in plants has been intensely studied for decades,
providing substantial molecular insight into plant lipid metabolism. Through a
combination of genetic and biochemical approaches, the genes encoding enzymes for
glycerolipid biosynthesis and fatty acid desaturation have largely been identified
(Benning and Ohta, 2005, Frentzen, 2004, Holzl and Dormann, 2007, Joyard et al., 1998,
Ohlrogge and Browse, 1995), and more recently, factors involved in endoplasmic
reticulum (ER)-to-plastid lipid trafficking have been discovered (Awai et al., 2006,
Benning, 2008, Lu et al., 2007, Xu et al., 2003, Xu et al., 2008). Furthermore,
sequencing of plant genomes, beginning with Arabidopsis (The Arabidopsis Genome
Initiative, 2001), followed by functional annotation of the encoded genes has allowed for
the identification of hundreds of candidate lipid metabolism and trafficking proteins that
await functional characterization (Beisson et al., 2003).
Chlamydomonas reinhardtii is a well established eukaryotic green microal gal
model organism, which has been utilized for the study of different processes including
photosynthesis (Niyogi, 1999), post-transcriptional gene silencing (Wu-Scharf et al.,
2000), cell biology of flagella (Silflow and Lefebvre, 2001), and microalgal metabolism
(Grossman et al., 2007). The availability of the complete Chlarnydomonas genome
sequence (Merchant et al., 2007), as well as a number of available molecular tools,
including facile nuclear insertional mutagenesis (Tam and Lefebvre, 1993), RNA
interference methods (Fuhrmann et al., 2001, Molnar et al., 2009), and a molecular map
(Kathir et al., 2003), make C hlamydomonas a promising model organism for the
discovery of genes involved in lipid metabolism and trafficking using both forward and
117
reverse genetics. Initial annotations of Chlamydomonas genes involved in lipid
metabolism have recently been published (Riekhof et al., 2005b, Riekhof and Benning,
2008), which highlight both the similarities and differences in land plant and green algal
glycerolipid metabolism.
With the resurgence of interest in using microalgae as a biomass feedstock for
renewable transportation fuels that does not have to compete with resources used in
agriculture (Hu et al., 2008), the availability of a suitable model microalgal organism is
timely. Even though Chlamydomonas reinhardtii is itself an unlikely candidate species
for production of biofuels, it is related to other unicellular algal species (e. g. Dunalliella
salina) that are used for commercial scale production of lipids (Hu et al., 2008).
Chlarnydomonas has been reported to accumulate triacylglycerols (TAGS) under
conditions of nutrient deprivation (Weers and Gulati, 1997) or high light (Picaud et al.,
1991), which has been reported in many other green algal species (Hu et al., 2008).
C hlamydomonas also synthesizes TAGS from lipids supplied in the medium (Grenier et
al., 1991). As such, insight into glycerolipid and TAG metabolism gleaned from studies
in Chlarnydomonas may be instructive in the rational engineering of other algal species
for biofuel production. Below, the current understanding of glycerolipid biosynthesis in
Chlarnydomonas and TAG metabolism in different algal species are discussed.
Fatty Acid Synthesis and Incorporation into Glycerolipids.
De novo synthesis of fatty acids is localized to the chloroplast of Chlamydomonas
cells (Sirevag and Levine, 1972), and the common ancestral origin of green algal and
seed plant plastids (Reyes-Prieto et al., 2007) is particularly apparent in many
homologous components of the fatty acid biosynthetic machinery. For example,
118
bioinformatic analysis of the Chlamydomonas genome has identified genes for the full
suite of enzymes required for the conversion of acetyl—CoA to acylated-acyl carrier
protein (ACP), including the multimeric bacterial-type acetyl-CoA carboxylase and fatty
acid synthase complexes (Riekhof et al., 2005b, Riekhof and Benning, 2008). These
enzymes are essential for fatty acid biosynthesis in plants (and presumably algae) which
predominantly produce 16:0-ACP and 18:1-ACP as the result of desaturation of 18:0-
ACP by a soluble stearoyl-ACP A9 desaturase (Browse and Somerville, 1991, Shanklin
and Somerville, 1991). As in plants, fatty acids are incorporated directly into chloroplast
membrane glycerolipids in Chlarnydomonas (Figure 1) by stepwise acylation of glycerol
3-phosphate to form phosphatidic acid (snI-18zl, sn2-16:0-PA) by glycerol 3-phosphate :
acyl-ACP acyltransferase (GPAT), which shows substrate specificity for 18:1-ACP, and
then by lysophosphatidate : acyl-ACP acyltransferase (LPAT, 16:0-ACP specific)
(Browse and Somerville, 1991, Kim and Huang, 2004, Kunst et al., 1988, Murata and
Tasaka, 1997, Xu et al., 2006). Fatty acids are also assembled into glycerolipids at the ER
where isofonns of the plastid acyltransferases are present and have been characterized in
Arabidopsis (Kim et al., 2005, Zheng et al., 2003). Putative orthologs of the plant GPAT
and LPAT genes are annotated in the final Chlarnydomonas genome draft (Riekhof et al.,
2005b, Riekhof and Benning, 2008). Candidates for the plastid PA phosphatase which
produces the diacylglycerol precursors for the biosynthesis of non-phosphorus lipids in
the plastid has been recently identified in Arabidopsis (Nakamura et al., 2007). However,
there is currently no good candidate in the Chlamydomonas genome predicted to encode
this enzyme (Riekhof et al. , 2005b, Riekhof and Benning, 2008).
119
Figure 4.1 Overview of glycerolipid biosynthesis in Chlarnydomonas. End-product
glycerolipids in the plastidic and endoplasmic reticulum (ER) pathways are shown in
bold. Abbreviations used are: ACP, acyl carrier protein; AdoMet, S-adenosylmethionine;
ASQD, 2'-O-acyl-sulfoquinovosyldiacylglycerol; CDP, cytidine-5'-diphosphate; CoA,
coenzyme A; CTP, cytidine-5'-triphosphate; DAG, diacylglycerol; DGDG,
digalactosyldiacylglycerol; DGTS, diacylglyceryl-N,N,N-trimethylhomoserine; Etn,
ethanolamine; FA, fatty acid; G-3-P, g1ycerol-3-phosphate; Glc, glucose; Ins-3-P,
inositol-3-phosphate; MGDG, monogalactosyldiacylglycerol; P-Etn,
phosphoethanolamine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PGP,
phosphatidylglycerolphosphate; PI, phosphatidylinositol; PA, phosphatidic acid; Ser,
serine; SQ, sulfoquinovose; SQDG, sulfoquinovosyldiacylglycerol; TAG, triacylglycerol;
UDP, uridine-5'-diphosphate. Key enzymatic steps (1-14, plastid; 15-23, BR) are
indicated by boxed numbers next to the arrows indicating substrates and products: 1,
acetyl-CoA carboxylase; 2, Malonyl-CoAzACP transacylase; 3, acyl-ACP thioesterase; 4,
G-3-P acyltransferase; 5, lyso-PA acyltransferase; 6, CDP-DAG synthase; 7, PGP
synthase; 8, PGP phosphatase; 9, PA phosphatase 10, sulfolipid synthase; ll, sulfolipid
2’-0-acyltransferase; 12, MGDG synthase; l3, DGDG synthase; l4, UDP-SQ synthase;
15, long chain acyl-CoA synthetase; l6, G-3-P acyltransferase; l7, lyso-PA
acyltransferase; 18, PA phosphatase; 19*; ER PG synthesis pathway (same as steps 6 to
8); 20, CDP-DAGzinositol phosphotransferase; 21, DAG acyltransferase; 22, betaine lipid
synthase; 23, CDP-EtnzDAG ethanolamine phosphotransferase.
120
~, / Plastid
Acetyl-CoA ASQD DGDG
IT El IT UDP-Gal
SQDG DAG—i MGDG
Malonyl-CoA UDP-Gal
UDP so n
1211”" 3032/. PA—C@—>CDP- DAG
MalonyI-ACP UDP-Glc El file-w
PAS Lyso-PA PGP
Acyl-ACP Ii
/ G- - PG
Free-FA
1 L r‘ifi
7 Cytoplasm/ER \T/
F FA
fee. G-3-P
Acyl-CoA/j-
TAG El Lyso-PA
Ser CDP-Etn
l DAG {-— pA
Etn—>P-Ern @lAdoMet P@/ Ext-P. -3-P
DGTS
121
Chloroplast Membrane Lipids. The overall structural organization of membranes in
the chloroplast of Chlamydomonas and seed plant chloroplasts is essentially identical,
where the inner and outer envelope membranes enclose an extensive thylakoid membrane
system in which the photosynthetic apparatus is embedded. Genetic studies of
Arabidopsis have identified many of the genes responsible for the biosynthesis of
chloroplast membrane lipids, and have revealed the essential role that lipid composition
plays in optimal photosynthetic function (Benning and Ohta, 2005, Dormann and
Benning, 2002, Vijayan et al., 1998, Wallis and Browse, 2002). Though the genetic study
of glycerolipid metabolism in C hlamydomonas has far fewer documented examples,
detailed biochemical analysis of this alga’s lipid composition has long confirmed the
presence of the major chloroplast membrane lipids found in land plants—including the
galactoglycerolipids mono- and di-galactosyldiacylglycerol (MGDG, DGDG),
sulfoquinovosyldiacylglycerol (SQDG), and the phosphoglycerolipid
phosphatidylglycerol (PG) (Giroud et al., 1988).
As in plants, galactoglycerolipids are the predominant membrane glycerolipid
class in Chlarnydomonas, where they make up a majority of the chloroplast membrane
lipids (Giroud et al., 1988, Janero and Barnett, 1981a). In Arabidopsis, the bulk of
galactolipid biosynthesis involves two enzymatic steps (Figure 1), whereby MGDG is
formed from diacyl glycerol (DAG) and UDP-galactose (UDP-Gal) substrates by MGDG
synthase (MGDl ), and DGDG is formed from MGDG and UDP-Gal by DGDG synthase
(DGDI) (Awai et al., 2001, Benning and Ohta, 2005, Dormann et al., 1999). Genes
encoding MGDG and DGDG synthases have been identified in the C hlamydomonas
genome as orthologs of the Arabidopsis genes MGD] and DGD], respectively (Riekhof
122
et al., 2005b, Riekhof and Benning, 2008). MGD] and DGD] are single-copy genes in
Chlamydomonas, which differs from that of the MGD], 2, 3 and DGD], 2 paralogs found
in the Arabidopsis genome. Molecular analysis of the Arabidopsis MGD2, 3 and DGDZ
genes has revealed their role in a galactolipid biosynthetic pathway which is
transcriptionally induced during phosphate deprivation, and is proposed to provide
galactolipids for extraplastidic membranes (Hartel et al., 2000, Jouhet et al., 2004, Kelly
et al. , 2003, Kelly and Dormann, 2002). The apparent lack of this induced galactolipid
pathway in Chlarnydomonas suggests a distinct lipid metabolic response to phosphate
limitation, or a lack of need for one. However, to date the galactolipid biosynthetic genes
of Chlamydomonas have not been studied in detail at the molecular level to test these
hypotheses.
The sulfolipid sulfoquinovosyldiacylglycerol (SQDG) has long been studied in
the context of its role in photosynthetic membranes, not only due to its prevalence in
photosynthetic eukaryotes and prokaryotes, but also because of its association with
photosynthetic pigment-protein complexes (Gounaris and Barber, 1985, Menke et al.,
1976, Pick et al., 1985, Stroebel et al., 2003). However, the more recent discovery of
SQDG and the genes and enzymes involved in SQDG biosynthesis in non-photosynthetic
bacteria and archea as summarized in (Benning, 2008, Cedergren and Hollingsworth,
1994), has clearly indicated that the role of sulfolipids is not limited to the function of
photosynthetic membranes. The biosynthesis of SQDG is carried out in two enzymatic
steps in Arabidopsis by (i) SQD] , which catalyzes the formation of UDP-sulfoquinovose
from UDP-Glc and sulfite, and (ii) SQD2, which transfers the sulfoquinovose moiety
from UDP-sulfoquinovose to DAG, forming SQDG (Essigmann et al., 1998, Sanda et al.,
123
2001, Yu et al., 2002). A single copy ortholog of SQDI is present in Chlamydomonas,
and two possible orthologs of Arabidopsis SQDZ (SQD2, 3) are found in the genome
(Riekhof et al., 2003, Yu et al., 2002). Recently, a Chlarnydomonas mutant deleted in
SQDI (Asqdl) and completely lacking sulfolipid has been studied (Riekhof et al., 2003).
Phenotypic analysis of Asqdl revealed a reduced growth rate during phosphate-limiting
conditions—under which the SQDG level was found to double in wild-type cells. This is
similar to what has been observed in sulfolipid-deficient mutants in other organisms such
as Arabidopsis, which showed impaired growth after severe phosphate limitation (Y u et
al., 2002), and in the photosynthetic purple bacterium Rhodobacter sphaeroides (Benning
et al., 1993). In addition, Asqdl showed sensitivity to a photosystem II inhibitor under
normal growth conditions (Riekhof et al., 2003). This is consistent with another
Chlamydomonas SQDG-deficient mutant, hf-Z, which was first discovered as a high
chlorophyll fluorescence mutant, and was later found to be impaired in photosystem II
stability and showed increased sensitivity to a PSI] inhibitor—which could be partially
restored by SQDG addition (Minoda et al., 2002, Minoda et al., 2003, Sato et al., 19953,
Sato et al., 1995b). However, whether the hf-2 mutant is impaired in grth during
phosphate limitation has not been reported, nor has the exact molecular defect in this
mutant been determined. Interestingly, detailed biochemical analysis of the Asqdl
mutant also lead to the discovery of the novel sulfolipid derivative, 2’-O-acyl-
sulfoquinovosyldiacylglycerol (ASQD), which was also not produced in Asqdl (Riekhof
et al., 2003). Due to the loss of both sulfolipids in Asqu, the specific roles played by
SQDG and ASQD in Chlarnydomonas and phenotypes associated with Asqdl can only be
124
fully interpreted after the identification and characterization of the acyltransferase
catalyzing ASQD production has been undertaken.
Phosphatidyl glycerol (PG) is presumably the only major phospholipid component
of seed plant thylakoid membranes, and biochemical analysis of thylakoid lipid
composition has confirmed this to be the case in Chlarnydomonas (Janero and Bannett,
I981b, Mendiola-Morgenthaler et al., 1985). While the gene encoding the final enzyme
in PG biosynthesis, phosphatidylglycerolphosphate (PGP) phosphatase, remains
unknown in plants and algae (Beisson et al., 2003), the putative genes encoding the
enzymes which catalyze the formation of the two intermediates, CDP-DAG synthetase
and phosphatidylglycerolphosphate synthase, have been identified (Riekhof et al., 2005b,
Riekhof and Benning, 2008), but not yet confirmed. While neither the single gene
encoding the CDP-DAG synthetase (CDSI) or the two putative plastid paralogs encoding
phosphatidylglycerolphosphate synthase (PGPI/Z) have been studied at the
molecular/genetic level, PG deficient mutants, mf I and mf 2, have been isolated and
studied in great biochemical detail (Dubertret et al., 1994, Dubertret et al., 2002, Gamier
et al., 1987, Gamier et al., 1990, Maanni et al., 1998, Maroc et al., 1987, Pineau et al.,
2004). The mf 1, 2 mutants were first isolated as low fluorescent strains lacking
fimctional PSII as well as an oligomeric form of the light-harvesting chlorophyll antenna
(CPII) (Dubertret et al., 1994, Maroc et al., 1987). It was also shown that both mfI and
mf 2 contained ~30 % of wild-type PG levels and lacked A3-trans-hexadecenoic acid
(16: 11‘3”” [carbons : double bonds Apos"'i°"""’]) (Dubertret et al., 1994, Maroc et al., 1987), a
fatty acid that is specifically esterified to chloroplastic PG in both Arabidopsis and
Chlarnydomonas (Browse et al., 1985, Gamier et al. , 1987, Giroud et al., 1988). Addition
125
A3trans
of a preparation of spinach leaf PG containing 16:1 to mf-2 cells restored the ability
to form oligomeric CP 11, while 18:0 PG additions did not, and 16:0 PG did only weakly
so (Dubertret et al., 1994, Gamier et al., 1990). A PG-deficient mutant in Arabidopsis,
pgpl , which is defective in the chloroplastid isoform of PGP synthase has been found to
be photosynthetically impaired with decreased quantum yield through PSII, but did not
lack 16:1A3'm’" PG (Babiychuk et al., 2003, Hagio et al., 2002, Xu et al., 2002). Similarly,
two Synechocystis PG deficient mutants showed altered PSII activity and required
exogenous PG addition for phototropic growth (Hagio et al., 2000, Sato et al., 2000).
Taken together with the contrasting findings from analyses of the Arabidopsisfad4
mutant, which lacks 16:1A3’m'” fatty acid, but is otherwise not affected in chloroplast PG
content and also shows no apparent photosynthetic defects, it can currently only be
concluded that in general PG plays an important role in photosynthetic membrane
biogenesis and function, and it seems possible that the 16:1A3'm'” PG form could be
essential in some organisms (e.g. Chlarnydomonas), but is of conditional importance or
dispensable in others. It is certain however, that the elucidation of the exact molecular
defects in the Chlamydomonas mfl, 2 mutants, and the identification of the genes
encoding FAD4 activity as well as the elusive plant/algal PGP phosphatase which
catalyzes the final step in PG biosynthesis, will be prerequisite to gaining a better
understanding of the roles PG plays in the photosynthetic membranes in various species.
Toward this end, the gene affected in the Arabidopsisfad4 16:1A'm’” deficient mutant was .
cloned (Gao et al., 2009), and appears to belong to a distinct class of desaturases, as
described below.
126
Extrachloroplastic membrane lipid metabolism. In eukaryotes the bulk of
extraplastidic membrane glycerolipids are assembled in the ER from acyl-CoA thioesters
which are formed from free fatty acids after their liberation from acyl-ACPS in the plastid
(see Figure 4.1). While other extraplastidic sites for lipid synthesis are known (e. g.
mitochondria), the ER-localized pathway is predominant, and in most plants the ER lipid
assembly pathway significantly contributes to thylakoid membrane biogenesis. As such, a
discussion of the analogous pathways in Chlarnydomonas is merited. As mentioned
above, C hlamydomonas lacks the capability for PC biosynthesis (Giroud et al., 1988) and
genes predicted to encode enzymes involved in PC biosynthesis are not present in its
genome (Riekhof et al., 2005b, Riekhof and Benning, 2008). Instead, it contains the non-
phosphorous zwitterionic betaine lipid DGTS in its membranes (Eichenberger and
Boschetti, 1977, Janero and Barrnett, 1982) which has similar structural (see Figure 2),
and hence biophysical properties to PC (Sato and Murata, I991). DGTS has also been
found in other algal species, (Eichenberger, 1982), prokaryotes like the purple bacterium
Rhodobacter sphaeroides, (Benning et al., 1995, Hofrnann and Eichenberger, 1996), and
in non-seed plants such as ferns, (Eichenberger, 1993, Sato and Furuya, 1983), but
appears to be absent in seed plants. Labeling studies suggest that the biosynthesis of
DGTS is similar in all organisms studied (Hofmann and Eichenberger, 1996, Sato, 1988,
Sato and Kato, 1988, Vogel and Eichenberger, 1992). It begins with the transfer of the 3-
amino 3-carboxypropyl residue from S-adenosylrnethionine (AdoMet) to DAG catalyzed
by AdoMetzDAG 3-amino-3-carboxypropyltransferase activity followed by successive
methylation of the amino group by an AdoMet-dependent N-methyltransferase . The two
genes encoding these catalytic
127
O
\ O\P/ i
+ \
/ \\/\O/ O/\l/\O R1
0 R2
PC \11/
O
000' o
\+NJ\/\O/\‘/\O/lkR1
0 R2
DGTS T
0
Figure 4.2. Structures of PC and DGTS. A comparison of DGTS and PC structure,
where sn-I and sn-2 acyl chains are represented by R. and R2, respectively.
128
activities, BtaA and BtaB, were first identified in R. sphaeroides (Klug and Bennin g,
2001, Riekhof et al., 2005a). More recently, a single gene sufficient for DGTS
biosynthesis in Chlamydomonas, Btal, was identified in the genome (Riekhof et al.,
2005b). The encoded protein Btal is similar in its N-terminal domain to bacterial BtaB
and in its C-terminal domain to BtaA and the predicted catalytic function of each Btal
domain was confirmed by mutagenesis (Riekhof et al., 2005b). The bifunctionality
observed in Btal as a fusion of two prokaryotic enzyme activities functional in the same
pathway into a single polypeptide is a common theme in plants (Moore, 2004), and could
represent an improvement in DGTS biosynthesis by eliminating the need for coordinated
regulation of two independent gene products or by permitting substrate channeling. In
addition, the presence of DGTS and concomitant lack of PC is perhaps related to the
absence of additional galactolipid biosynthetic pathways seen in seed plants, e.g., MGD2,
3 and DGD2 in Arabidopsis (Benning and Ohta, 2005). As this alternative galactolipid
pathway is believed to be involved in replacing phospholipids in extraplastidic
membranes with DGDG during phosphate deprivation, the constitutive replacement of
PC with the non-phosphorous DGTS has perhaps obviated the need to replace PC by
extraplastidic DGDG. Interestingly, DGTS was tentatively identified as a component of
purified Chlarnydomonas thylakoids (Janero and Barrnett, 1981b, J anero and Barrnett,
1982) and chloroplast envelope membranes (Mendiola-Morgenthaler et al., 1985).
However, whether DGTS is indeed present in the chloroplast membranes of
Chlamydomonas or plays a specific function in thylakoids remains to be confirmed.
Phosphatidylserine (PS) is a minor component of extraplastidic membranes of
plants and can be decarboxylated to phosphatidylethanolamine (PE) in plants and other
129
organisms (Nerlich et al., 2007, Vance, 1990). However, PS is absent in
C hlamydomonas membranes (Giroud et al., 1988) and genes encoding the
phosphatidylserine synthase or relevant phospholipid base exchange enzymes were not
detected in the genome (Riekhof et al., 2005b, Riekhof and Benning, 2008). AS such, the
biosynthesis of PE, which is a known component of extraplastidic membranes in
C hlamydomonas, is likely only carried out by a single pathway (Figure 4.1). Genes
encoding a serine decarboxylase which produces ethanolamine, an ethanolamine kinase
and CTP : phosphoethanolamine cytidylyltransferase—the combined activities of which
produce CDP-ethanolamine, and lastly, a CDP-ethanolamine : DAG ethanolamine
phosphotransferase which produces PE have been identified in the genome (Riekhof et
al., 2005b, Riekhof and Bennin g, 2008). Recently, the gene encoding the CTP :
phosphoethanolamine cytidylyltransferase has been characterized by heterologous
expression in Escherichia coli, and the expression of the respective gene was found to be
up-regulated during the reflagellation of Chlamydomonas cells (Yang et al., 2004).
Phosphatidylinositol (PI) is a minor component of Chlamydomonas membranes, and
genes required for its biosynthesis are present in the genome (Riekhof et al., 2005b,
Riekhof and Benning, 2008). The PI biosynthetic enzymes of Chlamydomonas have been
studied at the biochemical level, and PI synthesis was found to be highest in the
microsomal fraction, suggesting its association with the ER (Blouin et al., 2003).
Extraplastidic PG biosynthesis is known to be associated with both the ER and
mitochondria, and three isoforms of phosphatidylglycerolphosphate synthase are encoded
in the genome, each with differential targeting prediction probabilities for subcellular
localization to the mitochondria, chloroplast or cytosol (Riekhof et al., 2005b, Riekhof
130
and Benning, 2008). However, a detailed analysis of these proteins and their respective
genes is not yet available in C hlamydomonas.
Fatty Acid Desaturation. Biochemical studies in Chlamydomonas have indicated that
further desaturation of 16:0 and 18:1 acyl groups occurs after the production of the major
glycerolipids, in a manner similar to plants (Giroud et al., 1988). Furthermore, the
Chlarnydomonas fatty acid profile is known to change markedly in response to different
environmental conditions including C02 concentration as well as nitrogen and
phosphorous limitation (Tsuzuki et al., 1990, Weers and Gulati, 1997). The elucidation
of fatty acid desaturase (FAD) genes in Arabidopsis is a classic example of the power of
genetic and molecular biological approaches in solving biological problems which prove
to be largely intractable through a strictly biochemical approach (Browse et al. , 1985,
Wallis and Browse, 2002). The identification of many of the Chlamydomonas desaturase
gene candidates by their similarity to Arabidopsis orthologs, combined with a handful of
studies of C hlamydomonas desaturase mutants and characterization of cloned FAD genes,
has provided a reasonable picture of the fatty acid desaturation pathways in this alga
(Figure 4.3).
Putative orthologs for the plastidic desaturases encoded by Arabidopsis FAD5
(MGDG palmitate-A7-desaturase), FAD6 (m6-desaturase), and FAD? or FAD8 (encoding
m3-desaturase isozymes) are present in the C hlamydomonas genome (Riekhof et al. ,
2005b, Riekhof and Benning, 2008). Of these, only the Chlarnydomonas FAD6 gene has
been studied to date at the molecular/genetic level (Sato et al., 1995b, Sato et al., 1997).
131
Figure 4.3. Overview of acyl-chain desaturation in Chlarnydomonas. Glycerolipid
abbreviations are the same as those in Figure 4.]. Fatty acids are referred to by the
standard abbreviation “carbon atomszdouble bonds.” Specifically, common names of
these fatty acids include 16:0, palmitic acid; 18:0, stearic acid; 18:1, oleic acid; 16:1,
palmitoleic acid; 16:1”, A3-trans hexadecenoic acid; 18:2, linoleic acid; (118:3, linolenic
acid (18:3A9'12’”); i18:3, pinolenic acid (18:3A5‘9’”); 18:4, coniferonic acid. The
desaturases that catalyze the enzymatic steps in the plastid (24-29) and the ER (30-32) are
indicated by boxed letters: 24, stearoyl-ACP desatruase (des.); 25, PG-specific-16:O-
A3trans des.; 26, MGDG-specific-l6:0-A7 des.; 27, plastidic (0'6 des.; 28, plastidic (0-3
des.; 29, MGDG-16C-A4 des.; ER 03-6 des.; 31, A5 des.; 32, ER (0-3 des.
132
l
I
l
i
l
Lipid
Bros nthesls
PG SQDG MGDG DGDG
(16:0)18:1/16:O 18:1/16:0 18:1/16:0 18:1/16:0
ranger“ 1:33am 133237311115;7| refiso
18:2/16:1” 018:3/1620 18:2/16:2 0181311610
018:3/16z1A3‘ mas/16:3
lk Plastid 018:3116z4 J l
\\‘_ _ -_ _ .-_..-"/
ER
DGTS PE
16:0/18:1 18:0/18:1
16:0/18:2 182011822
fl 18:0-ACPE—b1821-ACP
momma
\a
16:0/i18:3 16:0/018:3
.VQ
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133
\IE
18:0/118:3 18:0/a
IE
18:0/18:4
18:3
El
The hf-9 mutant was first isolated by its high chlorophyll fluorescence phenotype, and
detailed lipid analysis revealed an apparent defect in m6-desaturase activity as it showed
marked decreases in both 16- and l8-carbon polyunsaturated fatty acyl groups, with
concomitant increases in 16:1A7 and 18:1 A9 (Sato et al., 1995b). The hf-9 mutant had an
increased doubling time and showed reduced photosynthetic 02 evolution as well as an
altered chloroplast ultrastructure (Sato et al., 1995b). The C hlamydomonas FAD6 gene
(first described as DES6) was subsequently cloned and found to be highly similar to
cyanobacterial A12- and seed plant (rm-desaturases, and was also Shown to complement
the hf-9 mutant desaturation defects (Sato er al., 1997). However, it did not restore the
photosynthetic defects, suggesting that these phenotypes arose from a mutation in another
gene (or possibly from multiple loci). As such, the roles of polyunsaturated fatty acids
(PUFAS) in the assemblage or maintenance of optimally functioning photosynthetic
membranes in Chlarnydomonas cannot be easily deduced from analysis of hf-9. The
function of PUFAS in this regard have been studied and found to differ in mutants of both
plants and cyanobacteria. The Arabidopsisfad6fad2 double mutant, which lacks both
plastidic and ER (rm-desaturases exhibited severe grth and photosynthetic defects
(McConn and Browse, 1998). In contrast, a mutant of Synechocystis Sp. PC6803 lacking
PUFAS had no observable photosynthetic defects under normal growth conditions
(Gombos et al., 1992). Thus, it still remains to be determined whether the importance of
PUFAS in photosynthetic membrane function in C hlamydomonas is more similar to that
of seed plants or cyanobacteria. As noted above, a candidate desaturase producing
16:1A3’m'" specifically on plastidic PG, was identified (Gao et a1. , 2009) by map-based
cloning of the genetic lesion in the Arabidopsisfad4 mutant (Browse et al., 1985). The
134
FAD4 protein is distinct from all other known acyl-chain desaturases with respect to
position and number of proposed catalytic residues, which may be related to the fact that
this enzyme produces a trans rather than cis desaturation product. A candidate FAD4
ortholog has been identified in the most recent draft of the Chlarnydomonas genome (v
4.0, http://genome.jgi-psf.org/chlamy/chlamyhome.html), but has not yet been
characterized (S. Zaiiner and C. Benning, pers. comm.) However, the gene list of
chloroplast desaturases in C hlamydomonas remains incomplete, as a gene encoding A4
desaturase activity, which is specific for MGDG-esterified l6-carbon acyl groups based
on biochemical studies (Giroud et al., 1988), and presumably produces both the
16:3’34’7‘lo and 16:4‘54’7"°'l3 in Chlarnydomonas is currently unidentified.
The extraplastidic 036- and co3-desaturases which produce 182“”12 and 18:3
”'2’” are encoded by FAD2 and FAD3, respectively in Arabidopsis, and putative
orthologs of these genes have been identified in the Chlamydomonas genome (Riekhof et
al., 2005b, Riekhof and Benning, 2008). The extraplastidic Chlarnydomonas lipids
A5,9,12
3
DGTS and PE have been shown to contain significant amounts of 18: and
1824559”15 esterified to the respective sn-2 positions of the glycerol back bone (Giroud
et al. , 1988); these A5-unsaturated fatty acids are also found in gymnosperms (Mongrand
et al., 2001, Wolff and Christie, 2002). Recently a Chlarnydomonas gene, CrDES,
encoding a “fi'ont-end” type AS-desaturase was identified by a bioinformatics approach
through its similarity to a known A5-desaturase from the liverwort Marchantia
polymorpha (Kajikawa et al., 2006). Heterologous expression of CrDES in Pichia
pastoris and analysis of desaturase activity indicated while the primary substrates were
18:21‘9’12 and 18:3 ”’12”, low but detectable levels of endogenous 18:1” desaturation
135
were also observed (Kajikawa et al., 2006). Transgenic tobacco plants constitutively
expressing the C rDES gene exhibited strikingly high levels of 18:3’35‘9‘12 and 18:4’35‘9‘n‘l5
(which are normally absent), with the highest combined yield reaching ~45% of leaf total
fatty acids, and no apparent morphological phenotypes (Kajikawa et al., 2006). The
biological roles of these A5-unsaturated fatty acids in the organisms which produce them
are largely unknown, and the identification of the CrDES gene responsible for 1823“"'2
and 18:4’35‘9’12’ls production in Chlarnydomonas will allow for this gene to be targeted for
suppression through RN Ai technology.
Differences in ER-to-chloroplast lipid trafficking in plants and Chlarnydomonas
The diacylglyceryl backbone moieties that feed into thylakoid lipid biosynthesis
in many plants are derived from two pathways (Mongrand et al., 1998), termed the
plastid and the ER pathways. This hypothesis was first formulated by Roughan and
coworkers based on labeling experiments (Roughan et al. , 1980, Roughan and Slack,
1982) and later confirmed by analysis of Arabidopsis mutants (Browse and Somerville,
1991, Wallis and Browse, 2002). The relative contribution of each pathway is evident in
the fatty acid composition of chloroplast-specific lipids (e. g. MGDG), where the ratio of
16:3 (plastidic pathway) to 18:3 (ER pathway) at the sn-2 position is diagnostic (Heinz
and Roughan, 1983), and fluxes through the two pathways have been determined
(Browse et al., 1986). Many plant Species have lost the plastidic pathway for providing
DAG moieties for biosynthesis of the dominant galactoglycerolipids, but nearly all plant
species reported derive at least a fraction of their thylakoid lipids from precursors
assembled at the ER (Mongrand et al., 1998) requiring lipid precursor import into the
plastid. Recent studies have identified genes encoding a putative ABC-type transporter
136
complex localized to the chloroplast inner envelope, which is required for ER-to-plastid
lipid trafficking in Arabidopsis, by analysis of mutants in the TGD1,2,3 genes (Awai et
al., 2006, Lu etal., 2007, Xu et al., 2003). In contrast, detailed compositional analysis of
lipids and labeling studies suggest that in Chlamydomonas all thylakoid lipids are
assembled de novo in the plastid (Giroud et al., 1988).
Intriguingly, orthologs of both the Arabidopsis TGDl and 2 proteins are encoded
in the Chlarnydomonas genome, suggesting they be essential for lipid trafficking within
the plastid in the absence of ER glycerolipid import, or alternatively, they are not
functioning in lipid transport in Chlarnydomonas. This could also be related to the lack of
PC biosynthesis in Chlamydomonas, and its (presumed) functional replacement with
DGTS (Eichenberger and Boschetti, 1977). PC is central to lipid metabolism in
developing seeds or leaves where it serves a substrate for fatty acid modifying enzymes
such as desaturases (Browse and Somerville, 1991, Ohlrogge and Browse, 1995, Wallis
and Browse, 2002) or possibly as the lipid transferred between the ER and the plastid
(Benning, 2008, Jouhet et al., 2007). Thus, it is possible that the lack of PC and the lack
of trafficking of lipid precursors from the ER to the plastid in Chlarnydomonas are related
if PC is a critical intermediate in ER-to-plastid lipid trafficking. Because in the betaine
lipid, DGTS, the head group moiety is ether-linked to the diacylglyceryl moiety,
Chlarnydomonas might lack an enzyme to break this ether linkage. This ether linkage is
more stable than the phosphate ester linkage in phosphoglycerolipids. Therefore, the
conversion of DGTS into the galactoglycerolipid precursor diacylglycerol might not be
possible in C hlamydomonas.
137
Neutral glycerolipid metabolism. Many algae accumulate significant quantities of
neutral lipids (predominately TAG) under certain conditions, and thus have long been
considered an attractive source for bio-based fuels. While there has been a resurgence in
interest in algal biofuel production (Hu et al., 2008), this concept was intensely studied
under the two decade spanning Dept. of Energy funded Aquatic Species Program (ASP)
that initiated in the late 1970’s (Sheehan et al., 1998). Through screening thousands of
algal strains, many species were identified that could accumulate TAG up to 75 % dry
weight (Ben-Amotz et al., 1989, Bigogno et al., 2002, Chisti, 2007).
In most algae species, TAG accumulates in response to environmental stress,
suggesting that triacylglycerol plays a role in microalgae beyond energy storage (Chisti,
2007, Hu et al., 2008, Shanklin and Somerville, 1991). Most studies focused on nutrient
deprivation, with an emphasis on nitrogen limitation, where it was found that the
chlorophytes Neochloris oleoabundans (Tomabene et al., 1983) and Parietochloris
incise (Merzlyak et al., 2007) accumulate TAGS. The eustigmatophyte Nannochloropsis
gave similar results (Suen et al., 1987), suggesting that this response to N-limitation may
be a common phenomenon in algae—given the large evolutionary divergence between
the Chlorophycae and Bustigrnatophycae (Reyes-Prieto et al., 2007). Other nutrient
deficiencies are also known to cause TAG accumulation. For example, silicon limitation
results in increased TAG content in the diatom Cyclotella cryptica (Roessler, 1988).
Light intensity and temperature are also known to affect the accumulation of TAG
in many algal Species. Changes in temperature result in species specific-alterations in
lipid accumulation, with some algae showing increased TAG content, and others showing
decreases (de Swaaf et al., 1999, Dempster and Sommerfeld, 1998, Richardson et al.,
138
1983). Growth of many algal species under high light intensity results in increased TAG
to total lipid ratios (Khotimchenko and Yakovleva, 2004, Khotimchenko and Yakovleva,
2005, Zhekisheva et al., 2002). The chlorophyte Dunaliella bardawil accumulates TAG
under high light Stress along with B-carotene, which was proposed to protect chloroplasts
from photo-oxidative damage (Ben-Amotz et al. , 1989, Rabbani et al., 1998). While
Chlamydomonas reinhardtii was largely considered a poor lipid accumulator, this species
can accumulate significant amounts of TAG during nitrogen or phosphorous deprivation
(W eers and Gulati, 1997). Despite these efforts, little is currently known about algal TAG
biosynthesis at the molecular level, including that in the genetic model alga
Chlarnydomonas. Given the above-described Similarities between Chlamydomonas and
plant membrane glycerolipid metabolism, it seems likely that many general aspects of
TAG metabolism are also shared.
A major pathway for TAG biosynthesis in many organisms, including plants,
involves the step-wise acylation of a glycerol backbone (i.e. the Kennedy pathway; see
Figure 1) (Kennedy, 1961). However, several studies in many different plant species
have also indicated that DAG derived from the PC pool also contributes substantially to
TAG biosynthesis (Bates et al., 2009, Ohlrogge and Browse, 1995). Clearly, PC plays no
role in TAG biosynthesis in C hlamydomonas, and its role in plants is not universal, as
studies of avocado mesocarp microsomes indicated that only the Kennedy pathway was
active (Stobart and Stymne, 1985). It may be possible that the assumed functional analog
of PC in Chlarnydomonas, DGTS, is an intermediate in the biosynthesis of TAG.
However, no biochemical studies to determine this have been undertaken to date, and
utilization of the DGTS pool to provide DAG precursors would require an as yet
139
unidentified enzyme to remove the ether linked trimethylhomoserine head group.
Regardless of the precursors used in forming DAG, the final step is catalyzed by
diacylglycerol acyltransferases, or DGATS, which transfer a fatty acid from acyl-CoA to
diacylglycerol. DGATs have been isolated and characterized from several plant species,
including Arabidopsis (Hobbs et al., 1999, Routaboul et al., 1999, Zou et al., 1999),
maize (Zheng et al., 2008) and castor beans (Kroon et al., 2006). The Chlarnydomonas
genome contains a number of putative DGAT isoforms, which all appear to be of the type
11 class of DGAT enzymes (Lardizabal et al., 2001), and remain to be studied in
molecular detail (Riekhof and Benning, 2008; Miller and Benning, unpublished).
Chlarnydomonas may also use an alternate pathway for TAG synthesis involving
the phospholipid : diacyl glycerol acyltransferases, or PDATS, to generate TAG using a
phospholipid as a fatty acid donor, rather than acyl-CoA (Dahlqvist et al., 2000). PDATs
have also been found in plants (Stahl et al., 2004), and a putative ortholog was also found
in Chlamydomonas (Riekhof and Benning, 2008). Given the induction of TAG
biosynthesis by different stresses, it is likely that the mechanism for the regulation of
TAG synthesis differs in Chlamydomonas fiom that in seed plants, which often produce
oil during a specific phase of their life-cycle and in specialized tissues. Intriguingly, a
recent study of chloroplast lipid remodeling during N-deprivation in Arabidopsis revealed
the accumulation of TAG and phytyl esters in the chloroplast (Gaude et al. , 2007),
suggesting that TAG accumulation may be a common response in photosynthetic
organisms to carbon-nitrogen imbalances brought about by N-limitation.
140
Perspectives
Interest in microalgal lipid metabolism is experiencing a renaissance due to the increasing
need for replacing fossil fuels with bio-based renewable resources. This, coupled with
the ever-decreasing cost of DNA sequencing technologies, will undoubtedly result in the
sequencing of a number of algal genomes. Thus, comparative genomics may become a
powerful tool in elucidating factors required for TAG biosynthesis in algae. Moreover,
such studies in other algal Species may reveal distinct mechanisms of lipid metabolism
that have evolved separately in this highly divergent group of organisms. The current
availability of the C hlamydomonas genomic sequence (Merchant et al., 2007), and its
corresponding molecular toolkit has indeed made the application of knowledge on well
studied lipid metabolism in seed plants to this model alga relatively facile using
comparative genomics. As such, the potential for applying insight gained from studying
lipid metabolism in Chlarnydomonas in gene discovery and the rational engineering of
other algal species to produce biofuels is promising.
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Chapter 5
Analysis of triacylglycerol accumulation in the green alga Chlamydomonas
reinhardtii during nitrogen deprivation: Identification and characterization of an
algal specific lipid droplet protein MLDPl
' This work was published in: Moellering, E. R. and C. Benning. (2010) RNA
interference Silencing of a major lipid droplet protein affects lipid droplet size in
C hlamydomonas reinhardtii. Eukaryot. Cell 9:97-106.
154
Abstract
Eukaryotic cells store oils in the chemical form of triacylglycerols in distinct organelles,
ofien called lipid droplets. These dynamic storage compartments have been intensely
studied in the context of human health and also in plants as a source of vegetable oils for
human consumption and for chemical or biofuel feedstocks. Many microalgae
accumulate oils, particularly under conditions limiting to growth and, thus, have gained
renewed attention as a potentially sustainable feedstock for biofuel production. However,
little is currently known at the cellular or molecular levels with regard to oil accumulation
in microalgae, and the structural proteins and enzymes involved in the biogenesis,
maintenance, and degradation of algal oil storage compartments are not well studied.
Focusing on the model green alga Chlamydomonas reinhardtii, the accumulation of
triacylglycerols and the formation of lipid dr0plets during nitrogen deprivation are
investigated Mass spectrometry identified 259 proteins in a lipid droplet enriched
fraction, among them a major protein, tentatively designated Major Lipid Droplet Protein
(MLDP). This protein is specific to the green algal lineage of photosynthetic organisms.
Repression of MLDP gene expression using an RNAi approach led to increased lipid
droplet size, but no change in triacylglycerol content or metabolism was observed.
155
Introduction
Triacylglycerols (TAGS) are stored in lipid droplets which are subcellular structures in
specialized cells ubiquitous to eukaryotes, but have more recently also been identified in
some prokaryotes (Murphy, 2001). In plants and animals, lipid droplets are surrounded
by cytosol and are believed to bud off the endoplasmic reticulum (Huang, 1992, Murphy,
2001). While traditionally considered merely as storage compartments, recent studies
suggest that lipid droplets in animals play important additional roles in lipid homeostasis,
and protein storage (Cermelli et al., 2006). In oilseed plants, TAG accumulated in seeds
is used as a reservoir of energy and membrane lipid building blocks to support rapid
growth after germination (Huang, 1992). Many green algae are capable of accumulating
large amounts of TAG in lipid droplets, particularly as a result of abiotic stresses such as
nutrient deprivation or high-1i ght exposure. Although TAG metabolism in algae has not
yet been extensively studied at the biochemical or molecular level, it is proposed that
TAG turnover contributes primarily to the assembly of membrane lipids to facilitate rapid
cell division after the cessation of nutrient limitation (Hu et al., 2008, Thompson, Jr.,
1996).
The general structure of lipid droplets is conserved in different species with a
globular neutral lipid core enclosed by a membrane lipid monolayer (Murphy, 2001). In
addition, Specific proteins are associated with lipid droplets and play important roles in
lipid droplet structure and function. A number of recent proteomic studies of lipid
droplets from different animals and tissues (Cermelli et al., 2006, Walther and Farese, Jr.,
2009), yeast (Athenstaedt et al., 1999), and from plants (Jolivet et al., 2004, Katavic et
al., 2006) have revealed that the lipid droplet-associated proteins of these organisms are
156
quite distinct. For example, the abundant lipid droplet proteins in animals—the so called
“PAT” family of proteins comprised of perilipin, adipose differentiation-related protein
(ADRP) and T1P47 (Londos et al., 2005), have no apparent orthologs in the desiccating
seed plant Arabidopsis thaliana; conversely, the oleosins which coat the oil bodies of
Arabidopsis and many other seed plants are not found in animals (Murphy, 2001).
Reverse genetic Studies of these proteins have helped to elucidate the role of A. thaliana
oleosins in regulating lipid droplet size and preventing droplet fusion (Shimada et al. ,
2008, Siloto et al., 2006), or that of mouse adipocyte perilipin in regulating lipolytic
activity at the lipid droplet surface (Tansey et al., 2001). Moreover, recent genome-wide
RNAi screens in Drosophila cells implicated 1.5-3.0 % of all genes as directly or
indirectly involved in lipid droplet formation and/or regulation, and resulted in the
identification of a new role for Arfl-COPI vesicular transport machinery in regulating
droplet morphology and lipid utilization (Beller et al., 2008, Guo et al., 2008). In
contrast, few molecular details are known about algal lipid droplet biogenesis although
many TAG-rich algal species have been described (Hu et al., 2008).
Our efforts to identify proteins related to the PAT protein family or oleosins in the
Chlamydomonas reinhardtii genome (Merchant et al., 2007) or genomes of other green
algal and diatom species including Thalassiosira pseudonana, Volvox carteri, and
Chlorella sp.NC64A revealed no putative algal orthologs. In order to identify both
potentially novel and conserved proteins which function in algal lipid droplet biogenesis,
we studied the accumulation of TAG in lipid droplets of nitrogen-(N)-lirnited C.
reinhardtii cells, and identified candidate lipid-droplet associated proteins by mass
spectrometry.
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Materials and Methods
Strains and growth conditions. Chlamydomonas reinhardtii strains dw15 (cw15, nit],
mt+) and CC-125 (nitl, nit2, mt+) were grown in Tris-acetate-phosphate (TAP) medium
with N114Cl at 10 mM, in liquid cultures or on agar-solidified plates under continuous
light (70—80 umol m"2 s") as previously described (Riekhof et al., 2005). Strain CC-125
was used in generating the data shown in Figure 5.1B and Figure 5.2, and de5 or
transgenic derivatives thereof were used in the remainder of this study. To induce N-
lirnitation, mid-log phase cells grown in 10 mM NH4C1 TAP were centrifuged for 5
minutes at 5,000 x g, washed once in TAP lacking NH4C1 or any other N-source (TAP-
N), and resuspended in TAP-N. Cells were counted with a hematocytometer.
Isolation of lipid droplets from N-limited C. reinhardtii. Cells grown in 500 ml TAP-N
medium for 24 h were centrifuged 3,000 x g for 5 min and resuspended in 10 ml of
isolation buffer (50 mM HEPES, pH 7.5, 5 mM MgC12, 5 mM KCl, 0.5 M sucrose, 1 mM
phenylmethylsulfonylfluoride, 1 mM 2,2’-dipyridyl, with 1x protease inhibitor cocktail
[Rocbe, Branchburg NJ ]). Cells were disrupted with the aid of a Potter homogenizer,
using 20 strokes as well as sonication at the lowest energy setting (3 W output, Misonex
Sonicator 3000, Newton, CT) for 5 sec. The homogenate was overlayed with buffer 2
(same as isolation buffer above, but lacking 0.5 M sucrose) and centrifuged at 100,000 x
g for 30 min. Lipid droplets floating on the overlay buffer were removed with a bent
Pasteur pipette and diluted lO-fold with washing buffer (same as isolation buffer, but
with KCl increased to 150 mM). The suspension was overlayed with buffer 2 and
centrifuged as described above. This washing process was repeated three times, and lipid
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droplets were suspended in buffer 2 and stored at -80 °C. Preliminary attempts to isolate
lipid droplets with this method from cells deprived of nitrogen for 2 or 3 days resulted in
lipid droplets with significantly increased chlorophyll contamination compared to lipid
droplets fiom 24 h -N cells (data not shown)»as such lipid droplets from 24-h -N grown
cells were used throughout in this study. Thylakoid membranes were isolated according
to (Chua and Bennoun, 197 5), and enriched eyespots were prepared as previously
described (Schmidt et al., 2006).
Mass spectrometric identification of lipid droplet proteins. Proteins were precipitated
as previously described (Schmidt et al., 2006) and solubilized in SDS sample buffer by
sonication. Proteins were separated on 4-20% SDS-PAGE gels (Biorad, Hercules, CA)
and stained with EZBLUETM gel staining reagent (Sigma-Aldrich, St. Louis, MO). The
lane containing lipid droplet proteins was sliced into 15 bands, which were processed for
in-gel trypsin digestion as previously described (Shevchenko et al., 1996) and analyzed
by LC-MS/MS by the Michigan State University Research and Technology Facility. The
extracted peptides were re-suspended in a solution of 2% acetonitrile/O.l% trifluoroacetic
Acid to 20 pl. From this 10 pl were automatically injected by a Waters nanoAcquity
Sample Manager (Waters, Milford, MA) and loaded for 5 minutes onto a Waters
Symmetry C18 peptide trap (Sum, l80um x 20mm) at 4 uI/min in 2% aceto nitrile/O. 1%
formic acid. The bound peptides were then eluted onto a Waters BEH C18 nanoAcquity
column (1.7 pm, lOOum x 100mm) and eluted over 35 minutes with a gradient of 2% B
to 35% B in 21min, 90% B from 21- 24 minutes and 5% B after 24.1 minutes at a flow
rate of 0.6 111/min. Separations were done using a Waters nanoAcquity UPLC (Buffer A
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= 99.9% Water/O. 1% F ormic Acid, Buffer B = 99.9% Acetonittile/O. 1% F onnic Acid)
and sprayed into a Thermo Fisher Scientific (Waltham, MA) LTQ mass spectrometer
using a Michrom ADVANCE nanospray source. Survey scans were taken in the ion trap
and the top five ions from each survey scan were automatically selected for high
resolution Zoom scans followed by low energy collision induced dissociation (CID). The
resulting MS/MS Spectra were converted to peak lists using BioWorkS Browser v 3.3.1
(Thermo Fisher Scientific) using the default parameters and searched against the
Chlamydomonas reinhardtii database, v3.0, downloaded from The DOE Joint Genome
Institute (www.jgi.doe.gov), using the Mascot searching algorithm, v 2.1
(www.matrixscience.com). The Mascot output was then analyzed using Scaffold
(wwwproteomesoftwarecom) to probabilistically validate protein identifications using
the ProteinProphet computer algorithm (Nesvizhskii et al., 2003). Assignments validated
above the Scaffold 95% confidence filter were considered true. Mascot parameters for_
database searching allowed up to 2 missed tryptic sites, variable modification of
oxidation of methionine and carbarnidomethyl cysteine, a peptide tolerance of +/- 100
ppm, a MS/MS tolerance of 0.8 Da, and a peptide charge state limited to +2/+3.
Fatty acid and lipid analysis. Various cellular and sub-cellular fractions were extracted
into methanol:chloroform:formic acid (2: 1 :0.1 vol) followed by phase separation by the
addition of l M KC1, 0.2 M H3PO4. The organic phase was spotted on ammonium sulfate
impregnated silica Si250 plates, and TAG was separated with petroleum etherzdiethyl
etherzacetic acid (80:20:1 vol). After visualization by brief iodine staining, TAG bands
were quantitatively scraped from the plates and processed for fatty acid methyl
160
esterification (FAME) and quantified by gas chromatography as previously described
(Rossak et al., 1997). For total cellular fatty acids, the total cellular extract from a known
number of cells was processed directly for FAME as described above.
Construction of vectors and transformation of C. reinhardtii. All PCR reactions were
performed using the Phusion® High-Fidelity DNA polymerase (Finnzymes, Wobum,
MA) according to the manufacturer’s instructions, all ligation reactions were performed
with T4 DNA ligase (Invitrogen, Carlsbad, CA), and the sequences of the modified
regions of all final expression vectors were confirmed by DNA sequencing at the
Michigan State University Research Technology Support Facility. In order to construct
the pNITPROl vector, the C hlamydomonas RBCSZ 3’UTR was amplified from genomic
DNA with primers P1 5’-GTTGGTACCTGATAAGGATCCCCGCTC-3’ and P2 5’-
GTTGTAAAACGACGGCCAGTGCC-3’ and ligated into Kpn] digested pBluescript II
KS+ (Stratagene, Agilent Technologies, Santa Clara, CA) after digestion with the same
enzyme; a clone containing the desired orientation of the insert was identified by PCR
amplification using P2 5’-GTTGTAAAACGACGGCCAGTGCC-3’ and P3 5’-
TAATACGACTCACTATAGGG-3’ (the T7 sequencing primer), and the resulting
plasmid was designated pEMl. The C hlamydomonas N] T l promoter region was
amplified from the vector pMN24 (Fernandez et al., 1989) using primers P4 5’-
CGGT'TCTAGACGTGTATGGCT'I'TGGCTAC-3’ and P5 5’-
GGTAACTAGTACTGGCAGGATTCGGCTG-3’. This amplicon was cut with Pcil and
Spel and ligated into pRBCSZ after its digestion with the same enzymes; the resulting
plasmid was designated pEM2. PCR primers P6 5’- 1'1 IGCTCACATGTACAGGTG-3’
161
and P7 5’-CCCCCATATGACCCGCTTCAAATACGC-3’ were used to produce a NIT 1
promoter-MCS-RBCS2 3’UTR region of pERM2 amplicon, which was cut with Pcil and
ligated into a Pch/Swal digested pSP124S—a plasmid containing the ble selectable
marker gene for Chlarnydomonas transformation.
To generate an MLDP RNAi construct, primers P8 5’-
GCGCACTAGTCTGGAAAGCCTCTGAAGCAC-3’ and P9 5’-
CCTTCCATGGGGTCGTAGTACTTGTCGGCG-3’ were used to amplify a sense
amplicon from genomic DNA, while P10 5'-
CCCCCTGCAGCTGGAAAGCCTCTGAAGCAC-3’ and P11 5’-
GGAACCATGGTCGGTGCTCCGGAGAATCTT-3 ’ were used to amplify the antisense
amplicon from cDNA. The sense and antisense amplicons were double digested with
Spel/Neal and PstI/Nco], respectively, and ligated into Spel/Pstl digested pBluescript II
KS+. The formation of the triple ligation product was confirmed by DNA sequencing,
and the resulting hairpin insert was excised by Spel/Pstl digestion, and ligated into
pNITPROl digested with the same enzymes. The resulting plasmid was designated
pMLDP-RNAi. C. reinhardtii strain dw15 was transformed as previously described
(Lumbreras et al., 2008), except that vectors were linearized with Pacl, and transformants
were selected and maintained on TAP agar plates supplemented with 1 % yeast extract
and 10 pg ml"l zeocin.
Quantitative Real-time RT-PCR. Total RNA was purified using the RNeasy plant mini
kit (Qiagen, Gennantown MA) including the on-column DNAse digestion. 500 ng RNA
was reverse transcribed using the RETROscript kit (Ambion, Austin, TX) in a 10 pl
162
reaction. Real-time PCR was performed on an ABI Prism 7000 (Applied Biosystems,
Foster City, CA) using the SYBR green PCR master mix with the equivalent input of 50
ng RNA and 0.3 pM each primer (MLDP- F 5’-GGATGCCTGGACCAAGTTC-3’ and
MLDP-R 5’-GAGTGCACCAGGAGGTCGT-3’; RACKI-F 5’-
GACCACCAACCCCATCATC-3' and RACK I -R 5’-AGACGGTCACGGTGTTGAC-
3’). In all experiments, expression of MLDP was normalized to RACK I (CBLP)
expression, a gene commonly used for normalization in C. reinhardtii (Allen et al.,
2007). Analysis of RACK] expression by the 2"“ method (Livak and Schmittgen, 2001)
resulted in the following fold-changes (average and standard deviation are indicated) in
expression when using the average threshold cycle (Ct) value of 10 mM N grown cells:
1.015: 0.17 for 10 mM NI-I4Cl grown cells, and 1.14 :1: 0.1 l, 1.79 :1: 0.72, and 1.62 i 0.28
for cells switched to 0 mM N for l, 2, and 3 days, respectively. The standard curve
method for quantification of mRNA abundance was used as previously described
(Larionov et al., 2005). Melting curves were performed after PCR to confirm that the
amplification product was unique.
Fluorescent, confocal, and electron microscopy. For confocal microscopy, cells were
re-suspended in phosphate buffered saline (PBS). Cells were stained with Nile Red at a
final concentration of 2.5 pg ml'l (from a stock of 50 pg ml’1 in methanol), followed by a
10 minute incubation in the dark. Stained cells were then mixed 1:1 (vol/vol) with 2%
low-temperature melting point agarose kept at 38 °C and applied to the microscope slide.
Afier the solidification of the agarose, images were captured using a ZeiSS 510 Meta
ConforCor3 LSM microscope (Zeiss, Maple Grove, MN). For neutral lipid specific
163
detection of Nile Red fluorescence, the 488 argon laser was used in combination with a
560-615 nm filter; for combined chlorophyll autofluorescence and Nile Red Fluorescence
of polar lipids, a 647—753 nm filter was used. Samples were viewed with a 63x oil-
irmnersion objective. Postacquisition image handling was done with Zeiss AIM software.
For fluorescence microscopy, a Zeiss Axio Irnager M1 microscope was used. Nile Red
fluorescence in neutral lipids was detected using a UV light source with a 500 :h 24
urn/542 d: 27 nm exitation/emission filter set. For the measurement of apparent lipid
droplet diameter micrographs were analyzed using Image J software (Abramoff et al. ,
2004). For electron microscopy, centrifuged cell pellets were processed as previously
described (Harris, 1989), and transmission electron micrographs were captured using a
JEOLlOO CXII instrument (Japan Electron Optics Laboratories, Tokyo, Japan).
Bioinformatics. Sequences were routinely searched using BLAST (Altschul et al., 1997).
Sequences were aligned using the Tcoffee multiple sequence alignment program (Poirot
et al. , 2003). Hydropathy plots were generated employing the Kyte-Doolittle algorithm
(Kyte and Doolittle, 1982) using the ProtScale program at
www.cxpasy.ch/tools/protscale.html. The value G in each graph is the grand average of
hydropathy value (GRAVY) for each protein and was calculated using the GRAVY
calculator program at http://gravylaborfrustdel.
164
Results
TAG accumulation following shift to N-limiting medium. While C. reinhardtii does
not accumulate TAG under nutrient-replete growth conditions, TAG has been reported to
accumulate in this genus after N or phosphorous limitation (Van Donk et al., 1997). In
order to optimize conditions for the isolation lipid droplets from C. reinhardtii cells, the
time course of N-deprivation and its effect on TAG accumulation and lipid droplet
formation was analyzed. The amount of fatty acid esterified in TAG steadily increased
over time to approximately 12 %, 45 % and then 65% of total cellular fatty acids 24, 48,
and 72 h, respectively, after resuspending the cells in TAP medium lacking a source of N
as shown in Figure 5.1A Total fatty acid profiles of N-limited cells showed moderate
changes in comparison to N-replete cells, with a slight increase in 16:0 (carbon number:
3A9"2"5 (superscript indicates
number of double bonds), and concomitant decrease in 18:
position of double bond counting form the carboxyl end); TAG fatty acid profiles of N-
replete and N-limited cells were more distinct with a decrease in 16:0 and an increase in
18:1/16:4 in N-lirrrited cells (Table 5.1). Total cellular fatty acids increased on a per cell
basis 48 and 72 h after imposing N-limiting conditions (Figure 5.1A), indicating that de
novo fatty acid synthesis contributes at least in part to TAG accumulation.
A time course of lipid droplet formation in N-deprived C. reinhardtii cells is shown
in Figure 5.18. Using the lipid-specific fluorescent dye Nile Red, lipid droplet formation
was observed by confocal microscopy adjusting excitation and emission filters to
predominately detect fluorescence from the neutral lipid fraction (Elsey et al., 2007).
First lipid droplets were observed after 12 h of N-deprivation in distinct locations within
165
>
FA (nmol)/ 10 cells
to
Figure 5.]. Accumulation of triacylglycerols in lipid droplets during N-limitation.
(A) Quantification of total cellular fatty acids (black bars) and fatty acids in
triacylglycerol (grey bars) of dw15 cells grown in medium with 10 mM NH; (+N), or
cells switched (arrow) to medium with no NH! (-N) and grown for 24, 48, or 72 h. (B)
Confocal microscopy images of Nile Red-stained cc-125 cells grown in medium with 10
mM NH..‘ (+N) or cells switched (arrow) to medium with 0 mM NH,“ (N) and grown for
12, 24 or 48 h. Differential interference contrast (D), chlorophyll autofluorescence (C),
Nile Red fluorescence indicating lipid droplets (L), and all three images merged (M) are
shown. The scale bars indicate 5 pm.
166
Table 5.1. Total fatty acid content of whole cells compared to fatty acids bound in
triacylglycerols isolated from cells which were grown in medium containing 10 mM
NH: (+N), or cells switched to 0 mM N114+ and grown for one day (-N).
a Whole cells Whole cells TAG TAG
Fattyacid +N -N +N -N
16:0 22.1 1 0.14 26.2 :1 0.70 51.3 1 1.30 37.6 31 3.99
16:1 3.4 1 0.02 2.9 1 0.01 5.4 1 2.37 3.7 :1 0.54
16:2 1.2 1 0.00 1.4 1 0.06 1.6 1 0.96 1.4 1 0.17
16:3 4.5 :1 0.15 3.4 1 0.25 13.9 :1 6.88 3.2 :1 0.27
18:0 2.4 1 0.02 2.5 :1 0.06 0.8 1 0.84 1.4 :1 0.17
18:1,16:4 21.9 1 0.13 21.2 1 0.14 7.9 :1 2.46 22.1 1 8.45
18:2 8.0 3; 0.02 9.9 1 0.06 6.1 :1 3.33 13.6 31 1.51
18:3A5'9‘” 6.7 1 0.01 6.5 1 0.12 4.6 .1 1.62 5.7 1 0.63
18:3A9"2"5 27.9 1 0.10 24.7 1 0.13 7.3 :1 3.69 9.9 31 1.20
18:4 1.9 1 0.02 1.5 :1 0.02 1.1 11 0.16 1.3 :1 0.16
aThe data are presented on a % mol basis where three replicates were averaged and the
standard deviation is shown.
167
the cell, and prolonged exposure of the cells to 24 or 48 h of N-deprivation caused an
increase in the size and number of lipid droplets. Similarly, transmission electron
micrographs revealed several ultra-structural changes that occur over a 72 h time course
of N-limitation, including a reduction of stacked thylakoid membranes, accumulation of
starch granules, and the appearance of lipid droplets (Figure 5.2).
Properties of lipid droplets. Lipid droplets were prepared from 24 h N-deprived cells
and compared to isolated thylakoid membranes and a fraction highly enriched in
eyespots—a chloroplast localized, carotenoid and lipid-rich organelle (Schmidt et al.,
2006). Spectral analyses of the various subcellular fractions indicated the presence of
carotenoids (Figure 5.3A). This result was similar to what was previously observed for
an enriched eyespot fraction (Schmidt et al., 2006), although when normalized on a per
fatty acid basis, carotenoids were much less abundant in lipid droplets. The near absence
of chlorophylls in isolated lipid droplets represented by the absorbance maximum at
approximately 660 nm visible in the thylakoid fraction (Figure 5.3A) suggested low
contamination of the lipid droplets with thylakoid membranes. The ratios of TAG-
esterified fatty acids to total fatty acids of lipid droplets and eyespots were approximately
88% and 25%, respectively (Figure 5.3B), corresponding to the higher relative membrane
abundance in the eyespots. The TAG fatty acid profile of isolated lipid droplets was very
similar to the profile of TAGS isolated from cells deprived of N for 1d, whereas the TAG
fatty acid profile of eyespots was quite distinct, being much more enriched for palmitic
acid on a mo] percent basis (Table 5.2). Taken together, these results indicated that the
lipid droplets were distinct from the lipid globules found in the eyespot fraction, with
168
Figure 5.2. Ultrastructure of cells during N-limitation. Electron micrographs are
shown of a representative cc-125 cell grown in medium containing 10 mM NH.+ (0d),
and representative cells switched to 0 mM NH4+ and grown for one (1d), two (2d), or
three (3d) days respectively. Scale bars represent 2 pm. The Arrows are pointing at: E,
eyespot; LD, lipid dr0plets; N, nucleus; P, pyranoid; S, starch granules; T, thylakoid
membranes; V, vacuoles.
169
Figure 5.3. Characteristics of isolated lipid droplets. (A) Absorbance spectra of
ethanol extracts (96%) normalized on an equal fatty acid basis from thylakoids of cells
grown in medium containing 10 mM NH4+ (+N), or cells switched to 0 mM NIH and
grown for one day (-N 1d), from enriched eyespot fraction, and from lipid droplets were
recorded. (B) Ratios of fatty acids derived from triacylglycerol (TAG FA) to total fatty
acids (Total FA) for extracts from whole cells grown in medium with 10 mM N11,.+ (+N),
or whole cells switched to and grown in medium with no Nl-I..+ (-N) for 24, 48, or 72 h,
the enriched eyespot fraction (Eye), and isolated lipid dr0plets (LD) as indicated. (C)
Proteins (30 pg) separated by gradient (4-20%) SDS-PAGE from whole cells (Tot. Prot.,
total protein), thylakoids, eyespots, and lipid droplets as indicated. The positions
molecular weight markers in kDa are indicated on the left and sections excised from the
gel separating lipid droplet associated-protein are indicated on the right.
170
UJ
TAG FA : Total FA
1...
1.0-
d
0.81
0.6-
0.4-
d
0.2‘
+N -N -N -N 0
O ' I ' 1 ‘ l
350 400 450 500 550 600 650 700
Wavelength (nm)
L0
24h 48h 72b 11’;
250-
150-
100-
80-
60-
50
40
30-
25-
20-
15-
10.
Cell
\- ~b
«‘9 g
«0"
".
171
Table 5.2. Fatty acid content of whole organelles (eyespot and lipid droplet) compared to
fatty acids bound in triacylglycerols isolated from these organelles.
Whole Whole
a organelle TAG organelle TAG
Fatty acid Eyespot Eyespot Lipid droplet Lipid droplet
16:0 35.5 1 0.67 67.0 1 7.55 32.2 1 1.24 41.8 1 4.33
16:] 3.0 1 0.01 4.5 1 1.08 3.4 1 0.37 3.1 1 0.58
16:2 3.4 1 0.02 3.0 1 0.30 1.2 1 0.15 0.9 1 0.06
16:3 15.7 1 0.18 7.8 1 0.45 6.4 1 3.03 5.1 1 1.70
18:0 3.0 1 0.01 1.9 1 0.50 2.1 1 0.87 1.5 1 0.77
1821,1624 18.4 1 0.39 7.9 1 1.99 27.2 1 1.74 27.3 1 2.86
18:2 10.9 1 0.71 4.7 1 1.77 10.7 1 0.67 10.0 1 1.67
18:31”12 5.6 1 0.09 1.4 1 0.57 7.7 1 1.59 5.9 1 2.62
18:3A9"2"5 3.3 1 0.47 0.8 1 0.41 6.8 1 4.69 2.9 1 0.34
18:4 1.1 1 0.02 1.1 1 0.47 2.3 1 0.92 1.4 1 0.38
a Organelles were isolated from cells grown switched to 0 mM NH: and grown for one
day. The data are presented on a % mol basis where two replicates were averaged for
eyespot samples and three for lipid droplet samples. The standard deviation is indicated.
172
minimal contamination of thylakoid membranes, the predominant membrane fraction in
C. reinhardtii.
Analysis of proteins from whole cells, thylakoids, the eyespots, and lipid droplets by
SDS-PAGE followed by total protein staining (Figure 5.3C) revealed a pattern for lipid
droplets that was distinct from the other protein fractions, with one major protein of
approximately 27 kDa, as well as several less abundant proteins, which were Specifically
enriched in lipid droplets compared to thylakoids and the eyespots. The lipid droplet
proteins were excised from the gel (as indicated in Figure 5.3C) and subjected to mass
Spectrometry for protein identification.
Proteins identified in the lipid droplet fraction. As summarized in Table 5.3, 259
proteins with two or more unique peptides were identified and fell into a broad range of
protein functional groups (Figure 5.4A). Sixteen proteins predicted to be involved in lipid
metabolism, including three presumed acyl-CoA synthetases and a predicted
lipoxygenase—proteins previously identified as lipid droplet associated in studies of
other organisms (Bartz et al. , 2007, Cermelli et al., 2006)—were among those proteins.
BTAI, the enzyme responsible for the synthesis of a major extraplastidic membrane lipid
diacylglyceryl-N-trimethylhomoserine (Riekhof et al., 2005) was also present. Several
predicted components of vesicular trafficking pathways were identified, including
subunits of the Coat Protein Complex I and its putative regulator ARF la, as well as other
small Rab-type GTPases. Orthologs of these proteins have also been identified in lipid
droplet proteomics studies done on different animal species (Bartz et al., 2007, Cermelli
et al. , 2006), and recent Drosophila cell RNAi inactivation studies targeting Arj79a
(ARFIa ortholog) and the Coat Protein Complex 1 encoding genes revealed a previously
173
Amino acid metabolismls) I (24) Other metabolism
Channels (21' c- (3) Oxidative pentose phosphate pathway
Cytoskeleton13)l 11(4) Oxidative stress related
.1 _ I(2) Photoreceptors
Function unknonwn (45)I I(24) Photosynthetic apparatus
I(2) Proteases and peptidases
Glycolysis (S)I
Ion transporting ATPaseS(5)I
Kinases and 14-3-3 proteinsl7)I
.(7) Protein translocation.
chaperones and
protein transport
Lipid metabolism (16)I . .
I(54) Ribosomes. translation
Microtubule motor ml and DNA related
Mitochondria (26) I
I(7) Transporters
I(1) Ubiquitin protein modification
I(15) Vesicle trafficking
Vesicle Trafficking (7) I I (2) Channels
I (3) Cytoskeleton
Transporters (S)I
I(10) Function Unknown
Ribosomes, Translation and DNA
Related (12) a I(3) Glycolysis
I (1) ion transporting ATPases
I(3) Klnases and 14-3-3 proteins
Protein Translocation, chaperones,
and protein transport (2)-
proteases and peptidases (1) I I (10) Lipid metabolism
Photosynthetic Apparatus (12)-
l (8) Mitochondria
Photorecptors (1) I
oxidative stress related (3) I . (8) Other Metabolism
l (2) Oxidative pentose phosphate
pathway
Figure 5.4. Distribution of lipid droplet associated proteins into functional groups.
(A) Functional distribution of all 259 proteins identified with 2 or more unique peptides
and (B) proteins (93) with greater than 12 spectral counts.
174
Table 5.3. Complete list of proteins identified in the Chlarnydomonas lipid droplet
fraction.
1 . . . . . .
Manual annotation and categorization of proteins into functional groups
2Number of Spectral counts or unique peptides identified by LC MS/MS
3Deduced protein molecular weight (kDa)
4Occurrence of identified proteins in C. reinhardtii MS-proteomes of the eyespot
(Schmidt et al., 2006), mitochondria (Atteia et al., 2009), or flagellum (Pazour et al.,
2005)
5Protein ID and model name for identified proteins (C. reinhardtii genome v. 3.0)
v
“‘ g"! 59.. 5' E {P '6
Functional category and protein description' g g a g .5 2 g g 3 m." 3':
§ ‘6 , g i g o g]. 3 Protein ID
6 l u. u.
2
WM
1 Glycine hydroxymethyitransferase 4 3 51845 - 107904
2 alutanb-ganma-semiaidehyde dehydrogenase 12 3 41039 130812
3 glutanine synthetase 12 6 42128 I 133971
4 aspartate semialdehyde dehydrogenase 11 6 39917 148810
5 cobalamin-independent methionine synthase 4 86756 I I 154307
6 Aeetylglutamate kinase-like protein 3 35955 78991
cum
7 Voltage-dependent anion-selective channel protein 16 7 28533 I 127172
8 Porin 21 8 281$ I 146232
9.1mm
9 alpha tubulin 1 47 14 49569 I 128523
10 beta tubulin 2 83 17 49601 I_ 129868
1 1 aclin 23 9 41819 I I 24392
W
12 Truncated Hemoglobin 7 2 15677 159380
13 VFL2 Centrin 2 2 19442 I 159554
14 Calmodulin related protein 6 3 36353 I 183554
15 Calreticulin 2, calcium-binding protein S 3 47311 I 78954
16 predicted protein 18 8 28992 103840
17 sinilar to Hismacro and SEC14 domain proteins 3 2 14867 107313
18 similar to adenylate kinase 8 3 53990 109953
19 similar to aspartate aminotransferase S 3 44429 11 1917
20 predicted protein 11 4 30564 118714
21 predicted protein 11 5 30582 121991
22 similar to cell division inhibitor 18 5 32540 122924
23 glycoside-hydrolase-like protein 4 2 100235 128630
24 Deflagellation inducible protein 4 3 12693 I 131284
25 Flagellar Associated Protein 3 2 67438 I 132213
26 predicted ER protein, similar to disulphide isomerase 4 3 58220 133800
27 s'milar to chloroplast outer envelope protein 7 4 73326 143879
28 Uncharacterized conserved protein 2 2 93303 149656
175
Table 5.3 (cont’d).
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
$8983.88?»
8839';
61
62
63
65
67
68
69
70
71
72
73
hypothetical protein
predicted protein
predicted protein
predicted protein
predicted protein
predicted protein
predicted protein
predicted protein
predicted protein
predicted protein
Flagellar Associated Protein
predicted protein
Chlamydomonas specific tamiy protein
predicted protein
mitochondrial trmscription termination factor
predicted protein
Anlryrin repeat protein
TBZIDP1 and HVA22 related protein
predicted protein
predicted protein
Flagellar Associated Protein
predicted protein
predicted protein
predicted protein
predicted protein
Flagellar Associated Protein
predicted protein
predicted protein
180660661
Hexokinase
Phosphoglycerate Kinase
GAP1A glyceraldehyde 3-phosphate dehydrogenase
Phosphoglycerate mutase
Enolase
W
Vacuolar ATP synthase subunit 8
plasma membrane-type proton ATPase
Vacuolar ATP synthase subunit E
vacuolar ATP synthase, srbunit A
calcirmr-transporting ATPase, ER
Win:
Calcium-dependent Protein Kinase 1
cyclic nucleotide dependent protein kinase
protein kinase
5‘-AMP-activated protein kinase. gamma subunit
14-3—3 protein
14-3-3 protein
Serine/threonine protein kinase
W
176
15
13
42
11
nmmawgwwwwnuN-huwwmmoumuhanew
$00100)
11
N#O’@Ul
“@0001
11
29855
38 1 91
1 521 1
36027
27659
2 1 028
231 30
27063
69974
21621
376%
36056
20380
1 3390
25701
1 3620
20319
19849
24678
42641
22 149
46163
27963
29581
25885
190$
47143
12745
29524
49015
39742
515$
55782
1 17266
26226
68378
1 14325
67431
81010
44181
29497
27986
49490
151400
151947
155900
166924
168924
170664
173167
175668
176621
176660
162502 ,
163446
183515
164901
185148
165214
165396
166150
166179
166261
169631
192470
192623
193410
194644
194731
195067
79613
123020
132210
140618
161085
83064
126382
182602
190371
54608
77047
128451
131695
153242
184895
185967
187228
93758
Table 5.3 (cont'd).
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
119
120
Long—chain acyl-00A synthetases
Long-chain acyl-00A synthetase
Cycloartenol synthase
coclauine N-methyltiansferase
Long-chain acyl-00A synthetases (AMP-forming)
VTE3 MPBQ/MSBQ methyltransferase
Lipoxygenase
pyruvate-tormate lyase
Esterase/Iipaselthioesterase
Predicted phosphate acyltransferase
4'-phosphopantetheinyt transferase
Phospholipase DfTransphosphatidylase
TEF21 phosphoinositide phosphatase
3-beta hyrkoxysteroid dehydrogenasefisomerase
Betaine lipid synthase
Predicted phosphate acyltransferase,
W
Cytoplasmic dynein 1b heavy chain
In ! I .
Mitochond' rial substrate cam'er
MPC1 Mitochondrial Phosphate Carrier 1
UQ:Cth oxidoreductase 50 kDa core 1 subunit
NADqubiqu'none oxidoreductase sibunit 10
mitochond' rial F1F0 ATP synthase 36.3 kDa protein
predicted mitochondrial substrate carrier
mrtochond' rial F1F0 ATP synthase 31.2 kDa protein
NADH:ubiquinone oxidoreductase N09 subuni
NADqubiqu'none oxidoreductase 16 kDa subunit
mitochond’ rial F1F0 ATP synthase 14.3 kDa protein
NADqubiquinone oxidoreductase 49 kDa subunit
Mitochond' rial F1F0 ATP synthase. delta subunit
cytochrome c oxidase submit
NADqubiquinone oxrdored' uctase 39 kDa subunit
mitochond' rial F1F0 ATP synthase 13.3 kDa protein
Mitochondna' ' lcytoctrorne c oxidase 13 kD sibun'l
Cytochrome c oxidase subunit ll. protein llb
mitochond' rial substrate carrier protein
mitochondrial substrate carrier protein
mitochondrial F1F0 ATP synthase 19.5 kDa protein
NADqubiquinone citidoreductase 18 kDa subunit
F1F0 ATP synthase gamma subunit
ATP synthase, alpha subunit
UQ:Cth oxidoreductase 14 kDa sibunit
beta subunit ot mitochondrial ATP synthase
mitochondrial F1F0 ATP synthase, 60.6 kDa protein
museum
NAB-dependent epimeraseldehydratase
uridine 5‘- monophosphate synthase
glycosyl transferase, group 1
S-Adenosyl homocysteine hydrolase
177
1 30
62
74
78
92
13
11
13
231
14
11
12
12
18
160
5.: N
cwwbwawwouw
b
0001
10
21
18
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#- 000163000116:- 0907.5th
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4.6
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.6
#0)
17
19
14
10875
55555
85316
41223
72232
37446
1 18099
91 132
271 14
40174
91780
245%
50595
41515
75764
23923
481450
28548
37422
55037
18104
39710
34000
32124
15253
14307
52591
21154
11665
43637
16166
19010
17232
31616
25861
22185
17995
35077
61497
14024
61804
63145
36459
50921
46161
52701
111160
113915
116558
119132
123147
129760
144677
146801
160378
189038
189764
190403
190968
58501
77062
98450
24009
118965
135199
138803
139850
152682
159938
182740
182980
184222
184815
185013
185200
185375
186185
186501
187882
190125
191516
191596
192157
193762
194183
76602
77031
78348
78831
111778
116267
122195
129593
Table 5.3 (cont'd).
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
165
166
cytochrome b5 protein
gamma carbonic anhydrase
type-ll calcium-dependent NADH dehydrogenase
Superoxide dismutase.
MAD-dependent malate dehydrogenase
Apoferredoxin
NADH-cytochrome b5 reductase
sinilar to glycosyl hydrolase
succinate dehydrogenase
soluble inorganic pyrophosphatase
S-adenosylmethionine synthetase
chitinase—related protein
UDP-glucose dehydrogenase
Malate dehydrogenase.
isocitrate lyase
gamma carbonic anhydrase
putative quinone oxireductase
Short chain dehydogenase/reductase family protein
nucleoside diphosphate kinase-lire
Malate dehydrogenase
Transaldolase
6-phosphogluconate dehydrogenase
2W
2-cys peroxiredoxin, chloroplastic
glutathione S-transterase
glutathione-S-transferase
Sinilar to peroxiredoxin 4 (Bos taurus)
EM!
COP2 chlamyopsin retinal binding protein
phototropin
W
Ferredoxin-NADP (+) reductase.
CFO ATP synthase subunit ll precursor
Photosystem l protein PsaD
Chlorophyll A-B binding protein
PSBO Oxygen-evolving enhancer protein 1 ll
Chlorophyll A-B binding protein
Chlorophyll A-B binding protein
LHCA6 light-harvesting protein of photosystem I
chloroplast ATP synthase delta chain
LCHAS light-harvesting protein of photosystem I
LHCA8
Photosystem II reaction center protein PsbP
PSBQ
Chlorophyll A-B binding protein
Chlorophyll A-B binding protein
Rieske ferredoxin
PSAH Subunit H of photosystem I
178
U'lmNhflmm
137
WM
12
21
30
32
14
10
16
17
19
73
10
10
37
59
10
14
15
29
10
46
16
0017019301“
(0
1
mObNN
1
(DON NWQONN§
0004501
14
("#030203
a
O
N§bwb00101NUlw
14988
31 1 50
66955
25898
32267
13442
31 135
74848
49697
22157
42566
22350
52707
36584
45732
24274
21427
24604
16523
77644
42419
53348
25944
16037
23941
21624
81450
38231
1 6554
20891
22827
30697
28670
27764
23977
2821 1
25904
25882
21807
23883
27549
14991
14134
131692
132797
133334
135320
158129
159161
163751
169029
172283
174103
182408
184328
185081
190455
191668
195016
24039
55666
58944
60444
133861
146574
15891 1
142363
154723
193661
53816
164843
183965
105589
105641
120177
128836
130316
130414
131 156
131602
132678
133575
134203
148057
153656
174723
178631
182093
182959
Table 5.3 (cont’d)
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
201
202
203
205
207
208
209
210
211
212
213
Chlorophyll A-B binding protein
LHCBMG chloropyll a-b binding protein of LHCll
LHCB4 chlorophyll a-b binding protein
Chlorophyll A-B binding protein
Chlorophyll A-B binding piotein
Chlorophyll A-B binding protein
PSBR 10 kDa photosystem II polypeptide
W
Predicted serine protease
Peptidase M14. carboxypeptidase A
LA
HSP70-like protein
Heat shock protein 90A
similar to Tic32-3
Translocon-associated complex alpha subunit
Heat shock protein 70A
SECS1-alpha subunit of ER-translocon
Peptidyl-prolyl cis-trans isomerase, cyclophilin type
W
Histone H4
Histone H2A
Ribosomal protein L10E
Histone-lire bacterial DNA-binding protein
R'bosomal protein 86
Ribosomal protein L7Ae
Receptor of activated protein kinase C 1
RNA-binding protein
Ribosomal protein S17e
ATP-dependent RNA helicase
Ribosomal protein $9
608 ribosomal protein L-11
Ribosomal protein 608
Ribosomal protein S4
Rbowmal protein L356
Ribosomal protein L13
eukaryotic translation elongation factor 1 alpha 2
Ribosomal protein 83
Ribosomal protein 810
Ribosomal protein L9
eukaryotic initiation factor
Ribosomal protein S19
Ribosomal protein 824
Ribosomal protein L186
Ribosomal protein L14
elongation factor EF-3
Ribosomal protein L18a
Ribosomal protein 810
Ribosomal protein 327 isofonn A
Ribosomal protein L28
Ribosomal protein L276
179
19
10
10
16
19
mfifllflflm
NWNUN‘U
(JINUlbUthUOUIQUéNUl @b‘lh #
.5
N
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26670
26944
29921
27363
28685
26205
13025
85142
281%
72477
80667
38224
27049
71 198
52488
22660
11440
13668
26534
8949
28418
27091
35127
25272
15880
63436
15916
19469
9948
21154
12040
24466
50807
25830
13290
21501
24231
16980
15105
23958
15324
115293
21354
19291
9497
19480
16373
184070
184490
18481 0
186064
24036\
78552
98021
147894
184660
133859
1381 17
145585
185608
185673
188207
118846
100020
100147
103135
104082
104348
105289
105734
107394
108082
111269
113739
116329
118268
122980
127247
129809
132905
134051
139083
139831
140593
141798
143072
144684
145271
145770
146844
149993
150644
153674
155455
Table 5.3 (cont’d)
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
237
238
239
240
241
242
243
245
247
248
249
251
252
254
255
256
257
258
259
Ribosomal protein S 18
Ribosomal protein L23
Histone H26 variait
Ribosomal protein L136
Acidic ribosomal protein P0
Ribosomal protein 83a,
Ribosomal protein L6
glycine rich RNA binding protein
Ribosomal protein S4
eukaryotic initiation factor 4A-Iike protein
Ribosomal protein L30
Ribosomal protein 65
Ribosomal protein L22
eukaryotic initiation factor
Ribosomal protein 814
Ribosomal protein L21
EFGZ Elongation Factor 2
Ribosomal protein L27
Ribosomal protein L24E
Ribosomal protein 65
Ribosomal protein S156
Ribosomal protein 88E
Glycyl-tRNA synthetase and related
1m
similar to 1m phosphate transporter
Adenine nucleotide translocator
Sunni warmer swerfamily
putative 2-oxoglutaratelmalate trmsporter
Nitrate transporter
Major facfl'tator superfamily protein
plastidic ADP/ATP translocase
Ubiquitin-conjugating enzyme E2
W
Clathrin propeller, N-terminal.
Ras GTPase
RA823 smal Rab-related GTPase
alpha-COP
small Rab-related GTPase
COPBZ Beta'-COP
small Art-related GTPase ARFA1a
similar to VAMP-associated protein
Dynamin GTPase effector
COPE1 Epsilon-COP
small Rab-related GTPase
COPG1 Gamma-COP
small Rab-related GTPase
Sar-type small GTPase
Dynamin. Dynamin central region
180
11
20
14
16
10
10
13
26
10
26
S3
19
32
111
51
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24355
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47037
27559
21690
14444
23079
16284
18300
94193
15248
13636
30273
14821
25176
76729
31 660
3351 1
33431
481 93
57394
62295
16038
190068
21959
19725
1381 16
23594
107719
20569
27963
60945
31578
22581
96947
22103
21847
67594
155649
159282
160509
162845
164097
168484
183518
184151
188837
188942
190273
191776
1951 31
24172
24344
24354
24524
260
44781
56455
59755
76860
82920
116544
138185
169636
178501
192090
193659
195230
187409
1 12083
1 16488
126840
143349
148836
154280
183074
183897
186497
190416
60490
78205
81259
82091
98432
unknown role for these proteins in lipid droplet formation and utilization (Beller er al.,
2008, Guo et al., 2008). The presence of presumed orthologs in C. reinhardtii lipid
droplets suggests the possibility of a similar role in algal lipid droplet biogenesis.
Other functional groups which were represented by many proteins included 54
predicted ribosomal and translation related proteins, 26 predicted mitochondrial proteins,
and 24 predicted proteins of the photosynthetic apparatus. Ribosomal and mitochondrial
proteins have been identified in lipid droplet proteomic studies of different animal species
(Cermelli er al., 2006, Walther and Farese, Jr., 2009) and plant oil bodies (in plants from
mitochondria only) (Katavic er al., 2006), and could be common contaminating proteins
in lipid droplet preparations, or alternatively could indicate the presence of ribosomal-
/mitochondrial-lipid droplet physical interactions, as has been previously postulated
(Cermelli er a1. , 2006). The identification of 24 photosynthetic apparatus proteins was
surprising, given the near absence of chlorophyll in purified lipid droplets, and it is likely
that these hydrophobic apo-proteins are contaminating artifacts that become associated
during the isolation procedure; however, the most abundant stromal enzyme, ribulose-
1,5-bisphosphate carboxylase/oxygenase, was not detected. A more stringent analysis of
93 proteins (identified by more than 12 spectral counts) resulted in 6 ~38 % decrease in
the number of lipid metabolism proteins retained in the “identified” group (from 16 to
10), whereas relative decreases in the more predominant contaminating groups were
generally higher (~78 % for ribosome/DNA/translation related, ~69 % for mitochondrial
and ~50 % for photosynthetic apparatus proteins—see Figure 5.4B). In addition, 45
proteins of unknown function were identified. Based on spectral counts for all unique
peptides for a given protein (Table 5.3), one of these was by a factor of lo the most
181
abundant protein in gel slice 9 (Figure 5.3C) which also contained the most abundant
protein in the lipid droplet fraction based on staining. Its calculated molecular mass is 28
kDa which approximately corresponded to its position in the gel (Figure 5.3C).
Therefore, this protein was presumed to be the most abundant protein in the lipid dr0plet
fraction and was tentatively designated major lipid droplet protein (MLDP—protein ID:
192823 in the C. reinhardtii v. 3.0 database http://genome.jgi- p
psf.org/Chlre3/Chlre3.home.html;_NCBl accession number XP_001697668). Notably, the
second most abundant protein based on spectral counts was BTAl, the enzyme catalyzing
the biosynthesis of betaine lipid. Both proteins were also identified in study of C.
reinhardtii lipid droplets conducted in parallel, but dismissed as contamination (Wang et
al., 2009).
MLDP suppression affects lipid dr0plet size. Steady-state mRNA abundance of MLDP
was determined by northern blotting and quantitative real-time RT—PCR in 10 mM N-
grown cells, and cells deprived of N for 24, 48 and/or 72 h as shown in Figure 5. The
widely used C. reinhardtii control gene RACK I (CBLP) was used as an internal control
for RT-PCR. Analysis of apparent fold change in RACK 1 expression by the 2"“ method
(see Materials and Methods) indicated that changes in its expression based on total RNA
input to the reverse transcriptase reaction were minor if any (~l .5 fold increases at 48 and
72 h -N). Previous studies have shown that during N-deprivation in C. reinhardtii, the
amount of chloroplast and cytoplasmic ribosomes and total RNA content decrease
(Martin 6101., 1976). While this can potentially add an additional level of difficulty in
identifying a suitable control gene in these experiments, the degree to which this
182
contributes to the apparent minor fold-changes in expression for RACK 1 is unknown.
With the above in mind, RACK 1 was deemed a suitable control gene for these
experiments, with the caveat that it may slightly underestimate the fold-changes for genes
induced by 48 or 72 h N—deprivation, and vice versa for down-regulated transcripts.
When normalized to RACK 1 , MLDP mRN A showed a 16-fold increase in abundance
after 24 h of N—deprivation and continued to be abundant even after 72 h of N-
deprivation. Moreover, MLDP mRN A abundance qualitatively corresponded during a
time course of N-deprivation with the accumulation of TAG (Figure 5.1A). To begin the
functional analysis of MLDP in C. reinhardtii MLDP RN Ai lines were generated. For
this purpose, a plasmid carrying an expression cassette which is inducible by N-
deprivation, pNITPROl , was constructed based on the nitrate reductase (NI T I ) promoter.
The pNITPROl vector was used to drive the expression of an MLDP genomic sense-
cDNA antisense RNAi hairpin (Figure 5.6A). Zeocin-resistant transformants were
screened for MLDP mRNA reduction at 48 or 72 h N-deprivation. Of 64 lines screened,
9 lines were identified with a range of MLDP transcript reduction between 20 and 60% of
empty vector controls. The four strongest MLDP-RNAi lines (A l , C7, E9, and G1 1)
showed a consistent 55-60% reduction of MLDP transcript measured after 72 h N-
deprivation from three independently grown cultures (Figure 5.6B).
The MLDP-RNAi lines were analyzed for possible lipid droplet morphological
phenotypes by fluorescence microscopy analysis of Nile-Red-stained cells deprived of N
for 3 days. An apparent increase in the average size of lipid droplets in the MLDP-RNAi
lines was observed (Figure 5.7). The apparent individual diameters of a population of
greater than 200 random and in focus lipid droplets per RNAi or control lines were
183
B
20 1
«$529 6““‘0’6 g '
«9 x‘ \9 x‘ '2 16:
3
.. 12 a:
MLDP 5 -
e. Z 8 J
1: .
E 4 -
i g .,
rRNA 0 .
+ N 24h 48h 72h
-N
Figure 5.5. MLDP mRNA is induced following nitrogen deprivation. (A) Northern
blot analysis of MDP, where 5 pg total RNA isolated from l0 mM NH4Cl grown cells
(10 mM N) and cells deprived of N for 2 days (-N 2 d) was processed for northern
blotting. Total RNA was stained with methylene blue and shown as a loading control
(rRNA). Two biological replicates of this experiment are shown (panels 1 and 2). (B)
Relative abundance of MLDP mRNA normalized to RA CK I (CBLP) in wild-type dw15
cells grown in medium with 10 mM NHf (+N), or dw15 cells switched to and grown in
medium with no NHf {-N) for 24, 48, or 72 h measured by QRT-PCR. Data are
expressed as a fold change compared to the +N grth condition which are given a value
of l. Standard deviation of three biological repeats, each done in triplicate is shown.
184
RBCSZ 3’UTR
NIT1Pr0 ll—
...5’UTR 1
91 ‘_ ‘
.nJJIlllllllll] i
.' ~ ’
91 310 422
"301R
.3
N
Rel mRNA Abundance w
.o .o
‘5 °.°
A
O
EV1 EV2 A1 C7 E9 G11
Figure 5.6 Generation of MLDP RNAi-mediated knockdown strains. (A) Diagram
showing the MLDP-RNAi construct designed with the NIT 1 inducible promoter (Pro) of
vector pNITPROl driving the expression of an MLDP genomic inverted repeat (numbers
indicate basepairs from predicted S’UTR), and the predicted transcribed RNA hairpin
product. (B) QRT—PCR analysis of MLDP mRNA levels in two empty vector control
lines (EVl and EV2), and four MLDP-RNAi lines isolated from four independent
transformation events (A1, C7, E9, 61 1). Data are represented as percent change
compared to RV] and normalized to RACK] as in (A). Three replicates were averaged
and standard deviation is shown.
185
>
L0 Diameter (pm)
OJ
E2511) E2613 (2213) (£56) (2E299) (2114}
A1...
Figure 5.7. MLDP RNAi knockdown lines exhibit a lipid droplet morphological
phenotype. (A) Analysis of apparent lipid droplet diameter in empty vector controls and
MLDP-RNAi lines shown in Figure 5.5. The average and standard deviation are shown
for n (indicated in parentheses below each bar) lipid droplets counted, and cell lines are
marked with an asterisk, for which P < 0.01 when compared to EV] in a two-tailed
Student’s t-test. (B) Fluorescent micrographs of representative cells of lines EVI and Al
stained for lipid droplets with Nile Red.
03
EV1
186
measured after three days of N-deprivation (Figure 5.7A). An approximate 40% increase
in the average lipid droplet diameter was determined in the MLDP-RNAi lines.
MLDP has orthologs only in green algae. The newly discovered MLPD protein has few
orthologs, none of which has been functionally characterized. When searched against the
National Center for Biological Information database of non-redundant proteins using
BLAST (Altschul er al., 1997) no sequence with significant similarity is identified.
However, several genome projects of green algal species are currently underway and
searching against green algal genomes at the respective websites, potential orthologs with
high sequence similarity were identified in Chlorella sp. NC64A, Chlorella vulgaris and
Volvox carterr'. No discemable ortholog was present in available diatom or red algal
genomes or the genomes of Osrreococcus species, which belong to a green algal lineage
distinct from C. reinhardtii. Apparently, MLDP is a rare protein in nature as we currently
know it at the molecular level and presumably arose during the evolution of green algae.
As such it also has no discernible domains to which functions have been assigned.
Grand average of hydropathy (GRAVY) indices and Kyte-Doolittle (Kyte and
Doolittle, 1982) hydropathy plots were generated for MLDP, as well as for mouse ADRP
and Perilipin, and Arabidopsis Oleosin l (OLEl) (Figure 5.8 A-D). All four proteins
were distinct with respect to extent and region of hydrophobicity, with the most
hydrophobic region for MLDP occurring between amino acid residues ~l 60-215. The
proteins ranked according to their GRAVY index (highest to lowest) are: MLDP (0.l l),
OLE] (0.17),
187
Figure 5.8. Hydropathicity plots of MLDP and other known lipid droplet binding
proteins. (A-D) Hydropathicity plots for the proteins (followed by NCBI accession
numbers in brackets) MLDP [XP_001697668] (A), mouse ADRP [NP_031434] (B),
mouse Perilipin [NP_783571] (C), and Arabidopsis OLE] [NP_l94244] (D). The value
G in each graph is the grand average of hydropathy value (GRAVY) for each protein and
was calculated using the GRAVY calculator program at http://gravyjaborfi’ustdel.
188
35-
236:041
15.
05.
415.
445
415.
3.5 v v r '
0 50 100 150 200 250
C3 ' 0 100 200 300 400
35-
2.5- = -0.40
[3 ' 0 100 200 300 400 500
0 50 100 150
189
ADRP (028), and Perilipin (-.40). Thus, MLDP is a very hydrophobic protein although
it lacks a specific hydrophobic core characteristic for oleosin.
Discussion
The potential of microalgae to provide sustainable feedstock for biofuel production has
led to renewed interest in the biology of this diverse group of organisms (Hu et al., 2008).
While C. reinhardtii is unlikely to become the microalgae of choice for the production of
biofuels, it is a well established genetic and genomic model for the study of unicellular
green algae (Merchant er al., 2007). However, very little is known about lipid metabolism
or the cell biology of lipid droplets and its regulation in this organism. The time course
and conditions for the induction of oil accumulation described here (Figs. 5.], 5.2) and
the detailed analysis of lipid droplets provides the basis for a molecular analysis of the
mechanisms of TAG accumulation in C. reinhardtii and possibly unicellular green algae
in general. As a logical first step, we focused on the TAG-storing compartment itself, the
lipid droplet. Its fatty acid and pigment composition is clearly distinct from that of other
organelles (Figure 5.3). This distinct composition and the very low chlorophyll content in
view of the predominance of thylakoid membranes in the cell suggested that the obtained
oil droplet preparation was only slightly contaminated with thylakoid membranes, the
largest membrane system in the cell. However, we cannot rule out some level of
contamination with ER—derived microsomes especially since lipid droplets are generally
thought to be derived from the ER. An increase in the total amount of fatty acids in N-
deprived cells indicated that the biosynthesis of TAGS is not merely due to conversion of
membrane lipids but the result of de novo synthesis. The specific fatty acid composition
190
of lipid droplet lipids tended towards shorter and more unsaturated fatty acids suggesting
that newly synthesized fatty acids were quickly incorporated into TAGs before fatty acid
modification by desaturases or elongases can occur; alternatively, reduced activities of
these fatty acid metabolic enzymes could account for this observation.
Lipid droplets in different organisms are increasingly recognized as dynamic
organelles that provide important nodes of cellular metabolic networks and participate in
intracellular lipid trafficking (e.g. Guo et al., 2008). In algae, induction of lipid droplet
formation is intricately regulated by nutrient deprivation and other abiotic stresses that
lead to a cessation of cell division or an over-reduction of the photosynthetic electron
transport chain (Hu et al., 2008). In this study, we sought to identify the proteins
associated with lipid droplets to determine factors involved in the biogenesis or turnover
of this organelle in C. reinhardtii. Aside from those of unknown function, of particular
interest are lipid metabolic enzymes and proteins involved in vesicular transport
Intriguingly, the betaine lipid (diacylglyceryl-MMN-trimethylhomoserine) biosynthetic
enzyme, BTAI (Riekhof er al., 2005), had the second highest spectral count of proteins in
the lipid droplet fraction. This protein was also identified as one of few in the lipid
droplet fraction in a parallel study, but considered a contamination (Wang et al., 2009).
This parallel study concluded that no protein was specifically associated with the
respective lipid droplet fi’action prepared and treated differently than in this study.
Here, a number of predicted acyl-CoA synthetases and acyltransferases were
present as well that could be involved in the biosynthesis of TAGs or the transfer of acyl
groups between membrane lipid and neutral lipid pools. In animal cells, enzymes
involved in phosphatidylcholine biosynthesis have been found critical for lipid droplet
191
formation and turnover. Lipid dr0plets in animal cells are surrounded by a monolayer of
phospholipids and the availability of those phospholipids affects lipid droplet size. When
the rate-limiting enzyme of phosphatidylcholine biosynthesis was inactivated in
Drosophila cells, lipid droplets increased in size presumably because the larger droplets
have a lower surface area to volume ratio and require less phospholipid to form the
phospholipid monolayer (Guo et al. , 2008). As C. reinhardtii lacks phosphatidylcholine,
it seems plausible that its role in the formation of lipid droplets is taken by betaine lipid,
which has similar properties and is thought to functionally substitute for
phosphatidylcholine in this green alga (Sato and Murata, 199]). A rich complement of
proteins predicted to be involved in vesicular trafficking including COP] homologs, Rab
GTPase, or ARF related GTPase were also identified in the proteomics data set.
Repression of the respective orthologs in Drosophila cells was shown to affect lipid
droplet formation and dynamics and TAG utilization (Guo et al., 2008). Our
understanding of the interaction of lipid droplets with the vesicular transport machinery is
only rudimentary at this time. The presence of the respective proteins in C. reinhardtii
follows a common theme also observed for animal cells and provides an opportunity to
study the role of the vesicular transport machinery in lipid droplet formation in a
unicellular model organism. However, the specific association of these proteins with
lipid droplets in C. reinhardtii remains to be confirmed.
The identification of proteins in the ribosome-ltranslation-lDNA-related,
mitochondria, and photosynthetic apparatus functional groups suggest a basic level of
contamination in our lipid droplet protein fraction. While more stringent filtering of the
proteomics data set to include proteins with more than 12 spectral counts resulted in a
192
larger decrease in the relative proportion of these apparent contaminating functional
groups, when compared to lipid metabolism proteins (Figure 5.4, Table 5.3), the
significance of such analyses should be regarded as circumstantial given the potential for
protein size bias in the method. Comparing the analysis of the lipid droplet proteins
presented here with proteomic studies of the eyespot (Schmidt etal., 2006), flagella
(Pazour er al., 2005), and mitochondria (Atteia et al., 2009) (Table 5.3) reveals that some
degree of contamination with proteins from specific functional groups is common (e.g.
ribosome/nanslation-IDNA-related, mitochondria and photosynthetic apparatus proteins
found in the eyespot study, or ribosome contamination in the mitochondrial and flagellar
studies). As such, apparent contamination of subcellular preparations in C. reinhardtii
seems to be a common challenge.
To begin the functional analysis of lipid droplet associated proteins in C.
reinhardtii we focused on the most abundant protein in the lipid droplet fraction based on
Coomassie Brilliant Blue staining (Figure 5.3C) and spectral counts (Table 5.3). The
identity of this protein was deduced by mass spectrometric analysis of all proteins in the
respective gel slice and MLDP was identified based on lO-fold overabundance of spectral
counts for its peptides. Repeated attempts to localize an MLDP-Green Fluorescent
Protein (GFP) fusion protein to the lipid droplets failed because none of the tested
constructs expressed in C. reinhardtii. Unlike plants, C. reinhardtii does not seem to have
an ortholog of oleosins, distinct structural proteins surrounding the lipid bodies in plant
oil seeds (Huang, 1992, Siloto er al., 2006). Interestingly, oleosins are also absent from
oil-rich mesocarp tissues of plants like oil palm or olives and their presence may be
related to special needs of seeds. Indeed, reduced oleosin content leads to delayed
193
germination and reduced freezing tolerance (Shimada er al., 2008, Siloto et al., 2006). At
the cellular level, a reduction in oleosin caused an increase in oil body size. Likewise,
when MLDP was inactivated in C. reinhardtii, an increase in lipid droplet size was
observed (Figure 5.7). Despite an extensive screening of transgenic lines expression of
the MLDP gene could only be reduced approximately 60% in the best RNAi lines leading
to a moderate but statistically significant increase in oil droplet size in these lines.
Because MLDP lacks any features that readily classifies it as a protein with known 1""
catalytic activity, it may function as a structural protein affecting lipid droplet dynamics
and size in C. reinhardtii. An increase in lipid droplet size has been shown in different
systems to delay turnover of TAGs. We measured TAG levels (at 3 d —N) and TAG ls
turnover in these lines following the resupply of N in the medium but could not detect a
statistically significant difference in the levels of TAG or rate of TAG metabolization in
these lines compared to the empty vector control lines. It seemed possible that the
observed moderate change in lipid droplet size would lead to only a small change in the
rate of TAG metabolization, too small for it to be detected within the statistical
limitations of our analytical method. This does not rule out the possibility that MLDP is
important in preventing lipid droplet fusion to provide a larger surface area per volume
ratio to increase the accessibility of enzymes metabolizing TAGS. Intriguingly, lipid
droplets isolated from N-deprived cells of the green alga Eremosphaera viridis
(Chlorophyceae) were found to have 26, and 28 kDa associated proteins (V echtel er al.,
1992); while these masses are similar to that of MLDP, the study was conducted prior to
the development of facile protein identification by mass spectrometric methods, and as
such, the identity of the E. viridis lipid droplet proteins remains unknown.
194
The newly discovered MLDP is a rare protein in nature unique to green algae.
The expression of its gene is strictly limited to conditions favoring TAG biosynthesis,
providing indirect evidence for a role of MLDP in lipid droplet formation or
maintenance. These properties potentially make MLDP a marker for lipid droplets and
TAG accumulation.
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Chapter 6
Conclusions and perspectives
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Conclusions and perspectives
During the past five years in which the research for this dissertation was
conducted, tremendous advances have been made in our understanding of glycerolipid
metabolism in plants. Lipid metabolic pathways are tightly regulated during normal
growth and development, but can be highly modulated in response to environmental
stress. A number of new enzymes and components of lipid trafficking pathways have
been identified, resulting in the emergence of a clearer overall picture of the interplay
between chloroplast and endoplasmic reticulum (ER) localized glycerolipid assembly
pathways (Benning, 2009, Wallis and Browse, 2010). The regulation of chloroplast
galactolipid metabolism has been a major focus in the field, not only due to the
predominance of these lipids in plant leaf membranes, but also because of their essential
role in providing a matrix for the thylakoid-localized photosynthetic machinery
(Diirmann and Benning, 2002). There are two well-studied pathways for galactolipid
biosynthesis in Arabidopsis: a constitutive and an alternative induced pathway activated
by stresses such as phosphate deprivation. A major question that existed at the onset of
this work (and one that still stands) is: what are the factors required for activating the
alternative galactolipid pathway during phosphate deprivation?
DGSI is not directly involved in the regulation of galactolipid biosynthesis in plants.
The work described in Chapter 2 was aimed at answering the above question.
Suppressor mutants in the dgdl mutant background, which is deficient in the constitutive
pathway for DGDG biosynthesis, were identified by Changcheng Xu and colleagues in
the Benning lab (Xu et al., 2003). The suppressors fell into two classes. First the tgd
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mutants, which constitutively accumulate oligogalactolipids through the activation of a
galactolipid:galactolipid galactosyltransferase (GGGT-further discussed below). Further
detailed analyses of the tgdl/2/3/4 mutants revealed an essential role for the TGDl/2/3/4
proteins in the chloroplast to ER lipid trafficking pathway (Awai et al., 2006, Benning,
2009, Lu et al., 2007, Xu et al., 2003, Xu et al., 2008). The second suppressor class
consisted of the dgdl suppressor I (dgsl-1) mutant (Xu et al., 2003). The gene affected
by the dgsl-l mutation was found to encode 6 ~70 kDa protein of unknown function,
DGSI , in which the mutation caused a D457N mutation. Work described in Chapter 2
describes the detailed characterization of DGSI through the analysis of the dgsl-l mutant
as well as a null insertion allele dgsl-2. The DGSI protein was found to be localized to
the mitochondrial outer membrane in a high MR complex. Importantly, the dgsl-I
mutation was found to behave semi-donrinantly, and the phenotypes observed in this
mutant background including activation of DGDl -independent galactolipid biosynthesis,
increased hydrogen peroxide levels, and lowered alternative oxidase protein levels were
not observed in the dgsl-2 null allele. Moreover, the alternative galactolipid pathway in
dgd1 dgsl-2 double mutants was not affected when compared to dgd1 plants, and no
clear phenotypes were established for dgsl-2 mutants. The dgd1 dgsl-l double mutant
also showed a larger increase in DGDG accumulation during phosphate deprivation when
compared to the dgdl control, indicating that dgsl-1 suppresses dgdl by activating a
galactolipid pathway that is distinct from the phosphate regulated one. With these
observations, it became evident that DGSI is not directly involved in the regulation of
galactolipid biosynthesis in plants, and the activation observed in the dgsl -1 background
is a secondary effect, which is commonly observed in dominant alleles (Veitia, 2007).
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If DGSl is not involved in regulating galactolipid metabolism, then what is its
function? A lack of observable phenotypes in the dgsl-2 null allele compounds the
difficulty in determining the function of this unknown protein. DGSl has orthologs in
other plants, algae, fungi, protists and yeast. The Saccharamyces cerevisae ortholog,
NCA2, has been studied as a mutant, but only shows phenotypes as a double mutant with
the gene encoding NCA3 (Carnougrand et al. , 1995). Lack of phenotypes in both
Arabidopsis and yeast mutants in shared DGSI orthologs could be due to conditional
importance of these proteins, where the right stress condition has not been investigated.
Though DGSland its orthologs have no apparent paralogs within any species, non-
homologous redundant proteins that have evolved through convergent evolution to have
the same function as DGSl-like proteins could also result in a lack of observable
phenotypes. While not found in all organisms, DGSI orthologs have been maintained in
genomes of highly diverse species, suggesting a potential for conditional importance of
DGSl. Future work on DGSI should focus on the null dgsl-2 allele, possibly running it
through a gauntlet of stress conditions to see if any phenotype can be associated with the
loss of DGSI. Furthermore, with ever-expanding genomic resources available, the use of
comparative genomics approaches (Hanson et al., 2010) may assist in developing
hypothesis-driven research in identifying the function of DGSI.
GGGT is encoded by the SFR2 gene in Arabidopsis and galactolipid remodeling in
the chloroplast envelope is essential for freezing tolerance.
Analysis of galactolipids in dgd1 dgsl-I plants during phosphate deprivation
revealed the condition-specific accumulation of oligogalactolipids (Chapter 2). This was
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similar to the above-described observations in the tgd class of dgdl suppressor mutants.
In both cases, the oligogalactolipids formed in these mutants were from the presumed
activity of GGGT, an enzyme capable of transgalactosylation between galactolipids,
which through processive activity is able to synthesize the oligogalactolipids tri- and
tetra-galactosyldiacylglyerol. GGGT remained unidentified in molecular/ genetic terms at
the onset of this study (Benning and Ohta, 2005). Chapter 3 describes a reverse genetics
approach that was successful in identifying the GGGT-encoding gene. Importantly, this
work revealed GGGT to be encoded by a previously described gene in Arabidopsis,
SENSITIVE T 0 FREEZING 2 (SFR2). SFR2 was first identified through a forward
genetic screen looking for mutants defective in acquired freezing tolerance, and found to
encode a glycosyl hydrolase family 1 (GH-l) enzyme of unknown function (Thorlby et
al., 2004). Work in Chapter 3 shows that sfi'2 null mutants are unable to produce
oligogalactolipids, which are formed at the expense of monogalactolipid (MGDG) during
freezing stress in wild-type plants. In addition, triacylglycerol accumulates during
freezing through the acylation of diacyl glycerol produced by the GGGT reaction. The
SF R2 protein was found to be highly active when expressed in yeast, and was sufficient
to catalyze GGGT activity (discussed in detail in Chapters 1 and 3). Considering this
data, along with the wealth of information generated in understanding the mechanisms of
freezing damage in plants (Steponkus, 1984, Thomashow, 1999), a model for the role of
SFR2 in membrane protection during freezing was formed. During freezing, membranes
are the site of damage due to cellular dehydration. SFR2 (GGGT) remodels the
chloroplast outer envelope lipid composition by converting non-bilayer forming MGDG
to bilayer forming digalactolipid and oligogalactolipids in order to prevent membrane
203
fusion in a dehydrated state. Triacylglycerol formed during freezing stress may act as a
means of removing membrane lipids and reducing the effective bilayer surface area of a
given membrane in order to cope with a shrinking cell volume during dehydration.
While the identification of GGGT and its role in freezing tolerance are a
significant advance in understanding galactolipid biosynthesis and its role in plant stress
responses, many questions about GGGT/ SF R2 remain. First among these is whether or
not the role of GGGT/SF R2 is restricted to freezing tolerance. Orthologs of SFR2 are
present in plants with no freezing tolerance (e.g. maize), suggesting that SFR2 may play a
more general role in lipid remodeling during severe water deficit—drought and freezing
tolerance responses in plants have long been known to have significant overlap
(Thomashow, 1999). Analysis of the 31?? mutants described in Chapter 3 for sensitivity
to drought or high salinity will be a straightforward approach in addressing this question.
In addition, the analysis of sfr2 tng/2/3/4 double mutants could reveal the significance of
the constitutive activation of GGGT in the tgd class of mutants.
Importantly, the mode of activation of GGGT/SFR2 activity is still unknown.
The SFR2 protein is constitutively present and not regulated by cold acclimation in
Arabidopsis leaves (Thorlby et al., 2004), and GGGT can be activated by a number of
hyperosmotic treatments of plant leaves (Chapter 3). All of this indicates a post-
translational mode of activating the enzyme during hyperosmotic stress. One potential
mode of activation could occur through post-translational modification (PTM), e. g.
reversible protein phosphorylation, through a signaling cascade that is activated by
cellular dehydration. The in vitro data suggest the possibility of a different mode of
activation—the enzyme is highly active in heterologous expression systems in assays
204
where the substrate MGDG is dispersed in detergent (Chapter 3). While an endogenous
protein present in the heterologous host (yeast) used to express the SFR2/GGGT protein
could provide the same activating PTM, this seems unlikely. Alternatively, the enzyme
could be directly activated by mechanosensation of membrane destabilization. For
example, when the GGGT/SF R2 protein is present in liquid crystal lamellar (bilayer)
phase biomembranes, it is inactive. When conditions change, such as the formation of
non-bilayer microdomains during membrane dehydration (e. g. during freezing), or
detergent destabilization (e.g. the in vitro assay), a dynamic shift in equilibrium could
result in the activation of GGGT/SFR2 to remodel MGDG into DGDG and
oligogalactolipids. Examples of direct mechanosensation-based activation or inhibition
of lipid metabolic enzymes, such as phospholipase A2, have long been known (Lehtonen
and Kinnunen, 1995). While only circumstantial evidence currently exists for the
possibility of GGGT/SFR2 to be activated through mechanosensation—this mode of
activation is simpler and more direct than activation through PTMs—which may not be
effective modes of regulation at sub-zero temperatures or states of severe dehydration
during which GGGT/SFR2 is activated. This aspect of GGGT activity could be
addressed by studying the activity in vitro as in Chapter 3, but where the enzyme has
been reconstituted into large unilamellar vesicles (Szoka, Jr. and Papahadjopoulos, 1980).
A preliminary experiment indicated that GGGT/SFR2 is inactive under these conditions
(data not shown). With GGGT/SFR2 starting out inactive on in vitro vesicle bilayers,
small molecules known to induce non-bilayer structure formation in vesicles could be
added and the change in GGGT activity observed. Point or truncation mutants of the
heterologous enzyme could be employed in identifying a potential mechanosensing
205
domain of GGGT, if initial results indicate its presence. Indeed, many other questions
about GGGT enzymology could shed light on its overall function in plants. For example,
GGGT/SFR2 is a glycosyl hydrolase, which has evolved high transglycosidase activity.
Mutagenesis of the protein, followed by the standard activity assay could reveal residues
or domains which are required for the high transglycosidasezhydrolase ratio exhibited by
this enzyme. Results from both continued genetic study of SFR2 in planta and detailed
enzymology of GGGT/SF R2 will provide great insight into the mechanisms through
which membranes are actively remodeled in order to cope with changing environmental
conditions.
Triacylglycerol accumulation in Chlarnydomonas reinhardtii: a role for MLDP in
lipid droplet biogenisis?
The accumulation of triacylglycerols (TAGS) and lipid droplet composition in the
green alga Chlamydomonas reinhardtii was investigated as described in Chapter 5. TAGS
were found to accmnulate substantially during nitrogen deprivation (to ~65 % of total
cellular fatty acids at 3 days -N), and accumulated in lipid droplets (LDs) within the cell.
An LD fraction was purified, and the associated proteins identified through a mass
spectrometric approach. An abundant LD protein, designated Major Lipid Droplet
Protein (MLDP) was selected for further study. MLDP encodes a ~28 kDA protein with
no known functional domains, and the MLDP mRNA was found to be strongly up-
regulated during nitrogen deprivation. RNAi-mediated knockdown of MLDP resulted in
a slight, but statistically significant increase in the average apparent lipid droplet diameter
when compared to empty vector controls.
206
While this indicates a possible role for MLDP in lipid droplet morphology in
Chlarnydomonas, this phenotype has been observed by knockdown of several different
proteins in Drosophila which fall into broad functional classes (Beller e101,, 2008, Guo et
al., 2008). As such, this phenotype on its own yields little mechanistic insight into the
function of MLDP. As such, much work remains to be done on MLDP. Numerous
attempts to over-express the MLDP protein as a GFP transgenic fusion failed—a
common drawback of working in Chlarnydomonas, where successful transgene over-
expression remains a community-wide bottleneck (Neupeit etal., 2009). As such, the
confirmation of MLDP as an LD-associated protein will have to be conducted through
more classical approaches, such as raising a specific anti-MLDP antibody for use in
histochemical microscopy-based localization of MLDP, or through immuno-enrichment
of the MLDP signal in LD fractions observed by western blot. Furthermore, potential
functions for MLDP might be discovered by trying to complement mutants defective in
various LD-associated proteins in other species such as yeast (Athenstaedt et al. , 1999),
plants (Siloto et al., 2006), or fruit flies (Guo et al., 2008). Lastly, the potential for using
homologous recombination in targeting MDP for insertional mutagenesis, though
challenging, may be possible (Zorin et al., 2009). A true null allele of MLDP could
provide greater insight into MLDP function, as the MLDP-RNAi knockdown level
achieved was only ~60 % (Chapter 5).
In addition to MLDP, many other factors are likely to be involved in TAG
accumulation and LD formation during nitrogen deprivation in green algae. Due to the
above-described limitations in reverse genetic approaches in Chlarnydomonas,
development of forward genetic screens for both chemical and insertion mutants that are
207
defective in TAG accumulation or LD morphology will likely play a key role in the
elucidation of the required factors for these proceses.
The work described in this dissertation adds to the wealth of knowledge generated
on plant lipid metabolism and how it is regulated during environmental stress.
Furthermore, it provides insight into lipid responses during stress in green algae, an
understanding of which is likely to play a key role in the development of algal species as
feedstocks for renewable bioenergy.
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