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MEMBRANE LIPID BIOSYNTHESIS IN
CHLA MYDOMONAS REINHARD T11
By

Wayne Russell Riekhof

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Graduate Program in Biochemistry and Molecular Biology

2004

ABSTRACT
MEMBRANE LIPID BIOSYNTHESIS IN
CHLAMYDOMONAS REINHARDTII
By
Wayne Russell Riekhof

Lipids play essential roles in all cellular organisms, most prominently as
components of cellular membranes, which delineate the boundary between the cell and its
environment, and between subcellular compartments. The synthesis of membrane lipids
in the unicellular flagellated alga Chlamydomonas reinhardtii was studied using
biochemical techniques, forward and reverse genetics, and genomics.

The major component of extraplastidic membranes in Chlamydomonas is the
betaine lipid diacylglyceryl-N,N,N—trimethylhomoserine (DGTS). As a model system for
the study of betaine lipid biosynthesis, the DGTS biosynthesis proteins BtaA and BtaB
from Rhodobacter sphaeroides were studied at the biochemical level, and were shown to
be responsible for all reactions of DGTS biosynthesis. Sequencing of the
Chlamydomonas genome revealed a protein containing BtaA- and BtaB- like domains on
the same polypeptide, leading to the hypothesis that this fusion protein, CrBTAl,
catalyzes all reactions of DGTS biosynthesis in this alga. This hypothesis was confirmed
by characterization of the recombinant enzyme, and site directed mutagenesis was used to
identify active site residues in the two domains. In vivo fimctional analysis of the
corresponding gene was carried out by using RNA interference to block gene expression,

which led to pleiotropic effects on grth rate and temperature sensitivity.

A forward genetic screen was carried out in Chlamydomonas by using plasmid
tagging insertional mutagenesis. This led to the discovery of a mutant defective in
synthesis of one of the thylakoid lipids, sulfoquinovosyldiacylglycerol. Taken together,
these results answer some previously unresolved questions regarding membrane lipid
biosynthesis in Chlamydomonas, and establish this organism as a useful complement to
the more thoroughly characterized models of seed-plant lipid metabolism, such as

Arabidopsis.

ACKNOWLEDGMENTS

My time at Michigan State has been the most exciting, stimulating, and interesting
four years of my life, and that fact is due to a great number of wonderful people with
whom I have interacted. First and foremost among them is my thesis advisor, Dr.
Christoph Benning. I owe a great deal to him for giving me a project which was a little
bit outside of the mainstream, allowing me a lot of freedom to roam with it, and
encouraging me to take my ideas and make something of them. I'm also indebted to him
for not charging me for all the broken glassware, dropped jugs of toluene, and similar
blunders I make on a regular basis.

Without exception, all of my co-workers have been wonderful, and coming to
work everyday was always exciting. The Benning lab has been my home away from
home, and my lab mates have been like a second family, so leaving is bittersweet.

The work on Chlamydomonas mutant screening was carried out by a motley crew
consisting of Todd Lydic, Mike Ruckle, and (sometimes) the author, with guidance from
Dr. C. Benning and Dr. Barbara Sears, MSU Department of Plant Biology. I will always
fondly recall the many hours spent standing idly by while we watched the Tecan robot
tirelessly spotting TLC plates. We all also owe a debt of gratitude to the custodial staff
for not emptying the trash on one particular Friday, thus allowing us to recover the
replica plate containing the mutant described in Chapter 5 on the following Monday.

I have had the good fortune to be involved with Dr's. Changcheng Xu and
Koichiro Awai in their studies on lipid trafficking mutants of Arabidopsis (Appendix B),

and I look forward to reading about the continuation of this exciting work after I leave the

iv

lab. Another project that was especially fun and interesting was the annotation of EST
sequences of the red alga Galdieria sulphuraria (Appendix A), conducted in
collaboration with Dr. Andreas Weber and his laboratory. Still another side project with
another experimental organism was conducted in collaboration with Dr. Helmut Bertrand,
MSU Dept. of Microbiology and Molecular Genetics, who I worked with in showing that
Neurospora crassa synthesizes betaine lipid under phosphate stress.

Chapters 3, 4 and 5 make use of data generated by the Chlamydomonas Genome
Project, and the work described in those chapters would have been made much more time
consuming and tedious (if not impossible) if it were not for this data being publicly
available. I was lucky enough to be involved on Christoph’s behalf in the annotation
phase of this project (described in Chapter 4), part of which was done at the “JGI Chlamy
Jamboree” held in December 2003 at the DOE- Joint Genome Institute in Walnut Creek,
CA.

My guidance committee, Drs. Gregg Howe, Bob Hausinger, Doug Gage, and John
Ohlrogge, have been very helpful and supportive over the years, and have given me good
feedback and advice from which I have benefited greatly. I also was given a good start at
MSU in the Plant Research Lab through rotations with Drs. Gregg Howe and Lee
McIntosh.

The work presented in this dissertation was funded by grants from the National
Science Foundation, the MSU Center for Novel Plant Products and Technologies, and
through a fellowship from the DOE-Plant Research Lab that funded my first two years, in

addition to an MSU Plant Science Fellowship which provided travel funds.

Last and most importantly, I want to thank my wife Stephanie, who makes me a
much better person than I would otherwise be, and without whom (according to my
mother, at least) 1 would not have made it in Michigan. I also want to thank my parents,
who recognized and encouraged my love of science from an early age, and have

encouraged me at all stages of my academic career.

vi

TABLE OF CONTENTS

List of Tables .......................................................................................... x

List of Figures ......................................................................................... xi

Chapter 1. Lipid biosynthesis in plants and algae ................................................. 1
Introduction ................................................................................... 2
Composition of plant cell membranes .................................................... 2
Fatty acid biosynthesis in the plastid ...................................................... 5
Incorporation of FA into the plastidic pathway of membrane lipid synthesis ....... 9
Galactolipid synthesis ..................................................................... lO
Sulfolipid synthesis and function ......................................................... 12
Phosphatidylglycerol synthesis and function ........................................... 15
Fatty acid export from the plastid ........................................................ l7
Phosphatidylcholine and phosphatidylethanolamine synthesis ...................... 20
Betaine lipid synthesis ..................................................................... 23
Cellular organization of lipid metabolism: The eukaryotic pathway of thylakoid
lipid synthesis ............................................................................... 27
Phosphate regulation of lipid metabolism ............................................... 30
Chlamydomonas and Arabidopsis as complementary models of plant lipid
metabolism .................................................................................. 31
Development of Chlamydomonas as a model system for lipid metabolism........33
References ................................................................................... 35

Chapter 2. Functional characterization of the bacterial betaine lipid biosynthetic enzymes

BtaA and BtaB ....................................................................................... 44
Abstract ...................................................................................... 45
Introduction ................................................................................. 46
Materials and Methods ..................................................................... 50

Materials ........................................................................... 50
Expression of RsbtaA and RsbtaB ............................................... 50
In vivo production of DGHS and DGTS in E. coli and R. sphaeroides....51
Subcellular localization of BtaA protein ....................................... 53
In vitro activity of RthaA ....................................................... 54
Effect of DAG production by phospholipase C on RthaA activity ....... 55
Results ....................................................................................... 56
RthaA and RthaB are sufficient to reconstitute DGTS biosynthesis in E.
coli ................................................................................... 56
RthaA is associated with membranes ......................................... 58
RthaA has AdoMetzdiacylglycerol 3-amino-3-carboxypropytransferase
activity .............................................................................. 60
BtaA is related to methyltransferases .......................................... 64
Discussion ................................................................................... 69
References ................................................................................... 74

vii

Chapter 3. Functions of the bacterial betaine lipid biosynthetic enzymes BtaA and BtaB

are carried out by the multifunctional polypeptide BTAl in Chlamydomonas

reinhardtii ............................................................................................ 78
Introduction ................................................................................. 79
Abstract ...................................................................................... 79
Materials and Methods ..................................................................... 80

Materials ........................................................................... 80
Cloning and expression of RsbtaA, CrBTAl and site directed mutants in
E. coli ............................................................................... 80
In vivo production of DGHS and DGTS in E. coli ........................... 82
In vitro activity assays of CrBTAl ............................................. 84
Mass spectrometry of DGTS .................................................... 85
Results ....................................................................................... 86
CrBTAl is a multifunctional protein constituting the entire DGTS
biosynthesis machinery in Chlamydomonas reinhardtii ..................... 86
Site directed mutagenesis of the AdoMet binding domains of BTAI ......89
Kinetic analysis of CrBTAl ..................................................... 93
Discussion ................................................................................... 93
References ................................................................................... 99

Chapter 4. Molecular biology of lipid metabolism in Chlamydomonas reinhardtii.

Analysis of the draft genome for pathways of lipid biosynthesis, and prospects for

functional characterization of metabolic pathways by RNA interference ................. 100
Abstract ..................................................................................... 101
Introduction ................................................................................ 102
Materials and Methods ................................................................... 103

Annotation of C. reinhandtii genes involved in lipid metabolism ......... 103
Construction of RNAi lines for suppression of CrB TAI ................... 104
Transformation with RNAi construct and growth of cells .................. 105
RNA blot analysis ............................................................... 106
Analysis of lipids ................................................................ 107
Results ...................................................................................... 108
Annotation of lipid biosynthesis gene in the Chlamydomonas genome..108
Lipid metabolism in plastids ................................................... 108
Lipid metabolism in the endomembrane system ............................ l 15
Fatty acid desaturases ........................................................... 1 16
Silencing of the ETA] gene is detrimental to growth ....................... 1 17
Discussion ................................................................................. 1 24
References ................................................................................. 1 31

Chapter 5. The sulfolipids 2’-0-acyl-sulfoquinovosyldiacylglycerol and

sulfoquinovosyldiacylglycerol are absent from a Chlamydomonas reinhardtii mutant

deleted in SQDI .................................................................................... 136
Abstract .................................................................................... 137
Introduction ................................................................................ 1 3 8
Materials and Methods ................................................................... 139

viii

Growth and labeling of Chlamydomonas reinhardtii cell cultures. . . . l 39

Quantitative lipid analysis ....................................................... 140
Generation and screening of insertional mutants ............................ 141
Identification and molecular characterization of the site of transgene
insertion ........................................................................... 142
Large scale isolation and extraction of compounds ......................... 143
Mass spectrometry ............................................................... 144
Fatty acid analysis ............................................................... 144
NMR spectroscopy ............................................................... 144
Results and Discussion ................................................................... 145
Accumulation of ASQD in C. reinhardtii .................................... 145
Isolation of a sulfolipid deficient insertional mutant of C. reinhardtii...152
CrSQDl clusters with bona fide plant SQDl proteins ..................... 157
The Asqdl mutant shows reduced growth under phosphate limitation and
increased herbicide sensitivity ................................................. 158
The presence of ASQD in lipid extracts of of C. reinhardtii is not an
extraction artifact ................................................................ 162
A hypothesis for the biosynthesis of ASQD in C. reinhardtii. . . . . . . . 1 63
The function of ASQD in C. reinhardn'i ..................................... 167
The identity of CrSQDl and evolution of sulfolipid biosynthesis. . . . . l 69
References ................................................................................. l 72

Appendix A. Analysis of lipid metabolism in a primitive red alga: EST analysis of the
thermoacidophilic red alga Galdieria sulphuraria reveals conservation of central

pathways and a complete pathway for lipid A biosynthesis ................................. 175
Introduction ................................................................................ 1 76
Materials and Methods ................................................................... 176
Results and Discussion ................................................................... 177

Fatty acid biosynthesis .......................................................... 177
Fatty acid catabolism ............................................................ 178
Membrane lipid biosynthesis ................................................... 178
Lipid A biosynthesis ............................................................ 179
References ................................................................................. l 84

Appendix B. Analysis of novel galactolipids accumulating in a lipid trafficking mutant of

Arabidopsis, and a hypothesis for their biosynthesis .......................................... 186
Introduction ................................................................................ l 87
Materials and Methods

Isolation of lipids ................................................................ 188
NMR .............................................................................. 188
Results and Discussion .................................................................. 190
References ................................................................................. 196

ix

LIST OF TABLES

Table 2.1. Strains and plasmids used in this work ............................................. 52
Table 3.1. Strains and plasmids used in this work ............................................. 83
Table 4.1. Identification of reactions diagrammed in Figures 4.1 and 4.2 ................. l l 1

Table 5.1. Comparison of the fatty acid composition of the two sulfolipids and cells... 1 51

Table 5.2. Lipid composition of dw15.l and Asqdl .......................................... 159

LIST OF FIGURES

Figure 1.1. Structure of polar lipid and fatty acids in plants and algae ........................ 3
Figure 1.2. Schematic for plastid membrane lipid synthesis .................................... 6
Figure 13. Synthesis of the 4 major plastid lipid classes ....................................... 7
Figure 1.4. Scheme for fatty acid incorporation into extraplastidic lipids ................... 18

Figure 1.5. Biosynthesis of PtdCho, PtdEtn, and DGTS in extraplastidic membranes. . ..19

Figure 1.6. Schematic representation of lipid transport pathways present in plants. . . .28
Figure 2.1. DGTS biosynthetic pathway and involvement of RthaA/B .................... 48
Figure 2.2. Reconstitution of DGTS biosynthesis in E. coli ................................... 57
Figure 2.3. The RthaA protein is associated with cellular membranes ..................... 59
Figure 2.4. In vitro assay of BtaA activity ....................................................... 61
Figure 2.5. pH dependence and affinity for AdoMet ............................................ 62
Figure 2 ..6 Multiple sequence alignment of bacterial BtaA orthologs and
methyltransferases ................................................................................... 65
Figure 2.7. Schematic representation of BtaA protein and active site ........................ 67
Figure 3.1. Identification of CrBTAl as a fusion of BtaA- and BtaB like domains ....... 87
Figure 3.2. CrBTAl catalyzes the full DGTS biosynthetic pathway ........................ 90
Figure 3.3. Analysis of BtaA- and and BtaB- like domains of BTAl by site directed
mutatgenesis .......................................................................................... 92
Figure 3.4. Biochemical properties of CrBTAl ................................................ 94
Figure 4.1. Pathways of lipid biosynthesis which are known or hypothesized to occur in
Chlamydomonas, and their subcellular localization .......................................... 109
Figure 4.2. Pathways of acyl chain desaturation .............................................. l 10
Figure 4.3. Schematic of RNAi construct ...................................................... l 18

xi

Figure 4.4. Growth curves of RNAi lines and vector controls .............................. 1 19

Figure 45. BTAI gene expression in RNAi lines .............................................. 121
Figure 4.6. Lipid composition of RNAi lines .................................................. 122
Figure 4.7. Temperature sensitivity of RNAi lines ............................................ 123
Figure 5.1 Separation of lipid extracts from 35S-labelled cells of C. reinhardtii wild-type
(dw15.l) and mutant (Asqdl) .................................................................... 146
Figure 52. Mass spectrometric analysis of ASQD ............................................ 149
Figure 5.3. 500 MHz proton NMR spectra of SQDG and ASQD ........................... 150
Figure 5.4. Comparison of genomic sequences surrounding the CrSQDl locus in the
wild-type (WT), the Asqdl mutant, and the rescued plasmid (qudI-R) .................. 154
Figure 55. Comparison of SQDI/SQDB from bacteria, archaea, and plants ............. 155

Figure 5.6. Growth of dw15.l and Asqdl under phosphate-replete and phosphate-limited
conditions ........................................................................................... 160

Figure 5.7. Growth of dw15.1 and Asqdl on agar solidified medium containing different

amounts of the herbicide DCMU ................................................................ 161
Figure 5.8. Hypothesis for ASQD biosynthesis in C. reinhardtii ........................... 164
Figure A.1. Subcellular organization of fatty acid biosynthesis and glycerolipid assembly
in Galdieria sulphuraria .......................................................................... 180
Figure A.2. Lipid A biosynthetic pathway, and ESTs present in the database. . . . . . . l 82
Figure B]. TLC of wild-type and tng-I lipid extracts ..................................... 189
Figure 3.2. NMR of galactolipids isolated from wild-type and mutants .................. 191

Figure B.3. Hypothesis for oligogalactolipid synthesis by a B-galactosidase
mechanism ......................................................................................... 194

xii

Chapter 1

Lipid biosynthesis in plants and algae

Introduction.

The molecular biology and biochemistry of membrane lipid biosynthesis in plants has
been extensively studied over the past several decades, leading to detailed hypotheses
regarding fatty acid synthesis, glycerolipid assembly, fatty acid desaturation, and
trafficking of lipids between membranes. (Somerville et a1., 2000). This chapter will
summarize the salient points of these critical cellular processes with a focus on recent
advances in the isolation of genes and proteins responsible for the biochemical activities,
and will also attempt to show which t0pics are in need of clarification and further study.
Additionally, the lipid metabolism of higher plants (typified by ArabidOpsis thaliana) and
green algae (e .g. Chlamydomonas reinhardtii) will be compared and contrasted, and I
will introduce the questions that have been addressed in the following chapters.
Composition of plant membranes.

The polar lipid and fatty acid composition of plant cells is complex, and is influenced
substantially by the presence of the plastid. This organelle was derived from an
endosymbiotic relationship between a cyanobacterium and the eukaryotic progenitor of
plant cells. As a consequence, most plants cells contain polar lipid species that are
characteristic of eukaryotes, such as phosphatidylcholine and phosphatidylethanolamine,
and also lipids characteristic of cyanobacteria, such as mono- and
digalactosyldiacylglycerol (MGDG and DGDG), and the sulfolipid
sulfoquinovosyldiacylglycerol (SQDG). The chemical structures of these polar lipids, as

well as other species commonly found in plants and algae, are presented in Figure 1.1.

Figure 1.1: Structures of polar lipids and fatty acids in plants and algae. MGDG,
monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; SQDG,
sulfoquinovosyldiacylglycerol, Pthro, phosphatidylglycerol; PtdCho,
phosphatidylcholine; DGTS, diacylglyceryl-N,N,N,-trimethy1homoserine; PtdEtn,
phosphatidylethanolamine; PtdSer, phosphatidylserine; PtdIns, phosphatidylinositol.
Fatty acids, from top down, (carbons :double bondsAWsmms) 16:0 (palmitate), 16:1A3”""‘,
16:1A7(palmitoleate), 16:247'10, 16:3A7"°"3, 18:0 (stearate), 18:1” (oleate), 1822”“

(linoleate), 18:3’39’12’15 (linolenate).

Major extraplastidic lipids Major plastidic lipids
Ptdlns 0

MGDG

Onyx/6R2 V6

" O . 0 R2

0 O\< 1 O ck<°
PtdEtn 1

 

Among those molecules not present in seed—plants are the betaine lipids, such as
diacylglyceryl-N,N,N-trimethylhomoserine (DGTS), and diacylglyceryl-N,N,N-
trirnethylhydroxymethy-B-alanine (DGTA). These lipids have been hypothesized to take
on the functions of PtdCho in species that synthesize them (Sato, 1992), and the
biosynthetic pathways involved in their synthesis will be discussed in detail.

The fatty acid complement of plant cells is also complex (Figure 1.1). The bulk of
fatty acids present in plant cell membranes consist of saturated, monounsaturated, and
polyunsaturated 16 and 18 carbon species, the unsaturated forms being derived from the
saturated chains by the action of soluble and membrane-bound fatty acid desaturases.
While the bulk of membrane fatty acids are made up of relatively simple linear chains,
other species, e.g. cyclopropene, hydroxyl, and epoxide containing derivatives are found

to occur predominantly in the seed oil of some species.

Fatty acid biosynthesis in the plastid.

Figure 1.2 gives a schematic representation of fatty acid biosynthesis and membrane lipid
assembly in the plastid. The plastid is known to be the major site of fatty acid synthesis
in plant cells (Ohlrogge and Browse, 1995) and most of the genes encoding enzymes
involved in plastidic fatty acid synthesis have been isolated and studied. Fatty acid
synthesis in the plastid occurs via a system very similar to that of bacteria. As the
committed step in the pathway, acetyl-CoA is carboxylated to form malonyl-CoA by the
action of acetyl-CoA carboxylase. The predominant plastidic isoform of ACCase is a
multimeric bacterial-type enzyme consisting of four proteins, including biotin

carboxylase, biotin carboxyl carrier protein, a-carboxyltransferase, and B-

m <— PthroP

m m
uoP Gal ’1 Limp- so 4— UDP Glc (3-3-p

 

 

m DAG — PtdOH — coP DAG
Aoetyl-CoA
UDP-Gal /
Malonyl—CoA
Iyso-F’tdOH A
Malonyl ACP ACP
Plastid
Acyl—ACP/
lMS
Cytosol

Figure 1.2: Schematic for plastid membrane lipid synthesis. Fatty acids are produced
in the plastid and incorporated into the 4 plastid lipids as outlined in this scheme.
PthroP, phosphatidylglycerolphosphate; UDP—Gal, uridine-S’-diphosphate-galactose;
DAG, diacylglycerol; UDP-Glc, uridine-S’-diphosphate-glucose; UDP-SQ, uridine-5’-
diphosphate-sulfoquinovose; CDP-DAG, cytidine-S’-diphosphate-diglyceride; G-3-P,
glycerol-3-phosphate; PtdOH, phosphatidic acid; ACP, acyl-carrier protein; IMS,

intermembrane space.

PAP

 

DAG < PtdOI-l
cos
MGD1l(UDP-Gal) (cm
COP-DAG
MGDG
PGP1l(Gro—3-P)
DGD1l(UDP-Gal) l
DGDG
Pthro
8001 3002
UDP-Glc ——-> UDP-SQ —-> SQDG
(sulfite) (DAG)

Figure 1.3: Synthesis of the 4 major plastid lipid classes. Major biosynthetic pathways
for the four major plastid lipids, as described in the text. MGDl, MGDG synthase;
DGDl, DGDG synthase; PGPl, phosphatidylglycerophosphate synthase; SQDl, UDP-

SQ synthase, SQD2, sulfolipid synthase.

carboxyltransferase subunits. ACCase is regarded as a control point in initiation of fatty
acid biosynthesis, and is under multiple regulatory controls (Somerville et al., 2000).

Malonate can be transferred from CoA to a small phosphopantetheine- containing
polypeptide called acyl carrier protein (ACP). This reaction is catalyzed by malonyl-
CoA: ACP transacylase, and the protein from Chlamydomonas has recently been
identified in a proteomic screen for thioredoxin interacting proteins (Lemaire et al.,
2004). This may indicate that its enzymatic activity is under the control of the plastidic
thioredoxin system, which would probably inhibit its activity through oxidation of a
disulfide bridge in the dark, when energy charge is low. Conversely, it would be
activated in the light during optimal growth conditions by reduction of this putative
disulfide bridge, and this cycle would represent another layer of regulation in fatty acid
biosynthesis.

After synthesis of malonyl-ACP, fatty acid synthesis in plastids continues by the
action of a type II (bacterial) multimeric fatty acid synthase, located in the chloroplast.
Using acyl-ACP and malonyl-ACP as substrates, the first component of this complex is
3-ketoacyl-ACP synthase, which catalyses a Claisen condensation of the malonyl and
acyl substituents, releasing carbon dioxide as a byproduct, and forming a carbon-carbon
bond to increase the chain length by two carbons. The next steps involve reduction of the
3-keto group, elimination of water from the resulting 3-hydroxy acyl species, and finally
reduction of the enoyl-ACP species by the activities of 3-ketoacyl-ACP reductase, 3-
hydroxyacyl-ACP dehydratase, and enoyl-ACP reductase (Somerville et al., 2000). This
sequence of reactions is repeated, resulting in the elongation of the growing fatty acid

chain by two carbons each cycle until the chain reaches 16 or 18 carbons. At this point,

the 18 carbon species, stearoyl-ACP, can be acted upon by a soluble fatty acid desaturase,
the strearoyl-ACP A9 desaturase (Shanklin and Somerville, 1991) giving oleoyl-ACP as a
product.

The long chain fatty acyl end products (predominantly palmitoyl-ACP and oleoyl
ACP) are then partitioned into one of two major pathways of polar lipid biosynthesis. The
acyl groups either become part of the plastidic membrane lipids by the action of the
plastidic glycerol-3-phosphate: acyl—ACP acyltransferase (GPAT, (Murata and Tasaka,
1997)) and lyso-phosphatidate acyltransferase (LPAAT, (Kim and Huang, 2004; Yu et
al., 2004)), or are directed to the cytosol by the action of fatty acyl-ACP thioesterases. In
the former pathway, fatty acids are retained in the plastid and used for plastidic
membrane lipid synthesis. In the latter pathway, free fatty acids are released by the
action of a thioesterase, and these are channeled across the inner and outer envelopes of
the chloroplast and incorporated into acyl-CoA by the action of a long chain acyl-CoA

synthetase (Koo et al., 2004).

Incorporation of FA into the plastidic pathway of membrane lipid synthesis.

As described above, the acyl-ACPs produced from plastidic fatty acid synthase may
become substrates for the plastidic GPAT and LPAAT enzymes. GPAT has been
isolated from a number of species (Murata and Tasaka, 1997) and shown to prefer oleoyl-
ACP as a substrate. Mutations in the Arabidopsis plastidic GPAT (encoded by the
ACT1(A T SI ) gene) give rise to plants that are unable to synthesise diacylglycerol
moieties in the plastid, thus forcing an increase in flux through the so called “eukaryotic”

pathway of thylakoid lipid synthesis (Kunst et al., 1988), which will be discussed below.

The second step in this sequence is the LPAAT (ACT2), and this enzyme shows higher
affinity for palmitoyl-ACP (Bourgis et al., 1999; Kim and Huang, 2004). Interestingly,
GPAT (actI) mutants are viable and grow essentially like wild type, presumably due to
the increase in flux through the eukaryotic pathway, whereas null alleles of the LPAAT
(ars2) gene cause an embryo lethal phenotype (Kim and Huang, 2004; Yu et al., 2004).
The combined substrate specificities of the two plastidic acyltransferases result in a
diglyceride sn 1/sn2 molecular species preference of 18:1/16:0, and this is the
predominant phosphatidic acid (PtdOH) species initially produced by the plastidic or
“prokaryotic” pathway. PtdOH produced by this sequence can then be converted to
CDP-diglyceride and used directly for phosphatidyl glycerol (Pthro) biosynthesis, or can
be dephosphorylated by the action of phosphatidic acid phosphatase, forming DAG as a

substrate for galactolipid and sulfolipid biosynthesis.

Galactolipid synthesis.

The most abundant membrane lipid in the biosphere is thought to be MGDG (Figures 1.1
and 1.3) (Dormann and Benning, 2002), which makes up about 50% of the lipid
component of the thylakoid membrane in most oxygenic photosynthetic organisms. The
biochemical activity responsible for synthesis of MGDG was characterized as a
component of the chloroplast inner envelope membrane, and a thorough biochemical
characterization was carried out on partially

purified fractions from spinach (Marechal et al., 1994). Cloning of the gene encoding
MGDG synthase was accomplished by purifying the enzyme from expanding cucumber

leaves, partial sequencing of the protein, and using this information to amplify a fragment

lO

of the gene by degenerate PCR (Shirnojirna et al., 1997). Sequence analysis revealed that
MGDG synthase is similar to the bacterial UDP-N-acetylglucosaminyl transferase MurG,
and that, by analogy with the MurG mechanism, MGDl condenses UDP-galactose
(produced by the action of UDP-glucose epimerase, (Dormann and Benning, 1996)) and
diacylglycerol with inversion of anomeric configuration (e.g. producing a B-galactoside
linkage). Subsequent studies in Arabidopsis identified two additional MGDG synthase
isoforms that are regulated strongly by phosphorus deprivation (Awai et al., 2001), and
these will be discussed later in the section on phosphate regulation of lipid biosynthesis.

The next most abundant lipid in the plastid is derived from MGDG by addition of
a second galactose moiety. DGDG (Figures 1.1 and 1.3) is a bilayer—forming lipid. That
is, when suspended in water, it tends to form a larnellar phase due to the packing together
of roughly cylindrical adjacent monomers. In contrast, MGDG is a non-bilayer forming
lipid, and its roughly conical shape allows pure preparations to attain inverted micellar
phases in aqueous solution (Sprague and Staehelin, 1983). The bilayerznonbilayer ratio
of biological membranes is typically under tight control (Vikstrom et al., 1999), therefore
it follows that the enzyme which forms DGDG from MGDG is a major control point in
determining this ratio in the thylakoid membranes, therefore determining a basic
biophysical property of the system.

The older literature on DGDG synthesis is mainly focused on the formation of
DGDG by a galactolipidzgalactolipid galactosyltransferase (GGGT) mechanism, by
which the galactose moiety of one MGDG is transferred to the 6’ hydroxyl of a second
MGDG molecule, with inversion of anomeric configuration (e .g. (Heemskerk et al.,

1988; van Besow and Wintermans, 1978)). This “classical” mechanism is at odds with

11

recent findings based on the cloning and characterization of DGDG synthase isoforms
from Arabidopsis.

A brute force biochemical screen involving TIC of lipids extracted from
mutagenized plants was used to isolate a line, designated (1ng , with a 90% reduction in
the amount of DGDG present in membranes (Dormann et al., 1995). Positional cloning
was used to isolate the DOD! gene, and the activity of its product was confirmed in E.
coli by co-expression of the MGDI gene from cucumber, which led to synthesis of
DGDG in E. coli. (Dormann et al., 1999). Although originally thought to be the GGGT,
DGDl is characterized by a glycosyltransferase motif in its C-terminal domain, implying
that it uses UDP-galactose as a substrate in a retaining glycosyltransferase reaction. The
role of the N-terminal domain of DGDl is unknown, but may have functions in lipid
trafficking (Dormann et al., 1999).

Since the mutation in dng is a null allele, the presence of DGDG at 10% of the
wild-type level was interpreted to mean that there is a second, DGDl independent
pathway of DGDG biosynthesis. Further studies identified this pathway as inducible by
phosphorus starvation (Hartel et al., 2000), and the enzyme responsible has recently been
identified as DGD2 (Kelly and Dorrnann, 2002). This aspect of galactolipid synthesis

will be discussed further below.

Sulfolipid synthesis and function.
The plant sulfolipid, SQDG (Figure 1.1), was described by Benson and co-workers nearly

50 years ago (Benson et al., 1959). As a component of the thylakoid membrane, it is one

12

of the most abundant sulfonate compounds in the biosphere, and much effort has been
dedicated to elucidating the molecular biology and biochemistry of its biosynthetic
pathway (Benning, 1998). Beginning with studies on Rhodobacter sphaeroides and
cyanobacteria, (Benning and Somerville, 1992a; Benning and Somerville, 1992b; Giiler
et al., 1996; Giiler et al., 2000), and later Arabidopsis (Essigmann et al., 1998; Sanda et
al., 2001; Yu et al., 2002) and spinach (Shimojima and Benning, 2003), it became
apparent that a sugar—nucleotide pathway is operative in which UDP-glucose and sulfite
act to form UDP sulfoquinovose (Pugh et al., 1995; Sanda et al., 2001), followed by
transfer of the sulfoquinovose moiety to diacylglycerol (Giiler et al., 2000; Yu et al.,
2002). This pathway is diagrammed in Figure 1.3.

The fimction of SQDG in photosynthetic reactions is still a contentious topic. It is
clear that sulfolipid deficient mutants of Rhodobacter, Synechococcus, and Arabidopsis
are photosynthetically competent, and present very little difference in photosynthetic
parameters relative to the respective wild-types, (Benning et al., 1993; Giiler et al., 1996;
Yu et al., 2002). However, a mutant of Synechocystis PCC 6803 in which the .9qu gene
is insertionally inactivated behaves as a sulfolipid auxotroph, indicating that SQDG is an
essential factor for growth of this species (Aoki et al., 2004). This leads to the conclusion
that there is no universal requirement for SQDG in photosynthesis, but that it is
conditionally important in a species-specific manner. This is also Due of photosynthetic
eukaryotes, such as Chlamydomonas, as discussed below.

The hj2 mutant of Chlamydomonas was isolated in a screen for strains showing
high chlorophyll fluorescence, and was later shown to be deficient in the biosynthesis of

SQDG (Sato et al., 1995b). Further characterization of this mutant has shown that it has

13

defects in the heat stability of PSII (Sato et al., 1995a; Sato et al., 2003a), and that it is
more sensitive to the diuron herbicide DCMU, a competitive inhibitor of the Q3 site of
PSII. Taken together, these data led to the conclusion that SQDG is involved in
maintaining the stability of PSII.

In addition to P81], a recent X-ray structural study of the cytochrome b4 complex
of Chlamydomonas reveals sulfolipid in a tight association with the cytochrome f subunit
(Stroebel et al., 2003). In this structure, the sulfonate group forms a salt bridge with a
lysine residue that was shown to be critical for the co-translational insertion of this
polypeptide into the thylakoid membrane (Choquet et al., 2003). Therefore, in
Chlamydomonas, sulfolipid has been implicated in the function of two of the core
components of the photosynthetic electron transport chain.

Another function of SQDG is in maintenance of the anionic character of the
thylakoid during phosphate deprivation (Benning, 1998). SQDG and the other anionic
lipid of the thylakoid, phosphatidylglycerol, are regulated in a reciprocal relationship
such that the net negative charge on the membrane is kept constant (Yu and Benning,
2003). This compensatory mechanism will be explored below.

Chapter 5 gives the results of a forward genetic screen for mutants of
Chlamydomonas with defects in lipid metabolism. Although a major focus of the work
presented here is on the elucidation of DGTS biosynthesis in Chlamydomonas (see
below), the most tractable line in which there were obvious differences in polar lipid
content was an SQDG deficient mutant. This line also lacked a previously unknown 2'-
O-acylated derivative of SQDG. Chapter 5 presents the structural elucidation of this

novel species, the characterization of the mutation in the SQDG deficient line as a

14

deletion of SQDI , and the functional consequences of SQDG deficiency on the growth of

Chlamydomonas.

Phosphatidylglycerol synthesis and function.

As mentioned above, Pthro is one of the two anionic phospholipids in the plastid, and
along with SQDG, it acts to maintain the negative charge density and proper function of
this membrane. Two PGP synthases (Figure 1.3) have been cloned from Arabidopsis and
biochemically characterized (Muller and Frentzen, 2001), and recent years have seen
several studies that show a direct link between Pthro deficiency and abnormal plastid
development and photosystem function (Xu et al., 2002; Gombos and Murata, 1998;
Hagio et al., 2002; Babiychuk et al., 2003). These studies have been based on a mutant of
Arabidoposis,pg21-I , which has an 80% reduction in the PGP synthase activity (Figure
1.3), and on T-DNA insertional mutants, which represent null alleles. Studies on the
pgpI -1 mutant showed that the pigment composition was altered, and that the plants were
generally smaller and less photosynthetically compentent than wild type (Xu et al., 2002).
Studies on T-DNA alleles of the PGP] gene have shown that its function is critical to
photosynthetic growth and plastid development, as the null mutant of this gene requires
sucrose for growth, and has severe defects in the organization of the photosynthetic
membranes (Babiychuk et al., 2003; Hagio et al., 2002). Additionally, PGPl has been
shown to be targeted to both plastids and mitochondria (Babiychuk et al., 2003). While
PGPl is essential for plastid function, the mitochondrial role appears to be redundant, as

the mitochondria in the jovtenky (pgpI) mutant are capable of forming Pthro and

15

cardiolipin, and there are no great dysfunctions in the mitochondrial electron transport
chain.

Pthro is unique among chloroplast lipids in that it contains a 16:1A3tralns acyl
species (Figure 1.1). This particular Pthro molecular species has been implicated in
maintaining the stability of the P811 complex of Chlamydomonas, and the photosynthetic
defects of the mutant lacking Pthro:16: 1A3""""s can be partially rescued by culturing this
strain in the presence of liposomes containing this lipid (Dubertret et al., 1994). Analysis
of these mutants, mfl and mf2, also implies a role for Pthro in the co-translational
insertion of PSII core polypeptides into the photosynthetic membrane (Pineau etal.,
2004), however the pleiotropic nature of these mutations calls into question whether a
Pthro biosynthesis gene or fatty acid desaturase is involved, or whether the lipid
phenotypes are an indirect result of some other lesion in a gene required for optimal
photosynthesis.

As alluded to above, Pthro and SQDG are linked by their anionic character, and
they exhibit an inverse relationship in terms of quantity. For example, in the .9qu mutant
of Arabidopsis, which lacks SQDG, Pthro content is maintained at a level so as to
replace the lost sulfolipid (Yu et al., 2002). Likewise, under phosphate deprived
conditions, Pthro levels drop, and SQDG biosynthesis is up-regulated to compensate
(Essigmann et al., 1998; Yu et al., 2002). It follows that a double mutant with lesions
affecting both pathways would lack this compensatory ability, and would be
compromised in growth. This was shown to be the case in a study (Yu and Benning,
2003) in which a double mutant of squ and pgplwas constructed by repeated back-

crossing (to normalize differences between the W5 and Col-O ecotypes) to create near-

16

isogenic lines segregating the wild-type, single, and double mutants. The double mutant
plants were severely compromised in autotrophic growth, and were hypersensitive to the
PSII inhibitor DCMU. Additionally, the single mutants of pgpI and squ showed
increased sensitivity to DCMU, implying that, while SQDG is not required for
photosynthetic growth and shows no growth phenotype under laboratory conditions, there
may be subtle changes in the organization of PSII that render it more sensitive to this

herbicide.

Fatty acid export from the plastid.

The section on fatty acid biosynthesis above gave a brief overview of the pathway, but
another critical step in plant lipid metabolism is the export of fatty acids and their
incorporation into extraplastidic membrane lipids such as PtdCho, PtdEtn, and in certain
non-seed plants, the betaine lipids, such as DGTS. Hypotheses regarding the steps in
acyl-chain export from the plastid have been tested using in vivo 180 labeling (Pollard
and Ohlrogge, 1999). This study showed that fatty acids produced in the chloroplast are
incorporated into plastidic lipids by direct acyl-transfer from acyl-ACP, but that those in
the extraplastidic membranes show a 50% reduction in 180 content. This result is
interpreted as being consistent with the model that the free fatty acids are released via
hydrolysis in the plastid, and transported out to the site of “eukaryotic” lipid synthesis.
At least two thioesterases in Arabidopsis, FatA and FatB, mediate the hydrolysis of
predominantly 18:1 and 16:0 ACPs, respectively (Salas and Ohlrogge, 2002). To further

elucidate the mechanism of fatty acid export from the plastid, (Koo et al., 2004) used a

17

Acyl-ACP
Plastid
FreeFA
IMS

C\

ytosol

 

m

Glc—1-P

c... 1
lns-3-P
Met Iyso—PtdOH 4——)—G-3-P

l t r-—
MOMOI PIdOH -’ COP-DAG
t
o—L—DAG PthroP

P-Etn —> CDP-Etk>-
t
Etn \

A P-Cho —> CDP-Cho —>
Ser

‘FA
1

E

Figure 1.4: Scheme for fatty acid export and incorporation into extraplastidic lipids.
IMS, intermembrane space; ACP, Acyl carrier protein; FA, fatty acid; PtdOH,
phosphatidic acid; CDP-DAG, cytidine 5’-diphosphate-diacylglycerol; Met, methionine;
AdoMet, S-adenosylmethionine; DGTS, diacylglyceryl-N,N,N-tn'methylhomoserine;
DAG, diacylglycerol; G-3-P, g1ycerol-3-phosphate; Pthro, phosphatidylglycerol;
PthroP, phosphatidylglycerophosphate; Glc-l-P, glucose-l-phosphate; Ins-3-P, myo-
inositol-3-phosphate; Ptdlns, phosphatidylinositol; Ser, serine; Etn, ethanolamine; P-Etn,
phosphoethanolamine; CDP-Etn, cytidine 5’-diphosphate-ethanolarnine; CDP-Cho,
cytidine 5’-diphosphate-choline; PtdCho, phosphatidylcholine; PtdEtn,

phosphatidylethanolamine.

l8

PtdCho

PtdEtn
EPT T(DAG) T CPT (DAG)
CDP-Etn CDP-Cho

ECTT T OCT
PEMT

P-Etn ———> P-Cho

EKI T
Etn
soc f
Ser
DAG 1R DGTS O
HO‘SVK/ BtaB | fifiz
\ + o 0 o
Wis , 9
R1 (AdoMet) (3 AdoMet) o 0.
Figure 1.5: Biosynthesis of PtdCho, PtdEtn and DGTS 1n the extraplastidic
membranes. Chemical species are identified in bold, and correspond to the
abbreviations in Figure 1.4. SDC, serine decarboxylase; EKI, ethanolamine kinase; ECT,
ethanolamine phosphate cytidylyltransferase; EPT, CDP-EtnzDAG
ethanolaminephophotransferase; PEMT, phosphoethanolamine methyltransferase; CCT,
phosphocholine cytidylyltransferase; CPT, CDP-ChozDAG cholinephosphotransferase;
BtaA, AdoMetzDAG 3-amino-3-carboxypropyltransferase (diacylglycerylhomoserine

synthase); BtaB, diacylglycerylhomoserine N-methyltransferase.

l9

labeling approach to determine whether fatty acids simply diffuse out of the plastid, or
whether there is a specific transport mechanism. The results indicate that the fiee pool of
fatty acid destined for export has a very short half life, on the order of <1 second, and that
there is a specific channeling of this pool to an acyl-CoA synthetase (ACS). This final
step in export, activation of the free fatty acid to an acyl-CoA, has been studied by
analysis of the Arabidopsis ACS gene family (Schnurr et al., 2002). This study showed
that the LACS9 isoform is likely to be the major activity in developing seeds and
expanding leaves, where the fatty acid synthesis rate is high. The protein is localized to
the chloroplast envelopes, consistent with its proposed function, however a T-DNA
insertional mutant showed essentially no effect on the composition of leaf lipids. Given
that there are multiple members of this ACS family, there is probably considerable
functional redundancy. Central metabolic pathways such as fatty acid synthesis and
export from the plastid are likely to be conserved among Arabidopsis and
Chlamydomonas, as evidenced by, for example, the similarity between Arabidopsis and

Chlamydomonas (0-6 fatty acid desaturases (Sato et al., 1997).

Phosphatidylcholine and Phosphatidylethanolamine synthesis.

The phospholipids PtdCho and PtdEtn are the most abundant lipids in yeast and
mammalian cells, and much is understood about their biosynthesis and function. In
Saccharomyces cerevisiae, there are two pathways of biosynthesis for PtdCho, referred to
as the methylation pathway, and the CDP-choline or Kennedy pathway. In the
methylation pathway, phosphatidylserine is produced in the ER by the action of PtdSer

synthase (Kohlwein et al., 1988) and then transported to the sites of the two PtdSer

20

decarboxylases (PSD) for conversion into PtdEtn. The two PSD activities are encoded by
the PSD] gene, encoding the mitochondrial isoform, and the PSDZ gene, which specifies
the golgi/vacuolar activity (Trotter et al., 1993; Trotter et al., 1995). The PtdEtn resulting
from PSD action is then transferred back to the ER for PtdCho production by N-
methylation via the PtdEtn methyltransferases Pemlp/ChoZp (catalyzing the first
methylation) and Pem2p/Opi3p (catalyzing the final two methylations) (Howe and
McMaster, 2001).

The Kennedy or CDP—Cho pathway is operative when yeast are fed Cho, but
under “normal” circumstances, the flux into this pathway is only a minor component of
PtdCho biosynthesis. This pathway involves phosphorylation of Cho, forming
phosphocholine (P-Cho), activation of P-Cho with CT P to form CDP-Cho, and finally
condensation of CDP-Cho with DAG to form PtdCho (Howe and McMaster, 2001). The
relative flux into the two pathways is largely dependant on the supply of choline,
however other factors such as temperature (Howe et al., 2002) and inositol supply
(Loewen etal., 2004) also play roles in the regulation of these pathways.

In plants, PtdCho is important as an intermediate in the eukaryotic pathway of
thylakoid lipid synthesis (Figure 1.6), and also in triacylglycerol biosynthesis in seeds
(Roughan and Slack, 1982). As in yeast, multiple pathways of PtdCho synthesis are
operative in plant cells, however the CDP-Cho pathway seems to be the major route to
synthesis of PtdCho, with methylation of PtdEtn being almost unknown in plants
(Somerville et al., 2000). Plants, however, have the capability to produce Em and Cho
directly for use as precursors. Unique to plants is a serine decarboxylase, which allows

the production of Etn from Ser (Rontein et al., 2001). The Etn may then be

21

phosphorylated by an ethanolamine kinase, followed by N-trimethylation of the phospho-
Etn to form P-Cho (the pathways outlined in Figure 15). The gene encoding the
trimethylation activity in this pathway was isolated from spinach and Arabidopsis by
cDNA complementation of Schizosaccharomyces pombe ch02 and Saccharomyces
cerevisiae opi3 mutants, respectively (Bolognese and McGraw, 2000; Nuccio et al.,
2000).

The control of PtdCho has been studied in mammalian cells, and it is widely
regarded that PtdCho synthesis is regulated at the level of formation of CDP-Cho in most
tissues. The major exception to this is in hepatic tissues, where formation of PtdCho by
the PtdEtn methylaan is the major route of synthesis (Vance et al., 1997). The major
isoform of CDP-Cho synthetase is the CT-a isoform, and its transcription is regulated
coordinately with the cell cycle (Golfrnan et al., 2001). The CT-a protein is also
regulated in a multivalent manner by phosphorylation (Arnold et al., 1997) and reversible
membrane binding. This last point has been the subject of some very elegant
biochemistry by Cornell and co-workers. It has been shown that membrane binding
activates CT -a (Cornell, 1991), and that the membrane binding is stimulated by non-
bilayer lipids in the membrane (Davies etal., 2001). This leads to the conclusion that
CT-a is itself a sensor of membrane bilayer/non-bilayer lipid ratios, in much the same
way that other biochemical pathways are allosterically regulated by their end products.
While the information on control of yeast and mammalian PtdCho synthesis is far from
complete, it is still much more than is known of the analogous regulatory processes in

plants.

22

Betaine lipid synthesis.

Another group of polar glycerolipids, the betaine lipids, consists of diacylglycerol bound
in an ether linkage to a quaternary amine alcohol. These lipids are widely distributed
among marine and freshwater algae, mosses, ferns, and other non-vascular plants, as well
as among many fungal species (Eichenberger, 1982; Sato and Furuya, 1984; Vaskovsky
et al., 1998). The structure of one member of this class, DGTS, is given in Figure 1.1.

The quaternary amine containing, zwitterionic betaine lipids are similar in overall
structure to PtdCho, and it has been postulated that betaine lipids such as DGTS can
functionally replace PtdCho in membranes (Sato, 1992). Evidence for this hypothesis is
given by the fact that organisms that contain betaine lipids, such as Chlamydomonas,
often lack PtdCho. Further correlative evidence comes from studies on Rhodobacter
sphaeroides which, when deprived of phosphate, decreases the amount of PtdCho and
other phospholipids in its membranes and induces de novo synthesis of DGTS and other
nonphosphorous lipids (Benning et al., 1995).

This last point is the key observation that spurred work on bacterial DGTS
biosynthesis. The initial discovery of phosphate-deprivation induced production of
DGTS in Rhodobacter was made in the context of studies to determine the function of
SQDG (Benning et al., 1993). In general, the complement of polar lipids in Rhodobacter
was observed to drastically change under phosphate limited growth conditions, and
several unidentified non-phosphorous lipids, one of which was later shown to be DGTS
(Benning et al., 1995), were produced concomitantly with a decrease in the amount of
phospholipids. A similar scenario has been shown to occur in several marine

pseudomonad species (Minnikin et al., 1972; Minnikin et al., 1974).

23

Studies following up on the discovery of DGTS in Rhodobacter focused on
defining the pathway by which this lipid is synthesized (Hofmann and Eichenberger,
1996), identifying the genes encoding enzymes of DGTS biosynthesis (Klug and
Benning, 2001) and defining the regulatory elements that control the induction of DGTS
accumulation in Sinorhizobium meliloti (Geiger et al., 1999). The DGTS biosynthetic
pathway has also been studied in the eukaryotic algae Ochromonas danica (Vogel and
Eichenberger, 1992) and Chlamydomonas reinhardtii (Moore et al., 2001; Sato, 1988),
and was proposed to follow the scheme outlined in Figure 1.5. DGTS biosynthesis in
Rhodobacter was also postulated to follow this pathway based on labeling studies
outlined in Hofrnann and Eichenberger ( 1996).

Geiger et al. (1999) showed that phosphate stress-induces DGTS production in
Rhizobium meliloti, and that this induction is dependent on phoB which, in E. coli,
encodes the response regulator of the two-component system controlling expression of
genes involved in phosphate acquisition and utilization under limiting conditions
(Wanner, 1993). This result showed that DGTS accumulation is a function of the pho
regulon, but the regulated genes encoding the proposed functions were not known at that
time. Studies described by Klug and Benning (2001) allowed the tentative placement of
two enzymes into the biosynthetic scheme, as outlined in Figure 1.5.

Briefly, screening of a mutagenized population of Rhodobacter grown under
phosphate limitation yielded several strains lacking the ability to accumulate DGTS, and
complementation cloning of one of the mutants revealed an operon consisting of two
open reading frames, designated btaA and (MB. The product of btaA was proposed to be

the S-adenosylmethionine (AdoMet) dependent 3-amino-3-carboxypropyl transferase

24

(see Figure 1.5), on the basis that over-expression of btaA alone leads to accumulation of
the precursor diacylglycerylhomoserine (DGHS), while disruption of the gene in the
RKL3 mutant abolishes DGTS (and DGHS) production. This function represents an
unusual reaction of AdoMet, which is primarily a donor of methyl groups, however
similar reactions have been noted in the biosynthesis of the antibiotic nocardicin A
(Reeve etal., 1998), and the plant siderophore nicotianamine (Ling et al., 1999). The
braB gene product is homologous to AdoMet-dependent methyltransferases, and
insertional disruption of the open reading frame leads to accumulation of DGHS under
phosphate limitation, hence the function of this protein is likely the catalysis of one or
more of the three methyl group transfers outlined in Figure 15. It is worth noting that the
analogous methyltransferase involved in PtdCho biosynthesis in Rhodobacter, encoded
by the pmtA gene (Arondel et al., 1993), is responsible for all three methylation events
leading to the formation of PtdCho from phosphatidylethanolamine (PtdEtn), so it is
feasible that the BtaB protein acts in a similar manner.

As mentioned above, the unicellular chlorophyte alga Chlamydomonas reinhardtii
has previously been shown to contain DGTS (Eichenberger and Boschetti, 1977) and to
lack PtdCho (Giroud et al., 1988). Aside from the replacement of PtdCho by DGTS, C.
reinhardtii has a complement of polar lipids quite similar to higher plants, and the
positional fatty acid analysis of individual lipid classes indicates that the eukaryotic
pathway of glycerolipid synthesis is involved only in assembly of extraplastidic
membranes, i.e. there is no transfer of diacylglycerol species from the endoplasmic
reticulum to the chloroplast (Giroud et al., 1988). This lack of a eukaryotic complement

of thylakoid molecular species contrasts with the situation in grasses and dicots such as

25

spinach and Arabidopsis, and correlates well with a lack of PtdCho, as this lipid is
thought to be important in lipid trafficking from the endoplasmic reticulum to the plastid
((Somerville et al., 2000); see Fig 1.6 below). Studies using [MG-methionine feeding
(Sato, 1988) and in vitro analysis of crude microsomal preparations (Moore et al., 2001)
indicated that the pathway for biosynthesis of DGTS is consistent with that given in
Figure 1.5. At the time of those studies, no genes or proteins specifying DGTS
biosynthesis had been isolated from any eukaryote, however a detailed analysis of the
fatty acid distribution in different lipids of C. reinhardtii presented strong evidence for an
extraplastidic origin for this lipid (Giroud et al., 1988), consistent with exclusion of the
biosynthetic activities from chloroplast membrane fractions (Moore et al., 2001).

The core focus of the work presented in the following chapters was to
biochemically characterize the BtaA and BtaB enzymes from Rhodobacrer, and to
identify the cognate eukaryotic activities. Chapters 2 and 3 present the biochemical data
addressing these points, and to this end the work was successful. Chapter 2 details the
functional analysis of BtaA and BtaB, details the development of an assay system for
BtaA, and proposes a model for the active site based on similarity to methyltransferases.
Chapter 3 follows with the discovery that the DGTS biosynthetic machinery in
Chlamydomonas (BTAl) is a multidomain fusion protein of BtaA- and BtaB- like
modules. Site-directed mutagenesis is also used to identify residues that are involved in
substrate binding by the individual domains, by analogy with the models proposed for

bacterial BtaA and BtaB.

26

Chapter 4 follows the biochemical data with an in vivo functional analysis, using RNAi to
reduce the accumulation of ET A] transcript and examine the effects of down-regulating

the machinery required for the bulk of extraplastidic membrane synthesis.

Cellular organization of lipid biosynthesis: The eukaryotic pathway of thylakoid
lipid synthesis.
Studies of lipid metabolism in plants in late 19705 and early 1980s revealed that there are
both eukaryotic and prokaryotic pathways of lipid biosynthesis involved in the
production of thylakoid lipid species, as reviewed by Roughan and Slack (1982) and
presented schematically in Figure 1.6. This scheme was proposed to involve the export
of fatty acid from plastids and their incorporation into PtdCho in the endomembrane
system, followed by the import of the diacylglycerol moiety of PtdCho back to the
plastid, and its incorporation into plastidic glycerolipid species. Further studies in
Arabidopsis by Browse, Somerville, and co—workers showed that the flux into the
prokaryotic and eukaryotic pathways of thylakoid biogenesis is approximately equal in
this species (Browse et al., 1986). As discussed above, corroborating evidence for these
pathways was obtained from studies on the act} mutant (Kunst et al., 1988), which is
deficient in the acyl-ACP dependent GPAT activity, the first step in the prokaryotic
pathway. The actual identity of the species being transported back to the plastid is still
controversial, but strong evidence points to the role of PtdCho as the donor of this
species.

The situation in Chlamydomonas is not as convoluted, in that this organism

contains DGTS in place of PtdCho, as described above, and this is in agreement with the

27

[net-re <- PthroP

Acetyl-CoA

Malonyl-CoA

Malonyl-ACP ACP

 

 

I
Cytosol ._
V
FreeFA
COP-Cho —> I
AcleoA
lyso-PtdOH G-3-P
P-Cho PtdOH
DAG

Figure 1.6: Schematic representation of lipid transport pathways present in plants.
Abbreviations for the metabolites are the same as Figures 1.3 and 1.5. Export of fatty
acid fiom the plastid is hypothesized to take place by direct channeling of free fatty acid
hydrolyzed fiom acyl-ACP to an acyl—CoA synthetase in the outer envelope membrane.
Import of the DAG moiety of PtdCho back into the chloroplast (the “eukaryotic pathway”

of thylakoid lipid synthesis) is also indicated, and takes place by an unknown mechanism.

28

fact that there are no eukaryotic galactolipid species in Chlamydomonas (Giroud etal.,
1988). This may not be true of all green algae, however, as recent evidence suggests that
the chlorophyte alga Chlorella kessleri possesses a eukaryotic pathway for thylakoid lipid
synthesis (Sato eta1., 2003b). This study was performed by the classical method of 14C
acetate pulse-chase labeling, followed by a time course of measurement of radioactivity
in each lipid class. Early in the chase, label was mostly detected in labeled MGDG and
DGDG species with a 16 carbon FA at the sn-2 position, which is typical of prokaryotic
lipids, and in the extraplastidic lipid PtdCho. Later in the chase, there was a reduction of
label in PtdCho, concomitant with an increase in MGDG and DGDG species with 18
carbon FA at the sn-2 position of glycerol. This is interpreted as the PtdCho DAG
moiety being transferred back to the chloroplast via the eukaryotic pathway.

This finding of a eukaryotic pathway in a Chlorophyte alga such as C. kessleri
raises questions about the evolutionary origin of this pathway. Apparently, either this
pathway was operative very early in the evolution of Viridiplantae, before the split
between Chlorophytes (such as Chlorella) and Streptophytes (such as Arabidopsis), or
this mechanism of lipid synthesis is the result of convergent evolution after this division.
Until there is a mechanistic molecular description of the eukaryotic pathway in both
phyla, this question will remain unresolved.

The enigma of the transport of eukaryotic fatty acid species to the plastid is
currently the subject of intense study, and the report of a mutant with a blockage in this
process has been reported (Xu et al., 2003). Data in Appendix B show that an atypical set
of galactolipids containing all-B-linked oligosaccharide chains accumulate in this mutant,

and a hypothesis for their biosynthesis is given.

29

Phosphate regulation of lipid metabolism.

As touched on above, the availability of phosphorus can have a profound impact on the
lipid composition of plant cell membranes. This is apparent in Arabidopsis, in which
phosphate limitation induces the synthesis of an extraplastidic DGDG species (Hartel et
al., 2000), and recent studies on plasma membranes from phosphate starved oat
(Andersson et al., 2003), and in maple suspension culture cells (Jouhet et al., 2003), show
that a major proportion of PtdCho is replaced by DGDG during phosphate starvation.
Since DGDG is a bilayer-forming lipid with roughly similar biophysical properties to
PtdCho, these studies imply that, under phosphate deprivation, PtdCho is replaced by
DGDG so that phosphate can be partitioned into more critical metabolites, such as
nucleic acids. This also implies a new or adaptable set of lipid trafficking mechanisms
which would be required to transport DGDG to the plasma membrane and ER (Dormann
and Benning, 2002).

As discussed briefly above, this lipid-substitution phenomenon is also observed to
occur in the plastid, e.g. substitution of Pthro with SQDG under phosphate limitation.
These lipid-substitution phenomena are inducible by phosphate deprivation, as are other
cellular responses to phosphate starvation such as the development of high affinity
transport systems, secretion of phosphatases to hydrolyze and release phosphate from
organic phosphate esters (Grossman, 2000), and increases in RNAse activity due to
induction of RNSI to increase turnover of cellular RNAs (Bariola et al., 1994). A Myb
domain transcription factor that potentially mediates the activation of a subset of the plant

PHO-regulon, and which is conserved between Chlamydomonas and Arabidopsis, has

30

been identified genetically in the respective organisms. The Chlamdomonas per
mutants were isolated based on their inability to induce phosphatase secretion and resist
killing by 32P04 (Shirnogawara et al., 1999), and the Arabidopsis mutant phrI was
isolated based on its inability to induce GUS expression from a phosphate-stress
inducible promoter (Rubio et al., 2001). The regulatory mechanisms involved in the
induction of the Arabidopsis SQD1/2, MGD2/3, and DGD2 genes are unresolved, and
may or may not be under the control of the PSRl/PHRl system. This point could be
resolved easily by analysis of lipid content and gene expression in these mutants grown in

phosphate limiting conditions.

Chlamydomonas and Arabidopsis as complementary models of plant lipid
metabolism.

The outline of plant lipid metabolism presented here has shown that there is fertile
ground for comparative biochemical genomics of Arabidopsis and Chlamydomonas.
These two organisms are representative of the two most deeply branching phylogenetic
clades in green plants, namely the Streptophyceae (Arabidopsis) and Chlorophyceae
(Chlamydomonas). Given that the genome sequences are available for both models
(Grossman et al., 2003; The Arabidopsis Genome Initiative, 2001), it could prove useful
to systematically compare the biosynthetic pathways involved in lipid metabolism in
these two organisms to find universal concepts applicable to all plants. It seems that
several are already evident, namely that the core pathways of fatty acid biosynthesis and
plastid membrane lipid assembly are the same, and that fatty acids are exported from the

plastid for the synthesis of extraplastidic lipids. These points are addressed in Chapter 4,

31

which, in addition to the functional analysis of BTAl, presents a set of genes that have
been annotated as having potential activities in lipid biosynthesis. This allows the
formation of hypotheses regarding lipid biosynthesis pathways, and reveals that some of
the components that are central to plastid fatty acid synthesis may also be targeted to the
mitochondrion.

This approach of using bioinformatics to identify similarities and differences
between Chlamydomonas and Arabidopsis shows several areas where apparently
conserved processes could perhaps be studied in Chlamydomonas more easily than
Arabidopsis. Fatty acid synthesis and export would be one example. It is known that
Chlamydomonas can take up and incorporate palmitate and oleate very rapidly into both
plastidic and cytoplasmic lipids (Giroud et al., 1988), and it is reasonable to think that
fatty acid auxotrophs could be isolated in this organism, as they have been for E. coli and
yeast. One would expect to find strains that are blocked at any of the various steps of
fatty acid biosynthesis outlined above, and also mutants defective in fatty acid export
from the plastid. Among these hypothetical mutants, those defective in fatty acid
synthesis could be identified by their inability to incorporate labeled acetate into fatty
acid, and fatty acid export mutants would be identified as those that incorporate this label
into plastidic lipids such as MGDG, but not extraplastidic lipids such as DGTS. The fact
that Chlamydomonas has fewer complicating factors than Arabidopsis (such as tissue
specific gene expression as in the LACS9 example (Schnurr et al., 2002) above), and that
a gene—tagging approach has been used to isolate lipid biosynthesis mutants (Chapter 5)

makes this sort of approach all the more attractive.

32

The differences in lipid metabolism between the two model organisms are also
informative. For example, the ER membranes of Arabidopsis and Chlamydomonas
should share many analogous functions, however PtdCho dominates the lipid
composition of the ER in Arabidopsis, while that of Chlamydomonas is composed
primarily of DGTS. The regulation of membrane composition in any organism,
especially in plants, is a topic that has proved fairly intractable, however, it may be the
case that biosynthesis of DGTS in Chlamydomonas and PtdCho in Arabidopsis are under
the control of a conserved membrane sensing mechanism. In other organisms, this sort of
membrane sensing takes the form of a membrane-bound transcription factor being
released and activated upon a change in membrane composition, as in the SREBP system
of metazoans (Brown and Goldstein, 1999; Dobrosotskaya et al., 2002), and the Opilp
protein in yeast (Loewen et al., 2004). Systems such as these are completely unknown in
plants, but it seems highly probable that an analogous system would be in play in the
plant kingdom. The identification of such a regulatory mechanism would certainly
benefit from a more thorough understanding of lipid biosynthesis and turnover in

multiple plant model systems.

Development of Chlamydomonas as a model system for lipid metabolism. Taken
together, the results in Chapters 3, 4 and 5 give the starting point for development of
Chlamydomonas as a model for plant lipid metabolism. Gene tagging will be useful in
forward genetic screens for lipid mutants or auxotrophs, as shown in Chapter 5, and
RNAi techniques are useful for reverse genetic experiments targeted to specific genes, as

outlined in Chapter 4. This last method may be especially helpful for determining the

33

functions of potentially essential genes, as is probably the case for BT A1 in Chapter 4.
These technologies, combined with biochemical approaches such as those in Chapter 3,
as well as the use of genomic resources, as demonstrated in Chapter 4 (and Appendix A,
in the case of Galdieria sulphuraria), will make Chlamydomonas a useful complement to

seed-plant models of lipid metabolism.

34

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51, 47-54.

Xu, C., Fan, 1., Riekhof, W., Froehlich, J.E., and Benning, C. (2003). A pennease-like
protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J. 22, 2370-
2379.

Xu, C., Harte], H., Wada, H., Hagio, M., Yu, 8., Eakin, C., and Benning,C. (2002). The

pgpI locus of Arabidopsis encodes a phosphatidylglycerol synthase with impaired
activity. Plant Physiol. 129, 594-604.

42

Yu, B. and Benning, C. (2003). Anionic lipids are required for chloroplast structure and
function in Arabidopsis. Plant J . 36, 762-770.

Yu, 3., Wakao, 8., Fan, J .L., and Benning, C. (2004). Loss of plastidic lysophosphatidic
acid acyltransferase causes embryo-lethality in Arabidopsis. Plant Cell Physiol. 45, 503-
510.

Yu, H., Xu, C ., and Benning, C. (2002). Arabidopsis disrupted in SQDZ encoding

sulfolipid synthase is impaired in phosphate-limited growth. Proc. Natl. Acad. Sci. U. S.
A. 99, 5732-5737.

43

Chapter 2
Functional characterization of the bacterial betaine lipid biosynthetic enzymes BtaA

and BtaB

Abstract

Betaine lipids are non-phosphorous glycerolipid analogs of phosphatidylcholine. The
biosynthesis of the betaine lipid diacylglyceryl-NJVJV-trimethylhomoserine (DGTS) has
previously been studied in phosphate-starved cells of the purple bacterium Rhodobacter
sphaeroides, and a genetic approach identified two proteins that are necessary for this
process. Here we show that all reactions of DGTS biosynthesis in R. sphaeroides are
attributable to RthaA and RthaB, as co-expression of the respective genes leads to
DGTS formation in E. coli, which normally lacks this lipid. The recombinant RthaA
protein was membrane-associated and showed S-adenosylmethionine:diacylglycerol 3-
amino-3-carboxypropyl transferase activity. RthaA directed the transfer of label from 1-
[”C]-S-adenosylmethionine (AdoMet) into the DGTS precursor
diacylglycerylhomoserine (DGHS) allowing an analysis of the biochemical and kinetic
properties of this enzyme. Comparative analysis of RthaA and its bacterial homologs
revealed a motif with similarity to the AdoMet binding pocket of methyltransferases, and

allowed the prediction of residues involved in substrate binding.

45

Introduction

Many organisms, such as bacteria, plants, and fungi, rely on mineral nutrients taken
directly from the soil or aquatic environment. As such, these organisms tend to have
exquisitely specialized mechanisms to cope with limitation of a given essential nutrient.
For example, most organisms have well defined responses to phosphate limitation,
including the induction of extracellular phosphatases and high affinity transport systems
(Grossman, 2000), the reorganization of fundamental biochemical pathways, e.g.
glycolysis (Plaxton, 1996), and even the replacement of cellular membrane phospholipids
with non-phosphorous lipids. This last point has been well documented in the plants
Arabidopsis (Essigmann et al., 1998; Hartel et al., 2000) and oat (Andersson et al., 2003),
and most pronouncedly in the a-proteobacteria Rhodobacter sphaeroides and
Sinorhizobium meliloti, which under phosphate stress become depleted of membrane
phospholipids and induce the synthesis of non-phosphorus lipids, such as glycolipid
species, the betaine lipid diacylglyceryl-NNN-trimethylhomoserine (DGTS), and
ornithine containing lipids. (Benning et al., 1995; Geiger et al., 1999).

DGTS, which was first discovered in the unicellular alga Ochromonas danica
(Brown and Elovson, 1974) and thereafter in Chlamydomonas reinhardtii (Janero and
Barrnett, 1982), and other lower plants and fungi (Furlong et al., 1986; Kunzler and
Eichenberger, 1997; Rozentsvet et al., 2000), has been proposed to take the place of
phosphatidylcholine (PtdCho) in membranes of these organisms. Studies of DGTS
biosynthesis using radioisotope feeding experiments identified methionine as a source of
both the C4 homoserine moiety and methyl groups of DGTS (Sato, l988)(see structure in

Figure 2.1), leading to the conclusion that S-adenosylmethionine (AdoMet) could be the

46

active donor of the four carbon unit of methionine to form the precursor
diacylglycerylhomoserine (DGHS), followed by lipid-linked methylation of DGHS by an
AdoMet dependent methyltransferase (Hofmann and Eichenberger, 1996)(Figure 2.1). A
closely related lipid, diacylglyceryl-NNN-trimethylhydroxymethyl-B-alanine (DGTA), is
derived from DGTS as judged by pulse-chase radiolabelling studies (Vogel and
Eichenberger, 1992).

Recently, Klug and Benning (2001) used a genetic approach in R. sphaeroides to
gain access to genes that are required for the biosynthesis of DGTS. This work resulted
in the identification of an operon containing two open-reading frames, designated btaA
and braB, which are necessary for DGTS accumulation during phosphate deprivation.
The product of btaA was proposed to function as an AdoMet-dependent 3-amino-3-
carboxypropyl-transferase, producing the intermediate DGHS. RthaB showed a high
degree of similarity to methyltransferases, and was proposed to be a trifunctional
AdoMet-dependent N-methyltransferase, adding 3 methyl units to the amino function of
DGHS to form DGTS (see Figure 2.1). In fact, RthaB shows a high degree of sequence
similarity to RsttA, which produces phosphatidylcholine by N-trimethylation of
phosphatidylethanolamine (Arondel et al., 1993), indicating that RthaB and RsttA
may share analogous functions in separate pathways. In order to understand DGTS
biosynthesis in greater detail, we initiated a biochemical characterization of the BtaA and
BtaB proteins fiom R. sphaeroides, focusing in particular on RthaA due to the unusual

nature of the proposed reaction.

47

Figure 2.1. DGTS biosynthetic pathway and involvement of RthaA/B-CrBTAl.
BtaA is proposed to catalyze the transfer of the 3-amino-3-carboxypropyl group of
AdoMet to the 3-hydroxyl of DAG to form the intermediate DGHS. DGHS is then
sequentially N—methylated by the BtaB methyltransferase to form DGTS. In C.
reinhardtii, these functions would be carried out by the BtaA- and BtaB- like domains of
the multifunctional CrBTAl protein. AdoHcy, S—adenosylhomocysteine; AdoMet, S-
adenosylmethionine; DAG, diacylglycerol; DGHS, diacylglycerylhomoserine; DGTS,
diacylglyceryl-(N,N,N)-trimethylhomoserine; MDO, membrane derived oligosaccharides;

MetK, AdoMet synthetase; 5'MTA, 5 '-methylthioadenosine.

48

0" Mets\
Phospholipid

OJYV CH3 ATP
NH3
S\Yhh'etK M00
AdoMet NH; DA
d 911;,
M9 (I: +
$2
BtaA/BTA1

5'MTA Rz

G

0 *0
HOWQTRI

233.1125; Bta B/BTA1

23333;; BtaB/BTA1

23333; f... BtaB/BTA1
R2

6 o’l‘o

DGTS * W0 R1

“'Y‘” I

 

49

Materials and Methods

Materials. Phospholipase C from Bacillus cereus was from Sigma (St. Louis, MO),
mouse monoclonal anti-His tag antibody was from Qiagen (Valencia, CA). Restriction
enzymes, T4 DNA ligase, calf-intestinal phosphatase, and Klenow fragment were fiom
New England Biolabs (Beverly, MA), Taq DNA polymerase was from Roche
(Indianapolis, IN), and Pfu DNA polymerase was from Stratagene (La Jolla, CA). All
other chemicals and solvents were of reagent grade and were from Sigma, EM Science

(Gibbstown, NJ) or J .T. Baker (Phillipsburg, NJ).

Expression of Rsth and RsbtaB. A summary of all strains and plasmids used in this
study is presented in Table l. The coding regions of Rsth and RsbtaB were PCR-
amplified from plasmid pRKL323 (Klug and Benning, 2001) for expression in pQE-3l
(Qiagen) or pACYC-31 (Dormann et al., 1999) respectively, with the following primers
(restriction sites underlined): btaA forward QM), 5’-
ACATGCATGCAGTGACGCAGTTCGCCCTC-3’; btaA reversem, 5’-
CGGGGTACCAGGACGATCCGCTCGAACCG-3’; btaB forward (Emil), 5’-
GCGGATCCGATGACCGACGCCACCCAT-3’; btaB reverse (HinDIII), 5’— “
GCAAGCTTCTCTCACCGCGTGAGCGTG-3’. PCR was carried out with Taq DNA
polymerase (Roche) according to the manufacturer’s specifications, except that 10%
DMSO (v/v) was added to each reaction to overcome difficulties in PCR due to the high
G 4C content of R. sphaeroides DNA. PCR products were first cloned into pCR2.l-
TOPO (Invitrogen, Carlsbad, CA) and sequenced at the MSU Genomics Technology

Support Facility, followed by subcloning into the expression vectors using restriction

50

 

 

sites as outlined in the PCR primers (see Table I). The resulting constructs were thus
designated thaA (btaA sequence in pQE-3l), thaB (btaB in pACYC-31), and thaA-
LC (btaA in pACYC-31). PCR2.1 and pQE-31 derivatives were propagated in LB
medium containing ampicillin at 100 rig/ml, and pACYC-31 derivatives in LB with
chloramphenicol at 25 ug/ml. Strains containing compatible combinations of plasmids as
indicated in individual experiments were constructed by co-transformation of heat-shock
competent cells of TOPIOF’ (lnvitrogen) with 100 ng of both plasmids followed by
selection on LB containing both ampicillin and chloramphenicol.

In vivo production of DGHS and DGTS in E. coli and R. sphaeroides. E. coli TOP10
F’ cells harboring compatible combinations of plasmids as described in individual
experiments were grown as 2 ml overnight cultures, and 0.1 ml of these cultures were
used to inoculate 10 ml LB containing appropriate antibiotics. After growing to OD
~0.6, the cultures were harvested by centrifugation, and dispersed into 10 ml M9 minimal
medium (Sambrook et al., 1989) containing 0.2 mM isoprpopyl-B-D-thiogalactoside
(IPTG) and 0.5 pCi 1-[14C]-methionine (American Radiolabeled Chemicals, St. Louis,
MO). Cultures were incubated for 3 h and harvested by centrifugation, followed by
extraction with 2 ml chloroformzmethanol (1:1, v/v) and phase separation by addition of
0.5 m1 1M KCI, 0.2 M H3PO4. R. sphaeroides wild-type 2.4.1 and its mutant btaB-dis
(Kan') (Klug and Benning, 2001) were grown to mid-log phase in 100 ml phosphate-
replete or phosphate-limited Sistroms medium (Sistrom, 1960; Sistrom, 1962) as

described (Benning et al., 1995; Klug and Benning, 2001). A 10 ml aliquot of each

51

Table 2.1. Strains and plasmids used in this work.

 

 

 

Strain or plasmid Description Source
R. sphaeroides
2.4.1 Wild type (Benning and

Somerville, 1992)

 

 

 

 

 

 

 

 

 

 

 

 

btaB-dis 2.4.1 with KanR cassette disrupting btaB (Klug and Benning,
2001)
E. coli
TOP] OF’ Cloning and expression strain, <D801acZAM15, Invitrogen
F’ {tactq Tn10(TetR)}
Plasmids
R2.l PCR product cloning vector (AmpR) Invitrogen
pBluescript SK + General cloningvector (AmpR) Stratagene
pRKL323 R. sphaeroides btaAB operon in pBluescript (Klug and Benning,
SK + 2001)
pACYC-31 Low copy E. coli expression vector (CamR) (Dormann et al.,
1999)
pQE-31 High copy E. coli expression vector (Ampk) Qiagen
thaA Sphl-Kpnl fragment of btaA amplified This work
from pRKL323 in pQE-3l
thaB BamHI-HinDllI fragment of btaB amplified This work

 

from pRKL323 in pACYC-3l

 

 

52

 

culture was then harvested by centrifugation and suspended in 10 ml fresh Sistrom’s
medium containing 05 uCi l-[14C]-Met, followed by 3 h incubation at 28° C and
harvesting and extraction as described for the E. coli cultures above. A portion of the
organic phase from each extraction was spotted onto activated (120° C, 2 h), ammonium
sulfate impregnated TLC plates (Silica-60, Baker) and resolved in acetone/toluene/water
(91 :30:7 , v/v) followed by autoradiography (method A) or onto untreated plates and
developed in the solvent system chloroform/acetone/methanollacetic acid/water

(lO:4:2:2:1, v/v) (method B).

Subcellular localization of BtaA protein. A 2 ml overnight culture of cells containing
thaA was used to inoculate 100 ml of LB-ampicillin, and then incubated at 37° C to an
OD600 of 0.7-0.9. IPTG was added at 50 DM to induce low level expression of RthaA
and to avoid the formation of inclusion bodies, and the culture was shifted to 28° C and
incubated a further 4 hours. Cells were harvested by centrifugation and suspended in 4
ml lysis buffer (20 mM KPi, pH 7.2, 500 mM NaCl, 15 % (v/v) glycerol), frozen in liquid
nitrogen and thawed slowly, and then lysed by sonication for 30 sec 3 times with a
microprobe tip. The crude lysate was centrifuged at 2,000 g for 10 min to remove
unbroken cells and cellular debris, and the supernatant was subjected to centrifugation at
20,000 g for 20 min. The soluble fraction was transferred to a new tube and the glassy
pellet of membranes was gently washed with lysis buffer, followed by addition of 400 pl
of lysis buffer and resuspension of the membrane fraction by sonication in a water bath
sonicator. The resuspended membranes were then subjected to another round of

centrifugation, washing and resuspension, resulting in a washed membrane fraction, and

53

20 pg of protein of each fraction (cell lysate, soluble, and membranes) were subjected to
SDS-PAGE on a 10% gel, followed by either Coomassie staining or western blotting

(Sambrook et al., 1989) with anti-His6 antibody (Qiagen) at a dilution of 1:3000.

In vitro activity assays of RthaA. E. coli TOP10 F ’ harboring thaA was grown in
250 ml LB ampicillin at 37° C to an OD of 0.7, and induced with 0.25 mM IPTG
followed by an additional 4 h of growth at 28° C. Cells were harvested by centrifugation
and the cell pellet was suspended in 10 ml of cold buffer (50 mM HEPES, 1 mM DTT, 1
mM EDTA, pH 7.3). The resuspended cells were sonicated 3—4 times, 30 seconds each
with a microprobe tip, and the lysate was centrifuged at 2000 g for 10 min to remove
unbroken cells and cellular debris. 1 ml aliquots of the cell-free extract were then frozen
in liquid N2 and stored at —80° C prior to use. Activity under these storage conditions did
not decrease appreciably for at least 1 month.

Assays for the pH versus activity profile were conducted in 100 pl final volume
by combining 48.75 p1 of cell free extract with 48.75 pl of 100 mM HEPES, TrisCl, or
MES, 1 mM DTT, 1 mM EDTA, at varying initial pH to give a final pH in the range of
5.5-8.6 when mixed with the cell free extract (initial pH 7.3). Reactions were initiated by
addition of 25,000 dpm 1-[14C]-AdoMet (American Radiolabeled Chemicals, 2.5 pl,
final concentration of AdoMet of 2.1 pM), incubated for 30 min at 28° C, and terminated
by addition of 400 p1 of chloroform/methanol (1:1, v/v) anleO pl 0.9% (w/v) NaCl to
separate aqueous and organic phases. The organic phase was transferred to a new tube,
dried under a stream of nitrogen, and dissolved in 50 p1 chlorofonn/methanol (1:1, v/v).

This lipid extract was then spotted onto silica-60 TLC plates (Baker) and developed with

54

the solvent chloroform/acetone/methanollacetic acid/water (10:4:2:2: 1 , v/v), followed by
quantification of the signal for DGHS (and methylated products in the case of CrBTA 1)
on a phosphorimager screen (Molecular Dynamics) with the ImageQuant software
package. Determination of KM was carried out at pH 7.6 for RthaA and 7.2 for
CrBTAl , with varying concentrations of AdoMet, and the value of KM was determined
by direct fitting of a hyperbola to the experimental data in the Origin software package
(OriginLab, Northampton, MA). Assays for the effect of proposed inhibitors were
performed with 4 pM AdoMet and 100 pM inhibitor added from a 1 mM stock solution

in water, and the same volume of water was added to the uninhibited control reaction.

Effect of DAG production by phospholipase C on RthaA activity. Phospholipase C
(PLC) from Bacillus cereus was obtained as a 32 M (NH4)ZSO4 suspension (Sigma) and
was desalted by several dilution-concentration cycles with assay buffer using an
Centricon-S microconcentrator (Amicon), and was diluted to a final concentration of -l
unit/pl. A portion of this PLC preparation was heated at 95° C for 5 min and served as a
heat inactivated control, i.e. the only difference between the experimental and control
assays was whether or not the PLC had been heat inactivated, thus allowing direct
comparison of the two treatments without consideration of residual salt or pH changes.
Assays were conducted in triplicate by pre-incubation of the reaction mix (85 pl cell
lysate, 10 pl PLC) for 10 min at 28°C followed by addition of 50,000 dpm of 1-[14C]-
AdoMet (final concentration of 4.2 pM), incubation for 30 min, and extraction and TLC

as described above.

55

Results

RthaA and RthaB are sufficient to reconstitute DGTS biosynthesis in E. coli. In
Figure 2.2, the reconstitution of DGTS biosynthesis in E. coli is shown, in agreement
with the proposed functions of the two putative enzymes. As positive controls for
accumulation of various lipids, the R. sphaeroides wild-type 2.4.1 and btaB—dis strains
(Klug and Benning, 2001) were grown in phosphate-replete or phosphate-limited
Sistrom’s medium to induce synthesis of DGHS in the btaB-dis strain, or DGTS in the
wild-type. Compatible combinations of expression plasmids were used to introduce
RthaA and RthaB singly or together into E. coli, followed by analysis of polar lipid
content.

To demonstrate the function of RthaA, we expected that E. coli expressing
RthaA could use endogenous DAG and AdoMet to accumulate DGHS, the non-
methylated precursor of DGTS (Figure 2.1). In Figure 22A we show that this is indeed
the case, as a primary-amine containing lipid which co-migrates with authentic DGHS
was present upon expression of btaA in E. coli. This result firmly establishes the role of
RthaA as the first step of DGTS biosynthesis in the pathway outlined in Figure 2.1, and
excludes the possibility that RthaA acts as a component of a heteromeric complex with a
protein specific to R. sphaeroides, or requires a substrate or co-factor present in R.

sphaeroides but not in E. coli.

56

Q\
‘3 '05 K ‘6'} ”5&3 C} i.) ‘37 83'?
$5? a? J 47 3 58‘!» «)5? f 03’ 55
49 Q9v (<93 «7’8 q?§ Q9 ((558 (003’ Q98 05,8
m '
Ml" < DGHS - W O G ,«DGHs

i w i . < DGTS

 

Irlit‘fhr-P-rtnxrvvi ‘23:. Lfihfjfigtfi‘*' ‘Wm-v'. , _ ___r_ _ ,

Figure 2.2. Reconstitution of DGTS biosynthesis in E. coli. A, E. coli expressing btaA
accumulate the DGTS precursor DGHS. Lipids were extracted and resolved as described
in the Experimental Procedures method A, and TLC plates were sprayed with ninhydrin
reagent to reveal primary amine-containing lipids. The btaA expression strain (lane 3)
shows a ninhydrin positive band co-migrating with authentic DGHS from a phosphate
stressed R. sphaeroides btaB KO strain (lane 1). Phosphate replete R. sphaeroides btaB
KO and empty pQE-31 serve as negative controls (lanes 2 and 4, respectively). B, BtaB
produced by expression of btaA on a compatible plasmid reconstitutes DGTS
biosynthesis. l-[14C]-methionine labels DGHS in the btaA expression strain, and co-
production of the BtaB protein converts a portion of this precursor to DGTS in cells

harboring both thaA and thaB.

57

To show the function of the putative trifunctional methyltransferase RthaB, a
compatible expression plasmid containing the RsbtaB gene was introduced into E. coli
along with the thaA expression plasmid. Our expectation was that a portion of the
DGHS produced by RthaA would be converted into N-methylated products as shown in
Figure 2.1. Figure 2.28 shows the results of a metabolic labeling strategy aimed at
testing the function of RthaB. R. sphaeroides and E. coli strains were incubated with 1-
['4C]-methionine, which is taken up and converted into AdoMet by the action of MetK
(AdoMet synthetase, see Figure 2.1 (Cohen and Saint-Girons, 1987)), and then used by
RthaA for DGHS synthesis. The fate of the radiolabel was determined by analysis of
the resulting lipids with TLC and autoradiography. The thaA/pACYC lane shows
accumulation of label in DGHS, and the thaA/thaB strain shows that a portion of
DGHS is converted to DGTS by the action of RthaB, as judged by co-migration of the
new species with authentic DGTS. A representative result using the solvent system
described in Experimental Procedures is given in Figure 2.28, however this species co-
migrated in several other solvent systems with authentic DGTS (data not shown). In
addition, a less abundant species migrating just below the DGHS spot presumably
represents a partially methylated intermediate in this particular solvent system (Figure
2.2B). These results suggest that RthaB acts as the sole methyltransferase in the
pathway.

RthaA is associated with membranes. Analysis of the localization of RthaA within
the E. coli cell is given in Figure 2.3. Prediction based on hydropathy analysis (Figure

2.3A), (Kyte and Doolittle, 1982) and with the T MHMM algorithm (not shown)

58

 

 

 

 

 

A 2
Hydrophobic
31
it
30
E4
autumn
’ eeeeeaea
BtaAresiduanumber
B Lys. 801. Mernb. C
<9 0‘” <9 {5‘ $193 6"
.9 a .0 6° 9° & s" or «9
_,_, ____
"wm 'Iv ~45KDa
9:” '2g
gr: 3*!!!

 

Figure 2.3. The RthaA protein is associated with cellular membranes. A,

Hydropathy analysis of RthaA. B and C, E. coli cells harboring meA or empty pQE-31

vector were induced, harvested, and separated into a soluble and membrane fraction as

described in the text. Twenty pg of protein of each fraction were electrophoresed on a

10% SDS-PAGE gel and (B) stained with Coomassie Blue or (C) transferred to a PVDF

membrane and probed with anti-His tag antibody. The position of Hiso-BtaA protein ( ~45

KDa) is indicated.

59

(Krogh et al., 2001) suggests that RthaA is devoid of membrane spanning helices and is
very hydrophilic throughout. Based upon this analysis, RthaA was predicted to be a
soluble protein. Figure 233 shows the results of the fractionation of a RsbtaA-expressing
E. coli culture by differential centrifugation with the parental pQE-31 vector serving as
control. Comparison of the cell lysate (Figure 238, left two lanes) and soluble (Figure
2.33, middle two lanes) fractions shows no discemable difference between the two
protein profiles, but the membrane fraction (right two lanes) shows enrichment of a ~45
kDa putative RthaA protein in the thaA membrane fraction (Figure 2.33, arrowhead).
The Hing-tagged recombinant RthaA was also identified by irnmunoblotting with anti-
Hi56 antibody as shown in Figure 2.3C. Empty vector controls were not present on the
immunoblot shown, but were routinely devoid of cross-reacting bands. Apparently,
RthaA is present in the cell lysate and membrane fractions, but absent from the soluble
fraction, suggesting that it is a membrane-associated protein. The presence of multiple
basic residues at the termini of some of the BtaA homologs suggests that the protein is
bound to the membrane though electrostatic interactions, similar to some peripheral
membrane proteins, e.g. (Wang et al., 2000). This hypothesis is presented schematically
in Figure 2.7A.

RthaA has AdoMet:diacylglycerol 3-amino-3-carboxypropyltransferase activity.
In vitro assays of RthaA enzyme activity were conducted using 1-[14C]-AdoMet to
follow product formation, and results are presented in Figures 2.4 and 2.5. Initially we

attempted to solubilize and purify RthaA utilizing the engineered His6-tag, but our

60

 

>
W

._a
O
O

DGHs— : - ‘-

DGHS synthesis
(% of control)
8

N
O

 

NO
8
I

DGHS synthesrs
(% of control)
8
O

0'1" u .-

06>, H (+)

.0
O
6i; 94 PLC

7
Figure 2.4. In vitro assay of BtaA activity. E. coli cells expressing RthaA were broken
by sonication and assayed for the ability to incorporate label from 1-['4C]-AdoMet into
DGHS as described in Experimental Procedures. A, Lysates from cells harboring pQE-3l
or thaA plasmids were incubated with labeled AdoMet and products analyzed by TLC.
B, Incubation of assay mixtures with potential inhibitors of the reactions. C, Generation
of DAG in situ by pre-incubation of assay mixtures with phospholipase C. Statistical
significance was assessed with a 2-tailed students-t test, and P< 0.05 is indicated by "*",
while P<0.0l is indicated by "**". AdoHcy, S-adenosylhomocysteine; AdoSme, 5'-

methylthioadenosine; DGHS, diacylglycerylhomoserine; PLC, Phospholipase C.

61

 

% of Maximum
8
J

 

 

 

 

 

 

 

40..
20 . . . .
6.5 7 7.5 8 8 5 9
pH
8
v0 5 - _
3?
e 4‘
g .
to 3 ‘
22
E 2-
C
3
.8. 1 -
5 -
0' - . . - fl .
O 20 40 60 80 100
[AdoMet] (pM)

Figure 25. pH dependence of BtaA and affinity for AdoMet. A, Assays were carried
out in HEPES or HEPES/Tris buffer at the indicated pH values as described in the text.
Reaction mixtures were incubated at 28° C for 1 h, followed by extraction, TLC (method
B), and quantification by exposure to a phosphorimager screen B, Assays were canied

out at pH 7.6 as in the determination of pH optimum, and the concentration of AdoMet

was varied.

62

attempts were unsuccessful. While we were able to purify small amounts of apparently
soluble RthaA protein using various detergents in the lysis buffer, we were unable to
demonstrate activity in a reconstituted liposome system. To circumvent this problem, we
developed a system to minimize the steps between expression and enzyme assay to
demonstrate the proposed reaction catalyzed by RthaA (Figure 2.1). E. coli membranes
contain small amounts of DAG derived from the synthesis of membrane-derived
oligosaccharides (Kennedy, 1987). Thus as the source of enzyme and DAG, a cell-free
lysate fiorn thaA expressing E. coli was used. As demonstrated in Figure 2.4A , label
from AdoMet was efficiently incorporated into DGHS by transfer of the 3-amino-3-
carboxylpropyl moiety to DAG. To probe the effect of potential inhibitors of the reaction
(Figure 2.4B), S-adenosylhomocysteine (AdoHcy), 5’—methylthioadenosine (5’MTA),
and methionine (Met) were tested for their ability to reduce incorporation of label into
DGHS. AdoHcy, a product and potent inhibitor of methyltransferases (Fauman et al.,
1999), and 5'-methylthioadenosine (the presumed product of RthaA) were slightly
inhibitory, but Met had no significant effect on the reaction.

In order to determine that DAG is the lipid substrate of BtaA, the cell lysate was
incubated with active or heat inactivated phospholipase C (PLC), thus generating DAG in
situ. Results of this experiment (Figure 2.46) demonstrate that RthaA activity is
stimulated at least two-fold in the assay mixtures that were pretreated with 10 units of
PLC, which corroborates the role of DAG as the lipid substrate.

A number of divalent metals including Mg“, Ca”, and Mn++ were tested at 1 mM
for their ability to stimulate the reaction, but no stimulatory effect was observed, with the

possible exception of slight increases in activity by calcium ions (data not shown). This

63

stimulation was variable between different preparations of enzyme, indicating that if CaH
does have a stimulatory effect, it might be indirect, e.g. Cchould act as a counter ion to
membrane phospholipids and thus stabilize the membrane upon which RthaA is acting.
The metal chelators EDTA and EGTA were tested and were found to have no effect on
enzyme activity, corroborating the idea that RthaA is probably not a metalloenzyme.
RthaA was found to have a slightly basic pH optimum of 4 .6 (Figure 2.5A), and the KM

toward AdoMet was 27 pM (Figure 2.58).

BtaA is related to methyltransferases. As shown in the multiple sequence alignment of
Figure 2.6.4 , BtaA from R. sphaeroides has orthologs in a number of other bacteria,
predominantly its relatives in the Gram-negative family of oI-proteobacteria. The
orthologs from S. meliloti and A. tumefaciens were cloned into pQE-31 essentially as
described above for btaA from R. sphaeroides, and recombinant E. coli carrying these
constructs also accumulated DGHS (data not shown), indicating that the bacterial btaA
genes are truly homologous in both sequence and function. All of these BtaA orthologs
are encoded in a putative operon of the same structure as that of R. sphaeroides
(predicted to be transcribed as a bicistronic btaAB message (Klug and Benning, 2001))
except for the BtaA-like protein of Xylella fastidiosa, which is not accompanied by a
btaB gene in the predicted operon. Given the fact that BtaB is similar to other small-
molecule methyltransferases, is was impossible to unequivocally identify the BtaB
ortholog elsewhere in this genome. Using PSI-BLAST(Altschu1 et al., 1997) to search

for other

64

Figure 2.6. Multiple sequence alignment of bacterial BtaA orthologs and
methyltransferases. BLAST searching using R. sphaeroides BtaA (Rsp, AAK53560) as
query was used to identify orthologous proteins in finished genomes of Agrobacterium
tumefaciens (Atu, NP_3 55081), Sinorhizobium meliloti (Sme, NP_3 86300),
Mesorhizobium loti (M10, NP_103130), Rhodopseudomonas palustris (Rpa, NP_946080).
PSI-BLAST revealed that a domain in the N-terrninal half of BtaA is similar to the
AdoMet binding motifs of some bacterial methyltransferases. Regions corresponding to
methyltransferase motifs I and II are indicated with (****), and GenBank protein

accession numbers are used as identifiers for the methyltransferases in the pileup.

65

AtuBtaA
SmeBtaA
MloBtaA
RsthaA
RpaBtaA
ZP_00081747
NP_616375
NP_637624
ZP_00010354
ZP_OOO98885
NP_442452
consensus

58
59
58
60
80

Motif I Motif II
**** *****
AMELGEGHRIVTIGSGGCNMLAYLSRNPASIDVVDLNPHHIAL
AMQIRPGHRIVTIGSGGCNMLTYLSAEPARIDVVDLNPHHIAL
AMQLGQGHRIVTIASGGCNILAYLTRSPARIDAVDLNAAHIAL
ALAIRPGDRLVAIASGGCNVLSYLTQGPGSILAVDLSPAHVAL
ALQLPIGSTIVTISSGGCNALSYLTAKPAQVFAVDLNEAHLAL
YLRSRPGGRLLEIGCGSGTMLSYLGSLGWRTEGIDVDPSAVAN
AAELSPQDTILEIGAGIGNLTERLARRAKKVIAVELDPALVSV
ARVALPGARVLDVGCGGGLLSEAMARLGAQVTAIDLAPELVKV
AGRLSPSDRVLEIGCGTGGTAIRLAPGVAQWTATDLSPEMVRI
SEVLRADMRVLEVATGTGLMALGIAKFVRQVEATDFSPKMIET
NLAISPGQKVLDLCCGGGQATVYLAQSGATVVGLDASPKALGR
a-—l--g-riv-ig-G---m-—-l----a-i-avdl-p--i-l

66

Figure 2.7. Schematic representation of BtaA protein and active site. A, BtaA is
proposed to be a peripheral membrane protein interacting with the negatively charged
membrane through basic regions present at the N- and C- termini (Figure 2.6A). 8,
Schematic model for the active site of BtaA, with AdoMet binding through motifs I and
II as has been shown for methyltransferases. The involvement of a hypothetical general

base (:B) to position and activate the 3-hydroxyl of DAG is also proposed.

67

  
    
 

AdoMet DAG
. O o

, if

   

++C
co.

2

    

i

  
    

    

G-loop/Motif I

 

68

protein families that might be related to BtaA, we discovered a number of related, but as
yet uncharacterized bacterial methyltransferase-like proteins. A sequence alignment of
the pertinent regions, as well as an analysis of putative motifs involved in AdoMet
binding (Martin and McMillan, 2002) is shown in Figure 2.68, leading to a prediction of

residues involved in AdoMet binding, as presented schematically in Figure 2.78.

Discussion

This work was undertaken to determine the biochemical function of two proteins, BtaA
and BtaB, which had both previously been shown by genetic means to be necessary for
phosphate starvation-induced DGT S biosynthesis in R. sphaeroides (Klug and Benning,
2001). A key result presented here is the demonstration that, not only are BtaA and BtaB
necessary for DGTS biosynthesis in R. sphaeroides, but that they are also sufficient to
provide all the activities in the biosynthetic pathway, as judged by their ability to convey
DGT S production in E. coli, which normally lacks this lipid. Genetic analysis following
the isolation of these genes left this point somewhat unresolved, in that disruption of the
RsbtaB gene with a kanamycin resistance cassette resulted in the accumulation of both
DGHS and a monomethylated form during phosphate starvation, leaving the possibility
open that the RthaB protein may only act as the methyltransferase for the second and/or
third N-methylation(s) of the precursor (Klug and Benning, 2001). A more detailed
analysis of RthaB was prevented by the apparent toxicity of this protein in E. coli cells.
While low expression sufficient for DGTS biosynthesis was achieved, it was not possible

to purify active RthaB in amounts pemritting further characterization.

69

However, the RthaA protein was conclusively shown to transfer the 3-amino-3-
carboxypropyl moiety from AdoMet to DAG. This result is in agreement with the role
that had been tentatively assigned based on mutagenesis and ectopic expression in the
previous work, and rules out any other contingencies for the activity of RthaA.
Attempts to solubilize, purify, and reconstitute BtaA into liposomes of defined
composition were unsuccessful. In fact, the BtaA activity is sensitive to low
concentrations of detergent when, e.g. 0.5 % (w/v) octyl-glucoside or Tween-20 was
added to an assay as described above. For this reason, we took the approach of keeping
the recombinant enzyme in an as native a state as possible and minimizing the steps
between expression and enzyme assay. E. coli membranes contain a small amount of
DAG derived from the synthesis of membrane derived oligosaccharides (Kennedy, 1987),
thus we used the endogenous E. coli DAG pool from membranes in the cell-free lysate as
substrate to obviate the need for reconstitution into liposomes.

Previous work in a cell-free system on the cognate BtaA-type transferase activity
in C. reinhardtii had given conflicting results as to the identity of the hydrophobic
substrate, indicating that DAG might not be the direct substrate because the addition of
exogenous DAG strongly inhibited the reaction (Moore et al., 2001). However, the point
was raised that excess DAG might simply disrupt the membrane environment in which
the enzyme(s) are working, and the decrease in activity might not be a result of enzyme
inhibition, per se. The results we present for recombinant RthaA seemingly provide a
solution to this question, in that incubation of our enzyme/membrane system with PLC
stimulated the rate of DGHS formation, and probably represents a milder way of

generating DAG in situ, versus adding this lipid directly to the reaction.

70

The kinetic properties of RthaA toward AdoMet and its analogs are similar to
those of methyltransferases. The apparent KM for AdoMet of 27 pM that was measured
for RthaA is in line with values obtained for methyltransferases, which range from ~5-
50 pM (Edwards and Dixon, 1991; James et al., 1995; Preisig et al., 1989). RthaA was
weakly inhibited by S-adenosylhomocysteine (AdoHcy), which lacks the activated
methyl unit of AdoMet and competitively inhibits methyltransferases (Fauman et al.,
1999), and also by its presumed product, 5’-methylthioadenosine, but not by methionine
itself. The pH profile showed an optimum at ~pH 7.6, which is similar to that measured
for the activity from C. reinhardtii (Moore et al., 2001).

Having established that RthaA is active in the proposed reaction, and given that
the substrate DAG is a minor component of bacterial cell membranes (Kennedy, 1987)
we wanted to determine if RthaA associates with membranes. Hydropathy analysis
using a standard Kyte-Doolittle algorithm (Kyte and Doolittle, 1982) (Figure 2.3A) or a
more sensitive hidden-Markov model based approach (TMHMM, (Krogh et al., 2001)) to
predict transmembrane helices revealed that RthaA is actually quite hydrophilic and
predicted to be a soluble protein. However fractionation by differential centrifugation
(Figure 2.38 and C) revealed that it is probably a peripheral membrane protein, as it is
not present in a soluble form in preparations that are active in the RthaA assay, but is
instead present solely in membrane containing fractions. Analysis of the multiple
sequence alignment in Figure 2.7A showed that a segment of 12-15 basic and
hydrophobic residues is present in the N—terminus of some of the homologs, as well as a
polybasic stretch at the C-tenninus. This finding led to the hypothesis that these regions

might target and anchor RthaA to the inner membrane by forming a basic amphipathic

71

a-helix and providing favorable electrostatic interactions with the anionic membrane
surface, as is the case for some other peripheral membrane proteins (Bernstein et al.,
2000; Wang etal., 2000).

The RthaA enzyme is a member of a small group of enzymes using the
ubiquitous alkylating agent AdoMet as a donor of a 3-amino-3-carboxypropyl moiety
rather than the methyl group. Other characterized enzymes that catalyze a similar
reaction include one involved in antibiotic biosynthesis in Nocardia uniformis (Reeve et
al., 1998), and spermidine synthase, which transfers the aminopropyl moiety of
decarboxylated-AdoMet to putrescine in the polyamine biosynthesis pathway (Korolev et
al., 2002). Sequence and structural similarity of these enzymes to AdoMet-dependent
methyltransferases has been demonstrated (Korolev et al., 2002; Reeve etal., 1998) and it
is clear from these results that these proteins share a methyltransferase fold (Fauman et
al., 1999; Martin and McMillan, 2002) with a conserved AdoMet binding pocket. The
identity of BtaA as a methyltransferase fold protein was not readily apparent on the basis
of sequence analysis, but a detailed comparison of the bacterial BtaA orthologs allowed
the identification of putative AdoMet binding motifs. The multiple sequence alignments
in Figure 2.6A and 8 show that residues corresponding to G75 and D94 in the R.
sphaeroides protein are completely conserved among the BtaA homologs, and
correspond to the conserved residues in AdoMet binding motifs I and II, analogous to the
“G-loop” and “D-loop” in other nucleotide binding motifs (Kagan and Clarke, 1994;
Martin and McMillan, 2002). This, coupled with the kinetic similarities of BtaA and
methyltransferases, provides strong evidence that BtaA also binds AdoMet in a

methyltransferase fold. A schematic representation of the active site of BtaA is presented

72

in Figure 2.78, and a model for the predicted topology and binding sites of DAG and
AdoMet are represented in Figure 2.7A . The involvement of a hypothetical general base
in activating the nucleophilic hydroxyl of DAG is also presented in Figure 2.7A , however
the presence and/or identity of this residue was not predictable from the sequence
analysis given the very limited similarity of BtaA to methyltransferases outside of the
putative AdoMet binding domain. Taken together, the results presented here represent an
initial mechanistic analysis of the enzymes of DGTS biosynthesis, identifying the key
step in the pathway, DGHS synthesis, as being catalyzed by a highly divergent

methyltransferase-like enzyme.

73

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77

Chapter 3
Functions of the bacterial betaine lipid biosynthetic enzymes BtaA and BtaB are

carried out by the multifunctional polypeptide BTAl in Chlamydomonas reinhardtii.

78

Abstract

In bacteria, biosynthesis of the betaine lipid diacylglyceryl-N,N,N-trimethylhomoserine
(DGTS) has been shown to be the function of two proteins, BtaA and BtaB, which
catalyze all steps of the biosynthetic pathway. Analysis of the genome sequences of the
green alga Chlamydomonas reinhardtii led to the discovery of a gene encoding a protein
with bacterial BtaA- and BtaB-like domains, suggesting that in this organism, DGTS
biosynthesis is the function of a single gene product. To test this hypothesis, the coding
region of the CrBTAl gene was expressed in E. coli, which led to DGTS accumulation.
Site-directed mutagenesis of the predicted active sites of the BtaA- and BtaB-like
domains of CrBTAl demonstrates the function of the individual domains, and also
demonstrates the importance of residues that had been predicted to be involved in
AdoMet binding. It is thus concluded that betaine lipid enzymes are highly divergent

members of the AdoMet-dependent methyltransferase family.

79

Introduction

The betaine lipid diacylglyceryl-N,N,N-trimethylhomoserine (DGTS) is a major
component of the extraplastidic membranes of Chlamydomonas reinhardtii (Giroud et al.,
1988). Due to its structural similarity to the phospholipid phosphatidylcholine (PtdCho),
and the lack of PtdCho in Chlamydomonas, DGTS is presumed to take on the functions
of PtdCho as the bilayer-forming component of the extraplastidic membranes (Moore et
al., 2001). Previous work has shown that DGTS biosynthesis in bacteria is the function
of two gene products, BtaA and BtaB, which are required for phosphate stress induced
DGTS production in Rhodobacter sphaeroides (Klug and Benning, 2001). This work
reports the identification and functional characterization of a protein from
Chlamydomonas, BTAl, containing BtaA and BtaB- like domains in a single
polypeptide. This leads to the conclusion that, in Chlamydomonas, a single protein is

responsible for the bulk of extraplastidic membrane biogenesis.

Materials and Methods

Materials. Restriction enzymes, T4 DNA ligase, calf-intestinal phosphatase, and
Klenow fragment were from New England Biolabs (Beverly, MA), T aq DNA polymerase
was from Roche (Indianapolis, IN), and Pfu DNA polymerase was from Stratagene (La
Jolla, CA). All other chemicals and solvents were of reagent grade and were from Sigma,

EM Science (Gibbstown, NJ) or IT. Baker (Phillipsburg, NJ).

Cloning and expression of RsbtaA, CrBTA I, and site directed mutants in E. coli. A

summary of all strains and plasmids used in this study is presented in Table 3.1. The

80

coding region of Rsth was PCR-amplified from plasmid pRKL323 (Klug and Benning,
2001) for expression in pACYC-3l (Dormann et al., 1999) with the following primers
(restriction sites underlined): btaA forward (Sgfl), 5’-
ACATGCATGCAGTGACGCAGT'TCGCCCTC-3’; btaA reverse(§;ll), 5’-
CGGGGTACCAGGACGATCCGCTCGAACCG—3’; PCR was canied out with T aq
DNA polymerase (Roche) according to the manufacturer’s specifications, except that
10% DMSO (v/v) was added to each reaction to overcome difficulties in PCR due to the
high G+C content of R. sphaeroides DNA. The PCR product was first cloned into
pCR2. l-TOPO (Invitrogen, Carlsbad, CA) and sequenced at the MSU Genomics
Technology Support Facility, followed by subcloning into pACYC-3l using restriction
sites as outlined in the PCR primers (see Table 3.1). The resulting construct was
designated thaA-LC (btaA in pACYC-3 l). PCR2.1 and pQE-3l derivatives were
propagated in LB medium containing ampicillin at 100 pg/ml, and pACYC-31
derivatives in LB with chloramphenicol at 25 pg/ml. Strains containing compatible
combinations of plasmids as indicated in individual experiments were constructed by co-
transfonnation of heat-shock competent cells of TOPIOF’ (Invitrogen) with 100 ng of
both plasmids followed by selection on LB containing both ampicillin and
chloramphenicol.

C. reinhardtii strain CC 125 (wild-type mt+, Chlamydomonas culture center, Duke
University) was grown to mid-log phase in 50 ml TAP medium (Harris, 1989) and
harvested by centrifugation. RNA was purified with Trizol reagent (Invitrogen)
according to the manufacturer’s instructions. 1.0 pg of total RNA was reverse-

transcribed with SuperScript II Rnase H' reverse transcriptase (Invitrogen), and the

81

coding region of CrBTAl was amplified with Pfu DNA polymerase (Stratagene) from an
aliquot of cDNA with the primers CrBTA] forward: 5’-
CAGGATCCAATGGGGTCGGGTCGT-3’; and CrBTAl reverse: 5’-
CAGGTACCGCCGCCAGCTGCTTA-3’. The native start codon of CrBTAl in the
forward primer is in bold, and BamHI and Kpnl sites are underlined in the forward and
reverse primers, respectively, and were used to clone the product in-frame with the N-
terrninal Hisé-tag of pQE-3l, giving rise to pBTAl.

Site directed mutagenesis of pBTAl was carried out with the Quikchange 11 XL
kit (Stratagene) according to the manufacturer’s instructions. Mutagenic primers were as
follows: for the V322A, D323A mutant version of thal (pBTAlpA), primers were pA-
fw: 5’-GCCAGGTGGTGTCGGCGGCATGCAACCCCGCGCAG-3’, and its reverse
complement, pA-rev. For the V103A, D104A mutant (pBTAlpB), primers were pB-fw:
5’-CAAGTCCATCTACGTGGCCGCCCTGTGCCACTCGCTG-3’, and its reverse
complement, pB-rev. Primers were designed such that, in addition to the missense
mutations for changing codons, a diagnostic restriction site was created or eliminated,
e.g. “pB” destroys a SalI site and “pA” introduces an Sphl site, in order to easily identify

strains carrying the mutated plasmids.

In vivo production of DGHS and DGTS in E. coli. E. coli TOPIO F’ cells harboring
compatible combinations of plasmids as described in individual experiments were grown
as 2 ml overnight cultures, and 0.1 ml of these cultures were used to inoculate 10 ml LB
containing appropriate antibiotics. After growing to OD ~0.6, the cultures were

harvested by

82

Table 3.1. Strains and plasmids used in this work.

 

Strain or Description Source
plasmid
C. reinhardtii
CC 125 Wild type Chlamydomonas
Genetics Center,
Duke University
E. coli
TOPIOF’ Cloning and expression strain, (1)801acZAM15, Invitrogen
F’ {laclq Tn10(TetR)}
Plasmids
thaA-LC SphI-Kpnl fragment of btaA amplified from This work
pRKL323 in pACYC-3l
pBTAl 8amHI-Kpnl fragment of CrBTA] coding This work
sequence in pQE-3l
PBTAl-pA V322A, D323A derivative of pBTAl This work
PBTAl-pB V103A, D104A derivative of pBTAl This work

 

83

centrifugation, and dispersed into 10 ml M9 minimal medium (Sambrook et al., 1989)
containing 0.2 mM isopropyl-B—D-thiogalactoside (IPTG) and 0.5 pCi l-['4C]-methionine
(American Radiolabeled Chemicals, St. Louis, MO). Cultures were incubated for 3 h and
harvested by centrifugation, followed by extraction with 2 ml chloroform:methanol (1 :1,
v/v) and phase separation by addition of 0.5 ml 1M KCl, 0.2 M H3PO4. A portion of the
organic phase from each extraction was spotted onto activated (120° C, 2 h) TLC plates
(Silica-60, Baker) and resolved in chloroform/acetone/methanol/acetic acid/water

(lO:4:2:2:1, v/v) followed by autoradiography.

In vitro activity assays of CrBTAl. E. coli TOPIO F’ harboring thaA was grown in
250 ml LB ampicillin at 37° C to an OD of 0.7, and induced with 0.25 mM IPTG

followed by an additional 4 h of grth at 28° C. Cells were harvested by centrifugation
and the cell pellet was suspended in 10 m1 of cold buffer (50 mM HEPES, 1 mM DTT, 1
mM EDTA, pH 7.3). The resuspended cells were sonicated 3-4 times, 30 seconds each
with a microprobe tip, and the lysate was centrifuged at 2000 g for 10 min to remove
unbroken cells and cellular debris. 1 ml aliquots of the cell-free extract were then frozen
in liquid N2 and stored at —80° C prior to use. Activity under these storage conditions did
not decrease appreciably for at least 1 month.

Assays for the pH versus activity profile were conducted in 100 pl final volume
by combining 48.75 pl of cell free extract with 48.75 pl of 100 mM HEPES, TrisCl, or
MES, 1 mM DTT, 1 mM EDTA, at varying initial pH to give a final pH in the range of
5.5-8.6 when mixed with the cell free extract (initial pH 7.3). Reactions were initiated by

addition of 25,000 dpm 1-['4C]-AdoMet (American Radiolabeled Chemicals, 2.5 pl,

84

final concentration of AdoMet of 2.1 pM), incubated for 30 min at 28° C, and terminated
by addition of 400 pl of chloroform/methanol (1:1, v/v) and 100 p1 0.9% (w/v) NaCl to
separate aqueous and organic phases. The organic phase was transferred to a new tube,
dried under a stream of nitrogen, and dissolved in 50 pl chloroform/methanol (1:1, v/v).
This lipid extract was then spotted onto silica-60 TLC plates (Baker) and developed with
the solvent chloroform/acetone/ methanol/acetic acid/water (10:4:2:2: 1, v/v), followed by
quantification of the signal for DGHS and methylated products on a phosphorimager
screen (Molecular Dynamics) with the ImageQuant sofiware package. Determination of
KM was carried out at pH 7.2 with varying concentrations of AdoMet, and determined by
direct fitting of a hyperbola to the experimental data in the Origin software package
(OriginLab, Northampton, MA). Assays for the effect of proposed inhibitors were
performed with 4 pM AdoMet and 100 pM inhibitor added from a 1 mM stock solution

in water, and the same volume of water was added to the uninhibited control reaction.

Mass spectrometry of DGTS. DGTS was isolated from lipid extracts of CrBTAl
expressing E. coli by preparative TLC on ammonium-sulfate impregnated silica-6O TLC
plates in the solvent chlorofomr/acetone/methanol/acetic acid/water (10:42:22 1, v/v).

The lipid spot was scraped from the plate and eluted in chloroform/methanol (1 :1, v/v)
and concentrated under a stream of nitrogen. F ast-atom bombardment-mass spectrometry
was as previously described (Benning et al., 1995) except that nitrobenzyl alcohol was

used as matrix.

85

Results

CrBTAl is a multifunctional protein constituting the entire DGTS biosynthesis
machinery in C. reinhardtii. The unicellular green alga C. reinhardtii is unusual among
eukaryotes in that its membranes are devoid of phosphatidylcholine, and contain DGTS
in its place. Given that a draft sequence of the C. reinhardtii genome is available
(genome.jgi-psf.org/chlamy, (Grossman et al., 2003)), we took a genomic approach to
elucidating the enzymes of DGTS biosynthesis in this organism. Figure 3.1A shows the
genomic structure of CrBTAl, (genome locus ID 15059) encoding a protein consisting of
an N-terminal domain with similarity to RthaB, and a C-terrninal portion with similarity
to RthaA (Figure 3.18 and C). The genomic locus is approximately 6 kb in length and
the transcript is interrupted 11 times by introns (Fig, 3.1A). The predicted mRNA also
contains a relatively long 3’ untranslated region of ~1 .6 kb, which is common to many C.
reinhardtii transcripts.

The residues identified for RthaA as being potentially important in AdoMet
binding are conserved in CrBTAl, and two residues in motif II (Kagan and Clarke, 1994)
of the BtaA-like domain (V322, D323) are highlighted in Figure 3. l C. They were chosen
for the mutagenesis experiments described below due to the fact that a V322A, D323A
mutant would likely be incapable of binding AdoMet. This mutation would render the
protein inactive in the DGHS synthesis reaction, but since the native conformation of the
protein would probably be retained, the BtaB-like portion should still be active in the
methylation steps. Additionally, the analogous positions in motif II (Kagan and Clarke,
1994) of the BtaB-like portion of the protein (V103, D104) were also targeted for alanine

mutagenesis (Fig 3.16). In this case, it was presumed that the

86

Figure 3.1. Identification of CrBTAl as a fusion of BtaA- and BtaB-like domains. A,
BLAST searching of the draft genome sequence of Chlamydomonas reinhardtii
(genomejgi-psf.org[chlamy) revealed a ~6 kb locus encoding a protein with BtaA- and
BtaB-like domains, referred to as BTAl. 8, Alignment of the BTAl sequence with BtaB

and BtaA.

87

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88

DGHS synthesis activity would be intact, but that the methyltransferase portion would be
unable to bind AdoMet and therefore inactive, as a result of the alanine substitutions.
Cloning of the CrBTAl coding sequence (AY656806) and expression in E. coli
led to accumulation of relatively large amounts of DGTS, allowing us to verify the
structure by mass spectrometry. Fig 3.2A shows a TLC of lipids from E. coli expressing
CrBTAl. It is clear that there is a new species accumulating in the expression strain, and
this lipid is subsequently identified as DGTS by positive ion FAB-MS (Fig, 3.28), and
consists of molecular species that are typical of E. coli membrane lipids, e.g. the species
at m/z 710.7 is DGTS carrying fatty acids totaling (carbonszdouble bonds) 32:1, and that
of m/z 736.7 is a 34:2 species of DGTS. This result verifies the hypothesis that CrBTAl
is capable of carrying out all reactions of DGTS biosynthesis in C. reinhardtii, given the
fact that it has the same function as co-expression of RthaA and RthaB in E. coli.
Site directed mutagenesis of the AdoMetbinding domains of BTAl.
Site-directed mutagenesis was used to change residues that are predicted to be critical for
AdoMet binding in the active sites of the BtaA-like and BtaB-like domains (Figure 3.3).
The CrBTAlpA protein is inactive in the synthesis of DGHS, as shown in Figure 3.3,
lane 2, with background levels of label in the DGTS or DGHS regions of the
chromatogram. BTAlpB, however, produces DGHS (Figure 3.3, lane 3), as confirmed
by co-chromatography with the product of a strain carrying RthaA on a low copy
plasmid, which causes accumulation of DGHS. Lane 5 (Figure 3.3) shows DGTS
accumulating in the pBTAl containing E. coli strain, and Figure 3.3, lane 6 shows that

the BtaB-like methyltransferase domain in pBTAlpA is still capable of methylating

89

Figure 3.2: CrBTAl catalyzes the full DGTS biosynthetic pathway. A. E. coli cells
harboring pBTAl were grown and expressed as described in Experimental Procedures,
and lipids were separated by TLC (method B) and visualized with iodine vapor. A
culture containing the empty pQE-3l vector served as a negative control. Pthro,
phosphatidylglycerol; PtdEtn, phosphatidylethanolamine; CL, cardiolipin; DGTS,
diacylglyceryl-N,N,N-trimethylhomoserine. 8. Fast atom bombardment mass
spectrometry (FAB-MS) in the positive ion mode was used to confirm the identity of the
putative DGTS spot in the TLC. Peaks at m/z 710.7 and 736.7 represent the most
abundant molecular species, with acyl chain lengths and total double bonds (carbons:

double bonds) of 32:1 and 34:2.

90

 

 

 

 

 

 

 

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91

fl" —DGTS

gain—fl”-

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Figure 3.3: Analysis of BtaA- and BtaB-like domains of BTAl by site-directed
mutagenesis. E. coli harboring combinations of plasmids as described in the text were
labeled with l-['4C]-Met and the lipids separated by TLC (Experimental Procedures,
method B), followed by exposure to a phosphorimager. DGHS, diacylglyceryl-

homoserine; DGTS, diacylglycerylWWW-trimethylhomoserine.

92

DGHS produced by RthaA. These results establish the function of the two domains of

C rBTAl as being comparable to those of the individual bacterial orthologs.

Kinetic analysis of CrBTAl. In Figure 3.4 the pH versus activity profile and
determination of KM for AdoMet for the DGHS synthesis reaction of CrBTAl are shown.
The extent of methylation changes with increasing pH (Figure 3.4A), which indicates a
difference in the pH optima between the BtaA-like and BtaB-like domains. Figure 3.48
shows the quantification of the total counts in each lane of Figure 3.4A, plotted versus the
pH, which gives the pH optimum of the DGHS synthesis reaction as ~7.2. Figure 3.4C
shows a plot of rate versus substrate concentration used to calculate the apparent KM (l 6

pM), which was determined by direct fitting of a hyperbola to the experimental data.

Discussion

This work was undertaken to determine whether DGTS biosynthesis in eukaryotes is
carried out in a similar manner as that of the BtaA/BtaB system of bacteria. Searching of
the whole-genome shotgun assembly of C. reinhardtii (www.genome.jgi-psf.org/chlamy)
with the RthaA sequence revealed a predicted protein (locus ID 15059), which we have
named CrBTAl, showing 27% identity and 40% similarity to RthaA over about 400
amino acids. In fact, not only was CrBTAl similar to RthaA, but it also contained an
N-terminal domain with similarity to RthaB, leading to the hypothesis that in C.
reinhardtii, DGTS biosynthesis is the function of a single protein with multiple activities.
To test this hypothesis, we expressed the coding region of the CrBTAl cDNA

(AY656806) in E. coli. This resulted in DGTS biosynthesis from expression of the single

93

Figure 3.4: Biochemical properties of CrBTAl. A. TLC of products of CrBTAl at
varying pH. Cell lysates were prepared and adjusted to the proper pH as described for
RthaA, followed by incubation with 1-['4C]-AdoMet (4.2 pM) for 1 hour. Products
were extracted and separated by TLC (Experimental Procedures, method B) followed by
exposure to a phosphorimager (Molecular Dynamics) screen. 8, The radioactivity in
each lane of the image in (A) was quantified in the Imageth software package
(Molecular Dynamics) and used determine the pH optimum for the DGHS synthase
activity. C, Determination of apparent KM (AdoMet) was carried out at pH 7.2 as in the
determination of optimum pH, and the concentration of AdoMet]was varied. The
apparent KM was determined by direct fitting of a hyperbola to the experimental data in

the Origin software package.

94

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95

protein, thus confirming that CrBTAl contains all activities of the pathway (Figure 3.2).
Expression of the BtaA-like portion of CrBTAl by itself was expected to result in DGHS
accumulation, as was shown for RthaA, however repeated attempts did not result in
DGHS synthesis activity (data not shown). As an alternative way of showing the
function of the two domains of CrBTAl and adding evidence to predictions of the BtaA
active site residues, we altered residues in the predicted AdoMet binding site of the BtaA-
and BtaB-like domains. The results of this experiment are informative on several levels.
First, the fact that the CrBTAl V322A, D323A mutant (CrBTAlpA) is inactive in
synthesis of DGHS and DGTS lends credence to the model that BtaA binds AdoMet in a
methyltransferase fold. Given the structure of the cognate binding site of
methyltransferases, V322 is expected to make favorable hydrophobic interactions with
the adenine moiety of AdoMet, and D323 is proposed to form hydrogen bonds to the 2’
and 3’ hydroxyls of the ribose portion. Mutation of these residues to alanine renders the
DGHS synthase function of CrBTAl inactive, or at least compromises it to below the
limits of detection. The DGHS tri-methylase function is, however, unaffected given that
the CrBTAl-pA protein is still capable of methylating DGHS produced by simultaneous
expression of RthaA (Figure 3.3, lane 6). This provides an additional piece of
information regarding the sequence of reactions, showing that the CrBTAl-B-domain is
able to take DGHS from the membrane as a substrate, rather than it being transferred
directly from the active site of the A-domain as would be the case for substrate
channeling. This is in agreement with in vitro assays conducted on isolated C. reinhardtii
membranes, as well as results presented in Figure 3.4, showing an accumulation of a

transient pool of DGHS.

96

CrBTAl activity was measured in a cell-free system as for RthaA, and similar
enzymatic parameters were observed. The analysis is somewhat complicated by the fact
that the methyltransferase is present on the same polypeptide, and thus multiple products
are formed. However, measurement of all products on the same TLC plate (Figure 3.4A)
made this a trivial point, in that the sum of DGHS and its methylated products gives the
total flux through the BtaA-like domain, and this quantity was used in the analyses of
Figure 3.48 and C. In fact, Figure 4A is informative as to the activity of the BtaB-like
(N-methyltransferase) portion, showing that the formation of the mono- and dimethylated
DGHS intermediates is preferred as the pH increases, and the final methylation is favored
at a higher pH still (~pH 8). The pH dependence of these subsequent methylation
reactions may reflect a change in pK, values of the amine species with higher degrees of
methylation.

The apparent KM for the recombinant CrBTAl activity is presumably directly
comparable with that of the activity that was measured in C. reinhardtii membranes
(Moore etal., 2001), however the value obtained for the recombinant enzyme (~16 pM)
was about 5 fold lower than that of the native activity, indicating that the membrane
composition and environment of the enzyme may affect the affinity for AdoMet of the
DGHS synthase module. This is not surprising, considering that a thorough kinetic
description would require both AdoMet and DAG concentrations to be varied, i.e. at
higher DAG concentrations (as might occur in E. coli membranes) the apparent KM of the
CrBTAl-A domain for AdoMet may be lower. This question remains unresolved given
the inability to reconstitute the proteins into the defined lipid environment of a synthetic

liposome.

97

Taken together, the results presented here show that the enzymes and mechanisms
of DGTS biosynthesis are conserved between eukaryotes and prokaryotes, with the
exciting result that the eukaryotic activities are fused into a single polypeptide. This
latter phenomenon is common among metabolic enzymes such as urease, which consists
of 3 separate structural genes in Klebsiella aerogenes that are fused together on a single
polypeptide in legumes (Goldraij et al., 2003), and acetyl-CoA carboxylase, which is
present as a multimeric complex in bacteria, but consists of a single multifunctional
polypeptide in eukaryotes (Nikolau et al., 2003). Apparently, the same regulatory
advantages that are in play in those systems are important in DGTS biosynthesis as well,
i.e. that having one gene to encode the machinery for the entire pathway presumably

alleviates the coordination required for proper expression of multiple individual genes.

98

References

Benning, C., Huang, Z.H., and Gage, DA. (1995). Accumulation of a novel glycolipid
and a betaine lipid in cells of Rhodobacter sphaeroides grown under phosphate
limitation. Arch. Biochem. Biophys. 31 7, 103-111.

DDrmann, P., Balbo, I., and Benning, C. (1999). Arabidopsis galactolipid biosynthesis
and lipid trafficking mediated by DGDl. Science 284, 2181-2184.

Giroud, C., Gerber, A., and Eichenberger, W. (1988). Lipids of Chlamydomonas-
reinhardtii - Analysis of molecular species and intracellular site(s) of biosynthesis. Plant
and Cell Physiology 29, 587-595.

Goldraij, A., Beamer, L.J., and Polacco, J.C. (2003). Interallelic complementation at the
ubiquitous urease coding locus of soybean. Plant Physiol 132, 1801-1810.

Grossman, A.R., Harris, E.E., Hauser, C., Lefebvre, P.A., Martinez, D., Rokhsar, D.,
Shrager, J., Silflow, C.D., Stern, D., Vallon, 0., and Zhang, Z. (2003). Chlamydomonas
reinhardtii at the Crossroads of Genomics. Eukaryot. Cell 2, 1137-1150.

Harris, EH. (1989). The Chlamydomonas Source Book. (New York: Academic Press).

Kagan, RM. and Clarke, S. (1994). Widespread occurrence of three sequence motifs in
diverse S- adenosylmethionine-dependent methyltransferases suggests a common
structure for these enzymes. Arch. Biochem. Biophys. 310, 417-427.

Klug, RM. and Benning, C. (2001). Two enzymes of diacylglyceryl-O-4'-(N,N,N,-
trimethyl)homoserine biosynthesis are encoded by btaA and btaB in the purple bacterium
Rhodobacter sphaeroides. Proc. Natl. Acad. Sci. U. S. A 98, 5910-5915.

Moore, T.S., Du, Z.R., and Chen, Z. (2001). Membrane lipid biosynthesis in
Chlamydomonas reinhardtii. In vitro biosynthesis of diacylglyceryltrimethylhomoserine.
Plant Physiol. 125, 423-429.

Nikolau,B.J., Ohlrogge,].B., and Wurtele,E.S. (2003). Plant biotin-containing
carboxylases. Arch. Biochem. Biophys. 414, 211-222.

Sambrook, J., Fritsch, BF, and Maniatis, T. (1989). Molecular Cloning: A Laboratory
Manual. (Plainview, NY: Cold Spring Harbor Laboratory Press).

99

Chapter 4
Molecular biology of lipid metabolism in Chlamydomonas reinhardtii. Analysis of
the draft genome for pathways of lipid biosynthesis, and prospects for functional

characterization of metabolic pathways by RNA interference. 1

 

' Parts of this work involving transformation of Chlamydomonas were performed in collaboration with Dr.
Barbara Sears, Department of Plant Biology, MSU. The annotation of genes involved in lipid metabolism
was canied out as part of the Chlamydomonas genome annotation project, under the direction of Drs.
Arthur Grossmann (Carnegie Institution of Washington, Stanford University) and Dan Rokhsar (DOE-Joint

Genome Institute).

100

Abstract

Lipid metabolism in seed-plants has been intensively studied at the biochemical, genetic,
and genomic levels, and knowledge regarding the identity of genes encoding components
of the major fatty acid and membrane lipid biosynthetic pathways is nearing completion.
We now present a study of fatty acid and glycerolipid biosynthesis in an algal model,
based on EST and genomic sequencing of Chlamydomonas reinhardtii. Genes encoding
proteins involved in membrane biogenesis were analyzed on the basis of similarity to
known proteins, and were organized so as to reconstruct the major pathways of
glycerolipid synthesis in Chlamydomonas. This analysis accounts for the majority of
genes involved in anabolic reactions of membrane lipid biosynthesis, and draws
comparisons and contrasts between these pathways in Chlamydomonas and higher plants.
As a case study in the functional analysis of the annotated genes, we used RNA
interference to silence the BTAI gene, encoding the machinery for synthesis of the
betaine lipid diacylglyceryl-N,N,N-trimethylhomoserine (DGTS), the major component
of extraplastidic membranes. Silencing of this gene resulted in an attenuation of growth
rate and increased sensitivity to heat stress, but showed little effect on the lipid
composition of the cells. This may indicate a strong tendency toward membrane
homeostasis, and that the reduced capacity for membrane biogenesis is compensated by a

reduction in growth rate.

101

Introduction

Knowledge of genes and proteins involved in lipid metabolism in plants has increased at
a phenomenal rate over the past decade. A large number of genes encoding enzymes of
glycerolipid biosynthesis and fatty acid desaturation have been identified by genetic and
biochemical means, e.g. references (Shimojima et al., 1997; Xu et al., 2002), and even
lipid trafficking between the endoplasmic reticulum and plastid is beginning to be
unraveled by forward genetic approaches (Xu et al., 2003). In addition, the availability of
the Arabidopsis genome sequence (The Arabidopsis Genome Initiative, 2001) has
facilitated the annotaan of many uncharacterized gene products which are likely to have
activity in lipid biosynthesis, trafficking, and catabolism (Beisson et al., 2003). Although
Arabidopsis is undoubtedly the most widely used higher plant model system currently
under investigation, the eukaryotic green alga Chlamydomonas reinhardtii is a well
established model for many processes such as photosynthesis (Niyogi, 1999), phototaxis
and flagellar function (Silflow and Lefebvre, 2001), post-transcriptional gene silencing
(Wu-Scharf et al., 2000), and nutrient acquisition (Davies et al., 1996). The recent large-
scale EST projects and sequencing of the C. reinhardtii genome (Shrager et al., 2003;
Grossman et al., 2003), as well as the development of insertional mutagenesis (Tam and
Lefebvre, 1993), RNA interference (RNAi) methods (Sineshchekov et al., 2002;
Fuhnnann et al., 2001), and a molecular map (Kathir et al., 2003) make Chlamydomonas
an attractive model in which to study gene function by both forward and reverse genetics
in a unicellular photosynthetic eukaryote. Additionally, Chlamydomonas is ideal as a
model organism in which to study the biosynthesis of non-phosphorous lipids such as the

plastidic galacto- and sulfolipids, and the extraplastidic betaine lipid diacylg1yceryl-

102

N,N,N-trimethylhomoserine (DGTS) (Giroud and Eichenberger, 1989; Moore et al.,
2001), as well as phospholipids such as phosphatidylethanolamine (PtdEtn)(Yang et al.,
2004). We have begun to use Chlamydomonas as a model for plant lipid metabolism,
first using a high-throughput robotic screening methodology to find mutants with defects
in lipid metabolism (Riekhof et al., 2003; Chapter 5), and using genomic resources to
identify and characterize the pathway of betaine lipid biosynthesis (Chapter 3). We now
present a study of genes involved in acyl-lipid metabolism, as gleaned from annotation of
the v2.0 draft genome sequence available at http://genome.jgi-psf.org/chlamy/. Enzymes
of fatty acid biosynthesis, glycerolipid synthesis, and fatty acid desaturation were
identified and organized so as to reconstruct the lipid biosynthesis pathways and posit
their subcellular localization. As a case study in the functional analysis of these annotated
genes, we used RNAi to suppress the expression of the BT A1 gene, encoding the protein
responsible for synthesis of the major extraplastidic glycerolipid DGTS (Chapter 3).
Suppression of BT A1 led to pleiotropic effects on growth, while showing little effect on
lipid composition. We discuss these results in the context of the coordinated regulation

of membrane biogenesis and cellular growth.

Materials and Methods

Annotation of C. reinhardtii genes involved in lipid metabolism. Version 2.0 of the C.
reinhardtii whole genome shotgun sequence assembly is available at http://genome.jgi-
psf.org/chlamy. Genes encoding enzymes and other components of lipid metabolism
(e.g. acyl-carrier proteins) were manually annotated based on keyword searches executed

against the automatically annotated gene loci (Grossman et al., 2003), followed by

103

verification of similarity by BLASTP searches (Altschul et al., 1997) and CLUSTALW
multiple sequence alignments (Thompson et al., 1994) with verified protein sequences.
In addition, direct BLASTP and TBLASTN searches of the v2 .0 assembly were carried
out with known protein sequences from plants, animals, fungi, and bacterial sources to
add additional evidence and to identify genes that were incorrectly annotated by the
automated system.

For prediction of subcellular localization, the programs TargetP and ChloroP were
used (Emanuelsson et al., 1999; Emanuelsson et al., 2000). In cases where the N-
terrninus was not available due to sequence gaps and/or ambiguity in prediction of the
first exon, tentative localization was predicted on the basis of their clustering with known

isoforms from other species.

Construction of RNAi lines for suppression of CrBTA I. As the basis for the target of
the anti-BTAI dsRNA construct, we chose a long EST from the middle section of the
gene (GenBank accession BU651274). Briefly, nucleotides 1-467 were fused in the
sense orientation to the RBCS2 5’ promoter element with overlap-extension (OE) PCR,
and this construct was ligated to the anti-sense oriented bases 1-697, forming a 467 bp
dsRNA stern and a 230 bp loop (Figure 4.3).

Pfu Turbo DNA polymerase (Stratagene) was used according to the
manufacturers protocol in all PCR reactions described below, except that 10% (v/v)
DMSO was added to all reactions to remedy problems associated with amplification of
the highly G -iC rich sequences of C reinhardtii. The RBCSZ 5’ element was amplified

from plasmid pSP124S (Lumbreras et al., 1998) with the primers RBCS fw-

104

CGAGCTCAAATGCCAGAAGGAGCG (SacI underlined), and RBCS rev-
CCTGGGTGTTCTGCTCC'TCACCTGGCCATI‘TI‘AAG. For RT-PCR of BT A1
sequences, RNA was isolated from mid-log phase C. reinhardtii strain CC125 with Trizol
reagent (Invitrogen) according to the manufacturer’s instructions, and 1.0 pg total RNA
was used for first-strand cDNA synthesis with SuperScript II RNase H' reverse
transcriptase (primed with oligo-dT') according to the manufacturer’s protocol. 2 pl
aliquots of the RT reaction were used as template in 50 pl PCR reactions. The sense-
oriented portion of CrB TA] was amplified with the primers “BTAI sense fw”-
CTTAAAATGGCCAGGTGAGGAGCAGAACACCCAGG; and “BTAI sense rev”-
CGGATCCGCGCT'TGGACCAGAAGT (BamHI underlined). For overlap-extension
PCR, the products of “RBCS fw/rev” and “BTAI sense fw/rev” were gel purified, and 10
ng of each product were mixed for use as the template in amplification of the fusion
product with “RBCS fw” and “BTAl sense rev” primers. Amplification of the antisense
portion was with primers “BTAI anti fw”- CGGATCCCACCAGGCTCACGAACTT
(8amHI underlined), and “BTAI anti rev”- CGAATT‘CGGAGCAGAACACCCAGG
(EcoRI underlined). All products were gel purified and cloned into the EcoRV site of
pBluescript SK +and sequenced at the MSU genonrics technology support facility. For
RNAi plasmid constmction, the RBCS::sense fusion and antisense segment were excised
and cloned into pSP124S by a three piece ligation as indicated by the Sacl, 8amHI, and

EcoRI sites in Figure 4.3, to give plasmid pBTAI-RNAi.

Transformation with RNAi construct and growth of cells.

105

The pSP124S plasmid and pBTAI-RNAi construct described above were used to
transform strain dw15.l (mt+, nit1-305, cw15) by the glass bead method (Kindle, 1998),
except that TAP medium was used for cell growth, and transformation mixtures were
plated in a slurry of cornstarch, as described (Shimogawara et al., 1998), onto TAP +1 %
yeast extract (TAPY) containing 10 pg/ml Zeocin (Invitrogen). Approximately 50
transformant colonies of each construct were propagated, and initial tests for growth
phenotype were carried out in 1.5 ml TAP cultures in 24 well nricrotiter dishes.

For all physiological and growth experiments, cultures were grown in TAP
medium with shaking (200 rpm) under constant cool white fluorescent light, at 24° C.
Analysis of growth rates was carried out by inoculation of cells into 50 ml TAP medium
to an initial density of OD750 =0.05, and growth was measured by taking the OD750 at
regular intervals.

Temperature sensitivity was assessed by streaking a loop of freshly grown cells as
patches onto TAPY plates, followed by incubation of the plates in the dark at 24, 37, or
42° C for 16 or 40 h. Following the challenge at high temperature, the plates were

incubated in the light at 24° C for 3 days to assess survival.

RNA blot analysis

For the analysis of transcript levels in the RNAi and wild type strains, 50 m1
cultures were inoculated and grown for two days, as in the growth rate analysis. Cultures
were harvested by centrifugation, and total RNA was extracted fiom cell pellets with
Trizol reagent (Invitrogen) according to the manufacturer’s instructions. 5 pg of RNA

was separated on a forrnaldehyde-agarose gel followed by blotting onto HyBond

106

(DuPont) membranes, and probing with the full length CrBT A1 open reading frame

(GenBank accession AY656806) (Sambrook et al., 1989).

Analysis of lipids

For the quantitative analysis of polar lipids, 50 ml cultures of the indicated strains
were grown to late log phase in TAP medium, as for the growth rate analysis. Cells were
collected by centrifugation and lipids were extracted from the cell pellets by vigorous
vortexing in 4 ml of chloroform: methanol (1:1 , v/v), followed by phase separation with 1
ml of 1 M KC1, 0.2 M H3PO4. For separation of polar lipid classes, a 2-dimensional thin-
layer chromatography (silica 60 plates, J .T. Baker) system was employed consisting of
chloroform: methanol: water (65/25/4, v/v) in the first dimension and chloroform:
acetone: methanol: glacial acetic acid: water (10/4/2/2/0.75, v/v) in the second dimension.
Polar lipid species were detected by brief exposure to iodine vapor, and the spots
containing each lipid were scraped into a tube with a razor blade, followed by addition of
5.0 pg myristic acid (C 14:0) as an internal standard. This preparation was then
transmethylated and analyzed by GC as described (Rossak et al., 1997). As previously
reported (Giroud et al., 1988), myristic acid is present in the total fatty acid pool (data not
shown), however this fatty acid was absent from the polar lipid classes that we measured,

and therefore 14:0 was suitable as an internal standard for quantification.

107

Results
Annotation of lipid biosynthesis genes in the Chlamydomonas genome.

We analyzed lipid biosynthetic pathways in Chlamydomonas by searching the
whole genome shotgun sequence for proteins with significant similarity to known
enzymes from plants and other organisms. Figures 4.1 and 4.2 and the accompanying
Table 1 present a scheme for membrane lipid biosynthesis, starting with fatty acid
synthesis in
the plastid (Figure 4.1, steps 1-4 and 16), plastidic glycerolipid biosynthesis (Figure 4.1,
steps 5-15), extraplastidic phospholipid and betaine lipid biosynthesis (Figure 4.1, steps

16-31), and fatty acid desaturation, (Figure 4.2, steps 32-40).

Lipid metabolism in plastids.

Genes encoding enzymes of plastidic lipid metabolism are readily identifiable in
the genome, and appear to be most similar to those of higher plants. For example, it is
clear that all components of the multimeric bacterial-type acetyl-CoA carboxylase and
fatty acid synthase (FAS) complexes are accounted for, similar to those that have been
described for higher plant plastids. Interestingly, many of the enzymes central to fatty
acid biosynthesis, e.g. the 3-hydroxyacyl-ACP-dehydratase and enoyl-CoA-reductase
components of the type II FAS, are apparently present in a single copy in the genome,
and have high and roughly equal probabilities of being targeted to both mitochondria and
chloroplasts as judged by TargetP analysis (Emanuelsson et al., 2000).

The pathway of sulfoquinovosyldiacylglycerol (SQDG) biosynthesis has been

thoroughly characterized in Arabidopsis and shown to be the function of two gene

108

13 9
—->- @«Hdcmr

8
UDP-Gal 1L1 Lu UDP.SQ 4—1 UDP- Gtc L G-3-P

fiDAG <I—1-o— PtdOH —-> COP-DAG

7 Acetyl—CDA
-UDP-Gai ‘ l 1
Melon
Iyso-PtdOH J 5 3 W
ACP(2)
Malon ACP
5 4 Y"

 

 

 

 

FreeFA
I17 Glc-i-P
AcleoA 30
Me! lyso-PtdOi-l G-3-P
AdoMet PtdOH 20-{1 COP-DAG 28

29
P-Etn —27-> 23co:.\2>-

Figure 4.1: Pathways of lipid biosynthesis which are known or hypothesized to occur in

Chlamydomonas, and their subcellular localization. Numbers in bold conespond to the

activities summarized in TABLE 1, which also correlates gene models with their

respective putative activities.

109

32
18:0—ACP —> 18:1-ACP
16:0-ACP}_1

l

 

 

Pthro MGDG
16:0/16:0 18:1/16:0\
I” SQDG I“ DGDG
18:1/16:1 18:1/16:0 18:1/16:1 18:1/16:0
135 351 I 35 135
1822/1610 18:2/16:2 18:2/16:0
133 361 I 36 l 36
18:3/16:0 18:3/16:3 18:3/16:0
I 37
Plastid 18:3/16:4
ER DGTS PtdEtn
16:0/18:1 18:0/18:1
133 Isa
16:0/18:2 18:0/18:2
V a V a
16:0/”8:3 16:0/(11823 1810/11813 1820/(11833
39 A 39 A
16:0/18:4 1820/1824

Figure 4.2: Pathways of acyl chain desaturation, with numbers in bold serving to cross

reference the specific reaction with the gene model(s) in Table 1.

110

Table 4.1: Identification of the reactions diagrammed in Figures 4.1 and 42, and
assignment of candidate genes on the basis of identity with known proteins and
subcellular prediction based on analysis of the N-tenninus. Candidates for dual targeting

to chloroplast and mitochondria are highlighted.

lll

PLASTID AND MITOCHONDRIAL PATHWAYS

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Reaction Description Gene model # ESTs
Multimeric Acetyl-CoA carboxylase
1 components:
13 Alpha-carboxyltransferase 0-160059 23
1b Beta-carboxyltransferase C_1 150054 39
1c Biotin carboxylase C_2010018 18
1d Biotin carboxyl carrier protein C_590042 16
C_250035 27
2 Acyl carrier protein C_650040 38
C_210028 51
3 Malonyl-CoA: ACP transacyiase C_240122 4
C_7350001 16
4 Type II Fatty acid synthase components:
48 3-ketoacyl-ACP synthase C_140066 17
C_720014 7
C_1320034 2
4b 3-ketoacyl-ACP reductase C_460059 19
4c 3-hydroxyacyl-ACP dehydratase C_120121 7
4d enoyl-ACP-reductase C_190023 64
5 Gcheroi-B-P: Acyl-ACP acyltransferase C_1210022 8
6 iyso-phosphatidate: Acyl-ACP acyltransferase no candidate
7 CDP-DAG synthetase C_30067 12
8 phosphatidylglycerophosphate synthase C_140036 21
C_2150006 4
9 phosphatidylglycerophosphate phosphatase no candidate
10 phosphatidate phosphatase no candidate
11 UDP-sulfoquinovose synthase C_60107 46
12 sulfolipid synthase C_250126 5
C_650023 0
13 sulfolipid 2'-O-acyltransferase no candidate
14 monogalactosyldiacylglycerol synthase C_21260001 3
15 digalactosyldiacylglycerol synthase C_230128 1
16 Acyl-ACP thiolase C_200208 7
CYTOSOLIC PATHWAYS
17 long chain acyl-CoA synthetase C_1580041 11
C_7940001 0
18 Glycerol-3-P: Acyl-CoA acyltransferase no candidate
19 Iyso-phosphatidate: Acyl-CoA acyltransferase no candidate
20 phosphatidate phosphatase C_1 170013 1
C_300097 4
21 CDP-DAG synthetase C_630023 4
22 AdoMet synthetase C_750005 82
23 Betaine lipid synthase C_570065 42
24 CDP-Etn: DAG Etn phosphotransferase C_960032 2
25 Serine decarboxylase C_250130 43
26 Ethanolamine kinase C_290062 0
27 CTP: phosphoethanolamine midylyltransferase C_960052 15
28 Phsphatidyiglycerophosphate synthase C_1350041 0

 

112

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

29 phosphatidylglycerophosphate phosphatase no candidate
30 lnositol-3-phosphate synthase C_860083 31
31 COP-DAG: lnositol phosphotransferase C_260084 3
FATTY ACID DESATURASES
Plastidic lsofonns (as judfl by ChloroP analysis)
32 Stearoyl-ACP-AQ—desaturase C_170008 10
33 Pthro palmitate-ABt-desaturase no candidate
34 MGDG palmitate-A7-desaturase C_330009 70
35 03-6 desaturase C_790066 264
C_290028 0
36 w—3 desaturase C_250125 39
37 MGDG 16 carbon A4—desaturase no candidate
Cytosolic isoforms
38 Oleate desaturase C_590003 20
C_4630001 0
39 Linoleate desaturase C_250037 4
40 Linoleate/linolenate-AS—desaturase C_1000063 3
11000033 4

 

113

products, namely SQDI and SQD2 (Essigmann et al., 1998; Yu et al., 2002). SQDI has
been characterized in Chlamydomonas by both insertional inactivation (Riekhof et al.,
2003) and complementation of a sulfolipid deficient squ cyanobacterial mutant (Sato et
al., 2003), and the SQDI gene is highly expressed as judged by EST abundance. This
apparent dispmportionately high mRNA abundance might be due to the relatively large
amount of SQDG in Chlamydomonas as compared to Arabidopsis, coupled with the fact
that Km for the reaction catalyzed by SQDI is very low (Sanda et al., 2001), which might
necessitate increased abundance of the enzyme.

Genes encoding enzymes of galactolipid biosynthesis are also present in the
genome, with MGDl (monogalactosyldiacylglycerol synthase) and DGDl
(digalactosyldiacylglycerol synthase) genes present in the genome as single copies, as
opposed to three MGDG and two DGDG synthase isoforms in Arabidopsis (Awai et al.,
2001; Kelly and Dormann, 2002).

Several functions in plastidic lipid biosynthesis are without candidate genes
including the phosphatidylglycerophosphate phosphatase (Figure 4.1, step 9), which is
unknown in plants (Beisson et al., 2003), and a putative activity utilizing SQDG and an
unidentified acyl donor to form 2’-0-acyl-SQDG (ASQD, Figure 4.1, step 13). This
species has been shown to be a minor component of Chlamydomonas lipids, however it is
apparently absent from higher plants (Riekhof et al., 2003; Chapter 5). While no
candidate can be predicted, Several serine carboxypeptidase-like enzymes responsible for
catalyzing similar transacylation reactions in phenylpropanoid metabolism have been
identified (Lehfeldt et al., 2000; Shirley et al., 2001), and it is conceivable that a similar

mechanism is operative in ASQD biosynthesis. Additionally, phosphatidate phosphatase

114

(Figure 4.1, step 10) has at least two recognizable homologs in the genome, however

neither of them are predicted to contain a plastid transit peptide.

Lipid metabolism in the endomembrane system.

Cytosolic membrane lipid metabolism in Chlamydomonas differs from that of
higher plants by both the presence and absence of certain central pathways and branches
of pathways. The most notable absence is phosphoethanolamine methyltransferase,
which provides the pathway for methylation of phosphoethanolamine to form
phosphocholine. This activity is responsible for the generation of phosphocholine for
conversion into CDP-choline and then PtdCho, and the enzymes from spinach and
Arabidopsis have been functionally identified by their ability to rescue a ch02 mutant of
Schizosaccharomyces pombe or an 0pi3 mutant of Saccharomyces cerevisiae,
respectively (Nuccio et al., 2000; Bolognese and McGraw, 2000). The apparent lack of a
pathway for PtdCho precursor biosynthesis in Chlamydomonas, as well as a lack of the
PtdEtn methylation pathway, correlates with a lack of this lipid from cellular membranes
(Giroud et al., 1988).

PtdSer is a major precursor of PtdEtn in yeast, bacteria, and plants through the
action of PtdSer decarboxylase (Voelker, 1997), however Chlamydomonas has been
reported to lack PtdSer as a component of its membranes (Giroud et al., 1988). This fact
is evident in the genome as an apparent lack of both phosphatidylserine synthase and
phospholipid base exchange enzymes. This suggests that the biosynthesis of PtdEtn is
the function of a single pathway, consisting of serine decarboxylase (Rontein et al., 2001)

to generate Etn (Figure 4.1, step 25) followed by phosphorylation of

115

Etn (Figure 4.1, step 26), activation with CTP to form CDP-Etn (Figure 4.1, step 27), and
transfer of the phosphoethanolamine moiety to DAG, forming PtdEtn (Figure 4.1, step
24). The Chlamydomonas CTP: phosphoethanolamine cytidylyl transferase enzyme has
recently been characterized through expression in E. coli and biochemical
characterization of the recombinant protein (Yang et al., 2004).

In the absence of PtdCho, the betaine lipid DGTS has been proposed as the
bilayer forming and major structural component of extraplastidic membranes (Giroud et
al., 1988; Sato, 1992). DGTS biosynthesis is catalyzed by a multifunctional enzyme
utilizing DAG and 4 equivalents of AdoMet to form this lipid (Chapter 3) and this gene is
annotated as BTAI (Figure 4.1, step 23).

Certain pathways are assumed to be present in multiple compartments. For
example, Pthro biosynthesis is proposed to be present in the plastid, mitochondria, and
endoplasmic reticulum, and 3 isoforms of the phosphatidylglycerophosphate synthase are
present in the genome. Two of these (Figure 4.1, step 8) are expressed as judged by EST
analysis and both contain a predicted plastid or mitochondrial targeting sequence. The
other is predicted to be cytosolic, but is not represented in the EST database. This may
correlate with the fact that the mass of Pthro present in the cytosol is predicted to be
much less than that in the plastid (Giroud et al., 1988), and thus the gene encoding the
cytosolic activity might not be as highly expressed as those directing Pthro synthesis in
the organelles.

Fatty acid desaturases.
In addition to the synthesis of the glycerolipids themselves, fatty acid desaturases

(FAD) were identified and annotated in this survey. Chlamydomonas contains several

116

fatty acids not found in Arabidopsis, namely 16:4A4’7'10‘l3, which is found predominantly
in plastidic MGDG, as well as 18:3’5‘5’9’12 and 18:4A5'9‘12'15 , the bulk of which are present
as components of DGTS (Giroud et al., 1988; Giroud and Eichenberger, 1989). The FAD
complement of Arabidopsis has been well characterized at the genetic and biochemical
levels (Ohlrogge and Browse, 1995), and homologs are present in the Chlamydomonas
genome, as outlined in Figure 4.2. Of note are the putative extraplastidic (06 and (n3
desaturases (Figure 42, steps 38-39), as well as two additional, apparently extraplastidic
FAD-like enzymes that are most similar to the plastidic palmitate specific A7 desaturase
(FADS).

Likewise, there are predicted plastidic homologs of the Arabidopsis FADS, FAD6
and FAD7/8 enzymes ((Ohlrogge and Browse, 1995); Figure 42, steps 34-36), and the
plastidic 03-6 desaturase of Chlamydomonas (Figure 42, step 35) has been previously
identified in a mutant screen for high-chlorophyll fluorescence mutants, and was
subsequently cloned and characterized (Sato et al., 1995; Sato et al., 1997). We were
unable to assign a candidate for the A4 desaturase involved in generation of 16:4’3‘1'7'10'l3
(Figure 4.2, step 37), although the gene encoding this protein would likely be expressed
at a high level, given the EST abundance for the other enzymes in this desaturation

pathway.
Silencing of the B TAI gene is detrimental to growth.

To begin the functional analysis of genes annotated as having putative functions in acyl—

lipid biosynthesis, we chose to target ET A] for silencing with a dsRNA hairpin

117

 

 

 

 

 

ir—neuw 1% n

I r r
Sacl BamHl EcoRl

11111111111: 9230bpstemloop
467bpdsRNA

 

 

Figure 4.3: Schematic of RNAi construct. A long EST from the middle of the BTAl
gene was used as the basis for design of a dsRNA hairpin, as in described Materials and
Methods. The parental vector was pSP124S, and the vector and RNAi constructs were
transformed into Chlamydomonas dw15.1, and transforrnants selected on Zeocin

(Invitrogen).

118

 

 
   
   

 

 

 

2 1 +pSP-1
+pSP-2
-o—RNAi-29

15‘ +RNAi-31

g +RNAl-35
o
o 1*

0.54

0 . .

0 20 4O 60 80
Time (h)

Figure 4.4: Growth curves of RNAi lines and vector controls. Cultures were inoculated

at an initial ODvso of 0.05, and growth was monitored by measuring the OD750 at regular

intervals.

119

construct (Figure 4.3). Transformation of this plasmid and the pSP124S vector into
Chlamydomonas strain dw15.l resulted in many Zeocin resistant colonies, and
approximately 50 transformants of both the vector and RNAi constructs were subcultured
and analyzed for defects in growth and lipid metabolism. Several lines of the RNAi
transformants, designated i29, BI, and i35, showed alterations in growth rate and
temperature sensitivity, growing at between 20 to 40 % of the rate of the control lines
containing only vector (Figure 4.4). The growth of these lines on agar plates containing 1
% yeast extract was also slower than the vector controls, but the growth phenotype was
slightly more pronounced in liquid culture. The growth rate defect also correlated with
more pronounced temperature sensitivity. Lines i29, i3 1 , and BS were unable to survive
incubation for 16 h at 42° C, while lines pSPl , pSP2 (vector controls), and i15
(containing the RNAi-construct but showing no growth phenotype) survived this
treatment. i35 was unable to survive 40 h at 37° C, whereas the other strains tolerated
this treatment, indicating that this line was both the most severely inhibited in growth rate
and tolerance to high temperature.

Northern analysis of steady-state mRNA levels was conducted on the slow-
growing RNAi lines and the vector controls. Cells were incubated for 48 h in 50 ml TAP
cultures as for the growth rate analysis, to ensure that each strain was growing at its
maximal rate. As shown in Figure 45, the growth phenotype correlates with a reduction
of mRNA levels. The silencing of this gene was not complete, but one would not expect
to recover strongly suppressed lines of what is likely to be an essential gene.

Figure 4.6 gives the polar lipid composition of the vector and RNAi lines. To our

surprise, the relative amounts of the different polar lipid classes were essentially

120

 

Figure 4.5: 87' A] gene expression in RNAi lines. Cultures of pSP124S transformed
controls and RNAi transforrnants were cultivated as in the growth curve experiment, and
the cells were harvested after 48 h, followed by RNA isolation and northern blotting with
10 pg of RNA per lane. The respective lines are indicated above each lane, and the
ethidium bromide stained gel serves as an indicator of equal loading as judged by rRNA

intensity.

121

 

 

Cl SP1

 

 

 

 

 

 

 

MGDG DGDG Pthro SQDG DGTS PtdEtn Ptdlns

Figure. 4.6: Lipid composition of RNAi lines. Cultures were inoculated and grown as for
the growth rate analysis, and were harvested after 72 h of incubation. Lipids were
extracted from the cell pellets and resolved by TLC, followed by quantitative analysis by
GC. Values are expressed as mole percentages (mol/ 100mol), and the quantities given
are averages of two independent determinations, with error bars representing the range of

the 2 values.

122

 

24° c

 

37°C

 

42°C

 

 

 

 

 

Figure 4.7: Temperature sensitivity of RNAi lines. The respective lines were freshly
streaked onto TAPY plates and kept in the dark for 16 or 40 h at the indicated
temperature. Following this treatment, plates were kept in the light at 24° C for several

days before photographing the plates.

123

unchanged in the RNAi suppressors of ET A]. In addition, the total fatty acid profile of

whole cells was largely unchanged in the 5 independent lines analyzed (data not shown).

Discussion

This work was initiated to explore lipid metabolism in Chlamydomonas by examination
of the whole genome shotgun sequence for predicted proteins with activity in acyl-lipid
metabolism. In addition, we used the hairpin RNAi method to investigate the functional
consequences of suppressing expression of the ETA] gene, encoding the multifunctional
enzyme for synthesis of the betaine lipid DGTS. This experiment also serves as a case
study in the reverse-genetic functional analysis of these annotated genes, and shows the
potential for rapidly determining the consequences of down-regulating other genes
annotated in this study.

During the course of the annotation and analysis phase, several themes became
apparent. First, the core pathways of fatty acid biosynthesis and glycerolipid assembly in
Chlamydomonas appear to be somewhat simpler than those of Arabidopsis, both in the
presence and/or absence of certain pathways, and in the size of the gene families that
represent the various activities. A case in point is that of galactolipid synthesis. In the
Chlamydomonas genome, only one MGDG-synthase (MGDI) and one DGDG-synthase
(DGDI) is present, as opposed to three MGD isoforms (Awai et al., 2001) and two DGD
isoforms (Kelly and Dormann, 2002) in the Arabidopsis genome. This may be thought of
as a direct result of (or accompaniment to) the lack of PtdCho and its presumed
replacement by the non-phosphorous analog DGTS in Chlamydomonas. The logic

behind this statement is as follows. In Arabidopsis, phosphate deprivation has been

124

shown to upregulate the expression of MGD2, MGD3, and DGD2, concomitant with a
loss of PtdCho and accumulation of DGDG in extraplastidic membranes (Ddrmann and
Benning, 2002). This scenario has also been observed in phosphate starved Oat, where
DGDG, presumably produced via the phosphate-starvation inducible biosynthetic
pathway, replaces a large proportion of PtdCho in the plasma membrane (Andersson et
al., 2003). This phenomenon is presumed to be an adaptation to phosphate limitation, in
that a nonphosphorous lipid is substituting for a phospholipid, thus allowing reallocation
of the limiting nutrient into more critical components, such as nucleic acids. Given that
Chlamydomonas has no need to replace PtdCho with DGDG due to the presence of
DGTS, the phosphate-inducible pathway of DGDG biosynthesis may have been rendered
superfluous and lost over the course of evolution (assuming it was present in the common
ancestor of angiosperms and Chlamydomonas), or may simply have evolved only in
higher plants.

Another theme is apparent in the biosynthesis of fatty acids, in that some of the
core components of this process appear to be targeted to both the plastid and
mitochondrion. Several lines of evidence support this conjecture. First, it is known from
studies on isolated organelles that fatty acid synthesis occurs in both plastids and
mitochondria, the plastid being responsible for supplying the bulk of fatty acid
biosynthesis to supply precursors for membrane lipid synthesis, and the mitochondrial
pathway being a function of lipoic acid biosynthesis, a critical prosthetic group for certain
mitochondrial enzymes. Several of the activities that are central to fatty acid synthesis,
e.g components of the multimeric acetyl-CoA carboxylase (ACCase) and the type II fatty

acid synthase, as well as acyl carrier proteins, are present in only one copy (two in the

125

case of ACP) in the Chlamydomonas genome. This leads to the conclusion that, since the
activities are present in multiple compartments, these proteins may be dually targeted
both to mitochondria and to plastids, a phenomenon with a growing number of
precedents, especially with proteins involved in core metabolic processes such as
nucleotide biosynthesis and DNA replication, e.g. (Chabregas etal., 2001; Wall et al.,
2004). Indeed, the Chlamydomonas proteins in question have a relatively high
probability of dual targeting to mitochondria and plastids, as judged by Target? analysis.
The recent finding of a potentially dual targeted multifunctional ACCase in grasses
(Focke et al., 2003) lends credence to this hypothesis. Additionally, Beisson et a1. (2003)
were unable to predict candidates for the mitochondrial 3-ketoacyl-ACP-reductase and 3-
hydroxyacyl-CoA-dehydratase activities of Arabidopsis. Given that these activities have
been demonstrated implicitly by the fact that mitochondria can synthesize fatty acids, it is
reasonable to posit that dual targeting of these proteins could be operative in Arabidopsis
as well.

Fatty acid desaturation has garnered much attention due to the potential for
industrial uses of novel plant fatty acids, and the analysis of Chlamydomonas FAD
enzymes presented here is based largely on work from Arabidopsis (reviewed by
Ohlrogge and Browse, ( 1995)). Clear homologs of the Arabidopsis FADs are present in
our survey. In addition, two putative cytosolic FADs are present which are somewhat
divergent from those that have clear similarity to characterized isoforms in other
organisms. Chlamydomonas synthesizes at least 2 extraplastidic fatty acids not found in
Arabidopsis, and these proteins might be involved in the synthesis of these atypical acyl

species (Figure 4.2, step 39). One approach to verifying hypotheses regarding the

126

function of these putative FADs would be to use RNAi to silence the genes or,
alternatively, to express the genes in Arabidopsis. For example, expression of the
putative Chlamydomonas ER desaturases (Figure 4.2, step 40) in Arabidopsis has the
potential to both show the function of these genes, and to demonstrate the consequences
of increasing the desaturation level of Arabidopsis ER lipids.

Given the similarities and differences between Chlamydomonas and Arabidopsis,
we used RNAi to investigate the consequences of down-regulating BTAI , encoding the
protein responsible for synthesis of the PtdCho analog DGTS, as described above. We
presumed that, given the role of PtdCho in animal and fungal cells, silencing of the ETA]
gene would be detrimental to the growth of the Chlamydomonas cells, and that this would
correlate with a decrease in DGTS content. In fact, most of the transformants showed no
phenotype and probably do not express the dsRNA hairpin. However, several strains
(approximately 10% of those propagated) showed a reduction in growth rate on both agar
plates and in liquid culture (Figure 4.4). Northern analysis (Figure 4.5) reveals a
significant decrease in the steady state transcript levels of the BT A1 gene in the RNAi
suppressor lines that show a growth phenotype. This rate of recovery of suppressors in
an RNAi experiment is in line with other values reported in the literature (Soupene et al.,
2004). The growth defects of the lines chosen for analysis probably represent
comparatively mild phenotypes, as there were also many very small transformant
colonies that we were unable to propagate or that died after several rounds of re-
streaking, and which may have represented more severe alleles. Nonetheless, we
investigated the growth properties and lipid profiles of three of the viable, yet poorly

growing lines.

127

PtdCho biosynthesis defects are often accompanied by temperature sensitive
growth and developmental phenotypes (Howe et al., 2002; Mon et al., 2002), and Figure
4.7 shows that this is also the case for suppressed expression of DGTS biosynthesis in the
BTAI-RNAi lines. The degree of temperature sensitivity correlated with the severity of
the growth rate defect, with lines i29, i3 1 , and BS being unable to survive 16 h at 42° C,
and line i35 showing the most stringent temperature sensitivity, losing viability after 40 h
at 37° C. Many studies have shown that a shift in temperature often induces changes in
the composition of membranes and in the activities of lipid biosynthetic enzymes, e.g in
S. cerevisiae, a shift in temperature from 24° to 37° was shown to induce an increased
capacity for PtdCho biosynthesis via the CDP-Cho pathway (Dowd et al., 2001; Howe et
al., 2002). In our experiment, by analogy with the situation in yeast, the temperature
sensitive phenotype may result from an inability of the cells to respond to the heat stress
by altering membrane lipid composition in a timely manner. While this seems a plausible
explanation, further studies are needed to define the changes in lipid metabolism upon a
shift in temperature in wild-type Chlamydomonas cells.

While there were definite consequences to growth when the BT A 1 gene is
silenced, there were no profound effects on polar lipid composition under the conditions
studied. Figure 4.6 shows the results of membrane lipid quantification by TLC followed
by GC of fatty acid methyl esters, indicating that there are no statistically significant
increases or decreases in the relative amounts of the polar lipid classes. This result was
somewhat unexpected, but may be explained by a strong tendency toward membrane
homeostasis. It appears that a reduced capacity for membrane biogenesis, as is

presumably the case in the BTAI-RNAi suppressors, can be partially compensated by

128

attenuation of growth rate. This phenomenon is similar to that observed in yeast with
compromised PtdCho biosynthesis, in which growth cessation was hypothesized to be a
mechanism to avoid cell death (Howe et al., 2002). In the yeast system, however, a
profound reduction in PtdCho mass was observed after inactivation of both of its
biosynthetic pathways. With proper experimental conditions, this might occur with
DGTS in Chlamydomonas, however at the present time we have no way to abruptly shut
off DGTS biosynthesis in rapidly growing cells. The RNAi lines under study are stably
transformed and, as discussed above, probably represent weak alleles with only partial
reduction in transcript levels (Figure 4.5).

The results of this study give a nearly comprehensive set of genes likely to be
involved in membrane lipid biosynthesis in Chlamydomonas. However, the assembly of
a biological membrane is more that the sum of its biosynthetic reactions, and there are
undoubtedly layers of complexity which this study fails to reach. One area in which this
gene set is lacking is lipid transport, which is likely to be critical in maintaining the
proper lipid composition of a given membrane. For example, the case of a lipid being
synthesized in one membrane and transported to another must be accounted for when
devising a mechanistic description of membrane biogenesis. Likewise, simply knowing
the genes involved in PtdEtn and DGTS biosynthesis (or any other set of lipids in a given
membrane) says nothing of how the final stoichiometric ratio between the two is
regulated so as to form a functional membrane with the correct biophysical properties.
Given the lack of redundancy in the pathways described above, the absence of
complicating factors such as tissue specificity, and the ease of functional analysis by gene

silencing as described for BTAI , Chlamydomonas is poised to be a useful model for

129

studying how plant cells sense and control the flux of precursors into different polar lipid

biosynthesis pathways in order to produce a functional membrane.

130

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135

Chapter 5
The sulfolipids 2'-0-acyl-sulfoquinovosyldiacylglycerol and
sulfoquinovosyldiacylglycerol are absent from a Chlamydomonas reinhardtii mutant

deleted in SQDI)

 

2 This work has been published: Riekhof,W.R., RuckleME., Lydic,T.A., Sears,B.B., and Benning,C.
(2003). The sulfolipids 2'-O-acyl-sulfoquinovosyldiacylglycerol and sulfoquinovosyldiacylglycerol are
absent from a Chlamydomonas reinhardtii mutant deleted in SQDI. Plant Physiol [33, 864-874.

136

Abstract
The biosynthesis of thylakoid lipids in eukaryotic photosynthetic organisms often
involves enzymes in the endoplasmic reticulum (ER) and the chloroplast envelopes. Two
pathways of thylakoid lipid biosynthesis, the ER and the plastid pathways, are present in
parallel in many species, including Arabidopsis, but in other plants, e.g. grasses, only the
ER pathway is active. The unicellular alga Chlamydomonas reinhardtii diverges from
plants like Arabidopsis in a different way because its membranes do not contain
phosphatidylcholine, and most thylakoid lipids are derived from the plastid pathway.
Here, we describe an acylated derivative of sulfolipid, 2'-0-acyl-
sulfoquinovosyldiacylglycerol (ASQD), which is present in C. reinhardtii. Although the
fatty acids of sulfoquinovosyldiacylglycerol (SQDG) were mostly saturated, ASQD
molecular species carried predominantly unsaturated fatty acids. Moreover, directly
attached to the head group of ASQD was preferentially an 18-carbon fatty acid with four
double bonds. High-throughput robotic screening led to the isolation of a plasmid
disruption mutant of C. reinhardtii, designated qu1 , which lacks ASQD as well as
SQDG. In this mutant, the SQDI ortholog was completely deleted and replaced by
plasmid sequences. It is proposed that ASQD arises from the sugar nucleotide pathway of
sulfolipid biosynthesis by acylation of the 2'-hydroxyl of the sulfoquinovosyl head group.
At the physiological level, the mutant showed increased sensitivity to a diuron herbicide
and reduced growth under phosphate limitation, suggesting a role for SQDG and/or
ASQD in photosynthesis as conducted by C. reinhardtii, particularly under phosphate-

lirnited conditions.

137

Introduction

The thylakoid membrane of oxygenic photosynthetic organisms provides the lipid matrix
in which the photosynthetic electron transport complexes are embedded. The lipid portion
of the thylakoid membrane consists of four glycerolipids, namely mono- and
digalactosyldiacylglycerol (MGDG and DGDG), phosphatidylglycerol (PG), and the
sulfolipid sulfoquinovosyldiacylglycerol (SQDG). The focus here is on SQDG and its
derivative 2'-O-acyl-sulfoquinovosyldiacylglycerol (ASQD). The genes encoding
enzymes for the biosynthesis of SQDG have recently been isolated in Arabidopsis
(Essigmann et al., 1998; Yu et al., 2002). Furthermore, sulfolipid-deficient mutants in
bacteria (Benning et al., 1993; Giiler et al., 1996; Weissenmayer et al., 2000),
Arabidopsis (Yu etal., 2002), and Chlamydomonas reinhardtii (Minoda et al., 2002; Sato
et al., 1995) have provided important clues toward the function of SQDG in
photosynthetic membranes. The growth of the sulfolipid-deficient mutants in the
aforementioned bacteria and Arabidopsis is impaired after phosphate starvation, whereas
the sulfolipid deficiency does not seem to have an adverse effect on growth under optimal
conditions. In the respective wild types, the biosynthesis of SQDG is up-regulated under
phosphorus limitation, apparently to compensate for a loss of the anionic phospholipid
PG. This adaptation results in conservation of phosphate, while maintaining the anionic
character of the thylakoid membrane. In the mutants, this compensatory mechanism is
inactivated and PG levels are maintained under phosphate deficiency. Experiments with a
C. reinhardtii mutant deficient in SQDG suggested that this lipid may be required for the
proper function of photosystem II (PSII) under optimal growth conditions (Minoda et al.,

2002; Sato et al., 1995). Sulfolipid might also be more critical for photosynthesis in the

138

cyanobacteria Synechocystis sp. PCC6803 because previous attempts to disrupt a
sulfolipid gene failed in this particular organism without exogenous supplementation of
SQDG (Giiler et al., 2000).

Sulfolipid biosynthesis in plants proceeds in two steps via the sugar nucleotide pathway
(Benning, 1998; Pugh et al., 1995). The first reaction, the biosynthesis of UDP-
sulfoquinovose from UDP-Glc and sulfite, is catalyzed by SQDI (Sanda et al., 2001) and
the second reaction, the transfer of sulfoquinovose from UDP-sulfoquinovose to
diacylglycerol, is carried out by SQD2 (Yu et al., 2002). Here, we describe a mutant of C.
reinhardtii in which the SQDI ortholog has been deleted and replaced by plasmid
sequences. To our surprise, not only was SQDG absent from this mutant, but also a
second minor sulfolipid, which has not been previously described, was missing. The
structure of this lipid was analyzed and its biosynthesis is discussed. Furthermore, the
absence of two lipids in this mutant raises questions regarding the interpretation of

physiological data obtained for sulfolipid mutants of C. reinhardtii.

Materials and Methods

Growth and Labeling of Chlamydomonas reinhardtii Cell Cultures

The cell wall-less strain dw15.1 (cw15, nitl , and Int-l) was obtained from Arthur
Grossman (The Carnegie Institution of Washington, Stanford, CA). This strain and its
derivatives were grown in Tris-acetate-phosphate (TAP) medium (Hanis, 1989) modified
to contain phosphate (Pi) at varying concentrations as indicated. For growth on plates,
this medium was solidified with 1% (w/v) purified agar. Small liquid cultures were grown

under constant illumination as 30- to 40-mL cultures in closed 50-mL conical centrifuge

139

tubes, or larger cultures as 5-liter cultures in 6-liter flasks under cool-white fluorescent
lights at 24°C under a 16-h day/8-h night regime. Cultures were agitated by shaking twice
daily, and cells were harvested by centrifugation in the late logarithmic phase of growth.
Growth rates were determined by measuring the optical density at 720 nm of 30-mL
cultures grown in TAP medium under Pi-replete (initial concentration of 1 mM Pi) or Pi-
limited (initial concentration of 0.02 IIIM Pi) conditions. 358- and l4C-labeling experiments
were carried out by the addition of 50 pCi [358] N82 804 (carrier-free 61606 Ci/mmol) or
25 pCi (47.5 mCi/mmol) 1-[14C]acetate to a 30 mL culture. Cells were also grown
photoautotrOphically on high salt (HS) medium agar plates (Harris, 1989) at 1/3 strength
and with varying concentrations of DCMU, added as a concentrated stock in dimethyl

sulfoxide.

Quantitative Lipid Analysis

Lipids from cell pellets of 358— or l4C-labeled 30-L cultures were extracted immediately
after harvesting with chloroform/methanol/88% formic acid (10/10/1, v/v) and the phases
were separated by addition of 3 volumes of 0.2 M H3PO4 and KCl. Radioactivity in an
aliquot of the chloroform phase was determined by scintillation counting and a fixed
amount of radioactivity was used for analysis. For l4C-acetate labeling, two-dimensional
TLC analysis (Benning et al., 1995) was followed by exposure to a phosphorimager
screen (Molecular Dynamics, Sunnyvale, CA), and for 35S-labeling, a one-dimensional
TLC system was used as described (Hartel et al., 2000) followed by quantification of the

signals on the phosphorimager.

140

Generation and Screening of Insertional Mutants

Strain dw15.1 was used for insertional mutagenesis with plasmid pMN24 containing the
wild-type nitrate reductase(N1Tl) gene (Fernandez et al., 1989). Transformation by the
glass bead method was essentially as described by Kindle (Kindle, 1998), except that
TAP medium containing nitrate as the sole N source (TAP-N03) was used for selection of
prototrophs, pMN24 was linearized with BamHI before transformation, and cells were
plated in 0.5 mL of a 50 % (w/v) slurry of corn starch in TAP (Shimogawara et al., 1998).
Suc was omitted to reduce fungal contamination of the plates.

Primary transformants were screened for lesions in lipid metabolism by a high-throughput
robotic screening strategy. Briefly, individual transformed colonies were picked with
sterile toothpicks, inoculated into 96-well plates containing 1.5 mL of TAP-N03 medium,
and grown to plateau stage. The cells in each 96-well plate were replicated onto two agar
plates with a 48-prong replica plating tool, followed by centrifugation of the remainder to
collect the cells. Cell pellets were extracted with 300 pL of chloroform/methanol/88 %
formic acid (10: 10:1, v/v) and the phases were separated with 150 uL of 1 M KCl and 0.2
M H3PO4, followed by centrifugation. Aliquots of the organic phase from each well were
then spotted in rows of 24 along each side of an activated, ammonium sulfate-
irnpregnated TLC plate (Benning and Somerville, 1992b) using a modified pipetting
robot (Genesis RSP 100, Tecan US, Research T rangle Park, NC) as previously described
(Xu et al., 2003). TLC plates were allowed to dry completely after spotting, cut in one-
half, and developed in a solvent system containing chloroform/acetone/methanollacetic

acid/water (10:4:2:2:1 , v/v). Lipids were visualized with a-napthol/sulfuric acid reagent

141

to identify the glycolipids (Benning et al., 1995) and prolonged heating at 120°C to char

the other polar lipids.

Identification and Molecular Characterization of the Site of Transgene Insertion
Plasmid rescue was accomplished essentially as described (Tarn and Lefebvre, 1993).
Briefly, genomic DNA was purified from the mutant by the method of Shimogawara et
a1. (Shimogawara et al., 1998). Subsequently,20 pg of genomic DNA was digested with
Sail and was purified by ethanol precipitation. This DNA was resuspended in 900 uL of
ligation buffer and was ligated for 16 h at 20°C with 10 units of T4 DNA ligase
(Invitrogen, Carlsbad, CA). The ligated DNA was precipitated with ethanol and was
suspended in water. A 50-pg aliquot was used to transform electrocompetent GeneHogs
(Invitrogen) Escherichia coli cells according to the manufacturer's instructions.
Transformants were selected on Lulia-Bertani broth-ampicillin (100 ug/ml) plates and
miniprep DNA was prepared and sequenced at the Michigan State University Genomics
Technology Support Facility using the primer 5'-ATGACCATGATTACGCCAAG-3'
designed to the pUC-119 multiple cloning site. The resulting sequence was used to query
the C. reinhardtii draft genome sequence (httpzllgenomejgi-

psf.org/chlre l/chlrel .home .htrnl) to identify the site of pMN24 insertion.

Restriction mapping of the rescued plasmid was canied out with Sail, HinD III, EcoRV,
Natl, Xbal, and SpeI. Because the rescued plasmid could only provide information to the
left of the insertion point, several PCR markers, designated A through B, were designed to
amplify regions of the chromosome to the right of the insertion site to determine the

extent of the deletion caused by the pMN24 insertion. Primer pair sequences were:

142

marker A, 5 '-GACAGCGCTACCGGTGTA-3 ', 5 '-TGTTCCTCCTCGCACGTA-3 ';
marker B , 5 '-AACGACATGTCGGGATCA-3 ', 5 '-ACGTGCATTTCCTGCAAT-3 ';
marker C, 5 '-CTCATCAAGCTGCTCCGT—3 ', 5 '-TTCAGCAGGAGGAGCAC-3 '; marker
D, 5 '-GTGCCTI‘GAATGCAAGCT-3 ', 5 '-GCAAACTCCTTGAGCTCG-3 '; and marker

E, 5 '-GTTAGCTGCTGCACATGC-3 ', 5 '-GCATAACACCGCAGACAA-3 '.

Large-Scale Extraction and Isolation of Compounds

Phosphate-limited cells from 20 liters of late log phase culture (TAP, 0.03 IIIM Pi, four
cultures, 5 liters each) Were collected by centrifilgation and were extracted with 150 mL
of chloroform/methanol/88% formic acid (10:10:1, v/v) followed by the addition of 37.5
mL of 02 M H3PO4 and l M KCl and were separated into two phases by centrifugation.
Anionic lipids were purified based on a procedure modified fiom O'Brien and Benson
(O'Brien and Benson, 1964). The lower organic phase was applied to a column (2 cm x
12 cm) of Florisil (J .T. Baker, Phillipsburg, NJ) equilibrated in CHC13. Pigments and
some MGDG were washed from the column with three column volumes of CHC13,and
the bound polar lipids were then eluted with two column volumes of CHCl3/CH3OH (2:1 ,
v/v). Eluate from the Florisil column was applied directly to a column (3 cm x 9 cm) of
diethylaminoethyl-cellulose (acetate form) pre-equilibrated in CHClg/CH3OH (2:1, v/v).
Uncharged polar lipids and residual pigments were washed fi'om the column with two
column volumes of the same solvent, and anionic lipids were eluted with
chloroform/methanol/concentrated ammonia (16:8: 1 , v/v), resulting in a preparation
enriched in anionic lipids, with traces of contaminating MGDG and DGDG. This eluent

was phase partitioned by the addition of water, followed by centrifugation, and the

143

organic phase was concentrated under a stream of N2 and was further purified by
preparative TLC (Benning et al., 1995). Compounds of interest were identified by light
staining with 12 vapor and were scraped from the plate into glass tubes,followed by

elution of the silica twice with chloroform/methanol (1:1 , v/v).

Mass Spectrometry

Fast atom bombardment-mass spectra were acquired in the negative ion mode on a mass
spectrometer (HX-l 10; J EOL, Peabody, MA) essentially as described (Gage et al., 1992).
Approximately 1 pg of sample dissolved in chloroform/methanol (1 :1, v/v) was mixed
with triethanolamine as matrix on the FAB probe tip. Ions were produced by
bombardment with a beam of Xenon atoms (6 KeV) and were accelerated by a voltage of
10 kV. For collisionally activated dissociation-tandem MS/MS, helilun was used as the
collision gas in a cell located in the first field—free zone,and a data system (DA-5000;

J EOL) was used to generate linked scans at a constant B/E ratio.

Fatty Acid Analysis

Samples were dried and suspended in 1 mL of methanolic HCl, incubated for 1 h at 80°C,
and the methyl esters were partitioned into hexane by the addition of 1 mL of hexane and
1 mL of 0.9 % (w/v) NaCl. Gas-liquid chromatography was as previously described

(Rossak et al., 1997).

NMR Spectroscopy

144

Samples were prepared by filtering the TLC-purified material through glass wool,
evaporating it to dryness under N2, and further drying it over desiccant under vacuum for
at least 3 h. Samples were then dissolved in 0.6 mL of chloroform-d/methanol-d4/acetic
acid-d4 (10: 10: 1, v/v) and dispensed into NMR tubes. Standard ‘H-NMR spectra were
acquired on a 500 MHz instrument (VXR-SOO; Varian, Palo Alto, CA) at 25°C, and 1H-
1H homonuclear decoupled spectra were recorded at 600 MHz on an [NOVA instrument
(Varian). Acquisitions were made by using the signal of internal methanol as lock signal,

and chemical shift values were referenced to the internal methanol resonance at 3.30 ppm.

Results

Accumulation of ASQD in C. reinhardtii

Thin-layer chromatograms of lipids from 35S-sulfate-labeled cells showed the expected
lipid band for the plant sulfolipid SQDG, and, in addition, a band of approximately 5 %
intensity of SQDG (Figure 5 .1 , ASQD). The chromatographic behavior of the compound
running with higher mobility suggested that it has a decreased polarity. The ratio of
ASQD to SQDG was comparable under normal and phosphate-depleted growth
conditions with increased labeling in the two compounds after phosphate starvation
(Figure 5.1).

To determine the identity of the unknown sulfur-labeled compound, we purified
approximately 100 pg fiom 20 liters of phosphate-starved C. reinhardtii cultures using a
combination of column and thinlayer chromatographic techniques. We hypothesized that
this compound may be a derivative of SQDG and we used a modification of a purification

procedure developed for sulfolipid (O'Brien and Benson, 1964). Fatty acid methyl ester

145

dw15.1 Asqd1
+Pi -_P.i_ +Pli -Pi
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= :s: '
: . ' * I
i:
D
.9
" , 1:1
-' 2.56.,

O

I

1
.mrmr ‘ ‘ .1: ‘1' is"? . .
' #1318“. ‘1 ”‘7‘; was. lit‘
.1; w 1 -. 4, -
M "m ..

Figure 5.1. Separation of lipid extracts from 35S—su1fate-labeled cells of C. reinhardtii
wild-type (dw15.1) and mutant (Asqu) cells. Cells were either grown in phosphate-
replete medium (+Pi) or phosphate-limited medium (-Pi). Lipids were visualized by
autoradiography. ASQD, 2'—0 acyl “ 1 ' . J ‘ " ,151, "”1, F, solvent front; 0,
origin; SQDG, “ ' . ‘ " ‘

I
1 J J‘b‘J VVAVA-

146

analysis was used to determine the fatty acid composition of ASQD, SQDG, and entire
cell extracts as shown in Table 5.1. Although SQDG carried mostly palmitate (16:0) and
oleate (18: 1), ASQD was strikingly different and contained more unsaturated fatty acids,
including an 18:4 fatty acid as subsequently confirmed by mass spectrometry (MS). This
fatty acid was originally identified in extracts of C. reinhardtii by Giroud et a1. (Giroud et
al., 1988) as 18:45’9’12’”. A corresponding 18:359‘12 fatty acid distinguishable in our

chromatograms from more typical a-linolenic acid (18:39'12'15

) was present as well. The
whole-cell extract composition was noticeably different from that observed for ASQD
and SQDG (Table 5.1). Negative-ion fast-atom bombardment (FAB)-MS (Figure 52A)
of the ASQD sample showed a major molecular ion species at approximately m/z 1074.
Closer inspection of this region (Figure 5.2, insert) revealed two major species at m/z
1073.5 and 1075.5 corresponding to triacylated SQDG molecular ions with total fatty
acid compositions (carbonszdouble bonds) of 52:7 and 52:6, respectively. Based on
analysis of fatty acid methyl esters (Table 5.1), it was hypothesized that the 52:7 species
represents a triacylated SQDG form composed of 18:3/16:0/18z4, and the 52:6 species
most likely represents a mixture of 18:3/16:0/18:3 and 18:2/16:0/18:4 molecular species.
The major peak of the FAB-MS at m/z 10735 was selected as the parent ion for FAB-
CAD-MS/MS (Figure 5.2B). Diagnostic secondary ions consistent with a triacylated
SQDG molecule were present throughout the spectrum. Only two in support of the
structure for ASQD are discussed here. The secondary ion at m/z 209 was tentatively
interpreted as a sulfoquinovose head group fragment (Figure 5.2, B and C, compound 2),

and a prominent fragment at m/z 543 and its neighbor at m/z 541 (not resolved in Figure

5.2B) corresponded to a monoacyl SQDG derivative after the loss of both diacylglycerol

147

acyl groups from a triacylated SQDG. The fatty acyl groups in these ions are 18:4 or
18:3, respectively, pr0posed to be esterifled to one of the sulfoquinovose hydroxyl groups

(Figure 5 .2C, compound 3).

To further corroborate the structure and to determine the position of the third acylation on
the sulfoquinovosyl head group, 1H-NMR spectra were recorded for ASQD and SQDG
(Figure 5.3). The ‘H spectrum of SQDG given in Figure 5.33 is essentially identical to
that described by Cedergren and Hollingsworth (1994). For ASQD (Figure 5.3C),
assignment of glycerol and carbohydrate ring proton resonances was aided by a lH-IH
homonuclear decoupling experiment (data not shown). This experiment revealed that the
2-sulfoquinovose proton is shifted down-field by approximately 12 ppm relative to
SQDG, and that the 1- and 3-protons are shifted down-field by approximately 0.17 ppm,
consistent with an acyl function esterified to the 2-hydroxyl. The region from 5.2 to 5.4
ppm provides information on the acyl substituents, as this region contains the vinyl proton
signals of the fatty acyl groups inaddition to the glyceryl-Z-proton signal. Integration of
this region (using the anomeric proton as reference) showed approximately three protons
for SQDG, indicating that the average SQDG molecule contains just one double bond
(two vinyl protons in addition to the glycerol-Z-proton). Conversely, ASQD had a much
stronger signal of approximately 14 protons (13 vinyl protons plus glycerol 2), in

agreement with the molecular species observed by MS and fatty acid analysis.

148

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8 it (f?
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.0 ’1 1' 1".
g 504 '~ 1 4:.”
.o ”i. 4 r . . '1'
< 0.0
O 3
°\ ‘ x100 F—T . i
0 100 200 300 400 500 600 700 800 900 1000
B 8.0- m/z .
o \ g
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:40‘ \ r
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0'07 I I ' I I I I r V I T , WI
0 100 200 300 400 500 600 700 800 900 1000
m/z
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Figure 5.2. Mass spectrometric analysis of ASQD. (A) Total ion negative FAB-MS with

 

insert showing an expansion of the molecular ion region. (B) FAB-CAD-MS/MS of the
molecular ion at m/z 1073.5. C. Structure prediction for the molecular ion and two
diagnostic fragments. The numbers in the two spectra refer to the three different

structures. In addition m/z values for the respective fragments are indicated.

149

 

sooc ASQD]

 

 

{ ASQD

' l 2' ; 3 . ’
' l r l 111,11," l.

~ee".%~J~«4-e/*¥H9”~”Iyihiv¥

Figure 5.3. 500 MHz proton NMR spectra of SQDG and ASQD. (A) Structures for both

lipids with proton designations as assigned to the signals in the spectrum for SQDG (B),

and ASQD (C).

150

a

Table 5.1. Comparison of the fatty acid composition of the two sulfolipids and cells

 

 

Fatty Acid ASQD SQDG Whole Cells
moI %

16:0 39.5 27.0 72.5 27.1 25.7 28.4
16:1 nd 1.4 20.2 7.9 :l:2.8
16:2 nd nd 02 20.1
16:3 nd nd 2.7 20.1
16:4 nd nd 11.1 205
18:0 4.5 21.0 2.5 21.6 3.7 20.0
18:1 2.5 21.6 205 255 19.2 20.9
18:2 6.6 21.0 1.1 20.1 7.4 20.7
18:3 A5912 10.3 21.6 0.9 20.1 9.2 23.6
18:3 ”'12:” 19.4 245 1.6 205 11.8 20.6
18:4 17.5 225 nd 1.1 205

 

a nd, Not detected. Two independent experiments were averaged (i-SD).

151

Isolation of a Sulfolipid-Deficient Insertional Mutant of C. reinhardtii

To begin to understand the biosynthesis of this new lipid and to identify genes essential
for its biosynthesis, a genetic approach was applied. For this purpose, we developed a
robotic high-throughput screening procedure (see "Materials and Methods" for details)
allowing the direct one-dimensional thin-layer chromatography (TLC) analysis of lipid
extracts from a large number of mutagenized C. reinhardtii colonies. To be able to
quickly identify a locus of interest, insertional mutagenesis by plasmid transformation
was used. Among approximately 10,000 primary transformants screened, one putative
sulfolipid-deficient strain was isolated along with a number of other putative mutant
strains showing abenant lipid profiles. To use the most stringent and sensitive test for the
presence of sulfolipids in the putative SQDG-deficient mutant, we labeled the cells with
3SS-sulfate. Cells were grown under replete and phosphate-limited conditions that
typically induce the synthesis of sulfolipids in plants (Essigmann et al., 1998; Yu et al.,
2002). As presented in Figure 5.1, C. reinhardtii dw15.1 showed increased labeling of
SQDG and ASQD under phosphate-limited growth conditions, with ASQD present at
approximately 5 % intensity of SQDG under both conditions, when approximately equal
amounts of total lipids were loaded. However, the sulfolipid-deficient mutant strain was
completely devoid of sulfolipid, and SQDG and ASQD could not be detected under either

growth condition (Figure 5.1).

A Putative SQDI Ortholog of C. reinhardtii is Deleted in the Sulfolipid-Deficient

Strain

152

Plasmid rescue was used to identify genomic DNA flanking the mutagenic plasmid
insertion. The resulting plasmid, qudI-R (Figure 5.4, bottom), carrying a greater than 10
kb insert, was partially sequenced in a single run from the right end of the pUC119 vector.
The resulting sequence could be placed on scaffold 22 of the recently released genomic
sequence of C. reinhardtii (http://genome.jgi-psf.org/chlre1/chlrel .home .html). This
sequence was approximately 30 kb downstream from a putative ortholog of the AtSQDI
gene, which is known to be essential for sulfolipid biosynthesis in Arabidopsis
(Essigmann etal., 1998). After restriction mapping of the plasmid insert and comparison
with the predicted map of the genomic sequence of scaffold 22 (Figure 5.4), the left end
of the plasmid vector was placed approximately 10 kb downstream of the putative
CrSQDI gene. To test genomic DNA flanking the inserted plasmid on the Opposite side, a
PCR marker-based strategy was used because these genomic sequences could not be
recovered by plasmid rescue. Oligonucleotides were designed to amplify segments of
DNA of defined length, which are denoted as markers A through E. Markers A through D
(Figure 5 .4) were present in DNA isolated from dw15.1 but could not be detected in
reactions using template DNA from the sulfolipid deficient mutant (data not shown).
However, Marker E (Figure 5.4) was present in both strains. Based on this analysis, the
sulfolipid—deficient strain carried a deletion of nearly 20 kb centered around the putative
CrSQDI gene. Therefore, the mutant line was designated Asqdl . The mutant is lacking
the entire CrSQDI gene as suggested by the negative result for PCR markers A and B
designed against the two ends of CrSQDI (Figure 5.4). Beyond the 3' end of this gene, no
additional predicted open reading frames were affected by the deletion (Figure 5.4).

However, beyond the 5' end of CrSQDI ,a repetitive element (Figure 5.4T), possibly the

153

 

 

 

qud1-R EisTJEf'i7."‘-“3i.; T": .* 4.42:. ‘—

Figure 5.4. Comparison of genomic sequences surrounding the CrSQDI locus in the wild
type (WT), the Asqd] mutant and the rescued plasmid (qudI-R). Five predicted
genes/elements are shown as black arrows/box. They are annotated to encode: CrSQDI,
SQDI/SQDB ortholog; T, repetitive element with similarity to transposases; ABC,
putative ABC lipid transporter. Plasmid sequences derived from pMN-24 are the pUCl 19
vector and the nitrate reductase marker gene (NI T I ). Diagnostic PCR products used to
delineate the extent of the deletion in 215qu are indicated by capital letters above the
wild-type genomic sequence. The arrow in qudI-R depicts the left end sequence used to
place the rescued genomic DNA onto the genome. Restricion enzymes indicated by two

vertical bars are: E, Eco RV; H, Hind III; N, Not I; S, Sal I; Sp, Spe I; X, Xba I.

154

Figure 5.5. Comparison of SQDI/SQDB from bacteria, archaea and plants. Unrooted tree
produced from the alignments using ClustalW software. Clusters of closely related
organisms are shaded. The following proteins were included in the analysis (GenBank
accession number): AtSQDl, Arabidopsis thaliana (T0531 1); SoSQDl, Spinacea
oleracea (AA039667); TaSQDB, Thermoplasma acidophilicum (NP_394533);
SultokSQDB, Sulfolobus tokodaii (NP_343918); SulsolSQDB, Sulfolobus solfataricus
(NP_343918); PCC6803SQDB, Synechocystis sp. PCC6803 (NP_440474); SmeSQDB;
Sinorhizobium meliloti O‘IP_386851); RspSQDB, Rhodobacter sphaeroides (B45729);
PCC794ZSQDB, Synechococcus sp. PCC7942 (AAC43899); TelSQDB,

T hermosynechococcus elongatus BP-l (NP_681188); PCC71208QDB, Nostoc sp.
PCC7120 (NP_485784). The Chlamydomonas reinhardtii ortholog, CrSQDl, was used
as predicted from the genome sequence

(http://genome.jgi-psf.org/chlre l/chlre l .home.html; scaffold_22, genie 22. 14).

155

A . . . v _ , 7 Archaea

a-Proteobacteria

 

156

remnant of a transposon, was completely missing as well. This element is present more
than 100 times in the available genomic sequence of C. reinhardtii and showed sequence
similarity to transposases. More importantly,the 5' end of a putative ABC transporter
gene with similarities to phospholipid transporters (Figure 5.4, A—C) was also deleted, as
indicated by a negative result for marker D in the mutant. Because marker E, just outside
the predicted ABC transporter open reading frame was present, it is concluded that the
deletion ends within this putative gene. Taken together, more than one gene is affected in
the qu1 mutant, complicating the interpretation of the resulting biochemical and

physiological phenotypes.

CrSQDl Clusters with Bona Fide Plant SQDI Proteins

As mentioned above, a putative ortholog of the plant SQDI gene, tentatively designated
CrSQDI , was found deleted in the sulfolipid-deficient qu1 mutant. Figure 5 shows a
lineup of representative SQDI/SQDB orthologs from plants, bacteria, and archaea, and an
unrooted tree with sequence similarity clusters. The orthologs from R. sphaeroides
(Benning and Somerville, 19923), S. meliloti (Weissenmayer etal., 2000), Synechococcus
PCC7942 (Giiler et al., 1996), and the two plants Arabidopsis (Essigmann et al., 1998)
and spinach (Shimoj ima and Benning, 2003) have been experimentally verified by mutant
and complementation analysis or by biochemical analysis of the recombinant protein. As
shown in Figure 5B, the known and presumed enzymes catalyzing the first reaction of
sulfolipid biosynthesis are highly conserved, even across kingdoms. The CrSQDl protein
clusters with the two plant proteins on a major branch of the unrooted tree that also holds

the cyanobacterial cluster (Figure 5 .5A).

157

The Asqdl Mutant Shows Reduced Growth under Phosphate Limitation and
Increased Herbicide Sensitivity

Relative lipid compositions of the control strain dw15.1 and the qu1 mutant derived
from dw15.1 were compared for cells grown under phosphate-replete and phosphate-
limited conditions (Table 5 .2). In the dw15.1 strain, the relative SQDG content drastically
increased under phosphate-limited growth conditions, whereas the amount of PG
decreased. In the .9qu mutant, which completely lacks SQDG and ASQD, PG amounts
were higher in cells grown in replete medium and did not decrease under phosphate
limitation to the same extent as observed for strain dw15.1. Qualitatively similar
observations have been made for SQDG-deficient mutants of Arabidopsis (Yu etal.,
2002) and bacteria (Benning et al., 1993; Gi'ller et al., 1996). Furthermore, the relative
content of DGDG increased in both strains under phosphate-limitation as was observed
for Arabidopsis (Hilrtel et al., 2000). Thus, glycolipid/phospholipid substitution as an
adaptive mechanism in response to phosphate deprivation is a common phenomenon in
species of bacteria, seed plants, and unicellular green alga. Interestingly, the relative
amount of the nonphosphorous extraplastidic betaine lipid (Table 5.2, DGTS) did not
change in either strain under the two growth conditions. The ASQD form of sulfolipid co-
chromatographed with DGTS in this experiment and could not be separately analyzed.
However, from our labeling experiments, we estimate that the relative amount of this
lipid did not represent more than 1% (w/v) of the polar lipids analyzed here. Thus, most

of the lipid in the DGTS/ASQD mixture was representative of the betaine lipid.

158

Table 5.2. Lipid composition of dw15.1 and Asqdl mutant under normal and phosphate-
deplived growth conditionsa

 

 

 

1.0 mM Pi 0.02 mM Pi

dw15.1 Asqdl dw15.1 Asqdl
MGDG 382 21:45 34.5 23.1 35.7 21:103 325 :t7.9
Diacylglycerol-N- 25.8 15.7 272 21:42 26.9 18.4 24.3 :32
trimethylhomoserine
(DGTS)/ASQD
Phosphatidylethanolamine 6.1 :06 7.0 :02 4.1 ;t1.9 5.0 $1.9
PG 7.7 il.1 12.4 1:0.6 3.6 il.8 10.6 i15
DGDG 14.8 1:23 163 i25 18.6 i3.4 23.2 $7.1
SQDG 5.0 :09 nd 12.6 $4.3 nd
Phosphatidylinositol 23 20.7 2.7 $05 01.8 1:09 4.4 i2.l

 

2' ASQD and DGTS were not separated in this experiment. Three independent
experiments were averaged (:60).

159

 

1.2 .
1.0 .
0.8
0.6 ,
0.4 .
0.2 .

OD720

 

 

 

0 20 40 60 80
Time (b)

Figure 5.6. Growth of dw15.1 (squares) and Asqdl (circles) under phosphate-replete

(closed symbols) and phosphate-limited (open symbols) conditions.

160

 

Figure 5.7. Growth of dw15.1 and Asqdl on agar-solidified medium containing different

amounts of the herbicide DCMU at concentrations as indicated.

161

 

Growth under the two conditions was compared for dw15.1 and Asqu , as shown
in Figure 5 .6. The growth of both strains was essentially indistinguishable under
phosphate-replete conditions. Growth of dw15.1 was delayed and growth of squ ceased
at much lower density under phosphate-limited conditions. Unfortunately, growth data on
phosphatereplete and -limited medium are not available for another SQDG-deficient
mutant of C. reinhardtii, hf-Z (Minoda et al., 2002; Sato et al., 1995). However, based on
the analysis of the hf-Z mutant, it was proposed that sulfolipid deficiency in general
causes an impairment of photosynthesis under standard growth conditions. As an indirect
test for PSII structure and function, we compared the growth of control strain dw15.1 and
the qu1 mutant on 3(3,4-dichlorophenyl)-l ,l-dimethylurea (DCMU),a herbicide
binding to the QB acceptor site of PSII and thereby blocking PSII activity. As shown in
Figure 5.7, the Asqdl mutant was more sensitive to DCMU than dw15.1, suggesting
alterations in PSII, thus making it more accessible to DCMU or less stable under the
experimental growth conditions. A similar result was observed for the hf12 mutant

(Minoda et al., 2002).

The Presence of ASQD in Lipid Extracts of C. reinhardtii is Not an Extraction
Artifact

Acylated glycolipids have been previously observed in plant extracts (Heinz, 1967), but it
was concluded that these are formed during extraction of the plant tissues at low pH by an
enzymatic transacylation reaction (Heinz et al., 1978). Several lines of evidence suggest
that the ASQD observed in extracts of C. reinhardtii was not the result of the extraction

conditions,but was present in the cell membranes. First, we tested several extraction

162

 

 

conditions, leaving formic acid out of our extraction solvent to keep the pH high or
extracting first with hot isopropanol, which should immediately destroy any enzymatic
activity, especially in a cell wall-deficient strain like dw15 .1. Applying these conditions
did not change the abundance of ASQD in the extracts (data not shown). Second, we did
not observe acylated galactolipids in the extracts as previously described for plant
extracts. Third, the diacylglycerol fatty acid composition of ASQD was highly specific
and very different from SQDG, making it unlikely that ASQD originated from the general
SQDG pool. Fourth, the third fatty acid attached directly to the head group of ASQD was
also very specific, inconsistent with a random transacylation of SQDG acyl groups from
other lipids. Taken together, these observations strongly suggest that ASQD is a minor

lipid in cellular membranes of C. reinhardtii.

A Hypothesis for the Biosynthesis of ASQD in C. reinhardtii

Important clues for the biosynthesis of ASQD can be derived from its fatty acid
composition, which differs fiom SQDG by its much higher ratio of unsaturated fatty acids
(Table 5.1). Analyzing gas chromatograms obtained for ASQD-derived fatty acid
methylesters, we could detect two unusual compounds with slightly smaller and slightly
larger retention times than that for 18:39'12'15 -linolenate methylesters (data not shown).
Two unusual fatty acids, 18:359'12 and 18:45'9'12’15, were previously reported for C.
reinhardtii (Giroud et al., 1988), and the observed retention times of the unknown
compounds would agree with retention times predicted for the methylesters of these two
unusual fatty acids. Although we did not obtain direct stnlctural data on the position of

double bonds in the acyl groups of ASQD, MS clearly suggested that the major molecular

163

 

Figure 5.8. Hypothesis for ASQD biosynthesis in C. reinhardtii. The width of arrows
indicates the extent of flux into the different molecular species. End products of the
pathway are boxed. Different molecular species can be distinguished based on their acyl
composition. Carbon number followed by double bond number are provided for the
respective acyl groups. Two acyl groups separated by / indicate the two acyl groups at the
511-1 and sn-2 positions of the diacylglycerol moiety, respectively. The head group acyl
groups are indicated by (2'). These two fatty acids are also marked with an asterisk to
indicated that they represent the two unsual acyl groups 18:3As'9'12and 18:4A5'9'12'15. The
block in the Asqu mutant is indicated. ASQD, 2'-0-acyl-sulfoquinovosyldiacylglycerol;
DAG, diacylglycerol; SQDG, sulfoquinovosyldiacylglycerol; UDP-Glc, UDP-glucose;

UDP-SQ, UDP-sulfoquinovose.

164

UDP-Glc + so;

18:1/16:0
DAG

   

UDP-SQ

16:0/16:0
DAG

     

 

16 0/160 1::l160
SQDG SQDG

 

 

 

183* (2) / 18- $80
or ‘__ - -
18:4* (2') SQDG

 

 

ASQD \ 1
18:3/16:0
SQDG

165

species has an 18:4 fatty acid attached directly to the head group. It seems likely that this
is the 18:459'12‘15 fatty acid, the only 18:4 fatty acid previously reported to be present in C.
reinhardtii. Furthermore, it may be inferred that the 18:3 fatty acid attached to the head

:359'12, also

group of the second most abundant molecular species is the 18:4 precursor 18
previously described. Our data do not allow us to conclude whether this fatty acid or
18:39‘12'15 -linolenic acid is present in the diacylglycerol moiety of ASQD. However, one
might predict that linoleic and -linolenic acids are esterified in the diacylglycerol moiety
of ASQD, whereas the two unusual fatty acids are esterified to the 2'-hydroxyl of the
sulfoquinovose head group.

A hypothesis for ASQD biosynthesis taking into account the different molecular species
for ASQD and SQDG is shown in Figure 5.8. There are clearly two pools of SQDG,
16:0/16:0 and 18: 1/16:0. ASQD biosynthesis seems to have access only to the latter pool,
which can give rise to the 18:3-containing diacylglycerol moiety by lipid-linked
desaturation. The most unsaturated 18:3/16:0 molecular species is also the most abundant
in ASQD. Interestingly, the two unusual fatty acids 183””12 and 18:459'12'15 proposed to
be esterified directly to the head group of ASQD are known as acyl groups primarily in
extraplastidic lipids, such as phosphatidylethanolamine and the betaine lipid DGTS
(Giroud et al., 1988) and it was proposed that these fatty acids are a product of the
endoplasmic reticulum (ER) desaturation pathway. A second aspect of the hypothesis
derives from the fact that the deletion of the CrSQDI gene in Asqu affects SQDG as
well as ASQD biosynthesis. If we assume that CrSQDl catalyzes the conversion of UDP-

Glc and sulfite to UDP-sulfoquinovose as described for its presumed spinach and

Arabidopsis orthologs (Sanda et al., 2001; Shimojima and Benning, 2003), the lack of

166

both lipids in the mutant suggests that ASQD is synthesized by acylation of UDP—
sulfoquinovose or by acylation of SQDG. The biosynthesis of SQDG occurs in the plastid
in all plants investigated and like its presumed orthologs, the CrSQDl sequence contains
a predicted N-terminal chloroplast transit peptide. Thus, SQDG biosynthesis in C.
reinhardtii is likely to be a function of chloroplasts, as it is in seed plants. Because UDP-
sulfoquinovose synthesis occurs in the plastid and the acyl groups found attached to the
sulfoquinovose head group of ASQD appear to be of extraplastidic origin, it seems
unlikely that UDP-sulfoquinovose is directly acylated, but that SQDG is the substrate for
acylation. Analysis of SQDG molecular species indicates that there are two pools of
SQDG, 16:0/16:0 and 18:1/16:0, of which only the second appears to be the substrate for
an acyltransferase forming ASQD (Figure 5.8). The acyltransferase involved in acylating
the sulfoquinovose head group apparently has access to the unusual fatty acids 18:35’9’12
and 18:459'12'15 derived from the ER pathway. Thus, it seems possible that this enzyme is
localized in the outer envelope of the plastid to gain access to SQDG made in the plastid
and to fatty acids derived from the ER. Whether ASQD itself is present only in plastid

envelopes or also in extraplastidic membranes remains to be investigated.

The Function of ASQD and SQDG in C. reinhardtii

To study the function of a particular membrane lipid, it would be ideal to have access to a
genetic null mutant, which is well defined at the molecular level such that effects from
secondary mutations can be ruled out. For a previously described SQDG-deficient mutant
of C. reinhardtii, hf-2 (Minoda et al., 2002; Sato et al., 1995), derived from UV

mutagenesis and identified based on its high chlorophyll fluorescence phenotype, the

167

exact molecular defect is not known. The physiology of this mutant was investigated in
great detail, and based on its photosynthesis related phenotypes, it was concluded that
SQDG plays a crucial role in photosynthesis of C. reinhardtii (Minoda et al., 2002; Sato
et al., 1995). However, whether this mutant is impaired in growth has not been reported.
Here, we describe a new SQDG-deficient mutant of C. reinhardtii, which provides a new
challenge for the interpretation of the physiological data obtained for this and possibly
other SQDG-deficient mutants. The Asqdl mutant completely lacks not only SQDG, but
also ASQD. Therefore, any physiological phenotype observed for the mutant could be
due to the lack of SQDG, ASQD, or both. To distinguish between the functions of ASQD
and SQDG, one would have to isolate a mutant specifically affecting ASQD biosynthesis.
It should be possible to isolate a strain carrying a mutation in the sulfoquinovose
acyltransferase gene proposed to be involved in ASQD biosynthesis, which would
specifically lack ASQD. On the other hand, because SQDG appears to be the precursor
for ASQD, we would predict that it will not be possible to isolate a strain lacking just
SQDG but still containing ASQD. It is not known at this time whether the independently
isolated SQDG-deficient mutant hf-2 also lacks ASQD.

The sqdl mutant, although well defined at the molecular level,carlies an extensive
deletion affecting not only the CrSQDI gene, but also one neighboring structural gene
(Figure 5.4). The specific effects of inactivating this second gene are not known,but as
Figure 5 .6 shows, growth under typical laboratory conditions in replete medium is not
affected by the deletion of either gene. In general, the analysis of the qu1 mutant
presented here concurs with previous findings in other organisms such as Arabidopsis,

which does not show a growth impairment unless the respective mutant becomes

168

phosphate deprived (Yu et al., 2002). When we grew the Asqdl mutant of C. reinhardtii
under phosphate—depleted conditions (Figure 5.6), we observed a reduction in growth and
no decrease in the relative amount of the anionic lipid PG, indicating its inability to adjust
the membrane lipid composition in response to phosphate starvation. However, Asqdl is
clearly more sensitive to DCMU than dw15.1, as was observed for hf-Z (Minoda et al.,
2002). Unfortunately, no DCMU sensitivity data are available for bacteria or Arabidopsis
for comparison. However, it should be pointed out that slight changes in the
photochemistry of PSII were observed for an SQDG-deficient mutant of Synechococcus
sp. PCC7942 without an effect on growth rates under optimal conditions (Giiler et al.,
1996). Therefore, SQDG deficiency in general may have subtle effects on PSII, which
leads to visible growth impairments only when additional stress factors such as phosphate

limitation are present.

The Identity of CrSQDI and Evolution of Sulfolipid Biosynthesis

Two arguments can be provided that CrSQDI represents an ortholog of SQDI/SQDB
found in plants and bacteria. First, its deletion causes the complete loss of sulfolipid in the
Asqdl mutant in agreement with the inactivation of a gene essential for sulfolipid
biosynthesis. Second, as the sequence lineup in Figure 55 shows, SQDI/SQDB proteins
are highly similar and display the expected relational clusters when presented as an
unrooted tree. The degree of experimentally confirmed orthology is high with five bona
fide members in different clusters. The presumed ortholog of C. reinhardtii, CrSQDl ,
clusters closely with the experimentally verified Arabidopsis ortholog, for which crystal

structure and reaction mechanism are known (Mulichak et al., 1999). All the unique

169

active site residues Thrl45, Tyrl82, and Hisl83 (numbering refers to the crystallized
recombinant AtSQDl protein) participating in the reaction mechanism are absolutely
conserved in all orthologs (Figure 55). Thus, the currently available evidence strongly
suggests that CrSQDl is the SQDI/SQDB ortholog of C. reinhardtii.

The close similarity of the SQDI/SQDB proteins suggests that the biosynthesis of UDP-
sulfoquinovose evolved only once, presumably in a common ancestor of bacteria,
archaea, and plant chloroplasts. Surprisingly, the SQDB protein from the cyanobacterium
Synechococcus sp. PCC7942 is more closely related to orthologs from the a-
proteobacteria group than those fiom the other cyanobacteria included in the analysis.
Considering that Synechococcus sp. PCC7492 is a simpler cyanobacterium relative to the
others, lacking tocopherols and higher unsaturated fatty acids, the clustering of its SQDB
protein with those from a-proteobacteria is revealing. It also supports the predictive value
of similarity tree building based on the SQDI/SQDB protein orthologs.

The presence of sulfolipid in archaea has not yet been described, despite the fact that
members of the archaea appear to have orthologs equally distantly related to the SQDB
proteins from bacteria and the SQDl proteins from plants. However, there is a report of a
sulfoquinovosylated protein in the archaeaon Sulfolobus acidocaldarr’us (L’ihringer et al .,
2000). Thus, archaea probably have the capability to synthesize UDP-sulfoquinovose
from UDP-Glc and sulfite catalyzed by their apparent SQDB orthologs,but they may lack
the glycosyltransferase needed to actually synthesize SQDG. Instead, they have a
transferase capable of adding sulfoquinovose as the terminal residue in an oligosaccharide

chain.

170

 

Lipid metabolism in the unicellular alga C. reinhardtii has several unique features
distinguishing it from other model systems such as Arabidopsis. Most prominently, it
lacks phosphatidylcholine but contains instead the betaine lipid diacylglycerol-N-
trimethylhomoserine. In addition to the previously described lipids, C. reinhardtii also
synthesizes a 2'-0-acyl derivative of the plant sulfolipid. Based on the fatty acids present
in ASQD, it was concluded that this lipid most likely arises from a distinct pool of SQDG
by acylation of the sulfoquinovose head group with fatty acids of extraplastidic origin. It
was proposed that the responsible enzyme may be located in the plastid outer envelope to
explain its access to plastidic and extraplastidic substrates. Plasmid insertional
mutagenesis was used in combination with a high-throughput (TLC) screen to isolate a
mutant, qu1 , carrying a deletion of the CrSQDI gene proposed to encode the UDP-
sulfoquinovose synthase of C. reinhardtii. This mutant lacks both sulfolipids, ASQD and
SQDG. Physiological analysis of the mutant led to the conclusion that the sulfolipid
SQDG and/or its derivative ASQD are required for optimal growth and photosynthesis
under phosphate-limited conditions. As previously concluded for photosynthetic bacteria
and plants, anionic sulfolipid compensates for losses in PG in C. reinhardtii as part of the

adaptive response to phosphate-limited growth conditions.

171

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Benning, C. and Somerville, CR. (1992a). Identification of an operon involved in
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Cedergren, RA. and Hollingsworth, RI. (1994). Occurrence of sulfoquinovosyl
diacylglycerol in some members of the family Rhizobiaceae. J. Lipid Res. 35, 1452-1461.

Essigmann, B., Gfiler, S., Narang, R.A., Linke, D., and Benning,C. (1998). Phosphate
availability affects the thylakoid lipid composition and the expression of SQDI , a gene
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(1989). Isolation and characterization of the nitrate reductase structural gene of
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Gage, D.A., Huang, Z.H., and Benning, C. (1992). Comparison of sulfoquinovosyl
diacyl glycerol from spinach and the purple bacterium Rhodobacter sphaeroides by fast
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Giroud, C., Gerber, A., and Eichenberger, W. (1988). Lipids of Chlamydomonas
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Giiler, 8., Essigmann, B., and Benning, C. (2000). A cyanobacterial gene, squ, required
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Gi’ller, S., Seeliger, A., Hirtel, H., Renger, G., and Benning, C. (1996). A null mutant of
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Harris, EH. (1989). The Chlamydomonas Source Book. (New York: Academic Press).

Hartel, H., Dbrmann, P., and Benning, C. (2000). DGDl-independent biosynthesis of
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Minoda, A., Sato, N., Nozaki, H., Okada, K., Takahashi, H., Sonoike, K., and Tsuzuki,
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173

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Xu, C., Fan, J., Riekhof, W., Froehlich, J .E., and Benning, C. (2003). A permease-like
protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J. 22, 2370-
2379.

Yu, B., Xu, C., and Benning, C. (2002). Arabid0psis disrupted in SQD2 encoding
sulfolipid synthase is impaired in phosphate-limited growth. Proc. Natl. Acad. Sci. U. S.
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4149.

174

Appendix A
Analysis of lipid metabolism in a primitive red alga: EST analysis of the
thermoacidophilic red alga Galdieria sulphuraria reveals conservation of central

pathways and a complete pathway for lipid A biosynthesis 3

 

3 Parts of this work have been published in: Weber, A.P.M., Oesterhelt, C., Gross, W., Briiutigam, A.,
lmboden, L.A., Krassovskaya, A., Linka, N., Truchina, J., Schneidereit, J., Vol], H., Voll, L.M.,
Zimmerrnann, M., Jamai, A., Riekhof, WR., Yu, B., Garavito, R.M., and Benning, C. 2003. EST-Analysis
of the Therrno-acidophilic red microalga Galdierr'a sulphuraria reveals potential for lipid A biosynthesis
and unveils the pathway of carbon export from rhodoplasts. Plant Mol. Biol. In press.

175

Introduction

Lipids play crucial roles in all cellular organisms, principally as components of
membranes that delineate the boundaries of the cell and its compartments, and also as
storage compounds, components of cuticular waxes, and as precursors to signaling
molecules in many organisms, among other functions. Lipid biosynthesis has been
studied extensively in higher plants and algae, leading to detailed models for biosynthesis
of fatty acids and their incorporation into glycolipids, phospholipids, and other fatty acid
derived compounds (Ohlrogge and Browse, 1995; Somerville et al., 2000). While
ubiquitous and critically important to life, lipid metabolism has traditionally been
difficult to characterize due to the inherent insolubility of many of the enzymes and
substrates involved. The combination of genetics, genomics, and biochemistry has
allowed the identification and characterization of most of the key enzymes in plant lipid
metabolism (Beisson et al., 2003; Frentzen, 2004; Ohlrogge and Browse, 1995;
Somerville et al., 2000), and the present appendix follows in this vein by using genomic

data to analyze lipid metabolism in a primitive red alga, Galdieria sulphuraria.

Materials and Methods

EST sequencing was carried out on phototrophic and heterotrophic cDNA libraries of
Galdieria sulphuraria as described (A .P.M. Weber et al., in press), and is available at
http://www.genomics.msu.edu/galdieria/index.cgi. Manual annotation of the G.
sulphuraria dataset was carried out by keyword searches of the automated annotations,
and by BLAST (Altschul et al., 1997) searching with relevant sequences of plant, animal,

fungal, or bacterial origin. Significance values used for determining homology were

176

generally set at a BLAST expectation value of less than Em, and were confirmed by

multiple sequence alignments, conservation of active site residues, etc. when possible.

Results and Discussion

Fatty acid biosynthesis. Based on the EST dataset, the fatty acid and lipid metabolism of
Galdieria is likely similar to that of seed plants. Production of malonyl-CoA through the
action of acetyl-coA carboxylase (ACCase) is regarded as a rate limiting and highly
regulated step in fatty acid biosynthesis (Somerville et al., 2000). In plants two ACCase
isozymes exist: a homomeric multi-domain protein in the cytosol, and a multisubunit
complex of four proteins in the plastid. Single ESTs for two of the subunits of plastidial
ACCase (biotin carboxylase, A4_40C10; and B-carboxyltransferase, HET__25A08) were
present in our database. Fatty acid synthase (FAS) in plants is also a plastidic multi-
protein complex, and ESTs for the 3-ketoacyl-ACP-reductase (gs01440) and enoyl-ACP-
reductase (HET_03C09) were present, representing clones fiom both the heterotrophic
and autotrophic libraries. Other FA biosynthetic activities such as FA desaturases, acyl-
CoA synthetases, and thiolases were also distributed in both libraries. These analyses
indicate that this red alga, like higher plants, most likely synthesizes fatty acids in the
rhodoplast via bacterial/plastidic type multimeric ACCase and FAS complexes,
incorporates some of these FA into plastidic membrane lipids, and exports a portion to

the cytosol for incorporation into extraplastidic lipids, as discussed below.

177

Fatty acid catabolism. B-oxidation, typically a function of the peroxisome, was
represented in the ESTs as at least two acyl-CoA oxidase and three acetyl-CoA thiolase
isozymes, while enoyl-CoA hydratase and 3-hydroxyacyl-CoA dehydrogenase, which are

often present as a multifunctional polypeptide, were not detected.

Membrane lipid biosynthesis. G. sulphuraria synthesizes phospholipids and glycolipids
typical of higher plants (T. Horlacher and C. Benning, unpublished results) and pathways
for their biosynthesis were readily identifiable hour the EST data. Two pathways for
phosphatidylethanolamine biosynthesis were identified, consisting of phosphatidylserine
decarboxylase (HET_44E08) and the CDP—ethanolamine (Kennedy) pathway
(ethanolamine kinase, HET10E09; ethanolamine phosphate cytidylyltransferase,
A4_10G06). Phosphatidylcholine is also apparently synthesized by two pathways in
which either phosphoethanolamine is methylated to phosphocholine
(phosphoethanolamine methyltransferase, A4_09F12) and then activated to CDP-Cho .
and used to form PtdCho, or by the lipid-linked methylation of PtdEtn to form PtdCho
(PEMT, A4_16B08.)

Phosphatidylglycerol is present in seed plants in the plastid, mitochondria, and ER
membranes, and phosphatidylglycerophosphate synthase (GSO6870, 2 ESTs) was
represented, although its localization could not be predicted, as was cardiolipin synthase
(HET_11H01), typically a mitochondrial enzyme. Phosphatidylinositol biosynthesis was
represented by inositol-3-phosphate synthase (HET_12A09), and implicitly by a PI-4-

kinase (HET_10C06) although the actual Ptdlns synthase was not readily identifiable.

178

 

 

Little is known about the biosynthesis and function of sphingolipids in plants,
however putative orthologs encoding serine palmitoyltransferase (HET_31C05) and
sphingolipid-A4 desaturase (6800710, 2 ESTs) were present.

Plastidial lipids typically consist of four species: mono- and
digalactosyldiacylglycerol (MGDG and DGDG), sulfoquinovosyldiacylglycerol (SQDG),
and Pthro. MGDG synthase (HET_20D10) and SQDI (HET_20F10) were represented,
as was the aforementioned PGP synthase, however cDNAs encoding other enzymes in
these pathways were not found. Typically, the expression of lipid genes is low and EST
coverage may not be sufficient for these to be included in the dataset.

Lipid A biosynthesis. Lipid A is the core membrane anchor of bacterial
lipopolysaccharide, the major component of the outer leaflet of the bacterial outer
membrane (Raetz and Whitfield, 2002). Surprisingly, genes encoding a partial lipid A
biosynthesis pathway are present in the Arabidopsis genome, although several steps are
missing and nether lipid A nor derivatives have ever been unequivocally identified in
seed plants. Here, we provide for the first time evidence for a complete lipid A
biosynthetic pathway in plants (Figure A2). In Galdieria, the first 2 steps in the pathway
(prA, prC, Figure A2) are likely to be encoded in the plastid genome by analogy with
C yanidioschyzon merolae and Cyanidium caldarium RKl (Glockner et al., 2000; Ohta et
al., 2003). Later steps in the pathway appear to be nuclear encoded, and corresponding
ESTs are present in our collection (prD, HET_35C04; prB, cn459 and
A4_T7comboD04; prK, A4_18A10; WaaA, A4_10A08; KDO synthetase, A4_17B04;
Figure A2), leading to the hypothesis that Galdieria has the capacity to synthesize a

molecule similar to Kd02-Lipid IVA. The pathway shown in Figure A2 represents the

179

 

 

Figure A.l: Subcellular organization of fatty acid biosynthesis and glycerolipid
assembly in Galdieria sulphuraria. EST sequences corresponding to the activities in
this scheme were annotated, and those present in the database are indicated by an

asterisk.

180

 

a

 

m _<— PthroP

UDP- Gal 3E L UDP- SQ <— UDP-Glc G-3.P

mfi DAG <— PtdOH ——> cor= DAG

   
 
    

 

 

UDP-Gal
lyso-PtdOH Acetyl-CoA
Acyl-00A K
1 Malonyl-CoA
Pl - -
35"“ Acyl-ACP
G-3—P
IMS
Cytosd\ GIc-r-P
Acyl-CoA 1
> lns-3-P
Iyso-PtdOH (3-3-1:
F
Ser PtdOH -> CDP—DAG ..
1 Die

5‘" PtdSer PthroP

P-Etn —> coo:$>+ @

1*
1* 1*

DAG
I at

P-Cho -—>coP-crto\>- PtdCho

181

 

Figure A2: Lipid A biosynthetic pathway, and ESTs present in the database. ESTs
encoding enzymes with similarity to those involved in bacterial lipid A formation were

annotated and indicated in the reaction scheme, as shown.

182

 

prA prC prD prH
(plastid encoded) (plastid encoded) (HET_35C04) (no candidate)

—-—> —> —-> —->
(cn459.
”"8 A4_T700mboDO4)
WaaA prK
(A4_10A08) (A4_18A10)
KdozLipid lVA +— 4—

183

 

“typical” case as elucidated in E. coli, however other bacteria accumulate lipid A species
with radically different structures, e. g. Rhizobium etli (Que et al., 2000a; Que et al.,
2000b). Also, the lipid A structures of cyanobacteria, from which the plastid derives, are
not well characterized, so while the core of the pathway appears to be present, the later
steps leading to the final products remain to be elucidated.

If the proposed pathway is indeed operational, the function of lipid A-like
molecules in plants is mysterious, and there are no published reports of lipid A or

derivatives being detected in higher plants. It is tempting to posit that, given the extreme

 

environment in which these primitive red algae live, lipid A is essential for stability of the
plastid outer envelope, and therefore part of the pathway has been retained in the plastid
genome. In fact, some of the ESTs annotated in the pathway show their greatest
similarity to cyanobacterial sequences, further supporting the idea that in red algae and
higher plants, the plastid may contain lipid A-like molecules as part of the plastid

envelope membranes. Given the recent report of chloroplast transformation in

 

Porphyridium (Lapidot et al., 2002), and nuclear transformation of Cyanidioschizon
merolae (Minoda et al., 2004), it could be feasible to create a genetic null mutant at any
point in the pathway to study the function of lipid A-like molecules in primitive red

algae.

184

References

Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J ., Zhang, 2., Miller, W., and
Lipman, DJ. (1997). Gapped BLAST and PSI-BLAST: a new generation of protein
database search programs. Nucleic Acids Res. 25, 3389-3402.

Beisson, F., Koo, A.1., Ruuska, S., Schwender, J ., Pollard, M., Thelen, J .J ., Paddock, T.,
Salas, J .J ., Savage, L., Milcamps, A., Mhaske, V.B., Cho, Y., and Ohlrogge, J .B. (2003).
Arabidopsis genes involved in acyl lipid metabolism. A 2003 census of the candidates, a

study of the distribution of expressed sequence tags in organs, and a web-based database.
Plant Physiol. 132, 681-697.

Frentzen, M. (2004). Phosphatidylglycerol and sulfoquinovosyldiacylglycerol: anionic
membrane lipids and phosphate regulation. Curr. Op. Plant Biol. 7, 270-276.

Glockner, G., Rosenthal, A., and Valentin, K. (2000). The structure and gene repertoire
of an ancient red algal plastid genome. J. Mol. Evol. 51, 382-390.

Lapidot, M., Raveh, D., Sivan, A., Arad, S., and Shapira, M. (2002). Stable chloroplast
transformation of the unicellular red alga Porphyridium species. Plant Physiol. 129, 7-12.

Minoda, A., Sakagami, R., Yagisawa, F., Kuroiwa, T., and Tanaka, K. (2004).
Improvement of culture conditions and evidence for nuclear transformation by

homologous recombination in a red alga, Cyanidioschyzon merolae 10D. Plant Cell
Physiol. 45, 667-671.

Ohlrogge, J. and Browse, J. (1995). Lipid biosynthesis. Plant Cell 7, 957-970.

Ohta, N., Matsuzaki, M., Misumi, O., Miyagishima, S., Nozaki, H., Tanaka, K., Shin-i,
T., Kohara, Y., and Kuroiwa, T. (2003). Complete sequence and analysis of the plastid
genome of the unicellular red alga Cyanidioschyzon merolae. DNA Res. 10, 67-77.

Que, N.L.S., Lin, S.H., Cotter, R.J., and Raetz, C.R.H. (2000a). Purification and mass
spectrometry of six lipid A species from the bacterial endosyrnbiont Rhizobium etli -

Demonstration of a conserved distal unit and a variable proximal portion. J. Biol. Chem.
2 75, 28006-28016.

Que, N.L.S., Ribeiro, A.A., and Raetz, C.R.H. (2000b). Two-dimensional NMR
spectroscopy and structures of six lipid A species from Rhizobium etli CE3 - Detection of

an acyloxyacyl residue in each component and origin of the arninogluconate moiety. J.
Biol. Chem. 275, 28017-28027.

Raetz, C.R.H. and Whitfield, C. (2002). Lipopolysaccharide endotoxins. Ann. Rev.
Biochem. 71 , 635-700.

185

Appendix B

Analysis of the structure of novel galactolipids accumulating in a lipid-trafficking

mutant of Arabidopsis, and a hypothesis for their biosynthesis4

 

*Ihis work was done in collaboration with Dr. Changcheng Xu, and data from parts of this appendix have
been published: Xu, C., Fan, J., Riekhof, W., Froehlich, J.E., and Benning, C. (2003). A permease-like
protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J. 22, 2370-2379

186

Introduction

In many plant species, biogenesis of the photosynthetic thylakoid membranes is reliant on
pathways of glycerolipid biosynthesis present in the endoplasmic reticulum (ER) and
plastid envelope membranes (Roughan and Slack, 1982). In the plastidic pathway (also
referred to as the “prokaryotic pathway” due to similarities with the lipid metabolism of
the cyanobacteria from which the plastid derives), fatty acids are synthesized in the
plastid and used by plastidic acyltransferases to generate phosphatidic acid (PtdOH).
PtdOH is then either used for phosphatidylglycerol (Pthro) synthesis, or
dephosphorylated to form diacylglycerol (DAG), which is then used for galactolipid and
sulfolipid biosynthesis (Somerville et al., 2000).

In the ER localized “eukaryotic pathway,” fatty acids are exported from the
plastid and converted into acyl-CoA species for use by ER localized acyltransferases,
giving rise to a cytosolic PtdOH pool. This pool of PtdOH becomes the precursor for
phospholipid biosynthesis in the ER, giving rise to the typical membrane lipids
phosphatidylcholine (PtdCho) and phosphatidylethanolamine (PtdEtn), among others.
Based on pulse-chase labeling kinetics and analysis of molecular species, it has long been
postulated that a portion of the cytosolic PtdCho pool is used as the donor of DAG
moieties for galactolipid biosynthesis in the plastid (Browse et al., 1986), thus giving rise
to “eukaryotic” thylakoid lipid species. While it is well established that the eukaryotic
pathway of thylakoid lipid biosynthesis is operational in many plant species, a
mechanistic description of the transfer of DAG moieties from cytosolic PtdCho into

plastidic galactolipids is lacking.

187

 

 

During a screen for suppressors of the Arabidopsis (1ng mutant (Ddrmann et al.,
1995; Ddrmann et al., 1999), approximately 25 mutant lines were discovered which
accumulate novel tri- and tetra galactosyldiglycerides in the intact leaf, and one of these
mutants was subsequently shown to have a defect in the eukaryotic pathway of thylakoid
lipid synthesis (Xu et al., 2003). This work describes the structural elucidation of these
oligo—galactolipids by nuclear magnetic resonance (NMR) as containing all-B(1—->6)
linked galactose residues, and a proposal for their biosynthesis via transgalactosylation by

a B-galactosidase type mechanism is presented.

Materials and Methods

Isolation of lipids.

Lipids of the wild type and tng -1 mutant were extracted and separated by preparative
TLC on silica-60 plates in the solvent system chloroform/ methanol/ acetic acid/ water
(85:15:10:5, v/v). Bands corresponding to the respective lipids were scraped from the
plate and eluted in chloroform/methanol (1 :1, v/v), followed by evaporation of the sample
to dryness under a stream of nitrogen.

NMR.

Lipids purified as described above were further dried under vacuum over dessicant for
several hours before dissolving in 0.7 ml of CDC13/CD3OD/CD3OOD(1021021, by vol .).
The addition of CD3OOD shifts the water signal to 5.5—6 p.p.m. Data were collected on a
Varian VXR-500 spectrometer at 500 MHz for protons and analyzed with the Varian
VNMR software package. Chemical shifts were measured in relation to residual methanol

as reference, defined as 3.30 p.p.m. For each sample, 128—5 12 Fle were collected,

188

 

 

- - -- -MGDG

W W

— -

w W
. I. I. 1111 1

WW

trimest- m.“

- - -DGDG

”w

in ------ TGDG

Figure B. l: TLC of tgdl and wild type. Lipids were extracted from the wild-type and

mutants, as indicated above the image, and resolved by thin layer chromatography as

outlined in the Materials and Methods section. MGDG, monogalactosyldiacylglycerol;

DGDG, digalactosyldiacylglycerol; TGDG, trigalactosyldiacylglycerol.

189

 

 

depending on concentration. Suppression of the water signal was accomplished using a

pre—saturating pulse in all mutant samples, which contained less lipid.

Results and Discussion

 

As shown in Figure 1, the tng -1 mutant accumulates novel glycolipid species, which are

absent in the wild-type. Gas-chromatography of alditol acetates derived from the

 

hydrolyzed TGDG sample confirmed the identity of the monosaccharides as exclusively
galactose, and fast-atom bombardment mass spectrometry (FAB-MS, data not shown)
was consistent with a trigalactosyldiglyceride structure. NMR was used to determine the
anomeric linkages of the trigalactosyl chain. Figure 2 shows the anomeric region of the
500 MHz NMR spectra of the galactolipids MGDG and DGDG isolated from the wild
type and DGDG and TGTG from the tng-l mutant. MGDG has a [Kl-’3) linkage to
diacylglycerol (Figure 1A), and DGDG has a Ga101( 1-—r6)-Gal[3( 1—>3)-DAG (Figure 2B,
aBDGDG) structure. Surprisingly, the structures of DGDG and TGDG isolated from the
mutant did not contain these typical anomeric linkages, as shown in the spectra in Figures
2C and 2D. Whereas we expected the TGDG lipid to contain at least one tit-linkage (that
is, we assumed it would be derived by further galactosylation of aBDGDG already
present in the membranes), we in fact observed three B-linkages. Given that the chemical
shifts of all three B—protons were nearly identical, a linear all-B( l—>6) chain is the most
likely structure. Since the TGDG of the tng -1 mutant is all B-linked, we assumed there
to be a pool of BBDGDG giving rise to all-BTGDG. NMR of the DGDG isolated from

the mutant (Figure 2B) shows clearly that it is a mixture of 018- and BBDGDG, due to the

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Figure B2: NMR of galactolipids isolated from wild-type and mutants. Lipids were
purified by TLC and NMR data was collected as described in Materials and Methods. A.
MGDG (monoglacatosyldiacylglycerol) from the wild-type (Col-2) background. B.
DGDG (digalactosyldiacylglycerol) from wild-type. C. DGDG from the tng-I mutant.

D. TGDG (trigalactosyldiacylglycerol) from the tgd] -1 mutant.

 

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presence of a B-anomeric resonance at about the same position as one of the TGDG
signals, and with about half the area as the a-anomeric resonance, indicating that the
mixture is about 2/3 018- and 1/3 BBDGDG.

The phenomenology of TGDG synthesis deserves comment. It has been observed
that Mg2+ions stimulate the formation of TGDG in isolated chloroplasts, and NMR
analysis of the TGDG accumulating in Mg2+treated isolated spinach chloroplasts is of
the all-B structure (data not shown), therefore both the TGDG being formed in the tng -1
mutant and in isolated chloroplasts may derive from the same enzyme(s). It is also of
note that the enzyme responsible for formation of TGDG resides on the outer chloroplast
envelope, as judged by its susceptibility to thermolysin treatment (Xu et al., 2003), and
also requires no addition of activated sugar donor (e.g. UDP-galactose). This leads to
several requirements: the TGDG synthesis activity is attributable to a Mg2+activated
enzyme capable of removing the galactose from one galactolipid molecule and
transferring it to a second, while retaining the B-configuration. This description is very
similar to that of a B-galactosidase such as LacZ from E. coli, and a hypothetical scheme
for the synthesis of all-B-oligogalactolipids by a galactosylhydrolase-like mechanism is
given in Figure 4. Briefly, a catalytic aspartate residue in the active site cleaves the
anomeric bond, resulting in a meta-stable a-linked galactosyl-enzyme intermediate (Juers
et al., 2001). During the normal galactosidase cycle, a water molecule hydrolyzes the
covalent intermediate, releasing galactose and free enzyme. If, however, the oxygen is
part of an alcohol (e.g. the 6’-hydroxyl of an MGDG molecule) then the
galactosylhydrolase functions as a galactosyltransferase. This activity for LacZ has been

known for decades due to the formation of allolactose (GalB(1—»6)Glc), which proceeds

193

 

 

Ho 1..

.. 1

Figure B3: Hypothesis for oligogalactolipid synthesis by a B—galactosidase
mechanism. As described in the text, (1) the catalytic aspartate in a B-galactosidase
cleaves the galactose-glycerol linkage, forming (2) a covalent enzyme intermediate. This
meta-stable intermediate is released by attack (3) of the 6' hydroxyl of a second MGDG

molecule, leading to the accumulation of (4) 138-DGDG. This cycle would create

194

by breaking of the GalB(1—+4)Glc bond of lactose, forming the a-galactosyl-enzyme
intermediate, followed by attack of the 6-hydroxyl of the same glucose on the galactosyl-
intermediate. This reaction happens with the same glucose molecule when the lactose
concentration is low, however at high concentrations of lactose, linear B( l—t6) linked
oligosaccharides are formed (Albayrak and Yang, 2002), i.e. the same structure as the
headgroup portion of the oligogalactolipids described above. Therefore it seems feasible

to conclude that the processive galactosyltransferase activity could be annotated as a

 

glycosyl hydrolase family enzyme in the Arabidopsis genome database, perhaps even as a

B-galactosidase.

 

195

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