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This is to certify that the
dissertation entitled

UNDERSTANDING THE ORIGIN AND FUNCTION OF
ORGANELLAR METABOLITE TRANSPORT PROTEINS IN
PHOTOSYNTHETIC EUKARYOTES: GALDIERIA SULPHURARIA
AND ARABIDOPSIS THALIANA AS MODEL SYSTEMS

presented by
MARC LINKA

has been accepted towards fulfillment
irements for the

Ph.D. Genetics

 

 

 

 

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12/05/2008

 

Date

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UNDERSTANDING THE ORIGIN AND FUNCTION OF ORGANELLAR
METABOLITE TRANSPORT PROTEINS IN PHOTOSYNTHETIC EUKARYOTES:
GALDIERIA SULPHURARIA AND ARABIDOPSIS THALIANA AS MODEL SYSTEMS
By

Marc Linka

A DISSERTATION

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

DOCTOR OF PHILOSOPHY
Genetics

2008

ABSTRACT
UNDERSTANDING THE ORIGIN AND FUNCTION OF ORGANELLAR
METABOLITE TRANSPORT PROTEINS IN PHOTOSYNTHETIC EUKARYOTES:
GALDIERIA SULPHURARIA AND ARABIDOPSIS IHALIANA AS MODEL SYSTEMS
By
Marc Linka
Membrane-bound compartments, especially the organelles mitochondria and plastids, are
a hallmark of eukaryotic cells. Organellar metabolite transport proteins facilitate the
exchange of metabolites across membranes in a cell and are crucial for connecting
biochemical pathways that operate in separate compartments. Of particular interest to
plant scientists is the plastid organelle of photosynthetic eukaryotes. Plastids synthesize
and deliver major biological molecule classes such as carbohydrates, fatty acids, amino
acids, and nucleic acids to the rest of the cell and thus the plastid has to be extensively
connected to the cytosol. Plastids originated fi'om an endosymbiotic cyanobacteria-like
ancestor about 1.6 billion years ago and in this thesis the differences between plastid
metabolite transporters of the eukaryotic model plant Arabidopsis thaliana and
prokaryotic cyanobacteria were investigated. A phylogenomic analysis of 83 predicted
plastid metabolite transporters from Arabidopsis thaliana has been conducted in
collaboration with Dr. Debashish Bhattacharya. These studies allowed the conclusion that
the majority of the transport proteins in extant plastids are absent fi'om free-living
cyanobacteria and originated from eukaryotic host genes. They represent true innovations
associated with organelle evolution. Transporters became likely targeted to the
endosymbiont via the endoplasmic reticulum of the host early in its evolution.

Furthermore, the results suggest that export of photosynthates from the plastid in form of

sugar-phosphates has been a selective advantage to set-up a permanent endosymbiosis
between the host and the endosymbiont. While these sugar-phosphate transport proteins
are conserved in all photosynthetic eukaryotes, their biochemical properties co-evolved to
meet the specific metabolic requirements in the distinct groups of the eukaryotic
kingdom. As reported in Chapter 3, further studies have shown that in contrast to higher
plants, the red alga Galdieria sulphuraria has a high affinity export system for tn'ose-
phosphates and lacks hexose-phosphates transport across the plastid envelope membrane.
This reflects an adaptation for an efficient export of photosynthates from the organelle
due to an absence of a plastidic starch pool in red algae.

Metabolite carriers facilitate also the transport of compounds in a single, highly
compartmentalized cellular pathway. A prime example is the photorespiratory pathway.
2-phosphoglycolate is produced by the oxygenase reaction of the enzyme ribulose- 1,5-
bisphosphate Carboxylase/Oxygenase (RubisCO) and subsequently recycled to 3-
phosphoglycerate in the compartments chloroplast, cytosol, peroxisome, and
mitochondrion. A reverse genetic approach was used to identify the transport proteins
involved in photorespiration and 32 candidates have been designated. Five of these were
genetically analyzed to test their role in the recycling of 2-phosphoglycolate, leading to
the discovery of a novel transporter required for a functional photorespiratory pathway.
This protein is localized to the inner envelope membrane of mitochondria and the
transporter most likely imports a cofactor from the cytosol, which is required for the

mitochondrial glycine decarboxylase enzyme complex.

ACKNOWLEDGEMENTS

I would like to sincerely thank Dr. Andreas Weber for this one-time opportunity to join
his research group and complete my graduate studies here at Michigan State University.
Under his supervision I developed my confidence and capabilities to inquire into the
phenomena on my daily scientific endeavor. He allowed me to be a student, an instructor,
a teacher and an independent thinker. He always encouraged me to broaden my scientific
knowledge and to enjoy research. I would also like to thank the Director Dr. Barbara
Sears and Associate Director Dr. David Amosti, the faculty and Jeannine Lee of the
Genetics Graduate Program for organizing such a great educational program at Michigan
State University. The fulfillment of the requirements during the first three years of my
graduate studies was of paramount importance for my scientific progress and will be for
my future goals. My committee members Dr. Robert Larkin, Dr. John Ohlrogge, Dr.
Steve van Nocker, and Dr. Richard Allison deserve a special note. They contributed by
discussion and opinions to my progress and allowed the transfer to Germany without any
concern during my last year. I would also like to thank all graduate students, postdocs
and faculty members for their interaction and discussion at seminars, in the laboratories,
and off—campus. In addition, these acknowledgements would not be complete if I did not
mention the former and recent members of “The Weber Lab”. Thanks for your support
and your tremendous scientific and non-scientific help. Most importantly, I would like to
thank my wife Nicole. You are the sunshine of my life. I thank my parents, for their
support to leave "Old Europe Into The West". But I have to confess that there are places

on this Earth, which are even farther away than Michigan.

iv

TABLE OF CONTENTS

LIST OF TABLES .................................................................................. vii

LIST OF FIGURES .................................................................................. viii

KEY TO ABBREVIATIONS ........................................................................ x

INTRODUCTION

PLASITD EVOLUTION FROM THE PERSPECTIVE OF METABOLITE

TRANSPORT PROTEINS .......................................................................... 1
References ........................................................................................ 25

CHAPTER 1

PHYLOGENY OF THE PLASTID PHOSPHATE TRANSLOCATOR FAMILY ....... 34
Abstract ............................................................................................. 35
Introduction ..................................................................................... 35
Material and Methods .......................................................................... 36
Results and Discussion ........................................................................... 37
References ....................................................................................... 45

CHAPTER 2

PHYLOGENY AND ORIGIN OF PLASTID METABOLITE TRANSPORT

PROTEINS FROM ARABIDOPSIS THALIANA ................................................. 47
Abstract .......................................................................................... 48
Introduction ....................................................................................... 49
Material and Methods ............................................................................ 52
Results and Discussion ........................................................................... 56
References ......................................................................................... 81

CHAPTER 3

ANALYSIS OF THE PLASTID PHOSPHATE TRANSLOCATOR FAMILY FROM

THE RED ALGA GALDIERIA SULPHURARIA ................................................ 87
Abstract .......................................................................................... 88
Introduction ..................................................................................... 89
Material and Methods .......................................................................... 92
Results ............................................................................................ 95
Discussion ..................................................................................... 117
References ..................................................................................... 127

CHAPTER 4

PROPOSED METABOLITE TRANSPORT PROCESSES FOR THE

RE-ASSIMILATION OF AMMONIA DURING PHOTORESPIRATION ............... 133
Abstract ....................................................................................... .134

Results .......................................................................................... 134
References ...................................................................................... 146

CHAPTER 5
A REVERSE GENETIC APPROACH TO IDENTIFY THE PROPOSED
METABOLITE TRANSPORT PROTEINS IN THE PHOTORESPIRATORY

PATHWAY.. .................................................................................................................. 149
Abstract ......................................................................................... 150
Introduction .................................................................................... 151
Material and Methods ........................................................................ 155
Results ........................................................................................... 168
Discussion ...................................................................................... 201
References ..................................................................................... 208

CHAPTER 6

CONCLUSION AND PERSPECTIVES ........................................................ 215
References ..................................................................................... 225

LIST OF TABLES

Table 2.1. List ofArabidopsis thaliana chloroplast solute transporters

analyzed in this study and their putative evolutionary origins ................................. 58
Table 3.1. Oligonucleotide primers designed in this study ................................... 97
Table 3.2. Molecular characteristics of the plastidic phosphate transporter

GsTPT, GsPPT and GSGPT from Galdieria sulphuraria ..................................... 100
Table 3.3. Comparison of kinetic constants from Galdieria sulphuraria and

higher plant pPT homologues ...................................................................... 1 13
Table 5.1. Primer sequences used in Chapter 5 ............................................... 158

Table 5.2. Co-expressed photorespiratory enzymes from Arabidopsis thaliana... . . . 170

Table 5.3. Putative transporter genes, which are transcriptionally co-regulated
with the photorespiratory pathway ............................................................... 171

Table 5.4. Systematic search for the BOU substrate ......................................... 196

Table 5.5. Presence of a membrane potential does not stimulate glycine of serine
transport by BOU .................................................................................. 197

Table 5.6. Several cofactors must be provided for the mitochondrial
glycine decarboxylase (GDC) .................................................................... 198

vii

LIST OF FIGURES

Figure l. The photorespiratory pathway ......................................................... 24

Figure 1.1. Maximum likelihood phylogeny of endomembrane and plastid
translocators .......................................................................................... 42

Figure 1.2. Model for plastid and translocator origin in the common ancestor of the

Plantae during primary endosymbiosis ........................................................... 43
Figure 1.3. Model for plastid and translocator origin in the common ancestor of the

chromalveolates during secondary endosymbiosis .............................................. 44
Figure 2.1. Origin of plastid targeted solute transporters in Plantae ........................ 61

Figure 2.2. Plastid targeted solute transporters of putative 'Host' origin in Plantae. . 64

Figure 2.3. Plastid targeted solute transporters of putative ’cyanobacterial'l‘plastid
endosymbiont' origin in Plantae ................................................................... 67

Figure 2.4. Plastid targeted solute transporters of putative 'Chlamydia-like'
origin in Plantae ...................................................................................... 74

Figure 2.5. Plastid targeted solute transporters of 'Other’ or 'PIantae-specific'
origin in Plantae ..................................................................................... 79

Figure 3.1. Protein alignment of plastidic phosphate transporters from

Arabidopsis thaliana and Galdieria sulphuraria ............................................... 102
Figure 3.2. Expression of plastid phosphate transporter (pPT) homologues

from Galdieria sulphurarz'a in yeast ............................................................... 105
Figure 3.3. Time-kinetics of pPT homologues from Galdieria sulphuraria .............. 108
Figure 3.4. Internal substrate dependence on the transport activities

of GsTPT, GsPPT, and GSGPT ................................................................... 112
Figure 3.5. Transcript abundance and protein activity of GsTPT,

GsPPT, and GSGPT in Galdieria sulphuraria cells ............................................ 1 16
Figure 3.6. Flexible transport of triose-phosphate across the plastid membrane is

central for the cellular carbon metabolism in the red alga Galdieria sulphuraria. . .. 125
Figure 4.1. Simplified scheme of the current textbook photorespiratory cycle. . . . . .138

viii

Figure 4.2. A citrulline/ornithine shuttle between mitochondria and plastids ............. 142

Figure 4.3. A glutamate/glutamine Shuttle between mitochondria and plastids .......... 145
Figure 5.1. Isolation of A. thaliana T-DNA insertion lines for At1g54350,

At2g42770, At2g48070, At3g26710, and At5g46800 ......................................... 176
Figure 5.2. Growth phenotype of T-DNA insertion lines from the candidate genes
Atlg54350, At2g42770, At2g48070, and At3g26710 ......................................... 178
Figure 5.3. Growth phenotype of T-DNA insertion line bouI ............................... 180

Figure 5.4. Concentrations of photorespiratory marker metabolites glyoxylate,
ammonium ions, glycine, and serine in bouI leaves ............................................ 183

Figure 5.5. Carbon dioxide-dependent effect on additional metabolite levels in b0ul...185

Figure 5.6. Molecular complementation of bouI .............................................. 188
Figure 5.7. Complementation of the bard mutant restores glycine concentrations

to wild type levels ................................................................................. 189
Figure 5.8. Mitochondrial glycine oxidation is impaired in bad ........................... 190
Figure 5.9. Irnmunoblot analysis of mitochondrial enzymes ................................. 1.91

Figure 5.10. Heterologous expression and purification of BOU protein in Escherichia
coli and Saccharomyces cerevisiae ............................................................... 195

Figure 5.11. BOU relevant branch from the maximum-likelihood unrooted tree of the
mitochondrial carrier family (MFC) 199

Figure 5.12. Reconstituted BOU protein exhibits ion channel properties .................. 200

ix

KEY TO ABBREVIATIONS
2-PG: 2-phospoglycolate
3-PGA: 3-phosphoglycerate
or-KG: alpha-ketoglutarate
ADP: adenine diphosphate
AMP: adenine monophosphate
ATP: adenine triphosphate
At: Arabidopsis thaliana
A.thaliana: Arabidopsis thaliana
CaClZ: calcium chloride
CCM: carbon concentrating mechanism
cDNA: complementatry DNA
cMDH: chloroplastic malate dehydrogenase
C02: carbon dioxide
DCT: dicarboxylate transporter
DHAP: dihydroxyacetone phosphate
DiT: dicarboxylate transporter
DNA: deoxyribonucleic acid
EGT: endosymbiont gene transfer
E. coli: Escherichia coli
ER: endoplasmic Reticulum
EST: expressed sequence tag

E-value: expectation-value

FAD: flavin adenine dinucleotide
FBPase: fructose-1,6-bisphosphatase
Fd: ferredoxin

Fd-GOGAT: glutamate synthase
Frcl,6BP: fructose-1,6-bisphosphate
Frc6P: fi'uctose-6-phosphate

Gal: galactose

GallP: galactose-l-phospbate
GAPDH: glyceraldehydes-B-phosphate dehydrogenase
GDC: glycine decarboxylase

gDNA: genomic DNA

GFP: green fluorescent protein
GGT: glutamatezglyoxylate aminotransferase
Glc: glucose

Glc 1 P: glucose- 1 -phosphate

Glc6P: glucose-6-phosphate

Glu: glutamate

Gluc6P: gluconate-6-phosphate
Gly3P: glycerol-3-phosphate
GLYK‘ glycerate kinase

GOGAT: glutamate synthase

GOX: glycolate oxidase

GPT : glucose-6-phosphate/phosphate translocator

xi

Gs: Galdieria sulphuraria

G. sulphuraria: Galdieria sulphuraria

GS: glutamine synthetase

H+: proton

H20: water

H202: hydrogen peroxide

HGT: horizontal gene transfer

HPR: hydoxypyruvate reductase

IPTG: isopropyl B-D-l-thiogalactopyranoside
Ki: inhibitory constant

KM: Michaelis-Menten constant

Li: lithium

LiAc: Lithium acetic acid

Mal: malate

MCF: mitochondrial carrier family

MgCl2: Magnesium chloride

mMDH: mitochondrial malate dehydrogenase
MP: maximum parsimony

NAD(P): nicotinamide adenine dinucleotide (phosphate)
NAD(P)H: reduced nicotinamide adenine dinucleotide (phosphate)
NJ: neighbor joining

NDP: nucleotide diphosphate

NH3: ammonia

xii

NH4+: ammonium ion

02: oxygen

0AA: oxaloacetic acid

OPPP: oxidative pentose phosphate pathway
pABA: para-aminobenzoic acid

PCR: polymerase chain reaction

PGK: 3-phosphoglycerate kinase

PEP: phosphoenolpyruvate

PGP: phosphoglycolate phosphatase

Pi: orthophosphoric acid

PLP: pyridoxal-phosphate

pMDH: peroxisomal malate dehydrogenase
pPT: plastid phosphate translocator

PPT: phosphoenolpyruvate/phosphate translocator
PR: photorespiration

Pyr: pyruvic acid

RibSP: ribose-S-phosphate

RNA: ribonucleic acid

RubisCO: ribulose- l ,5-bisphosphate carboxylase/oxygenase
RuBP: ribulose- 1 ,5-bisphosphate

S. cerevisiae: Saccharomyces cerevisiae
SGT: serinezglyoxylate aminotransferase

SHMT: serine hydoxymethyltransferase

xiii

TCA: tricarboxylic acid

THF: tetrahydrofolate

Tic: translocon at the inner chloroplast envelope
Toc: translocon at the outer chloroplast envelope
TP: triose-phosphate

TPT: triose-phosphate/phosphate translocator
UDP: uridine diphosphate

UMP: uridine monophosphate

XPT xylulose-S-phosphate/phosphate translocator

xiv

INTRODUCTION

PLASTID EVOLUTION FROM THE PERSPECTIVE OF METABOLIT E
TRANSPORT PROTEINS

INTRODUCTION

Approximately three billion years ago, an ancestor of the extant free-living cyanobacteria
introduced oxygenic photosynthesis to a literally oxygen-free atmosphere (Blankenship
and Hartman, 1998). Descendants of these early oxygenic photosynthetic organisms are
sustaining the oxygen concentration in the present Earth’s atmosphere (Blankenship,
1992; Cavalier-Smith, 2006; Falkowski and Godfrey, 2008). Sunlight is absorbed and
utilized by the photosynthetic apparatus for withdrawing electrons fi'om water, thereby
generating a proton gradient and oxygen (02) as a byproduct. The electrons and protons
are stored as usable chemical energy to assimilate inorganic nutrients (i.e., C, N, P and S)
into organic matter. Thus, autotrophic organisms are of central importance for the Earth,
providing the essential 02 for cellular respiration of aerobic organisms, and are the
primary producers at the base of almost every food chain (Field et al., 1998).

More than one billion years ago it is likely that a phagotrophic eukaryote gained
the ability to conduct photosynthesis by engulfing a cyanobacterium-like prokaryote, thus
changing fiom a heterotrophic to a mixotrophic and subsequently autotrophic lifestyle
(Yoon et al., 2004; Cavalier-Smith, 2006). This permitted the invasion of and thriving in
new habitats. Over millions of years the endosymbiotic cyanobacterium evolved into the
organelle plastid (Reyes-Prieto et al., 2007). Today’s photoautotrophic organisms exist
ubiquitously on the Earth with a remarkable diversity of species and appearance
(Bhattacharya et al., 2004). The smallest and largest photosynthetic eukaryotic organisms
described to date are the green micro-alga Ostreococcus tauri (Lanier et al., 2008) and
the Giant Sequoia tree, respectively. The red alga Galdieria sulphuraria and green alga

Chlamydomonas species adapted to the upper (56°C) (Gross, 2000) and lower

temperature limits (Morgan-Kiss etal., 2006) of eukaryotic life. Phototrophic organisms
recently attracted much attention as renewable and sustainable sources for biomass and
energy production and as alternative bioreactors to the established eubacteria, yeasts and
human cell lines (Gordon and Polle, 2007; Field et al., 2008; Raja et al., 2008; Rubin,
2008). Plastids are the site of carbon dioxide, nitrite, and sulfur assimilation (Calvin,
1962; Crawford, 1995; Kopriva, 2006). They partially or fully conduct the biosynthesis
of various metabolite classes, such as fatty acids, starch, amino acids, nucleic acids,
isoprenoides (chlorophyll, carotenoids, gibberelline) or phenylpropanoids (cell wall
components, pesticides, anthocyane) (Weber et al., 2005). To understand and fully utilize
the potential of photosynthetic eukaryotes, it is helpful to reconstruct the events that led
to plastid origin and decipher the common features and specific adaptations in extant,
distantly related photosynthetic organisms.

Plastid origin from a cyanobacterial—like prokaryote was postulated already at the
turn of the 19th and 20th century by microscopists Sachs, Altrnann, and Schimper
(reviewed in Kutschera and Niklas, 2005) and led in 1971 to the “Theory of
Endosymbiosis” by Lynn Margulis (Margulis and Nealson, 1989). Based on numerous
single and multi-gene phylogenetic analyses fiom diverse plastid genomes and nuclear-
encoded plastid-targeted genes (Martin et al., 2002; Reyes-Prieto and Bhattacharya,
2007a), it is now commonly accepted that the primary endosymbiosis had been a single
event in eukaryotic evolution and all photosynthetic species containing primary plastids
have been recently summarized in the supergroup Archaeplastida (Adl, 2005). Within
approximately 150 million years, the unicellular primordial alga (i.e., the protoalga) had

been established, and it gave rise to the three major lineages of the Archaeplastida:

Glaucophyta (glaucophyte algae), Rhodophyceae (red algae) and Chloroplastida (green
alga/plants) (Y oon et al., 2004;Ad1, 2005; Reyes-Prieto and Bhattacharya, 2007b).
Multicellularity evolved independently in the Rhodophyceae and in the Chloroplastida,
whereas only the Chloroplastida mastered the colonization of the land. This was
accompanied by an extraordinary radiation of the existing species (Bhattacharya et al.,
2004). Heterotrophic eukaryotes later captured green or red algae, respectively,
transforming them to secondary plastids of green algal or red algal origin (Gould et al.,
2008). The process of secondary endosymbiosis gave rise to a plethora of new species
mainly micro- and macro-algae, dominating aquatic environments as major primary
producers (Field et al., 1998). Serial secondary endosymbiosis by replacing a secondary
plastid by another one and even tertiary endosymbiosis via an engulfinent of algae
harboring a secondary plastid by another heterotrophic eukaryote are evident (Gould et
al., 2008). Also of interest is the complete or partial loss of plastid functions in several
species (Fast et al., 2001). Noteworthy is also continuing evolution of novel
photosynthetic organisms of primary endosymbiotic origin, as it lms been recently
reported for the host species Paulinella (Nowack et al., 2008).

The antiquity of the last common ancestor, the recurrent acquisition of the plastid
and species radiation has scrambled the evolutionary signal of plastid origin. In the last
decade, plastid-, EST- and complete genome sequence projects fi'om a diverse group of
photosynthetic organisms were initiated by the research community to clarify the
complex host-endosymbiont partnership. The green lineage has been followed from the
unicellular and multi-cellular green algae to mosses (Cove, 2005; Merchant et al., 2007),

gymnosperms and several mono- and dicotyledonous plants (Arabidopsis Genome

Initiative, 2000; Goff et al., 2002; Tuskan et al., 2006; Pavy et al., 2007). A catalog of
sequences fi'om diverse micro- and macro-a1 gae with primary, secondary and even
tertiary plastids was initiated to compile red algae evolution (Matsuzaki et al., 2004). The
genome of the glaucophyte alga Cyanophora paradoxa is currently sequenced to
complement the available sequence resources with the third major lineage of the
Archaeplastida (Reyes-Prieto et al., 2006; Loeffelhardt, 2007). Besides all the diversity,
common themes and specialization in plastid evolution can be now validated and
addressed:

Endosymbiont gene transfer (EGT) to the host nucleus massively reduced the
plastid genome size (Tirnmis et al., 2004). The model-plant Arabidopsis thaliana has a
chloroplast genome of ~155 kb with 87 protein-coding genes (Sato et al., 1999), which is
just a small fraction of extant cyanobacteria! genomes. On the other hand, bioinformatic
and proteomic predictions assume nearly 4,500 proteins in the organelle (Martin et al.,
2002; Sun et al., 2008). Expression and re-import of these nucleus-encoded plastid
targeted proteins has been predicted for a long time to be a hallmark for a successful
organelle establishment (Dyall et al., 2004). An intriguing question is to what extent and
for what purpose cyanobacteria] genes had been transferred and maintained in the host
nucleus. According to Sato et a1. (2005) and Martin et al. (2002), the A. thaliana nuclear
genome harbors between 1,200 — 4,500 genes of cyanobacteria] origin, respectively. The
majority of these cyanobacteria] genes have non-plastid function (Martin et al., 2002),
hence, the present plastid proteome evolved from multiple sources including
cyanobacterial, host proteins and eubacterial genes acquired by horizontal gene transfer.

Data from glaucophytes and red algae suggest a much smaller portion of transferred

cyanobacterial genes and in contrary to A. thaliana, the majority of these candidates are
plastid targeted (Reyes-Prieto et al., 2006). Although these findings are not in full
agreement, they still demonstrate a Significant and complex contribution of EGT to the
evolution of the host cell. Host genes and endosymbiont genes have been replaced by one
another, duplicated, diverged and several times redirected to a new compartment or even
existing homologous genes recombined with each other. A challenge will be to follow up
these rearrangements in numerous model organisms and assemble the tree of
photosynthetic eukaryotes.

Prerequisites for the gene transfer to the nucleus were (i) the regaining
transcriptional and translational activation and (ii) the retargeting of plastid-needed
proteins (Martin, 2003). Recent studies demonstrated the transfer of plastid-encoded
antibiotic resistance genes that were introduced by plastome transformation to the
nucleus, including gene activation, within a short timefi‘ame (Huang et al., 2003;
Stegemann et al., 2003). More elaborate must have been the evolution of the plastid
protein import machinery. The sophisticated Toc (Translocon at the outer chloroplast
envelope) and Tic (Translocon at the inner chloroplast envelope) system evolved not only
to import the majority of the proteins across both envelope membranes, but also to
distribute them to one of the six possible destinations (outer and inner envelope
membrane, interrnembrane space, plastid stroma, thylakoid membrane and lumen) (Inaba
and Schnell, 2008). Comparative analysis of the Toc and Tie apparatus across many
photosynthetic eukaryotes points again to a monophyly of plastids and suggests its early

establishment, likely already in the protoalga. Core parts of the cyanobacterial protein

transport machinery have been recruited and were functionally replaced or improved with
new elements, which are absent in the cyanobacterial membrane (Gould et al., 2008).

Another major control point in plastid biogenesis was the establishment of
coordinated division of endosymbiont and host cell (Miyagishima, 2005). Unrestricted
growth of the endosymbiont would harm the host and inhibiting its division would
ultimately lead to its loss. The plastid division apparatus follows the same evolutionary
path described earlier for the protein import apparatus (Miyagishima, 2005). The
organelle division is based on ancient bacterial binary fission and several genes with
cyanobacterial and other eubacterial origin are still conserved in extant plastids. The
nuclear genome took control over the expression of most of the genes (Glynn et aL,
2007). In land plants, the transfer is completed and none of the proteins are encoded on
the organelle genomes sequenced to date, but some algal genomes sporadically still
encode division factors (Miyagishima, 2005; Glynn et al., 2007). Analogous genes
replaced or had been added as division components originated from eukaryotic and
eubacterial sources. Even after the split of the Archaeplastida, lineage-specific
remodeling occurred. This makes it difficult to provide a universally valid plastid
division model and in particular challenging to find new division related genes via
sequence-based comparative analysis (Glynn et al., 2007).

From the engulfinent of the plastid ancestor to the split into the Glaucophyta, red
algae and green algae/plants, a period of 150 million years has been estimated (Y oon et
al., 2004). All above-mentioned major events in plastid origin are results of a complex
multi-gene acquisition and build the foundation of all descendants of the proto-alga. To

date, it is not satisfactorily answered how the two organisms managed to coexist long

enough to provide sufficient time for the transformation of the endosymbiont into an
organelle.

In the following paragraph and in chapters one and two I will outline the potential
of metabolite exchange and integrating metabolisms to set-up a strong mutual symbiosis,
which ultimately led to an enslavement of the cyanobacterimn by the protist. This initial
metabolic interaction might have provided the time needed for the emergence of a full-
fledged organelle, including large scale EGT and establishment of effective protein
import apparatus and division machinery. My aim was to use comparative genomics to
unravel the differences of the metabolite transport protein composition between the
plastid inner envelope membrane and the plasma membrane of free-living cyanobacteria.
In this work, I could Show that recruitment of host-derived metabolite transporters to the
plastid envelope membrane was very likely a driving force in the transition of a
mutualistic endosymbiosis to the proto-alga. Chapters one and two had been prepared in
collaboration with the laboratory of Debashish Bhattacharya (University of Iowa) and
were published in two different scientific journals. In

In the proposed scenario, a complete cyanobacterium was engulfed by a
phagotrophic eukaryote and escaped digestion in the food vacuole. Eventually, the
membrane surrounding the cyanobacterium was lost or merged with the cyanobacterial
outer membrane (Cavalier-Smith, 2000), resulting in two membranes marking the barrier
to the new environment: the cytosol of the host. The composition of the outer membrane
changed considerably over evolutionary time scales (Block et al., 2007). Eubacterial
lipoproteins and lipopolysaccharides are absent fi'om higher plant chloroplasts and the

phospholipid phosphatidylcholine was specifically introduced into the outer envelope

membrane from the endomembrane system of the host (Douce and Joyard, 1990;
Cavalier-Smith, 2000). Extant Archaeplastida display an intimate connection between the
Endoplasmic Reticulum (ER) and the envelopes of the plastid. Lipids are synthesized in
the plastid stroma and a sophisticated traffic system has been established between plastid
and ER in both directions for the biogenesis of plastidial and extra-plastidial membranes
(Benning, 2008). As a conserved feature of extant primary plastids, the Tic/Toe protein
import apparatus recognizes most nuclear-encoded genes via a N—terminal leader
sequence of 25 - 125 amino acids (Soil and Schleifi’, 2004). Still, some plastid-targeted
proteins do not exhibit an obvious leader sequence and an alternative protein
translocation route via the ER and Golgi vesicle sorting system has been recently
discovered (Villarejo et al., 2005). The host-plastid interaction was repeatedly established
through endosymbiosis in secondary plastid evolution. Three to four envelope
membranes surround secondary plastids (Bhattacharya et al., 2004) and nuclear-encoded
proteins are delivered by a multi-partite leader sequence through the concerted action of
the ER sorting system and the Tic/Toc related import machinery of the organelle (Kilian
and Kroth, 2003 ).

These direct contacts between ER and plastid support the idea that vesicle
transport to and from the plastid was initiated at an early stage of endosymbiosis. ER or
Golgi-derived vesicles initially might have fused randomly with the outer membrane of
the cyanobacterium, releasing their contents into the periplasmatic space and the proteins
were inserted into the cyanobacterial plasma membrane or taken up by the cyanobiont.
Thus, even if insertion of host metabolite transporters into the cyanobiont’s plasma

membrane were initially not very efiicient, it would have given the host the opportunity

to connect the cyanobiont with its cytosol and tap into photosynthates fi'om the
cyanobacterium. This could have become a selective advantage for the phagotrophic
eukaryote and the endosymbiont and favored the establishment of a permanent
partnership.

In chapter one we address the phylogenetic origin of the plastid-targeted triose-
phosphate/phosphate transporter (TPT). As a starting point, I used the available EST
database from the unicellular red alga G. sulphuraria to annotate and assemble the
homologous plastid phosphate translocator (pPT) genes. This work was part of the
publication: “EST analysis of the thermo-acidophilic red microalga Galdieria
sulphuraria reveals potential for lipid A biosynthesis and unveils the pathway of carbon
export from rhodoplasts” (Weber et al., 2004). The results led us to hypothesize that the
ancestor protein of the TPT has evolved fi'om a host-derived endomembrane transporter,
which was “mistargeted” to the cyanobacterial membrane and thus enabled a metabolic
connection between the endosymbiont and the host. The TPT as a member of the pPT
family was the first solute transport protein identified on a molecular basis in the inner
envelope membrane in higher plants (F liigge, 1999). This transporter is of particular
interest, because it is the major route for export of assimilated carbon from the plastid
into the cytosol dining the day in seed plants. Our analysis in chapter one revealed that
homologous TPT genes are widespread in the red algal lineage including chromalveolate
algae with secondary plastids, but related genes are absent from cyanobacteria and other
prokaryotes. Export of triose-phosphate is truly an innovation of plastid evolution.

The pPT family originated from nucleotide-sugar transporter present in the

endomembrane system of extant eukaryotes. However, none of the extant pPT members

10

do transport nucleotide sugars, thus leading to the question how a nucleotide sugar
transporter might have fimctioned in the initial establishment of a metabolic connection
between cyanobiont and host. Nucleotide-sugars are the substrate for literally all
glycosylation reactions in a cell. They are incorporated into disaccharides, oligo- and
polysaccharides, which are utilized as compounds for transport, storage, structural
integrity and post-translational modifications in the cell. Recently, (Deschamps et al.,
2008) highlighted the importance of the nucleotide-sugar transporters for merging the
storage polysaccharide metabolism in host and cyanobiont. According to their hypothesis,
both organisms initially harbored complete and independent sets of enzymes to
synthesize, store and mobilize carbon in their respective compartments. The insertion of
an NDP-sugar transporter would have allowed the host to withdraw the cyanobacterial
starch precursor ADP-glucose fiom the cyanobiont. The key assumption of this
hypothesis is that ADP-glucose represents an activated form of organic carbOn that has
already been committed to storage, thus it can be withdrawn from the evolving plastid
without disturbing its primary carbon metabolism. However, ADP-glucose is a poor
substrate for the soluble starch synthase in heterotrophic eukaryotes. To counter this
argument, it has been argued that the gene encoding soluble starch synthase of
cyanobacterial origin has been transferred to the host nucleus at an early stage and was
subsequently expressed in the cytosol of the host. Importantly, this would only require
the transfer of a single gene to the host. It is not necessary for the gene product to be
retargeted to the cyanobacterium and proteins for mobilization of the host a-glucans are
already present. Thus, the proposed scenario is a very simple one in evolutionary terms —

it does require only a single gene transfer and ‘mistargeting’ of a single endomembrane-

11

derived transporter. The scenario would be even simpler if the ancestral host starch or
glycogen synthase would have accepted ADP-glucose as precursor, even with low
affinity. In this case, the initial connection would not have required EGT or evolutionary
innovations.

Nucleotide-sugar transport proteins operate in a strict counter-exchange mode and
the transport direction depends on the concentration of the substrates on either sides of
the membrane. The export of NDP—sugar is compensated with an import of nucleotide-
monophosphates, such as AMP or UMP (Handford et al., 2006; Rollwitz et al., 2006). In
the endosymbiosis context, this counter-exchange mode of transport would have
prevented the depletion of the cyanobacterial nucleotide pool. A relatively higher
concentration of ADP—glucose at its side of biosynthesis inside the endosymbiont would
favor its eXport and immediate consumption in the cytosol. As outlined above, the
transporter was likely inefficiently inserted and only carbon committed to storage was
drained, thus it did not affect the viability of the symbiont (Deschamps et al., 2008).
However, to date, a paralogous NDP-sugar export system has not been identified in the
inner envelope membrane of any extant photosynthetic eukaryote. Identification of such a
transporter would represent the missing link that would lend strong support to the
“metabolic symbiosis and birth of the Plant kingdom” hypothesis put forward by
Deschamps et a1. (2008). According to my work presented in chapter one, the pPT gene
family with at least three distinct members already evolved in the last common ancestor
of the red algae and the green algae and plants, shortly after the initial endosymbiosis was
established (Weber et al., 2006). They all are equipped with an N—terminal leader

sequence for an efficient retargeting and insertion by the Toc and Tic system. As a result,

12

the major flux of sugar-phosphates across the membrane of plastid in higher plants was
reorganized fi'om an NDP-sugar transporter towards a triose-phosphate (TP)/phosphate,
phosphoenolpyruvate (PEP)/phosphate and glucose-6-phosphate (Glc6P)/phosphate
shuttle system (Flfigge, 1999). The transport systems fit perfectly into metabolic
compartrnentalization of higher plants: triose-phosphate is exported to the cytosol and is
mainly used for sucrose or cell wall biosynthesis. PEP is taken up from the cytosol to
drive fatty acid biosynthesis and synthesis of compounds by the shikimate pathway (i.e.
aromatic amino acids and a heterogeneous group of phenolics). The glucose-6-phosphate
transporter (GPT) provides glucose-6-phosphate (G6P) for starch synthesis and the
oxidative pentose phosphate pathway (OPPP) in heterotrophic tissues, or the Ribulose-
1,5-bisphosphate Carboxylase/Oxygenase (RubisCO) bypass reaction in mixotrophic
tissues.

In summary, the most likely scenario in evolutionary terms is that the NDP-sugar
transporter represented the starting point for connecting the previously independent
carbon metabolisms of host and cyanobiont. Gene duplications and further radiation of
the gene family delivered the genetic material for the evolution of functional
diversification, eventually promoting the evolution of a more efficient targeting system to
the plasma membrane of the endosymbiont.

The findings about the phylogenetic origin of the plastid phosphate transporter
family encouraged me to extend the phylogenomic analysis of the repertoire of plastid
metabolite transporters to a broad range of different photosynthetic organisms. In present-
day plastids of seed plants, photosynthesis provides the energy for the Calvin-Benson-

Cycle and several other anabolic pathways, including the synthesis of starch, fatty acids,

13

several amino acids, nucleic acids, and the reductive assimilation of inorganic ions like
nitrate and sulfate. For each pathway, a number of precursors and intermediates have to
be transported across the envelope membrane (Weber et al., 2005). Plastids must be
extensively connected to the cytoplasm to distrrbute the numerous products within the
cell. In contrast, the cyanobacterial membranes generally prevent leakage of their primary
metabolites to the surrounding The plasma membrane is mainly configured for the
uptake of inorganic and organic nutrients and the excretion of waste products or
components for the periplasmic space, the outer membrane, and cell wall (Kaneko et al.,
1996).

In chapter two we addressed two questions concerning the transporter evolution
of the plastid inner envelope membrane. First, we sought to estimate the contribution of
host and symbiont on the transporter repertoire in photosynthetic eukaryotes. Is the pPT
the rule or the exception for host proteins, which had been inserted into the
cyanobacterial plasma membrane? Second, we determined the differences between the A.
thaliana transporter set and photosynthetically active prokaryotic and eukaryotic
organisms. This would give us for the first time an insight into the conserved and lineage-
specific metabolite “permeome” of higher plant plastids. To address these hypotheses, we
analyzed the phylogenetic origin and distribution of well-annotated transporter proteins
from A. thaliana. Transport proteins have characteristic hydrophobic transmembrane
spans to reside in the membrane as well as hydrophilic domains to allow interaction and
transport of their substrates (Schwacke et al., 2003). In silica searches for genes in the A.
thaliana genome with a predicted plastid target sequence and hydrophobic domains

provided an initial catalogue of candidate genes (Weber et al. 2005). Several plastid-

l4

Specific proteomic studies confirmed the presence and extended the list of proteins
targeted to inner envelope membranes of plastids to approximately ~130 putative
metabolite transporter proteins (Sun et al., 2008). This number does not include genes
from the Toc, Tic and thylakoid-related protein import machinery and prominent
thylakoid-localized electron-transporting proteins of the photosynthetic core complex,
such as D1, D2, cytochrome b6, cytochrome f, or proton-coupled ATP—synthase subunits.
Our results for the first time revealed that the host has been the major source for
metabolite transporter acquisition by the plastid. We also found an tmexpectedly high
proportion of Chlamydiae-derived genes encoding plastid metabolite transporters. The
group of Debashish Bhattacharya examined this result in more detail and identified a total
of 55 genes in Plantae with chlamydial origin (Moustafa et al., 2008). A similar result
was demonstrated for the red alga Cyanidioschyzon meralae (Huang and Gogarten,
2007). This provides a minor but crucial third genome source for plastid evolution by
horizontal gene transfer or even an “endosymbiotic” gene transfer from a parasitic
Chlamydiae bacterium. Intriguingly, most of the metabolite transport proteins were
distributed in both green and red algae and to a much lesser extent present in only one of
them. This suggests that the initial phase of the proto-alga, before the split into the
Archaeplastida lineages, was accompanied by a massive invasion of the envelope
membrane with host-proteins.

As mentioned earlier, the extant pPT family evolved from endomembrane NDP-
sugar transporters. The three pPT subfamilies do not simply encode differentially
expressed or targeted gene products with identical functions. They evolved distinct

substrate specificities and define the major flux of carbon across higher plant plastid

15

envelope membranes (F lfigge, 1999). The phylogenetic tree shown in chapter one argues
for highly conserved orthologous sugar-phosphate transporters with the preferred
substrates Pi, TP, PEP, and Glc6P in the red alga G. sulphuraria. This is in contrast to the
enormous differences in how starch and soluble sugars are synthesized in the green and
the red lineages (Viola et al., 2001). Red algae harbor an UDP-glucose specific starch
synthase of ancient eukaryotic origin, which is localized in the cytosol. The soluble starch
synthase fiom green algae and plants is closely related to cyanobacterial genes and
incorporates ADP-glucose into the polysaccharide chain in the stroma of the organelle
(Deschamps et al., 2008). Probably, UDP- and ADP-glucose dependent starch synthesis
was still manifested in the genome of the proto-alga. By the time the common ancestor
split up into the Rhodophyceae and Chloroplastida, the carbon metabolism was far from
being established as we find it in extant species. The pathway has been relocated to the
plastid stroma in green plants and algae (Deschamps et al., 2008) or stayed and gradually
developed in the cytosol of Rhodophyceae. In addition, red algae produce the
predominant soluble carbohydrate floridoside, composed of a glycerol and galactose
moieties (Viola et al., 2001) and plants synthesize the disaccharide sucrose from UDP-
glucose and fi'uctose-6-phosphate (Huber and Huber, 1996).

Based on phylogenomic analysis, I developed the hypothesis that transport of
glycolytic intermediates, i.e. PEP, TP, and Glc6P, was a successfirl invention in plastid
biogenesis and is conserved in primary photosynthetic eukaryotes. Hence, if the
hypothesis is correct, the homologues in G. sulphuraria should represent truly
orthologous genes with identical substrate specificities as their counterparts in the green

lineage. However, the major differences in carbon metabolism between photosynthetic

16

eukaryotes challenge the hypothesis. Possibly, the ancient NDP—sugar transporters
continued to evolve divergently in red algae from the TPT, PPT and GPT of plants. To
find an answer to these contradictory hypotheses, I functionally tested the pPT genes
from G. sulphuraria by recombinant expression and biochemical characterization, and
comparison to their counterparts in higher plants. The results are presented in detail in
chapter three. I discussed alternative mechanisms in green plants and red algae to
organize their primary carbon partitioning between rhodoplasts and cytosol. Prior to this
study, carbon assimilation and especially the metabolic fate of the photosynthates in
Rhodophyceae did not receive great attention compared to green algae and land plants.
This does not match with the major role of red algae in the global carbon cycle, since
they dominate carbon fixation in aquatic environment, which approximately accounts for
half of the global photoautrophic C02 assimilation (Field et al., 1998). In chapter three, 1
demonstrate that the red alga G. sulphuraria, in comparison to the Plantae, possesses an
export system for triose-phosphates with very high specificity and afiinity. In red algae,
TPT controls the flux of carbon during autotrophic and heterotrophic growth conditions
and, in contrast to plants, glucose-6-phosphate transport activity does not exist in the red
alga as a major import route for sugar-phosphates. The lower affinity TPT in green plants
might represent an adaptation to maintain a relatively high concentration of TP that is
required for driving plastidial starch biosynthesis. The TPT protein fiom G. sulphuraria
is a good example for the biochemical diversity of known proteins in distantly related
photoautotrophic organisms and implies that this gene pool is an underestimated toolbox

to engineer plants for applied sciences.

17

Besides understanding of the origin and contribution of metabolite carriers for
plastid evolution, a major challenge is linking the catalogue of uncharacterized transport
proteins to their biological function. A defining feature of eukaryotes is their cellular
compartrnentation. The cytosol is separated from the internal space of mitochondria,
plastids, vacuole, peroxisomes, nucleus, and ER by biological membranes. Lipid bilayers
create a diffusion barrier for hydrophilic and charged molecules and define separate
compartments for enzymatic reactions. In a single cell, pathways can be confined in a
specific or even partitioned in more than one compartment. In multi-cellular organisms,
several pathways are connected between different cells, tissues or even organs and often
separated by more than one membrane. Thus, metabolite transport proteins are required
for nutrient and metabolite uptake, secretion of toxic by-products, to sustain and even
regulate the metabolic flux through pathways and distribute a single metabolite to distinct
compartments at the same time (Kunze et al., 2002). One quarter of the A. thaliana
genome encodes for proteins with at least one transmembrane span that are predicted to
be embedded in one of the cellular membranes (Schwacke et al., 2003; Komatsu et al.,
2007). This nrunber includes numerous receptor kinases, electron transport chain or
protein import components fi'om mitochondria and chloroplasts. Nevertheless, it is
estimated that up to 40% of these membrane-localized proteins could represent
metabolite transporters (Ward, 2001). Most of the predicted transporters lack homology
to known proteins from other species and less than 20% of the putative carriers are
characterized at the biochemical and molecular level (Barbier-Brygoo et al., 2001).

During the day, the major flux across plastid membranes is the export of triose-

phosphate, which is catalyzed by the TPT (Flitgge, 1999). TP is the product of the

18

reductive part of the Calvin-Benson-cycle (Calvin, 1962). Carboxylation of ribulose-l,5-
bisphosphate (RuBP) by the RubisCO results in two molecules of 3-phosphoglycerate (3-
PGA) and the following two enzymatic reactions generate triose-phosphate, consuming
ATP and NADPH provided by the photosynthetic light reactions.

Atmospheric oxygen (02) competes with C02 for the acceptor molecule RuBP,
yielding significant amounts Of 2-phosphoglycolate (2-PG) and 3-phosphoglycerate (3-
PGA) (Ogren, 1984). In contrast to 3-PGA, 2-PG cannot be metabolized further in the
Calvin-Benson-cycle. It is hydrolyzed in the chloroplast to glycolate and exported to the
cytosol to be recycled by the photorespiratory pathway (F igrue l) (Douce and Neuburger,
1999). Measurements on chloroplast membranes demonstrated the presence of a transport
system, which is able to export glycolate with rates required to cope with the flux through
the photorespiratory pathway (Howitz and McCarty, 1991). Two molecules of the 02
carbon glycolate are converted to one 03 molecule glycerate in a complex cycle
organized in peroxisomes, mitochondria and cytosol. Glycerate is taken up by the
chloroplast, phosphorylated and reduced to triose-phosphate (Howitz and McCarty, 1991;
Schwarte and Bauwe, 2007). Numerous studies identified the enzymes, which hydrolyze
2-PG to glycolate (Somerville and Ogren, 1979), oxidize and transaminate glycolate to
glycine in the peroxisomes (Kendall et al., 1983; Liepman and Olsen, 2001; Igarashi et
al., 2003; Queval et al., 2007). Two molecules of glycine are converted to serine in the
mitochondria (Somerville and Ogren, 1981; Vol] et al., 2006; Engel et al., 2007), and
serine is deaminated to hydroxypyruvate and reduced to glycerate in the peroxisomes
(Liepman and Olsen, 2001; Igarashi et al., 2003) (Murray et al., 1989). During the 1970s

and 19803, forward genetics screens with mutants of A. thaliana and Hordeum vulgare

19

has been a powerful tool to identify many of the photorespiratory genes (Ogren, 1984).
Isolated mutants grew normally under elevated carbon dioxide levels, but the enzymes
were essential under photorespiratory conditions (Ogren, 1984). Actually, genetic
dissection of photorespiration using Arabidopsis mutants paved the way for A. thaliana
as a model organism in plant biochemical genetics and plant physiology (Somerville,
2001). Surprisingly, however, map-based cloning of the photorespiratory A. thaliana
genes did not reveal a single transporter of the mandatory shuttle systems needed for the
described conversion of glycolate to glycerate between chloroplast, peroxisomes, and
mitochondria.

The only isolated transport protein linked to the photorespiratory pathway and
identified by forward genetic screen is the dicarboxylate transporter DCT 1/DiT2.l in the
inner envelope membrane of chloroplasts (Figrn'e 1) (Somerville and Ogren, 1983; Renne
et al., 2003). During glycine oxidation in the mitochondria, two molecules of glycine are
converted to one molecule of serine, NADH, C02 and ammonia. The DCT protein is
required in the so-called nitrogen cycle of the photorespiratory pathway to re-assimilate
the released ammonia (Keys, 2006). The concerted activity of glutamine synthetase (GS)
and glutamate synthase (GOGAT) incorporates ammonia into 2-oxoglutarate in the
chloroplast yielding glutamate. Export of glutamate depends on the activity of the
DCT/DiT2.l protein. An A. thaliana mutant defective in this carrier cannot export
glutamate to the cytosol and the amino-donor for the peroxisomal aminotransferase
reaction of glyoxylate to glycine is depleted fast (Renne et al., 2003). Similarly, impaired

GS or GOGAT enzymes short-circuit the re-assimilation of 2-PG, demonstrating that the

20

nitrogen cycle is essential for the growth under photorespiratory conditions (W allsgrove
et al., 1987; Coschigano et al., 1998).

With a concentration of 380 ppm C02 and 210,000 ppm of 02 in the air, the
carboxylation to oxygenation ratio of RubisCO is approximately 3:1 in C3 plants without
a carbon concentrating mechanism (CCM) (Sharkey, 1988). Hence, a massive flux
through the carbon and nitrogen cycle of photorespiration results (Figure l). The protein-
mediated transport of glycerate/glycolate (Howitz and McCarty, 1991),
glycolate/glycine/serine/glycerate/glutamate/Z-oxoglutarate (Reumann et al., 1998) and
glycine/serine (Walker et al., 1982; Yu et al., 1983) is documented on isolated
membranes from chloroplast, peroxisomes and mitochondria, respectively. However the
genes encoding the corresponding transporters have not yet been identified at the
molecular level. Several reasons could explain why the forward genetic screens were
unsuccessful to identify additional metabolite transporters. So far, only plants that show a
severe decrease in chlorophyll content within four days after shifting fiom high C02 to
ambient air have been further analyzed (Somerville, 2001). Prolonging the period of
photorespiratory conditions could serve to identify weaker alleles for the pale green
phenotype. Overlapping substrate specificity of metabolite transporters could mask the
phenotype of a single gene mutant. The mitochondrial glycine decarboxylase complex for
glycine oxidation is absolutely essential for the fitness of the plant. The gdc knockout
mutant is lethal at ambient and elevated carbon dioxide and thus demonstrate that not
every gene in the photorespiratory pathway is dispensable under elevated C02 conditions
(Engel et al., 2007). It is tempting to speculate that this could be true for other proteins,

such as the porin-like channel of the peroxisomes, which facilitates the transport of a

21

broad range of mono- and dicarboxylate anions (Reumann et al., 1998). A loss of this
activity would presumably interfere with the peroxisomal glyoxylate cycle to mobilize
fatty acids (Graham, 2008) and the oxaloacetate/malate Shuttle to exchange reduction
equivalents with the cytosol (Reumann and Weber, 2006).

In chapter four and five I revisited the highly compartmentalized photorespiratory
pathway and initiated a search for the long outstanding metabolite transporters. In chapter
four I summarize putative alternative pathways leading to ammonia re-assimilation in the
organelles mitochondria and chloroplast. I emphasize the required, but not yet identified
transport systems. In chapter five I took advantage of publicly available bioinforrnatics
databases to screen for genes with two or more predicted transmembrane spans, which
are highly co-expressed with known photorespiratory genes (Schwacke et al., 2003;
Steinhauser et al., 2004). I initiated a reverse genetics candidate approach for five of the
best 32 co-expressed putative metabolite transporters and report on the discovery of a
novel transporter involved in the photorespiratory pathway. The long-term goal will be to

identify the transporters involved in the photorespiratory pathway in higher plants.

22

Figure 1: The photorespiratory pathway

The enzymes involved in photorespiratory pathway are compartmentalized between
chloroplasts, peroxisomes, and mitochondria (cMDH, chloroplastic malate
dehydrogenase; GDC, glycine decarboxylase; GGT, glutamatezglyoxylate
aminotransferase; GLYK, glycerate kinase; GOX, glycolate oxidase; GS/GOGAT,
glutamine synthetase/ glutamate synthase; HPR, hydoxypyruvate reductase; mMDH,
mitochondrial malate dehydrogenase; PGP, phosphoglycolate phosphatase; pMDH,
peroxisomal malate dehydrogenase; RubisCO, ribulose-l,5-bisphosphate
carboxylase/oxygenase; SGT, serine:glyoxylate aminotransferase; SHMT, serine
hydoxymethyl transferase). Photorespiratory metabolites are abbreviated as follows: 3-
PGA, 3-phosphoglycerate; Glu, glutamate; KG, or-ketoglutarate; Mal, malic acid; 0AA,
oxaloacetic acid; RuBP, ribulose-bisphosphate; THF, tetrahydofolate. The transport
proteins represented in this scheme have been characterized biochemically (white box) or
the corresponding genes have been cloned (black box). The scheme was adapted from

Reumann and Weber (2006). The figure in this dissertation is presented in color.

23

    
   
          
  

  
  

 

 

If Peroxisome
Glycerate 2x Glycolate

NAD a\“ 02
-MDHX J HPR GOX ‘ -CAT H20
NADH

V H202 —-> +

2x Glyoxylate
Glu

   
          
  
    

 

 

 

Hydroxy- 0.5 02

pyruvate

 

 

 

2x Glycine

 

 

 

Figure 1: The photorespiratory pathway

24

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33

CHAPTER 1

PHYLOGENY OF THE PLASTID PHOSPHATE TRANSLOCATOR FAMILYl

 

I This work has been published in Weber A.P.M., Linka M., and Bhattacharya D. (2006)
Single, ancient origin of a plastid metabolite translocator family in Plantae from a
endomembrane-derived ancestor. Eukaryotic Cell 5:609-612.

34

ABSTRACT

Phylogenetic analyses Show the single origin of a plastid metabolite translocator family in
the Plantae from a gene encoding an existing endomembrane-derived protein. Red algal
secondary endosymbiosis has spread a translocator gene into the ancestor of the

“chromalveolate” protists, where it has diversified into a novel clade of proteins.

INTRODUCTION
The photosynthetic organelle (plastid) in red, green, and glaucophyte algae (Plantae)
likely had a single origin fiom a cyanobacterial endosymbiosis (Cavalier-Smith, 1992;
Palmer, 2003). This ancient event (ca. 1 — 1.5 billion years ago) (Yoon et al., 2004;
Douzery et al., 2004) was followed by the transfer of genetic material from the
endosymbiont to the nuclear genome of the host, the evolution of a protein import
apparatus for the plastid, and targeting sequences for nucleus-encoded plastid-targeted
proteins, and the establishment of genome-plastome intracellular communication and
regulation (Brown et al., 2001; Fey et al., 2005; Strand, 2004). Current hypotheses
regarding plastid origin and evolution provide plausible explanations for the later stages
of organelle establishment (e.g., gene transfer to the nucleus), but do not specifically
address the initial formation of the endosymbiosis.

We hypothesize that the insertion of a metabolite antiporter into the ancestral
plastid membrane was essential for establishing the primary endosymbiosis, allowing the
ancestor of the Plantae to profit immediately from cyanobacterial carbon fixation. This

antiporter likely evolved from an existing host metabolite translocator with either

35

mitochondrial function (Bullerwell and Gray, 2004; Gray et al., 2004) or the
endomembrane system. In contrast, autotrophic free-living cyanobacteria have no
obvious need for such antiporters (i.e., bacterial translocators are not detectable with
BLAST searches). Members of the nucleotide-sugar/triose phosphate translocator gene
family (Knappe et al., 2003) are likely ancestors to plastid translocators because (i) genes
encoding these proteins are found in all sequenced eukaryotic genomes, (ii) orthologous
proteins are absent from prokaryotes, and (iii) some gene family members are targeted to

plant plastids.

MATERIAL AND METHODS

To address plastid translocator origin, we gathered available sequences (genome and

expressed sequence tag data) from the NCBI WWW, DOE Joint

Genome Institute (hgp'J/wwwjgidoegovb, Michigan State University Galdieria

. ,z r m ,, , Porphyra yezoensis

expressed sequence tag index thgp://www.kamsa.or.jp/en/plant/mrphjga/ESTO, and the

Cjzam‘dioschyzon merolae Genome Project (http://merolae.biol.s.u-tokyo.ac.jp/).

 

Homologs were identified using BLAST searches

W with an E-value cut-off of 5 0.0001. A
translocator phylogeny was inferred using protein maximum-likelihood (ML) method

(PHYML V2.4.3 WWW, neighbor joining (NJ) (PHYLIP V3.63,

, m ), maximum parsimony (MP)

 

(PAUP*V4.0blO, hgpzl/paupesitfsueduh, and Bayesian inference (MrBayes V3.0b4,

36

WEI—928121112) (for details of phylogenetic approach, see Hackett
et al., 2004). A total of 250 amino acids from 65 plastid phosphate translocators from red

and “chromalveolate” (i.e., chlorophyll c—containing) algae and land plants and their non-

plastid homologs were included in these analyses.

RESULTS AND DISCUSSION

The resulting tree (Figure 1.1) shows that plastid translocators are monophyletic and had
a single origin from an existing endomembrane translocator (bootstrap support:

ML, 100%; NJ, 98%; MP, 93%), consistent with our model. Each major group of plant
plastid translocators (i.e., TPT, triose-phosphate/phosphate translocator; GPT/XPT,
glucose-6-phosphate and xylulose-S-phosphate/phosphate translocator, respectively; PPT,
phosphoenolpyruvate/phosphate translocator, shown in green text in Figure 1.1) form
robustly supported lineages (ML and MP bootstrap values > 90%) that are sister to
homologs in the red algae (Figure 1.1, shown in red text). This result supports the
monophyly of red and green algae and land plants (e.g., Rodriguez-Ezpeleta et al., 2005)
and is consistent with the origin of the different plastid translocators in their common
ancestor; i.e., all plastid translocators are'monophyletic and each is divided into sister
groups comprised of the red and green clades. Presumably, these Plantae translocators
diversified because of the selective advantage they offer; i.e., to harvest the different
fixed carbon products of the cyanobacterial primary endosymbiont. This phylogeny
provides strong evidence for a Single primary endosymbiosis in the studied Plantae,

reflecting a critical and early step in plastid evolution. The monophyly of GPTS and

37

XPTS has previously been reported (Knappe et al., 2003) and likely reflects a more recent
plant-specific gene duplication event.

The chromalveolates are a taxonomically diverse assemblage of protists
comprised of alveolates (dinoflagellate algae, ciliates, and parasitic apicomplexans) and
chromists (cryptophytes, haptophytes, and stramenopiles) that are thought to have
ancestrally contained a plastid of red algal origin (Cavalier-Smith, 1999). This group is
yet to be substantiated in a global eukaryotic phylogeny. It is therefore of interest that the
available (i.e., apicomplexan, haptophyte, and stramenopile) chromalveolate sequences
are monophyletic in our tree (Figure 1.1) and solidly associated (ML, 80%, NJ, 95%)
with one clade of red algal proteins (i.e., C. merolae CMK114C, G. sulphuraria
HET‘39C12). The addition of cryptophyte and dinoflagellate plastid translocators is
needed to verify this result. Interestingly, preliminary biochemical analysis of the G.
sulphuraria HET39C12 translocator suggests that it is a TPT (Marc Linka unpublished
data); therefore, the ancestral chromalveolate plastid translocator was likely of this type.

The topology of the chromalveolate subtree supports a single origin of the plastid
translocator gene in the common ancestor of these species (i.e., supporting their
monophyly) from the nucleus of the red algal endosymbiont. It is notWorthy that PPT
and GPT/XPT fi'om Plantae are absent from the chromalveolates. In organisms
containing plastids of secondary endosymbiotic origin with three or four plastid envelope
membranes, transfer of a plastid phosphate translocator-related gene to the host nucleus
and retargeting to the inner plastid membrane alone are not sufficient for the export of
photosynthates to the host. Additional translocators with identical transport properties

would be required in the third, the remnant of the endosymbiont plasma membrane, and

38

the fourth membranes, the plastid endoplasmic reticulum, to connect the metabolism of
the host and the endosymbiont. This essential metabolic connection has been
accomplished in the ancestor of the chromalveolates in our tree with the red algal gene
Imdergoing diversification through duplication and divergence giving rise to the complex
branching pattern shown in Figure 1.1. The specific functions and the detailed membrane
localizations of the different chromalveolate translocators remain to be determined.

We analyzed chromalveolate translocators with significant N-terminal extensions
to determine if they are putatively plastid (or apicoplast) targeted. Using PATS

. no) and PlasmoAP

 
 

' r m), apicoplast targeting is
supported for the apicomplexan Plasmodium spp. (e.g., P. falciparum NP_703 643; PATS
probability = 0.954, PlasmoAP = 5/5 tests positive). Similarly, the N-terminal extension
in the diatom (stramenopile) Thalassiosira pseudonana Sol 5 (gene located on Scaffold
15) sequence contains a typical bipartite plastid-targeting sequence (Lang et al., 1998)
comprised of a signal sequence using SignalP WW
probability = 0.997; predicted length is 18 amino acids) followed by a transit peptide
(using TargetP, probability = 0.740). These results suggest that the chromalveolate clade
in our tree includes plastid-targeted translocators, although it is possible that others in this
group may have lost this function. For example, the translocator from Iheileria annulata
apparently does not encode a N-terrninal extension, but is clearly related to the other
apicomplexan sequences. Although we do not have glaucophyte translocators in the tree,
the signature activity of plastidic phosphate translocators has been detected in isolated

cyanelles of Cyanophora paradoxa, indicating the presence of plastidic phosphate

39

translocators (Schlichting and Bothe, 1993). We therefore hypothesize that the insertion
of a phosphate translocator into the plasma membrane of the endosymbiont occurred
before the split of the Plantae and was probably a critical step in rendering the
association between the cyanobacterium and the mitochondriate eukaryote irreversible.
The proposed sequence of events for plastid establishment (Figure 1.2 and Figure 1.3)
does not involve major evolutionary innovations. The basic components of a plastid
envelope protein import apparatus were present in the cyanobacterium (Reumann et al.,
2005) and it is reasonable to assume that this machinery was capable of importing
proteins from the host cell cytosol, albeit with low efficiency; i.e., the orrter leaflet of the
plastid outer envelope membrane consists mainly of ER—derived phospholipids (Douce
and J oyard, 1981; Cavalier—Smith, 2000), indicating a close interaction between plastid
envelope membranes and the host endomembrane system. It is likely that the
cyanobacterium was engulfed as a prey item (as often occurs today) through
phagocytosis, an event that likely occurred countless times and in some of these cells, the
prey was not digested in the food vacuole but rather maintained as an endosymbiont. A
descendant of these cells gave rise to the Plantae. The origin (or replacement) of plastids
through cell (or organelle) engulfinent has occurred several times in evolution and can be
found in taxa such as the filose amoeba Paulinella chromatophora (Bhattacharya et al.,
1995) and in several dinoflagellate lineages (Schnepf and Elbrachter, 1999; Saldarriaga et
al., 2001; Hackett et al., 2004). The position of the Plantae in the tree of life (Baldauf,
2003; Rodriguez-Ezpeleta et al., 2005) suggests that its common ancestor was a highly
developed flagellate (or had flagellated stages) that most likely had the capacity for

phagocytosis and endomembrane formation (Cavalier-Smith, 2000).

40

Figure 1.1. Maximum-likelihood phylogeny of endomembrane and plastid translocators
ML bootstrap values (200 replications) are shown above the branches on the left of the
slash mark, whereas the values to the right are from a NJ analysis (100 replications).
Only bootstrap values 2 60% are shown. The different plastid translocators are GPT
(glucose-6-phosphate/phosphate translocator), PPT (phosphoenolpyruvate/phosphate
translocator), TPT (triose-phosphate/phosphate translocator), and XPT (xylulose-S-
phosphate translocator). Apicomplexans: Iheileria annulata, Plasmodium falciparum,
Toxoplasma gondii; HaptophyteszEmiliana huxleyi; Stramenopiles: Thallassiosira
pseudonana. The numbers in the circles indicate translocators to the right that resulted
from primary ( l) and secondary (2) endosymbiosis. T. pseudonana: Ihallassiosira

pseudonana; A. thaliana: Arabidopsis thaliana; C. merolae: Cymidioschyzon merolae

41

 

Theileria annulata
Plasmodiumfalciparwn NP_703428
Taxoplasma gondii

Plasmodium falciparum NP_703643
100 r—Emiliam'a huxleyi
L_Thalassiosira pseudonana Sc153
T. pseudonana Sc5

T. pseudonana Sc9
Thalassiosira pseudonana Sc15 / Sc44

Galdieria sulphuraria HET39C12
Physcomitrella patens 4787

Arabidopsis thaliana AAC83815

Zea mays P491 33

Galdieria sulphuraria A414H8

7 Physcomitrella patens 824

100 100 Arabidopsis thaliana AAL153 10 GPT
‘ Arabidopsis thaliana AAC28500
Arabidopsis thaliana K10A8 (XPT)
Arabidopsis thaliana AAF01540

Arabidopsis thaliana AAM91743

Chlamydomonas reinhardtii AA020101
Galdieria sulphuraria A416F5
Homo sapiens KIAA044
100 C. meralae CMS488C
72 Karena brevis C01596
Karena brevis C00075
A. thaliana AtGONST4

85 _lllQ C. merolae CMOSSC
64 Cyanidioschyzon merolae CMT467C
199 Phytophtora ramorum 8384
Galdieria sulphuraria A413D9
Phytophtora ramorum 7748
Homo sapiens BAB5530

‘_1m_l—Neurospora crassa CAB9145

Saccharomyces cerevisiae Sly4l

   
 
 
 
 

 

 

 

TPT

88

 

  
 
 
 
 
 

 

84

 

 

 

 

 

 

 

 

 

 

 

 

 

Figure 1.1. Maximum likelihood phylogeny of endomembrane and plastid translocators

42

Acquisition of a
cyanobacterium by a
mitochondriate protist.

Endomembrane-derived
translocator of fixed
carbon originates
(triose-phosphate, TPT).

Origin of plastid targeting
system and
ATP/ADP transporter
to supply plastid
with energy for e.g.,
fatty acid biosynthesis.

Organellar genome reduction
to the nucleus. Origin of
additional translocators;

e.g., dicarboxylate, hexose
through gene duplication
and divergence
by a mitochondriate protist.

Glaucophytes Red algae Radiation of the Plantae.

Green algae
Land plants

 

Figure 1.2. Model for plastid and translocator origin in the common ancestor of the

Plantae during primary endosymbiosis

CB: cyanobacterium; ATP: Adenine triphosphate; TPT: Tr'

translocator. The figure in this dissertation is presented in color

43

Acquisition of a red alga
by the mitochondriate
ancestor of the
chromalveolates

Transfer Of the red algal
plastidic translocator
(and other) genes(s) to the
chromalveolate nucleus.
Duplication and divergence
of the chromalveolates
translocator gene family

 

4-

Radiation of the
chromalveolates

Cryptophytes

    
   

Haptophytes Alveolates
Stramenopiles

Chromalveolates

Figure 1.3. Model for plastid and translocator origin in the common ancestor of the
chromalveolates during secondary endosymbiosis

RA is red alga. The chromalveolate tree reflects the present understanding of the
phylogeny of this group (e.g., Bhattacharya et al., 2004; Harper et al., 2005). The figure

in this dissertation is presented in color.

44

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Brown E.C., Somanchi A., and Mayfield SP. (2001) Interorganellar crosstalk: new
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Bullerwell C.E., and Gray MW. (2004) Evolution of the mitochondrial genome: protist
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Cavalier-Smith T. (2000) Membrane heredity and early chloroplast evolution. Trends
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Cavalier-Smith T. (1999) Principles of protein and lipid targeting in secondary
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Douce R., and Joyard J. (1981) Does the plastid envelope derive from the endoplasmic
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Douzery B.J., Snell E.A., Bapteste E., Delsuc F., and Philippe H. (2004) The timing of
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Fey V., Wagner R., Brautigam K., and Pfannschmidt T. (2005) Photosynthetic redox
control of nuclear gene expression. Journal of Experimental Botany 56:1491-1498.

Gray M.W., Lang B.F., and Burger G. (2004) Mitochondria ofprotists. Annual Review
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Hackett J.D., Yoon H.S., Soares M.B., Bonaldo M.F., Casavant T.L., Scheetz T.E.,

Nosenko T., and Bhattacharya D. (2004) Migration of the plastid genome to the
nucleus in a peridinin dinoflagellate. Current Biology 14:213-218.

45

Harper J.T., Waanders E., and Keeling PJ. (2005) On the monophyly of
chromalveolates using a six protein phylogeny of eukaryotes. International Journal of
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Knappe S., Fltigge U.l., and Fischer K. (2003) Analysis of the plastidic phosphate
translocator gene family in Arabidopsis and identification of new phosphate translocator-

homologous transporters, classified by their putative substrate-binding site. Plant
Physiology 131:1178-1190.

Lang M., Apt KE., and Kroth P.G. (1998) Protein transport into "complex" diatom
plastids utilizes two different targeting signals. Journal of Biological Chemistry
273:30973-30978.

Palmer J.D. (2003) The symbiotic birth and spread of plastids: How many times and
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Reumann S., Inoue K., and Keegstra K. (2005) Evolution of the general protein import
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Rodriguez-Ezpeleta, N., H. Brinkmann, Burey S.C., Roure B., Burger G.,
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Saldarriaga J.F., Taylor F.J., Keeling P.J., and Cavalier-Smith T. (2001)
Dinoflagellate nuclear SSU rRNA phylogeny suggests multiple plastid losses and
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Schlichting R., and Bothe H. (1993) The cyanelles (organelles of a low evolutionary
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Evolution 21:809-818.

46

CHAPTER 2

PHYLOGENY AND ORIGIN OF PLASTID METABOLITE TRANSPORT
PROTEINS FROM ARABIDOPSIS THALIANA 2

 

2 This work has been published in Tyra B.M., Linka M., Weber A.P.M., and Bhattacharya D.,
(2007) Host origin of plastid solute transporters in the first photosynthetic eukaryotes. Genome
Biology 8:R212-224. Taira HM. and I contributed equally to this work. Taira H.M. wrote the
initial draft of the manuscript. Dr. D. Bhattacharya was responsible for the final phylogenetic
trees.

47

ABSTRACT

Background: It is generally accepted that a single primary endosymbiosis in the Plantae
(red, green (including land plants), and glaucophyte algae) a common ancestor gave rise
to the ancestral photosynthetic organelle (plastid). Plastid establishment necessitated
many steps, including the transfer and activation of endosymbiont genes that were
relocated to the nuclear genome of the 'host' followed by import of the encoded proteins
into the organelle. These innovations are, however, highly complex and could not have
driven the initial formation of the endosymbiosis. We postulate that the retargeting of
existing host solute transporters to the plastid fore-runner was critical for the early
success of the primary endosymbiosis, allowing the host to harvest endosymbiont
primary production.

Results: We tested this model of transporter evolution by conducting a comprehensive
analysis of the plastid permeome in Arabidopsis thaliana. Of 137 well-annotated
transporter proteins that were initially considered, 83 that are broadly distributed in
Plantae were submitted to phylogenetic analysis. Consistent with our hypothesis, we find
that 58% of A. thaliana transporters, including all carbohydrate transporters, are of host
origin, whereas only 12% arose fiom the cyanobacterial endosymbiont. Four transporter
genes are derived from a Chlamydia-like source, suggesting that establishment of the
primary plastid likely involved contributions fi'om at least two prokaryotic sources.
Conclusion: Our results indicate that the existing plastid solute transport system shared
by Plantae is derived primarily from host genes. Important contributions also came from
the cyanobacterial endosymbiont and Chlamydia-like bacteria likely co-resident in the

first algae.

48

INTRODUCTION

Plastids in eukaryotes that contain chlorophyll are capable of carrying out photosynthesis,
a process that converts light energy, carbon dioxide, and water into organic compounds.
The evolutionary history of this organelle unfolded over a billion years ago when a
previously non-photosynthetic protist engulfed and maintained a fi'ee-living
cyanobacterium in its cytoplasm (Y oon et aL, 2004). It is hard to over-state the
importance of this ancient and extraordinarily rare primary endosymbiosis because
plastids allowed the evolution of algae and the plants that form the base of the food chain
for many ecosystems on Earth. Current data suggest that the primary endosymbiosis
occurred once in the common ancestor of the red, green (including land plants),
glauCOphyte algae, the Plantae (Rodriguez-Ezpelata et al., 2005; Weber et al., 2006;
Reyes-Prieto and Bhattacharya, 2007), with the Original plastid and the nuclear-encoded
machinery for running the organelle spreading in subsequent cell captures to other
branches of the eukaryotic tree (Cavalier-Smith et al., 1994; Mo Fadden, 1999;
Bhattacharya et al., 2004). The only other known case of a potential bonafide
cyanobacterial primary endosymbiosis occurred relatively recently in the thecate amoeba
Paulinella chromatophora (Marin et al., 2005; Yoon et al., 2006).

The gradualist view of evolution through mutation-selection suggests that it
would have taken millions of years for the captured prokaryote to become fully integrated
into the 'host' eukaryote, ultimately becoming the site not only for carbon fixation but
also for other complex functions, such as lipid, isoprenoid, and amino acid biosynthesis
(Weber et al., 2005). These processes were associated with the migration of much of the

cyanobacterial genome to the host nucleus'and development of the complex protein

49

import system that are key shared features among all canonical plastids (Timmis et al.,
2004; Weber et al., 2006; Gutensohn et al., 2006). A remarkable exception to the view
that endosymbiosis was a gradual process of integration is offered by the katablepharid
protist 'Hatena', which undergoes large-scale morphological changes following the
engulfment of a green alga (Okamato and Inouye, 2005).

Regardless of whether the ancient primary endosymbiosis fostered an accelerated
rate of morphological evolution in the Plantae ancestor or whether general cell
morphology was unchanged as in the Paulinella example (Johnson et al., 1919), one
thing is clear - in the absence of rapid benefits to the host it is unlikely that the
endosymbiosis would long have been sustained. Given the need for short-term survival, a
key feature of early success for the endosymbiosis must have been the integration of the
metabolism of the two cells. A solution for this process would have been solute
transporters that regulate the flux of metabolites (for example, ATP, phosphate, sugars
and sugar phosphates, metal ions, and other important ions) across the organelle
membranes. Controlled exchange in response to environmental factors such as changes in
light intensity and trace metal availability (Raven et al., 1999; Renne et al., 2003; Reiser
et al., 2004) is decisive because the unregulated flux of metabolites would have had
detrimental effects and, thereby, lowered the evolutionary fitness of the endosymbiosis. A
complex system of solute transporters is in place today in extant plastids that provides the
link between this organelle and the surrounding cytosol (F erro et al., 2003; Weber, 2004;
Weber and Fischer, 2007). Here we focus on the evolutionary history of these plastid

metabolite transporters to infer early events in plastid evolution.

50

We make two assumptions in this study. First, a system of metabolite transporters
was a critical and early development in plastid evolution to supply the endosymbiont with
essential nutrients and to enable the host to reap immediate benefit from photosynthetic
primary production. It is rmclear why the cyanobacterium that was destined to become
the plastid escaped digestion in the host but this scenario has also played out in 'Hatena'
and in Paulinella. Second, whereas the genome of the previously flee-living
cyanobacterium encoded all the transport systems required for the uptake of essential
inorganic nutrients, it most likely did not harbor genes encoding transporters for the
export of organic solutes to the host - this would have served no obvious pre-existing
purpose in the prokaryote. Precisely how the plastid solute transport system was
established is unknown. One possible model involves a primarily cyanobacterial origin,
in which the plastid continued to utilize its own original cyanobacterial solute
transporters with their evolution over time into proteins that perform most or all currently
known plastid permeome functions. An alternative model involves a host—driven solute
transport system, likely derived from the vacuolar envelope that initially surrounded the
endosymbiont after its engulfinent (Weber et al., 2006). And finally, both of the new
partners could have contributed proteins equally to this machinery, resulting in a chimeric
system composed of the most beneficial combination possible of prokaryotic and
eukaryotic transporters. To determine which of these competing hypotheses best explains
plastid transporter evolution, we undertook an initial bioinformatics analysis of 137
A. thaliana solute transporters and then a detailed phylogenetic analysis of a subset of 83
conserved proteins that included available data from other Plantae. The A. thaliana

transporters are either predicted or have been shown to be chloroplast targeted and are

51

ideal for tracking plastid permeome evolution. Using these data we demonstrate that over
one-half of Plantae plastid targeted transporters are putatively of host origin whereas less
than a quarter arose from the cyanobacterial endosymbiont. This suggests that primarily
host genes made the lasting contribution to the Plantae host-endosymbiont relationship
with regard to the plastid solute transport system. We also find evidence for the origin of
four transporter genes or gene families from a Chlamydia-like source. This latter result
raises the possibility that establishment of the ancient primary plastid may have involved
contributions from at least two prokaryotic sources, perhaps explaining its singular
nature. This hypothesis received substantial support from the recent finding of at least 21
genes of Chlamydia-like origin in the nuclear genome of the extremophilic red alga

Cyanidioschyzon merolae (Huang and Gogarten et al., 2007).

MATERIAL AND METHODS

Initial transporter analyses

As a starting point for the compilation of a conservative set of predicted or confirmed
plastid envelope membrane transporters, we used a previously published list of 137
plastid-targeted membrane proteins that was based on predicted plastid localization and
classification by the transporter classification system (Weber et al., 2005). This list was
manually curated to remove proteins from the list if published evidence indicated that
they were localized to a cellular location other than chloroplasts, if they represented
membrane-bound enzymes, or if they were annotated as components of the Tic/Toe

protein import apparatus, the photosynthetic machinery of the thylakoid membrane, or

52

the Sec or Tat protein targeting pathways. This curated list of candidate genes was
updated and amended with recently published chloroplast envelope membrane
transporters, such as AtFOLTl , a plastid localized transporter belonging to the
mitochondrial carrier family that does not contain a plastid targeting signal (Bedhomrne
et al., 2005) and was thus not included in previous lists. The final list contained 83 A.
thaliana predicted or confirmed chloroplast solute transporters.

The sequence for each protein was obtained fi'om The Arabidopsis Information Resorn'ce

website (11W). These protein sequences were used as
queries in the BLAST program “blastp” and “tblastn” searches of the NCBI Database

WWW), the plant and algal genomes available through
the Joint Genome Institute Eukaryotic Genomics Website (hmflgmejgi;

W), the Cyanidioschyzon merolae Genome Project website
W the Galdieria sulphuraria Genome Project website
(m://genomics.msu.edu/ga_ldieria_/),_and Dragonblast V2.1 (Ruemmele SE, unpublished
data), a web based database in Bhattacharya’s lab that contains EST datasets for several
chromalveolates, Plantae, excavates, Rhizaria, and Amoebozoa. We used the predicted
protein sequences for the following species for our analysis whenever available:
Arabidopsis thaliana, Oryza sativa, Physcomitrella patens, Chlamydomonas reinhardti i,
Ostreococcus tauri, Ostreococcus Iucimarinus, Cyanidioschyzon merolae, Galdieria
sulphuraria, cyanophora paradoxa, Dictyostelium discoideum, Strongylocentrotus
purpuratus, Xenopus Iaevis, Danio rerio, Mus musculus, Canisfamiliaris, and Homo
sapiens. In addition, we included at least one insect, three fungal species, and a broad

range of Bacteria and Archaea in our analysis. The BLAST searches used an E-value cut-

53

off < 0.00001. If a translated EST sequence was not available, the nucleotide sequence
was translated over Six frames using the EXPASY translate tool
(hm://ca.expa§y.grgtools/dna.htrnl). The resulting protein sequences were used in a
BLAST search against the NCBI protein database to ensure the correct translation was
obtained.

We used the ClustalW feature included with BioEdit V7.0.5.3 to generate protein
alignments (Hall, 1999). Alignments were visually inspected and manually corrected if
necessary. Trees were generated under maximum likelihood using PHYML V2.4.4
utilizing the WAG model of amino acid substitution and estimating both the proportion
of invariable sites and the alpha parameter (that is, WAG +1 + I') (Guindon and Gascuel,
2003). We performed non-parametric bootstrap analysis with 100 replicates for each
PHYML analysis. The resulting trees were analyzed to determine the origin of the
_ transporter in A. thaliana and other Plantae.

The designation of gene origin was done as follows. When the Plantae solute
transporter formed a well-supported (usually > 70% bootstrap support) monophyletic
group with homologs in opisthokonts (that is, animals and fungi) and secondarily with
other eukaryotes such as excavates and chromalveolates (if present), then it was classified
as having a 'Host' origin. Under this scheme, no bacterial sequences interrupted the
eukaryotic domain. 'Cyanobacterial' or 'Chlamydia-like' origin was inferred if the Plantae
sequence formed a monophyletic group with protein sequences finm either of these
lineages with strong bootstrap support. Other bacterial or eukaryotic sequences could (not
necessarily) be in these trees but there had to be a robust separation of the Plantae and

cyanobacteria or Chlamydia-like clade from all other homologs. We had two other

54

categories of gene origin that likely reflected a lack of phylogenetic resolution or
pervasive horizontal gene transfer (HGT) among taxa that defied a clear inference of
origin. The first was the 'Other' category in which the Plantae transporter formed a well-
supported monophyletic clade but its position relative to available bacterial data was
Imresolved, thereby not allowing us to identify the donor taxon. The second, 'PIantae—
specific', was for transporters that had no significant bits to sequences in GenBank or
other databases and appeared to be limited to the Plantae. The protein alignments are

available in the download section of the Bhattacharya Lab website

 

Detailed phylogenetic analyses

For eight representative transporters fiom the five categories described above we inferred
a maximum likelihood phylogeny using RAxML (RAxML-VI-HPC, v2.2.1) (Stamatakis
et al., 2005) and the WAG + F evolutionary model. The specific transporters were: 'Host'
- At4g39460 (SAMT, S-adenosylmethionine transporter, 236 amino acids), At5g663 80
(AtFOLTl, Arabidopsis thaliana folate transporter l, 268 amino acids); 'Cyanobacterial‘
- Atl g19800 (T GDl, trigalactosyldiacylglycerol l, permease-like protein for lipid
transport, 234 amino acids), At5g64940 (ABC transporter protein, 487 amino acids);
'Chlamydia-like' - Atlg15500 (plastidic ATP/ADP transporter 2 (AtNT'T 2), 436 amino
acids), At4g37270(HMA1, copper exporting ATPase, 444 amino acids); 'Other' -

Atl g32080 (putative membrane protein, 211 amino acids); 'Plantae-specific' -
At5g24690 (hypothetical expressed protein, 274 amino acids). These detailed analyses

used a random starting tree (one round of taxon addition) and the rapid hill-clirnbing

55

algorithm (that is, option -f d in RAxML). To generate bootstrap values for these
phylogenies, we used RAxML with the same settings and 100 replications. In addition,
we used Bayesian inference (MrBayes V3.0b4) (Huelsenbeck and Ronquist, 2001) with
each of the eight data sets using the WAG +1 + I‘ model to calculate posterior
probabilities for nodes in the RAxML trees. Metropolis-coupled Markov chain Monte
Carlo from a random starting tree was used in this analysis with two independent runs
and 1 cold and 3 heated chains. The Bayesian analyses were run for two million
generations each with trees sampled every 100th generation. To increase the probability
of chain convergence, we sampled trees after the standard deviation values of the two
runs were < 0.01 to calculate the posterior probabilities. We also ran the Bayesian
analysis for the remaining two putative 'Chlamydia-like' genes in Plantae (dicarboxylate
translocators DiTl, DiT2.1, and DiT2.2, and the phosphate transporter PHT2;1) to assess
the topologies. We incorporated a representative diversity of available sequences in all of

these trees.

RESULTS AND DISCUSSION

Distribution of transporters within Plantae

Phylogenetic analysis of the best-annotated transporter data that are currently available
from Arabidopsis was used to identify and putatively annotate homologs fiom other
Plantae. Of 137 transporter proteins that were initially considered, BLAST and
phylogenetic analyses and manual curation of recently available data led to the

identification of 83 proteins that were of sufficient conservation and broad distribution

56

among Plantae to be used for firrther analyses. Each of these 83 proteins that included
gene families (that is, representing 63 distinct, ancestral genes; Table 2.1) was used as
input in BLAST and PHYML bootstrap analyses to infer the trees. This approach
identified 41 proteins that are present in both red and green algae (including land plants)
and, therefore, were likely found in the Plantae ancestor (glaucophyte homologs were
found for some of these genes; for example, ADP/ATP translocase, hypothetical protein
At3g45890). Eleven proteins were restricted to green algae and land plants, seven were
plant-specific, and two were limited to red algae and land plants. The distribution of these
proteins with respect to their putative origin in Plantae is shown in Figure 2.1(A). Given
the lack of evidence for widespread horizontal gene transfer in extant Plantae, which
most likely lost the capacity for phagotrophy early in its evolution (Archibald et al., 2003;
Reyes-Prieto et al., 2007), we postulate that the patchy distribution for many plastid
targeted transporters primarily reflects differential gene loss over the greater than one
billion years that has passed since the primary endosymbiosis (Y oon et al., 2004). Under
this interpretation, the large set of shared transporters among Plantae lineages provides

resounding support for the monophyly of this supergroup (Hackett et al., 2007).

57

Table 2.1. List of Arabidopsis thaliana chloroplast solute transporters analyzed in this
study and their putative evolutionary origins

 

Arabidopsis thaliana solute transporters

 

Putative evolutionary origin: Host

AGI code

Atl g05580
At1g54320
Atl g59870
Atl g61 800
Atl g64150
At1g66950
Atl g70610
Atl g79450
At2g04620
At2g13 100
At2g27810
At2g28070
At2g29650
At2g3 8060
At2g38330
At2g40420
At3g01550
At3g12740
At3g17690
At3g17700
At3 g45 890
AB g523 10
At4g00370
At4g l 3590
At4g 1 7 340
At4g25 750
At4g32400
At4g32650
At4g383 80
At4g39460

Gene Annotation

Cation/hydrogen exchanger

Ligand-effect modulator 3 (LEM3) family

ABC transporter

Glucose-6-phosphate/phosphate translocator 2 (GPT2)
Expressed protein

ABC transporter

Transporter associated with antigen processing protein 1 (AtTAPl)
Ligand-effect modulator 3 (LEM3) family

Cation efilux family protein

Glycerol-3-phosphate transporter

Xanthine/uracil perrnease

ABC transporter

Na+-dependent inorganic phosphate cotransporter
Na+-dependent inorganic phosphate cotransporter
Multi antimicrobial extrusion (MATE Efflux) protein
Amino acid transporter
Phosphoenolpyruvate/phosphate translocator 2 (PPT2)
Ligand-effect modulator 3 LEM3 family

Cyclic nucleotide-binding transporter 2

Cyclic nucleotide-binding transporter l

Expressed protein

ABC transporter

Anion transporter 2 (ANTR2)

Expressed protein

Major intrinsic family protein

ABC transporter

Adenine nucleotide uniporter

Arabidopsis thaliana K+ rectifying channel 1 (ATKCl)
Multi antimicrobial extrusion (MATE Efflux) protein
S-adenosylmethionine carrier 2 (SAMT)

 

58

Table 2.1. (cont’d).

 

Arabidopsis thaliana solute transporters

 

Putative evolutionary origin: Host

AGI code

At5g04770
At5g05630
At5g13550
At5g14040
At5gl6150
At5gl7630
At5g19410
At5g19600
At5g22830
At5g26820
At5g33320
At5g42130
At5g45450
At5g46l 10
At5g52860
At5g54800
At5g59250
At5g66380

Gene Annotation

Amino acid permease

Amino acid permease

Sulfate transporter

Mitochondrial phosphate transporter

Hexose transporter

Xyluose-S-phosphate/phosphate transporter l (XPT)
ABC transporter (White)

Sulfate transporter

CorA-like magnesium transporter

F erroportin-related protein
Phosphoenolpyruvate/phosphate translocator (PPTl)
Mitochondrial substrate carrier family

Iron transporter-related

Triose phosphate/phosphate translocator (TPT)
ABC transporter (White)
Glucose-6-phosphate/phosphate transporter 1 (GPTl)

Sugar transporter
Folate transporter l (AtFOLTl)

Putative evolutionary origin: Cyanobacteria

AGI code

Atl g045 70
Atl g08640
At1g19800
Atl g78620
At2g32040
At3g51 140
At3g60590
At4g33520
At5g12470
At5g64940

Gene Annotation

Integral membrane family protein

Expressed protein

Trigalactosyldiacylglycerol 1, permease-like protein (TGDl)
Integral membrane family protein

Folate monoglutamate transporter, FT

Expressed protein

Expressed protein

Metal-transporting P-type ATPase (PAAl)

Expressed protein

ABC] -family protein

 

59

Table 2.1. (cont’d).

 

Arabidopsis thaliana solute transporters

 

Putative evolutionary origin: 'ChIamydia-like'

AGI code

At1g15500
Atlg80300
At3g26570
At4g37270
At5g12860
At5g64280
At5g64290

Gene Annotation

Adenine nucleotide translocase 2 (AtNTT 2)
Adenine nucleotide translocase 1 (AtNTT 1)
Low affinity phosphate transporter (PHT2;1)
Heavy metal ATPase HMAl

Dicarboxylate translocator 1 (DiTl)
Dicarboxylate translocator 2.2 (DiT2.2)
Dicarboxylate translocator 2.1 (DiT2. l)

Putative evolutionary origin: ‘Other’

AGI code

Atlg01790
Atlg32080
Atlg44920
At1g54350
At1g78560
At2g02590
At2g21340
At2g26900
At3g25410
At4g30580
At5g03555
At5gl3720
At5g52540
At5g62720

Gene Annotation

Potassium transporter

Membrane protein

Expressed protein

ABC transporter

Bile acid:sodium symporter

Expressed protein (Putative small multi-drug export)
Enhanced disease susceptibility protein

Bile acid:sodium symporter

Bile acid:sodium symporter

l-Acylglycerol-3 -phosphate O-acyltransferase (ATS2)
Cytosine/purines, uracil, thiamine, allantoin family permease
Expressed protein

Expressed protein

Integral membrane HPP family protein

Putative evolutionary origin: 'Plantae-specific'

AGI code

At2g38550
At3g57280
At5gl7520
At5g24690

Gene Annotation

Expressed protein
Expressed protein
Maltose transporter (MEXl)
Expressed protein

60

 

 

30
25 I Host
I Chlam dia—like
20 I Cyano acteria
I Other + Plantae-specific

 

 

Number of Genes
6i

 

 

 

0
Red and Green Green Red and Plant Plantae-specific

 

B 12%
8%
5%
I Host
17% El Chlamydia-like
1 5% I Cyanobacterial
7% I Plantae-specific
D Other
50% 7%

      

20%

Figure 2.]. Origin of plastid targeted solute transporters in Plantae

(A) Gene distribution among Plantae and gene origin for 63 distinct transporters
considered in this study. (B) Summary pie charts showing the origin of all the 83
transporters (top chart) and the 63 distinct genes (lower chart) considered in this study.

The figure in this dissertation is presented in color.

Most proteins of the plastid envelope permeome are host-derived

Analysis of the phylogenetic data supports the notion that the host drove the integration
of plastid and host metabolism. We find that the majority (58%, when considering all 83
genes; Figure 21(3) and Table 2. l) of the plastid solute transporters were most likely
derived from existing host membrane proteins. These 48 proteins are diverse in nature,
including several ATP binding cassette (ABC) transporters, nucleotide and amino acid
permeases, sulfate, potassium, magnesium, and iron transporters, and cation efflux
proteins (see Figure 2.2 (A) for S-adenosylmethionine transporter (SAMT) and Figure
2.2 (B) for A. thaliana folate transporter 1 (AtFOLTl) as representative trees).

Of particular interest is the finding that in addition to the members of the
nucleotide-sugar/triose phosphate translocator gene family previously reported to be of
host origin (Weber et al., 2006), all other carbohydrate transporters included in our
analysis were derived from existing host proteins. This result strongly suggests that the
host utilized existing eukaryotic transport proteins pre-adapted to this function to 'tap'
into the photosynthates produced by the captured cyanobacterium. In addition, the
Plantae host also provided transporters to facilitate the movement of valuable nutrients
such as magnesium, potassium, iron, and phosphate into the captured prokaryote. The
replacement of pro-existing cyanobacterial anion and cation transporters with host
derived proteins again suggests that there was strong selection to rapidly establish control
over and utilize the endosymbiont. This process was most likely accomplished by using

transporters derived fi'om the host vacuolar envelope (Weber et al., 2006).

62

Figure 2.2. Plastid targeted solute transporters of putative host’ origin in Plantae

These are RAxML trees with the nrunbers above the branches inferred fi'om a RAxML
bootstrap analysis and the thick branches showing significant (P > 0.95) support fi'om a
Bayesian phylogenetic inference. Only bootstrap values 2 60% are shown. Branch
lengths are proportional to the number of substitutions per site (see scale bars). The filled
magenta circle shows the node that unites the Plantae taxa within the eukaryotic domain.
The different algal groups are shown in different text colors: red for red algae, green for
green algae and land plants, and brown for chromalveolates. The inclusion of
chromalveolates within the Plantae is believed to reflect horizontal or endosymbiotic
gene transfer events (for example, Li et al., 2006). The two transporters are: (A) SAMT,
S-adenosylmethionine carrier transporter; and (B) AtFOLTl, A. thaliana folate
transporter 1. The name of the A. thaliana solute transporter used for the query is
indicated for both trees shown in this figure. The figure in this dissertation is presented in

color.

63

Danio rerio
Xenopus laevis
Mus musculus
Homo sapiens .
Strongylocentrotus purpuratus Oplsthokonts
Drosophila melanogaster

C oprinopsis cinerea

87 Schizosaccharomyces pombe
Edmowia lipolytica

Candida albicans

98 Piraeodacty/um tricornutum
T/ralassiosira pseudonana

.-'l lexrmdrium tamarense
98 PhyIO/J/rt/zuru ramorum

Plrr'mphllzora sofas
' 100 Cyanidimchvzon mero/ue
Galdieria sulphuraria
88 Plasmodimn fir/Cipurum
_'1 Theileria (mu/ma
C h/amr'dmmmas rein/IuI'c/Iii
()strcuc'occus quri
Ostreat/occur lucimurimrs
P/II'xcomiIrc/lu pawns
()rr':a saliva
Papa/us Irit'lzoc'w‘pa
A-"icn/iunu hen/lumziuna
Capsicum ummum
Arabidopsis thaliana At4g39460

 
  
   
  

 

 

 

      
   
 
  

 

 

 
  
 
  

 

98

— 0.1 substitutions/Site

 

B Milago maydis
Cryptococcus neoformans
_ 97 Aedes aegypti
100 Tribolium castaneum
r—Strongylocentrotus purpuratus .
Xenopus laevis OplSthOKOIltS
100 Gallus gallus

 

Danio rerio

T etraodon nigroviridis
100 Homo sapiens

Mus musculus

Cyanidirrs‘c/ryzr)n INC/‘0](lc’
Phaeou'actr'lum Iricormrmm
P/zvtop/rt/rora rumarum
(71/chI'dmmmas reinlzm'dlii
Oslrem'nt't'us quri
()sIrcuc‘oc't'us lrlt'imurinus
Papa/us Ir'ir'lmt'urpu
Arabidopsis t/Iu/iuml r‘\15}.16038()
()rr':u saliva

 

   
  
   

 

 

 

100 100

—— 0.1 substitutions/site

Figure 2.2. Plastid targeted solute transporters of putative 'host' origin in Plantae

64

The cyanobacterial contribution

The cyanobacterial endosymbiont putatively contributed ten solute transporters to the
plastid transport system (Table 2.1). These proteins include permease-like protein TGDl
(trigalactosyldiacylglycerol 1, Figure 2.3), which is required for integrating the
prokaryotic (that is, cyanobacterial) with the eukaryotic (that is, endoplasmic reticulum)
pathway for lipid biosynthesis (Xu et al., 2003; Awai et al., 2006; Benning et al., 2006),
the metal-transporting P-type ATPase PAAl (Shikanai et al., Abdel-Ghany et al., 2005),
and a transporter required for folate/biopterin biosynthesis (Klaus et al., 2006). The
remaining seven proteins of unknown function that are localized to the chloroplast inner
membrane were included in the cyanobacterial group. Whereas the predicted secondary
structure of most of these proteins indicates they represent transporters (that is, they
contain at least four transmembrane domains that are connected by short loops), some,
such as the ABCl-farnily protein At5g64940 (F igure 2.3) contain only one or two
predicted transmembrane domains and may thus have functions other than metabolite
transport. It is also intriguing that with the exception of the PAAl copper transporter the
only cyanobacterial transport proteins apparently retained by Arabidopsis are those for
which the host lacked a suitable replacement. For example, the initial steps of folic acid
biosynthesis in plants are confined to the chloroplast; the final steps are localized in the
cytosol and in mitochondria (Sahr etal., 2006; Basset et al., 2004a; Basset et al., 2004b).
Plastids thus depend on an external folate supply and require an uptake system for this
important metabolite. Interestingly, redundant systems for folate uptake exist in
Arabidopsis chloroplasts, consisting of the cyanobacterial-derived folate transporter FT

(Klaus et al., 2005) and the host-derived transporter AtFOLTl (Bedhomrne et al., 2005).

65

Figure 2.3. Plastid targeted solute transporters of putative 'cyanobacterial'l'plastid
endosymbiont' origin in Plantae

For details of tree building see Figure 2.2. The filled magenta circle shows the node that
unites the Plantae taxa as sister to cyanobacteria. The different photosynthetic groups are
shown in different text colors: blue for cyanobacteria, red for red algae, green for green
algae and land plants, and brown for chromalveolates. The inclusion of chromalveolates
or Euglenozoa (Eugl.) within the Plantae is believed to reflect horizontal or
endosymbiotic gene transfer events (for example, Li et al., 2006). The two transporters
are: (A) TGDl , trigalactosyldiacylglycerol l, lipid transporter; and (B) ABCl-farnily
transporter protein The name of the A. thaliana solute transporter used for the query is
indicated for both trees shown in this figure. The figure in this dissertation is presented in

color.

66

 

T hermosinus carboxydivorans Norl
* Solibacter usitatus Ellin6076
Mari rtzfimdus ferrooxydans PV-l
ic ettsia bellii RML369-C

 

      
   
 
  

100

Rickettsia prowazekii str. Madrid E Non-
Parvibaculum lavamentivorans DS-l Cyano-
Acidi hilium crv tum JF-S bacteria

Iasto ire/ ula marina DSM 3645
Syntrophus acidilrop icus SB
Wolinella succinogenes DSM 1740
Myxococcus xanthus DK 1622
Thermos}'nec'llacocc'us e/(mgatus BP—l
_{——— Sl'llé’t'lhu:l‘SliS sp. PC'C 6803 (2)
Naxa‘wpza'zc'Ii/armu PC‘C 73 l 02 (2)
— Crocmplzacra walsanii WH 8501
S)'nec-lmuu'cus elongatus PCC 6801
Prat/z/arar'ac'r'u.\' marinas 511'. 921 l
Si'HUL‘IIUCUL'L'IIA' Sp. \‘Vll 57Ul
Nos/ac- Hindi/army PCC 73 l ()2 ( l )
Nada/aria spamigcna CC Y9—1 I 4
.V'asmt' sp. P('(‘ 7120
Ana/menu variabi/ix' ATCC 294 l 3
Tric/zm{osmium uni/Human N S l 0|
Lyng/n‘a sp. P('(' 8 I 06
G/(M'U/‘lclt'ft’l' via/accus PCC 743|
Sl'llt’L'/2(I(.’()('(‘ll.\' Sp. JA-3-3 Ab
Sl'm’c'llm'm't'ux S . JA-2-3B‘a(2- l 3)
.Sl‘fl(’(‘/l()(:l'.\‘fi.9 Sp. PC‘C 68 )3 ( l )
83

 

 

 

 

 

100

 

  
 
     
  

 

    

 

 

 

 

Ci'anidiasC/ntan mcrala
Grad/aria Iwzuislipilala var. liui
Porphyra purpurca

Parp/rvra _1*e:()ensis

O.s'lrca(-au'u.s lat'imarimzs
()xlrcacm'(‘Im‘ {am'i
PratoI/wca u'ic'lx'cr/zamii
Cit/am}'damanas I't'in/zardlii
Ph}'sc'amilrclla palms

()rym saliva

Pa m/us Iridmcarapa

A cc/ic'aga a'mzc'aIu/a

Arabidopsis thaliana AI l g [9800

        
    
  
 
 
 
  
  
 

 

100

 

Nuclear

79
— 0.1 substitutions/site

Figure 2.3. Plastid targeted solute transporters of putative 'cyanobacterial'l'plastid

endosymbiont' origin in Plantae

Figure 2.3 continues on the following page.

67

Sl‘llcé“/2()(‘()('L‘ll.8' (”la/lgalus PCC630|

 
 
 
 
 
 
 
 
  
 

lerznast'nec‘lzac'aa'us' elanglus BP-l
Trir'lzmlcsmium (’l'l‘l/ll‘rlt’lllll [MS 1 0|
Ana/mwza \‘al'ia/vilis ATCC 29413
Nr‘mlm' Sp. PCC 7 120
Noslac' pzmcli/armc PCC 7310?.
Mala/aria .s‘pzmzigcna (CY 94 I 4
Cracmp/zaura l'l'c'lIS'Olll-l' WH 850i
Sl'llct'lllu‘lNUS 5]). PC‘C 6303
Glue/ulc‘lw' Via/accux PCC 7421
Sl'llc’t‘llrlt'lu‘t‘l(.\' Sp. .1 A-JBB‘aQ-U)
Lyngbm 3;). PCC 8106

95
fi 100 I Galdieria sulplna'al'ia (I)
Galdieria .S'lllp/llll'ul‘ic’l (2)

.8_5‘ 7 Cyanidioscllyzon memlae (I)

 

 

 

 

 
 
  
 
  
 
  
 
  

Cyanidi().s'('/1_1.‘:an mam/ac) (2)

 

Porphyrc'l I)’L’:()c'l7.8‘i5

Plzaeoa’acl)‘lum lricormllum
T l?c7/d.§'.$'i05il‘('l pseua’mzana
Euglena gracil is

 

 

Chlam_\‘d(mmnas rcin/zardlii
100 Oslrc'ac'ac'cuslac'imarinus
()slreac'ac'c'as lam'i
Pl?):s‘comilrc/la palms

Papa/us Il‘it'llUCtll'pa

Arabidopsis l/zaliana A15g649—10
Orin: saliva

— 0.1 substitutions/site

Figure 2.3. (cont’d).

68

'Chlamydia-like' transporters

In addition to the host and cyanobacteria, a third significant contributor to the Plantae
plastid solute transport system is the Chlamydiae. A surprisingly high number (four) of
plastid envelope membrane transporters have been contributed by these prokaryotes. The
presence of plant-like genes in Chlamydia has been noted in the past, sparking debate
over whether their presence indicated a transfer fiom the ancestral plant to Chlamydia, an
evolutionary relationship between cyanobacteria and Chlamydia, or a horizontal gene
transfer (HGT) from a chlamydial parasite to the plant ancestor (Brinkman et al., 2002;
Wolf et al., 1999; Schmitz-Esser et al., 2004). Phylogenetic analysis of plastid,
Chlamydiae, and Rickettsiae ADP/ATP translocases (Schmitz-Esser et al., 2004) supports
an ancient Chlamydia-to-Plantae direction of transfer. This explanation for the origin of
the ADP/ATP translocase gene (and other ChlamydiaI-like genes) in Plantae was
strongly supported by the phylogenomic analysis of Huang and Gogarten (2007). We
found a monophyletic relationship between the AtNTTl and AtNTT2 (the A. thaliana
plastid ADP/ATP translocases l and 2) and Chlamydiae ATP/ADP translocases (Figure
2.4) (Greub and Raoult, 2003; Linka et aL, 2003). In addition, the copper transporter
heavy metal ATPase 1 (HMAl; Figure 2.4), the dicarboxylate translocators (DiT) DiTl,
DiT2. l, and DiT2.2 (data not shown), and the low affinity phosphate transporter PHT2;]
(data not shown) apparently has a chlamydial origin in Plantae (Huang and Gogarten,
2007). All of these trees provide bootstrap (except for the DiT tree) support for the
monophyly of the 'ChIamydia-like' and plastid transporters. In the case of HMAl there
are two ancient paralogs in plants, one of cyanobacterial likely endosymbiotic origin and

one from a Chlamydia-like source that is shared with red and green algae. The DiTs

69

(Weber and Flugge, 2002) are present only in green algae, plants, and bacteria (that is,
not in red algae). Whereas genomic data for glaucophytes are not yet available, transport
experiments using isolated Cyanophora cyanelles showed that this glaucophyte uses a
transport system for glutamine and 2-oxog1utarate that is distinct from green plant DiTs
(Kloos et al., 1993). Taken together, these data indicate that 'Chlamydia-like'
dicarboxylate translocators have likely been lost from red algae and glaucophytes. An
alternative explanation is that the gene was acquired by the green lineage after the split of
Chlorophyta and Rhodophyta. A DiT2 gene was also found in the dinoflagellates
Amphidim'um carterae and Heterocapsa triquetra, which likely originated from an
independent horizontal gene transfer (HGT). Several 'green' genes have been found in
dinoflagellates and other chromalveolates that could have either originated from multiple
independent HGTs or an ancient green algal endosymbiosis (for discussion, see Nosenko
et al., 2006).

In summary, it is surprising that bacteria not putatively involved in the
endosymbiosis contributed 8% of the transporters that we have identified. When one
considers the functions of these transporters, the chlamydial contribution becomes more
important. HMAl increases copper and/or zinc transport into the plastid under conditions
of high light, facilitating the production of copper/zinc superoxide dismutase
(CuZnSOD), which protects the plant from superoxide radicals produced under high light
conditions (Seigneurin-Berny et al., 2006, Lee et al., 2007). PHT2;], a phosphate
transporter, controls phosphate allocation under conditions of phosphate-starvation
(Versaw and Harrison, 2002). The DiT transporters are involved in assimilating nitrogen

and recovering carbon lost to photorespiration, a process that is initiated by the

70

oxygenation reaction of RubisCO that primarily occurs under conditions when a high
022C02 ratio is present in the vicinity of RubisCO. Mutants lacking these transporters are
tmable to survive in ambient C02 concentrations (Renne et al., 2003; Reumann and
Weber, 2006; Schneidereit et al., 2006). Finally, the AtNTTl and AtNTT2 transporters
are required for ATP import into the plastid during the dark (that is, in the absence of
photosynthetic ATP production), particularly during lipid and chlorophyll biosynthesis.
Although AtNTT2 mutants are still capable of producing lipids, indicating that the plastid
has an alternative method for generating the ATP required for lipid biosynthesis, the
production is significantly reduced and mutant plants have a sharply reduced growth rate
Reiser et al., 2004). Arabidopsis mutants deficient in both AtNTTl and AtNTT 2 develop
necrotic lesions when grown under short days, accumulate H202, and, strikingly, show
constitutive expression of CuZnSOD2 and ascorbate peroxidase (Reinhold et al., 2007).
The phenotype of the mutant was linked to reduced magnesium chelatase activity and it
was concluded that ATP import into plastids in the dark is required for chlorophyll
biosynthesis and for preventing photooxidative damage (Reinhold et al., 2007). The
import of ATP into plastids in the dark is thus clearly a case in which the endosymbiont
benefits from host metabolism. The ancient origin of these transporters in the tree of
photosynthetic eukaryotes (Figure 2.4) is indicative of an essential role of this uptake
system in the formation of the endosymbiosis. With the exception of the DiT
translocators, each of these transporters appear to perform somewhat redundant functions
(that is, copper and phosphate transport) but in a way that permits the plant to adapt to

stresses involved in life on the land (that is, high light and 02 levels or low phosphate

71

availability). This may explain why the genes encoding these four plastid transporters
have been retained in the Arabidopsis genome.

How the 'ChIamydia-like' genes entered into the Plantae ancestor is unclear but it is
possible that both the cyanobacterial endosymbiont and chlamydial parasites may have
co-existed in the cell. Many environmental Chlamydia are known today that are broadly
distributed in animals and protists (Molmeret et al., 2005). The co-existence of these two
distinct prokaryotes may have provided the genetic 'toolkit' to make permanent the
endosymbiosis with gene transfer from each cell providing essential functions for
endosymbiont utilization. An alternative explanation is that the cyanobacterial
endosymbiont was itself highly chimeric (that is, the 'fluid chromosome model') (Embley
and Martin, 2006) and contained genes of chlamydial origin that had been gathered
through HGT. Although possible, this scenario seems less plausible because it invokes,
for example, the presence of an ATP/ADP translocator (a gene typical for 'energy
parasites' such as Rickettsiae) in the genome of an oxygenic photosynthetic cell that is
unlikely to encounter high concentrations of ATP in the surrounding environment; To
this end, a 'Chlamydia-like' ATP/ADP translocator is absent from all studied

cyanobacteria.

72

Figure 2.4. Plastid targeted solute transporters of putative 'Chlamydia-like’ origin in
Plantae

For details of tree building see Figure 2.2. The filled magenta circle shows the node that
unites chlamydial taxa with plastid targeted Plantae transporters. The different
photosynthetic groups are shown in different text colors: blue for cyanobacteria, red for
red algae, green for green algae and land plants, magenta for glaucophytes, and brown for
chromalveolates. The inclusion of chromalveolates within the Plantae is believed to
reflect horizontal or endosymbiotic gene transfer events (for example, Li et al., 2006).
The two transporters are: (A) ATP/ADP translocator (NTT); and (B) Heavy metal
ATPase (HMAl) copper transporter. The name of the A. thaliana solute transporter used
for the query is indicated for both trees shown in this figure. The figure in this

dissertation is presented in color.

73

A 100 Rickettsia prowazekii

Rickellsia bellii RML363-C

Rickeltsia prowazekii C AA 1 4826
Rickellsia sibirica 246

Caedibacler caryophilus

100 Rickettsia belli RML369-C

Ricketlsia montanensis

82 Lawsom'a inlracellularis PHE/MNl -00

Candidalus Paracaedibacter symbiosis
Candidalus Protochlamydia amoephila

78 Candidalus Protochlamydia amoephila UWE25

93 Neochlamydia hartmannellae
100 Parachlamydia sp. Hall’s coccus
100 Chlamydia lrachomalis D/UW-3/CX
Chlamydophila pneumoniae 1138
Glaucoc'vslis noslochincarum
96 100 Fragilariapsis (iv/indl'us‘
97 T/ta/assiasil'a pseuc/(mana
Plzamclac'lt‘lum ll'lt‘Ol'llllllHN
Parp/n‘ra t'ciacnsis
8. Galdieria su/p/mraria
Ct’anic/imc‘lzt'mn mam/ac
Pralalheca wicker/lama
98 66 Dlllla/ic/la s‘a/illa
C/I/cllllt‘a’amanas rcin/zardlil'
Torin/a rural/'5‘
93 Plans law/a
84 Onta saliva
Salalmm lulwraxum
" :‘ Cilrus Irt'hrid cullit‘ar
.~ll'(ll)l(.l()p.\‘l.\' l/Ia/iana Al I g80300
lllawn/72')‘anllzcmum ('lft‘xlal/inmn

  
 
 
    

 

 

 

 

 

100

 

— 0.1 substitutions/site

Figure 2.4. Plastid targeted solute transporters of putative 'Chlamydia-like' origin in

Plantae

Figure 2.4 continues on the following page.

74

 
   
  

lauri ..
remhardm
( 2)

Host

    
 
 
 
 
 
  
 
 
   
 
 
 
  
    
 
 
 
 

ramorum.
hansevm

purpuratus

 

JA-3-3 Ab

walsonii
1

CC93Il

Cyanobacterial

reinhardlii

 

amoebophila

, sp. PCC 8l06
Chlamydia pneumoniae
trachomatis

. . . At4g37270
— 0.1 substrtutrons/srte
merolae

Figure 2.4. (cont’d).

75

'Other' and 'Plantae—specific' transporters

We were unable to conclusively determine the origin of 18 transport proteins. Fourteen of
these data sets resulted in PHYML trees in which the Plantae transporters were rooted
within prokaryotes but without bootstrap support for a specific afi'rliation. An excellent
example is provided by At] g32080 (Figure 2.5), which is a putative membrane protein
conserved among Plantae, chromalveolates, and a diverse set of Eubacteria and Archaea
(that is, the Thermococcus and Pyrococcus clade). Although the prokaryotic source of
this gene in Plantae is unclear with the available data, the eukaryotic clade is clearly
monophyletic, which is consistent with a single gene origin in the Plantae ancestor and,
thereafter, transfer to chromalveolates (for example, diatoms in this tree) via secondary
endosymbiotic gene transfer (Li et al., 2006). The unresolved provenance of At1g32080
and the 'Other' set of transporters in Plantae can be explained by pervasive HGT
followed by full or partial gene replacement or differential gene loss among prokaryotes
that has erased the ancient phylogenetic signal. Alternatively, these results may indicate
erratic rates of sequence divergence that make it impossible to model protein evolution
for these sequences. Given the growing evidence, however, for recurring HGT among
bacteria (Dagan and Martin, 2007), it is likely that genes in the 'Other' category have
reticulate evolutionary histories. In this regard it is noteworthy that the likely frequent
HGTs seen in Figure 2.5 (A) among prokaryotes and other genes in this category
contrasts starkly with the apparent single origin and vertical inheritance in Plantae. This
result suggests a clear difference in rates of HGT for these genes with elevated rates in

prokaryotes relative to eukaryotes.

76

Of the remaining transporters, four fell in the 'Plantae-specific' category because
they lacked identifiable homologs outside of this supergroup and may simply be too
divergent to determine their origin. This includes At5g24690 (Figure 2.5 (B), a
hypothetical expressed protein) and the plastidic maltose exporter l (MEXl; data not
shown). The latter is required for export of maltose resulting from starch breakdown from
plastids at night in green plants. Storage of starch inside the chloroplast is exclusively
found in the green linage. Therefore, MEXl has likely co-evolved with plastid-based
starch biosynthesis and breakdown since it can be detected only in members of the
Viridiplantae with one gene found in the dinoflagellate Karloa'inium micrum, which, as

described above for DiT2, likely has resulted from a HGT.

77

Figure 2.5. Plastid targeted solute transporters of 'Other' or Plantae-specific' origin in
Plantae

For details of tree building see Figure 2.2. The filled magenta circle shows the node that
unites the Plantae taxa. The different algal groups are shown in different text colors: red
for red algae, green for green algae and land plants, magenta for glaucophytes, and brown
for chromalveolates. The inclusion of chromalveolates within the Plantae is believed to
reflect horizontal or endosymbiotic gene transfer events (Li et al., 2006). (A) A. thaliana
solute transporter in the 'Other' category: putative membrane protein (Atl g32080) (B) A.
thaliana solute transporter in the 'PIantae-specific' category: hypothetical expressed

protein (At5g24690). The figure in this dissertation is presented in color.

78

Del lia acidovorans SPH-l
Bur holderia multivorans ATCC176 l 6
Ralstom'a eutropha H16
77 Ralstonia eulropha JMP134
Ralslonia pickettli l2]
Ralstonia solanacearum UW551
Polynucleobacler if. gLW-Pl DMWA-l
Dechloromonas aromatica C
Paracoccus denitrificans PD 1 222
Chloroflexus aglgregans DSM 9485
_ Roseiflexus sp. RS-
100 Geobacler bemidjiensis Bem
Geobacler metallireducens GS-15
100 T hermococcus kodakarensis KODl
'—EE Pyrococcus horikoshii 0T3
Pyrococcusfuriosus DSM 3638
Desulfitobaclerium hafm'ense Y5]
100 r— Pseudomonas pulida GB-l
l-Pseudomonas enlomophila L48
Clostridium cellulol licum H10
Enlerococcus faeca is V583

   
    
 
  
 

 

 

 

   
  
 

 

 

 

__ Sfiphylococcus haemolyticus JC SC 1435
100 aemophilus in uenzae Rd KW20
-—-1_‘:- Vibriofischeri ES] 1
Photobaclerium s . SKA34

G aa’iel'ia su/ 7hza'aria
Ct'anidiosr' titan mero/ac
Pllc‘lé’0(l'a('l_l‘lllm

T/m/assiasil‘a
(ll/am}'danmnas rein/zardlii
()slrcac'm'("us lat'imarinus
O.s'll'ca('()('c'zl.s' lam'i
Pit)ascomilrc/la palms
98 Papa/us ll'lt'llUt'tl/‘(lpa
Ol't'za .s'alit‘al I)
()ljt'3a sail/val?)
LAl'ahidapxis llza/iana At 1 g32080

 

 
  
 

 

100

 

100

    
 
 
 
 

 

— 0.1 substitutions/site

Figure 2.5. Plastid targeted solute transporters of 'Other' or 'PIantae-specific' origin in
Plantae

Figure 2.5 continues on the following page.

79

 

C t‘anap/mra paradm'a

k Ct'alzidimc/irtan mam/ac

Glac/iw'ia Slilp/llll't'll'la

 

 

Cli/(lllll't/UNIUHHS i'cin/larc/lii

 

62‘

100 O.\'ll't’(l('()('(’ll.\' luvinmrinus

 
   
 
 
   

 

Oslrcat'mt'us lauri

 

P/zit'sc'amilrt'l/a palms

99 013:4: salim

Zea mars

Medicaga lrum'ala/a

Papa/us ll'lt'llfK'tll‘U/Hl
Arabidopsis thaliana A15g24690
- 0.1 substitutions/site

Figure 2.5. Plastid targeted solute transporters of 'Other' or 'Plamae-specific' origin in

Plantae

80

Conclusion

Here we determined the phylogeny of 83 Arabidopsis plastid solute transporters to
determine whether they are of endosymbiotic origin from the captured cyanobacterium,
of host origin, or of a ‘mixed' origin fi'om both of these sources. Our analysis has afforded
a rare look at early, critical events in primary plastid evolution and support the notion that
integration of plastid-host metabolism was primarily driven by host-derived transporters
with important contributions coming from the cyanobacterial endosymbiont and
Chlamydia-like bacteria. Another class of proteins of currently unknown origin included
plant specific transporters such as MEX] . Despite the power of our comparative
approach, our work has some important limitations. One is that because we used the
Arabidopsis transporter set, we most certainly have missed a number of lertae
transporters that are specific to red orgreen algae and have been lost from the
Arabidopsis genome. In addition, we lack significant data fi‘om glaucophytes, but the
upcoming Cyanophora paradoxa (glaucophyte) nuclear genome sequence
WWW will allow us to incorporate this lineage
into future inferences about transporter evolution. It is reasonable to assume, however,
given the wealth of data supporting Plantae monophyly (McFadden, 1999; Rodriguez-
Ezpeleta et al., 2005; Weber et al., 2006; Reyes-Prieto and Bhattacharya, 2007), that our
inferences regarding the red and green lineages also apply to their glaucophyte sisters.
Despite these limitations and the fact that phylogenetic signal is imperfectly maintained
over a billion years of evolution, our comprehensive analysis of the chloroplast solute
transport system will likely hold up and can be further tested as other genome sequences

become available.

81

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86

CHAPTER 3

ANALYSIS OF THE PLASTID PHOSPHATE TRAN SLOCATOR FAIVIILY
FROM THE RED ALGA GALDIERIA SULPHUR/1M3

 

3 This work has been published in Linka M., Jamai A., Weber A.P.M. (2008) Functional
characterization of the plastidic phosphate translocator gene family from the thermo-acidophilic
red alga Galdieria sulphuraria reveals specific adaptations of primary carbon partitioning in
green plants and red algae. Plant Physiology 148: 1487-1496. I performed all of the experiments
shown here.

87

ABSTRACT

In chloroplasts of green plants and algae, C02 is assimilated into triose-phosphates; a
large part of these triose-phosphates is exported to the cytosol by a triose-
phosphate/phosphate translocator (TPT) whereas some is stored in the plastid as starch.
Plastidial phosphate translocators (pPT) have evolved from transport proteins of the host
endomembrane system shortly after the origin of chloroplasts by endosymbiosis. The red
microalga Galdieria sulphuraria shares three conserved putative orthologous transport
proteins with the distantly related seed plants and green algae. However, red algae, in
contrast to green plants, store starch in their cytosol, not inside plastids. Hence, due to the
lack of a plastidic starch pool, a larger share of recently assimilated C02 needs to be
exported to the cytosol. We thus hypothesized that red algal transporters have distinct
substrate specificity in comparison to their green orthologs. This hypothesis was tested by
expression of the red algal genes in yeast and assessment of their substrate specificities
and kinetic constants. Indeed, two of the three red algal pPT candidate orthologs have
clearly distinct substrate specificities when compared to their green homologs. GsTPT
displays very narrow substrate specificity and high affinity; in contrast to green plant
TPTs, 3-phosphoglyceric acid (3-PGA) is poorly transported and thus not able to serve as
a triose phosphate/3-PGA redox shuttle in viva. Apparently, the specific features of red
algal primary carbon metabolism promoted the evolution of a highly efficient export
system with high affinities for its substrates. The low-affinity TPT of plants maintains
triose phosphate levels sufficient for starch biosynthesis inside of chloroplasts, whereas
the red algal TPT is optimized for efficient export of triose phosphate from the

chloroplast.

88

INTRODUCTION

In plants, the photosynthetic light reactions provide the energy for major plastid localized
pathways, such as C02 assimilation, the synthesis of starch, fatty acids, several amino
acids, nucleic acids, and the reductive assimilation of inorganic ions like nitrate and
sulfate (Weber et al., 2005; Zrenner et al., 2006). To supply the cell and the organism
with these primary metabolites, a large number of precursors, end products, and
intermediates have to be transported across the organelle envelope membrane and
therefore present-day plastids are extensively connected to the cytoplasm by metabolite
transporters that reside in the envelope membranes (Weber and Fischer, 2007; Tegeder
and Weber, 2006).

Chloroplasts originated approximately 1.6 billion years ago through a single
primary endosymbiosis between a non-photosynthetic primitive mitochondriate
eukaryote and a cyanobacterium (Y can et al., 2004; Bhattacharya et al., 2007; Reyes-
Prieto et al., 2007). Within a period of 0.15 billion years, establishment of the plastid and
divergence of the three major lineages of the Archaeplastida (Adl et al., 2005), that is the
red algae (Rhodophyceae), green a1 gae/ land plants (Chloroplastida), and glaucophytes
(Glaucophyta), began (Bhattacharya et al., 2004; Yoon et al., 2004). Establishment of the
chloroplast within the host cell required massive remodeling of its membrane proteome;
novel transport proteins to connect its metabolism with the metabolic network of the host
cell had to be acquired (Bhattacharya et al., 2007 ; Weber and Fischer, 2007).
Phylogenetic and phylogenomics analyses recently revealed that a large portion of these
plastid-resident transporters is host-derived, indicating that integration of the chloroplast

with host metabolism was predominantly a host-driven process (Tyra et al., 2007). Genes

89

encoding these transporters are not present in extant cyanobacterial genomes but most are
conserved throughout the Archaeplastida, indicating they have been established at an
early stage during formation of endosymbiosis, likely already at the stage of the protoalga
(Tyra et al., 2007).

Of particular importance for establishment of chloroplasts was an efficient and
controlled export of photoassimilates from the endosymbiont to its host cell (Weber et al.,
2006). In the Chloroplastida, a triose-phosphate/phosphate transporter (TPT) exports a
significant amount of the dihydroxyacetone phosphate (DHAP) that is generated by the
Calvin-Benson cycle to the cytosol; there it mainly serves as precursor for sucrose and
cell wall biosynthesis (Flugge, 1999; Reiter, 2002). However, part of the triose
phosphate is retained in the plastid stroma and used to fuel starch biosynthesis and other
intraplastidial biosynthetic pathways (Zeeman et al., 2007). TPT candidate orthologs are
highly conserved throughout the photosynthetic eukaryotes (Weber et al., 2006) and
transport experiments with isolated organelles or reconstituted organellar membranes
demonstrated triose-phosphate transport activity in the red algae and glaucophytes
lineages, respectively (Schlichting and Bothe, 1993; Weber et al., 2004). Apparently,
triose-phosphate (TP) already served as the main export photoassimilate at the stage of
the proto-alga (Weber et al., 2006). In the plant model organism Arabidopsis thaliana,
additional sugar phosphate transporters with specialized functions have been
characterized (Flugge, 1999). Import of phosphoenolpyruvic acid (PEP) fi'om the cytosol
by the PEP/phosphate transporter (PPT) drives fatty acid biosynthesis and synthesis of
compounds by the shikimate pathway (i.e. aromatic amino acids) (Fischer et al., 1997;

Knappe et al., 2003a; Voll ct aL, 2003) and a glucose-6-phosphate/phosphate transporter

90

(GPT) provides g1ucose-6-phosphate (Glc6P) for starch synthesis in heterotrophic
plastids (Kammerer et aL, 1998; Niewiadomski et al., 2005). Lastly, a protein closely
related to the GPT, the pentose-phosphate/phosphate transporter (XPT) connects the
oxidative pentose phosphate pathways (OPPP) in cytosol and plastid (Eicks et al., 2002;
Fliigge and Gao, 2005).

Phylogenetic analysis showed that candidate orthologs for the PPT and the
GPT/XPT type of translocators are present in the genome of the ancient red microalga G.
sulphuraria, which is separated from green plants by an evolutionary distance of at least
one billion years (Y oon et aL, 2004). However, reconstituted membrane fractions from G.
sulphuraria showed neither significant PEP nor Glc6P transport activity (Weber et al.,
2004). This raises the intriguing question as to whether the genetic repertoire present in
the last common ancestor of Rhodophyceae and Chloroplastida was functionally
maintained or whether it diverged after separation of the two phyla To address this
question, we used G. sulphuraria as a model for the Rhodophyceae. The draft genome of
G. sulphuraria is publicly available and the unicellular organism shares core features of
carbon metabolism with other red algae (Viola et al., 2001; Barbier et al., 20053). In
contrast to higher plants, starch in red algae is produced in the cytosolic compartment,
using UDP-Glucose (UDPGlc) as precursor (Coppin et al., 2005; Patron and Keeling,
2005). Cell wall polysaccharides as well as the major soluble carbohydrate floridoside
(a-D-galactopyranosyl-l-2’-glycerol) are produced in the same compartment. Notably, in
addition to photoautotrophic growth, G. sulphuraria is also able to grow mixo- or
heterotrophically on more than 50 different carbon sources (Gross and Schnarrenberger,

1995; Barbier et al., 2005b). Carbon partitioning thus has to be coordinated not only

91

during alternating light and dark periods but also under continuous heterotrophic growth
conditions. It is not known in detail how the carbon allocation between the plastid and the
cytosol is accomplished in red algae. Based on phylogenetic data, the plastid phosphate
translocators presumably are the major routes for metabolite exchange. They have to
balance a high demand for photoassimilates in the cytosol with maintaining sufficient
levels of triose-phosphates for rhodoplast—localized pathways. Here we report the
heterologous expression and biochemical characterization of the plastid phosphate
transporter family fi‘om G. sulphuraria. With regard to higher plants we will discuss an

alternative strategy to fine tune carbon flux across the plastid membrane in

photosynthetic eukaryotes.

MATERIAL AND METHODS

Growth and sampling of G. sulphuraria cells

Galdieria sulphuraria strain 074W (Gross et al., 1999) was cultured autotrophically in a
minimal salt medium at 37°C with a light intensity of 80 umol photons per square meter
and second. The culture was grown at pH 2.0 in Erlenmeyer flasks under vigorous
shaking at ambient air conditions. Heterotrophic growth of the algal cells was performed
at 37°C in culture flasks in complete darkness in the identical salt medium containing

25 mM glucose as sole carbon source. Cells were harvested at the late logarithmic phase
by centrifugation (3,000 g, 5 min, 4°C), washed with leE buffer (10 mM Tris-HCl

pH 7.5, 1 mM EDTA), flow in liquid nitrogen and stored at -80°C or directly used for

nucleic acid extraction (Barbier et al., 2005) or total membrane enrichment.

92

Isolation of genomic DNA and total RNA from G. sulphuraria

Isolation of genomic DNA was performed as previously described (Barbier et al., 2005).
Briefly, cells were ground in liquid nitrogen and proteins were removed by ‘
Phenol/Chloroform/Isoamylalcohol (PCI) extraction. Nucleic acids were precipitated by
ethanol and RNA was removed by a DNAse-free RNAse treatment and purified by an
additional PCI and ethanol precipitation step. Total RNA was isolated with an acid
guanidium isothiocyanate-phenol/chloroform solution as described by (Chomczynski and

Sacchi, 1987).

Enrichment of total membrane fractions from G. sulphuraria

Harvested cells were resuspended in breaking buffer (100 mM HEPES/KOH, pH 7.6,

1 mM EDTA, 5% glycerol, 5 mM ascorbic acid, 5 mM DTT, 1 mM PMSF) to yield an
OD600 of ~100 (0.5 mL) and an equal volume of acid-washed glass beads (0.4-0.6 mm
size, Sigma Aldrich) were added. Cells were lysed with a mixer mill (MM301, Retsch
GmbH, Germany) for 4 minutes. Broken cells were diluted with 10 mL breaking buffer
and centrifuged (2,000 g, l min, 4°C). The supernatant contained the total membrane
fractions. Membranes were pelleted from the supernatant by ultracentrifugation
(100,000 g, 50 min, 4°C), resuspended in 0.4 mL 10 mM I-IEPES/KOH (pH 7.6), 1 mM

MgC12, and 0.05 mL aliquots were reconstituted into liposomes.

Expression of GsPTs in S. cerevisiae
The coding sequences of all three GsPT were amplified from G. sulphuraria cDNA by

PCR (Platinum Pfic polymerase, Invitrogen, Carlsbad, CA) using the primer combinations

93

listed in Table 3.1. PCR products were subcloned into the pGEM-T Easy vector system
(Promega, Madison, WI) and sequenced. Forward and reverse primers for the genes
GsHET39C12 and GsAl6F5 were designed with BamHI and XhoI restriction recognition
sites, respectively. The GsA14H8 specific forward primer had a KpnI and the reverse
primer an XhoI restriction site. Each cDNA was ligated in frame with an N-terminal
polyhistidine-tag into the yeast expression vector pYESZ/NT (Invitrogen). Standard
molecular methods were applied for DNA restriction and cloning (Sambrook et al.,
1995). The resulting constructs were transformed into competent Saccharomyces
cerevisiae INVScl cells (Invitrogen). Selection, maintenance of the transformants, and
galactose-inducible expression of the recombinant proteins was done according to the
manufacturer’s instructions (pYESZ/NT expression system, Invitrogen). Preparation of
yeast membranes containing heterologously expressed GsI-IET3912, GsAl6F 5, and
GsA14H8 proteins, respectively, was done as described previously (Bouvier et al., 2006),
except that the expression was induced at an OD600 of 0.6 in the presence of 2%
galactose and cells had been cultured for eight hours at 30°C. Presence of recombinant
proteins was verified by standard SDS-PAGE and immunoblot analysis (Sambrook et aL,
1995) using an anti-penta-His antibody (Qiagen, Hilden, Germany) and a secondary anti-

mouse IgG antibody conjugated with an alkaline phosphatase (Promega).

Reconstitution into liposomes and transport activity assays
Acetone-washed bet-phosphatidylcholine (PC) (Sigma-Aldrich, St. Louis, MO) had been
sonicated (5 min on ice, Branson Sonicator 250, output 2, duty cycle 30%) to a final

concentration of 2% (w/v) in 120 mM Tricine-KOH (pH 7.5) and 30 mM internal

94

substrate. 50 uL yeast membrane fraction was reconstituted with 950 uL liposome
suspension by the freeze-thaw procedure (Kasahara and Hinkle, 1977). After thawing, the
proteoliposomes were pulsed 20 times on ice to yield unilamellar vesicles. PDlO-
desalting columns (GE Healthcare) were pre-equilibrated with 150 mM Tricine-KOH
(pH 7.5) and used to remove non-incorporated substrate from the external medium. If not
stated otherwise, all transport studies were initiated with radiolabeled [32P]-
orthophosphate at a final concentration of 0.25 mM. For each data point, 190 uL
proteoliposomes were terminated with 10 uL 10x inhibitor stop solution (200 mM PLP,
20 mM DIDS, 100 mM Tricine-KOH, pH 8.0). Control experiments with membranes
from yeast cells harboring the empty vector were performed in parallel. Adding of
inhibitor mix before addition of radiolabeled substrate was used to monitor unspecific
binding. External phosphate was removed by strong anion-exchange chromatography
with AG-l X8 resin (Acetate form, pre-equilibrated with 150 mM sodium acetate,
BioRad, Hercules, CA). The flow-through of the anion-exchange columns containing the
proteoliposomes with the imported radiolabeled phosphate was quantified by a liquid
scintillation counting. All substrates used for uptake studies were purchased from Si grna-
Aldrich. Kinetic constants were determined by measuring the initial velocity of each
experiment. Michaelis-Menten constant (KM) has been analyzed with at least six external
Pi concentrations ranging between 0.05 mM — 10 mM and competitive inhibition of Pi
transport was assessed by the inhibitor constant Ki as described by Dixon (Dixon, 1953).
GraphPad-Prism Software was used for non-linear regression fitting of all enzyme kinetic

data.

95

RT-PCR gene expression study
DNAse-treated RNA was used for first strand cDNA synthesis (Superscript II F irst-
Strand Synthesis System, Invitrogen). Oligonucleotide primers for expression studies of

the G. sulphuraria genes are summarized in Table 3.1.

Other methods

Protein concentrations were determined with a standard Bradford assay. Proteins in the

membrane fractions were delipidated prior to determination (Shultz et al., 2005).

96

Table 3.1. Oligonucleotide primers designed in this study
Amplified Galdieria sulphuraria gene products were used for cloning heterologous
expression (HE) constructs or Reverse-Transcriptase PCR (RT-PCR) expression
analysis. Underlined nucleotides highlight restrictions sites for cloning into expression

vector system pYES. Fwd, forward primer", Rev, Reverse primer.

 

 

Primer Sequence (5'—3') Direction Purpose
Gs39C12
ML58 TAWATGAACCTTCTAAAGAATCCA fwd HE
TCA
ML59 ATCIEQAQATGAACCCAGAGTCT ACT'IT AT rev HE
TT
ML 1 37 AGCACTTGAACCCI‘TTGTCG fwd RT-PCR
ML 1 3 8 TGCATTGGCAGGAGTGATAG rev RT-PCR
Gsl6F5
ML56 TAQGATCCCAAGGAAACTCACTATTCGTC fwd HE
ML66 GTCT CGAGTCTATTCT ATI'I‘TCTCC'ITCI' TC rev HE
TTGG
ML135 CAACCATGGCTACCTITCGT fwd RT-PCR
ML136 CGTCAAAAAGCCAATCCAAT rev RT-PCR
Gsl4H8
ML94 TGGTACCCAAAGGAGAAAAGGATATCATA fwd HE
AGAGC
ML95 ATCLQQAGACTACTGC’ITI‘TCTCGCTTCG rev HE
ML133 AGAGTGGTGGCTCTCCTCAA fwd RT—PCR
ML134 GATGAATCCGGTCCAGGTAA rev RT-PCR
GsActin
ML132 CACAGCATCTGGCCAAAC'IT fwd RT-PCR
ML13 1 GACCCACATI‘TGTI‘GGAAGG rev RT-PCR

 

97

RESULTS

Molecular features of the plastidic phosphate translocator (pPT) homologues from
G. sulphuraria

Three genes encoding proteins with significant similarity to higher plant pPT have
previously been identified in the genome of the red alga G. sulphuraria (Weber et aL,
2006). Based on detailed phylogenetic analysis, GsHET39C12 is the candidate ortholog
to the plastid triose-phosphate/phosphate translocator (TPT) and will thus be called
GsTPT. GsAl4H8 and GsAl6F5 cluster with the functionally characterized green plant
plastid glucose-6-phosphate/phosphate transport proteins (GPT), and
phosphoenolpyruvate/phosphate translocator proteins (PPT), respectively. They were
thus assigned with the acronyms GSGPT and GsPPT. The coding sequences of GsTPT
and GsPPT contain 1224 nucleotides, corresponding to 407 amino acid residues and
computed molecular masses of 45.83 kDa (T PT) and 44.98 kDa (PPT). GsGPT has a
calculated molecular mass of 45.45 kDa, consisting of 410 amino acid residues that are
encoded by 1233 nucleotides. The N-terminal regions of all three proteins represent
putative plastid target sequences that display only low sequence similarity with the pPTs
from higher plants (Figure 3.1) and the red alga Cyanidioschyzon merolae. The G.
sulphuraria proteins show an average sequence identity of 37% and a 55% similarity
with their A. thaliana homologs and a slightly higher identity of 48% to the
corresponding C. merolae proteins (Table 3.2). Six lysine and two arginine residues that
are invariantly embedded in conserved motifs in all fimctionally characterized pPT
proteins to date (Knappe et al., 2003a), are also found in all three G. sulphuraria

homologues. Relative to the GsTPT sequence, the positively charged amino acids are at

98

positions K123, K199, R261, K266, K267, K361, R362, and K399 (Figure 3.1). All three
putative transport proteins are highly hydrophobic proteins that are predicted by ConPred
Il (Arai et al., 2004) to contain nine to ten membrane spanning alpha-helices. The GsTPT
and GSGPT genes contain one intron that separates a short first exon from the residual
coding sequence (Table 3.2). GsPPT has a total of four exons. The last exon covers
approximately 50% of the CDS. All three sequences have been submitted to GeneBank
and can be also found at the MSU Galdieria genome database (http://genomicsmsuedu/

galdieria). Accession numbers and annotations are listed in Table 3.2.

99

Table 3.2. Molecular characteristics of the plastidic phosphate transporter GsTPT,
GsPPT and GSGPT from Galdieria sulphuraria.

 

 

GsTPT GsPPT GSGPT
Accession number EU853171 EU853172 EU853173
. . . Stig_15 Stig_53 Stig_45
Galdrerra Genome Browser Burld 3.0 Gs21 6 60 Gs53050 Gs48050
Amino acid statisitc report [%]
Identity to AtTPT 33 30 32
Similarity to AtTPT 53 51 53
Identity to AtPPT 31 41 31
Similarity to AtPPT 52 57 52
Identity to AtGPT 33 33 37
Similarity to AtGPT 56 51 55
# of Exons 2 4 2
Exon length (nt) 119 244 1 12
l 105 174 l 121
191
615
# of Introns 1 3 1
Exon length (nt) 53 55 50
47
70
# of transmembrane spans 9 10 9

 

100

Figure 3.1. Protein alignment of plastidic phosphate transporters (pPT) from Arabidopsis
thaliana (At) and Galdieria sulphuraria (Gs)

Multiple sequence alignment program ClustalW2 (Larkin et al., 2007) at EMBL-EBI
lined up the sequence similarities and the tool GeneDoc WWW
WWI) visualized and edited the higher plant pPT proteins and their red
alga] homologues Gs39C12 (GsTPT), Gsl6F5 (GsPPT) and Gsl4H8 (GSGPT). AtTPT,
triose-phosphate/phosphate translocator, AGI genome code At5g46110; AtGPT l,
glucose-6-phosphate/phosphate translocator, At5g54800; AtPPT] ,

phosphoenolpyruvate/phosphate translocator, At5 g33320.

101

* 20 * 4O

 

AtTPT -MESRVLLRATANVVGIPKLRRPIGAIHRQFSTASSSSFSVKPIG 44
Gs39C12 ------------------------------- MRKFDWITRQYLVA 14
AtPPTl MQSSAVFSLSPSLPLLKPRRLSLRHHPITTAASSSDLNVSPNVVS 45
GsAl6F5 MIAFATPLGNTSWKLGYRSLKSVIYFQSTTSSHPRKLTIRPSIQP 45
AtGPTl -MVLSVKQTLSP ------- KIGLFRRNP ----- SSSLGRSPVSLS 32
GsAl4H8 MMIEAAFVPLSCSLFSKQHRWTVVSRNKNKNSISSHMEGSKKLLG 45
* 6O * 80 *
AtTPT GIGEGANLISGRQLR ----- PILLLDSSAINGGEKREILKPVKAA 84
Gs39C12 KARRLPQFYCLANTS ------ LDEPSK ------------------ 35
AtPPTl IPSLSRRSWRLASSD----SPLRAWSGVP--SPISHSLDTNRFRT 84
GsAl6F5 WLPFVQ---NQKTVD----HPSHLTSSFS--SPMMTLGDG----A 77
AtGPTl FPSTELPKRTVLAVS----KPLHLSSSLRAKSPVVRCEAYEADRS 73
GsAl4H8 TPRFTLSRSQFLNVSYLRTKYNNVASSSKGEKDIIRAACRRAE-S 89
100 * 120 *
AtTPT AAEGGDTAGDAKVGFLAKYP TGFFFF F :. 129
Gs39C12 ESIKVTEASQPSQNTASWKR VASYFF I Ii- 80
AtPPTl AATAVPESAEEGDNSGKLTK ELGLLF L I EK' 129
GsAl6F5 ESSTGTSSSNVRQPVQSLQK LTFYIGC 1 LL 122
AtGPTl EPHPIGDDAAAAETKSEAAK IGIYFA :K' 118
GsAl4H8 GGSPQ~--KSSVGVSPTLVH VGFYFF F F E ‘ 131
140 * 160 * 180
AtTPT YNYFPYPYFVS HL YCLIS SV PKRAPIDSNLLK I 174
Gs39C12 LNAYPFPWTVA QLA FYVVP LLH KAPHIPLEDIK L 125
AtPPTl LKALHAPMTVT QFA SVLITI L YKRPKISGAQL L 174
GsAl6F5 LKVFPLFATVT QFL SLVGLA IS HRFQKASLEDLK 167
AtGPTl LNAYPYPWLTS SL SLMMLIS V ETPKTDFDFWK F 163
GsAl4H8 LNMWKYPWVLS QLG LYCTF L TKPNVSKKLI I 176
AtTPT 218
Gs39C12 169
AtPPTl 218
GsAl6F5 212
AtGPTl 207
GsAl4H8 220

 

Figure 3.1. Protein alignment of plastidic phosphate transporters from Arabidopsis
thaliana (At) and Galdieria sulphuraria (Gs)

The Figure 3.1 continues on the following page.

102

AtTPT GQSIPITLWLSA 1L
Gs39C12 RSVFPZPVYLS} CG
AtPPTl GEKPTPWVLG‘. f
GsAl6F5 GTAYTLWVYLSn 1C
AtGPTl GETFPTSVYLSC ,
GsAl4H8 GEFFEPLTYL vs
280 *
AtTPT TD —————— MDSTN
Gs39C12 Is DQTSYKHMSPAN
AtPPTl KK--DDSLDNIT
GsAl6F5 K KG----VQFDNLN
AtGPTl K K—---GKSVSGMN
GsAl4H8 F DFKNEKTLIAQNT
320 * 340
AtTPT ———PKLLNHGFADAIAKVGMTKFISDLFWVGMFYH
Gs39C12 ———PKLYQGWILATSGKTTSMQLZTGLLTSGLFFY
AtPPTl ————— IKFTPSYIQSAGVNVKQIYTKSLIAALCFH
GsAl6F5 GRWREMASVATHIGSEGCT:PVLLLRIAIAGFLHF
AtGPTl ———PQMWVDGWQTALATVG-PQFVWWVVAQSVFYH
GsAl4H8 -——FPP;VS-———A:AGVSKAKLFGSIMFCSLFYH
* 380
AtTPT
Gs39C12
AtPPTl
GsAl6F5
AtGPTl
GsAl4H8
* 420
AtTPT v II KIEEEKRQGKKA 410
Gs39C12 v L YYYSQKIK ————— 362
AtPPTl v R IKPKPKTA---— 408
GsAl6F5 v Q ISTKKKEKIE-- 407
AtGPTl T Q L ———————————— 388
GsAl4H8 T L KLPSKREKQ——- 411
Figure 3.1. (cont’d).

* 240

 
   
   
 

 
  
  
   
 

103

 

   
   
    
   
   
 

 

263
214
263
257
252
265

302
259
306
298
293
310

344
301
346
343
334
348

389
346
391
388
379
393

Heterologous expression of GsTPT, GsPPT, and GsGPT

To assess the substrate specificities of the G. sulphuraria pPTs, we cloned the
corresponding cDNAs into the yeast expression vector pYES/NT under control of the
galactose-inducible GAL4 promoter. The regions of the cDNAs encoding for the putative
target sequences of each protein were removed and instead fused to an N-terminal hexa-
histidine tag. After transformation of the corresponding constructs into the yeast strain
INVScl , all three pPT homologues could be successfully expressed and accumulated in
the membrane fraction (Figure 3.2). Immunoblot analysis with an anti-polyhistidine tag
antibody verified the galactose-inducible accumulation of the pPT proteins (Figure 3.2,
lane 2-4) compared to controls, which maintained the empty expression vector (Figure
3.2, lane 1). The calculated molecular masses of the N-terminal His-tagged proteins were
41.6 kDa, 45.2 kDa and 41.2 kDa for GsTPT, GsPPT and GsGPT, respectively. The
presence of recombinant protein was verified for all biological replicates by Western blot

prior to reconstitution.

104

 

 

Figure 3.2. Expression of plastid phosphate transporter (pPT) homologues from
Galdieria sulphuraria (Gs) in yeast

Upper part: Membrane proteins from galactose-induced S. cerevisiae cells transformed
either with an unmodified expression vector (Lane 1) or with constructs for GsTPT (lane
2), GsPPT (lane 3) and GsGPT (lane 4) were separated by SDS-PAGE and stained with
Coomassie Blue R250. Lower part: Immunoblot analysis of an identical SDS-
polyacrylamide-gel as described above. N-terminal poly-histidine tagged proteins were
colormctrically verified with anti-His antibody and anti-mouse-IgG antibody linked to an
alkaline phosphatase. TPT, triose-phosphate/phosphate translocator; GPT, Glucose-6-

Phosphate-Translocator; PPT, L L ',,......;.."L L ‘ translocator; kDa, kilo

r r K

 

Dalton.

105

Functional analysis of three putative transport proteins from G. sulphuraria

All functionally characterized plastidic phosphate translocators of higher plants catalyze,
in addition to their characteristic substrates, also a strict homo-exchange of ortho-
phosphate (F liigge, 1999). To test whether the yeast-expressed, recombinant GsPTs are
functional, we thus first examined their ability to catalyze the signature Pi homo-
exchange. To this end, they were reconstituted into liposomes that were preloaded with
30 mM Pi (i.e., the liposomes contained 30 mM Pi inside). Then radiolabeled [32P]-
phosphate was added to the liposomes and the uptake kinetics were recorded. In GsTPT
and GsPPT containing proteoliposomes, protein-mediated isotope equilibrium was
reached within ten minutes, using an external Pi concentration of 0.25 mM (Figure 3.3).
GsGPT mediated Pi uptake equilibrated after 15 minutes with a four times higher
external [32P]-Pi concentration of 1 mM. Ortho-phosphate exchange followed a first-
order kinetic with an equilibrium plateau (Vmax) of 400:13.74, 187.3133] and
237.1:702 nmol per mg protein and initial rates of 60.691248, 30.94:0.43 and
23.23:].27 nmol per mg protein and minute for GsTPT, GsPPT, and GSGPT,
respectively. Control membranes from yeast cells transformed with an empty pYES/N T
vector or boiled recombinant proteins showed negligible accumulation rates over time
(data not shown). PLP and DIDS also effectively inhibited GspPT mediated import of
[3 2P]-phosphate (data not shown). Marginal uptake of radiolabeled [3 2P] -phosphate
compared to control experiments was measured when vesicles did not contain Pi as a
counter substrate (Figure 3.3). In summary, all putative phosphate transport proteins from
G. sulphuraria were frmctionally expressed in yeast and are able to catalyze the signature

Pi homo-exchange activity, similar to their higher plants homologs.

106

Figure 3.3. Time-kinetics of pPT homologues fiom Galdieria sulphuraria (Gs)
Liposomes were reconstituted with membrane proteins fi'om S. cerevisiae cells
expressing GsTPT (panel A), GsPPT (panel B) or GsGPT (panel C). Uptake of ortho-
Phosphate (Pi) was measured in the presence (0) or absence (0) of intraliposomal Pi
(30 mM). Transport was initiated with a final concentration of 0.25 mM [32P]-Pi (panel
A & B) or 1 mM [32P]-Pi (panel C). Diagrams represent a typical result of at least three

independent studies.

107

400-
300-
2004

100-

 

Uptake of Pi
[nmol/mg protein]

0 2 4 6 s in
Time [min]

    

 

Uptake of Pr
[nmol/mg protein]
3
f

 

 

i

0 2 4 6
Time [min]

A

‘1
I
8

 

Uptake of Pi
[nmol/mg protein]

 

 

 

Time [min]

Figure 3.3. Time-kinetics of pPT homologues from Galdieria sulphuraria (Gs)

108

Transport properties of GsTPT, GsPPT and GsGPT
To assess the substrate specificity of the recombinant GspPT proteins, vesicles were
preloaded with saturating concentrations (i.e., 30 mM) of various counter-exchange
substrates and initial rates of [32P]-Pi uptake were determined.

GLUE Relative to the Pi/Pi homo-exchange experiment, the uptake rate of [32P]-
Pi into proteoliposomes reconstituted with GsTPT was 86.7:13.3% when
proteoliposomes were preloaded with the triose-phosphate DHAP. Uptake rates into
liposomes preloaded with 3-phosphoglyceric acid (3-PGA), phosphoenolpyruvic acid
(PEP), or glucose-6-phosphate (Glc6P), respectively, were low (Figure 3.4A). Uptake
was also negligible when GsTPT liposomes were preloaded with glucose-l-phosphate
(Glc l P), fructose-6-phosphate (Frc6P), galactose-1-phosphate(GallP), glycerol-3-
phosphate (Gly3P) (Figure 3.4A), adenosine-monophosphate (AMP), uridine-
monophosphate (UMP), gluconate-6—phosphate (Gluc6P), ribose-S-phosphate (Rib5P),
ADP-glucose (ADP-Glc), UDP-glucose (UDP-Glc), UDP-galactose (UDP-Gal), alpha-
ketoglutaric acid (or-KG), malic acid (Mal), pyruvic acid (Pyr), or oxaloacetic acid
(0AA) (data not shown). GsTPT has an apparentKM value of 0.33:0.07 mM for Pi and
DHAP competitively inhibited Pi uptake with a Ki value of 0520.04 mM (Table 3.3). 3-
PGA and PEP inhibited [32P]-Pi uptake only at non-physiological concentrations of 8
and 9 mM, respectively, and Glc6P did not show significant inhibition of Pi transport at
any of the tested concentrations.

GsPPT: Reconstituted GsPPT protein efficiently used Pi and PEP as a counter

 

substrate for the import of radiolabelled [32P]-phosphate (Figure 3.43). 18 additional

substrates, as listed above for the GsTPT, showed marginal initial Pi uptake rates of less

109

than 30% of the Pi/Pi homo-exchange rate (Figure 3.4B and data not shown). GsPPT has
an apparent KM value of 0.76:0.075 mM for Pi and a Ki value of 0.36:0.04 mM for
PEP. Compared to PEP, ll-times higher 3-PGA and 22-times higher DHAP levels,
respectively, were needed to inhibit Pi uptake by 50% (Table 3.3). Glc6P had no affinity
for the Pi binding site.

@911: Vesicles reconstituted with the GPT homologue from G. sulphuraria were
preloaded with the same 20 substrates as given above; in all cases the [32P]-Pi uptake
was much lower than for liposomes preloaded with 30 mM Pi (Figure 3.4C). GsGPT has
raflier low affinity to ortho-phosphate (KM: 5.07:0.4 mM) (Table 3.3). Compared to
non-preloaded proteoliposomes (l3.03:5.9%), approximately three- to fourfold higher
uptake rates were measured when liposomes were preloaded with either 3-PGA or
DHAP. Importantly, however, even high concentrations of 3-PGA and DHAP did not
significantly inhibit Pi homo-exchange (Table 3.3). Glc6P and PEP are non-relevant

substrates for the GsGPT protein.

110

Figure 3.4. Internal substrate dependence on the transport activities of GsTPT, GsPPT,
and GsGPT

Liposomes were preloaded with various substrates (30 mM) and reconstituted with
membrane proteins from yeast cells expressing GsTPT (panel A), GsPPT (panel B) or
GsGPT (panel C). Uptake experiments were initiated with 0.25 mM [32P]-Pi (panel A &
B) or 1 mM [32P]-Pi (panel C) and terminated afier 1.5 min (GsTPT), 2 min (GsPPT) or
4 min (GsGPT). As negative control, proteoliposomes were preincubated (2 min) with
inhibitor stop solution to calculate net uptake activity. Relative uptake activities were
compared to the Pi/Pi counter-exchange experiment, which was set to 100%. Data are
summarized as the arithmetic mean 1 SE of at least three independent experiments. Pi,
ortho-phosphate; DHAP, dihydroxyacetone phosphate; 3-PGA, 3-phosphoglyceric acid;
PEP, phosphoenolpyruvic acid; Glc6P, glucose-6-phosphate; GlclP, glucose-1-
Phosphate; F rc6P, fructose-6-phosphate; GallP, galactose-l-phosphate, Gly3P, glycerol-

3-phosphate.

111

Internal substrate

 

0 20 40 60 80 100
Relative uptake [%]

B

 
   
  
  
  
 
 
 
 

Gly3P
GallP
Frc6P
GlclP
Glc6P
PEP
3-PGA
DHAP
Non
Pi
0 20 40 60 80 100
Relative uptake [%]

Internal substrate

 
   
 

Gly3P C
GallP
Frc6P
GlclP
Glc6P
PEP
3-PGA
DHAP
Non

Pi

0 20 40 60 80 100

Relative uptake [%]

  
  
  
 
 
 
 

Internal substrate

Figure 3.4. Internal substrate dependence on the transport activities of GsTPT, GsPPT

and GsGPT

112

Table 3.3. Comparison of kinetic constants from G. sulphuraria and higher plant pPT
homologues

The Michaelis-Menten constant (KM) for Pi was determined using various external
[32P]-Pi concentrations (0.05 mM - 10 mM). The competitive inhibition constant (Ki) of
[32P]-Pi uptake was assayed at two different external Pi concentrations with increasing
inhibitor concentrations (0.05 mM — 20 mM) (Dixon, 1953). All proteoliposomes were
preloaded with 30 mM Pi. Data represent the arithmetic mean :1: SE of at least three
independent experiments. Recombinant higher plant pPT data are taken from Flugge
(1999). “None” refers to: No competitive inhibitory constant could be measured under
the given experimental conditions. Abbreviations are listed in Figure 3.4.

 

 

Kinetic TPT homologue PPT homologue GPT homologue
Constants [mM] [mM] [HIM]
Plants Gs Plants Gs Plants Gs
KM (Pi) 1.0 0.33 2 0.07 0.8 0.76 :r: 0.08 1.] 5.07 a: 0.4
Ki (DHAP) 1.0 0.5 :t 0.04 8.0 7.93 :1: 0.23 0.6 None
Ki (3-PGA) 1.0 7.71 a 0.47 4.6 3.95 :r: 0.1 1.8 None
Ki (PEP) 3.3 8.75 = 0.40 0.3 0.36 a: 0.04 2.9 None
Ki (Glc-6P) None None None None 1.1 None

 

Transcript levels and protein activity of the plastidic phosphate translocator
homologues in G. sulphuraria cells

In land plants, TPT expression is confined to photosynthetic tissues whereas GPT
expression is highest in heterotrophic tissues (Kammerer et al., 1998; Flugge, 1999;
Niewiadomski et al., 2005). PPT-genes are expressed at low levels in various tissues
(Knappe et al., 2003b). To determine the expression pattern of GspPTs, semi-quantitative
RT-PCR was used. Figure 3.5 shows that Gs TPT, GsPPT, and GsGPT transcripts are
clearly detectable and abundant under both, autotrophic and heterotrophic growth
conditions. The steady state transcript levels of all three genes were higher than those of

the reference gene actin (G515600; Galdieria Genome Browser Build 3.0). Liposomes

113

reconstituted with total membrane fractions exhibit high [32P]-Pi import using DHAP as
a counter substrate under both culturing regimes (Figure 3.5). Exchange rates with PEP
and 3-PGA as counter-exchange substrates were much lower as compared to DHAP.
Exchange of glucose-6-phosphate with Pi was close to the background values of

imloaded vesicles (Figure 3.5).

114

Figure 3.5. Transcript abundance and protein activity of GsTPT, GsPPT, and GsGPT in
G. sulphuraria cells

Upper part: Total RNA from auto- and heterotrophically cultured G. sulphuraria cells
was isolated, treated with DNase and used for first strand cDNA synthesis. Expression of
GsTPT, GsPPT, GsGPT and actin transcripts was amplified by PCR with 25, 30 or 35
elongation cycles and visualized on an ethidium bromide-stained 1.25% (w/v) agarose-
gel. Lower part: Relative [32P]-Pi uptake rates into liposomes reconstituted with total G.
sulphuraria membranes isolated from heterotrophically (black bar) or autotrophically
(white bar) grown cells, respectively. Membrane fractions were reconstituted into
liposomes preloaded with 30 mM various substrates. Experiment was performed and

abbreviations were used as given in the legend to Figure 3.4.

115

Autotrophic Heterotrophic

 

PCR m m
GsTPT
GsPPT
GsGPT «II-I-
GsActin
120- D Autotrophic

—

100. - Heterotrophic

W
O
I

ch
6
1

Relative uptake [%
N 65
e G
l I

 

 

 

 

ml—

(““399 “0193 .5308 9"? (X6?

Figure 3.5. Transcript abundance and protein activity of GsTPT, GsPPT, and GsGPT in

G. sulphuraria cells

116

DISCUSSION

In this study, we report the molecular and biochemical characteristics of the plastidic
phosphate translocator family from the unicellular red alga G. sulphuraria. All three
members of the GsPT gene family were heterologously expressed in S. cerevisiae,
functionally reconstituted into liposomes (Figure 3.2 and Figure 3.3), and their substrate

specificities and kinetic constants were assessed (Figure 3.4 and Table 3.3).

The G. sulphuraria genome encodes a high-affinity TPT with narrow substrate
specificity

Recombinant G. sulphuraria TPT mediates a strict counter-exchange of the triose-
phosphate DHAP with ortho-phosphate (Figure 3.4). Interestingly, the kinetic constants
of GsTPT considerably differ from those of the green plant TPT (Table 3.3). The red
algal transporter exhibits a three times higher affinity for Pi and, as indicated by its low
Kr value, the substrate DHAP has a twofold higher affinity to the Pi binding site. Further,
GsTPT is highly specific for DHAP. In contrast to the green plant ortholog, 3-PGA
poorly acts as a counter substrate, even under saturating conditions (Figure 3.4A). In
addition, 3-PGA is a weak competitive inhibitor of Pi import with an eightfold and
fifteen-fold higher Ki value as compared to the TPT fi'om plants and the Ki (DHAP)
constant from GsTPT, respectively (Table 3.3). These results argue strongly against 3-
PGA as a physiological relevant substrate of GsTPT. PEP and Glc6P were even less
efficient exchange substrates and inhibitors of Pi transport (Figure 3.4 and Table 3.3).
Very likely, the distinct kinetic properties of GsTPT represent an important adaptation to

red algal carbon metabolism. In contrast to the Chloroplastida, red algae store an

117

insoluble starch-like polymer called floridean starch and synthesize cell wall building
blocks and the main soluble carbohydrate floridoside, composed of UDP-Gal and Gly3P
moieties in their cytosol (Viola et al., 2001; Collen et al., 2004; Barbier et al., 2005a).
Hence, primary carbon partitioning in red algae is predominantly organized within the
cytosol and GsTPT enables solely the export of triose-phosphates, even at low stroma]
concentrations, to cope with the massive cytosolic demand of recently fixed carbon
(Figure 3.6). On the other hand, GsTPT is highly expressed and active in
heterotrophically cultured G. sulphuraria cells (Figure 3.5). Under these conditions,
GsTPT presumably mediates TP import into the plastid, thus mediating carbon flux
between cytosol and non-photosynthetic rhodoplasts. In contrast, land plants export TP
from photosynthetic chloroplasts via TPT and import Glc6P into heterotrophic plastids
via GPT. The TPT is an integral part of the photoassimilate partitioning in plants
(Hausler et al., 2000; Smith and Stitt, 2007). Accumulation of sucrose and hexose-
phosphates in the cytosol sequesters cytosolic Pi and thus slows down the export of
triose-phosphate. A resulting increased stroma] 3-PGA/Pi ratio allosterically activates
ADP-glucose pyrophosphorylase, which is the committing step of plastidic starch
biosynthesis (Ballicora et al., 2004). Plants have to maintain sufficiently high levels of TP
in the plastid stroma to enable both the regeneration of the C02 acceptor ribulose-1,5-
bisphosphate and starch biosynthesis during the light period (Zeeman et al., 2007). The
apparent lower affinity of the plant transporter for Pi and TP thus ensures higher steady

state levels of TP in the stroma to drive these reactions.

118

G. sulphuraria has a PPT with similar properties as its green plant ortholog

PEP uptake from the cytosol is required for stroma localized fatty acid and shikimate
biosynthesis (F lfigge, 1999). Previously published results have been ambiguous about the
existence of a putative PEP/Pi transporter (PPT) in G. sulphuraria. Phylogenetic analysis
revealed an orthologous candidate gene (Weber et al., 2006), although isolated membrane
fractions did not exhibit pronounced PEP uptake activity (Weber et al., 2004). Here, we
show that EST GsAl6F5 (Weber et al., 2006) represents a highly conserved PPT with
almost identical kinetic constants in both the green and red lineages (Table 3.3). Both
transporters are highly specific for Pi and PEP and poorly accept 3-PGA, DHAP, or any
other of the 16 tested metabolites as a substrate (Figure 3.4B, data not shown). In
addition, we now employed an improved reconstitution method for total membranes from
autotrophic and heterotrophic cell cultures that yielded lower background and a weak but
clearly detectable PEP/Pi counter-exchange activity (Figure 3.5). The low PEP transport
capacity (Figure 3.5) mirrors published data from chloroplasts of C3 land plants (Flugge
and Weber, 1994). TPT is by far the most abundant transport protein in C3 chloroplasts
and accounts for a high DHAP exchange capacity (F lugge, 1999). Fatty acid biosynthesis
and the shikimate pathway in red algae are predicted to be similar to that of seed plants
and are presumably localized in the plastid, based on phylogenetic analyses and N-
terminal target sequence predictions (Weber et al., 2004; Richards et al., 2006). A
question that cannot be conclusively answered at the moment is whether rhodoplasts,
analogous to chloroplasts of green plants, also have negligible activities of plastid
phosphoglyceromutase and enolase to produce PEP from plastidial 3-PGA (Trimming

and Emes, 1993). The G. sulphuraria genome encodes for two phosphoglyceromutase

119

isozymes (Gs04140 and Gs52680) and a single enolase (Gs21490), which are all three
computationally predicted as cytosolic enzymes (Nielsen et al., 1997) without any N-
terminal targeting sequences, when compared with their homologous genes from A.
thaliana (phosphoglyceromutase: At4g09520, At3g08590, At1g09780; enolase:
At2g36530, At2g29560, At1g74030) (Friso et al., 2004; Larkin et al., 2007).
Alternatively, PEP could be generated fi'om pyruvic acid via the pyruvate orthophosphate
dikinase (PPDK) reaction. The annotated PPDK gene Gs42070 is computationally
predicted to be localized in the cytosol, exhibiting a high similarity with cyanobacterial
PPDK genes, and is not phylogenetically related to the dual targeted enzyme from A.
thaliana (At4g15530) (Parsley and Hibberd, 2006). While computational targeting
predictions have to be takenwith a grain of salt, bioinformatics analysis indicates that
conversion of TP to PEP is not possible in G. sulphuraria and rhodoplasts thus depend on

PEP import fi'om the cytosol to drive PEP-depended reactions in the stroma (Figure 3.6).

G. sulphuraria does not possess a plastidic hexose phosphate importer

Reconstituted recombinant putative GsGPT mediated a strict Pi/Pi homo-exchange that
followed a first order rate kinetics (Figure 3.3). An unusually high KM value of 5 mM for
Pi (Table 3.3) indicates that GsGPT under physiological conditions is not able to compete
with GsTPT and GsPPT for the common substrate Pi. While liposomes preloaded with
high (i.e., saturating) concentrations of DHAP or 3-PGA (i.e., 30 mM internal substrate
concentration) showed Pi uptake rates of approximately 50% of Pi/Pi homo-exchange, it
is important to notice that neither 3-PGA nor DHAP significantly inhibited Pi/Pi homo-

exchange either at physiological concentrations or at lOO-fold excess (Table 3.3). The

120

prime substrate candidate glucose-6-phosphate was poorly accepted under saturating
conditions (Figure 3.4) and it did not inhibit Pi import in the Pi/Pi homo-exchange setup
(Table 3.3). These results strongly indicate that 3-PGA, DHAP, and Glc6P are not
physiologically relevant substrates of the putative GsGPT. We also tested various
additional metabolites, such as nucleotide-sugars, mononucleotides, hexose-phosphates
(GlclP, Frc6P, and Gall P), pentose-phosphates and a precursor of floridoside
biosynthesis, glycerol-3-phosphate. All of these are poorly exchanged with [32P]-Pi and
the physiological substrate of GsGPT thus remains unidentified. In non-green plastids of
plants, Glc6P is the preferred precursor for starch synthesis and NADPH generation via
the oxidative pentose phosphate pathway (OPPP) due to the absence of a fructose-1,6-
bisphosphatase (FBPase) activity (Flugge, 1999). In photosynthetic tissues, plastidic

F BPase is inactivated at night and thus hexose phosphates cannot be generated fi'om
triose phosphates (and vice versa). In contrast, red algal starch biosynthesis is cytosolic
and plastidic FBPase fiom G. sulphuraria is not subject to a strict redox regulation
(Reichert et al., 2003; Oesterhelt et al., 2007). Importing triose-phosphates via the GsTPT
could thus sustain the production of hexose-phosphate for carbon and NADPH supply in
the rhodoplast during the night or prolonged heterotrophic growth conditions, thus
bypassing the requirement for a plastidic hexose phosphate translocator (Figure 3.6).

Further work will be required to pinpoint the catalytic activity of this enigmatic protein.

An operative DHAP/3-PGA reduction shuttle is unlikely in G. sulphuraria
3-PGA is a major substrate for the TPT in higher plants (F lfigge, 1999). In spinach and

other model plants, 3-PGA levels in the plastid and the cytosol frequently exceed DHAP

121

levels by several fold (Gerhardt et al., 1987). However, the affmity of the spinach TPT is
identical for both metabolites. It has been proposed that in vivo a significant amount of 3-
PGA transport occurs across the envelope membrane; the TPT could thus operate as a
NAD(P)H reduction equivalent shuttle between stroma and cytosol (Heineke etal.,
1991). The substrate specificity and the kinetic constants of recombinant proteins, as
discussed in detail earlier, and total membrane fiactions from G. sulphumria presumably
cannot sustain a physiological relevant exchange rate of 3-PGA across the rhodoplasts
envelope membrane. Still, DHAP offers a possibility to export reduction equivalent to the
cytosol. The non-phosphorylating glyceraldehyde-3 -phosphate dehydrogenase (NP-
GAPDH) (Rius et al., 2006), which is also present in the draft genome sequence of the
red alga (Gs00840) or the combined glycolytic reaction sequence of glyceraldehyde-3-
phosphate dehydrogenase (GAPDH) and 3-PGA kinase (PGK) (Plaxton, 1996) generates
NADPH, or NADH and ATP, respectively. However, in contrast to green plants, not 3-

PGA but Pi must be postulated as the alternative counter-substrate for DHAP.

Impact of subtle differences of TPT in Chloroplastida and Rhodophyta
Triose-phosphate is the main entry and branching point in the glycolytic network in
photosynthetically active cells (Plaxton, 1996). Metabolic flux has to be regulated
between generating hexose-phosphates via gluconeogenesis and glycolytic “downstream”
products, such as 3-PGA, PEP, and pyruvic acid. In higher plants, the redox transfer by a
TP/3-PGA shuttle via (NP)-GAPDH and PGK is proposed to be marginal and GAPDH
expression is mamly induced under stress conditions, such as anaerobiosis and heat shock

or sucrose feeding (Yang et al., 1993; Bustos et al., 2008; Holtgrefe et al., 2008). On the

122

other hand, glycolysis has to provide adequate amounts of PEP to replenish TCA
intermediates used for biosynthesis of, e.g., amino acids, to produce pyruvic acid for
mitochondrial metabolism, or to fuel plastid fatty acid biosynthesis (Plaxton, 1996).
Because 3-PGA transport is seemingly a specific adaptation in plants, we hypothesize
that 3-PGA transport across the chloroplast envelope membrane might provide a bypass
of the complex regulation of the cytosolic triose-phosphate pool that permits supporting
cytosolic PEP biosynthesis and serves under stress conditions as an additional redox

shuttle system.

123

Figure 3.6. Flexible transport of triose-phosphate across the plastid membrane is central
for the cellular carbon metabolism in the red alga G. sulphuraria

In photosynthetically active cells, triose-phosphate is exported via the GsTPT from
rhodoplasts to sustain soluble and insoluble carbohydrate synthesis (floridoside and
floridean starch) in the cytosol. Under heterotrophic growth conditions the same
transporter supplies the rhodoplasts with triose-phosphate for, e.g., hexose-phosphate
synthesis. Rhodoplasts are not capable to produce PEP via partial glycolysis and are thus
dependent on the import of PEP from the cytosol via the PPT protein for fatty acid and
aromatic amino acid biosynthesis. The red algal plastid envelope does not contain any 3-
PGA and Glc-6—P transport activities, which is a specific trait of green plants. PPT,
PEP/phosphate-translocator; TPT, triose—phosphate/phosphate-translocator; F rcl ,6BP,
fructose-1,6-bisphosphate; Gal, galactose; Glc, glucose; Ru-l,SBP, ribulose-1,5-
bisphosphate; other acronyms are listed in the legend to Figure 3.4. The figure in this

dissertation is presented in color.

124

 

C I
m” Floridoside

KRhodoplast_ .. _____ —L UDP-Gal

 

  
       

 

 

 

 

 

 

UDP-
Glc
Glc-6-P HGlc-1 -P
GIc-6-P l Glucose UDP-
Glc
PEP T /
Floridean
Starch

 

 

 

 

Figure 3.6. Flexible transport of triose-phosphate across the plastid membrane is central

for the cellular carbon metabolism in the red alga G. sulphuraria

125

Conclusions
Although the primary structures of plastidic phosphate translocators are highly conserved
between green plants and red algae, these proteins have evolved quite distinct
biochemical‘characteristics in the different lineages of the Archaeplastida. Comparative
biochemical analysis of candidate orthologs fi'om distantly related organisms thus
provides novel insights into alternative modes for regulating “ancient” metabolic
pathways. While the triose-phosphate/phosphate antiport activity across the plastid
envelope membrane is highly conserved across all photosynthetic eukaryotes analyzed to
date, the red algal transporters catalyze neither 3-PGA nor Glc6P transport (Figure 3.6).
G. sulphuraria must have evolved alternative mechanisms to distribute triose-
phosphate into various pools ('Le. floridean starch, floridoside, fatty acid and amino acid
synthesis), a less stringent redox control of Calvin cycle, oxidative pentose phosphate
pathway, and glycolysis (Oesterhelt et al., 2007). Plastidial phosphate translocators thus
represent crucial components of primary carbon partioning in higher plants and red algae
that have evolved to the specific requirements of each lineage through modulation of

substrate specificities and kinetic constants.

126

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132

CHAPTER 4

PROPOSED METABOLITE TRANSPORT PROCESSES FOR THE RE-
ASSIMILATION OF AMMONIA DURING PHOTORESPIRATION1

 

4 This work has been published in Linka M., and Weber A.P.M. (2005) Shufiling ammonia
between mitochondria and plastids during photorespiration. Trends in Plant Science 10:46] -465.

133

ABSTRACT

Surprisingly, it was recently found that glutamine synthetase is dual targeted to
chloroplasts and mitochondria in Arabidopsis thaliana leaves. Mitochondrial glutamine
synthetase likely assimilates ammonia, which is generated in large amounts in
mitochondria during photorespiration. However, ammonia assimilation is a two-step
process and the second step, catalyzed by glutamate synthase, is exclusively located in
plastids. Hence, a shuttle for ammonia, possibly in the form of amino acids, is required
between mitochondria and plastids. We discuss two alternative shuttles, an ornithine-
citrulline shuttle and a glutamine-glutarnate shuttle. Both shuttles allow the safe transport
of the toxic metabolite ammonium in the form of amino acids. The ornithine—citrulline
shuttle also provides a means for the transport of carbon dioxide from mitochondria to
plastids, but this shuttle requires more energy than the alternative glutamate-glutamine

shuttle.

RESULTS

Photorespiration is a complex, multi-compartrnent biochemical pathway
Ribulose-1,5-bisphosphate carboxlyase/oxygenase (RubisCO) is a bifimctional enzyme
that catalyzes both the carboxylation of ribulose 1,5-bisphosphate (RuBP) and its
oxygenation. Oxygenation of RuBP produces one molecule of 3-PGA and one molecule
of 2-phosphoglycolate (2-PG). 2-PG cannot be directly metabolized inside chloroplasts;
instead, two molecules of 2-PG are recycled to one molecule of 3-PGA by the complex,

multi-compartrnent photosynthetic carbon oxidation cycle (Figure 4.1). Because this

134

pathway leads to consumption of oxygen and production of carbon dioxide in the light, it
is also called photorespiration. The photorespiratory pathway involves chloroplasts,
peroxisomes, and mitochondria. 2-PG is dephosphorylated by phosphoglycolate
phosphatase (PGP) in the chloroplast and the resulting glycolate is shuttled to
peroxisomes where it is oxidized to glyoxylate, which is subsequently transaminated to
glycine. Glycine is transported to the mitochondria where two molecules of glycine are
converted to one molecule each of serine, ammonia, and carbon dioxide by glycine
decarboxylase (GDC) and serine hydroxymethyltransferase (SHMT). Serine is
transported back to the peroxisomes, converted to glycerate by serine glyoxylate
aminotransferase and hydroxypyruvate reductase (HPR) and glycerate is finally shuttled
to the chloroplasts where it is phosphorylated to 3-PGA by glycerate kinase (GLYK). The
reaction sequence converts two molecules of 2-PG into one molecule of each 3-PGA and
carbon dioxide and thus leads to the net loss of one carbon (W ingler et al., 2000; Douce
and Neuburger, 1999). The oxidation of glycine by GDC in mitochondria during
photorespiration generates massive amounts of the cytotoxic metabolite ammonia,
exceeding ammonia production fi'om nitrate via the concerted action of nitrate reductase
and nitrite reductase by at least one order of magnitude (Ogren, 1984). It was assumed
that photorespiratory ammonia would be either actively or passively transported to
plastids to be assimilated into glutamate by the glutamine synthetase 2 (GS2) and Fd-

dependent glutamate synthase (F d-GOGAT) system (Figure 4.1).

135

Glutamine synthetase is dual targeted to plastids and mitochondria

The surprising discovery of dual targeting of GS2 to both plastids and mitochondria

(T aira et al., 2004) has changed this view - the presence of G82 in mitochondria opens
the possibility of ammonia assimilation by GS in the mitochondrial matrix, directly at the
site of ammonia generation by GDC. However, to maintain the nitrogen stoichiometry of
the photorespiratory pathway, glutamine (Gln) needs to be converted to glutamate (Glu),
which serves as amino donor for the peroxisome-localized glyoxylate aminotransferase
reaction. The conversion of glutamine and alpha-ketoglutarate (or-KG) to glutamate is
catalyzed by F d-GOGAT and this enzyme is exclusively localized in the plastid stroma
(Lancien et aL, 2002; Coschigano et al., 1998). Alternatively, it could be envisaged that
ammonia would be directly assimilated into glutamate in mitochondria by glutamate
dehydrogenase (GDH). However, GDH mutants do not show a photorespiratory
phenotype in ambient air (Melo-Oliveira et al., 1996), whereas Fd-GOGAT mutants show
a strong photorespiratory phenotype and are not viable in ambient air (Coschigano et al.,
1998; Somerville and Ogren, 1980). Hence, Fd-GOGAT is essential and glutamine (or
ammonia) needs to be shuttled fi'om mitochondria to chloroplasts to close the

photorespiratory nitrogen cycle.

136

Figure 4.1. Simplified scheme of the current textbook photorespiratory cycle

The oxygenation of ribulose-1,5-bisphosphate (RuBP) leads to the production of 2-
phosphoglycolate (2-PG) which is dephosphorylated to glycolate (Glt). After transfer to
peroxisomes, glycolate is oxidized to glyoxylate (Gox) and aminated to glycine (Gly),
which is then transported to the mitochondria. Gly is oxidatively decarboxylated by
glycine decarboxylase (GDC) in the mitochondrial matrix, yielding Methylen-THF, C02,
and ammonia. Ammonia and C02 arrive back in the chloroplasts, presumably by
diffusion, to be re-assimilated by glutamine synthetase (GS) and glutamate synthase
(GOGAT) and RubisCO, respectively. Glutamate, the amino donor for the amination of
Gox in the peroxisomes and the reaction product alpha-ketoglutarate (or-KG) are shuttled
between plastids and peroxisomes by a plastidic malate-coupled two-translocator system
(Weber and Flugge, 2002; Weber et al., 2004) and unknown transporters in the
peroxisomal membrane. For sake of simplicity, the conversion of glycine to serine, and
the downstream reaction sequence leading to the conversion of serine to 3-
phosphoglycerate is omitted. (1) plastidic glycolate transporter; (2) peroxisomal glycolate
transporter; (3) plastidic glutamate/malate translocator; (4) plastidic cr-KG/malate
translocator; (5) peroxisomal glutamate transporter; (6) peroxisomal a-KG transporter;
(7) peroxisomal glycine transporter", (8) mitochondrial glycine transporter. The figure in

this dissertation is presented in color.

137

     
   

Peroxisomes

02
_ Glt H202

Figure 4.1. Simplified scheme of the current textbook photorespiratory cycle

A citrulline-omithine shuttle between plastids and mitochondria

To couple mitochondrial ammonia assimilation with plastidic glutamate biosynthesis,
Taira et a]. (2004) have suggested a citrulline-omithine shuttle between chloroplasts and
mitochondria (Figure 4.2) and have supported this concept by demonstrating that isolated
leaf mitochondria are able to catalyze the glutamine- and ATP-dependent conversion of
ornithine to citrulline. Ornithine-citrulline transporters are well characterized in yeast and
mammals. For example, two closely related human omithine-citrulline transporters have
been cloned and functionally characterized; they transport arginine, lysine, ornithine, and

citrulline by exchange and unidirectional mechanisms (F iermonte et al., 2003). Less is

138

known about mitochondrial ornithine-citrulline transporters in plants. The closest
relatives of mammalian ornithine-citrulline transporters in Arabidopsis are the basic
amino acid carriers AthACl (At2g33820) and AthACZ (Atl g79900) and it was
shown that AthACl and AthAC2 are able to relieve arginine-auxotrophy of a yeast
mutant that is deficient in the mitochondrial ornithine/arginine transporter (Catoni et al.,
2003; Hoyos et al., 2003). Recombinant reconstituted AthACl catalyzed the counter-
exchange of arginine with lysine, ornithine, arginine, and histidine, but transport of
citrulline was marginal (Hoyos et al., 2003). The substrate spectrum of AthAC2 has not
yet been tested in a reconstituted system. Hence, the Arabidopsis ornithine-citrulline
transporter has not yet been unequivocally identified. Robert Ludwig (1993) also
previously showed that isolated Arabidopsis chloroplasts are able to convert externally
supplied citrulline into ammonium, bicarbonate, ornithine, and ATP. This indicates the
presence of a citrulline transporter in the chloroplast envelope membrane, although
uptake of citrulline into chloroplasts has not been directly shown by, e.g., the silicon-oil
filtration-centrifugation technique. In summary, evidence derived from experiments with
isolated chloroplasts and mitochondria indicates that the components of the proposed
citrulline-ornithine shuttle seem to be present and active in leaves but the corresponding
proteins remain to be identified. Importantly, the ornithine-citrulline shuttle would not
only (indirectly) transfer ammonia fi'om mitochondria to plastids, but also simultaneously
carbon dioxide, and thus both products of the glycine decarboxylase reaction that need to
be re-assimilated in the chloroplast stroma. Although this concept is appealing, it comes
at a price: Whereas fixing one molecule of 02 by RubisCO and salvaging of the resulting

2-phosphoglycerate (2-PG) by a textbook C2-cycle (not including re-assimilation of C02

139

that is lost during the process) costs approximately 9.5 ATP (Canvin and Salon, 1997),
two additional ATP would be required in the proposed citrulline-ornithine cycle due to
mitochondrial carbamoylphosphate synthase activity (the additional ATP required by
GS2 in mitochondria is offset by production of one ATP by carbamate kinase in plastids),

thus raising the overall ATP-consurnption by 20% to 11.5 ATP.

140

Figure 4.2. A citrulline/ornithine shuttle between mitochondria and plastids

Glycine (Gly), produced in peroxisomes , is decarboxylated and deaminated by GDC in
the mitochondrial matrix. In contrast to the scheme shown in Figure 4.], GS assimilates
ammonia into glutamate (Glu) directly within the mitochondrial matrix, yielding
glutamine (Gln). Carbamoylphosphate (Caer) is synthesized from Gln and C02 by
carbamoylphosphate synthetase (CPS), thereby regenerating the ammonia acceptor Glu.
Citrulline (Cit) is generated from ornithine (Om) and Caer by Om carbamoyltransferase
(OCT) and Cit is shuttled to plastids were it is converted back to Om and CarP by
catabolic OCT. Carbamate kinase (CK) generates C02, ammonia, and ATP from Caer.
For clarity, the generation of glycine, its conversion of glycine to serine, and the
downstream reaction sequence leading to the conversion of serine to 3-PGA is omitted.
(8) mitochondrial glycine transporter; (9) mitochondrial omithine/citrulline translocator;
(10) plastidic omithine/citrulline translocator. The figure in this dissertation is presented

in color.

141

 

Figure 4.2. A citrulline/ornithine shuttle between mitochondria and plastids

A less costly alternative — the glutamate-glutamine shuttle

Alternatively, a glutamate/glutamine shuttle between mitochondria and plastids can be
envisaged that is compatible with ammonia assimilation by GS2 in mitochondria and that
does not require more ATP than a textbook C2-cycle (Figures 4.1 and Figure 4.3). In this
cycle, ammonia would be assimilated by GS2 in mitochondria, yielding glutamine.
Glutamine is exported from mitochondria, imported into plastids, and the amido group
from glutamine is transferred to or-KG by GOGAT, yielding two molecules of glutamate.
One molecule of glutamate is shuttled back to mitochondria where it serves as substrate
for the GS-reaction. The ATP expense for ammonia assimilation would thus be shified

from plastids to mitochondria, but would remain otherwise unchanged. This hypothetical

142

glutamine/ glutamate shuttle between mitochondria and chloroplasts would require the
presence of glutamine and glutamate transporters in both organelles. The activity of a
glutamine-glutamate antiporter has been described in isolated chloroplasts (Y 11 and Woo,
1988); hence chloroplasts are able to take up glutamate in counter-exchange with export
of glutamine. Relatively little is known about amino acid transporters of plant
mitochondria. It was shown that glutamate enters beetroot and cauliflower bud
mitochondria; the uptake required the presence of both phosphate and a permeable
dicarboxylic anion (Day and Wiskich, 1977). The activity of a glutamate-aspartate
exchanger was demonstrated in isolated pea mitochondria, the partially purified protein
was constituted into liposomes, and capable to facilitate the exchange of glutamate for
both aspartate and glutamate (Vivekananda and Oliver, 1989). Mammalian mitochondria
also possess a mitochondrial glutamate-aspartate exchanger (Palmieri et al., 2001), and in
addition related proteins that catalyze the co-transport of glutamate with H+ (or counter-
exchange with a hydroxyl ion) (Fiermonte et al., 2002). The mitochondrial aspartate-
glutamate carrier from Saccharomyces cerevisiae, unlike its human orthologs, catalyzes
both aspartate-glutamate exchange and substrate uniport (Cavero et al., 2003). It is not
known whether the plant glutamate-aspartate carrier is able to catalyze glutamate uniport
or whether plant mitochondria possess a glutamate carrier. It is also not known whether
glutamine can be transported across the membrane of plant mitochondria; however, in
analogy to mammalian mitochondria, it is likely that plant mitochondria possess a
transport system for glutamine. For example, rat liver mitochondria have a glutamine-
transporter that is able to catalyze both the unidirectional flux of glutamine and

glutamine-glutamine counter-transport (Indiveri et al., 1998). After reconstitution into

143

liposomes, this protein also catalyzes the counter-exchange of glutamine with glutamate
or aspartate, albeit this activity was one order of magnitude lower than glutamine-
glutamine counter-transport (Indiveri et al., 1998). In summary, with the exception of the
mitochondrial glutamine transporter, all components of a glutamate/glutamine shuttle
between mitochondria and plastids have been experimentally verified in leaf cells. In
contrast to the above-described Orn/Cit shuttle, though, the Glu/Gln shuttle only provides
a vehicle the transport of NHB between both organelles, whereas the transport of carbon
dioxide would be dependent on passive diffusion. On the other hand, shuffling NH3
between mitochondria and chloroplasts by the Glu/Gln shuttle requires two ATP less than

by the Om/Cit shuttle.

Safety first - a bucket chain for intracellular ammonium transport

Assimilation of photorespiratory ammonia in mitochondria by GS requires a shuttle
system for ammonia transport between mitochondria and plastids. Both, an
ornithine/citrulline shuttle (Figure 4.2) or a glutamate/glutamine shuttle (Figure 4.3)
could fulfill this function. In both cases, no transport of free ammonia would occur;
instead, this toxic metabolite would be transiently stored as amino acids and would be
transported safely in this form. Future research will be required to examine whether either
of the two alternative shuttles is physiologically relevant (if not both). Likely, the
photorespiratory cycle just became more complex, involving even more metabolite

transporters than previously assumed (Weber, 2005).

144

  
 

 
 

Peroxisomes.

02.
G., H202

  

 

 

Figure 4.3. A glutamate/glutamine shuttle between mitochondria and plastids

In contrast to scheme depicted in Figure 4.1, ammonia generated by GDC in
mitochondria is not assimilated into Glu in the plastids, but by mitochondrial GS and the
resulting Gln is shuttled to the plastid stroma were it is used for Glu biosynthesis fi'om or-
KG by GOGAT. The ammonia acceptor Glu is returned to mitochondria by the same
shuttle system. For clarity, the conversion of glycine to serine, and the downstream
reaction sequence leading to the conversion of serine to 3-PGA is omitted. (1‘) plastidic
glycolate transporter; (2) peroxisomal glycolate transporter; (3) plastidic
glutamate/malate translocator; (4) plastidic or-KG/malate translocator; (5) peroxisomal
glutamate transporter; (6) peroxisomal a-KG transporter; (7) peroxisomal glycine
transporter; (8) mitochondrial glycine transporter; (l l) mitochondrial Glu/Gln
translocator; (12) plastidic glutamate/glutamine translocator. The figure in this

dissertation is presented in color.

145

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mechanisms. In Plant Metabolism (2nd edition) (Dennis, D.T., et al., editors), Addison
Wesley Longman.

Catoni E., Desimone M., Hilpert M., Wipf D., Kunze R., Schneider A., Flugge U.l.,
Schuhmacher K., and Frommer WB. (2003) Expression pattern of a nuclear encoded
mitochondrial arginine-ornithine translocator gene from Arabidopsis. BMC Plant Biology
3: l.

Cavero S., Vozza A., del Arco A., Palmieri, L., Villa L., Blanco E., Runswick M.J.,
Walker J .E., Cerdan S., Palmieri F., and Satrustegui J. (2003) Identification and
metabolic role of the mitochondrial aspartate-glutamate transporter in Saccharomyces
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Coschigano K.T., Melo-Oliveira R., Lim J., and Coruzzi G.M. (1998) Arabidopsis gls
mutants and distinct F d-GOGAT genes: implications for photorespiration and primary
nitrogen assimilation. Plant Cell 10:741-752.

Day D.A., and Wiskich, J.T. (1977) Glutamate transport by plant mitochondria. Plant
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Douce R., and Neuburger, M. (1999) Biochemical dissection of photorespiration.
Orrrent Opinion in Plant Biology 2:214-222.

Fiermonte G., Dolce V., David L., Santorelli F.M., Dionisi-Vici C., Palmieri F., and
Walker J .E. (2003) The mitochondrial ornithine transporter. Bacterial expression,

reconstitution, functional characterization, and tissue distribution of two human isoforms.
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Fiermonte G., Palmieri, L., Todisco S., Agrimi G., Palmieri F., and Walker J.E.
(2002) Identification of the mitochondrial glutamate transporter. Bacterial expression,
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Indiveri C., Abruzzo G., Stipani 1., and Palmieri F. (1998) Identification and
purification of the reconstitutively active glutamine carrier fiom rat kidney mitochondria.
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Hoyos M.B., Palmieri L., Wertin T., Arrigoni R., Polacco J.C., and Palmieri F.
(2003) Identification of a mitochondrial transporter for basic amino acids in Arabidopsis
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Lancien M., Martin M., Hsieh M.H., Leustek T., Goodman H., and Coruzzi G.M.
(2002) Arabidopsis gltI T mutant defines a role of NADH-GOGAT in the non-
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Ludwig RA. (1993) Arabidopsis chloroplasts dissimilate L-arginine and L-citrulline for
use as N source. Plant Physiology 101:429-434.

Melo-Oliveira R., Oliveira LC, and Corn- G.M. (1996) Arabidopsis mutant analysis
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Ogren W.L. (1984) Photorespiration: Pathways, regulation, and modification. Annual
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Palmieri L., Pardo B., Lasorsa F.M., del Arco A., Kobayashi K., Iijima M.,
Runswick M.J., Walker J .E., Saheki T., Satrustegui J., and Palmieri F. (2001) Citrin
and aralarl are Ca(2+)-stimulated aspartate/ glutamate transporters in mitochondria.
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Somerville CR, and Ogren, W.L. (1980) Inhibition of photosynthesis in Arabidopsis
mutants lacking leaf glutamate synthase activity. Nature 286:257-259.

Taira M., Valtersson U., Burkhardt B., and Ludwig RA. (2004) Arabidopsis
thaliana GLNZ-encoded glutamine synthetase is dual targeted to leaf mitochondria and
chloroplasts. Plant Cell 16:2048-2058.

Vivekananda J., and Oliver, DJ. (1989) Isolation and partial characterization of the
glutamate/aspartate transporter fi'om pea leaf mitochondria using a specific monoclonal
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Weber A.P.M. (2005) Synthesis, export, and partitioning of the end products of
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Weber A.P.M., Schneidereit J., and Voll L.M. (2004) Using mutants to probe the in
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Weber A., and Fliigge U.I. (2002) Interaction of cytosolic and plastidic nitrogen
metabolism in plants. Journal of Experimental Botany 53:865-874.

Wingler A., Lea P.J., Quick W.P., and Leegood RC. (2000) Photorespiration:

metabolic pathways and their role in stress protection. Philos. Trans. R Soc. Lond. Ser.
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147

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148

CHAPTER 5

A REVERSE GENETIC APPROACH TO IDENTIFY THE PROPOSED
METABOLITE TRANSPORT PROTEINS IN THE
PHOTORESPIRATORY PATHWAY‘

 

5 Dr. Nicole Linka and Andrea Brautigam contributed to Table 5.3 and Dr. David Gagneul
provided the metabolite data for Figures 5.4, 5.5, and 5.7.

149

ABSTRACT

In C3 plants, the oxygenation reaction of Ribulose-1,5-bisphosphate Carboxylase/
Oxygenase (RubisCO) produces a significant amount of 2-phosphoglycolate, which must
be converted to 3-phosphoglycerate in the photorespiratory pathway. The enzymatic steps
of this pathway are remarkably coordinated although they are distributed in the three
compartments plastids, peroxisomes, and mitochondria. To cope with the high flux of the
intermediates between these compartments during the day, a significant number of
metabolite transporters are required. To date, forward genetic screens have identified
only a single plastidial transporter DiT2.1/DCT1 (dicarboxylate transporter) as a classical
photorespiratory mutant. The mutant is non-viable under ambient air conditions and
indistinguishable from wild type under C02 enriched air. To identify additional
metabolite transporters, publicly available microarray datasets were screened for putative
transporter genes that are co-expressed with fifteen known photorespiratory enzymes. We
identified 32 candidate genes and present the initial genetic analysis Of five genes. One
mutant shows a delayed pale green phenotype after 14 days in ambient air. The
corresponding gene was previously identified as a putative mitochondrial carnitine
carrier, designated BOU (Bout de Souffle). However, our results indicate that instead of
carnitine, BOU presumably imports Vitamin B6 as an essential cofactor required for the

function of the photorespiratory enzyme glycine decarboxylase.

150

INTRODUCTION

Photorespiration (PR) and the Calvin-Benson-Cycle are closely linked pathways via the
bifunctional enzyme ribulose-1,5-bisphosphate carboxylase/oxygenase (RubisCO)
(Ogren, 1984; Tolbert, 1997). Under ambient atmospheric conditions, oxygen (02)
competes with carbon dioxide (C02) for the reaction with ribulose-1,5-bisphosphate
(RuBP) catalyzed by RubisCO. Carboxylation of ribulose-1 ,5-bisphosphate yields two
molecules of 3-phosphoglyerate (3-PGA) that are converted to triose phosphates (TP)
using chemical energy in the form of ATP and NADPH provided by the photosynthetic
light reactions (Calvin, 1962). However, oxygenation yields only one molecule of 3-PGA
and one molecule of 2-phosphog1ycolate (2-PG). The latter molecule represents a
metabolic dead end in the chloroplast stroma. It must thus be removed from the
chloroplast and recycled to 3-PGA in a complex, highly compartmentalized pathway
named photorespiration. In photorespiration, two molecules of 2-PG are recycled to one
molecule of 3-PGA performed in a series of enzymatic steps, which are distributed in
three different subcellular compartments (Figure l; Reumann and Weber, 2006). 2-PG is
hydrolyzed to glycolate in the chloroplast (Somerville and Ogren, 1979; Schwarte and
Bauwe, 2007), and metabolized to glyoxylate and further to glycine in the peroxisomes
(Volokita and Somerville, 1987 ; Igarashi et al., 2003). Glycine oxidation and serine
synthesis takes place in the mitochondria (Somerville and Ogren, 1981; Vol] et al., 2006;
Engel et aL, 2007) and serine is shuttled back to the peroxisomes to be deaminated and
reduced to glycerate (Murray et al., 1989; Liepman and Olsen, 2001). Glycerate is
imported and finally phosphorylated to 3-PGA in the chloroplast (Boldt et al., 2005). The

resulting 3-PGA is then further metabolized in the Calvin-Benson-Cycle. In summary,

15]

two molecules of 2-PG are converted to one molecule each of, 3-PGA, C02, and NH3.
The pathway leads to a net loss of C02 and ammonia and therefore, in addition to a
carbon cycle, photorespiration is intertwined with a nitrogen cycle (Ogren, 1984; Keys,
2006). Ammonia (NH3) is released during glycine oxidation in the mitochondria (Keys et
al., 1978). It has been widely accepted that NH3 enters the chloroplast and is re-
assimilated by the concerted action of glutamine synthetase (GS) and glutamate synthase
(GOGAT) (Somerville and Ogren, 1980; Coschigano et al., 1998; Wallsgrove et al.,
1987). The product glutamate is guided to the peroxisomes and directly utilized to
produce glycine fiom glyoxylate — In this way, a new round of nitrogen cycling is
initiated. The flux of NH3 re-assimilation exceeds by far the rate of de novo ammonium
ion production and has been demonstrated to be absolutely required under
photorespiratory conditions to circumvent toxic glyoxylate accumulation (Sharkey, 1988;
Keys, 2006).

It has been recemly demonstrated that GS is dually targeted to chloroplasts and
mitochondria (T aira et al., 2004). This would allow the re-assimilation of the NH3 into
glutamine on location. Still, the enzyme GOGAT is exclusively active in the chloroplast
(Coschigano et al., 1998) and glutamine instead of ammonia has to be transported (Linka
and Weber, 2005). In another scenario, ammonia could be incorporated into carbamoyl
phosphate and subsequently into ornithine yielding citrulline in the mitochondria.
Citrulline is broken down in the chloroplast to ornithine and ammonia enters the
GS/GOGAT cycle (Ludwig, 1993; Linka and Weber, 2005).

The identification of enzymes that are required for conversion of 2-PG to 3-PGA

has been one of the earliest and, in retrospect, most successful forward genetic screens in

152

the Arabidopsis thaliana research field (Somerville and Ogren, 1979; Somerville, 2001).
Classical photorespiratory mutants are indistinguishable from wild-type plants in C02
enriched air but not viable under ambient air conditions (Somerville and Ogren, 1979).
PR mutants significantly contributed to establishing A. thaliana as a model organism for
plant biochemical geneticists and plant physiologists (Meyerowitz and Pruitt, 1985).
Almost three decades later, the enzyme glycerate-3-kinase as the last enzymatic step in
the series was identified on the molecular level and published as the “last unknown
enzyme in the photorespiratory cycle” (Boldt et al., 2005). As outlined above,
photorespiration is a prime example for pathway compartrnentalization in eukaryotic
cells, since enzymes for the carbon and nitrogen cycle are partitioned almost in equal
parts in chloroplasts, peroxisomes and mitochondria (Kunze et al., 2002). Despite the
excellent progress in understanding the photorespiratory pathway, none of the metabolite
transporters navigating the photorespiratory intermediates through the cell are mapped on
the genome of A rabidopsis thaliana (Reumann and Weber, 2006). Forward genetic
screens identified only one afi'rliated transport system: the malate/ glutamate transporter
DiT2.l/DCT1 (dicarboxylate transporter; Somerville and Ogren, 1983; Renne et al.,
2003). This exports the amino-donor glutamate fi'om the chloroplast for the
glyoxylate: glutamate aminotransferase (GGT) reaction in the peroxisomes as described
earlier. The released product a-ketoglutarate (or-KG) has to be shuttled back to the
chloroplast and converted to glutamate by the GOGAT reactions (Weber and Flugge,
2002)

The list of A. thaliana photorespiratory mutants has been continuously expanded.

For example the peroxisomal catalase detoxifies H202 released by the glycolate oxidase

153

(Queval et al., 2007) and malate dehydrogenases sustain the NADH pool via an
oxaloacetate/malate redox-system for the reduction of hydroxypyruvate to glycerate
(Cousins et al., 2008). The mitochondrial glycine decarboxylase is a multienzyme
complex consisting of four different subunits (P protein, H protein, T protein, and L
protein) and several cofactors [lipoic acid, tetrahydrofolate (T HF), NAD, FAD, and
pyridoxal-phosphate (PLP)] (Douce et al., 2001). A null mutant for the P protein subunit
is inviable and reduced GDC activities result in classical PR phenotypes (Heineke et al.,
2001; Engel et al., 2007). It has been demonstrated that impaired biosynthesis of the
cOfactor lipoic acid (Ewald et al., 2007) or THF recycling in mitochondria causes a
decrease in GDC activity (Collakova et al., 2008). Import of malonic acid and pyruvate is
a prerequisite for lipoic acid synthesis and the three precursors glutamate, pABA (para-
aminobenzoic acid) and pterin have to be provided for THF synthesis inside the
mitochondria (W ada et al., 1997; Rebeille et al., 2007).

Apparently, primary photorespiratory metabolites as well as a great number of
additional compounds have to be distributed within the cell to cope with the high
photorespiratory rates timing the day. In this paper we attempted to establish a catalog of
metabolite transporter candidates by a reverse genetic approach. Routine transcriptional
profiling is providing an enormous data set, which is becoming a valuable tool to
associate proteins of unknown ftmction to known pathways. We used the comprehensive
system-biology database (CSB.DB) to identify the best 5% of transcripts co-regulated
with a set of 15 highly co-expressed photorespiratory genes. Candidates, which are co-

expressed with several PR genes and contain at least two predicted hydrophobic

154

transmembrane spans, were selected for genetic analysis to identify classical

photorespiratory mutants.

MATERIAL AND METHODS

Chemicals

The radiolabeled compounds [l4C(U)]-arginine, [l4C(U)]-g1utamic acid, [14C(U)]-
glycine, [l4C(U)]-L-serine, [N-methyl-14C]-L-camitine, [2-14C]-malonic acid and [1-
14C]-L-ornithine were purchased from Moravek Radiochemicals and Biochemicals
(Brea, CA, USA). Sigma-Aldrich (St. Louis, MO, USA) supplied the chemicals and the
company Bio-Rad (Hercules, CA, USA) provided the Bio-Beads SM-2 hydrophobic
adsorbent and the ion exchange resins Dowex AGl-X8 and AG50-X8. Reagents and
enzymes for nucleic acid amplification, purification and cloning were purchased fiom
Promega (Madison, WI, USA), Qiagen (Hilden, Germany) and Invitrogen (Carlsbad, CA,

USA).

Plant Material and Growth Conditions

Seeds of the A. thaliana ecotype Columbia-0 and T-DNA insertion lines for At1g54350
(SALK003 891 & SALK069087), At2g42770(SALK118636), At2g48070
(SALK066383) and At3g26710 (SALK055566) were obtained fiom the Arabidopsis
Biological Resource Center (ABRC) at Ohio State University (USA). The A. thaliana
GABI-Kat line 079D12 was ordered from the Center for Biotechnology (CeBiTec),

Bielefeld University (Germany) and the SHMT (shmI) line was previously established in

155

our lab (V 011 et al., 2006). Surface-sterilized seeds were germinated on petri dishes
containing half-concentrated MS medium supplemented with 1% (w/v) sucrose and 0.8%
agar and plantlets were transferred on soil after two weeks and watered regularly with
Hoagland solution. Unless stated otherwise, the plants were kept under a 12 h-light

(120 uEinstein per square meter and minute)/ 12 h-dark regime at 22°C in growth
chambers with elevated C02 concentrations (0.3%) throughout their complete life cycle.
Selection of the transgenic A. thaliana plants was performed with kanamycin (50 ug/mL)

or hygromycin (30 ug/mL).

Transcript co-response analyses

The publicly available comprehensive system-biology database (CSB.DB) (Steinhauser
et al., 2004) was used to identify genes transcript, co-expressed with 15 genes known to
be involved in the photorespiratory pathway in A. thaliana. AtGenExpress matrices
atge0100 and atge0200 comprised of 63 experiments with 12,200 genes of the
developmental expression series, and the abiotic stress series (abovegromrd samples) with
13,197 genes, respectively (Schmid et al., 2005), were analyzed. The NASC Arrays
database (Nottingham Arabidopsis Stock Centre) with microarray experiments from the
matrix nasc02 71 was also included (Craigon et al., 2004). Based on the Spearman’s
correlation, the best 5% of co—responding genes from each single gene query were further
analyzed with the ARAMEMNON database to identify putative membrane proteins with
at least two predicted alpha helix transmembrane spans (Schwacke et al., 2003;

Steinhauser et al., 2004).

156

Phylogenetic tree analysis

The putative mitochondrial carnitine carrier, designated BOU (Bout de Souffle), belongs
to the eukaryotic mitochondrial carrier family (MCF). Homologous sequences of the
MCF from yeast and human and Arabidopsis thaliana were obtained from the National
Center for Biotechnology Information (NCBI) (mzllwwwncbinlmnihggv). The
BLAST program at NCBI (Altschul et al., 1990) identified homologous genes in the
genome of the red alga Galdieria sulphuraria (http://genomics.msu.edu/galdieria_/1 using
the human and yeast proteins as a query with a cut-off expectation value (E-value) of

_<_ 0.0001. The phylogenetic analysis was performed with full-length amino acid

sequences of 208 proteins according to Weber et al., 2006.

Isolation of homozygous T-DNA insertion lines

Genomic DNA was isolated and T-DNA insertions were determined by standard PCR
with the following primers: T-DNA specific primer LBal and ML116 were used to
amplify fiom the T-DNA left border in SALK lines and the GABI-Kat line, respectively,
and the At3g] 8780 (AtACT2)-specific primer pair ML40/ML41 served as a quality
control. At1g54350 gene-specific primer pair ML102/ML103, At2g42770 gene-specific
primer pair ML104/ML105, At3g26710 gene-specific primer pair ML108/ML109,
At2g48070 gene-specific primer pair ML107/ML]20, and At5g46800 gene-specific
primer pair ML114/ML115 were designed to analyze the transgenic lines. Primer
sequences are shown in Table 5.1. Amplified PCR fi'agments were cloned into the
pGEM-T Easy vector (Promega) and the T-DNA insertion sites were verified by

sequencing (Research Technology Support Facility, Michigan State University, USA).

157

Homozygotes were further propagated and heterozygous mutants were self-pollinated

and analyzed again for homozygous T-DNA insertion.

Isolation of total RNA and RT-PCR analysis

The method by Chomczynski and Sacchi (1987) was used to extract total RNA from 6-
weeks old A. thaliana rosettes. After DNAse treatment, cDNA was synthesized
(Superscript II F irst-Strand Synthesis System, Invitrogen). The transcript levels were
examined from the putative metabolite transporter BOU (At5g46800) and the control
gene AtACT7 (At5g09810) by PCR analysis. The PCR reaction was an initial
denaturation step at 94°C for 2 min, followed by 35 cycles of 94°C for 35 s, 58°C for 30
s, 72°C for 2 min, and final extension of 72°C for 3 min and products were visualized on

an ethidium brornide-stained, 1% agarose gel.

Table 5.1. Primer sequences used in Chapter 5

Primers were used for cloning of heterologous expression (HE) constructs, Reverse-
Transcriptase PCR (RT-PCR) expression analyses, complementation studies (COMP) or
T-DNA insertion line screens (TDNA). Underlined nucleotides highlight restrictions sites
used for cloning.

 

Primer Sequence (5'—3')

 

LBal TGGTTCACGTAGTGGGCCATCG
ML 40 GTTGGGATGAACCAGAAGGA
ML 41 GAACCACCGATCCAGACACT
ML 90 TATAGQAICCAAATGGCGGATGCGTGGAAG
ML 91 ATATCTCGAGTTTATCCCAAGCTTGACCT
ML 92 TATACflIACQACCATGGCGGATGCGTGGAA
ML 93 ATATIQI‘AQATCCCAAGCTTGACCTTGTCA

sea
asssgggg

 

158

Table 5.1. (cont’d).

 

 

Primer Sequence (5'—3') Purpose
ML 102 TAGCT CTT CT GATCCACGGAG TDNA
ML 103 TCAATGGTGGCATTAACAAGC TDNA
ML 104 CCCAACACCCACAAGTAGTTG TDNA
ML 105 TTTCC'ITTGAAGCAAGCTGTG TDNA
ML 107 AGAGAAAGAAACCAACCTGCC TDNA
ML 108 TTCGACCTGAACATCATCTCC TDNA
ML 109 TTGGTCTGATTTTATGTCGGG TDNA
ML 114 CAAAGGTTTTGATI'TTGGTTTG RTPCR
ML 115 CAAACCCC'ITAGCGAACTCTT TDNA
ML 116 CCCATTTGGACGTGAATGTAGACAC TDNA
ML 120 TTGCCTTAAACCCACGTACC TDNA
ML 143 CGCACAATCCCACTATCCTT TDNA
ML 164 CAATGCGGTTTTG'ITCACTG RT-PCR
ML 165 CCCAAGCTTGACCTTGTCAT RT-PCR
ML 166 AAAAGAAGCACCAGCGACTC RT-PCR
ML 167 TTCAATGTCCCTGCCATGTA RT-PCR
ML 168 TGAACAATCGATGGACCTGA RT-PCR
ML 175 TACQAILKECGGATGCGTGGA COMP
ML 176 ATM TCCCAAGCTTGACCTTGTC COMP
ML 180 CACACICIAQAAGCCGGATCTCAGTG HE
ML 181 CACACGAGATATACCCATfiCGGATG HE

 

159

Complementation of the GABI-Kat line 079D12

For the molecular complementation of the T-DNA insertion line GABI-Kat 079D12
(bad) the hill-length cDNA of BOU (At5g46800) was amplified by PCR with the
primer pairs ML175 and ML176 (Table 5.1), cloned into pGEM-T Easy (Promega) and
verified by sequencing. The BOU gene was excised with the designed restriction sites
Nhe I and Nco 1 and ligated into the plant expression vector pCAMBlA 1302 after
removal of the GFP reporter gene with the same restriction enzymes. Homozygous bouI
plants were transformed by the Agrobacterium-mediated floral dip method (Clough and
Bent, 1998) and several independent lines were selected on MS agar plates containing
20 ug/mL hygromycin. The 35S:BOU construct in the T1 generation was confirmed on
the genomic DNA level by PCR with the primers ML143/ML165 and BOU transcripts
were analyzed with the primers ML164/ML165 and ML114/ML166. The control
transcripts AtACT7 were amplified with the primer ML167 and ML168 (Table 5.1). T2
generations were screened by segregation analysis and homozygous lines were
established in the following generation. The standard PCR reaction was an initial
denaturation step at 94°C for 2 min, followed by 35 cycles of 94°C for 35 s, 58°C for 30
s, 72°C for 2 min, and final extension of 72°C for 3 min and products were visualized on

an ethidium bromide-stained, 1% agarose gel.

Antibody production and protein analysis
To use BOU as an antigen and for biochemical characterization, the protein was
expressed in Escherichia coli strain BL2] Codon Plus (Stratagene, La Jolla, USA).

Coding sequence was amplified with the primers ML180/ML181 (Table 5.1) by PCR

160

fi'om cDNA templates. The fiagment was subcloned into pGEM-T Easy (Promega) and
via Nde I and Barn HI restriction sites introduced into the pET16b vector (Merck
Chemicals Ltd., Darmstadt, Germany). Expression was induced with 1 mM IPT G
(isopropyl B-D-l-thiogalactopyranoside) at an OD600 of 0.6 and cells were harvested
after two hours. Inclusion bodies containing recombinant BOU protein translationally
fused with an N-terminal hexahistidine tag were isolated according to Schroers et al.,
1997, and further purified using nickel-affinity chromatography under denaturing
conditions (Qiagen). In detail, 100 mL cultures of E. coli cells were harvested,
resuspended in 10 mL Buffer A (20 mM TRIS/HC] pH 7.8, 1 mM EDTA. 1 mM PMSF),
incubated with lysozyme (1 mg/mL) and burst by a subsequent sonication step. The
insoluble inclusion bodies were pelleted by centrifugation (12,000 g, 10 min, 4°C),
dissolved by sonication in buffer A and centrifuged again at 1,100 g for 5 minutes. The
supernatant (containing the inclusion bodies) was transferred to a new tube, and inclusion
bodies were collected by centrifugtion (12,000 g, 5 min, 4°C), and washed three times
with a buffer containing 2% (v/v) Triton X-100, 20 mM TRIS/HC] pH 7.8, 0.1 mM
EDTA. The inclusion bodies were solubilized in 400 uL SLS buffer [1.67 % (w/v)
sodium lauroylsarcosine, 20 mM TRIS/HCl pH 7 .8, 0.1 mM EDTA] and diluted with
800 uL deionized water. The expression and purification steps were colorimetrically
confirmed by Coomassie-stained SDS-PAGE and by immunoblot analysis using a
monoclonal anti-penta-His antibody (Qiagen, Germany) combined with an alkaline
phosphatase (AP)-conjugated secondary antibody (Promega). Cocalico Biologicals Inc.
(Reamstown, PA, USA) produced the polyclonal antibodies against the purified BOU in

rabbits. Polyclonal antibodies raised against lipoic acid, H- and P-protein subunits of

161

glycine decarboxylase (GDC), and serine-hydroxymethyl-transferase (SHMT) were
kindly provided by Prof. Bauwe from the University of Rostock (Germany). For the
protein analysis, 5 g of 4-5 weeks old A. thaliana rosettes were harvested and
mitochondrial fiactions were prepared as described by Keech et al., 2005. Fifty
micrograms of enriched mitochondrial proteins were resolved on a 15% SDS-PAGE,
immobilized on PVDF membranes (Bio-Rad) and analyzed with the Amersham ECL
(Enhanced Chemiluminescence) Western Blotting detection system with a secondary
anti-rabbit antibody conjugated to HRP (horseradish peroxidase, GE Healthcare UK
Limited, Buckinghamshire, UK). Protein concentrations were determined with the Pierce

BCA protein assay (Pierce Biotechnology, Rockford, IL, USA).

Metabolite analysis

For time course experiments, mutant and wild type plants were grown for four weeks
under high C02 concentrations (0.3% C02) and transferred to ambient air

(0.038% CO2). Entire rosettes were harvested prior to and two, four and seven days after
the shift. Rosette leaves from control plants, which were kept at 0.3% C02, were sampled
in parallel. Experiments were set up in a completely randomized manner. Start of the
experiment and sampling were done at midday. Shoots were collected, immediately
frozen, and subsequently powdered in liquid nitrogen, and stored at -80°C tmtil firrther
processing. Glyoxylate and ammonium were extracted and quantified according to
Brautigam et al., 2007. For proteinogenic and non-proteinogenic amino acid analyses,
100 mg fresh tissue were extracted with 700 RL pre-chilled 100% methanol (-10 to -

20°C) supplemented with phenylalanine-d5 20 rtM (internal standard for amino acid

162

profile normalization). Samples were shaken at 70°C for 15 min. After cooling on ice,
700 uL de-ionized water was added to the mixture. Subsequently, 375 uL pre-chilled
chloroform was added and the samples were shaken again for 30 min at 4°C. The samples
were centrifuged for 10 min at 14,000 g, and the upper, aqueous phase was used for
metabolite analysis. Amino acids were characterized and quantified by LC-MS/MS using
a Waters Symmetry C18 column (Milford, MA) with a Shimadzu LC-20AD HPLC
system (Columbia, MD) coupled to a Waters Quattro micro mass spectrometer (Gu et al.,
2007). Amino acid concentrations were calculated by making reference to external
calibration curves, and recovery of successive procedures was assessed using the internal

standard phenylalanine-d5.

Measurement of glycine decarboxylase activity

Mitochondria fi'om wild type and ban] mutant plants were isolated as described above
(Keech et al., 2005) except that the final mitochondria pellet was resuspended in 500 uL
of 50 mM TES/KOH (pH 7 .5), 2 mM EDTA, 5 mM MgC12, 30% (v/v) glycerol and
0.1% (v/v) Triton X-100 to permeabilize the membrane and release the matrix proteins.
The 100 RL activity assay contained 50 uL mitochondrial preparation and final
concentrations of 50 mM TES/KOH (pH 7.5), 1 mM EDTA, 2.5 mM MgC12, 15% (v/v)
glycerol and 0.01% (v/v) Triton X-100, 2 mM B-mercaptoethanol, 0.03 mM pyridoxal-
phosphate (PLP), 0.5 mM tetra-hydrofolate (T HF), 1 mM NAD+ and 0.2 mM [l4C(U)]-
glycine. The reactions were conducted in an airtight vial. After 15 min the reaction was
terminated by the addition of 1 M sulfirric acid (0.5 mL). GDC activity was linear for at

least 30 minutes. The released [14C]-C02 was transferred with a stream of nitrogen gas

163

into a new vial filled with 5 mL of a 1 M KOH solution to trap the carbon dioxide as
hydrogen bicarbonate (Walker and Oliver, 1986). The released [l4C]-C02 was
quantified by liquid scintillation counting (LSC) with a Beckrnan LS6000 counter

(Beckman Coulter Inc., Fullerton, CA).

Heterologous expression of BOU in Saccharomyces cerevisiae

To clone BOU into the yeast expression system pYESZ/NT (Invitrogen) the full-length
sequence was amplified from cDNA by PCR as described earlier. The primer pairs
ML90/ML91 and ML92/93 (Table 5.1) were used to yield recombinant BOU protein with
an N- and C-terminal hexa-histidine epitope. The fiagment obtained with the first primer
set was digested with Barn HI and Xho I and cloned into the pYES/NT-A vector. For the
C-terminal His-tag fusion, the plasmid pYES/NT-A was digested with Hind III, treated
with T4 polymerase (Promega) and subsequently incubated with Xba I. The BOU
fragment was processed consecutively with the enzymes Kpn 1, T4 polymerase and Xba I
and ligated with the prepared pYES/NT-A vector. Transformed INVScl yeast
(Invitrogen) cells were selected and the recombinant protein was expressed and partially
purified as described by Linka etal., 2008. Production of the BOU protein was confirmed
by immuoblot analysis using an anti-His antibody (Qiagen, Germany) and an AP-

conjugated secondary antibody (Promega).

Reconstitution into liposomes

Two methods were used in this study to functionally reconstitute recombinant BOU

protein into liposomes. For the freeze-thaw-sonicate procedure (Palmieri etal., 1995), a

164

2.5% (w/v) Lon-phosphatidylcholine (PC) suspension was prepared in 120 mM
TRIS/BC] (pH 8.0) with 30 mM of the substrate or in 150 mM TRIS/HC] (pH8.0). The
phospholipids were dissolved by sonication (Branson Sonicator 250, output 2, duty cycle
30%) on ice for three minutes. 50 rtL of the yeast extract or 30 pg of the recombinant,
solubilized BOU protein were mixed with the liposome suspension to a final volume of

1 mL. The protein-phospholipid mixtm'e was frozen in liquid nitrogen and thawed at
room temperature. To obtain unilamellar and sealed vesicles, the suspension was
sonicated with 25 pulses (output 2, duty cycle 30%). Alternatively, in the “cyclic
detergent removal procedure” (Kramer and Heberger, 1986), 30-50 ug of the
recombinant BOU protein was mixed with 110 uL 10% Triton X-100. A 10% (w/v) PC
suspension was prepared in deionized water and 160 RL was added to the .
protein/detergent mixture. The concentrated micelles were diluted to a volume of 1 mL
with TRIS/HC] (pH 7.8) and the putative substrate to a final concentration of 120 mM
and 30 mM, respectively. After an incubation of 15 min on ice, the detergent was
gradually removed by passing the mixture twelve times over a column filled with 1 g of a
hydrophobic absorbent (Bio-Beads SM-2, Bio-Rad). External substrate was removed by
an additional chromatography step using a desalting Sephadex PD-10 column (GE

Healthcare), previously equilibrated with 150 mM TRIS/HCl (pH 7.8).

Transport assay

W: Uptake of 0.1 mM [N-methyl-14C]-L-camitine into

proteoliposomes was tested with and without preloading (L-carnitine; 30 mM). After

165

defined time intervals (up to 30 minutes) the vesicles were passed through a PD-lO
column to separate the external radiolabeled substrate from the vesicles. Liposomes were
collected from the flow-through and the imported [N-methyl-l4C]-L-carnitine was
quantified by LSC.

Glutamate and malonic acid transmit assay: Liposomes were prepared with 30 mM
glutamate, glutamine or malonic acid and tested for the uptake of radiolabeled [l4C(U)]-
glutamic acid (0.75 mM) or [2-14C]-malonic acid (0.25 mM), respectively. External
radiolabeled substrate was removed by binding to an anion-exchange column (Dowex
AGl-X8 Resin acetate form, 100-200 mesh, Bio-Rad), which had been pre-equilibrated
with 10 mM TRIS/HCl (pH 7.8) and 140 mM sorbitol. The proteoliposomes-specific
radioactivity was quantified by LSC.

WW: Putative antiport and uniport transport activities
of BOU with the substrates arginine, citrulline and ornithine were investigated. In both
cases the experiments were initiated with 0.25 mM [14C]-radiolabeled substrate and
stopped by cation-exchange chromatography (AG50W-X8 Resin, 100-200 mesh,
BioRad). Prior to the experiment, the hydrogen ion activated resin columns were
converted to its lithium form by washing the resin with five volumes of 1 M lithium
acetic acid (LiAc). Due to the greater affmity of arginine and ornithine compared to
lithium-ions, both substrates could be efficiently removed fi'om the external medium.
9W: As described above, proteoliposomes were produced
in the presence of 150 mM Tris/KOH (pH 7.8) or with 120 mM Tris/KOH (pH 7.8) and
either 30 mM glycine or serine. A concentration of 0.25 mM [l4C(U)]-L-serine and

[14C(U)]-glycine were used to initiate the transport assay. External radiolabeled serine

166

and glycine were removed from the uptake medium by anion-exchange chromatography.
Because the affinity of the resin was rather low for these substrates, the hydroxyl ion of
the anion-exchange resin AGlW-X8 was replaced with the bufler molecule tricine.
Liposomes were eluted from the column with 10 mM Tris/HC] (pH 7.8) and 140 mM
sorbitol and quantified by LSC. To study the dependence of glycine and serine import on
an electrochemical gradient across the membrane, artificial pH and potassium (K+)
gradients were created in the presence of the ionophores nigericin (0.1 pig/mg PC) and
valinomycin (1.5 rig/mg PC), respectively, as described by Indiveri et al., 1997.
Alternatively, internal or external pH values were adjusted with the buffer compormd
MES/KOH (pH 6.0) to analyze the effect of serine and glycine transport by a pH gradient

across the liposomal membrane.

Giant liposomes preparation and Patch-Clamp recording

Recombinant BOU protein was produced in E. coli and inclusion bodies purified as
described above. 100 uL of solubilized protein was diluted with 1.4 mL resin buffer

[0.1 M sodium phosphate buffer (pH 8), 0.3 mM sodium chloride, 0.0] mM irnidazole,
2% (v/v) Triton X-100]. Nickel-NTA agarose (Qiagen) was added (100 uL) and gently
shaken for one hour at 4°C. The agarose beads were pelleted (1,000 g, 0.5 min, 4°C), the
supernatant discarded, and the beads were washed five times with resin buffer. The
protein was eluted with 1 mL resin buffer containing 250 mM imidazole. The initial
mixture for functional reconstitution contained 10 ug protein, 0.25% (v/v) Triton X-100,
0.35% (w/v) PC, 0.1 M KC] and 0.02 M MOPS/KOH (pH 8). Proteoliposomes were

prepared by hydrophobic chromatography (Kramer and Heberger, 1986). To yield giant

167

liposomes suitable for patch-clamp measurements, the vesicles were centrifuged at
100,000x g for 1 hour at 4°C and resuspended in 40 RL MOPS/KOH (pH 7.4) containing
5% (v/v) ethylene glycol. Half of the liposomes were partially dehydrated on a
microscope slide in a desiccator containing anhydrous CaC12. After 3 hours the
proteoliposomes were rehydrated with 20 uL 0.1 M KC1 in a humidified Petri dish
overnight to form giant liposomes (Criado and Keller, 1987; Keller et al., 1988; Delcour
et al., 1989; Riquehne et al., 1990). Patch-pipettes were prepared fi'om 1.5 mm
borosilicate glass with a P-87 F laming/Brown mircopipette puller (Sutter Instrument
Company, Novato, CA). A Nikon Eclipse TE2000-S microscope (Nikon Instruments
Europe B.V., Amstelveen, The Netherlands) and the Eppendorf PatchMan NP2
micromanipulator (Eppendorf AG, Hamburg, Germany) was used to achieve gigaohm
seal formation with the giant liposomes. Patch-clamp recording was performed using the
HEKA EPC10 patch clamp amplifier combined with the PatchMaster software (HEKA

Instruments Inc. Bellrnore, NY).

RESULTS

Putative metabolite transporter co-expressed with the photorespiratory pathway
The publicly available Comprehensive System-Biology DataBase (CSB.DB) (Steinhauser
et al., 2004) provides a statistical tool to scan transcript profiles fi'om the NASC’s
Affymetrix facility (matrix nasc02 71; Craigon et al., 2004) and from the AtGenExpress
consortium (matrix atge0100 and atge0200, Schmid et al., 2005) for co-expressed genes

in the model organism A. thaliana. In particular, we were interested in the enzymes

168

participating in the recycling of 2-phosphog1ycolate to 3-phosphoglycerate. We showed
that with the exception of glycerate kinase (GLYK), catalase (CAT2), and the subunits
L1 and H2 of the glycine decarboxylase, all known photorespiratory genes form a robust
cluster of 15 transcripts (data not shown) and thus represent a highly co-regulated
metabolic pathway. We used the CSB.DB platform to retrieve additional candidates that
co—respond with the strong gene cluster. Based on the Spearman's rank correlation
coefficient (SRCC) the best 5% of each of the 15 individual gene queries from each of
the three microarray matrices (atge0100, atge0200, and nasc02 7 I ) were screened for
hydrophobic membrane proteins with at least two putative alpha helix transmembrane
regions using the plant membrane database ARAMEMN ON (Schwacke et al., 2003).
These proteins represent candidate metabolite transporters, which could enable the high
flux of metabolites between the subcellular compartments during photorespiration.
Initially, 162 different hydrophobic proteins were identified in the top 5% of positively
co-regulated transcripts. 32 out of the 162 hydrophobic genes Show at least 30 out of the
45 possible co-responses with the fifteen query genes (Table 5.2) in the three microarray

matrices, and are summarized in Table 5.3.

169

Table 5.2. Co-expressed photorespiratory enzymes from Arabidopsis thaliana
Fifieen out of twenty known photorespiratory genes are highly co-regulated.

 

 

AGI Gene Description Localrz' ation Reference
At5g36700 2-Phosphoglycolate Phosphatase Chloroplast Schwarte and
(PGLP l) Bauwe (2007)
At5g35630 Glutamine synthetase Chloroplast Somerville et a1.
(GS2/GLN2) (1980)
At5g12860 2-Oxoglutarate:malate antiporter Chloroplast Tanigushi et a1.
(DiTl/OMT 1) (2002)
At5g64290 Glutamatezmalate antiporter Chloroplast Renne et a1.
(DiT2.l/DCT1) (2003)
At3g14420 Glycolate oxidase Peroxisomes Volokita and
(GOX2) Somerville
(1987)
At1g23310 Gluzglyoxylate aminotransferase Peroxisomes Igarashi et a1.
(AOATl/GGAT) (2003)
At2gl3360 Ala/Ser:glyoxylate Peroxisomes Liepman et a].
aminotransferase (AGT] /SGT 1) (2001)
At] g68010 Hydroxypyruvate reductase Peroxisomes Murray et a1.
(HPR) (1989)
At2g35370 Glycine decarboxylase Mitochondria Bauwe et a].
H protein (GDHl) (2003)
At1g32470 Glycine decarboxylase Mitochondria Bauwe et a].
H protein (GDH3) (2003)
At1g48030 Glycine decarboxylase Mitochondria Bauwe et a].
L protein (LPD2) (2003)
At4g33010 Glycine decarboxylase Mitochondria Engel et a].
P protein (GDPl) (2007)
At2g26080 Glycine decarboxylase Mitochondria Engel et a].
P protein (GDP2) (2007)
At1g11860 Glycine decarboxylase Mitochondria Bauwe et a].
T protein (GDTl) (2003)
At4g3 7930 Serine hydroxymethyltransferae Mitochondria Vol] et a1.
(SHMT 1) (2006)

 

170

Table 5.3. Putative transporter genes, which are transcriptionally co-regulated with the
photorespiratory pathway

The comprehensive system-biology database was used to identify the top 5% of genes,
which are co-expressed with 15 genes of the photorespiratory pathway (Table 5.2).
Putative membrane proteins and transmembrane spans were extracted from this list using
the ARAMEMNON database (Schwacke et al., 2003). Genes highlighted in bold were

firrther tested in this study. Subcellular localization was experimentally demonstrated
(Sun et a1, 2008; Haezlewood et al., 2004) for those proteins with an asterisk (*).

 

 

Candidate Gene annotation Putative Predicted
gene transmembrane subcellular
spans localization
1 At1g10830 Sodium symporter-related 6 Chloroplast/
Mitochondria
2 At1g22850 Hypothetical protein 4 _5 Chloroplast
3 At] g32080 putative membrane protein (MEPl) 12 Chloroplast“
(Envelope)
4 At1g44920 Hypothetical protein 4 Chloroplast“
5 At] g5 0730 Hypothetical protein 2 Mitochondria
6 At] g52870 Putative peroxisomal PMPZZ-type Chloroplast/
protein of unknown function 2 Mitochondria
7 Atlg54350 Putative plastidial ABC-type Chloroplast"
transporter (AtPMPI) 5 (Envelope)
8 At] g7 8180 Putative mitochondrial carrier 3 5 Chloroplast!
protein - Mitochondria
9 At2g42770 Putative-peroxisomal PMP22-type 2 Chloroplast“
protern of unknown function (Envelope)
10 At2g48070 Protein of unknown fimction 3 _4 Chloroplast
function 10
12 At3 g2 15 80 Hypothetical protein 6 Chloroplast
l3 At3 g25805 Hypothetical protein 7 Chloroplast/
Mitochondria
14 At3g26710 Hypothetical protein 4 Chloroplast“
(Thylakoid)
15 At3g48200 Hypothetical protein 9 Secretory
pathway
16 At3 g50685 Hypothetical protein 3 Chloroplast
17 At3 g5 5250 Hypothetical protein 2 Chloroplast/
Mitochondria
18 At3g56160 Putative bile acid:sodium symporter 7 Chloroplast

 

171

Table 5.3. (cont’d).

 

 

Candidate Gene annotation Putative Predicted
gene transmembra subcellular
ne spans localization
19 At3g59780 Protein of unknown function 3 Chloroplast/
Nucleus
20 At3g61870 Hypothetical protein 5 Chloroplast
21 At4g02920 Hypothetical protem 2 Unknown
22 At4g] 641 0 Hypothetical protein 3 Chloroplast/
Mitochondria
23 At4g24090 Hypothetical protein 3 Chloroplast
24 At4g24460 Protein of unknown function 10 Chloroplast
25 At4g35760 Hypothetical protein 4 Chloroplast
26 At5g13720 Hypothetical protein 3 Chloroplast
27 At5g27290 Hypothetical protein 3 Chloroplast/
Mitochondria
28 At5g38660 APE] (Acclirnation of *
Photosynthesis to environment) - 2 Chloroplast
unknown function
29 At5g42130 Putative mitochondrial carrier 3 6 Chloroplast‘
protein ' (Envelope)
30 At5g46800 Putative mitochondrial carrier Mitochondria“
protein (BOU) 3-6
31 At5 g5 7345 Protein of unknown fimction Chloroplast/
2-3 . .
Mrtochondna
32 At5g64970 Putative mitochondrial carrier 3 6 Mitochondria“
protein - + chloroplast“

 

172

Phenotype of T-DNA insertion lines from candidate genes

Classical mutants in the photorespiratory pathway grow asymptomatically under elevated
levels of carbon dioxide (C02); however, they turn chlorotic under ambient air
conditions (Somerville, 2001). To test the hypothesis that the 32 candidate genes are
involved in the photorespiratory pathway, T-DNA insertion lines for several candidate
genes were established and their ability to grow under ambient C02 levels were
evaluated. To this end, homozygous T-DNA exon insertion lines were established for the
putative plastidial ABC-type transporter At1g54350 (pmpI-I & pmpI-Z) and the putative
peroxisomal PMP22-type protein of unknown function At2g42770 (SALK118636)
(Figure 5.1). Germination under elevated C02 levels for ten days and shifting the
seedlings to ambient air for two weeks induced the photorespiratory phenotype in the
known serine-hydroxymethyltransferase (SHMT) mutant shmI-Z, which was used as a
positive control (Figure 5.2; Vol] et al., 2006). In contrast, mutant lines pmpI-I, pmpI-Z
and SALK118636 had no obvious grth phenotype compared to A. thaliana wild type
Columbia-0 plants (Figure 5.2). Seeds had also been germinated under ambient air
conditions and in the presence and absence of sucrose as an external carbon source.
Under all conditions tested, the three mutant lines were indistinguishable from the wild
type (data not shown).

Heterozygous T-DNA insertion lines SALK055566 (At3g26710, hypothetical
protein) and SALK066383 (At2g48070, protein of unknown function) were self-fertilized
(Figure 5.1) and repeatedly screened for a T-DNA linked homozygous genotype. In both
populations, only heterozygous and wild type-like lines germinated and developed

normally, independent of the C02 concentration in the growth chamber (data not shown).

173

Seedlings from homozygous T-DNA insertions lines of these genes developed white
cotyledons and did not establish any new true leaves (Figure 5.2). An external carbon
source in the MS medium or different C02 levels in the growth chamber did not rescue
the lethal phenotype (data not shown).

The homozygous T-DNA knockout line bouI (Figure 5.1) showed a typical
photorespiratory phenotype (Figure 5.3). In ambient air, two week-old boul seedlings
displayed a pale green phenotype and were significantly smaller than the wild type Col-0
plants. However, bouI seedlings were indistinguishable from wild type when germinated
at 0.3% C02 (Figure 5.3). The C02 dependent phenotype could also be observed after
shifting 6 week old plants to ambient air conditions. After 14 days, bouI rosettes
displayed a pale green phenotype and enhanced senescence, especially in the older rosette

leaves (Figure 5.3). This prompted me to focus on the explanation of the bard phenotype.

174

Figure 5.1 Isolation of A. thaliana T-DNA insertion lines for At1g54350, At2g42770,
At2g48070, At3g26710, and At5g46800

Upmr part: Gene structure and relative position of T-DNA insertions. A black box refers
to an exon. T-DNA insertion sites are indicated by triangles and arrows indicate the
relative positions of the primer for PCR analysis. M Genomic DNA PCR
analysis of wild type A. thaliana Col-0 plants (WT) and T-DNA insertion lines. Lane G
represents gene specific regions surrounding the T-DNA insertion and lane T shows the
PCR reaction with primers specific for T-DNA/ gene flanking region. Lane C refers
always to AtACT 2 control gene amplification with primer pair ML40/ML41. (A) PCR
analysis of the (At1g54350) gene in pmpI -1 (SALK003891) and pmpI-Z (SALK069087).
Gene specific products with primer pair ML102/ML103 are absent in the homozygous
pmpI -1 and pmpI-Z mutants and present in WT. Primers LBal/ML103 were used to
amplify the T -DNA specific fragment. (B) At2g42770: T-DNA insertion (SALK118636)
was verified with primers LBal/MLIOS and absence of the amplified gene fi'agment
At2g42770 in the mutant line (ML104/ML105). (C) PCR analysis of the heterozygous
T-DNA insertion line SALK055566 (At3g26710). Only plants with amplification of
target gene (primer ML108/ML109) and T-DNA flanking region (LBal/ML109) were
isolated. (D) PCR analysis of the T-DNA insertion line SALK0663 83 (At2g48070).
GenePrimers ML107/ML]20 and T-DNA insertion was verified with primers
LBal/ML107. (E) Putative mitochondrial acyl-carnitine carrier protein (BOU)
At5g46800. Homozygous T-DNA insertion line boul (GABI Kat line 079D12).
ML116fMLl 14 verified T-DNA specific insertion and primer pair ML115/ML114 were

used for genomic bou fiagment.

175

 

A pmp1-1 pmp1-2

IML102 ML102
‘—

ML103 ML103

   

 

 

 

wr wr
pmp1-1 pmp1-2
B Salk118636 c Salk055566
ML1 04 ML1 09
i—i'YI-I— :1.“—
4— <-
ML1 05 ML1 08
c r c
wr
Salk
055566

   

 

 

 

 

 

 

Figure 5.1. Isolation of A. thaliana T-DNA insertion lines

The Figure 5.1 continues on the following page.

176

 

n Salk066383

Salk
066383

 

 

 

 

Figure 5.1. (cont’d).

177

 

 

E bou1

ML114
+
4- 4- 4-
ML ML ML
166 165 115
c r c

 

bout

 

 

 

Salk
1 18636

pmp1-2

 

Salk066383

.> 1. x:

 

Figure 5.2. Growth phenotype of T-DNA insertion lines from the candidate genes
At1g54350, At2g42770, At2g48070, and At3g26710

(A) Seedlings from A. thaliana Col-0 (WT), T-DNA insertion lines from Atlg54350
(pmpI -1 , pmpI-Z), At2g42770 (SALK118636) and the photorespiratory mutant shmtI-Z
(Voll et a1. 2006) were germinated under elevated C02 concentrations (0.3%) for ten
days and the picture was taken two weeks after transfer to ambient air (0.038% C02). (B)
Homologous T-DNA insertion lines from gene At3g26710 (SALK055566) and (C)
At2g48070 (SALK066383) were sown under elevated C02 concentrations and grown for

two weeks. The figures in this dissertation are presented in color.

178

Figure 5.3. Growth phenotype of T-DNA insertion line bouI

(A) Seedlings from Arabidopsis thaliana Col-0 (WT) and the T-DNA insertion line bouI
were germinated at ambient air (0.038% C02) (upper part) or under elevated C02
concentrations (0.3%) (lower part). (B) Eight-week old WT and ban] plants had been
growing under high C02 levels (upper part). In parallel, plants were transferred to
ambient air after six weeks (lower part) and photographed two weeks later. The figures in

this dissertation are presented in color.

179

ambient air
(0.038% C02)

elevated C02
(0.3% C02)

   
 
 

elevated coz
(0.3% coz) s.

Shift to
ambient air
(0.038% 002)

 

Figure 5.3. Growth phenotype of T-DNA insertion line boul

180

Rosette leaves of boul accumulate glycine under ambient and elevated C02 levels
For several photorespiratory mutants it had been demonstrated that one or more
intermediates of the pathway accumulate after transfer to ambient air (Somerville, 2001).
To corroborate the phenotypic analysis of bar], we quantified photorespiratory marker
metabolites, such as glyoxylate, glycine, serine, and NH3 in Col-0 (WT) and ban] upon
shift from 0.3% C02 to 0.038% C02 (Figure 5.4). The most dramatic change was
observed for the amino acid glycine. Within 48 hours the glycine concentration in boa]
rapidly increased from 3.43 20.44 nmol per gram fiesh weight to 25.64 11.76 nmol per
gram fresh weight (8-fold). Even more dramatic is the comparison between wild type and
boul plants: After two days in ambient air, glycine levels in the mutant were l30-times
higher than in the wild type (Figure 5.4). Interestingly, even in high C02 conditions, the
glycine content in the mutant was 25 — 30 times higher than in the wild type. We
observed a constant increase of glyoxylate over the time course experiment under
ambient air from 31.61 :4.03 nmol per gram fresh weight to 312.8 :65.8 nmol per gram
fresh weight. This was not observed under elevated C02 conditions, where glyoxylate is
even less abundant compared to WT. The metabolites ammonium ion and serine

(Figure 5.4) and glutamate, glutamine and ornithine (Figure 5.5) did not change
dramatically during the first five days of the Shift experiment in WT and haul,
respectively. After seven days of ambient air the compounds serine, glutamine and
ornithine are slightly higher in the bard plants. Citrulline levels (Figure 5.5) were always
higher in bad plants. However, the 10-fold difference in C02 concentration between the

growth chambers did not have a large effect on the absolute citrulline levels.

181

Figure 5.4. Concentrations of photorespiratory marker metabolites glyoxylate,
ammonium ion, glycine, and serine in bar] leaves

Arabidopsis thaliana Col-0 (WT, black bars) and ban] (white bars) plants were grown at
elevated C02 concentrations (0.3%) for four weeks. Metabolites were analyzed in plants
maintained at elevated C02 levels (left panel) or moved to ambient air (right panel).
Samples were taken prior to and two, four and seven days after shifting to ambient air ((1).
Results are given as the mean :SD of three biological replicates in nmol per gram fresh
weight for glyoxylate and NH4+ or in umol per gram fresh weight in the case of glycine

and serine. FW, fresh weight; C02, carbon dioxide.

182

Elevated COZ Ambient air

ann
'VUU

 

 

 

 

 

 

 

 

 

 

.wr
2£3oo.Dbou1
22
>5!
53200-
go.
’o 100-
E ILILILEL
C
0d 2d 4d 7d on 2d 4d 7d
:E SOC-WT Dbou1 800 IWT Elba”
fig soe 600
3L
§34oc 400
58
s- 2 200
<2 00‘
5- e 0
0d 2d 4d 7d 0d 2d 4d 7d
E ”um 28 WT Elbou1
E 8‘Db0u1 24
3* 2 l. [’1
a? 6‘
30
or: 4' 3.
O
2- 4-
s .11 Jill . I]
" o'd 2'd 4'd 7'd " o'tr 21d 4'd 7'd
— 1o-
EW-wr I-wr
E 84:! bou1 8-l:lbou1
I3
£5 5 e
'6:—
m8 4- 4-
T:
E 2' 2.
H o. o-
oo 2d 4d 7d 0d 2d 4d 7d

Figure 5.4. Concentrations of photorespiratory marker metabolites glyoxylate,

ammonium ion, glycine, and serine in boul leaves

183

Figure 5.5. C02-dependent effect on additional metabolite levels in boul and WT

The identical samples as described in Figure 5.4 were analyzed and glutamate, glutamine,
ornithine, and citrulline levels were determined. Concentrations are shown as the mean
:SD of three biological replicates in nmol per gram fresh weight (F W). C02, carbon

dioxide.

184

Elevated COZ

WT
1:1 bou1

1c

 

Glutamate
[nmol per gram FW]
e 9’

 

 

 

or _
0d 2d 4d 7d

IWT
Dbou1

.3
G

 

Glutamine
[nmol per gram FW]

 

 

0d 2d 4d 7d

 

Ornlthlne
r

 

 

 

Citrulline

 

 

0.
0d 2d 4d 7d

Ambient air

IWI'
Elbou1

 

1C

 

 

3.5?
E

2‘ l |
0'

0d 2d 4d 7d
1c

84

 

 

 

ever
Ell
Egg

 

0d 2d 4d 7d

 

B
0.0.. - WT
0-04‘ I: bout

0,03-
0.02'
0.01-

 

 

i

0d 2d 4d 7d

 

0.5
I WT
0.4- 1:1 mm

0.3.
0.2-
0.1‘

 

 

0d 2d 4d 7d

Figure 5.5. C02-dependent effect on additional metabolite levels in boul and WT

185

Molecular complementation of the boul mutant

Although a truncated cDNA of the BOU gene could still be identified by RT-PCR in
boul (Figure 5.6), immunoreactive protein was not detectable in the mitochondrial
fiaction (Figure 5.9) of boul plants. A boul mutant expressing BOU under the control of
the 358 promoter (boul :BOU) (Figure 5.6) germinated and developed normally under
ambient air conditions (data not shown) and no significant glycine accumulation was

observed (Figure 5.7), showing that the boul mutation causes the C02-dependent

phenotype.

Glycine decarboxylase (GDC) activity is reduced in boul

Elevated levels of glycine and unaltered concentrations of serine and ammonium ions in
the boul mutant suggested impaired photorespiratory glycine oxidation due to either a
deficiency in glycine uptake into the mitochondria or a lack of GDC activity. Therefore,
mitochondria were isolated from high C02 adapted WT and boul plants, the envelope
membranes were permeabilized with detergent, and the rate of [U-14C] -glycine
decarboxylation to [14C]-CO2 by GDC was determined (Figure 5.8). In boul, the
apparent rate of the GDC reaction was only 15.2 1 5.3% of that Observed for wild type.
We further tested the abundance of the GDC and SHMT proteins and the presence of the
crucial GDC cofactor lipoic acid (Figure 5.9). The GDC H protein subunit, SHMT and
lipoylated proteins were not affected in mitochondrial boul fiactions. However, a subtle
negative effect on the stability of the PLP-binding GDC subunit P was observed in the

boul mutant (Figure 5.9).

186

Figure 5.6. Molecular complementation of boul

The boul mutant plants were transformed with BOU cDNA driven by the constitutive
CaMV 358 promoter. (A) PCR analysis of genomic DNA (gDNA) from A. thaliana Col-
0 (WT) and a representative complementation line boul/BOU. Lane 1: Amplification of
an endogenous BOU fragment with a BOU promoter-specific primer ML114 and primer
MLl66. Lane 2: Primer pair ML1 14/ML165 was used to confirm the boul background in
the complementation line. Lane 3: 35S-specific primer (ML143) and primer ML166
demonstrate the presence of the complementation construct in the gDNA. (B) RT-PCR
fragments from boul and a representative complementation line. Lane 1: cDNA
amplification with primer pair ML164/ML165 demonstrates restored transcript ofBOU.
Lane 2: Tnmcated BOU cDNA still present in the boul mutant. RT-PCR was done with
primer ML164 and ML166. Lane 3: RT-PCR control with AtACT7 (actin) specific

primer set m167M168.

187

 

A gDNA B RT-PCR

ML114 ML143 mm
—> z -> ->
-I ——

 

 

4- 4— 35S promoter
ML166 ML165
1 2 3

WT”

bou1/
BOU

  
   

 

 

 

 

 

Figure 5.6. Molecular complementation of boy]

188

 

03
O

E 501 IL
a, E 4 _, I wr
.E 3, 3.. D bou1
g h I bou1lBOU #1
5 g s- l I bou1/BOU#2
_. m bou1lBOU #3
O 4-
E
.3 2-

 

 

 

 

 

 

Elevated C02 Arn-bient air

Figure 5.7. Complementation of the bard mutant restores glycine concentrations to wild
type levels

A 35S:BOU construct was introduced into the bou1 mutant line and several independent
homozygous complementation lines (bou1/BOU) were established. A fraction of high
C02 adapted wild type (WT, black bars), boul (white bars) and bou1/BOU (grey bars)
plants were shifted to ambient air and glycine concentrations were determined after seven
days. Glycine concentrations are given as the mean :SD of three technical replicates in

nmol per gram fresh weight. C02, carbon dioxide; FW, fresh weight.

189

 

.9.
N
(TI

1 00-

GIN
?‘l‘

Glycine decarboxylase
activity [% of WT activity]
N
9'

Ill

Boiled WT bou1

 

 

 

‘E

Figure 5.8. Mitochondrial glycine oxidation is impaired in bou1

Arabidopsis thaliana Col-0 (WT, black bar) and ban] (white bar) plants were grown
under elevated C02 concentrations. Mitochondria were isolated fi'om four week-old
leaves and permeabilized with the detergent Triton X-100. An aliquot was incubated with
0.2 mM [U-14C]-glycine for 15 min and the released [14C]-C02 was determined. As a
control, boiled mitochondrial proteins were monitored to rule out non-enzymatic
breakdown of glycine. Relative rates are given as mean a: SD. Relative glycine

decarboxylase activity (GDC) of 100% refers to 110.29 nmol €02 per gram protein.

190

Mitochondria
Spec'fic wr bou1
l"—'_| l—‘l

 

antibody
a. , t B
BOU _> ‘7 Mitochondria
GDC ' '* ”EM“ 3 SP°°ifi° WT bou1 Protein

P protein _> " " antibody t—t PDH E2a
I - Lipoic Acidla Eilg PDH E2b

KGDH E2

‘P “- <- H protein

GDC
H protein _) . . '

 

GDC
H protein

 

 

 

 

 

SHMT +9.5

 

Figure 5.9. Immunoblot analysis of mitochondrial enzymes

(A) A mitochondrial preparation from A. thaliana Col-0 (WT) and two preparations of
the bou1 mutant line as described in Figure 6 were analyzed for the subunits P and H of
the glycine decarboxylase (GDC), BOU and serine methyltransferease (SHMT). (B) A
lipoic acid specific antibody analyzed the presence Of the lipoylated subunits E2a and 2b
from pyruvate dehydrogenase (PDH), the H subunit fi'om GDC, and the E2 subunit of the
a-ketoglutarate dehydrogenase (KGDH). Protein extracts were separated on a 15% SDS-

PAGE, immobilized on a PVDF membrane and analyzed with the protein-specific

antibodies and ECL.

191

Search for the substrate of the recombinant BOU protein
To investigate the transport activities of the putative carnitine carrier BOU, the protein
was expressed in E. coli and S cerevisiae (Figure 5.10). The recombinant transport
proteins were tagged with a N-terminal 6x His-epitope for detection and purification from
the cell extracts. In addition, a C-terminal His-tagged construct was produced in yeast
(Figure 5.10). The three constructs were reconstituted by either the fieeze/thaw method
(FT) or the cyclic detergent removal procedure (CDRP) resulting in six different
proteoliposome preparations. Each was tested with a number of putative substrates in a
homo-exchange (identical substrate inside and outside), hetero-exchange (external
substrate different fi'om the internal), or 1miport (no counter-substrate inside) experiment
(Table 5.4). Because the substrate carnitine was hypothesized earlier by Lawand et al.
(2002), our observation that the bou1 mutant accumulates glycine (Figure 5.4), and the
fact that serine is capable to inhibit glycine uptake into isolated pea mitochondria (Y 11 et
al., 1983), these three substrates were the primary targets of analysis. However, none of
these metabolites was taken up into BOU-reconstituted liposomes with significant rates
(data not shown). Electrochemical gradients across the liposomal membrane had no
positive effect on the negligible glycine or serine transport activity catalyzed by
recombinant BOU (Table 5.5).

In a search for other candidates, we next employed a phylogenomics approach.
The mitochondrial carrier family (MCF) is widely distributed in eukaryotic organisms. A
maximum-likelihood phylogenetic tree was generated with the MCF members from A.
thaliana, yeast, human, and the red algae Cyanidioschyzon merolae and Galdieria

sulphuraria (Figure 5.11). BOU belongs to a sub-clade of the MCF in which all

192

functionally tested members transport several amino acids (summarized in Table 5.5).
We hence tested uptake of these substrates into liposomes reconstituted with BOU.
Neither the proteinogenic amino acids glutamate, glutamine, arginine, nor the non-
proteinogenic amino acids ornithine, citrulline were accepted as a substrate. Since GDC
activity is diminished in the bou1 mutant, it is reasonable to hypothesize, that BOU could
be involved in the supply of GDC cofactors or precursors for cofactor biosynthesis,
which are needed in the mitochondrial matrix. Table 5.6 lists putative substrates, which
likely have to be transported across the mitochondrial membranes to provide the
biological active cofactors PLP, NAD, FAD, THF and lipoic acid for glycine oxidation
(Douce et al., 2001). Except for malonic acid, none of the compounds are available in a
radiolabeled form. Thus, flux analysis in a reconstituted liposome system cannot be
applied to these substrates. Hence, a method had to be established that permits the
screening on non-radiolabeled BOU substrates. A promising approach is the patch-clamp
technique, which however requires reconstitution of BOU into giant liposomes that can
be visualized in the light microscope. To this end, E. coli expressed BOU protein was
purified to homogeneity by Nickel-NTA chromatography (Figure 5.10) and reconstituted
into liposomes. Giant liposomes were then produced from small, unilamellar liposomes
by a dehydration/rehydration step. Using these giant liposomes, pipette-membrane seals
could be reproducibly obtained with a resistance of 1-4 giga ohm, which is suitable for
patch-clamp recording. Figure 5.12 shows that BOU reconstituted into giant liposomes
has voltage-dependent ion channel properties with discrete ion current levels between the
applied voltages +100 mV and +200 mV. Seals with resistances of 5-10 GS2 were also

achieved with protein-flee liposomes, however no voltage-dependent ion channel

193

activities could be detected between membrane potentials of -140 mV and +200 mV (data
not shown). This experimental set-up enables us to test the effect of putative substrates on
the channel activity of BOU. The current hypothesis is, that the correct substrate has a
high affinity for its binding site and will thus block the voltage-dependent ion channel
activity of the BOU transporter, which would be detected by a decreased open probability

of the channel. The substrates in Table 5.4 and Table 5.6 are under investigation.

194

PDVF
SOS-PAGE membrane

kDa 1 2 1 2

85— ‘
49— -. ..

 

 

26—

 

 

 

 

 

B PDVF
SOS-PAGE membrane

85—“? v
49" law may w

 

 

zs—r‘

 

 

 

 

 

Figure 5.10. Heterologous expression and purification of BOU protein in E. coli and S.
cerevisiae

(A) Inclusion bodies of BOU producing E. coli cells (lane 1) and Ni-NTA purified BOU
protein (lane 2) were separated by SDS-PAGE and stained with Coomassie Blue. The
proteins of an identical gel were blotted on a PDVF membrane and the N-terminal His-
tagged BOU was verified with an anti-His antibody. (B) Membrane proteins from yeast
cells were separated on a 12% SDS-polyacrylamide gel and expression of N-terminal-
His-tagged (lane 1) and C-terminal-His-tagged (lane 2) recombinat BOU protein was
demonstrated with an anti-His antibody. Protein extracts from yeast cells, which had been

transformed with an empty expression vector are shown in lane 3.

195

Table 5.4. Systematic search for the BOU substrate

N-terminal His-tagged BOU protein was expressed in yeast and E. coli. In addition, BOU
was also produced with a C-terminal fused His-tag in yeast and tested. The purified
protein was reconstituted into liposomes by the freeze/thaw (FT) procedure or by the
cyclic detergent removal procedure (CDRP) as described in “Material and Methods”.

 

 

Radiolabelled . Functional
external Liposomgtplreloaded Recognblipant pill-loteln reconstitution
substrates ‘1’ essed procedure
:[14C1' :mmfine Y & E r FT & CDRP
i tine None east . co 1
Glycine
{MC} Serine Y & E 1' FT & CDRP
Gl cine ea“ "'0' ,
y None
4 Serine
[1 91' Glycme Yeast & E. coli FT& CDRP
Senne None
[14C] Glutamate
' Glutamine
Glutamate None Yeast FT
[14C] Glutamine
f Glutamate Yeast FT
Glutamlne None
Arginine
[14C]- Omithine
None
Omithine
14C _. Citrulline
OE'nithine Arginine Yea“ FT
None
[14C]- Malonic acid .
M alonic acid None Yeast & E. colt FT & CDRP

 

196

Table 5.5. Presence of a membrane potential does not stimulate glycine or serine

transport by BOU

The ionophore valinomycin induced a potassium (K+) diffusion potential gradient to
analyze its influence on glycine or serine transport To generate an artifical pH gradient
with the K+/H+ exchanger nigericin, the internal (in) medium (outward K+ directed
gradient) or the external (out) medium (inward K+ directed gradient) was not buffered

 

 

(NB), respectively.
. . K+ (in) /
Ll somes H rn/
lpo . p K" (out) Valinomycin N igericin
pre oaded Wlth pH out i “,li
. . 8 / 8 l / l
Glycme, serine, none
8 / 8 50/ 1
8 / 8 l / 50
. . NB / 7 1 / l
Glycrne, serine, none
NB / 7 50/ 1
, . 7 /NB 1 / 1
Glycme, senne, none
7 / NB 1 / 50
. _ 8/6
Glycme, serine, none
6/8

 

197

Table 5.6. Several cofactors must be provided for the mitochondrial glycine
decarboxylase (GDC)

For cofactor biosynthesis inside the mitochondria, precursors have to be imported If a
cofactor biosynthesis occurs outside the mitochondria, the biologically active form must
be provided from the cytosol. All steps require protein-mediated transport steps across
the inner envelope mitochondrial membrane. Abbreviations: PLP, pyridoxal 5-phosphate;
SAM, S-Adenosylmethionine; NaMN, nicotinate mononucleotide; pABA, para-
aminobenzoic acid.

 

 

GDC Co factor Final step cofactor Predicted transport processes
subunit biosynthesis across mitochondrial membrane
PLP
P protein PLP Cytosol Pyridoxine
Pyridoxal
Malonic acid
. . Pyruvate
rtftlein 1:12:30 Cytosol “Sulfur” donor (cysteine, SAM)
p CoenzymeA (Valine,
ketopantoate, CoA)
T protein THF Mitochondria Pterin, pABA, glutamate
L protein FAD Cytosol FAD
Quinolinate
L protein NAD Mfioygi‘t’lld/fia NaMN
‘ NAD

 

198

AthAC1 (At2933820)

YMC1/YPRO58W
YMC2/YBR104W
Gs20540.1
AtBOU (At5946800) |
'— HsORNTZ/SLC25A2
HsORNT1/SLC25A15
HsCAC/SLC25A20
HsC14orf68/NP997000
AthACZ (At1g79900)
ORT1IARG11/YOR1BOC
CRC1/YOR1OOC

 
 
   
 
 
 
 
 
  
 

 

 

 

Figure 5.11. BOU relevant branch from the maximum-likelihood unrooted tree of the
mitochondrial carrier family (MF C)

MCF sequences from yeast, plants and human were obtained from NCBI
1hng/wwwncbi.nlm.nih.gov[). The red algae sequences were extracted from the
Galdieria sulphuraria (http://genomics.msu.edu/galdieria) and Cyanidioschyzon merolae
(httns/l’ ‘ hinl s H-tokvo.ac.ip) genome project database. Protein annotations from
yeast, human, plants and red algae are written in brown, blue, green and red, respectively.
Bootstrap values are pointed out at the corresponding nodes. The figure in this

dissertation is presented in color.

199

v [mV]
C3 wwwele - WWW it“ __________________________
—————— fiTH-——LJLJe‘w-———-—————————————————-

. ' ewwefifw «W ------ +200

C5

 

MWMMW++ ————————————————————————————————

(:2 "L ------ ”V'mfwmw ww+180
J. r... 1....1 “W

———————— 3—14——TpL————————L—-————
C4 —————————— ___..r ————————————————————————

 

 

 

 

C4 ' T7‘7“"""""'1+160
........................ 1"-.'.".'- Weevil

ca ........................ J ....... an-.."
”Titiamv‘v‘liwi'fif" N.v.v'.w..|..mwiilir .. fili‘u: L41? quym m?»
_..__'._,_ _____ +W ._"'“‘ AMT". _____ Memw-__q_.w+l40
_______________ L..- ___L,__________..___~u_

C2 _________________:i _________________ ext.

 

 

 

 

 

 

 

 

 

00080

 

 

L- -L 4‘; 1 H A m A A. n A. . ~1 _ ‘4 -
v... .. re . a“: , . r . -‘ Y , 1,. w e —b'-hvw¢1-|o‘qu—apfirb~m
0.49.»...Am-WM-J.‘ unatvon¥ t .---I - *AA ‘4-—~ ‘4- A— -‘ ‘__fl“u~”~d*’+fi-*’WM‘W.TI~N~‘V mural—m +40
A A r— ,_ .— mucv-M.u-_u_.s-. ---— 8......
0.. — ~.I~.~..».u.--.— Mn..."— .._...-.._. _~..~.A — .- v u... .- ~--~n-‘-_.o~.~ “II—u. mM.—.0 m.-—- ...-¢ 0
--...~_-.--—-—.--c.‘_-...~~. on. .— ,__,... --. . - . .. ». .0 ~.—.—_#. ---.._.._—.— --- ~.- -- --——. . -....-..._ u. .. _,__,.'~____ *u ..__. ---.......--.. _ ....——~—~—
»¢-~.-— pen-so .W a' v ‘ v -— a ,. W. A A w-w
W'»~w-' — w . " v . v—L “‘7 v _— V v— . fl", . ‘fid~~v‘-~um~*vw-qew-—-rw-wwa

 

m‘vmflfinvwwmmm-aww- - 8 0

5......“ we..." ~~m~rnmmopaa unwifiwww

 

 

1 sec
Figure 5.12. Reconstituted BOU protein exhibits ion channel properties

Purified BOU protein was reconstituted into giant liposomes. Single channel currents
were recorded in a symmetrical 100 mM KCl buffer between negative (-80 mV) and
positive (+200 mV) pipette potentials (V). Currents levels of open state (0) and closed

state (C) channels are shown.

200

DISCUSSION

Identification of membrane proteins co-expressed with photorespiration

The carbon and nitrogen cycles of the photorespiratory pathway are distributed in three
different compartments and acting jointly to recycle two molecules 2-phosphoglycolate to
3-PGA (T olbert, 1997). A prerequisite to facilitate the traffic of the photorespiratory
intermediates are numerous shuttles inserted into the plastid, peroxisomal and
mitochondrial membranes. As an alternative to identify these transporters by forward
genetics, the CSB.DB resource was screened for hydrophobic membrane proteins that are
highly co-regulated with known photorespiratory genes (Steinhauser et al., 2004). As
commented by Wei et al. (2006) the photorespiration is embedded in the expression
pattern of genes from the photosynthetic apparatus and the Calvin-Benson-cycle. Several
components of the photosynthetic electron transport chain were indeed found to be
repeatedly co—expressed with photorespiration, since the light reaction overlaps
temporarily and spatially with the photorespiration and the initial substrate ribulose-1,5-
bisphosphate and final product 3-PGA are directly connected to the Calvin-Benson-cycle.
The expression profiles of several metabolite transporters were also tested. The Calvin-
cycle associated triose-phosphate/phosphate transporter, which exports recently fixed
C02 into the cytosol, was retrieved from our CSB.DB search. Other transporter genes
such as the mitochondrial basic amino acid transporter (AthACl) or the peroxisomal
nucleotide transporter AtPNCl do not cluster with the query genes and point out the
applicability to screen for photorespiratory candidate transport proteins. To prioritize the
candidates for reverse genetic analysis, already characterized carrier proteins and

thylakoid-localized membrane proteins were neglected. Despite the progress in proteomic

201

analysis of all three organelles and GFP studies (Miller et al., 2001; Reumann et al.,
2004; Sun et al., 2008), the localization of more than two-third of our candidate genes (23
out of 32) is predicted solely by in silico targeting programs. Seven out of the nine
candidate proteins were found by Arabidopsis chloroplast proteomic studies (Sun et al.,
2008), one is postulated to be dual targeted to mitochondria and plastid (At5g64970)
(Dunkley et al., 2006) and the remaining candidate is inserted into the mitochondrial

membrane (At5g46800) (Lawand et al., 2002).

Photorespiratory transport systems across the plastid inner envelope membrane
Besides the two-translocator shuttle for 2-OG/ glutamate recycling in the plastid (Renne et
al., 2003), a single transport protein for glycolate/glycerate antiport or their uniport was
demonstrated on intact chloroplasts (Howitz and McCarty, 1991). This flexibility in the
transport mode (i.e. uniport vs. antiport) is important, because for every recycled and
finally imported glycerate two molecules glycolate have to be exported fi'om the plastid
(Figure 1). The nitrogen cycle would require a transport system for the ammonium ion,
glutamate/glutamine or ornithine/citrulline performed by three to maximal five proteins
across the chloroplast inner envelope membrane (Linka and Weber, 2005). Transport of
glutamine and citrulline was demonstrated on intact chloroplast - its relevance in
photorespiration, however, is speculative (Y u and Woo, 1988; Ludwig, 1993).

Eighteen candidate proteins have a robust plastidial computer-predicted targeting
and seven of them were in addition identified by proteomics. T-DNA insertion lines for
four plastid-targeted genes did not show a conditional chlorotic phenotype under ambient

air conditions (Figure 5.2). Homozygous mutant lines from At2g48070 and At3g26710

202

lack pigments and seedlings did not develop beyond the cotyledon stage. At2g48070 has
been identified by a recent proteomics study as a protein targeted to thylakoid membranes
(Zybailov et al., 2008). These results direct its function towards the biogenesis of the
photosynthetic apparatus. In the case of SALK line 055566 (At3g26710) only
heterozygous mutant lines fully developed Exon-insertion lines for Atlg54350 and
At2g42770 grew normally in ambient air (Figure 5.2). Atlg54350 (AtPMPl) is annotated
as one of 129 ABC transporter proteins in A. thaliana (Sanchez-Fernandez et al., 2001).
At2g42770 is related to the peroxisomal pore-forming 22 kDa membrane protein PMP22
from human, however, not further characterized yet (T ugal et al., 1999; Visser et al.,
2007). Seven additional proteins are annotated as PMP22 or PMP22-related proteins and
targeted to plastid, peroxisome and mitochondria in the genome of A. thaliana
(ARAMEMNON database; Schwacke et al., 2003). At this point it cannot be ruled out
that the PMP22 members, AtPMPl or gene product At3g26710 are involved in
photorespiration and it is tempting to speculate that genetic analyses are masked by
functional redundancy or by the severity of the loss of function mutant, respectively. The
majority of photorespiratory mutants tln'ive under elevated C02 concentrations. This is
not true for mutants deficient in the enzyme glycine decarboxylase (Engel et al., 2007). A
complete loss of the enzyme activity is lethal and alternative strategies such as inducible
RNAi targeted gene silencing might be necessary to aim for novel photorespiratory

mutants.

203

The mitochondrial envelope membrane transport protein BOU is required for
photorespiratory function

BOU (At5g46800) was the only evident mitochondrial transport candidate that was
retrieved from the co—expression data (Table 5.3). The gene belongs to the mitochondrial
carrier family (MCF) and was previously published as a putative carnitine/acylcarnitine
carrier providing alternatively acetate as a carbon and energy source to the mitochondria
(Lawand et al., 2002). Under long-day conditions, the seedlings were not capable to
establish photoautotrophic growth. In this study, we isolated a second mutant allele
(bou1) and demonstrated that “non-photorespiratory” conditions can completely rescue
the seedling’s development (Figure 5.3). The classical pale green phenotype was not
apparent until the 14th day, which might explain why Somerville & Ogren (1979) did not
isolated an EMS mutagenized plant with a deficient BOU gene. Their genetic screen
concentrated on visible chlorotic effects after four days. Importantly, metabolic analysis
of boy] demonstrated a massive accumulation of glycine already under 0.3% C02 and
even more dramatic increase at the second day at ambient air (Figure 5.4). This suggests
that metabolic profiling of photorespiratory marker metabolites is a powerful tool to
identify additional photorespiratory mutants that do not display an apparent phenotype.
Glycine import and serine export are the most prominent transport processes during
photorespiration in the mitochondrial envelope membrane. Two studies on isolated
mitochondria showed that both substrates could interfere with each other for uptake
(Walker et al., 1982; Yu et al., 1983). Analogous to the plastid glycolate/glycerate
shuttle, a single protein is hypothesized to mediate a glycine/serine transport in a

stoichiometry of 2:1. Numerous attempts with distinct experimental conditions failed to

204

show transport activity of glycine, serine, or carnitine with recombinant BOU (Table 5.4
and Table 5.5). Elevated citrulline levels (Figure 5.5) and the phylogenetic relation to
basic amino acids transporters (Figure 5.11) directed the analysis to test the hypothesized
ornithine/citrulline and glutamine/ glutamate shuttle system without any evident activity.
Intriguingly, bou1 exhibits a similar glycine accumulation in leaves as knockout mutants
affected in the provision of cofactors lipoic acid and THF for the glycine decarboxylase
complex (Ewald et al., 2007; Collakova et al., 2008). The GDC activity was diminished
(Figure 5.8) in haul and the PLP dependent P protein subunit was partially degraded
(Figure 5.9). The Vitamin B6 derivative PLP is synthesized in the cytosol (T ambasco-
Studart et al., 2005) and GDC is the most abundant soluble protein in leaf mitochondria -
therefore a major sink for PLP (Sarojini and Oliver, 1983; Douce et aL, 2001; Bauwe and
Kolukisaoglu, 2003). Taking both arguments into account, we hypothesize that BOU
provides PLP to the mitochondria. Since PLP covalently binds to the P protein subunit
and is an integral part of its active site, it is tempting to speculate that a distorted P
protein lacking PLP is prone to degradation (Figure 5.9). Since additional enzymes such
as amino-transferases are dependent on PLP and a GDC knockout is lethal, an impaired,
but not complete lack of PLP provision, has to be proposed at this point. Using the patch-
clamp technique we demonstrated that the BOU is functional when reconstituted into
liposomes and displays voltage-dependent ion channel activity (Figure 5.12). This
suggests that the previously tested metabolites, which were reconstituted with the same
procedure, are not the substrates of BOU. Future studies will show if PLP, pyridoxal or
pyridoxine can inhibit the voltage-dependent ion channel activity of BOU. Identification

of the major mitochondrial glycine import protein will be challenging. The A. thaliana

205

genome predicts at least 53 selective or broad-spectrum amino acids transporters
distribute in every plant tissue and in most subcellular compartments including plasma
membrane, mitochondria, plastid and peroxisomes (Wipf et al., 2002). Complete loss of
glycine import would stall the essential GDC activity and in a broader view the
mitochondrial protein synthesis (Engel et al., 2007). For protein synthesis a complete set
of amino acids is required and a sophisticated and partial overlapping import system for
mitochondria can be postulated. Mutant analysis of promising amino acid transporters
combined with metabolite profiling might help to resolve the major route for glycine

import.

Photorespiratory transport systems in the peroxisomal membrane

In comparison to mitochondria and plastids, very little is known about peroxisomal
membranes (Reumann et al., 2004; Reumann and Weber, 2006). The isolation of pure
peroxisomal membrane fraction for proteomic studies is challenging and a target signal of
peroxisomal membrane proteins for computer-based analysis is not deciphered yet
(Reumann et al., 2004). A porin-like channel is postulated to facilitate the flux of the
mono- and dicarboxylates glycolate, oxaloacetate, malate, 2-OG and glycerate

(Reumann et al., 1996; Reumann et al., 1998). Transport of amino acids glycine, serine,
or glutamate and the bulky cofactors NAD, NADP, or CoenzymeA remains completely
unanswered. We could extract from the co-expression analysis a member of the PMP22-
related protein family in A. thaliana. However, an obvious visible phenotype could not be
detected in the corresponding mutant. The twelve PMP22-related genes might be

promising targets for metabolic profiling and multiple gene knockout analyses.

206

OUTLOOK

Despite the overrepresentation of plastid-targeted proteins, co-response analysis is a
valuable tool to associate genes of tmknown function with photorespiration. BOU was
identified as a new player in the photorespiratory network. An initial genetic analysis of
the remaining 27 lines is feasible in the model organism A. thaliana. Detailed metabolite
analysis of the mutant plants and a selective transcript profiling of the candidate genes in
wild type plants adapting to high and low C02 concentration will add important

information to identify missing transporters from the photorespiratory network.

207

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CHAPTER 6

CONCLUSIONS AND PERSPECTIVES

215

CONCLUSIONS AND PERSPECTIVES

The long-term goal of studying metabolite transporters is to link the uptake, distribution
and secretion or excretion of a metabolite with its particular transport protein. This
knowledge is essential for understanding the cellular metabolic networks by providing
insights into the control of fluxes of key metabolites between metabolic pathways -from
intracellular compartments up to the organismal level. This knowledge is also required
for rational engineering of valuable natural products and plant biomass. Proteins, fatty
acids, starch, and cellulose have been our central agricultural food and energy resource in
the past and will be in firture. The importance of solute carriers for the synthesis of the
latter metabolites is underestimated mainly due to the limited molecular, biochemical,
and structural information on many of these transport processes, as compared to soluble
proteins (Xie, 2008). The findings presented in this dissertation validate that transport
systems are rather specific enzymes, which are precisely placed in a eukaryotic cell to
connect or even regulate the compartmentalized metabolic network, and not simply pore-
like structures in biological membranes that are permeable for most of the small
molecules.

In chapters one and three I characterized an ancient sugar phosphate transporter
system in the plastid envelope membrane of the red alga G. sulphuraria. These
transporters originated during the endosymbiosis between a photosynthetic
cyanobacterium and its eukaryotic host. The insertion of a single pre-existing carrier into
the inner membrane of the endosymbiont provided a selective advantage by reallocating
photosynthates into the cytosol of the host, thereby directing the symbiosis to a

permanent enslavement of the cyanobiont The bacterial envelope membrane had been

216

massively altered with transport proteins predominantly derived fiom the host.
Importantly, the plastid phosphate translocator family (pPT) evolved fi'om an ancient
nucleotide-sugar transporter to a versatile group of transporters mediating the exchange
of Pi, triose-phosphate (TP), phosphoenolpyruvate (PEP), and glucose-6-phosphate
(Glc6P) between cytosol and plastid of higher plants (F lfigge, 1999). It testifies the
dynamic adaptation and the potential of carrier proteins to influence metabolic networks.
The identification of the triose-phosphate/phosphate transporter fiom G.
sulphuraria (GsTPT) with a high specificty and high affinity for its substrate triose-
phosphate opens up a new tool to investigate carbon partioning into soluble and insoluble
sugars in higher plants. The GsTPT has been introduced into the genome of A. thaliana

wild type and (pt knockout plants (Sclmeider et al., 2002) and stable transformants are
currently being established. In green plants, carbon partitioning is primarily organized in
the plastid. Recently fixed carbon has three major routes in the Calvin-Benson—cycle. It is
exported to the cytosol in form of triose-phosphates, further metabolized to produce
hexose-phosphates for starch synthesis, or recycled to the C02 receptor ribulose-5-
phosphate. The TPT is positioned at the metabolic branch of TP. Triose-phosphate
export supplies carbon for sucrose synthesis during the day and on the other hand starch
reserves are built up for carbon provision during the night. This assures continuous
growth and development of the plant (Smith and Stitt, 2007). Over-expressing a TPT
protein in tobacco plants did not significantly alter carbon partitioning, but showed that
TPT activity limits the mardmal rate of photosynthesis (Hausler et al., 2000a; Hausler et

al., 2000b). The GsTPT will allow us to investigate the capacity of TP export to

217

influence sucrose and starch synthesis. Accumulation of sucrose and hexose-phosphate in
the cytosol sequesters cytosolic Pi, which is needed for the counter-exchange with the
plastid-localized triose-phosphate. As a result, TP transport rate slows down and 3-PGA
levels increase relatively to Pi in the plastid. The altered 3-PGA/Pi ratio activates
allosterically the enzyme ADP-glucose pyrophosphorylase (AGP), the committed step
in starch synthesis (Ballicora et al., 2004). A three and two-fold higher affinity for Pi and
triose-phosphate, respectively, could increase and extend TP export during the day,
thereby promoting prolonged sucrose synthesis. TPT is not the solely regulator to
control the carbon flux. Thioredoxin and trehalose-6-phosphate or the regulatory molecule
fructose-2,6-bisphosphate influence the activity of AGP and sucrose synthesis,
respectively (Stitt et al., 1984; Tiessen et al., 2002; Lunn et al., 2006). Monitoring the
rate of sucrose and starch accumulation throughout the day and a comprehensive analysis
of enzyme activities and levels of soluble sugar and sugar-phosphates will demonstrate if
modulating TPT kinetic constants can indeed influence the robust carbon assimilation
network. Activities of AGP (Srnidansky et al., 2002) or the sucrose degrading enzymes
invertase and sucrose synthase in storage organs such as seeds, fruits, and tubers (Herbers
and Sonnewald, 1998) contribute to their sink strength and have 3 effect on organ number,

size and dry weight.

The triose-phosphate transport protein fi-om G. sulphuraria may be a promising
candidate for manipulating the source strength of a crop plant. If this is true, the additive

effects of an increased source and sink strength are worth to be evaluated in future. Sink

strength is a quantitative trait and an ultimate goal will be to identify the “least gene

218

alleles combination” to improve the agricultm‘al yield.

Another difference between the TPT proteins of higher plants and the red algae
are their affinity to the metabolite 3-PGA. G. sulphuraria membranes lost or never
evolved the capability for 3-PGA transport. This clearly points towards an important
physiological role of 3-PGA transport in higher plant chloroplasts. By comparing the A.
thaliana wild type and tpt knockout plants expressing the red algal protein, we will be
able to assess the role of 3-PGA transport across the chloroplast envelope inA. thaliana
leaves. Based on the results I provided in chapter three, the GsTPT cannot restore the 3-
PGA transport activity in the knockout plants. We can use this approach to investigate
the proposed TP/3-PGA reduction equivalent shuttle and in addition the postulated
provision of 3-PGA for the cytosol to fuel the glycolytic pathway for PEP and pyruvate
metabolism. We will focus our investigation on the NADP-dependent malate
dehydrogenase in the chloroplast and the glycolytic enzymes in the cytosol. Reduction of
oxaloacetate (0AA) to malate regenerates the terminal electron acceptor NADP+ of the
photosynthetic system and malate is exported to the cytosol and subsequently oxidized
to 0AA (Scheibe, 2004). Thus, the redox status of the plastid is tightly connected to the
NAD(P)H pool of cytosol and the mitochondria (Raghavcndra and Padmasree, 2003).
The so-called “malate/0AA valve” and the TP/3-PGA shuttle are the main routes to
export reduction equivalent during the day (Heineke et al., 1991). The effect of the

impaired TP/3-PGA shuttle on the photosynthetic performance can be studied in detail.

In chapter three I could not assign glucose-6-phosphate as an in vivo substrate for

219

the homologous GPT protein fi'om G. sulphuraria. This finding strongly suggests that
GsTPT is a versatile exporter and importer of triose-phosphate depending on the
autotrophic or heterotrophic growing conditions of the cell, respectively. I tested more
than 20 commercially available sugar-phosphates without success. It will be challenging to
identify the biological substrate of the “enigmatic” transport protein. Metabolic profiling
is currently being pursued in the laboratory and not yet tested metabolites, such as
phosphorylated disaccharides, trehalose, floridoside or sulfated soluble sugars might be

recognized in the analysis as putative candidate.

The characterization of the pPT family and the comprehensive phylogenetic
analysis of the metabolite transport proteins demonstrated the plasticity of plastid
organelle evolution. The majority of the plastid metabolite transport systems have been
established in the last common ancestor of the Archaeplastida. Nevertheless, distantly
related photosynthetic organisms evolved alternative solutions to organize and optimize
their compartmentalized metabolic network within the last one billion years. Investigation
of these homologous proteins offers a great potential to discover a variety of regulatory
mechanisms to fine tune the cellular metabolism and increase agricultural productivity and

efficiency.

In addition, chapter two provides a new perspective on plastid evolution.
Connecting the cytosol of host and cyanobacterium by existing host solute transporters
was a key feature for the early success of the endosymbiosis or even reflects a turning
point to a permanent partnership of the endosymbiont by the protist. The export of

photosynthates was a central element in the integration of plastid-host metabolism.

220

Importantly, the pPT family was not an exception. Transporters derived primarily fiom
eukaryotic genes and provide a selective advantage for the protist. When this analysis
was initiated, several studies investigated the result of the endosymbiotic gene transfer
(EGT) on the nuclear genome of photosynthetic eukaryotes (Timmis et al., 2004). Here
we demonstrated that the eukaryotic host made a major contribution for the proteome of
the organelle. Host genes diversified and were targeted to new locations in the cell with
alternative functions as described for cyanobacterial genes by EGT. In essence, we
concluded that the initial connection of independent metabolite entities (i.e.,
cyanobacterium and eukaryotic host cell) by metabolite transport proteins, was a crucial
early event in endosymbiosis, providing the selective advantage that permitted the step-
wise completion of the process by EGT and establishment of an efficient protein

translocation machinery.

Identification of metabolite transporters participating in the photorespiratory
pathway

As stated in the first paragraph, the challenge will be to characterize the metabolite
carrier families predicted fiom computer-based analyses of several genomes, membrane-
specific proteomics approaches and the transport capacity of different membrane
fractions (Barbier-Brygoo et al., 2001). Chapter two, four and five represent typical
examples of the discrepancy between the scientific progress made for soluble and
insoluble proteins participating in a specific pathway. One limitation of our
phylogenomic analysis is the not yet characterized physiological roles of the majority of

the transport proteins. By knowing the substrate of each individual carrier we could

221

functionally categorize the host, cyanobacterial, and chlamydial contribution for the
transport of inorganic ions and metabolites across the envelope of the endosymbiont and
draw the “ancient plastid-cytosol interaction map” of the proto-alga.

The enzymatic steps to recycle 2-phosphoglycolate in the photorespiratory
pathway are described in detail, however, the molecular nature of the transport systems in
the plastidial, peroxisomal and mitochondrial membranes are mainly unknown (Reumann
and Weber, 2006). A first attempt has been already published to successfully apply the
basic knowledge of glycolate metabolism and design an alternative pathway to recycle 2—
phosphoglycolate and increase biomass (Kebeish et al., 2007). The glycolate catabolic
pathway from E. coli was targeted to the plastid inA. thaliana to short-circuit the
endogenous photorespiratory pathway. Is the photorespiratory pathway and the
identification of the transport proteins already “a thing of the past” (Sage and Coleman,
2001)? A detailed investigation of the transporters can contribute to the understanding of
the photorespiratory network. For example, if the plastid glycolate exporter would have
been identified, a targeted reduction presumably increases the flux through the
biotechnologically introduced prokaryotic glycolate pathway. We could challenge the
biotechnological approach by testing the capacity of glyoxylate detoxification in the
plastid. Is the engineered photorespiratory bypass able to cope with an increased
glycolate oxidation and result in an enhanced biomass production?

In chapter four I addressed the nitrogen cycle during photorespiration. The
mitochondrial glycine decarboxylase reaction releases C02 and ammonia. It is the “most
inefficient step” in terms of the recycling of glycolate during the photorespiration.

Nevertheless, GDC is essential and knockout mutants are lethal even under non-

222

photorespiratory conditions (Engel et al., 2007). An alternative approach to improve
photorespiration could be the re-assimilation of CO2 and ammonia on site. Future
experiments have to elucidate, if glutamine synthetase (GS) and carbamoyl-phosphate
synthase are required for the assimilation of both gases during the day (T aira et al., 2004;
Keys, 2006). To this end, it is still controversial if a glutamine synthetase (GS) is active
in the mitochondria. 35S-driven GF P analysis of GS and purified mitochondria exhibit
GS activity (Taira et al., 2004). On the other hand, to cope with the significant ammonia
release, GS should be an abundant protein in the mitochondrial matrix. However,
numerous organelle specific proteomics studies could not identify a mitochondrial GS
(Heazlewood et al., 2004; Eubel et al., 2007). Over-expressing GS alone or in
combination with the carbamoyl-phosphate synthase in the mitochondria could be an
alternative approach to decrease the loss of the ammonia and C02 gas to the
environment.

The classical forward genetic approach identified only a single metabolite
transporter linked to photorespiration (Somerville, 2001). The aim in chapter five was to
initiate an alternative reverse genetic approach to identify membrane protein candidates
mediating the flux through the photorespiratory pathway. We isolated 32 putative
metabolite transporters and I initiated the genetic analysis of five of them. Two knockout
mutants were indistinguishable from wild type in ambient air. Two insertion lines were
lethal and did not develop beyond the cotyledon stage. The phenotype was independent
of the C02 concentrations in the growth chamber. Lastly, the bou1 mutant showed a
delayed classical photorespiratory phenotype. Similar to the GsGPT protein, despite the

knowledge gained fi'om genetic, bioinformatic and biochemical analyses of the bou1

223

mutant, it has been difficult to assign a substrate for the mitochondrial transporter. The
next step will be to demonstrate the vitamin B6 transport by the BOU protein across the
mitochondrial membrane. A future goal will be to establish the remaining knockout
plants and systematically screen for accumulation of photorespiratory marker metabolites
to prioritize the candidate gene list. Essential genes could be targeted with an inducible
RNAi approach.

To characterize transport proteins, the genes are heterologous expressed, purified,
and ftmctionally reconstituted in artificial vesicles (Palmieri et al., 1995). During my PhD
thesis, I established additional systems to fimctionally characterize transport proteins in
the laboratory of Prof. Andreas Weber. We are able to express the proteins in yeast, E.
coli and in cell-free systems based on extracts of wheat germ and E. coli. I established an
alternative method, the so called “cyclic detergent removal procedure” (Kraemer and
Heberger, 1986), to reconstitute membrane proteins in the laboratory. Transport assays
are also limited due to availability of radiolabeled substrates. With the patch-clamp
technique it will be possible to test non-radiolabeled substrates. This will allow uptake
assays with candidate metabolite, such as vitamin B6 derivatives, which are not
commercially available radiolabeled. A combination of all methods will help to overcome
the difficulties in transport research to express, preserve the activity of the purified

protein and identify the in vivo substrate.

224

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