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The anaerobic life of the photosynthetic alga

Chlamydomonas reinhardtii

Photofermentation and hydrogen production upon

sulphur deprivation

Das anaerobe Leben der photosynthetischen

Alge Chlamydomonas reinhardtii

Photofermentation und Wasserstoffproduktion unter

Schwefelmangel

Dissertation zur Erlangung des Grades

eines Doktors der Naturwissenschaften

der Fakultät für Biologie

an der Internationalen Graduiertenschule Biowissenschaften

der Ruhr-Universität Bochum

angefertigt am Lehrstuhl für Biochemie der Pflanzen

in der Arbeitsgruppe Photobiotechnologie

vorgelegt von

Anja Christine Hemschemeier

aus Engelskirchen

Bochum

August 2005

Meinen Eltern und Meike.

In Liebe und Dankbarkeit.

„Wer sich nicht mehr wundern und in Ehrfurcht verlieren kann,

ist seelisch bereits tot.“

Albert Einstein

Teile dieser Arbeit wurden bereits veröffentlicht:

Happe T., Hemschemeier A., Winkler M. and Kaminski A. (2002) Hydrogenases in green

algae: Do they save the algae´s life and solve our energy problems? Trends Plant Sci 7,

246-250

Winkler M., Hemschemeier A., Gotor C., Melis A. and Happe T. (2002) [Fe]-

hydrogenases in green algae: Photo-fermentation and hydrogen evolution under sulfur

deprivation. Int J Hydrogen Energy 27, 1431-1439

Hemschemeier A. and Happe T. (2005) The exceptional photofermentative hydrogen

metabolism of the green alga Chlamydomonas reinhardtii. Biochem Soc Trans 33, 39-41

Fouchard S., Hemschemeier A., Caruana A., Pruvost J., Legrand J., Happe T., Peltier G.

and Cournac L. (2005) Autotrophic and mixotrophic hydrogen photoproduction in sulfur-

deprived Chlamydomonas cells. Appl Environ Microbiol, in press

Referent: Prof. Dr. Thomas Happe, Lehrstuhl für Biochemie der Pflanzen,

Arbeitsgruppe Photobiotechnologie

Koreferent: Prof. Dr. Franz Narberhaus, Lehrstuhl für Biologie der

Mikroorganismen

Tag der Abgabe: 26. August 2005

Diese Arbeit wurde gefördert durch die Studienstiftung des deutschen Volkes

Table of Contents

Table of contents 1

1 Introduction 1

1.1 Hydrogenases and hydrogen metabolism in unicellular green algae 1

1.2 Hydrogen production of sulphur-deprived C. reinhardtii cells 6

1.3 Fermentation: anaerobic energy production 11

1.4 The pyruvate formate-lyase system in E. coli 12

1.5 Objectives of this work 15

2 Materials and Methods 18

2.1 Organisms and growth conditions 18

2.1.1 Green algae 18

2.1.2 E. coli 19

2.2 Plasmids 20

2.3 Oligonucleotides 21

2.4 DNA and RNA techniques 21

2.5 Expression of C. reinhardtii pfl and pflA in E. coli 23

2.6 Western Blot Analyses 24

2.7 Physiological analyses of algal cultures 25

2.7.1 Quantification of hydrogenase activity 25

2.7.2 Measuring oxygen exchange with a Clark-type electrode 26

2.7.3 Detection of fermentative products and starch 26

2.7.4 Chlorophyll fluorescence measurements 27

2.7.5 Mass spectrometric analyses of the gas exchange in algal cultures 32

3 Results 36

3.1 Analysing the reasons leading to hydrogen production 36

3.1.1 Mass spectrometry as a tool for analysing gas exchange in C. reinhardtii 36

3.1.2 The analysis of C. reinhardtii mutant strains revealed three special phenotypes 40

3.1.3 A PSII-mutant offers clues to the electron source of hydrogen production 41

3.1.4 Is acetate essential for hydrogen production? 44

3.1.5 Hydrogen metabolism is delayed in a cytochrome oxidase deficient strain 46

3.1.6 A Rubisco-deficient strain produces hydrogen in the presence of sulphur 56

3.2 C. reinhardtii has an exceptional fermentative metabolism 63

Table of Contents

3.2.1 C. reinhardtii has several ethanol producing pathways 63

3.2.2 Fermentation is impaired in mutant strains FuD7 and CC-2803 65

3.2.3 The C. reinhardtii pfl and pflA cDNAs were isolated and characterised 66

3.2.4 C. reinhardtii PFL is functionally synthesised in E. coli 69

3.2.5 The algal PflA fails to activate E. coli PFL 73

3.2.6 Expression studies on selected genes 74

3.3 Overview of results 77

4 Discussion 78

4.1 What are the factors that finally lead to hydrogen production? 78

4.2 What kind of fermentative system is active in C. reinhardtii? 98

5 Summary 109

6 Zusammenfassung 111

7 References 113

8 Appendix 125

8.1. Assembly of the pflA-cDNA and deduced oligonucleotides 125

8.2. Alignments of PFL and PflA polypeptides 126

8.3. Annotated sequences encoding fermentative enzymes in C. reinhardtii 127

9 Curriculum vitae 129

Abbreviations

Abbreviations

aas amino acids

ACK acetate kinase

ADH Zn-containing alcohol dehydrogenase

ADP/ATP adenosine diphosphate / adenosine triphosphate

AOX alternative oxidase

bps base pairs

CDD Conserved Domain Database

cDNA “copy” desoxyribonucleic acid

CDS coding sequence

Chl chlorophyll

CoA coenzyme A

COX cytochrome oxidase

DBMIB 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone

DCMU 3(3,4-dichlorophenyl)-1,1-dimethylurea

DNA desoxyribonucleic acid

EDTA ethylenediamine tetraacetic acid

FHL formate hydrogen lyase

FNR ferredoxin-NADP-oxidoreductase

GPD glycerol-3-phosphate dehydrogenase

GPP glycerol-3-phosphate phosphatase

HRP horse reddish peroxidase

HydA1 , 2 [Fe]-hydrogenases 1, 2 from C. reinhardtii

IPTG isopropyl-β-thiogalactoside

JGI Joint Genome Institute

KOG clusters of euKaryotic Orthologous Groups

LHC I, II light harvesting complex I, II

MOPS 3-(N-morpholinopropane)-sulfonic acid

mRNA messenger ribonucleic acid

NAD(P)H reduced nicotinamide adenine dinucleotide(phosphate)

NCBI National Center for Biotechnologial Information

Ndh1 rotenone-sensitive class I NAD(P)H dehydrogenase

Ndh2 rotenone-insensitive class II NAD(P)H dehydrogenase

PAGE polyacrylamide gel electrophoresis

PAM pulse amplitude modulating

PBS phosphate buffered saline

PCR polymerase chain reaction

PDC pyruvate decarboxylase

PetF algal ferredoxin

PFL pyruvate formate-lyase

PflA PFL activating enzyme, PFL activase

PFO pyruvate ferredoxin/flavodoxin-oxidoreductase

Abbreviations

PQ plastoquinone

PSI, PSII photosystem I, II

PTA1, 2 phosphotransacetylase 1, 2

PTOX plastidic terminal oxidase

QA, QB primary, secondary electron acceptor of P680 (PSII)

RACE rapid amplification of cDNA ends

RNA ribonucleic acid

ROS reactive oxygen species

RT-PCR reverse transcriptase PCR

Rubisco ribulosebisphosphate-carboxylase/oxygenase

SAM S-adenosylmethionine

SDS sodium dodecylsulphate

SHAM salicyl hydroxamic acid

SQDG sulphoquinovosyl diacylglyceride

TAE Tris-acetate-EDTA

TAP Tris-acetate-phosphate

TdcE E. coli 2-ketobutyrate formate-lyase

Units of the International System of Units (SI) are not separately listed.

Gene designations

gene encoded protein

ack acetate kinase

adh1 alcohol- and acetaldehyde-dehydrogenase, PFL deactivase (Adh1/AdhE)

aox1, 2 alternative oxidase (AOX) 1, 2

atpB subunit of plastidic ATPase

coxI subunit COXI of mitochondrial cytochrome oxidase

cox90 subunit COX90 of mitochondrial cytochrome oxidase

hydA1, 2 [Fe]-hydrogenases HydA1, 2

pdc pyruvate decarboxylase

petA cytochrome f apoprotein of cytochrome b6f complex

pfl pyruvate formate-lyase (PFL)

pflA PFL activating enzyme, PFL-activase

psbA core protein D1 of PSII

pta 1, 2 phosphotransacetylase (PTA) 1, 2

rbcL large subunit of Rubisco (RbcL)

sac1 regulator of specific responses to sulphur-deprivation (SAC1)

Introduction

1

1 Introduction

Research on the unicellular chlorophyte alga Chlamydomonas reinhardtii (fig 1-1) is

almost one century old now, and this organism, sometimes called the “photosynthetic

yeast” (Rochaix, 1995), has become an important model (Harris, 2001). A simple life cycle

that can be easily manipulated, rapid growth and a haploid genome in vegetative cells are

characteristics that make this alga an ideal model system. Although the Volvocales are

considered to be a side-branch of the phylogenetic tree leading to higher land plants

(Chapman and Buchheim, 1992), the photosynthetic apparatus is nevertheless highly

conserved, making C. reinhardtii an excellent model system for the elucidation of

photosynthesis in vascular plants. Due to its ability to grow heterotrophically on acetate,

C. reinhardtii is useful for the study of photosynthesis and chloroplast biogenesis, since

photosynthetic mutants are viable on acetate (Harris, 1989). C. reinhardtii, however, also

has its very own mysteries, since it possesses features that are quite unusual for a

eukaryotic photosynthetic organism. This alga differs from other eukaryotes by having a

complex fermentative metabolism that is

marked by the production of hydrogen gas and

formate.

Fig 1-1: Light microscopic photograph of the unicellular green alga C. reinhardtii (A. Kaminski, Rheinische Friedrich-Wilhelms-Universität Bonn).

1.1 Hydrogenases and hydrogen metabolism in unicellular green algae

In the first half of the last century it was observed that the unicellular green alga

Scenedesmus obliquus, after adaptation to anaerobic conditions, develops a hydrogen

metabolism. It can use the reductive power of hydrogen for the assimilation of carbon

dioxide (Gaffron, 1939), and it is also able to photoproduce hydrogen (Gaffron and Rubin,

1942). Other investigators observed photosynthetic hydrogen evolution in anaerobic

cultures of Chlorella fusca (Spruit, 1958) and Chlamydomonas moewusii (Frenkel, 1952).

In 1974, it was concluded that many species of unicellular green algae have a hydrogen

metabolism (Kessler, 1974). One of the most efficient hydrogen producers was

10 µm

Introduction

2

C. reinhardtii (Stuart and Gaffron 1972; Ben-Amotz et al., 1975). Its hydrogenase enzyme,

HydA1, was purified to homogeneity and biochemically characterised in the 1990s (Happe

and Naber, 1993). It turned out to be a small iron-containing protein of 48 kDa, which is

localised in the chloroplast stroma. In all likelihood, the photosynthetic ferredoxin PetF is

the natural electron donor to the hydrogenase (Happe et al., 1994). HydA1 is very sensitive

to molecular oxygen and only synthesised under anaerobic conditions. Hydrogenase

activity is detectable very soon (~ 5 min) after oxygen has been removed from a

C. reinhardtii culture and Northern blot analyses indicated transcriptional regulation of the

hydA1 gene in response to anaerobiosis (Happe and Kaminski, 2002). More recent research

using reporter gene assays confirmed these findings and showed that the hydA1 promoter is

regulated in response to levels of oxygen within the cells (Stirnberg and Happe, 2004).

Indeed, isolation of the hydA1 gene, which had proved unsuccessful until recently, was

achieved by making use of the anaerobic expression of hydA1. Applying the method of

suppression-substractive hybridisation PCR, which is a tool to identify differentially

expressed genes, a cDNA sequence encoding the [Fe]-hydrogenase of C. reinhardtii could

be identified in total mRNA isolated from an anaerobic algal culture (Happe and Kaminski,

2002). The deduced amino acid sequence of HydA1 revealed the highly conserved

C-terminal domain characteristic for [Fe]-hydrogenases, including four cysteine residues

that coordinate the active site, the so-called H-cluster (Peters et al., 1998).

Based on these fundamental data, the hydA genes of further algal species were isolated in

the following years (Florin et al., 2001; Winkler et al., 2002a; Winkler M., personal

communication). It turned out that the hydrogenase enzymes of unicellular green algae

represent a novel type of [Fe]-hydrogenases (Happe et al., 2002). Chlorophycean type

[Fe]-hydrogenases are small proteins (44 to 48 kDa) and lack the N-terminal ferredoxin-

like domain present in all other [Fe]-hydrogenases isolated to date (Vignais et al., 2001).

This so-called F-domain harbours two or more additional Fe-S-clusters which are thought

to be involved in the transfer of electrons between the external electron donor / acceptor

and the catalytic centre (Peters et al., 1998; Nicolet et al., 1999). The absence of any

additional redox clusters in the hydrogenases of green algae indicates a novel electron

transfer pathway for these enzymes and suggests direct electron transfer between the

electron donor and the H-cluster of HydA1. Protein modelling studies indicate a single

positively charged binding site in an otherwise negatively charged molecule. A polypeptide

stretch in the C-terminal part of the primary structure, which is unique for algal

Introduction

3

hydrogenases, appears to form a positively charged loop. This loop is proposed to support

the electrostatic interaction with ferredoxin (Winkler et al., 2002a) (fig 1-2).

Fig 1-2: Model of the hypothetical interaction between C. reinhardtii HydA1 and ferredoxin PetF (Winkler M., personal communication). The model of HydA1 was generated on the basis of the known three-dimensional structure of the [Fe]-hydrogenase CpI of Clostridium pasteurianum (Peters et al., 1998). C. reinhardtii PetF was modelled according to the crystal structure of the respective Chlorella fusca protein (Bes et al., 1999). The left side of the graphic (A) shows the modelled ribbon structures of HydA1 and PetF. The right side of the drawing (B) displays the surface charge of the enzymes (red = negatively, blue = positively charged). The positively charged, blue coloured binding site is discernible on the lower surface of HydA1 (B).

Interestingly, all unicellular green algae whose hydA genes have been isolated possess a

second hydrogenase gene. To distinguish between the two very similar genes, they were

named hydA1 and hydA2 (Forestier et al., 2003). HydA1 was the first hydrogenase to be

isolated from C. reinhardtii (Happe and Naber, 1993). The function of HydA2 has yet to

be elucidated.

The metabolic function of algal hydrogenases was unclear for a long time. The occurrence

of an oxygen-sensitive enzyme in a photosynthetic and thus oxygen-producing organism

was regarded as a biological curiosity. Nevertheless, research on this paradoxical feature of

unicellular green algae has continued and is revealing more and more details about the

complexity of photosynthetic electron transport pathways.

Introduction

4

After the light dependence of algal hydrogen metabolism was demonstrated (Gaffron,

1939; Gaffron and Rubin, 1942), the participation of the two photosystems in light-driven

hydrogen evolution was analysed in C. reinhardtii. In anaerobically adapted cells,

hydrogen production was observed in the presence of DCMU, an inhibitor of electron

transport at photosystem II (PSII), but it was not detected after application of DBMIB,

which blocks electron transport at the site of the cytochrome-b6f-complex (Stuart and

Gaffron, 1972). Despite the fact that PSII is dispensable for photo-hydrogen evolution, it

can nevertheless function as an electron source for the hydrogenase (Gaffron and Rubin,

1942; Bishop and Gaffron, 1963), resulting in the simultaneous generation of hydrogen and

oxygen (Spruit, 1958; Greenbaum et al., 1983; Greenbaum and Lee, 1998). However, this

process acts only transiently (60 – 90 s), because the hydrogenase is rapidly inactivated by

photosynthetically produced oxygen (Ghirardi et al., 1997). A second pathway that

provides electrons for photo-hydrogen production couples the oxidative degradation of

organic substrates with photosynthetic electron transport by transferring electrons from

NAD(P)H to the plastoquinone (PQ) pool (Bamberger et al., 1982; Gfeller and Gibbs,

1985; Godde and Trebst, 1980). The latter process probably depends on the so called

chlororespiratory pathway.

The concept of a respiratory chain in the chloroplast is more than 40 years old (Goedher,

1960; Bennoun, 1982). It suggests at least two additional electron transport components

able to exchange electrons with the photosynthetic electron transport chain, commonly

accepted as the Z-scheme (Trebst, 1974). One of these additional components, the

NAD(P)H-plastoquinone oxidoreductase, performs a non-photochemical reduction of the

PQ-pool using stromal reductants. The second enzyme is a quinole-to-oxygen

oxidoreductase that is able to oxidise PQ by transferring electrons to oxygen. Dark

oxidation of the PQ pool was observed in the early 1960s. It was proposed to occur

through “some kind of chloroplast respiration” (Goedher, 1963). This hypothesis was

supported by Bennoun (Bennoun, 1982), who analysed the effects of respiratory inhibitors

on chlorophyll fluorescence. Non-photochemical reduction of the PQ-pool was described

in the early 1980s (Godde and Trebst, 1980; Godde, 1982).

More recent work using genetic and biochemical approaches provided the model of

chlororespiration. In several higher plant chloroplasts, functional NAD(P)H dehydrogenase

complexes (called Ndh1), which are homologous to bacterial or mitochondrial respiratory

complex I, have been characterised (Cuello et al., 1995; Guedeney et al., 1996; Sazanov et

Introduction

5

al., 1998; Casano et al., 2001). Most subunits of Ndh1 are encoded by the plastid genome

(Ohyama et al., 1986; Ohyama et al., 1988; Sugiura, 1992). Some years ago, an oxidase

capable of PQ oxidation was shown to be inactivated in the Arabidopsis thaliana mutant

immutans (Carol et al., 1999; Wu et al., 1999). This oxidase shows similarity to

mitochondrial alternative oxidases (Wu et al., 1999) and was termed plastid terminal

oxidase, PTOX (Cournac et al., 1998).

Heterologous synthesis of the IMMUTANS (IM) protein in Escherichia coli revealed it to

be a cyanide-resistant quinol oxidase that is sensitive to the alternative oxidase inhibitor

propyl gallate (Cournac et al., 2000). The same study showed that chlororespiratory

oxygen uptake by C. reinhardtii is also inhibited by propyl gallate. In thylakoid

membranes of the alga, a protein of 43 kDa cross-reacted with an antibody raised against

the recombinant A. thaliana IM-protein. Later, the sequence of a putative quinol-to-oxygen

oxidoreductase of C. reinhardtii was submitted to the PubMed database (AAM12876;

Chen H.-C. and Melis A., direct submission, 2002). The deduced molecular mass of the

pre-protein is 54 kDa.

However, genes encoding subunits for the Ndh1 complex could not be identified in the

chloroplast genome (Maul et al., 2002), or in the recently published genome sequence of

C. reinhardtii (www.jgi.doe.gov/, C. reinhardtii release 2.0, February 14th 2004). Very

recent research indicates that the NAD(P)H-PQ oxidoreductase of C. reinhardtii belongs to

the monomeric, rotenone-insensitive class II NADH dehydrogenases (Ndh2), which are

found for example in yeast (Small and Mc Alister-Henn, 1998). NAD(P)H-dependent

reduction of the plastoquinone pool and light-dependent hydrogen evolution in

C. reinhardtii are insensitive to rotenone, the classic inhibitor of class I NAD(P)H

dehydrogenases, but are sensitive to several compounds known to inhibit rotenone-

resistant class II NAD(P)H-ubiquinone oxidoreductases [e.g., to the flavoprotein inhibitor

diphenyleneiodonium (DPI) or flavone] (Mus et al., 2005). In the C. reinhardtii genome,

seven sequences that encode Ndh2-like proteins have been identified. Several of them

contain a putative chloroplast transit peptide (Mus et al., 2005). Fig 1-3 summarises the

current model of the involvement of HydA1 in the photosynthetic electron transport chain.

It has been suggested that the chlororespiratory pathway originated from the cyanobacterial

ancestors of chloroplasts (Scherer, 1990). In today’s cyanobacteria, respiratory pathways

are also found in the thylakoid membranes, where they share redox proteins with the

Introduction

6

photosynthetic apparatus (Scherer, 1990). In chloroplasts, one possible function of

chlororespiration could be the reoxidation of NADPH originating from plastid starch

breakdown (Singh et al., 1992). Generally, the plastid oxidase seems to be significantly

engaged when the plastoquinone pool is highly reduced (Cournac et al., 2002).

Fig 1-3: Schematic diagram of the photosynthetic electron transport chain and the proposed

involvement of the hydrogenase (HydA1). Electrons can be provided by the splitting of water at PSII and these are transported via the PQ-pool, cytochrome b6f complex (Cyt b6f), plastocyanine (PC), PSI and ferredoxin (Fd) to HydA1. Ferredoxin is also an electron donor to ferredoxin-NADP-oxidoreductase (FNR). The resulting NADPH can be used for reductive biosynthetic processes, e.g. carbon dioxide assimilation. Additionally, electrons originating from oxidative degradation of organic matter (e.g. starch) can be transferred to the photosynthetic chain. This is catalysed by a putative NADPH-PQ-oxidoreductase (Ndh). A plastid terminal oxidase (PTOX) can use electrons from the PQ-pool to reduce oxygen.

1.2 Hydrogen production of sulphur-deprived C. reinhardtii cells

The chlororespiratory pathway was assumed to play a major role in the so-called two stage

hydrogen production process of C. reinhardtii, which was discovered by Melis and co-

workers in 2000 (Melis et al., 2000). This process suggested an important physiological

role for the hydrogenase, since hydrogen metabolism developed without artificial

manipulation and was sustained for several days. The key feature of this metabolism is a

form of metabolic switch that causes an algal culture to change its physiology from aerobic

photosynthetic growth to an anaerobic resting state accompanied by hydrogen evolution.

The trigger for this radical physiological change is the absence of any exogenous source of

sulphate.

PQ

NADPH NADP+

Ndh

starch

O2 H2O

PTOX

H2O O2

PSII Cytb6f

PSI

PC

Fd

NADP+ NADPH

FNR

H+

H2

stroma

lumen

HydA

Introduction

7

Sulphur is an essential constituent of proteins (methionine, cysteine), lipids (above all in

sulfoquinovosyl diacylglycerides, which account for 5 % of the thylakoid lipids),

coenzymes (e.g. coenzyme A or lipoamide), molecules involved in photoprotection

(glutathione) and electron carriers (e.g. in Fe-S clusters). In most cells, there is no specific

sulphur storage, thus the growth of an organism mostly depends on sulphur that is

available externally. C. reinhardtii probably senses sulphur limitation by the sac1-gene

product (sac stands for sulphur acclimation) (Davies et al., 1994, 1996), a putative

regulatory ion transporter. The algal reactions to limited sulphate include the synthesis of

high-affinity sulphate transporters (Yildiz et al., 1994), the release of an extracellular

arylsulfatase capable of cleaving sulphate from aromatic compounds (Lien and Schreiner,

1975), and the enhanced expression of enzymes involved in the sulphate assimilation

pathway (Yildiz et al., 1996; Ravina et al., 1999). Furthermore, the cells economise their

use of sulphate. They synthesise new, almost sulphur-free cell wall proteins (Takahashi et

al., 2001) and make internal sulphur resources available, e.g. by degrading sulpholipids

(Sato et al., 2000). General responses of C. reinhardtii to nutrient deprivation are cessation

of growth (Lien and Knutsen, 1973; Davies et al., 1996), the accumulation of starch (Ball

et al., 1990) and a decrease in and alteration of photosynthetic activities (Wykoff et al.,

1998).

The latter is the main reason for the development of anaerobiosis in sulphur-deprived

C. reinhardtii, and it is a good example of how photosynthetic organisms can adjust their

rate of photosynthesis (and concomitantly the formation of reactive oxygen species)

according to the environmental conditions. A detailed analysis of the effect of

macronutrient starvation on photosynthesis in C. reinhardtii revealed that the light-

saturated rate of photosynthesis, as measured by oxygen evolution, declined by 75 % after

one day of sulphur deprivation and by 90 % after two days (Wykoff et al., 1998).

Phosphorous starvation also caused a reduction in photosynthesis, but its effect was much

less dramatic. Oxygen evolution rates decreased gradually and were reduced by 75 % only

after four days. The decrease in photosynthesis in cells starved for sulphur (examined after

one day) or phosphorus (four days) was proposed to be due to a reduction in

photosynthetic electron transport, and not to a lowered efficiency of the Calvin cycle. The

decline of photosynthetic electron transport, in turn, was essentially caused by a loss of

PSII activity. Quantification of PSII complexes indicated that less than half of the decrease

of in vivo oxygen evolution was due to a reduced number of PSII centres. Rather, so-called

Introduction

8

QB non-reducing centres – PSII centres capable of charge separation but unable to reduce

the PQ pool and non-productive in water splitting (Ort and Whitmarsh, 1990; Melis, 1991)

– accumulated. Whereas during nutrient replete growth, these centres accounted for 29 %

of total PSII centres, they accumulated up to 65 % in sulphur starved C. reinhardtii cells.

Furthermore, light-energy was directed away from PSII by state transition and the

accumulation of xanthophylls. The cells changed from state 1, in which light energy is

mostly directed to PSII, to state 2, in which excitation is mostly focussed on PSI (for

details see paragraph 2.7.4). On a chlorophyll a basis, the amount of antheraxanthin,

zeaxanthin and lutein strongly increased (Wykoff et al., 1998).

All these events leading to dissipation of light energy and a significant decrease in oxygen

evolution are critical for survival of sulphur-depleted C. reinhardtii cells. This is shown by

the sac1 mutant of C. reinhardtii, which is defective in both specific and general responses

to sulphur starvation (Davies et al., 1994, 1996). The mutant dies within two days after

transfer to sulphur-limited medium. This phenotype, however, can be rescued by keeping

the cells in the dark or by adding the PSII inhibitor DCMU, indicating that the decreased

viability of this strain is due to enhanced synthesis of reactive oxygen species or altered

redox conditions (Davies et al., 1996).

In view of the dramatic decrease in photosynthetic oxygen evolution, it was reasoned that a

sealed, sulphur-deprived C. reinhardtii culture would become anaerobic due to respiratory

activity and might induce the hydrogenase pathway. Indeed, sulphur-depleted algal

cultures start to evolve hydrogen within one or two days after sulphur has been withdrawn

from the medium (fig 1-4), and hydrogen production is maintained for several days (Melis

et al., 2000). Analysis of the three photosynthetic complexes PSII, cytochrome b6f complex

and PSI confirmed that the number of active PSII centres strongly decreases, while the

activity of the other two complexes declines only gradually. The algal cells accumulate

starch and proteins in the first one or two days of sulphur deprivation, but upon the onset of

anaerobiosis and hydrogen production, both starch and proteins are degraded again. It was

proposed that electrons for H+ reduction originate from degradation of these organic

reserves, and that they could be transferred to the photosynthetic electron transport chain

via the chlororespiratory pathway (Melis et al., 2000; Zhang et al., 2002). Hydrogen

evolution in sulphur depleted C. reinhardtii is accompanied by carbon dioxide production,

further supporting the assumption that degradation of organic matter is linked with

Introduction

9

hydrogen production. In contrast, anabolic metabolism seems to be greatly reduced. The

cells cease to grow, and ribulose-bisphosphate carboxylase/oxygenase (Rubisco) is

degraded. After 24 h of sulphur starvation, the amount of Rubisco, quantified by Western

blot analyses with antibodies raised against the large subunit, is almost halved. After 48 h,

no more Rubsico protein can be detected (Zhang et al., 2002). The Rubisco protein, which

is rich in sulphurous amino acids, might serve as an intracellular sulphur source, as

suggested for Lemna minor (Ferreira and Teixeira, 1992).

Fig 1-4: Photograph of a C. reinhardtii culture deprived of

sulphur. Little hydrogen bubbles are visible at the side of the bottle.

The “single organism, two stage hydrogen production

method” (Melis et al., 2000), separating photosynthetic

oxygen evolution and carbon dioxide assimilation (stage

1) from hydrogen production at the expense of

accumulated carbon (stage 2), has attracted much interest. Besides suggesting an important

physiological role for the algal hydrogenase, it holds the promise of being a possible

renewable energy source, which uses sunlight as the main supply of energy, and water,

directly or indirectly, as the donor of electrons (Melis and Happe, 2001). Furthermore, this

complex algal metabolism is an excellent model for studying the interactions of several

metabolic functions, which cooperate to enable C. reinhardtii to acclimate to the

unfavourable conditions of sulphur starvation.

Two features of the hydrogen metabolism of sulphur deprived C. reinhardtii cells have

received only minor attention: the first is the role of acetate, and the second the function of

mitochondrial respiration. It became quickly apparent that sulphur depletion does not lead

to hydrogen generation when the cells are cultivated photoautotrophically, without acetate

as organic carbon source (Hemschemeier, 2002). C. reinhardtii can assimilate acetate and

incorporate it into cellular compounds, probably via both the citric acid cycle and the

glyoxylate cycle (Gibbs et al., 1986). This alga belongs to the so-called “acetate

flagellates”, which live in environments that are rich in organic compounds, where oxygen

can be limited and carbon dioxide concentrations high (Pringsheim, 1937). Indeed, the

Introduction

10

ancestor of the common laboratory strains of C. reinhardtii was isolated in 1945 from

nutrient-rich soil collected near Amherst, Massachusetts by G.M. Smith (Smith, 1946). It is

common to culture C. reinhardtii in acetate-containing medium, although it is known that

acetate has a significant influence on several metabolic activities. Most obvious are the

inhibitory effects of acetate on photosynthesis (Endo and Asada, 1996, Heifetz et al., 2000)

and the expression of nuclear-encoded genes involved in photosynthesis (Goldschmidt-

Clermont, 1986; Kindle, 1987). Furthermore, stimulation of respiratory oxygen uptake by

acetate was reported (Fett and Coleman, 1994; Endo and Asada, 1996). In sulphur depleted

C. reinhardtii, absolute photosynthetic oxygen evolution is higher and respiratory rates are

lower in the absence of acetate, which prevents the establishment of anaerobic conditions.

If acetate is added to an autotrophic sulphur-deprived culture, anaerobic conditions and a

concomitant hydrogenase activity are established within a couple of hours (Hemschemeier,

2002).

The key for the establishment of anaerobic conditions is the removal of oxygen. Because

photosynthetic activity decreases upon sulphur starvation, respiratory oxygen uptake

exceeds oxygen production, resulting in net oxygen consumption (Melis et al., 2000). It is

remarkable that respiratory activity remains high during the progressive adaptation of

C. reinhardtii to limitations in sulphur. As an alga, C. reinhardtii has several possibilities

to reduce oxygen, either in the mitochondrium or in the plastid. Mitochondrial oxygen

uptake is catalysed by complex IV (cytochrome oxidase), which allows ATP-generating

respiratory electron transport, or by alternative oxidase (AOX), which bypasses the proton

pumping complexes III and IV. AOX is believed to catalyse a form of short circuit,

allowing the disposal of excess electrons, often accompanied by heat production (Moller,

2001). C. reinhardtii possesses two aox genes. The aox1 gene is transcribed at higher

levels than aox2. Generally, the capacity for alternative respiration is constitutive in

C. reinhardtii (Goyal and Tolbert, 1989). Upon sulphur deprivation, transcription of aox1

is down-regulated, while the transcript level of coxI, a subunit of mitochondrial

cytochrome oxidase, remains constant (Hemschemeier, 2002).

Plastid terminal oxidase could constitute a third possibility to consume oxygen. Research

on this topic in C. reinhardtii is comparatively recent, thus little is known about the

regulation and activity of PTOX in this model alga. It is currently unknown how the

respiratory activities in sulphur starved C. reinhardtii cultures are regulated.

Introduction

11

1.3 Fermentation: anaerobic energy production

In the 1980s it was shown that hydrogen production in C. reinhardtii is embedded in a

complex fermentative metabolism, which is another exceptional characteristic of this green

alga. A detailed analysis of the anaerobic starch breakdown in C. reinhardtii revealed that

formate, acetate, ethanol, carbon dioxide, hydrogen, as well as traces of glycerol and D-

lactate are produced (Gfeller and Gibbs, 1984). This fermentative pattern is more similar to

the mixed acid fermentation of some bacteria than to the rather simple anaerobic

metabolism of higher plants (Kennedy et al., 1992). The production of formate was

ascribed to pyruvate formate-lyase (PFL), an enzyme previously thought to be restricted to

prokaryotes. In view of the severe anaerobiosis established in sulphur-deprived

C. reinhardtii, the culture medium was analysed for soluble fermentation products. Indeed,

significant amounts of ethanol and formate were detected. Furthermore, the accumulation

of these products was accompanied by strongly increased levels of the pfl transcript

(Hemschemeier, 2002; Winkler et al., 2002b). These findings showed that sulphur-starved

C. reinhardtii cells perform a marked fermentative metabolism, which was termed photo-

fermentation because it develops upon full illumination in an organism performing

oxygenic photosynthesis.

Generally, the term fermentation describes the oxidation of organic compounds in the

absence of an appropriate exogenous electron acceptor. Classic fermentation is usually

based on glycolysis, in which glucose is oxidised to pyruvate. Phylogenetically, glycolysis

is the oldest energy-producing bioprocess, which evolved in times when the earth’s

atmosphere did not contain oxygen. For most animal and plant cells, glycolysis is a prelude

to further oxidative processes in the tricarboxylic acid cycle, followed by mitochondrial

respiration. In the absence of oxygen, however, substrate-level phosphorylation is still the

major ATP source for many organisms. Electrons that result from these oxidative

processes have to be dissipated, and NAD+ as the electron acceptor has to be recycled

continuously. For organisms that are adapted to aerobic life, fermentation is always a

compromise between maximal energy yield and maintenance of redox balance. The

tolerance of aerobic organisms towards anaerobiosis is thought to correlate with the type of

fermentative products and with the energy yield. Some fermentative products, such as

ethanol, can become toxic to cells, while others, like lactate, acidify the cytoplasm

(Roberts et al., 1984a, 1984b). In organisms adapted to aerobic growth, the amount of ATP

that can be generated by fermentative pathways usually does not suffice to maintain

Introduction

12

membrane gradients and to recycle essential membrane compounds, thereby leading to a

loss of homeostasis.

Since organisms with an oxygenic photosynthesis produce oxygen, they are commonly not

associated with anoxia and fermentation. However, there are many situations in which

photosynthetic cells encounter anaerobiosis. This is the case for water-living organisms

like algae or cyanobacteria during the night, when photosynthesis does not take place and

oxygen is removed by respiratory activity. In higher plants, roots and seeds can become

anaerobic in waterlogged soils. Most of the higher plants are relatively intolerant to

hypoxia (Kennedy et al., 1992). Under anaerobic conditions, cotton roots survive only for

30 – 60 min, potato tubers and maize roots for about one day. Higher plants usually carry

out ethanol or lactate fermentation (Kennedy et al., 1992) and it is only some wetland

species that are adapted to survive and even grow in hypoxia. The best known example is

probably rice (Oryza sativa), which performs a complex fermentative metabolism and

produces ethanol, alanine, γ-amino butyrate, succinate and lactate (Fan et al., 1993). Later

it was proposed that the unusual flood tolerance of rice could be a consequence of its

ability to carry out nitrate respiration, a process consuming additional NADPH and protons

(Fan et al., 1997).

1.4 The pyruvate formate-lyase system in E. coli

The fermentative metabolism of C. reinhardtii is not comparable to that of higher plants,

not even to that of very hypoxia tolerant plants, such as rice. Rather, the anaerobic

production of formate is typical for facultative anaerobic bacteria. Usually, formate and

ethanol production are associated with the activity of PFL. Probably the best studied PFL

system is that of E. coli. The fermentation products of E. coli are ethanol as well as acetic,

formic (hydrogen and carbon dioxide), lactic and succinic acid (Clark, 1989). They have

different oxidation states and can be produced in variable amounts to modulate

fermentative metabolism. The conversion of pyruvate into acetate and ethanol “is the

backbone of the cellular machinery for the anaerobic life of E. coli” (Kessler and Knappe,

1996), and the enzyme which catalyses the first step of anaerobic pyruvate degradation,

PFL, is a key element of glucose fermentation. Fig 1-5 shows an overview of the PFL-

catalysed pyruvate fermentation in E. coli.

Introduction

13

Fig 1-5: Schematic representation of the

PFL fermentation pathway in E. coli.

Pyruvate formate-lyase (PFL) cleaves pyruvate, originating from glycolytic glucose degradation, into formate and acetylCoA (CoA, coenzyme A). Formate can be disproportionated to carbon dioxide and hydrogen by the formate hydrogen lyase complex (FHL). AcetylCoA can be reduced to ethanol by the multifunctional AdhE protein (combining acetaldehyde dehydrogenase and alcohol dehydrogenase activities), or converted to acetate via phosphotransacetylase (PTA) and acetate kinase (ACK).

PFL cleaves pyruvate into acetylCoA and formate. The synthesis of PFL is increased 10 to

15-fold in the absence of oxygen. Regulation of this process occurs at the transcriptional

level (Sawers and Böck, 1988; Sawers and Suppmann, 1992). PFL is a homodimer

comprising two polypeptides of 759 amino acids (fig 1-6). It is a radical enzyme, whose

active form harbours an α-carbon-centered glycyl radical at position 734 in the polypeptide

chain (Knappe et al., 1984; Wagner et al., 1992). The radical is stable under anaerobic

conditions, but oxygen causes a specific scission of the polypeptide backbone at position

734. The catalytic cycle of PFL comprises two distinct half reactions. First, pyruvate is

cleaved, formate is released and the acetyl moiety is covalently bound to the enzyme at

Cys-418 (Knappe et al., 1974). It is assumed that the adjacent Cys-418 and Cys-419

residues form intermediary thiyl radicals that govern the cleavage of the carbon-carbon

bond in pyruvate (Knappe et al., 1993). Probably, the radical is located at Gly-734 in the

resting state of the enzyme, and it is transferred to Cys-419 via Cys-418 upon substrate

binding (Becker and Kabsch, 2002). Then, the acetyl group is transferred to coenzyme A,

and the catalytic cycle returns to its starting point. Both acetyl phosphinate (a pyruvate

analogue) and hypophosphite (a formate analogue) are mechanism-based irreversible

inhibitors of PFL (Knappe et al., 1984).

formate acetylCoA

CO2 + H2

NADH

NAD+ + CoA

NADH

NAD+

ethanol

PFL

FHLAdhE

acetate

Pi

CoA

PTA

ADP

ATPACK

pyruvate

acetaldehyde acetylphosphate

glucose

AdhE

CoA

Introduction

14

Fig 1-6: 3D-model of homodimeric E. coli PFL in complex with the substrate analogue oxamate (structure file from Becker et al., 1999). The structure model was modified with Cn3D 4.1 (NCBI). The oxamate molecule is coloured in green.

The glycyl radical has to be inserted post-translationally into the PFL polypeptide. This is

catalysed by PFL activase (PflA). PflA requires S-adenosylmethionine (SAM) and reduced

flavodoxin as substrates for PFL-activation. PflA is a member of the group of so-called

radical SAM proteins (Sofia et al., 2001). All radical SAM proteins use SAM for reactions

that involve the formation of radicals. They possess a conserved Cys-x-x-x-Cys-x-x-Cys

motif (x stands for any amino acid), which coordinates a 4Fe-4S-cluster. Generally, an

amino acid with an aromatic residue is found adjacent to the third cysteine. A second

conserved region contains a glycine rich sequence resembling the SAM-binding site in

methyltransferases (Niewmierzycka and Clarke, 1999).

There also is a distinct PFL deactivase, which can transform PFL back to its non-radical

form. PFL deactivase is identical to the AdhE protein, which also reduces acetylCoA to

ethanol (see below). In anaerobic E. coli cells, PFL exists almost completely in its radical

form, although large amounts of AdhE are present. Probably, the high intracellular levels

of pyruvate and NADH suppress the PFL deactivase activity of AdhE (Kessler et al.,

1992). Conversely, in aerobically growing E. coli, PFL (constituting about 10% of the

level observed upon anaerobiosis) is present in its non-radical form. This seems to be a

consequence of the lack of dihydroflavodoxin reductant necessary for PFL-activation.

AcetylCoA, which is produced by non-oxidative conversion of pyruvate via PFL, can be

metabolised by two pathways. A reaction catalysed by phosphotransacetylase (PTA) and

acetate kinase (ACK) is one of the major pathways used by fermenting bacteria to generate

Introduction

15

ATP (Thauer et al., 1977). First, acetylCoA is converted to acetyl phosphate by PTA,

replacing coenzyme A by inorganic phosphate. Then, ACK uses the energy-rich acetyl

phosphate bond to transfer the phosphate group on ADP, generating ATP. Both reactions

are reversible, allowing aerobically growing E. coli to use acetate as the sole carbon source

(Brown et al., 1977). PTA and ACK are expressed constitutively.

A second pathway allows the re-oxidation of two molecules of NADH through sequential

reduction of acetylCoA to ethanol via acetaldehyde. Genetic and biochemical analyses

have shown that the acetaldehyde dehydrogenase and alcohol dehydrogenase functions are

localised on one homopolymeric enzyme complex called AdhE (Goodlove et al., 1989). As

mentioned above, a third function of AdhE is the specific quenching of the PFL radical

(PFL deactivase; see above) (Kessler et al., 1991, Kessler et al., 1992). Expression of the

adhE gene is induced by anaerobiosis.

In E. coli, formate can be further disproportionated into carbon dioxide and dihydrogen by

the formate hydrogen lyase (FHL) complex. The intracellular formate concentration is the

crucial determinant for the formation of the FHL complex. When E. coli grows at neutral

pH, formate is excreted from the cells, and the FHL complex is poorly synthesised. When

the extracellular level of fermentative acids increases, the medium pH decreases and

formate is re-imported into the cells, triggering a strong expression of the FHL complex

(Birkmann et al., 1987; Mnatsakanyan et al., 2002; Sawers, 2005).

In C. reinhardtii, a PFL system was proposed when fermentative analyses of dark-adapted

algae revealed a ratio of formate, ethanol and acetate of 2:1:1 (Gfeller and Gibbs, 1984;

Kreuzberg, 1984). The inhibition of formate production by hypophosphite further

supported this assumption (Kreuzberg, 1984). Since then, little research on this exceptional

feature of C. reinhardtii has been conducted.

1.5 Objectives of this work

In the absence of sulphur, the unicellular chlorophyte alga C. reinhardtii changes its

physiology from aerobic assimilatory growth to an anaerobic resting state. The transition

from one to the other condition is caused and accompanied by radical changes of the whole

metabolism. The most remarkable effect of sulphur deprivation is a strong decrease of

photosynthetic activities within one day of sulphur starvation. Since respiratory activity is

Introduction

16

less affected, oxygen uptake exceeds photosynthetic oxygen evolution after a certain point

in time, resulting in sustained anaerobic conditions. Once oxygen is removed from the

algal culture, the cells start to produce hydrogen and also accumulate ethanol and formate.

The drastic physiological changes that occur in the algal cells upon sulphur deprivation are

still unclear in many aspects. Deeper insights into different aspects of this metabolism

would contribute to the understanding of several physiological processes, since all of the

major cellular pathways are concerned. Furthermore, this system aroused the public

interest since the hydrogen yields of sulphur-deprived C. reinhardtii cultures are relatively

high, promising a biotechnological applicability of this system.

This work aims to analyse several aspects of this complex and fascinating metabolism of

C. reinhardtii. It concentrates on the examination of photosynthesis, respiration,

chlororespiration, starch accumulation and degradation, hydrogen production and

fermentation, which all intercommunicate. Fig 1-7 shows a rough summary of the complex

processes that will be analysed in this work.

These physiological processes of C. reinhardtii are to be analysed by a broad spectrum of

methods. The progressive development of sulphur-deprived algae under different

conditions is examined by a special mass spectrometric set-up, which is present in CEA

Cadarache, France, and which allows the coincident determination of the relevant gas

exchange rates. Chlorophyll fluorescence measurements are performed to characterise the

status of the photosynthetic apparatus of the cells. Determination of medium supplements

and cellular metabolites is done to characterise the nutritional status of the cells. Moreover,

several C. reinhardtii mutant strains deficient for central metabolic pathways are examined

by the same methods to get information about the role of the respective pathway. The

major questions addressed by these experiments are:

1. What are the reasons and adaptations that lead to this radical change of physiology?

2. What is the major electron source for hydrogen production by sulphur-deprived

C. reinhardtii?

Moreover, the exceptional fermentation system of C. reinhardtii, which is quite unusual

among eukaryotes, is examined. The production of formate under anaerobic conditions

indicates that C. reinhardtii possesses a bacterial-type fermentation system catalysed by

Introduction

17

pyruvate formate-lyase. This study aims to analyse the PFL system of C. reinhardtii in

physiological, molecular biological and biochemical detail.

Fig 1-7: Highly simplified scheme of the metabolic interactions in sulphur-deprived C. reinhardtii cells. Electrons for hydrogen production by HydA1 can either originate from splitting of water at PSII or from the oxidative degradation of endogenous substrates, such as starch. In the latter case, electrons are transferred to the photosynthetic chain via a putative NADPH-PQ oxidoreductase (Ndh). Photosynthetic electrons might also be used for carbon dioxide fixation in the Calvin cycle via Rubisco (Rbc). Oxygen might be removed from the chloroplast by PTOX, using electrons provided by Ndh or PSII, or by mitochondrial respiration. The latter comprises cytochrome oxidase (COX), allowing the generation of ATP, and alternative oxidase (AOX), allowing for the disposal of electrons. The respiratory substrate could be starch, which is accumulated from inorganic or organic (acetate) carbon sources, or acetate, which is present in the medium. Fermentation via PFL produces formate, which accumulates in the medium, and acetylCoA. The latter can be reduced to ethanol or recycled in respiration. For simplification, the PFL pathway is drawn in the cytoplasm. It is not yet known, where it takes place, but it might be localised in the mitochondrion or the plastid (Kreuzberg et al., 1987).

Materials and Methods

18

2 Materials and Methods

2.1 Organisms and growth conditions

2.1.1 Green algae

Wild type strain C. reinhardtii CC-125 (wild type mt+ 137c) was originally obtained from

the Chlamydomonas Culture Collection at Duke University, Durham, NC, USA. The

following mutant strains of C. reinhardtii were investigated additionally:

name affected gene deficient for reference

FuD7 psbA PSII Bennoun and Delepelaire, 1982

CC-2803 rbcL Rubisco (carbon dioxide

fixation)

Chlamydomonas Center

(www.chlamy.org/)

M.90 cox90 mitochondrial cytochrome

oxidase

Lown et al., 2001

FuD50 atpB plastidic ATPase Chlamydomonas Center

(www.chlamy.org/)

PM9.5A petA cytochrome b6f complex Lown et al., 2001

These mutant strains were a gift from Dr. Saul Purton, University College London, UK.

C. reinhardtii cultures were grown photoheterotrophically in Tris-acetate-phosphate (TAP)

medium (Harris, 1989). Batch cultures were shaken (100 rpm) at 20°C under continuous

illumination (100 µmol photons x m-2 x s-1). Since mutant strains CC-2803 and PM9.5A

are light-sensitive, they were grown under the same conditions as the wild type but covered

with paper towels. For sulphur deprivation, cells were harvested by centrifugation (3 min,

3.500 rpm) in the mid-exponential stage of growth, washed with TAP-S medium (TAP in

which all sulphate compunds are replaced by the chloride counterparts) and resuspended in

TAP-S medium. Cells were placed into squared glass bottles sealed with a gas-tight septum

(red rubber Suba seals 37, Sigma-Aldrich, St. Louis, Mo, USA) and incubated under

continuous one-site illumination of 100 µmol photons x m-2 x s-1 (fig 2-1). In some

experiments, the Rubisco-deficient strain CC-2803 was incubated anaerobically in TAP

medium. In this case, cells were harvested as described above, resuspended in fresh TAP

medium, and placed into the same flasks that were used for sulphur deprivation

experiments.


19

Fig 2-1: Illustration of the incubation conditions of sulphur-deprived C. reinhardtii cultures.

2.1.2 E. coli

The following E. coli strains were used in this study:

strain genotype reference / company

DH5α F‘/end A1 hsd 17 (rk- mk

+) sup E44 thi-1 rec A1 ∆

(laclzyA-argF) u 169 deoR

Raleigh et al., 1989;

Woodcock et al., 1989

BL21 (DE3) pLysS F-, ompT, hsdSB (rB-, mB

-), dcm, gal, λ(DE3), pLysS,

Cmr

Novagen Brand, EMD

Biosciences Inc., Madison,

WI, USA

BL21(DE3) F-, ompT, hsdSB (rB-, mB

-), dcm, gal, λ(DE3) Novagen

BL21(DE3)∆pfl BL21(DE3) carrying a CmR cassette in the pfl-gene Sawers G., personal

communication

234M11 like MC4100 (F- araD139 (argF-lac) U169 ptsF25

deoC1 relA1 flbB530 rpsL 150 λ-1) but ∆act

Ω(act::cat pACYC184) ∆(srl-recA)306::Tn10

Sauter M. and Sawers R.G.

(1990)

E. coli BL21(DE3)∆∆∆∆pfl and 234M11 were thankfully obtained by Dr. Gary Sawers

(Department of Molecular Microbiology, John Innes Centre, Norwich, UK). The pfl-gene

of E. coli BL21∆pfl is disrupted by a chloramphenicol resistance cassette (G. Sawers,

personal communication). Strain BL21∆pfl was used for testing the in vivo activity of

C. reinhardtii PFL, and PflA-deficient strain E. coli 234M11 was used for testing the in

vivo functionality of C. reinhardtii PflA.

E. coli cells were usually grown in liquid LB-medium (25 g x l-1 Luria Broth Base,

Invitrogen Gibco, Carlsbad, CA, USA) or on LB agar plates (32 g x l-1 Luria Agar,

Invitrogen Gibco) at 37°C over night. Liquid cultures were shaken at 130 rpm. Antibiotics

were added to final concentrations of 100 µg x ml-1 (ampicilline), 50 µg x ml-1

(tetracycline), 10 µg x ml-1 (kanamycine), or 25 µg x ml-1 (chloramphenicol).


20

2.2 Plasmids

plasmid description

pGEM-T® Easy cloning vector (Promega, Madison, WI, USA), ampR

pASK-IBA7 vector for recombinant expression of proteins with N-terminal strep-tag in E. coli

controlled by tet-promoter, ampR (IBA, Göttingen, Germany)

pET9a vector for recombinant expression of proteins in E. coli controlled by T7-promoter, kanR

(Novagen)

pAH15 5’-RACE fragment of pflA (618 bps), in pGEM-T Easy

pAH17 C. reinhardtii pfl cDNA with EcoRI restriction sites in pGEM-T Easy

pAH18b C. reinhardtii pfl cDNA with NdeI- and BamHI-restriction sites in pGEM-T Easy

pAH20 3’-RACE fragment of pflA (1,6 kb), in pGEM-T Easy

pAH21 C. reinhardtii pfl cDNA in pASK-IBA7 (pAH17 as donor)

pAH23 C. reinhardtii pflA cDNA amplified with oligonucleotides Act-total-3-nest and Act-total-

5-nest, in pGEM-T Easy

pAH24 C. reinhardtii pflA CDS with Eco31I restriction sites in pGEM-T Easy

pAH25 C. reinhardtii pflA CDS in pASK-IBA7 (pAH24 as donor)

pAH26 C. reinhardtii pflA CDS with NdeI- and Bpu1102I restriction sites in pGEM-T Easy

pAH27 C. reinhardtii pflA CDS in pET9a (pAH26 as donor)

pAH28 C. reinhardtii pfl CDS in pET9a (pAH18b as donor)

pAH29 C. reinhardtii pfl CDS without putative signal peptide coding sequence with NdeI- and

BamHI-restriction sites in pGEM-T Easy

pAH30 C. reinhardtii pflA CDS without putative signal peptide coding sequence with NdeI- and

Bpu1102I restriction sites in pGEM-T Easy

pAH32 C. reinhardtii pfl CDS without putative signal peptide coding sequence in pET9a

(pAH29 as donor)

pAH33 C. reinhardtii pflA CDS without putative signal peptide coding sequence in pET9a

(pAH31 as donor)

pAH34 E. coli pfl cDNA with NdeI- and Bpu1102I restriction sites in pGEM-T Easy

pAH36 E. coli pfl cDNA in pET9a (pAH34 as donor)

pAH45 E. coli pflA with NdeI and Bpu1102I restriction sites in pGEM-T Easy

pAH46 E. coli pflA in pET9a (pAH45 as donor)

Tab 2-1: List of plasmids used or created in this study.


21

2.3 Oligonucleotides

name sequence

PFL-Race-back1 CTCGATCTCGGTGGCAATCA

PFL-Race-back2 AAGGACGAGTTGCCGGCGTA

PFL-Anf-Nde CATATGAGCCAGATGCTGCTGGAG

PFL-pET-ohneSP CATATGCTCCCGGTGGCACCCAG

PFL-End-BamHI GGATCCTTACATGGTGTCGTGGAAGGTG

PFLStrep_ATG GAATTCATGAGCCAGATGCTGCTGGAG

PFLStrep_End GAATTCCATGGTGTCGTGGAAGGTG

PFL-Seq-Mitte TGGTGTTCGCGTACATGGAG

PFL-Ecoli-Nde CATATGTCCGAGCTTAATGAAAAG

PFL-Ecoli-Bpu GCTCAGCTTACATAGATTGAGTGAAGG

Ecoli_PFL_SeqMitte AGGCTTCGTAGCTGTACTTG

PFL-Act-1 GTTCTGCCGCAATACGAGCC

PFL-Act-2 CCTGGAACACGGTGGACACA

PFL-Act-3 CAGCCGAAGCTCTCGACGAA

PFL-Act-4 CCGCGTGGCTTGAGGTAGTT

PFL-Act-5 TTGATGTCCGCCGCGATCTC

PFL-Act-6 TCTCCTTGCTGCTGGTCTTG

Act-IBA7-5 ATGGTAGGTCTCAGCGCATGTTGAGGGCTGCGTTGCCAC

Act-IBA7-3 ATGGTAGGTCTCATATCACTCGGCGCAGATGACGGGAAC

Act-pET-Nde GTAGCATATGTTGAGGGCTGCGTTG

Act-pET-ohneSP CATATGCCTGAGGTTTTCGGAAACG

Act-pET-Bpu-2 GCTCAGCTCACTCGGCGCAGATGAC

Act-total-5 TCTCGACCGCACAAGTGCTA

Act-total-3 TTAAGAACCACGCGCCTGAC

Act-total-5-nest GACCGCACAAGTGCTATGTT

Act-total-3-nest CACACGCCGTTAGTTGAATC

Ecoli_PflA_Nde CATATGTCAGTTATTGGTCGC

Ecoli_PflA_Bpu GCTCAGCTTAGAACATTACCTTATGACC

Tab 2-2: List of oligonucleotides created and used in this study. Underscores indicate non-matching nucleotides that encode specific restriction sites.

2.4 DNA and RNA techniques

Genomic DNA and total RNA were isolated according to standard methods as described

previously (Hemschemeier, 2002). mRNA was isolated with the QIAGEN Oligotex

mRNA Mini Kit (QIAGEN, Hilden, Germany). Gel electrophoresis of DNA occurred in


22

1 % TAE (Tris-acetate-EDTA) agarose gels. RNA was separated in formaldehyde

containing 1 % MOPS agarose gels. RNA samples were treated with a special loading

solution (2 µl 5x MOPS; 3,1 µl 37 % formaldehyde; 8,9 µl formamide per 20 µg of total

RNA) and heated for 10 min at 65°C. Nucleic acids were stained with ethidium bromide

(0,5 µg x ml gel-1). Gels were analysed and photographed with Gel Max from INTAS,

Göttingen, Germany.

Polymerase Chain Reaction (PCR) was performed with Taq-polymerase (Fermentas,

Burlington, Canada) in control experiments and with proof reading PfuUltraTM Hotstart

High-Fidelity DNA-Polymerase (Stratagene, La Jolla, CA, USA) for the accurate

amplification of DNA sequences. Oligonucleotides were obtained from Sigma-Genosys

Ltd (Sigma-Aldrich) and applied in final concentrations of 0,4 µM. cDNA-synthesis was

either performed with the QIAGEN OneStep RT-PCR Kit, excluding the PCR-step, or by

the ProSTARTM Ultra HF RT-PCR System from Stratagene. 5’/3’-RACE experiments

were conducted using the SMARTTM RACE cDNA Amplification Kit (Clonetech

Laboratories, Palo Alto, CA, USA) according to the manufacturer’s recommendations.

Depending on the purity of the PCR-product, the PCR sample was directly used for

ligation, or the respective fragment was purified from an agarose gel using the GFXTM

PCR DNA and Gel Band Purification kit from Amersham biosciences (GE healthcares,

Chalfont, UK). Most often, cDNA sequences were pre-cloned in pGEM-T® Easy

(Promega). Restriction occurred with restriction endonucleases obtained by Fermentas.

Sequencing of PCR-products was conducted by MWG Biotech AG, Ebersberg, Germany.

1-2 µg of DNA were dried and sent to the company. If necessary, 20 µl containing 10

pmoles of a specific oligonucleotide were included.

Original sequence data provided by MWG were analysed with Chromas DNA sequence

analysis software (Conor Mc Carthy, Griffith University, Brisbane, Australia). DNA and

protein sequences were analysed and compared with SECentral (Sci Ed Central for

Windows, Scientific and Educational Software). Specific sequence searches and

comparative blast analyses were done in the PubMed databank of NCBI (National Centre

for Biotechnological Information; www.ncbi.nlm.nih.gov/entrez/query.fcgi). Analyses of

the C. reinhardtii genome were conducted on JGI (DOE Joint Genome Institute)

Chlamydomonas reinhardtii v1.0: http://genome.jgi-psf.org/chlre1/chlre1.home.html and

v2.0: http://genome.jgi-psf.org/chlre2/chlre2.home.html.


23

2.5 Expression of C. reinhardtii pfl and pflA in E. coli

To test the in vivo activity of C. reinhardtii PFL and PflA, the algal pfl- and pflA cDNAs

were cloned into the bacterial expression vector pET9a (Novagen) and introduced into the

E. coli mutant strains BL21∆pfl and 234M11. The pET system was developed for the

expression of recombinant proteins in E. coli. pET vectors were originally constructed in

the 1980s (Studier and Moffatt, 1986; Rosenberg et al., 1987; Studier et al., 1990) and

optimised for easier handling by Novagen. Target genes are under the control of

bacteriophage T7 transcription signals. Therefore, T7-polymerase has to be provided for

expression. E. coli BL21 strains possess a chromosomal copy of the T7-polymerase gene

under the control of lacUV5 promoter (Studier and Moffatt, 1986). The pET system is

usually applied for overproduction of target proteins. T7-polymerase is very selective and

active. After some hours of full induction, the recombinant protein can comprise more than

50 % of the total cell protein. In this study, the system was chosen because of direct

availability. cDNAs encoding C. reinhardtii pfl and pflA were cloned into pET9a, either as

the complete sequence (pfl, pAH28; pflA, pAH27) or in the form of a truncated sequence

which lacked the first 210 bps in case of pfl (pAH32) or the first 267 bps in case of pflA

(pAH33). E. coli pfl (pAH36) and pflA (pAH46) were also cloned into pET9a and served

as controls. The resulting strains are listed in tab 2-3.

plasmid containing sequence transformed strain resulting strain

pAH28 complete C. reinhardtii pfl BL21∆pfl BL21.28

pAH32 truncated C. reinhardtii pfl

(lacking the first 210 bps)

BL21∆pfl BL21.32

pAH36 E. coli pfl BL21∆pfl BL21.36

pAH27 complete C. reinhardtii pflA 234M11 234M11.27

pAH33 truncated C. reinhardtii pflA

(lacking the first 267 bps)

234M11 234M11.33

pAH46 E. coli pflA 234M11 234M11.46

Tab 2-3: List of the plasmids that were used in subsequent expression studies, and the strains that resulted from transformation with each plasmid.

For the examination of anaerobic growth and fermentation of E. coli strains, cells were pre-

cultured in liquid LB-medium over night and then inoculated in 50 ml fresh LB-medium in

100 ml squared Schott bottles. To establish anaerobic conditions, the bottles were closed

with gas-tight septa (red rubber Suba seals 37, Sigma-Aldrich) and flsuhed with argon for


24

10 min. Afterwards, E. coli cultures were incubated anaerobically for several hours.

Samples were taken with a syringe at different points in time to measure the optical density

as well as formate and hydrogen production. Formate and hydrogen were detected as

described below (2.7.1, 2.7.3).

2.6 Western Blot Analyses

To obtain crude protein extracts of algae or bacteria, cells were harvested by

centrifugation. The cell pellet was resuspended in lysis buffer (2 M urea; 0,5 M Tris/HCl

pH 5,8; 20 % glycerol; 7 % SDS; 2 % mercaptoethanol; 0,05 % bromphenol blue) and the

lysate was heated for 5 min at 95°C. After centrifugation for 1 min at high speed, warm

protein extracts were loaded onto the gels. SDS-polyacrylamide-gelelectrophoresis (SDS-

PAGE) was conducted as described before (Laemmli and Favre, 1973) using 10 %

separating and 5 % collecting gels. Proteins were transferred to a nitrocellulose membrane

by electro blotting. Blocking and wash steps were performed in 1x PBS (phosphate

buffered saline; 4 mM KH2PO4; 16 mM Na2HPO4; 115 mM NaCl) with 0,1 % Tween 20

and, in case of blocking, 2 % non fat skimmed milk powder (Biomol Feinchemikalien

GmbH, Hamburg, Germany). The first antibody was recognised by goat-anti-rabbit

antibody conjugated with horse reddish peroxidase (HRP) in a dilution of 1:5.000.

Substrate of HRP was the Super Signal West Dura Extended solution from Pierce

(Rockford, IL, USA). Chemiluminescence was detected by the FluorChem 8800 apparatus

from Alpha Innotech, San Leandro, CA, USA.

antibody specific for dilution reference

rabbit-anti-AdhE E. coli AdhE 1:5.000 Kessler et al., 1991

rabbit-anti-TdcE E. coli TdcE and PFL 1:5.000 Sawers G., personal

communication

rabbit-anti-D1 C. reinhardtii D1 (PsbA) 1:150 Johanningmeier, 1987

Tab 2-4: List of antibodies used in this study. AdhE antibody was a gift from D. Kessler. TdcE antibody was thankfully received from G. Sawers.


25

2.7 Physiological analyses of algal cultures

2.7.1 Quantification of hydrogenase activity

Three different assays were performed to determine hydrogenase activity and hydrogen

accumulation in C. reinhardtii: in vivo and in vitro hydrogenase activity tests as well as

analyses of the gas phase. Hydrogen and other gases were analysed by gas chromatography

(Shimadzu GC-2010, Kyoto, Japan; equipped with a varian capillary column, plot fused

silica 10MX0.32MM, coating mol sieve 5Å) or by mass spectrometry (Prisma QMS 2000,

1-100 amu, quadrupole mass spectrometer from Pfeiffer Vacuum, Asslar, Germany;

mounted on a turbo-molecular pumping station, and equipped with an inlet made from a

UDV 146 metal valve) in case of experiments conducted in CEA Cadarache, France.

A rough indicator for the different phases of the metabolism of sulphur deprived

C. reinhardtii cultures is the presence of oxygen, hydrogen and carbon dioxide in the gas

phase above the cultures. To analyse the composition of the gas phase, 200 µl of gas were

taken from the culture vessel with a gas-tight syringe (SampleLockTM Syringe, Hamilton,

Reno, NV, USA) and injected into the gas chromatograph or the mass spectrometer.

In vivo test were performed to quantify the actual hydrogen producing activity of the cells.

2 ml of algal suspension were transferred to an oxygen free 10 ml headspace bottle closed

with a red rubber Suba seal. The vessel was placed into a shaking water bath (20°C) above

a light source (100 µmoles of photons x m-2 x s-1). The first gas sample was taken after

10 min. The second sample was taken after further 60 min of incubation. The difference

between the hydrogen concentration present in the second and the first sample is the real

hydrogen producing rate of the algae in one hour.

An in vitro test system containing reduced methyl viologen as an artificial electron donor

to the hydrogenase was used to detect active hydrogenase in an algal sample. The reaction

mixture contained final concentrations of 100 mM Na-dithionite, 1 % Triton X-100 and

10 mM methylviologen in 60 mM potassium phosphate buffer pH 6,8. Buffer, triton and

methyl viologen were mixed in a 10 ml headspace bottle and closed with a Suba seal.

Oxygen was removed by flushing the mixture with argon for 5 min. Then, Na-dithionite

was added. 500 µl of cells were taken from the culture vessel with a syringe and injected

into the reaction mixture. Rigorous mixing was performed to disrupt the cells in the

detergent containing suspension. Before incubation, the solution was flushed with argon

for 2 min to remove hydrogen that could be dissolved in the culture. After 20 min of


26

incubation at 37°C, 200 µl of the gas were taken from the reaction vessel and analysed by

gas chromatography.

The hydrogen yields of in vivo and in vitro tests were related to the chlorophyll content of

the respective algal sample. For chlorophyll determination, 800 µl of acetone were added

to 200 µl of C. reinhardtii suspension. After vigorous mixing, the cell debris was spun

down. The extinction of the green supernatant was measured at λ = 652 nm against 80 %

acetone. Total chlorophyll was calculated according to the equation E652 x 27,8 = cChla+b

[µg x ml-1].

2.7.2 Measuring oxygen exchange with a Clark-type electrode

Oxygen exchange activities were measured with a Clark-type oxygen electrode (model

respire 1 from Hansatech, Norfolk, UK). 2 ml of a C. reinhardtii culture were taken from

the sealed incubation bottle using a syringe and injected into the reaction chamber of the

electrode. Photosynthetic oxygen evolution was measured with a slight projector lamp as

light source (100 µmol photons x m-2 x s-1) and respiratory oxygen uptake was detected in

the dark. The rate of each process was recorded for several minutes until a straight line

appeared. The oxygen electrode was calibrated with air saturated water (= 100 % oxygen)

and Na-dithionite treated water (0 % oxygen). The Clark type electrode was used to

perform inhibitor studies on respiratory activities to determine the oxygen uptake capacity

of the three major oxygen consuming pathways in C. reinhardtii. Algae can reduce oxygen

by mitochondrial complex IV (cytochrome oxidase, COX) or alternative oxidase (AOX). A

competent and specific inhibitor of the COX pathway is myxothiazol, which inhibits

electron transfer at the site of the cytochrome bc1 complex. AOX is effectively inhibited by

salicylhydroxamic acid (SHAM). C. reinhardtii also possesses a plastidic terminal oxidase

(PTOX) (Cournac et al., 1998, 2000; PubMed AAM12876) that is inhibited by propyl

gallate (Cournac et al., 2000). The potential participation of each pathway in respiratory

oxygen consumption was determined by recording the dark rate of oxygen uptake in

C. reinhardtii and sequentially adding myxothiazol (2 µM), SHAM (0,4 mM) and propyl

gallate (0,2 mM).

2.7.3 Detection of fermentative products and starch

Fermentative products and acetate were quantified with test kits from Boehringer

Mannheim / r-biopharm, Darmstadt, Germany, following the instructions of the supplier.

The tests make use of the absorbance peak of NADH (but not NAD+) in ultra violet light


27

(340 nm). For detection of soluble fermentation products, a cell sample of 1 ml was spun

down. Ethanol, formate and acetate were quantified in the supernatant using the specific

test kit. Starch was determined as described before (Gfeller and Gibbs, 1984) with slight

modifications. Chlorophylls of 2 ml algae were extracted by acetone. The cell pellet was

washed and finally resuspended in 1 ml Na-acetate buffer (pH 4,5). To disrupt the cells and

solubilise starch, samples were autoclaved for 10 min. Afterwards, starch was determined

with the starch assay kit SA-20 from Sigma-Aldrich in the supernatant.

2.7.4 Chlorophyll fluorescence measurements

Measuring chlorophyll fluorescence dynamics in plant or algal cells in vivo is a fast, non-

invasive technique for the evaluation of electron transfer reactions and energy distribution

within the photosynthetic apparatus (Keren and Ohad, 1998). When a chlorophyll molecule

absorbs a light quantum, an electron is raised from the ground state to an excited state.

Upon de-excitation of the chlorophyll molecule, a small proportion of the absorbed light

energy is dissipated by red fluorescence that can be measured above λ = 680 nm. This

proportion constitutes 3 – 5 % of absorbed energy in vivo. Dissipation of excess energy by

fluorescence occurs mostly at PSII. It competes with the two major ways of energy

dissipation, photochemistry and heat dissipation. The dimension of photochemistry

depends on light-energy as well as on the redox state of intermediate electron carriers and

terminal electron acceptors and therewith on the metabolic state of a plant cell. Energy

dissipation by heat is mainly due to carotenoids and can be modulated by changes in the

membrane carotenoid composition. Generally, fluorescence is highest when

photochemistry and heat dissipation are lowest and vice versa. This fact allows chlorophyll

fluorescence to be an indicator for the state of the photosynthetic apparatus.

An important parameter that influences chlorophyll fluorescence at PSII is a process

commonly known as state transition. It balances light utilisation between the two

photosystems, based on the reversible transfer of a fraction of light-harvesting complex II

(LHCII) from PSII to PSI. The LHCII complex can be associated with PSII (a condition

which is referred to as state 1), so that a big part of light energy is directed towards the

PSII reaction centre and the cells show a high PSII fluorescence. When the mobile fraction

of the LHCII antennae is dissociated from PSII and associated with PSI, cells are in state 2,

and they emit a lower PSII fluorescence. State transition from state 1 to state 2 occurs via

phosphorylation of LHCII proteins by a membrane-bound protein kinase which is activated


28

upon reducing conditions (i.e., when the PQ pool is reduced). Transition back to state 1

occurs upon oxidising conditions (PQ-pool oxidised), which deactivate the kinase and

activate a phosphatase (for more details see Keren and Ohad, 1998).

In higher plants, only a 15-20 % fraction of LHCII participates in state transition. In

C. reinhardtii, a much larger fraction of PSII antennae migrates during state transition

(Bassi and Wollman, 1991), and a much larger decrease in PSII energy capture is observed

(Delosme et al., 1994, 1996). In C. reinhardtii, the cytochrome b6f complex accumulates in

the unstacked thylakoid lamellae in state 2 (Vallon et al., 1991), indicating that state

transition in this alga does not only aim to balance light energy between the two

photosystems, but to favour cyclic electron flow around PSI. It was shown that in maximal

state 2, electrons for reducing the cytochrome b6f complex do not originate from PSII, but

from PSI (Finazzi et al., 1999). This indicates that upon optimal state 2, PSII is not

connected to the remainder electron transport chain. In fact, in C. reinhardtii cells which

are in maximal state 2 conditions, no photosynthetic oxygen evolution can be observed

(Finazzi et al., 1999, 2002). The following table lists several conditions, the according state

and the fluorescence level that can be observed:

condition PQ-pool state fluorescence

light preferentially exciting PSII (λ= 650 nm) reduced state 2 low

light preferentially exciting PSI (λ= 710 nm) oxidised state 1 high

light intensities that saturate the electron flow to

the electron sinks

reduced (Wollman and

Delepelaire, 1984)

state 2 low

aerobic darkness, autotroph oxidised (Wollmann and

Delpelaire, 1984)

state 1 high

darkness, heterotroph reduced (Endo and Asada,

1996)

state 2 low

inhibition of mitochondrial respiratory chain in

the light

reduced (Gans and

Wollman, 1995)

state 2 low

Tab 2-5: List of conditions that influence the state in C. reinhardtii cells.

In this study, three commonly used methods were applied to characterise the status of the

photosynthetic apparatus in sulphur-depleted C. reinhardtii wild type and mutant strains.

Pulse amplitude modulated (PAM) and fluorescence induction measurements were

conducted to evaluate the status of PSII and the degree of photochemistry. Fluorescence


29

emission spectra at low temperature (77 K) were recorded to assess the state of the LHCII

mobile antennae.

PAM measurements show the kinetics of fluorescence in a minute timescale (fig 2-2).

The sample is excited by light of λ < 600 nm and the emitted fluorescence above λ =

680 nm is measured. To excite the sample, the PAM system uses a continuous very weak

modulated light that does not result in the closure of PSII centres (PSII centres are “closed”

when the primary electron acceptor QA is in its reduced state). The fluorescence measured

under this condition is termed F0, and it is defined as the initial fluorescence level

following dark adaptation, when PSII centres are open. Short saturating light pulses of 100

ms result in the maximal fluorescence level Fm, since they cause the transient closure of

all PSII centres. This can also be achieved by adding DCMU, which inhibits the transfer of

electrons between QA and QB, the secondary electron acceptor of PSII, and therefore

results in complete reduction of QA. Upon continuous illumination with actinic, non-

modulated light, the fluorescence reaches the steady state level Ft. Ft is related to the

steady-state electron flow and represents the ratio of closed to open PSII centres. Ft is

higher in dark-adapted than in light-adapted plant cells. This may be due to the fact that

Calvin cycle enzymes have to be activated in the light, so that it needs some time until the

electron-flow reaches the equilibrium.

Fig 2-2: Graph obtained by PAM

measurements of dark adapted

cells. The ground fluorescence F0 is visible after the onset of the measuring beam. A saturating light flash is accompanied by a fast rise of fluorescence to the maximal value Fm. Actinic light results in photochemistry and causes F0 to increase to the steady state level Ft. In light adapted cells, the maximal fluorescence Fm’ caused by a saturating flash is usually lower than in dark adapted cells.

The measurement of fluorescence rise kinetics (fluorescence induction) records

fluorescence in a second scale. It uses a continuous weak actinic light (λ < 600 nm) and

measures the emitted fluorescence above λ = 680 nm. Changes in the fluorescence signal


30

show changes in the redox state of the primary electron acceptor of PSII, QA, which in turn

is affected by the redox status of the PQ pool (Melis, 1991). The fluorescence that can be

detected after the rapid onset of the actinic light is shown in fig 2-3. After the opening of

the shutter, a fast rise of fluorescence is observed which parallels the increasing light

intensity. The changes in fluorescence that follow the complete opening of the shutter are

known as the Kautsky effect (Kautsky and Hirsch, 1931; Govindjee, 1995). Within

0,5 – 1 s, the fluorescence rises to a higher level that depends on the actinic light intensity

and decreases thereafter to a steady-state level (Ft). As described for the PAM

measurements, the intensity rise and subsequent decrease might be due to a form of lag

phase of the Calvin cycle. It cannot be observed in light adapted cells. In fluorescence

induction experiments, the maximal fluorescence level Fm is obtained by the addition of

DCMU.

Fig 2-3: Graph obtained by

fluorescence induction experi-

ments in dark adapted cells. Maximal fluorescence Fm (all PSII centres closed) is reached by the addition of DCMU. Without DCMU, fluorescence is less and follows a typical progress. Fluorescence intensity rises and decreases thereafter to a steady state level Ft.

Fluorescence parameters are relative and depend on light sources, chlorophyll content of

the sample and the measuring apparatus. Therefore, the values are normalised to Fm or Fm’.

Tab 2-6 lists the typical parameters obtained by PAM (or fluorescence induction) and their

meanings:

parameter interpretation calculation

F0 = ground state of fluorescence in very weak modulated light; PSII centres

open (difficult to determine with fluorescence induction measurements)

Fm = maximal fluorescence upon saturating light flash in dark adapted cells;

QA fully reduced, PSII centres closed (in fluorescence induction

experiments obtained by adding DCMU)


31

FV = variable fluorescence; allows determination of maximum quantum

efficiency of PSII primary photochemistry

(Fm – F0) / Fm = FV /

Fm

Ft

= steady state fluorescence upon constant moderate illumination; steady-

state electron flow

Fm’ = maximal fluorescence in light adapted cells; usually lower than in dark

adapted cells; QA fully reduced, PSII centres closed

FV’ = variable fluorescence in light-adapted cells; allows the determination of

open reaction centres in the light

(Fm’ – F0’) / Fm’ =

FV’ / Fm’

Tab 2-6: Parameters recorded by PAM or fluorescence induction measurements, their interpretation and their calculations.

In this study, PAM measurements were conducted with a chlorophyll fluorometer (model

101ED, Walz, Effeltrich, Germany) that was connected to an oscilloscope (Gould Classic

6100, digital storage oscilloscope, 200 MHz). The intensity of the measuring beam was

1 µmol of photons x m-2 x s-1. The actinic light source was a red LED light of 50 µmoles of

photons x m-2 x s-1. The saturation light pulse exceeded 15.000 µmoles of photons x

m-2 x s-1 and was applied for 100 ms.

Fluorescence induction was measured in a laboratory-build set-up at 23°C using a 1 x 1 cm

cuvette with 2 ml sample volume. The light source was a halogen lamp with a filter

combination that produced light with a centre wavelength of 445 nm and an intensity of

100 µmoles photons x m-2 x s-1. Illumination was controlled by an electronic shutter

(opening time 2 ms). Fluorescence was measured at 686 nm with a photomultiplier. The

output of the photomultiplier was recorded with a digital storage oscilloscope (the same

type as was used for PAM measurements) in 2 ms intervals.

Cells were taken from the incubation vessels and directly transferred to the respective

measuring chamber. Since the chlorophyll content does not significantly change during

sulphur depletion (Melis et al., 2000), it was not adjusted to keep the disturbance of the

system as low as possible and not to change the nutritional conditions. In PAM

experiments, cells were first incubated in actinic light for 10 min and F0’, Ft and Fm’ were

recorded. Afterwards, cells were incubated in the dark for 10 min, and F0 and Fm were

measured. For fluorescence induction measurements, two cell samples were taken because

DCMU was included in the analyses. Cells were either pre-incubated in the light or in the

dark for 10 min. Afterwards, fluorescence kinetics was recorded first in the absence and

then in the presence of 2,5 µM DCMU.


32

The recording of fluorescence emission spectra at 77 K allows a great improvement of

the resolution of emission spectra. At room temperature, fluorescence occurs mostly at

PSII. Only at cryogenic temperatures, fluorescence emission of PSI is significant

(Gershoni et al., 1982). In C. reinhardtii cells that are cooled with liquid nitrogen (77 K),

PSI gives a significant fluorescence signal peaking at 715 nm, while PSII fluorescence

appears mostly at 686 nm (fig 2-4). The ratio between fluorescence emission at 686 nm

(F686, PSII) and 715 nm (F715, PSI) is commonly used as an indicator for state transitions.

A decreasing F686/F715 ratio indicates a transition from state 1 to state 2, directing more

excitation energy towards PSI, at the expense of PSII.

Fluorescence emission spectra were determined with a single-beam luminescence

spectrometer (Aminco Bowman Series 2, Spectrometric Instruments, Rochester, NY,

USA) connected to a personal computer containing the software for AB2 luminescence

spectrometer (v4.2). To prepare the cells, chlorophyll content of the respective sample was

determined. Afterwards, a fresh cell sample was taken, adjusted to a chlorophyll content of

5 µg x ml-1 with fresh medium, transferred to a measuring cuvette and directly cooled in

liquid nitrogen. The frozen cuvette was placed into the measuring chamber of the

luminescence spectrometer. The samples were excited at 435 nm, and the fluorescence

emission was recorded between 650 and 750 nm.

Fig 2-4: Characteristic graphs

obtained by recording 77 K

fluorescence spectra. Cells in state 1 show a higher fluorescence at 686 nm (PSII), cells in state 2 show a higher peak at 715 nm (PSI).

2.7.5 Mass spectrometric analyses of the gas exchange in algal cultures

In this study, special mass spectrometric analyses were performed with the help of Laurent

Cournac in the working group of Prof. Gilles Peltier (Départment d’Ecophysiologie

Vegétale et de Microbiologie, CEA Cadarache, France). This working group has

established a unique set-up to record the in vivo gas exchange of a cell suspension almost

without time delay. The merit of this set-up is the possibility to measure several gases at

670 695 720 745 770

emission wavelenght [nm]

flu

ore

sce

nce

(n

orm

alis

ed

)

state 1

state 2

686715


33

once. Therefore, different metabolic processes can be observed at the same time, allowing

a direct comparison without the need to take account for different measuring conditions

and set-ups (e.g. light intensity, temperature, disturbance of the system by entry of air etc.).

The basic principle of mass spectrometry is to produce ions from certain substances, to

separate these ions according to their charge and their mass and to record the abundance of

the respective masses qualitatively and quantitatively with a suitable recording system. The

production of ions from uncharged molecules can occur thermically, electrically or by

bombardment of the sample with electrons, ions or photons. The classical methods of

analyte ionisation are electron ionisation (EI) and chemical ionisation (CI). In case of

electron ionisation, the analyte is fired with high-energy electrons (70 eV) in the nearly

vacuum (~ 10-6 mbar) so that one or two electrons are extracted of the molecule, leaving a

positively charged radical ion (Schröder, 1991). Separation of ions is performed by static

or dynamic electrical or magnetic fields (Kienitz, 1968). Mass spectrometry destroys and

thereby consumes the analyte, but because of its extreme sensitivity, the consumption can

usually be neglected.

The CEA Cadarache working group uses a measuring chamber of the Hansatech electrode

type that is connected to the vacuum of a mass spectrometer (model MM 8-80, VG

instruments, Cheshire, UK) by a Teflon membrane at the bottom of the chamber. Gases

dissolved in the liquid above the membrane diffuse through the thin Teflon layer and are

directly introduced to the ion source of the mass spectrometer through a vacuum line. The

measuring chamber is thermostated by a water jacket, and the liquid is continuously mixed

by a magnetic stirrer. Light is applied by a fibre optic illuminator (Schott, Mainz,

Germany) (fig 2-5).

In this study, sulphur-deprived C. reinhardtii cultures were prepared as described above.

Samples were taken with a gas tight syringe from the incubation bottle and transferred to

the measuring chamber of the mass spectrometer. To keep the entry of air as low as

possible, nitrogen was blown above the cells during injection. After the closure of the

chamber, it was waited until the cells had stabilised, that means, until the curves for each

recorded gas were stable. Cells were continuously illuminated with 100 µmoles of photons

x m-2 x s-1.


34

Fig 2-5: Photograph (left) and schematic draw (right) of the measuring chamber connected to a mass spectrometer in CEA Cadarache, France. Note that the light source has not been included into the scheme. It is attached laterally and gets through the water jacket (white arrow in photograph).

This experimental set-up is ideal for analysing the photosynthetic and hydrogen

metabolism of C. reinhardtii, since oxygen and carbon dioxide producing and consuming

activities can be monitored in parallel. By photosynthetic activity, the algae produce 16O2,

and respiration generates 12CO2. If the heavy isotopes 18O2 and 13CO2 are added to the

cells, their consumption reflects respiratory oxygen uptake and photosynthetic carbon

dioxide assimilation, respectively. Using a mixture of isotopes, photosynthetic and

respiratory activity of C. reinhardtii can be recorded in one sample and upon illumination,

whereas in a Clark type oxygen electrode respiratory activity has to be followed in the

dark. Thus, the results obtained by this mass spectrometric set-up are more consistent to

the conditions in continuously illuminated sulphur-deprived algal cultures. . 13CO2 was applied as NaH13CO2 (0,3 mM final concentration; 99 % 13C isotope content,

Euriso-Top, Les Ulys, France). 18O2 (95 % 18O2 isotope content, Euriso-Top) was applied

in low amounts that were just enough to detect a significant signal, but which did not have

an obvious effect on hydrogenase activity. Both isotopes were injected through the

capillary of the plug. For the calculation of the gas exchange rates in the presence of 13CO2

and 18O2, the parallel uptake of the unlabelled gas species has to be considered. The

general equations are as follows (S is the unlabeled substrate, which can be either produced

or taken up, and S* is the added labelled substrate, which is considered to be only taken up

in this time-scale):

Uptake: (dS*/dt) x ((S+S*)/S*)

Production: (dS/dt) + uptake x (S/(S+S*))

plug

mobile bulb

O-ring seal

water jacket

magnetic stirrer

Teflon membrane

support by steel fritexit to the mass spectrometer

inlet for gases and reagents


35

For the determination of the real carbon dioxide exchange rates, the equilibrium between

dissolved carbon dioxide and bicarbonate (HCO3-) has to be noticed. The pKa is around

6,35 at 25°C, which means that at pH 6,35, 50 % of dissolved inorganic carbon is in the

form of CO2 and 50 % exists as HCO3-. Consequently, to infer carbon uptake rates from

decrease in carbon dioxide concentration, the apparent rates have to be multiplied by 2. At

increasing pH, the CO2/HCO3- ratio decreases and the factor to apply increases. An

additional complication is that not only the pH, but also the ionic strength of the medium

will affect the CO2/HCO3- equilibrium. In a solution with an ionic strength of 0,1 M, the

concentrations of CO2 are 33 % (pH 6,5), 13,5 % (pH 7), 9 % (pH 7,2), 4,7 % (pH 7,5) and

1,5 % (pH 8).

The fast time response of the measuring system allowed furthermore the determination of

the direct effect of the PSII inhibitor DCMU on hydrogen production. If PSII activity

provides electrons for hydrogen production, hydrogen evolution rates will decrease in the

presence of DCMU (fig 2-6).

02

46

8

0

2

4

6

8

10

12

14

nm

ole

s H

2 x

h-1

x µ

g C

hl-1

time [min]

DCMU

light off

Fig 2-6: Schematic illustration of the hydrogen evolution rates which were recorded by the mass

spectrometric set-up described above. The diagram reflects the actual hydrogen evolution rate of a sulphur-deprived C. reinhardtii culture. After the transfer of the cells to the measuring chamber, the hydrogen graph was recorded until it was stable. Then, 2,5 µM of DCMU were added, which caused a drop of the hydrogen graph. After the line was stable on this new level, light was shut off to measure the background of dark hydrogen evolution.

Results

36

3 Results

C. reinhardtii is a photosynthetic eukaryote having some remarkable characteristics. It

possesses a very oxygen sensitive [Fe]-hydrogenase coupled to the photosynthetic electron

transport chain (Stuart and Gaffron, 1972; Happe and Naber, 1993; Happe and Kaminski,

2002) and several studies indicate that this freshwater alga has a bacterial-type

fermentation system (Gfeller and Gibbs, 1984; Kreuzberg, 1984; Happe et al., 2002;

Hemschemeier, 2002). In former times, the anaerobic metabolism of C. reinhardtii was

induced by removing oxygen artificially. Five years ago, it was published that sulphur

deprivation causes the algae to dramatically re-organise the whole metabolism and to

establish anaerobic conditions without further manipulation (Melis et al., 2000). Under

these conditions, relatively large amounts of hydrogen (Melis et al., 2000) and smaller, but

significant amounts of ethanol and formate are produced (Hemschemeier, 2002; Winkler et

al., 2002b). This is all the more remarkable since hydrogen production and classic

fermentation occur in an organism with oxygenic photosynthesis upon illumination. This

special kind of metabolism was termed “photofermentation” (Winkler et al., 2002b).

3.1 Analysing the reasons leading to hydrogen production

3.1.1 Mass spectrometry as a tool for analysing gas exchange in C. reinhardtii

The first figures summarise the characteristic development of a sulphur-depleted

C. reinhardtii culture. In contrast to all other studies published to date, the results

presented here were obtained by mass spectrometry. This technique allows the detection of

the three gas species which are relevant for the characterisation of sulphur-deprived green

algae: hydrogen, oxygen and carbon dioxide. Furthermore, the use of different oxygen and

carbon dioxide isotopes allows the simultaneous determination of gas producing and

consuming rates upon continuous illumination.

The analysis of the gas phase above a sulphur-deprived C. reinhardtii culture reveals a

characteristic progression of the concentrations of oxygen, carbon dioxide and hydrogen

(Melis et al., 2000). Fig 3-1 presents the concentrations of oxygen, carbon dioxide and

hydrogen in the gas phase above a sealed, sulphur starved C. reinhardtii suspension

(chlorophyll content was 14 µg Chl x ml-1). In the first 45 h, the oxygen concentration in

the gas phase transiently increases above the air value (21 %) (note that all percent

Results

37

specifications shown in this study represent mol-%). This shows that in sulphur-free

medium, the cells first pass an aerobic phase. In parallel to the rising oxygen concentration,

the carbon dioxide content of the gas phase decreases from 0,035 % to almost zero. In this

phase, no hydrogen can be detected. After the aerobic phase, which lasts for 45 h in this

experiment, oxygen is removed rapidly. This is accompanied by a fast increase in carbon

dioxide concentration. Soon after the oxygen concentration has started to decrease,

hydrogen can be detected in the gas phase. The hydrogen production phase sustains for

several days. After 210 h, hydrogen and carbon dioxide concentrations have reached 75 %

and 0,22 %, respectively (note that the difference to 100 % is due to residual nitrogen) (fig

3-1).

Fig 3-1: Hydrogen (),

oxygen () and carbon

dioxide () concentration in the gas phase above a sulphur-deprived C. reinhardtii culture (14 µg Chl x ml-1). At the depicted time points, gas samples were taken with a gas-tight syringe from the culture vessel by piercing through the septum. Analysis of the gas samples was conducted with a mass spectrometer equipped with an injection valve.

It should be noted, that the physiology of sulphur starved C. reinhardtii cultures is

dependent on many influences. The most important parameters are probably the light

intensity, the cell density of the sulphur depleted culture and also the age of the pre-culture.

As an illustration, fig 3-2 shows the development of the in vitro hydrogenase activity in

four C. reinhardtii cultures of different chlorophyll contents (10 to 28 µg Chl x ml-1).

In vitro hydrogenase activity assays measure the presence of active hydrogenase enzyme in a cell extract. The cells are disrupted by detergence, and the hydrogenase enzymes are supplied with excess electrons by an artificial electron donor (reduced methyl viologen). Therefore, the measured hydrogenase activity is independent from cellular in vivo hydrogen evolving capacity. It is, however, a sensitive method to detect active hydrogenases. This, in turn, is an indicator for the developmental stage of a sulphur-starved algal culture. In the C. reinhardtii wild type, in vivo hydrogen production usually parallels in vitro hydrogenase activity (data not shown).

0

20

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60

80

100

0 50 100 150 200 250

time [h]

H2,

O2 [

%]

0

0.05

0.1

0.15

0.2

0.25

CO

2 [

%]

Results

38

Fig 3-2 shows that a dense C. reinhardtii culture (28 µg Chl x ml-1) has synthesised active

hydrogenase enzyme already 6 h after the transfer to sulphur free medium. A cell

suspension of 24 µg Chl x ml-1 exhibits hydrogenase activity after 15 h, a culture of 18 µg

Chl x ml-1 has just started to produce enzyme after 24 h, and a quite thin culture (10 µg Chl

x ml-1) has still not expressed active hydrogenase after 55 h of sulphur starvation. When

hydrogenase activity is once induced, it increases steadily in all four cultures.

Fig 3-2: In vitro hydrogenase

activity in sulphur-deprived C. reinhardtii cultures of different chlorophyll contents ( = 28, = 24, = 18 and = 10 µg Chl x ml-1). 500 µl of cells were taken from the culture vessel at the depicted time points and directly injected into the reaction mixture. After rigorous mixing to disrupt the cells, the suspension was flushed with argon to remove residual hydrogen. Then, the sample was incubated at 37°C for 20 min. Hydrogen was quantified with a gas chromatograph thereafter.

To circumvent the arising problem of generalisation, the results presented in this study

show the values of one measurement most often. However, each experiment was

conducted several times, so that the trend of the physiological development was

demonstrated. The illustrated measurements represent these general metabolic progresses.

Fig 3-3 and 3-4 display the development of in vivo evolution and uptake rates of oxygen

and carbon dioxide in sulphur-depleted C. reinhardtii cells. The special mass spectrometric

set up which was used in CEA Cadarache, France, allows the concomitant analysis of all

four activities by applying heavy isotopes of oxygen and carbon dioxide.

PSII activity produces 16O2 and respiration evolves 12CO2. 18O2 and 13CO2 are consumed by

respiration and carbon dioxide fixation, respectively. The rates which are recorded by the mass spectrometer have to be corrected for the gas consumption of the device and for the parallel uptake of 16O2 and 13CO2 (for details, see materials and methods, paragraph 2.7.5).

0

150

300

450

600

0 25 50 75 100time [h]

in v

itro

hyd

rog

en

ase

activity

[mo

les H

2 x

h-1

x µ

g C

hl-1

]

10

18

24

28

Results

39

Fig 3-3 shows the rates of photosynthetic 16O2 production and respiratory 18O2 uptake in

illuminated cells. Photosynthetic oxygen evolution decreases strongly in the time course of

sulphur starvation. In the figured experiment, the rate of photosynthesis as measured by in

vivo 16O2 evolution has decreased by 66 % after 25 h and by 95 % after 48 h. In the

following 48 h, there is no further significant decline of the oxygen production rate.

Respiratory rates as measured by 18O2 uptake also decline in the time course of sulphur

starvation. Only after 5 h of sulphur deprivation, 18O2 consumption is slightly increased.

The 24 h value is significantly lowered again. In the further development of the culture,

respiratory rates decrease significantly less than photosynthetic rates. Because of the

different progress of oxygen producing and consuming rates, oxygen uptake overcomes

oxygen evolution after about 30 h of sulphur starvation in this experiment.

Fig 3-4 shows the development of 13CO2 uptake and 12CO2 production rates in sulphur-

deprived C. reinhardtii cells. Carbon dioxide exchange was determined in the same

samples that were used for the measurement of oxygen uptake and evolution rates. The

recorded rates are corrected for the pH of the medium, which increases from pH 7,2 to pH

8 in the first 24 h of sulphur starvation and slowly decreases thereafter (Kosourov et al.,

2002). Carbon dioxide exchange rates parallel the respective oxygen production and

consuming activities depicted in fig 3-3. Carbon dioxide fixation strongly declines in the

first 48 h of sulphur starvation. After two days, carbon dioxide uptake can not be detected

anymore (fig 3-4). Carbon dioxide production also decreases in the first 24 h, but stays

constant during the last three days of the experiment (fig 3-4).

Fig 3-3: Oxygen evolution () and

uptake () in sulphur-deprived C.

reinhardtii cells (17 µg Chl x ml-1). Cell samples were taken from the incubation bottle at the depicted time points and transferred to a measuring chamber which was connected to the vacuum of a mass spectrometer. The cell suspension was continuously illuminated with 100 µmoles of photons x m-2 x s-1. Oxygen evolution and uptake rates were detected by 16O2 production (photosynthesis) and 18O2 uptake (respiration).

0

20

40

60

80

0 25 50 75 100

time [h]

oxyge

n e

xcha

nge

[nm

ole

s O

2 x

h-1

x µ

g C

hl-1

]

Results

40

Fig 3-4: Carbon dioxide uptake

() and evolution () in sulphur-starved C. reinhardtii cells. Rates were detected by 13CO2 uptake and 12CO2 evolution in the same cell samples described in fig 3-3.

3.1.2 The analysis of C. reinhardtii mutant strains revealed three special phenotypes

The first attempt to analyse the complex metabolic adaptation of C. reinhardtii to sulphur depletion was to examine several mutant strains. Powerful techniques of molecular genetics and transformation of C. reinhardtii have greatly improved the alga as a system for analysing bioenergetic processes (Hippler et al., 1998) and hundreds of specific mutant strains deficient for several metabolic pathways can be obtained at the Chlamydomonas Genetic Centre (www.chlamy.org/).

The mutant strains analysed in this study were deficient for several parts of the

photosynthetic and respiratory electron transport chains (PSII, cytochrome b6f complex,

plastidic ATPase, Rubisco, cytochrome oxidase). It was shown that all of these mutant

strains have an impaired hydrogen production when deprived of sulphur. Strains FuD7

(PSII-deficient) and PM9.5A (deficient for cytochrome b6f complex) do hardly produce

any hydrogen. Strain FuD50 (deficient for plastidic ATPase) produces some hydrogen for

two days but hydrogen evolution stops thereafter (data for ATPase- and cytochrome b6f

complex-deficient strains not shown). The cytochrome oxidase deficient strain M.90 shows

a similar behaviour as the wild type, but its temporal development is significantly delayed

(see paragraph 3.1.5). Strain CC-2803 (Rubisco-deficient) distinguishes itself from all

other examined strains by producing hydrogen also in the presence of sulphur (see

paragraph 3.1.6). The three mutant strains FuD7, CC-2803 and M.90 were of special

interest, since their behaviour addressed major questions regarding the hydrogen

metabolism of sulphur-starved C. reinhardtii: What is the electron source for hydrogen

production (PSII versus chlororespiration)? What is the role of mitochondrial respiration?

0

10

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30

40

50

0 25 50 75 100

time [h]

carb

on d

ioxid

e e

xch

an

ge

[nm

ole

s C

O2 x

h-1

x µ

g C

hl-1

]

Results

41

How do the competing pathways carbon dioxide fixation and hydrogen production

interfere?

3.1.3 A PSII-mutant offers clues to the electron source of hydrogen production

One of the most disputed details concerning hydrogen evolution by sulphur-deprived

C. reinhardtii cultures is the nature of the electron source for H+ reduction. It can either be

water oxidation at PSII or the oxidation of organic matter and subsequent transfer of

electrons into the PQ pool by a yet unidentified NAD(P)H-plastoquinone-oxidoreductase.

Examining the hydrogen evolution of a PSII mutant or treating the wild type with the PSII

inhibitor DCMU are valuable tools to analyse the participation of each possible electron

source.

Little hydrogen evolution occurs if PSII activity is blocked from the beginning

When deprived of sulphur, the C. reinhardtii wild type starts to evolve hydrogen after a

certain lag phase, in which photosynthetic activity is down regulated and the culture

becomes anaerobic due to mitochondrial respiration (Wykoff et al., 1998; Melis et al.,

2000; fig 3-1 and fig 3-3). Hydrogen accumulates in the gas phase above a sulphur

depleted culture and can make up to 80 % of the gas mixture. The PSII mutant strain FuD7

as well as the wild type treated with DCMU immediately after the transfer to sulphur-free

medium do not produce significant amounts of hydrogen (fig 3-5), although they become

anaerobic only several hours after sealing the cultures (data not shown).

Fig 3-5: Hydrogen accumulation of sulphur-deprived C. reinhardtii wild type (), wild type treated with DCMU (), and PSII deficient strain FuD7 (). Cultures were transferred to sulphur-free medium, and DCMU was added to one wild type culture in a final concentration of 2,5 µM (arrow). Samples of the gas phase were taken with a syringe and injected into a gas chromatograph. Cultures were of the same chlorophyll content (20 µg x ml-1).

0

20

40

60

80

0 20 40 60 80 100

time [h]

H2 [

%]

Results

42

A remarkable effect of the severe anaerobiosis of DCMU treated cells is a fast developing

and very high in vitro hydrogenase activity (tab 3-1). Already 5 h after the cells have been

transferred to sulphur depleted medium and sealed, the in vitro hydrogenase activity of

DCMU-treated cells reaches 170 nmoles H2 x h-1 x µg Chl-1. This is about half of the

highest activity detected in control cultures. During the whole time course of sulphur

starvation, DCMU treated cells show a much higher hydrogenase activity than untreated

cells. In view of the low hydrogen accumulation of DCMU-treated cultures this shows that

the presence of active hydrogenase enzyme is not necessarily accompanied by net

hydrogen evolution.

time [h] in vitro hydrogenase activity [nmoles H2 x h-1 x µg Chl-1] wild type -DCMU wild type + DCMU 0 0 0 5 0 170

24 180 375 48 300 700 72 350 880 96 360 920

Tab 3-1: In vitro hydrogenase activity of sulphur-depleted C. reinhardtii wild type with and without DCMU. An algal culture was transferred to sulphur free medium and distributed to two incubation bottles. Chlorophyll content of the culture was 20 µg x ml-1. Samples were taken at the depicted time points and injected into an anaerobic buffered solution containing Triton X-100 and reduced methyl viologen within a gas-tightly sealed reaction vessel. After 20 min of incubation at 37 °C, the gas phase above the solution was analysed by gas chromatography.

If DCMU is added to a sulphur-deprived C. reinhardtii culture at the time point when

photosynthesis is already down-regulated (as measured by oxygen evolution in the light in

an oxygen electrode), a significant hydrogen evolution can be observed. After 96 h of

incubation in sulphur-free medium, the hydrogen yield of a DCMU treated culture reaches

about one third of the amount produced by untreated cells (fig 3-6).

Results

43

Fig 3-6: Hydrogen accumulation of sulphur-depleted C. reinhardtii wild type without () and with DCMU (). A C. reinhardtii culture was transferred to sulphur-free medium at h = 0 and distributed to two incubation bottles. Chlorophyll content of the culture was 20 µg x ml-1. One of the cultures was treated with DCMU as photosynthesis was already down (h = 17; arrow). Samples were taken as described for fig 3-5.

The direct effect of DCMU on hydrogenase activity is only partial

Using mass spectrometric real-time analyses in CEA Cadarache, France, the direct effect of DCMU on hydrogen production rates could be determined. For this purpose, in vivo hydrogenase activity of sulphur-depleted C. reinhardtii was recorded for several minutes until DCMU was added. The drop of hydrogenase activity caused by DCMU is equivalent to the amounts of electrons that were supplied by water-splitting at PSII.

Experiments were done in several cultures of different chlorophyll contents. It turned out

that the effect of DCMU on hydrogen production is always partial and varies strongly.

Inhibition of DCMU depends on the time point of the measurement within one sulphur

deprivation experiment and on the culture density. Fig 3-7 shows one experiment in which

the DCMU effect was intermediate. Here, the involvement of PSII in hydrogen production

lies between 20 % (24 h) and 35 % (48 h). In other experiments, the inhibitory effect of

DCMU varied between 0 % and 60 %. A comparison of the DCMU effect with the

respective chlorophyll content of the cultures indicated a connection between both. The

lowest inhibitory effect of DCMU was observed in thin cultures (~ 17 µg Chl x ml-1),

whereas hydrogenase activity was much more affected by DCMU in dense cultures (~ 27

µg Chl x ml-1) (data not shown). These results show nevertheless that hydrogen production

depends both on PSII activity and on electron supply by non-photochemical PQ reduction.

0

10

20

30

40

50

60

0 20 40 60 80 100 120

time [h]H

2 [%

]

Results

44

Fig 3-7: In-time effect of DCMU on in vivo hydrogenase activity in a sulphur-depleted C. reinhardtii culture of 22 µg Chl x ml-1. Measurements were performed in a special mass spectrometric set-up as described in detail in materials and methods (see 2.7.5). Samples were taken from the incubation bottle with a gas-tight syringe and transferred to the illuminated measuring chamber of the mass spectrometer at the depicted time points. Hydrogenase activity was recorded for several minutes until the rate was stable (). Then, DCMU was added, and the hydrogen evolution rate was again recorded for several minutes (). Finally, light was shut off to measure the background of dark hydrogen production. The effect of DCMU was controlled by recording the oxygen evolution rate, which became zero after the application of DCMU (data not shown).

3.1.4 Is acetate essential for hydrogen production?

Acetate is necessary to establish anaerobic conditions

When C. reinhardtii cultures are starved for sulphur, anaerobiosis and subsequent

hydrogen evolution are only established in the presence of acetate. Commonly, TAP and

TAP-S medium, respectively, contain 20 mM of acetate. Acetate seems to be necessary to

keep algal photosynthesis low and respiration high enough for an efficient removal of

oxygen from the culture medium (Hemschemeier, 2002). In this study, the minimal acetate

concentration for the establishment of anaerobic conditions and the effect of additional

acetate on hydrogen yields were analysed. Sulphur-deprived C. reinhardtii cultures which

are not supplemented with acetate do not establish anaerobic conditions (Hemschemeier,

2002). In the presence of 5 mM acetate, however, C. reinhardtii cells behave the same. The

oxygen concentration in the gas phase above the culture rises above 25 % in the first 24 h

and remains high during the next 48 h of sulphur depletion (fig 3-8). Afterwards, it

decreases gradually and reaches 10 % after about 200 h. No hydrogen evolution is

detectable under these conditions within 200 h of sulphur depletion (fig 3-9). Sulphur-

depleted C. reinhardtii cultures which are supplemented with 10, 15 or 20 mM of acetate

0 24 48 72 96

11.1

12.2

7.8

2.9

9.0

7.9

5.8

2.00

2

4

6

8

10

12

14

in v

ivo

hyd

rog

en e

vo

lution

rate

s

[nm

ole

s H

2 x

h-1

x µ

g C

hl-1

]

time [h]

+DCMU

-DCMU

Results

45

do not exhibit significant differences in their behaviour. The oxygen concentrations of the

gas phases above the cultures rise within the first 24 h of sulphur starvation. But

afterwards, they decrease rapidly and reach concentrations below 5 % in the next two days

(fig 3-8). Hydrogen evolution of all three cultures starts after 48 h and hydrogen

concentrations increase thereafter in almost the same rates (fig 3-9).

Fig 3-8: Oxygen concentration in the gas phases of sulphur-deprived C.

reinhardtii cultures (14 µg Chl x ml-1) supplemented with different amounts of acetate. One algal culture was harvested in the mid logarithmic phase of growth and transferred to sulphur-free medium without any acetate. It was distributed to four bottles afterwards, and to each bottle, the depicted amount of acetate was added (5 mM , 10 mM , 15 mM , 20 mM ). Gas samples were taken with a gas-tight syringe and directly injected into a gas chromatograph.

Fig 3-9: Hydrogen concentration in the gas phases of the same cultures described in fig 3-8 (acetate concentrations were 5 mM , 10 mM , 15 mM , 20 mM ). Hydrogen was detected by gas chromatography at the same time as oxygen.

Acetate is no direct electron source for hydrogen production

The results described above indicate that, above a certain threshold, the concentration of

acetate has no influence on the hydrogen yield. To confirm this, further experiments with

higher acetate concentrations were performed to observe the effect of additional acetate on

hydrogen production. In these experiments, four cultures were compared. One culture did

not contain any acetate but it was flushed with argon to remove oxygen when

0

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25

30

0 50 100 150 200

time [h]

O2 [%

]

5 mM

10 mm

15 mM

20 mM

0

10

20

30

40

50

60

70

0 50 100 150 200

time [h]

H2 [

%]

5 mM

10 mM

15 mM

20 mM

Results

46

photosynthetic activity was already low. Two cultures contained 20 mM acetate and to one

of these cultures, additional 20 mM acetate were added after 96 h of sulphur depletion. A

fourth culture contained 40 mM acetate right from the beginning on. It turned out that

hydrogen evolution of an acetate-free culture is very low. There is no significant

accumulation of hydrogen even after 200 h of sulphur depletion (fig 3-10), although

oxygen concentration in the gas phase remains low after the removal of oxygen by argon-

flushing (data not shown). The other three cultures behave very similar to each other. No

significant differences can be observed, neither in the development of oxygen

concentration (data not shown) nor in hydrogen accumulation (fig 3-10). The addition of

acetate to an already hydrogen evolving culture has no visible effect.

Fig 3-10: Hydrogen concentration in the gas phases of sulphur-depleted C.

reinhardtii cultures that were supplemented with no (), 20 mM (), 20 mM plus additional 20 mM (arrow upward) (), or 40 mM acetate (). One algal culture (17 µg Chl x ml-1) was transferred to sulphur- and acetate-free medium. It was distributed to four bottles and the depicted amount of acetate was added to each. The autotrophic culture was flushed with argon after 24 h (white arrow downward) to remove oxygen. Samples were taken as described for fig 3-8.

3.1.5 Hydrogen metabolism is delayed in a cytochrome oxidase deficient strain

Upon sulphur starvation, photosynthetic oxygen evolution of C. reinhardtii is strongly

down regulated, but respiratory activity decreases only gradually (fig 3-3). Oxygen uptake

in the first one or two days of sulphur depletion plays a significant role for the

establishment of anaerobiosis. As an alga, C. reinhardtii possesses three major oxygen

reducing components, mitochondrial cytochrome- and alternative oxidase (COX and

AOX), and most probably a plastid terminal oxidase (PTOX) (Mus et al., 2005; PubMed

AAM12876). Further oxygen consuming activities, such as Mehler reaction or the

oxygenase function of Rubisco are neglected here. It was shown before that the transcript

level of coxI (encoding subunit I of COX) remains constant during 96 h of sulphur

deprivation in the C. reinhardtii wild type, whereas the amount of aoxI transcript strongly

0

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30

40

50

0 50 100 150 200time [h]

H2 [%

]

Results

47

decreases (Hemschemeier, 2002). Sulphur-deprived cultures of the COX-deficient

C. reinhardtii strain (M.90) start to produce hydrogen significantly later than wild type

cultures, indicating a significant role of mitochondrial respiration for the progress of this

metabolism.

Anaerobiosis is established later in strain M.90

The COX-deficient strain M.90 shows a delay in the typical development of sulphur-

deprived C. reinhardtii cultures. The removal of oxygen from the gas phase is significantly

slower than in case of the wild type (fig 3-11). Accordingly, hydrogen accumulation in the

gas phase of strain M.90 is observed later (fig 3-11).

Fig 3-11: Oxygen and hydrogen

concentration in the gas phases above sulphur-starved C. reinhardtii wild type (H2 , O2 ) and the COX deficient strain M.90 (H2 , O2 ). Chlorophyll content of both cultures was 16 µg x ml-1. At the depicted time points, gas samples were taken with a gas-tight syringe and analysed by gas chromatography.

Down-regulation of photosynthesis is decelerated in strain M.90

Photosynthetic oxygen evolution and respiratory oxygen uptake rates were analysed in the

COX mutant strain M.90 by determining 16O2 producing and 18O2 consuming activities (fig

3-12). In contrary to the expectations, the retarded establishment of anaerobiosis in strain

M.90 is not due to lower respiratory rates, but to higher photosynthetic activity. Fig 3-12 A

presents the photosynthetic 16O2 evolution rates in sulphur-starved C. reinhardtii wild type

and strain M.90. After 24 h of sulphur deprivation, oxygen production has decreased much

less in C. reinhardtii M.90 than in the wild type. Tab 3-2 summarises the decline of

photosynthetic oxygen evolution rates in wild type and strain M.90 during 96 h of sulphur

starvation in percent.

0

20

40

60

80

100

0 50 100 150time [h]

O2,

H2 [

%]

Results

48

Fig 3-12 B shows the development of respiratory activity in both strains. 18O2 uptake rates

of strain M.90 are lower in the beginning and constitute 75 and 89 % of the wild type rates

at 0 and 24 h of sulphur starvation, respectively. But the respiratory rates of the wild type

decrease gradually in the time course of sulphur deprivation, whereas oxygen uptake

activity of strain M.90 slightly increases. After 48 h of sulphur starvation, 18O2 uptake rates

of this strain are already higher than in the wild type (108 %), and in the end of the

experiment, they are almost double as high (170 %) as in the wild type.

Fig 3-12: Photosynthetic 16O2

production (A) and respiratory 18O2

uptake rates (B) in sulphur-deprived C. reinhardtii wild type () and COX-deficient strain M.90 (). Cells were harvested in the mid exponential stage of growth and transferred to sulphur-free medium as described in materials and methods. Cultures were of the same chlorophyll content (16 µg x ml-1). Samples were taken at the depicted time points and transferred to a measuring chamber connected to the vacuum of a mass spectrometer. Cells were continuously illuminated with 100 µmoles of photons x m-2 x s-1. After the cells had stabilised, 18O2 was added and the rates for 18O2 uptake and 16O2 production were recorded.

[%] [h]

Wt M.90

24 68 23 48 78 38

72 97 76

96 98 85

Tab 3-2: Decrease of oxygen evolution rates in sulphur-starved C. reinhardtii wild type and COX-deficient strain M.90 in percent. Chlorophyll content of the cultures was 16 µg Chl x ml-1. The decline in percent is related to the respective control value at h = 0.

0

10

20

30

40

50

60

oxyg

en

pro

ductio

n r

ate

s

[nm

ole

s 1

6O

2 x

h-1

x µ

g C

hl-1

] A

0

10

20

30

40

50

60

0 20 40 60 80 100time [h]

oxygen

upta

ke

rate

s

[nm

ole

s 1

8O

2 x

h-1

x µ

g C

hl-1

]

B

Results

49

To examine which respiratory pathway is responsible for oxygen uptake in C. reinhardtii M.90, the effects of specific inhibitors on respiratory oxygen uptake rates in the dark were analysed in a Clark type electrode. Myxothiazol was used to block the COX pathway, SHAM inhibits AOX, and propyl gallate inhibits both AOX and PTOX. Inhibitor studies do not necessarily reflect the real situation in the cells, since one pathway can be rapidly replaced by another. In fact it turned out in preliminary experiments, that sometimes neither myxothiazol nor SHAM had a significant effect when added alone.

Only in the first 6 h of sulphur depletion experiments myxothiazol has a strong effect on

respiratory activity when added as the only inhibitor. This shows that in the C. reinhardtii

wild type a big part of oxygen consumption (56 %) is due to the COX pathway and can not

be replaced by other pathways in the beginning of sulphur starvation. In fig 3-13 this is

indicated as low AOX participation. In all following samples, neither myxothiazol nor

SHAM had an effect when applied solely, indicating that the affected pathway could be

rapidly replaced by the other pathway(s). This suggests that sulphur-deprived C.

reinhardtii wild type cells exhibit a marked flexibility in respiratory oxygen uptake

pathways. However, to keep the illustration of the results clear, the decrease in oxygen

consumption rates that were observed after the addition of myxothiazol AND SHAM are

designated as potential AOX activity in fig 3-13. The results show that at least the capacity

for AOX respiration has markedly increased after 24 h of sulphur depletion. Respiratory

oxygen uptake which is left after the addition of myxothiazol and SHAM and which can be

inhibited by propyl gallate is due to plastidic respiration via PTOX.

As expected, myxothiazol as inhibitor of the COX pathway has no effect on oxygen

consumption rates in a C. reinhardtii COX-deficient strain. In strain M.90, the biggest part

of respiratory oxygen uptake is due to the mitochondrial alternative pathway via AOX (fig

3-13). In both wild type and strain M.90, the potential participation of AOX decreases

gradually after the 24 h value. At the same time, the capacity for oxygen uptake by PTOX

increases. In the wild type, the potential involvement of PTOX increases three fold from

11 % (0 h) to roughly 30 % (96 h). In strain M.90, PTOX capacity is higher from the very

beginning on (17 %) and increases 2,5 fold to 42 % (fig 3-13).

Results

50

0 24 48 72 96 0 24 48 72 96 wildtype M.90

0

25

50

75

100

0 20 40 60 80 100

time [h]

O2 u

pta

ke

ca

pa

city [

%]

Fig 3-13: Potential participation of AOX and PTOX in respiratory oxygen uptake in percent. Cell samples of sulphur-starved C. reinhardtii wild type and COX-deficient strain M.90 were taken at the depicted time points and transferred to the measuring chamber of a Clark type oxygen electrode. Respiratory oxygen consumption was recorded in the dark until the rate was stable (= 100 %). Then, myxothiazol (2 µM) was added and the rate was again recorded until it was stable. A decline of the oxygen uptake rate after the addition of myxothiazol reflects the potential capacity of the COX pathway (data not shown). A further decrease of oxygen consumption rates after the addition of SHAM (0,4 mM) represents the potential participation of the alternative pathway via AOX (wild type , M.90 ) (see text for further explanation). Propyl gallate was always added at last, since it also inhibits AOX. A further drop of respiratory activity caused by propyl gallate is due to the inhibition of PTOX activity (wild type , M.90 ).

The amount of D1 (PsbA) protein decreases later in strain M.90

The retarded removal of oxygen from sulphur-deprived C. reinhardtii M.90 correlates with

its photosynthetic oxygen evolution rates, which are higher than in the wild type during the

whole experimental period (96 h). The higher in vivo oxygen production of strain M.90

demonstrates that photochemistry is maintained longer than in the wild type. Western Blot

analyses with an antibody raised against the core protein of PSII, D1 (PsbA), indicate that

in strain M.90, PSII degradation is a bit slower than in the wild type. In the COX-deficient

strain, a decrease of the amount of D1 occurs only after 72 h of sulphur depletion. In the

wild type, some degradation of D1 is visible already after 48 h (fig 3-14).

Fig 3-14: Western Blot analysis using D1 antibody against crude protein extracts from sulphur-starved C. reinhardtii wild type and strain M.90 cultures. Samples were taken at the indicated time points. Cells were harvested, resolved in lysis buffer and heated at 95°C for 5 min. The equivalent of 8 µg chlorophyll was loaded onto the gel.

Results

51

The differences of D1 amounts in the sulphur-deprived C. reinhardtii wild type and COX-

deficient strain M.90 as examined by Western Blot analysis are not significant to explain

the very distinct photosynthetic activity of strain M.90. Therefore, the deviating phenotype

of this strain is probably due to altered photochemistry.

Three different types of fluorescence measurements were conducted to analyse the status of the photosynthetic apparatus in sulphur-deprived C. reinhardtii wild type and COX deficient cultures in more detail. 77K measurements, recording fluorescence emission spectra at low temperature, allow the examination of the state of the mobile LHC II antennae by determining fluorescence of PSII (F686) and PSI (F715). Chlorophyll fluorescence measured with a PAM fluorometer allows a characterisation of status and efficiency of PSII photochemistry. Fluorescence induction experiments resolve electron transfer events from the PSII reaction centre to its primary electron acceptor, QA.

C. reinhardtii M.90 is predominantly in state 2 conditions

Strain M.90 is mainly in state 2, in which PSII is less excited than PSI. This is shown by

results obtained by fluorescence emission spectra recorded at 77K. Fig 3-15 shows the

ratios of fluorescence at 686 nm (caused by PSII) over fluorescence at 715 nm (caused by

PSI).

The F686/F715 ratio indicates the state of the mobile LHCII antennae. Generally, cells are in state 1 (LHCII associated with PSII) when the PQ pool is oxidised and in state 2 (LHCII associated with PSI) when it is reduced. In acetate supplemented C. reinhardtii, maximal

state 1 is achieved by illuminating the cells with far red light (λ > 714 nm, “PSI light”), and it is characterised by a F686/F715 ratio of ~ 2 (Wykoff et al., 1998). In continuous white light, this value accounts for ~ 1,8.

Sulphur-deprived C. reinhardtii wild type and COX-deficient strain M.90 show a different

level and a different development of the F686/F715 ratio. In the wild type, the F686/F715 ratio

increases from 1,5 to almost 1,9 in the first 4 h of S starvation, reflecting state transitions to

state 1 and an oxidised PQ pool. In the next 20 h, the F686/F715 ration decreases and lies

around 1,5 for the remaining three days. In strain M.90, the F686/F715 ratio is close to 1 at 0

h, showing marked state 2 conditions and a strong excitation of PSI at the expense of PSII.

In the measurements at 4 and 24 h, the ratio has decreased to 0,8. Afterwards, it rises up to

1,2 (72 h) and declines again to 1 (96 h). Overall, these results show that the wild type is in

an intermediate state in which PSII is still significantly excited, whereas strain M.90 is in

strong state 1 conditions.

Results

52

Fig 3-15: Ratio of fluorescence at

686 nm (PSII) and 715 nm (PSI) in sulphur-starved C. reinhardtii wild type () and COX-deficient strain M.90 () measured at 77 K. Samples were taken at the depicted time points, rapidly diluted to a chlorophyll content of 5 µg x ml-1, transferred to a measuring cuvette and directly cooled in liquid nitrogen. Fluorescence emission was recorded immediately thereafter by exciting the samples with 435 nm and measuring the emission between 650 and 750 nm. The depicted values are means of three independent measurements with cultures of similar chlorophyll content (15 to 18 µg Chl x ml-1).

The PSII centres of strain M.90 remain open

It was observed that PSII centres are degraded in the C. reinhardtii wild type during

sulphur deprivation (Melis et al., 2000). However, PSII oxygen evolution activity

decelerates sooner than the absolute number of PSII centres decreases. This was shown to

be due to a closure of PSII centres (Wykoff et al., 1998; Antal et al., 2003). In this study,

the status of the primary electron acceptor of PSII, QA, was comparatively analysed in

C. reinhardtii wild type and strain M.90 by fluorescence induction measurements.

The kinetics of fluorescence rise as a response to a rapid onset of light reflects the status of PSII and the PQ pool. Changes in the fluorescence signal show changes in the redox state of the primary electron acceptor of PSII, QA, which in turn is affected by the redox status of the PQ pool. A high fluorescence level can be attributed to highly reduced QA and/or a reduced PQ pool. A rising signal shows the process of QA/PQ reduction, a decreasing signal reflects oxidation. It has to be noted that even in light-adapted cells, some seconds of darkness precede the proper experiment. Therefore, PQ reducing or oxidising processes that occur in the darkness have to be considered. A maximal fluorescence can be achieved by the addition of DCMU, which causes a complete reduction of the primary electron acceptors (QA) of all PSII centres.

Fig 3-16 presents the results obtained by fluorescence induction measurements. The

diagrams show the fluorescence induction curves of C. reinhardtii wild type and strain

M.90, which were incubated in the light for 10 min before the experiment, in the absence

0.0

0.5

1.0

1.5

2.0

0 20 40 60 80 100time [h]

ratio

E6

86 :

E71

5

Results

53

or the presence of DCMU. Light was shut off immediately before the experiment (the

opening of the shutter).

Fig 3-16: Fluorescence induction curves upon the rapid onset of light (100 µmoles photons x m-2 x s-1) in sulphur-deprived C. reinhardtii wild type (A, B) and strain M.90 (C, D). Samples were taken from the

cultures (18 µg Chl x ml-1) after the indicated time points (−−−−6, −−−−24, −−−−48, −−−−72 h) and directly transferred to the measuring cuvette of the apparatus. Fluorescence kinetics were recorded after 10 min of light adaptation either in the absence (A, C) or the presence of 2,5 µM DCMU (B, D).

The fluorescence curves of the wild type and strain M.90 differ significantly. In the light-

adapted wild type, a marked difference between the signals obtained in the absence or the

presence of DCMU can only be observed in the 6 h sample, showing that a big part of PSII

centres is still open in these cells. Furthermore, only this wild type sample shows a slight

rise of fluorescence in the absence of DCMU (fig 3-16 A), indicating that some PQ

oxidation occurs in the dark. After this first sample taken early from a sulphur-deprived

culture, the fluorescence signals of wild type cells incubated 24 h or longer in the absence

of sulphur increase significantly and reach the same heights as are observed in the presence

of DCMU (fig 3-16 B). This shows that PSII centres of the wild type are almost

Results

54

completely closed after 24 h of sulphur starvation. Only in the 48 h sample, some

difference between the fluorescence level in the absence or the presence of DCMU is

visible.

In strain M.90, the fluorescence curves rise only gradually in each measurement performed

in cells that were incubated in sulphur-free medium for 6 to 48 h. This indicates a rapid

oxidation of QA/PQ in this strain within the few seconds of darkness that have to be

applied before the experiment. When light is turned on, the PQ pool is only gradually

refilled by PSII activity and/or non-photochemical PQ reduction. Light-adapted cultures of

strain M.90 show a rapid fluorescence rise (a rapid reduction of QA/PQ) not before the 72 h

measurement (fig 3-16 C). Furthermore, throughout the experimental period, the

fluorescence signals of DCMU treated cells are higher than in untreated cells, indicating

that a significant proportion of PSII centres is still open and active (fig 3-16 D).

FV reflects the maximum quantum efficiency of PSII primary photochemistry. It is about 0,8 in healthy plants and if the value is lower, some PSII are damaged (Krause and Weis, 1992). FV’ reflects the efficiency of open PSII reaction centres. A high FV’ value indicates a constant and high-level electron flow from functional PSII reaction centres to the PQ pool, allowing a high degree of water splitting and oxygen evolution, respectively.

Fig 3-17 A represents the development of FV, which reflects the functional status of PSII in

the wild type and in strain M.90. In the wild type, the relative number of functional PSII

centres decreases significantly and steadily in the time course of sulphur deprivation, as

indicated by a strong decline of FV. In strain M.90, in contrast, PSII centres are not

significantly damaged within 60 h of sulphur starvation, since FV stays nearly constant.

Only thereafter, a rapid drop of functional PSII is visible.

FV’ shows a somewhat different development (fig 3-17 B). In the first 24 h of sulphur

starvation, FV’ develops similar in both wild type and strain M.90. FV’ increases in the first

7 h from 0,33 to 0,5 and decreases to 0,4 in the next 17 h. After this time point, FV’

develops differently in C. reinhardtii M.90 and the wild type. In the latter, a rapid and

strong decrease of the activity of open reaction centres is observed between 24 and 29 h,

where FV’ decreases from 0,4 to 0,2. This drop is not due to an equivalent loss of intact

PSII centres, since FV’ decreases significantly faster than FV. Rather, the strong decline of

FV’ indicates a fast developing “tailback” of electrons, which causes an inhibition of

electron transport events from PSII to the PQ pool. The strong decline of FV’ in the wild

type is paralleled by the onset of anaerobiosis and hydrogen production (black arrow

upward). Afterwards, a more gradual decrease of the efficiency of open PSII centres is

Results

55

observed. In strain M.90, FV’ stays high (around 0,4) for 54 h in sulphur-free medium.

Between 54 and 72 h, a significant descent of PSII efficiency occurs also in sulphur-

deprived cultures of strain M.90. In this strain, the rapid decrease of FV’ is also

accompanied by the establishment of anaerobic conditions and subsequent hydrogen

evolution (indicated by the grey arrow downward) (fig 3-17 B). The decrease of FV’ is

followed by a drop of FV, indicating that the loss of electron transport in PSII promotes its

damage.

Fig 3-17: Variable fluorescence in dark-

(A: FV) and light-adapted cultures (B:

FV’) of sulphur-starved C. reinhardtii wild type and mutant strain M.90 (both cultures had a chlorophyll content of 19 µg x ml-1). FV allows determination of maximum quantum efficiency of PSII primary photochemistry, FV’ allows the determination of open reaction centres in the light. Fluorescence parameters were obtained by PAM fluorometry. Cells samples were taken from the incubation flask at the depicted time points and directly transferred to the measuring chamber of the PAM apparatus. F0 and Fm were measured in dark-adapted cells, Fm’ and F0’ in light-adapted cells. FV was calculated as (Fm – F0) / Fm, FV’ as (Fm’ – F0’) / Fm’. The arrows indicate the start of hydrogen accumulation in the wild type (black arrow upward) and in strain M.90 (grey arrow downward).

Carbon assimilation and acetate uptake by M.90 are not conspicuous

The rates of carbon dioxide fixation in the COX-deficient strain M.90 are in the same order

of those of the C. reinhardtii wild type. But whereas 13CO2 uptake of the wild type ceases

after 48 h of sulphur starvation, (fig 3-4), strain M.90 continues to consume carbon dioxide

for one day (data not shown). These results correspond to the accumulation and subsequent

degradation of starch in strain M.90. Like the wild type (Melis et al., 2000; fig 3-18), this

mutant strain accumulates starch when starved for sulphur (fig 3-18). When the sulphur-

deprived wild type already starts to degrade starch, strain M.90 continues to accumulate

this carbon reserve for one day. Remarkably, the onset of starch degradation is

0.0

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0.2

0.3

0.4

0.5

0.6

rela

tive

eff

icie

ncy (

FV)

A: FV

0.0

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0.3

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0.5

0 20 40 60 80 100time [h]

rela

tive e

ffic

ien

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FV')

B: FV'

Results

56

accompanied by the start of hydrogen accumulation in both strains (indicated by arrows in

fig 3-18).

Fig 3-18: Starch accumulation in sulphur-deprived C. reinhardtii wild type () and strain M.90 ().Cells were harvested by centrifugation, and the pellets were frozen away until starch determination, which was prformed as described in materials and methods (paragraph 2.7.3) with a starch assay kit from Sigma-Aldrich. The presented values are means of three experiments with cultures of similar chlorophyll content (15 to 17 µg x ml-1). The arrows indicate the onset of hydrogen production in the wild type () and strain M.90 ().

Strain M.90 consumes acetate in the same rates and amounts as the wild type

The uptake of acetate is energy dependent, and the COX-deficient strain M.90 might have

a lower ATP/ADP ratio because two proton-pumping respiratory complexes are bypassed.

It was analysed therefore if this mutant strain shows an impaired uptake of acetate.

However, no difference could be observed in acetate uptake rates or absolute acetate

uptake between the C. reinhardtii wild type and strain M.90 (data not shown).

3.1.6 A Rubisco-deficient strain produces hydrogen in the presence of sulphur

All data presented to date indicate that the algal hydrogenase is localised in the chloroplast

stroma and coupled to the photosynthetic electron transport chain by ferredoxin (Stuart and

Gaffron, 1972; Happe et al., 1994; Florin et al., 2001). Therewith, the hydrogenase and

ferredoxin-NADPH-reductase (FNR), which uses photosynthetically provided electrons for

the reduction of NADP+, should theoretically compete for electrons. The function of FNR,

in turn, is dependent on the consumption of reductive equivalents in the chloroplast, which

is mainly due to the carbon dioxide assimilating pathway via Rubisco. The analysis of the

hydrogen metabolism of a Rubisco-deficient strain should provide some information about

the interaction of hydrogen production and reductive biosynthetic pathways.

0 24 48 72 96

0

2

4

6

8

10

12

sta

rch

con

ten

t [µ

g x

µg

Ch

l-1]

time [h]

wt

M.90

Results

57

Rubisco-deficient strain CC-2803 (spr-u-1-6-2 sr-u-2-60 mt+, disrupted rbcL) is listed in the Chlamydomonas Genetics Center (www.chlamy.org/). Chloroplast translation of this strain is resistant to streptomycine due to a mutation of the 16S rRNA. Because this strain can not synthesise the large subunit of Rubisco, it is unable to assimilate carbon dioxide by the Calvin cycle.

The most remarkable characteristic of the Rubisco-deficient strain CC-2803 is its ability to

produce hydrogen in the presence of sulphur (fig 3-19). Both in the presence or the absence

of sulphur, hydrogen accumulation can be observed very soon after the culture has been

sealed gas-tightly. The mutant strain produces less hydrogen than the wild type and more

hydrogen in the presence than in the absence of sulphur. After 96 h of sulphur starvation,

hydrogen concentrations in the gas phases above cultures of strain CC-2803 reach 40 %

and 60 % in TAP-S and TAP+S, respectively, and nearly 80 % in the gas phase above the

wild type. In both cultures of strain CC-2803, the hydrogen level stagnates after 48 h of

incubation, whereas it steadily increases in the case of the wild type (fig 3-19).

Fig 3-19: Hydrogen concentrations

in the gas phases above the sulphur-starved C. reinhardtii wild type () and the Rubisco-deficient strain CC-

2803 in sulphur-free medium (∆), and strain CC-2803 in fresh sulphur-containing medium (). Cultures were of the same chlorophyll content (20 µg Chl x ml-1). Strain CC-2803 was pre-cultured in one flask and distributed to the different media thereafter. The gas phase was analysed by mass spectrometry.

According to the hydrogen accumulation in the gas phase, in vitro and in vivo hydrogenase

activity of strain CC-2803 can be detected very soon after the closure of the incubation

vessel (data for the former not shown). Fig 3-20 shows the development of in vivo

hydrogen producing rates in strain CC-2803 and the C. reinhardtii wild type. In vivo

hydrogenase activities of CC-2803 are quite similar in both media in the first 10 h of

incubation (fig 3-20). They reach 1,8 nmoles H2 x h-1 x µg Chl-1 just after centrifugation of

the culture (0 h) and rise to almost 4 nmoles H2 x h-1 x µg Chl-1 in the next 10 h.

Hydrogenase activity of C. reinhardtii CC-2803 in sulphur containing medium rises further

to 5,2 nmoles H2 x h-1 x µg Chl-1 in the next 14 h, whereas the rate of the sulphur starved

0

20

40

60

80

100

0 20 40 60 80 100

time [h]

H2 [

%]

Results

58

culture already decreases. After this time point, in vivo hydrogen production rates of both

cultures of strain CC-2803 decrease steadily. The observed hydrogen evolving activities of

strain CC-2803 in sulphur replete medium correspond to the amount of hydrogenase

(HydA1) protein as detected by Western blot analyses (fig 3-20, right upper corner). It is

highest after 24 h of incubation and decreases thereafter. In vivo hydrogenase activity of

the wild type starts after 24 h and increases to a value of 10 nmoles H2 x h-1 x µg Chl-1

after 48 h. Thereafter, hydrogen evolution rates of the wild type also decrease, but they

stay significantly higher than the rates observed in CC-2803 cultures (fig 3-20).

Fig 3-20: In vivo hydrogen production rates of the cultures described in fig 3-19 (C. reinhardtii wild type

() and CC-2803 (∆) in sulphur-free medium, CC-2803 in fresh sulphur-containing medium ()). Samples were taken at the depicted time points and transferred to the measuring chamber of a mass spectrometer. Cells were incubated in the sealed chamber upon continuous stirring in the light (100 µmoles of photons x m-2 x s-1) until the hydrogen evolution rate was stable. In the right upper corner of the diagram, a Western blot analysis of the HydA1 protein of strain CC-2803 in sulphur containing medium is shown (for sample treatment see paragraph 2.6).

In strain CC-2803, oxygen evolution rates are very low

Hydrogenase activity requires anaerobic conditions. The Rubisco-deficient strain CC-2803

passes into anoxia within a couple of hours after the culture vessel has been sealed (fig 3-

21). Photosynthetic oxygen evolution activities of strain CC-2803 are very low when

compared to the wild type. They amount to 13 nmoles 16O2 x h-1 x µg Chl-1 after 6 h of

incubation. This is only 25 % of the corresponding wild type rate (fig 3-22, fig 3-3).

Oxygen uptake rates of strain CC-2803 are higher than oxygen evolution rates throughout

the whole experiment, either in the presence or the absence of sulphur. Both oxygen

0

2

4

6

8

10

12

0 20 40 60 80 100

time [h]

nm

ole

s H

2 x

h-1

x µ

g C

hl-1

CC-2803 +S

CC-2803 -S

wt -S

0 24 48 72 96 h

Results

59

producing and consuming activities decrease steadily in the course of the experiment, and

they are lower in sulphur-free medium (fig 3-22). The ratios of oxygen uptake and

consuming activities are summarised in tab 3-3. In strain CC-2803, the ratio is always

higher than 1, showing that oxygen is removed more efficiently than it is produced. In the

sulphur-depleted wild type, oxygen producing rates are higher than consuming rates in the

first 24 h. Oxygen uptake exceeds oxygen production only thereafter (tab 3-3).

Fig 3-21: Oxygen content of the gas phase above sulphur-deprived C. reinhardtii wild type (), Rubisco-deficient strain CC-2803

(∆) and the latter in sulphur replete medium (). Samples were taken after the indicated time points with a gas-tight syringe and analysed by mass spectrometry.

Fig 3-22: Oxygen producing

and consuming activities in the Rubisco-deficient strain CC-2803 in the presence or absence of sulphur. Net oxygen uptake rates were determined by the

uptake of 18O2 (-S ∆, +S), net oxygen evolution rates by the production of 16O2 (-S , +S ) in the same CC-2803 samples described in fig 3-20.

[h] ratio of O2 uptake over O2 production CC-2803 +S CC-2803 -S Wt -S

0 2,53 2,2 0,67 5 2,27 2,23 0,85

24 2.27 3,25 0,83 48 2,86 10,00 4,9 72 5,00 7,50 4,32 96 5,78 20,00 6,08

Tab 3-3: Ratio of 18O2 uptake over 16O2 evolution rates in the Rubisco-deficient strain CC-2803 in sulphur containing (+S) or sulphur-free (-S) medium, and the wild type upon sulphur depletion (Wt -S).

0

10

20

30

0 20 40 60 80 100

time [h]

O2 [%

]

0

10

20

30

40

0 20 40 60 80 100time [h]

oxyg

en

exch

an

ge

ra

tes

[nm

ole

s 1

6/1

8O

2 x

h-1

x µ

g C

hl-1

]

Results

60

Hydrogen production of strain CC-2803 is largely dependent on PSII

The results presented above already show that hydrogen metabolism of the Rubisco-

deficient C. reinhardtii strain CC-2803 is different from that of the wild type. It is

remarkable that in vivo hydrogen production rates of strain CC-2803 (fig 3-20) parallel

photosynthetic oxygen evolution rates (fig 3-22) both in the time lapse as in the height.

This suggests a direct connection between hydrogen evolution and photosynthetic oxygen

evolution in strain CC-2803. In the C. reinhardtii wild type, PSII activity and hydrogenase

activity develop reversely.

If the Rubisco-deficient strain CC-2803 is treated with DCMU at h = 0 of sealed

incubation, no hydrogen can be detected in the gas phase. If DCMU is added to an already

hydrogen accumulating culture, no further increase of the hydrogen concentration can be

observed (data not shown). The latter result indicates that the DCMU effect on hydrogen

production of strain CC-2803 is almost complete. Indeed, the examination of the real-time

effect of DCMU on in vivo hydrogen producing rates revealed that in strain CC-2803,

hydrogen evolution is almost completely inhibited when PSII activity is blocked (fig 3-23).

Fig 3-23: Inhibitory effect of

DCMU on in vivo hydrogen production in the Rubisco-deficient strain CC-2803 (20 µg Chl x ml-1). Measurements were conducted in a measuring chamber connected to a mass spectrometer (for details, see paragraph 2.7.5 and fig 3-7). (Note that the x-axis is not linear for the first five values.)

Sulphur-deprived cells of strain CC-2803 do not accumulate starch

C. reinhardtii wild type is known to accumulate starch upon nutrient deprivation (Ball et

al., 1990). It was shown, that the starch content of sulphur-starved C. reinhardtii cultures

increases significantly in the first 24 to 40 h, but decreases thereafter (Melis et al., 2000;

fig 3-18). From a basal level of about 2 µg starch x µg Chl-1, the amount of starch increases

fourfold to 8 µg x µg Chl-1 in the wild type. Afterwards it declines gradually to 4 µg x µg

0 2 4 10 24 48 72 96

0

1

2

3

4

5

6

in v

ivo

hyd

rog

en

pro

du

ctio

n

[nm

ole

s H

2 x

h-1

x µ

g C

hl-1

]

time [h]

-S DCMU

+S DCMU

-S

+S

Results

61

0 24 48 72 96

0

2

4

6

8

10

sta

rch

co

nte

nt

[µg

x µ

g C

hl-1

]

time [h]

Chl-1. In the Rubisco-deficient strain CC-2803, no starch accumulation can be detected in

the presence or in the absence of sulphur (fig 3-24). The mutant strain contains a certain

level of starch (1,5 µg starch x µg Chl-1), which increases slightly to 2,3 (-S) or 2,2 (+S) µg

starch x µg Chl-1 in the first 24 h of incubation. Afterwards, it decreases again to a level

around 1,2 µg starch x µg Chl-1 (fig 3-24).

Fig 3-24: Starch content of sulphur-deprived C. reinhardtii wild type () and Rubisco-deficient strain CC-2803 in the presence () or the absence of sulphur (). Samples were taken at the depicted time points and treated as described for fig 3-18.

CC-2803 becomes anaerobic and produces hydrogen without acetate

The effect of acetate on the establishment of anaerobiosis in C. reinhardtii CC-2803 was

tested. To confirm that possible different effects of acetate on the C. reinhardtii wild type

and strain CC-2803 are not due to a different uptake rate of acetate, the latter was tested by

measuring the acetate concentration in the medium of both sulphur-depleted strains. As

was published before (Melis et al., 2000), the C. reinhardtii wild type consumes the most

part of acetate in the first 24 h of sulphur starvation. Afterwards, the acetate content of the

medium (data not shown) and the acetate uptake rates remain more or less constant (fig 3-

25). The Rubisco-deficient strain CC-2803 shows a lower acetate uptake rate in the

beginning of the experiment (37 mg acetate consumed per 24 h and per µg Chl).

Afterwards, acetate uptake rates of strain CC-2803 decrease strongly, as already observed

in the wild type. However, whereas a low but significant acetate uptake can be observed in

the wild type throughout the experiment (150 h), the Rubisco-deficient strain shows no

more acetate consumption after 72 h.

Results

62

Fig 3-25: Acetate uptake in sulphur-starved C. reinhardtii wild type () and strain CC-2803 (). Samples were taken at the depicted time points, cells were harvested and the supernatant was frozen away. Acetate was determined in the collected samples with a test kit from Boehringer / r-biopharm. Daily uptake rates were calculated by dividing the difference of acetate concentrations by the chlorophyll content.

When strain CC-2803 is incubated in sulphur containing medium without acetate, the

removal of oxygen from the gas phase is delayed for one day (data not shown). But in

contrast to the wild type, this strain becomes anaerobic under autotrophic conditions and

starts to produce hydrogen, even if the hydrogen yield is lower than in the presence of

acetate (fig 3-26). To test whether this difference in hydrogen concentration was due to a

lost of reductive power during the anaerobisation phase, the experiment was repeated, but

the cells were flushed with argon to remove oxygen at the beginning of the experiment.

However, an autotrophic CC-2803 culture from which oxygen was removed artificially

also exhibits a lower hydrogen accumulation, which amounts to approximately 50 % of the

hydrogen yield of a mixotrophically grown culture (fig 3-26).

Fig 3-26: Hydrogen concentration in the

gas phase above strain CC-2803 in sulphur-containing medium with () or

without acetate (∆, ). One pre-culture was harvested and resolved in fresh medium without acetate (10 µg Chl x ml-1). Cells were distributed to three flasks. To one culture, 20 mM acetate were added. Oxygen was removed from the acetate supplied () and one autotrophic culture

(∆) by argon flushing in the beginning of the experiment (grey arrow). From the second autotrophic culture oxygen was not removed artificially (). The gas phase was analysed by gas chromatography.

0

5

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15

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25

30

35

0 20 40 60 80 100

time [h]

H2 [%

]

24 48 72 96 122 146

0

10

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30

40

50

60

70

aceta

te u

pta

ke

[mg a

ceta

te x

24

h-1

x µ

g C

hl-1

]time [h]

Results

63

Additionally, the effect of additional acetate on hydrogen production in the Rubisco-

deficient strain CC-2803 was tested. There was no obvious difference in oxygen removal

or in hydrogen accumulation between three cultures supplemented with 20 mM, 20 mM

plus additional 20 mM after 96 h, or 40 mM acetate (data not shown).

3.2 C. reinhardtii has an exceptional fermentative metabolism

In the 1980s it was shown that one of the fermentative products of C. reinhardtii is formate

(Gfeller and Gibbs, 1984; Kreuzberg, 1984). Later it was observed that sulphur-starved

C. reinhardtii cultures accumulate ethanol and formate (Hemschemeier, 2002; Winkler et

al., 2002b). The fermentative accumulation of formate is ascribed to pyruvate formate

lyase (PFL). A fragment of a cDNA encoding a putative PFL was found in the databank of

NCBI (PubMed X66410). Northern Blot analyses with a probe raised against this fragment

showed a parallel transcriptional induction of hydA1 and pfl during sulphur depletion

(Hemschemeier, 2002).

PFL and the corresponding fermentation metabolism are typical for (facultative) anaerobic bacteria such as Enterobacteriaceae or Clostridiae (Böck and Sawers, 1996). PFL systems are usually not found in eukaryotes. The only two eukaryotic lineages in which a PFL metabolism has been described are the chytridiomycetes Neocallimastix frontalis and Piromyces sp. E2 (Marvin-Sikkema et al., 1993; Akhmanova et al., 1999) and some chlorophyte algae as Chlorogonium elongatum and C. reinhardtii (Kreuzberg, 1984).

This exceptional characteristic of green algae has not been characterised in detail to date.

In the following, the results of a detailed physiological, biochemical and genetic analysis

of this special fermentative pathway of C. reinhardtii are presented.

3.2.1 C. reinhardtii has several ethanol producing pathways

Hypophosphite is a specific mechanism based inhibitor of PFL. If hypophosphite (10 mM)

is added to sulphur-deprived C. reinhardtii wild type cultures, no formate accumulation

occurs (fig 3-27 A; hypophosphite was added after 20 h of incubation, when a small

amount of formate was already produced). Ethanol production, however, is not inhibited

by hypophosphite (fig 3-27 B). This suggests an additional ethanol producing pathway in

C. reinhardtii.

Results

64

0

10

20

30

40

50

60

0 24 48 72 96

time [h]

eth

ano

l [n

mole

s x

µg C

hl-1

]B

Fig 3-27: Formate (A) and

ethanol accumulation (B) in sulphur-deprived C. reinhardtii cells with () or without () hypophosphite (10 mM). Cell samples were taken after the depicted hours of sulphur starvation. Cells were spun down and the supernatant was frozen away until the analysis was performed. Quantification of formate and ethanol was done in the collected samples with test kits from Boehringer / r-biopharm. The columns represent the means of five measurements with cultures of similar chlorophyll contents (17 – 20 µg Chl x ml-1). The bars indicate the standard deviation.

The typical enzyme responsible for ethanol production in plants and yeast is pyruvate

decarboxylase (PDC). This enzyme decarboxylises pyruvate to acetaldehyde, which is

subsequently reduced to ethanol by alcohol dehydrogenase. C. reinhardtii possesses a PDC

(PubMed E15259). In many cyanobacteria and other bacterial species, pyruvate can be

degraded by pyruvate ferredoxin-oxidoreductase (PFO) (e.g., Pieulle et al., 1997; Stal and

Moezelaar, 1997). PFO decarboxylises pyruvate to acetylCoA, and reduces ferredoxin at

the same time. AcetylCoA can either be reduced to ethanol, or converted into acetate. A

sequence with homology to known PFOs is found in the genomic sequence of

C. reinhardtii (see paragraph 3.2.6 and appendix 8.3 tab 8-1). Ethanol production in

hypophosphite treated C. reinhardtii could therefore be due to PDC or PFO, provided that

acetylCoA which is formed by the latter is converted to ethanol. If PDC or PFO are

responsible for ethanol production in hypophosphite treated C. reinhardtii, these cultures

should exhibit a stronger carbon dioxide production. In fact, when the gas phases of

sulphur-deprived C. reinhardtii cultures were analysed by mass spectrometry, it turned out

that cultures in which PFL is inhibited produce more than doubled amounts of carbon

dioxide (fig 3-28).

0

10

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30

40

50

60

form

ate

[n

mo

les x

µg

Ch

l-1]

A

Results

65

0 24 48 72 96

0

10

20

30

40

50

form

ate

[n

mole

s x

µg C

hl-1

]

time [h]

A

0 24 48 72 96

0

10

20

30

40

50

eth

an

ol [n

mo

les x

µg

Ch

l-1]

time [h]

B

Fig 3-28: Carbon dioxide

accumulation in sulphur-deprived C.

reinhardtii in the presence () or absence () of 10 mM hypophosphite. The presented areas are means of two experiments with cultures of similar chlorophyll content (18 and 20 µg Chl x ml-1). At the indicated time points, gas samples were taken with a gas-tight syringe and injected into a mass spectrometer. (Note that the x-axis is not linear for the first two time points.)

3.2.2 Fermentation is impaired in mutant strains FuD7 and CC-2803

Fermentation in C. reinhardtii strains deficient for Rubisco (CC-2803) or PSII (FuD7) is

significantly impaired. Both produce roughly 25 % of the amounts of formate and ethanol

that are produced by the wild type (fig 3-29). Since both strains pass into anoxia very soon

after the closure of the incubation vessel, this reduced fermentation is probably not due to

the oxygen level of the cultures. Rather, a connection between the accumulation of starch

and the extent of fermentation seems likely. Fermentation in C. reinhardtii appears to be

directly connected with starch breakdown (Gfeller and Gibbs, 1984). Neither strain

CC-2803 (fig 3-24) nor FuD7 (data not shown) produce significant amounts of starch.

Fig 3-29: Accumulation of formate (A) and ethanol (B) in sulphur-deprived C. reinhardtii wild type () and mutant strains FuD7 (PSII deficient) (), CC-2803 (Rubisco deficient) () and M.90 (COX deficient) (). Cultures were of the same chlorophyll content (19 µg x ml-1). Samples were taken at the depicted time points and formate and ethanol were quantified in the supernatants using test kits from Boehringer / r-biopharm.

0 5 24 48 72 96

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

CO

2 [%

]time [h]

Results

66

COX-deficient strain M.90 produces ethanol and formate in equivalent amounts as the wild

type (fig 3-29), but the accumulation of fermentative products appears later. This is in

accordance with the generally retarded anaerobic metabolism of sulphur-deprived

C. reinhardtii M.90.

3.2.3 The C. reinhardtii pfl and pflA cDNAs were isolated and characterised

All the physiological data presented above indicate that the photosynthetic eukaryotic alga

C. reinhardtii has a bacterial-type PFL fermentation system. Aim of the following part of

this study was to characterise the algal PFL system genetically and biochemically and to

demonstrate the functionality of the protein. For this purpose, C. reinhardtii cDNAs

encoding PFL and PFL activase (PflA) were isolated and characterised. The in vivo and in

vitro activities of both enzymes were tested by complementation of E. coli mutant strains

and by purification of the protein and subsequent in vitro assays (the latter was done by a

diploma student; Jacobs, 2005). Finally, the whole enzyme system involved in the PFL

fermentation pathway was analysed by investigations of the C. reinhardtii genome and

expression studies of some of the participating enzymes.

C. reinhardtii PFL and PflA are homologous to the E. coli enzymes

A fragment of a PFL encoding cDNA of C. reinhardtii was already filed in the internet

databank of NCBI (accession number X66410). With this fragment, a blast search was

conducted against the first version of the recently published sequence of the C. reinhardtii

genome on JGI (v1.0) to obtain the whole sequence of the cDNA. A single annotated

sequence and the deduced cDNA sequence were found on scaffold 8 (256002-259292)

(note that meanwhile, v2.0 of the C. reinhardtii genome is published, in which pfl is found

on scaffold 133 (238755-245249)). A protein-protein blast using the deduced amino acid

sequence was conducted on NCBI. It showed that the C. reinhardtii PFL sequence

comprises typical PFL domains, including the conserved region of the glycyl radical. The

algal PFL exhibits high similarity to several known bacterial PFL proteins, except an

approximately 90 bps long N-terminal sequence. An alignment of several PFL-proteins is

presented in appendix 8.2, fig 8-3. The conserved regions around the two catalytic

cysteine residues and the glycyl radical are identical in the PFL enzymes of C. reinhardtii

and E. coli (fig 3-30).

Results

67

The pfl sequence found on JGI was used to deduce oligonucleotides for the amplification

of the whole open reading frame. Suitable restriction sites for cloning the pfl cDNA into

the expression vectors pET9a and pASK-IBA7 were added (see below). RT-PCR using

mRNA from a sulphur-deprived C. reinhardtii culture as template resulted in the expected

fragment of 2,5 kb. Sequencing revealed the identity of the sequence.

In case of PflA, only a small fragment of 559 bps designated as a putative PflA encoding

sequence was filed in the databank of PubMed (BE025007). A blast search on JGI (v2.0)

showed that a part of this sequence is present in the published genome sequence, but it is

not annotated. Therefore, 3’ and 5’ RACE PCR experiments were performed to obtain the

whole pflA transcript. Oligonucleotides were deduced from the known fragment (for

position of the primers, see appendix 8.1, fig 8-1). 3’-RACE was conducted with the

QIAGEN one step RT-PCR kit, using an oligo(dT)18 primer and oligonucleotide PFL-Act-

3. A nested PCR with primers oligo(dT)18 and PFL-Act-1 resulted in a defined fragment of

1680 bps.

For carrying out 5’-RACE, the SMARTTM RACE cDNA Amplification Kit (Clonetech)

was used. Primer PFL-Act-2 was applied for first strand synthesis. Three nested PCR

reactions using oligonucleotides PFL-Act-4, -5 and -6 were necessary to obtain a defined

fragment of 540 bps. The resulting fragments of both RACE-PCR reactions were

sequenced and showed the expected overlapping regions with the original 559 bps

fragment used for the creation of oligonucleotides (see a schematic view in appendix 8.1,

fig 8-2). Assembling of the fragment revealed a 2020 bps fragment with a 1011 bps open

reading frame embracing bases 105 to 1115. The sequence was submitted to the PubMed

databank of NCBI (AY831434).

The deduced amino acid sequence of this open reading frame shows similarity to other

known PFL activating enzymes. Fig 3-30 shows a schematic comparison between

C. reinhardtii and E. coli PflA proteins. The presence of the conserved Cys-x-x-x-Cys-x-x-

Cys motif, coordinating the FeS-cluster of PflA (Sofia et al., 2001), and a conserved

glycine rich region both classify the protein to be a member of the superfamily of radical

SAM proteins (Sofia et al., 2001). See an alignment of several PflA-proteins in appendix

8.2, fig 8-4.

Results

68

Fig 3-30: Schematic comparison between E. coli and C. reinhardtii PFL and PflA polypeptides. Both algal proteins, shown as bars, harbour the highly conserved sequence motifs that are characteristic for each enzyme. In case of PFL, the two catalytic cysteines and the glycyl radical are found in highly conserved regions. PflA contains a characteristic Cys-x-x-x-Cys-x-x-Cys motif and a glycine rich region (indicated by SGGEA in this scheme). The length of the sequences is given at the end of the bars. The positions of the conserved regions which are pictured in this graphic are indicated by black lines and the respective amino acid number. The possible signal peptides of C. reinhardtii proteins are assigned by “SP?”.

The C. reinhardtii pfl and pflA cDNAs were cloned into pASK-IBA7 to allow tightly

controlled expression in E. coli and subsequent purification of the enzymes by an N-

terminal strep-tag. The respective plasmids were constructed in this work, whereas

purification of PFL and in vitro activity assays were conducted by a diploma student

(Jacobs, 2005).

The strategy for cloning the C. reinhardtii pfl cDNA into pASK-IBA7 was worked out

with the programme IBA Primer D’Signer 1.1 (download from web page www.iba-

go.com/download.html). The recommended Eco31I (BsaI) restriction sites could not be

used since there is a single Eco31I site in the pfl coding sequence. The only suitable

restriction site was EcoRI, which allowed an undirected cloning only. The resulting

plasmid pAH21 contained the algal pfl cDNA in the right orientation in pASK-IBA7. PflA

could be cloned into pASK-IBA7 utilising Eco31I sites. This resulted in plasmid pAH25.

The activity of algal PFL and PflA was also analysed in vivo by complementation of

E. coli strains that are deficient for either PFL or PflA. For this purpose, the respective

Results

69

sequences were cloned into the bacterial expression vector pET9a. For cloning into pET9a,

in frame orientation is reached by integrating the ATG start codon of the insert into an

NdeI restriction site (CATATG). The restriction site for ligation of the 3’ end can be either

BamHI or Bpu1102I. Both sites are also found once within the C. reinhardtii pfl cDNA.

Therefore, a partial restriction was conducted with BamHI. As a control, E. coli pfl was

cloned into pET9a, too. In this case, the sequence contains an NdeI restriction site.

Therefore, this sequence was partially restricted, too. All plasmids that resulted from the

ligation of several sequences with pET9a for subsequent complementation assays are listed

in tab 3-4.

plasmid contained sequence resulting E. coli strain

pAH28 complete C. reinhardtii pfl BL21.28

pAH32 truncated C. reinhardtii pfl (lacking the first 210

bps which encode a putative signal peptide)

BL21.32

pAH36 E. coli pfl BL21.36

pAH27 complete C. reinhardtii pflA 234M11.27

pAH33 truncated C. reinhardtii pflA (lacking the first 267

bps which encode a putative signal peptide)

234M11.33

pAH46 E. coli pflA 234M11.46

Tab 3-4: Summary of the PFL and PflA encoding sequences that were cloned into pET9a to be used in subsequent complementation studies, and the E. coli strains that resulted from transformation with the respective plasmids..

3.2.4 C. reinhardtii PFL is functionally synthesised in E. coli

To test the in vivo activity of C. reinhardtii PFL and PflA, E. coli strains deficient either

for PFL (BL21∆pfl) or PflA (234M11) were transformed with the respective C. reinhardtii

sequences which were present in pET9a (tab 3-4). As a control for the test system, the

E. coli mutant strains were also transformed with E. coli pfl and pflA sequences.

Preceding tests were conducted to characterise the fermentative metabolism of E. coli

“wild type” and mutant strains. E. coli cultures were treated as described in materials and

methods to induce fermentation.

Results

70

Anaerobic E. coli cultures degrade pyruvate via PFL and secrete formate until a certain threshold is reached. Then, formate is re-imported into the cells, where it triggers the expression of the FHL complex (Mnatsakanyan et al., 2002; Sawers, 2005). This complex disproportionates formate into carbon dioxide and hydrogen. Therefore, the activity of PFL can be indirectly followed by hydrogen accumulation.

The analysis of fermenting E. coli DH5α and BL21(DE3)pLys revealed that the former

strain behaves as described in literature, but the latter strain does not re-import formate and

shows no hydrogen production (data not shown). It is not clear why BL21 behaves

different. The consequence for this study was to measure formate excretion as an indicator

for PFL activity, and not hydrogen production.

A further characteristic of PFL deficient E. coli strains is their poor growth in the absence

of oxygen (Varenne et al., 1975). Therefore, anaerobic growth of the complemented

BL21∆pfl and 234M11 should be restored if the introduced C. reinhardtii PFL and PflA

proteins, respectively, are fully active.

Before testing the activity of the algal PFL enzyme, a control strain of BL21∆pfl

complemented with the E. coli pfl sequence was analysed to secure the functionality of the

pET-system for this object. Since overproduction of the proteins was not desired in this

case, no or very low concentrations (0,1 mM) of the inductor IPTG were added.

Additionally, the effect of glucose was examined, since glucose should have a negative

effect on the lacUV5 promoter due to catabolite repression. On the other hand, glucose

could have a positive effect on the growth rate of anaerobic E. coli cultures.

These first experiments revealed that the expression system was functional. Growth of

BL21∆pfl complemented with the E. coli PFL (termed BL21.36) is restored (fig 3-33), and

the cells produce formate (fig 3-31, 3-32). It turned out that neither 0,1 mM IPTG nor

0,2 % glucose have a significant effect on formate accumulation of BL21.36. In the

presence of glucose, the culture reaches a somewhat higher optical density (data not

shown).

According to the results of the preceding experiments, complementation experiments with

the algal proteins were always conducted in LB medium without IPTG, but with 0,2 %

glucose to promote growth of the cells. It was shown in parallel experiments that the

heterologous expression of the algal strep-tagged PFL enzyme in E. coli does not yield

soluble protein when the cultures are incubated at 37°C (Jacobs, 2005). Therefore, pre-

cultures and anaerobic cultures were kept at room temperature (23°C).

Results

71

0.00

0.05

0.10

0.15

0.20

0.25

0,1 mMIPTG

0,2% Glc

BL21 wt BL21-pfl BL21.36 BL21.36 BL21.36

form

ate

[g x

l-1

]

BL21∆ pfl

0.0

0.1

0.2

0.3


form

ate

[g x

l-1

]

BL21∆pfl

Fig 3-31: Formate concentration in the medium of anaerobically growing E. coli strains. Aerobically growing pre-cultures were transferred to 50 ml LB medium and flushed with argon for 10 min to remove oxygen. BL21(DE3)pLys (BL21 wt) and

BL21∆pfl served as positive respective negative controls.

BL21∆pfl which was complemented with E. coli PFL (BL21.36) was either incubated without supplements, or treated with 0,1 mM IPTG or 0,2 % glucose. Anaerobic cultures were incubated at 37°C, and samples for formate detection were taken after 5 () and 22 h ().

The complementation studies revealed that C. reinhardtii PFL is functionally expressed in

E. coli. Cultures of BL21∆pfl strains equipped with the algal PFL proteins excrete formate

(fig 3-32). No significant difference is observable if BL21∆pfl is transformed with the

complete pfl sequence (BL21.28) or with the truncated sequence lacking the first 210 bps

which might encode a transit peptide (BL21.32) (fig 3-32). BL21.28 and BL21.32 produce

less formate than the “wild type” or BL21.36.

However, the growth of BL21.28 and BL21.32 is not restored by the algal PFL proteins

(fig 3-33) Whereas the “wild type” BL21(DE3)pLys as well as BL21.36 grow up to an

OD600 of around 0,6, BL21.28 and BL21.32 show the same impaired growth as the PFL

deficient strain BL21∆pfl (fig 3-33) and only reach an OD600 of 0,3.

Fig 3-32: Formate accumulation of several

E. coli strains (BL21∆pfl, BL21.36 =

BL21∆pfl transformed with E. coli pfl,

BL21.28, BL21.32 = BL21∆pfl transformed with complete and truncated C. reinhardtii pfl). BL21wt = BL21(DE3)pLys was used as control. Samples were taken after 8 () and 24 h () of anaerobic incubation with a syringe. Cells were spun down and the supernatant was frozen away until the quantification of formate occurred with a test kit from Boehringer / r-biopharm.

Results

72

kDa

– 130

– 100

– 72

– 55

BL21.36 BL21 wt BL21∆pfl BL21.32 BL21.28

2 4 20 2 4 20 h 2 4 20 2 4 20 h

KkDa

– 130

– 100

– 72

– 55

Fig 3-33: Anaerobic growth as measured by the optical density (OD600) of the same

E. coli strains as described for fig 3-32. Cell samples were taken with a syringe after the indicated hours of anaerobic cultivation and measured photometrically

at λ = 600 nm against LB medium.

To detect E. coli and C. reinhardtii PFL proteins in the examined strains, Western Blot

analyses were performed with anti-E. coli-TdcE antibodies.

TdcE is a 2-ketobutyrate formate-lyase that also exhibits PFL activity (Heßlinger et al., 1998; Sawers et al., 1998). Anti-E. coli-TdcE-antibodies cross react with E. coli PFL (G. Sawers, personal communication).

85 kDa PFL proteins were detected in BL21 “wild type” and in significantly higher

amounts in BL21.36, but not in BL21∆pfl extracts (fig 3-34). In BL21.28 (complemented

with complete C. reinhardtii pfl cDNA), a ~91 kDa protein could be detected. In BL21.32

(truncated C. reinhardtii pfl), a ~85 kDa protein reacts with anti-TdcE antibody (fig 3-34).

In all cases, the specific two-band pattern of PFL from aerobically prepared protein

extracts is visible. The lower band represents the bigger PFL fragment that arises upon

scission of activated PFL polypeptide at the position of the glycyl radical in the presence of

oxygen (Heßlinger et al., 1998).

Fig 3-34: Western Blot analysis of PFL expression in anaerobically growing E. coli strains: Crude protein extracts from various E. coli strains were separated in SDS-gels, and after transfer to nitrocellulose membrane, the polypeptides were challenged with anti-E. coli-TcdE antibodies. E. coli strains were BL21

“wild type” (BL21 wt) and BL21∆pfl as well as BL21∆pfl complemented with E. coli PFL (BL21.36) and C. reinhardtii PFL with (BL21.28) and without putative signal peptide (BL21.32). C. reinhardtii PFL purified by strep-tag chromatography served as a control (K; Jacobs, 2005). Samples for protein extracts were taken after 2, 4 and 20 h of anaerobic growth. The expected sizes for the PFL proteins are 85 kDa (E. coli), 84 kDa (C. reinhardtii, without putative signal peptide) and 91 kDa (C. reinhardtii, complete PFL).

0

0.2

0.4

0.6

0.8


OD

60

0

0h

5h

8h

24h

BL21∆pfl

Results

73

3.2.5 The algal PflA fails to activate E. coli PFL

E. coli strain 234M11 is deficient for PFL activase (PflA). It was transformed with

plasmids pAH27 (complete C. reinhardtii pflA), pAH33 (truncated C. reinhardtii pflA that

lacks the first 267 bps encoding a putative signal peptide) and pAH46 (E. coli pflA). E. coli

strain 234M11 is a derivative of MC4100 that does not possess a chromosomal gene for

viral T7-polymerase. Therefore it seemed necessary to provide an extra source for T7-

polymerase via a second plasmid. However, co-transformation of 234M11 with the pET9a

derivates constructed in this study and a plasmid harbouring a gene which codes for T7-

polymerase failed. It turned out nevertheless that T7 polymerase is dispensable for the

transcription of genes under the control of the T7-promoter. In preceding experiments,

234M11.46 exhibited a strong hydrogen production, showing that PFL is activated in this

strain. This, in turn, is only possible if the introduced pflA gene under the control of T7

promoter is expressed. Furthermore, the behaviour of 234M11.46 demonstrates that strain

MC4100, the origin of 234M11, behaves as it is described in literature (see above).

Parallel experiments showed that strep-tagged C. reinhardtii PflA is synthesised as soluble

protein only at 4°C (Jacobs, 2005). Therefore, to test C. reinhardtii PflA activity in E. coli,

pre-cultures of the 234M11 derivates transformed with pAH27, pAH33 and pAH46 were

grown at room temperature. After the transfer to anaerobic medium, the cells were pre-

incubated at 4°C for 3 h. Strain 234M11.46, the PflA deficient strain 234M11 transformed

with E. coli pflA, is clearly rescued from the phenotype of 234M11 that does not grow

under anaerobic conditions and shows no formate or hydrogen production (fig 3-35, tab 3-

5). Strain 234M11.27 (complete C. reinhardtii pflA) and 234M11.33 (truncated C.

reinhardtii pflA) show the same growth rates as strain 234M11, and they do not produce

formate or hydrogen in the absence of oxygen. This indicates that C. reinhardtii PflA is not

functionally expressed or not able to activate E. coli PFL.

Tab 3-5: Hydrogen and formate that are present in the gas phase and the medium, respectively, after 20 h of anaerobic incubation in the same E. coli strains that are described in the legend of fig 3-35.

H2 (µmoles x ml-1) formate (g x l-1)

DH5αααα 2,69 0,103

234M11 0 0

234M11.27 0 0

234M11.33 0 0

234M11.46 6,56 0,026

Results

74

Fig 3-35: Anaerobic growth of

several E. coli strains. Aerobic growing pre-cultures were transferred to 50 ml LB medium, sealed with a Suba seal and flushed with argon for 10 min. To allow the synthesis of soluble C. reinhardtii PflA, cells were kept at 4°C for 3 h. Afterwards they were incubated at

room temperature (20°C). DH5α and 234M11 served as positive and negative controls, respectively. 234M11.27, .33 and .46 = 234M11 transformed with pET9a vectors harbouring the complete (27) or truncated (33) C. reinhardtii pflA or E. coli pflA (46).

3.2.6 Expression studies on selected genes

The PFL pathway involves, besides PFL and PflA, three further enzymes: AdhE, a

multifunctional enzyme comprising of acetaldehyde and alcohol dehydrogenase function,

phosphotransacetylase (PTA) and acetate kinase (ACK). As mentioned above (paragraph

3.2.1), C. reinhardtii possesses further pathways for ethanol production. Pyruvate could be

either decarboxylised by pyruvate decarboxylase (PDC) or pyruvate ferredoxin

oxidoreductase (PFO). It was described before that anaerobic C. reinhardtii cultures also

produce traces of glycerol (Gfeller and Gibbs, 1984; Kreuzberg, 1984). Little amounts of

glycerol were also detected in sulphur-deprived C. reinhardtii cultures (data not shown).

The enzymes responsible for glycerol fermentation are glycerol-3-phosphate

dehydrogenase (GPD) and glycerol-3-phosphatase (GPP) (Påhlman et al., 2001). Glycerol-

3-phosphate, which is produced by GPD, can also be used for the biosynthesis of

glycerolipids.

To examine if C. reinhardtii possesses all the listed enzymes, blast searches were

conducted with known protein sequences against the algal genome. Indeed, C. reinhardtii

has all genes which encode for fermentative enzymes involved in the PFL system.

Additionally, gene sequences that encode proteins of the PDC and the PFO pathway as

well as enzymes of glycerol fermentation were found (tab 3-6). Some of the sequences are

found several times in the genome. For example, five alcohol dehydrogenase coding

sequences are present in the C. reinhardtii genome. It is remarkable that all of the PFL

0.0

0.2

0.4

0.6

0.8

1.0

1.2

DH5 234M11 234M11.27 234M11.33 234M11.46

OD

60

0

2 h

4 h

20h

DH5α

Results

75

pathway enzymes, AdhE, PTA and ACK, are found twice in the genome, whereas PFL and

PflA seem to exist as single copy genes only. It should be noted here that the genomic

sequence of C. reinhardtii is not completed yet. There are still gaps. For example, the

complete PflA coding sequence could not be found in the genome. After the isolation of

the cDNA during this study, it turned out that the main part of the pflA gene covers a gap

between two scaffolds. Because of still missing sequences, the following table may not be

complete.

enzyme number reaction

PFL 1 non-oxidative cleavage of pyruvate into formate and acetylCoA

PflA 1 PFL activating enzyme

AdhE 2 trifunctional: alcohol and acetaldehyde dehydrogenase, PFL

deactivase

PTA 2 conversion of acetylCoA into acetyl phosphate

ACK 2 phosphorylation of ADP with the phosphogroup of

acetylphosphate

PDC 1 decarboxylation of pyruvate to carbon dioxide and acetaldehyde

ADH 5 reduction of acetaldehyde to ethanol

PFO 1 decarboxylation of pyruvate to carbon dioxide and acetylCoA,

reduction of ferredoxin

GPD 6 interconversion of dihydroxyacetone phosphate and glycerol-3-

phosphate

Tab 3-6: List of the fermentative enzymes involved in the PFL (pyruvate formate-lyase), PDC (pyruvate decarboxylase) or PFO (pyruvate ferredoxin oxidoreductase) pathway and their presence (as indicated by the corresponding gene sequence) in C. reinhardtii. The name of the respective enzyme is written in the first column. The second and third columns present the number of sequences found in the genome and the catalysed reaction. (AdhE = tri-functional enzyme complex harbouring acetaldehyde and alcohol dehydrogenase function as well as PFL deactivase; PTA = phosphotransacetylase; ACK = acetate kinase; ADH = zinc containing alcohol dehydrogenase; GPD = glycerol-3-phosphate dehydrogenase). See a detailed table in appendix 8.3, tab 8-1.

RT-PCR studies were performed to examine the expression of several of the genes

encoding fermentative enzymes. Using mRNA that had been isolated daily from sulphur-

deprived C. reinhardtii wild type cultures, RT PCRs were conducted with oligonucleotides

specific for hydA1 (hydrogenase HydA1), pfl (PFL), pflA (PFL activase), pta1

Results

76

(phosphotransacetylase 1), adh1 (AdhE 1) and pdc (pyruvate decarboxylase). For all these

genes, the respective transcript could be detected (fig 3-36).

Fig 3-36: RT-PCR on mRNA isolated from

sulphur-deprived C. reinhardtii. Total RNA was isolated from algal cultures at the depicted time points of sulphur starvation. mRNA was isolated from total RNA thereafter. Specific oligonucleotides raised against hydA1, pfl, pflA, pta1, adh1 (AdhE) and pdc were used to check each mRNA sample for the presence of the corresponding transcript.

Furthermore, Western Blot analyses confirmed the expression of PFL and AdhE in

sulphur-deprived C. reinhardtii. PFL was detected with an antibody raised against E. coli

TdcE. Applied to algal crude protein extracts, one significant band of the expected size

(~ 91 kDa) and some further signals appeared. The multifunctional AdhE protein of

C. reinhardtii was detected with anti-E. coli-AdhE antibodies, which showed excellent

cross reaction with an algal protein of the expected size (~ 100 kDa).

Upon sulphur deprivation, PFL protein accumulates in the C. reinhardtii wild type in a

similar pattern as HydA1. AdhE seems to be constitutively expressed in C. reinhardtii, as

indicated by the significant signal in the control sample (0 h). Upon shifting to

anaerobiosis, the algae enhance AdhE synthesis (fig 3-37).

Both PFL and AdhE are also significantly expressed in the Rubisco-deficient strain CC-

2803, though this strain does not produce significant amounts of ethanol and formate (see

paragraph 3.2.2) (fig 3-37).

Fig 3-37: Western Blot analyses to examine the expression of fermentative proteins in sulphur-deprived C. reinhardtii wild type and Rubisco deficient strain CC-2803. Crude protein extracts from the algal cultures were separated in SDS-gels, and after transfer to nitrocellulose membrane, the polypeptides were treated with anti-C. reinhardtii-HydA1, anti-E. coli-AdhE or anti-E. coli-TcdE antibodies. Crude protein extracts were produced at the indicated time points of sulphur depletion.

HydA

PFL

AdhE

HydA

PFL

AdhE

0 24 48 72 96 h

C. reinhardtii wild type

C. reinhardtii CC-2803

hydA1

pfl

pflA

pta1

adh1

pdc

0 24 48 72 96 h

Results

77

3.3 Overview of results

The presented study aimed to characterise the photofermentative metabolism of sulphur-

deprived C. reinhardtii cultures. The following results were obtained:

• Hydrogen evolution of sulphur-deprived C. reinhardtii wild type cultures is indirectly

dependent on PSII activity. Active PSII is essential during the first hours of sulphur

starvation to build starch. Its direct contribution to hydrogen evolution is only partial.

• The C. reinhardtii wild type is dependent on a certain amount of acetate as a

respiratory substrate to establish anaerobic conditions. Even if oxygen is removed

artificially, only a little amount of hydrogen is produced in the absence of acetate. Once

a hydrogen metabolism is established, acetate has no further effect on hydrogen yields.

• As shown by the deviating development of a C. reinhardtii mutant deficient for

cytochrome oxidase (M.90), the length of the aerobic phase is correlated with

respiratory events that, in turn, influence the status of the photosynthetic electron

transport chain. Strain M.90 cells are mainly in state 2 and PSII centres remain open.

• A Rubisco-deficient strain (CC-2803) produces hydrogen in the absence and in the

presence of sulphur. In both media, oxygen is removed as soon as the incubation flask

is sealed. This is due to low photosynthetic rates that are always exceeded by

respiratory activity. Cultures of CC-2803 start to produce hydrogen only a few hours

after closure of the culture vessel. Hydrogen evolution of this strain, which hardly

accumulates starch, is mainly dependent on PSII activity.

• C. reinhardtii disposes of a bacterial-type fermentation system catalysed by PFL.

Hypophosphite treated cultures do not produce formate. However, ethanol production

is not affected, and inhibited cultures produce larger amounts of carbon dioxide. This

indicates the presence of additional pathways of ethanol fermentation.

• The cDNAs encoding C. reinhardtii PFL and its activating enzyme (PflA) were

isolated. The deduced polypeptide sequences are homologue to other PFL and PflA

enzymes. Investigations on the C. reinhardtii genomic sequence show that this green

alga disposes of all genes encoding enzymes involved in the PFL pathway.

Furthermore, sequences that encode enzymes of the PDC, PFO and glycerol

fermentation pathways were characterised.

• C. reinhardtii PFL is functionally expressed in E. coli, while C. reinhardtii PflA was

not functional in activating E. coli PFL.

Discussion

78

4 Discussion

The unicellular chlorophyte alga C. reinhardtii has a remarkable complex fermentation

system that is pronounced anaerobically and which is established in the absence of sulphur.

This work aims to provide deeper insights into several aspects of this physiology, which

was termed “photofermentation” (Winkler et al., 2002b) since it develops in illuminated

algae.

4.1 What are the factors that finally lead to hydrogen production?

When deprived of sulphur, cultures of the unicellular green alga C. reinhardtii radically

change their physiology. Within one or two days, the metabolism of the alga changes from

aerobic photosynthetic growth to an anaerobic resting state.

The composition of the gas phase above a sulphur-starved C. reinhardtii culture is a rough

indicator of the metabolic processes in the cells. The real-time analysis of gas exchange

rates in vivo gives detailed information about the physiological adaptations of the algae. It

reveals that photosynthetic oxygen evolution rates of C. reinhardtii cells decline drastically

within one or two days as a reaction to sulphur depletion (fig 3-3; Wykoff et al., 1998).

Respiratory oxygen uptake activity remains significantly higher which results in a net

consumption of oxygen (fig 3-1, 3-3; Melis et al., 2000).

The special mass spectrometric set-up that could be used in this study allowed the

recording of gas uptake and production rates simultaneously in one illuminated sample.

The technique applied in this work confirmed the existing data regarding oxygen uptake

rates, showing that respiratory rates in the light are highly similar to oxygen uptake rates

measured in the dark. Carbon dioxide exchange rates for sulphur-deprived C. reinhardtii

have not been published before. The results obtained by mass spectrometric experiments

reveal that carbon dioxide fixation activity is high at the beginning of sulphur deprivation,

but strongly decreases in the first one or two days until it reaches zero (fig 3-4). This is in

accordance with phenomena described previously: The high initial carbon dioxide uptake

rates account for the accumulation of starch in the first hours of sulphur depletion (Melis et

al., 2000; Zhang et al., 2002; fig 3-18). The decrease in in vivo uptake rates thereafter is

paralleled by the degradation of Rubisco protein (Zhang et al., 2002) and net production of

carbon dioxide (Melis et al., 2000; fig 3-1) due to low but significant carbon dioxide

production rates throughout the sulphur deprivation experiment (fig 3-4). Basically, the

Discussion

79

decrease in carbon dioxide uptake rates parallels the decrease in photosynthetic oxygen

evolution. However, whereas carbon dioxide uptake becomes completely diminished, a

low but significant level of photosynthesis is maintained. This indicates that another

electron sink has replaced the Calvin cycle.

Many aspects of the drastic physiological changes that occur in C. reinhardtii upon sulphur

deprivation are still unclear. The results obtained in this study contribute to the

understanding of some of the major aspects of these complex metabolic adaptations.

The main factor that causes PSII activity to decrease is photoinhibition which results

in reversible “closure” and irreversible damage to PSII centres. In a cytochrome

oxidase-deficient C. reinhardtii strain, PSII inactivation is decelerated because light

energy is mainly focussed on PSI.

In the C. reinhardtii wild type, the decrease of photosynthetic activity as a reaction to

sulphur starvation has been extensively studied (Wykoff et al., 1998; Melis et al., 2000;

Zhang et al., 2002). Summarising existing data and the results presented in this work, it

can be stated that a process commonly known as photoinhibition is responsible for the loss

of PSII oxygen evolving activity.

The term photoinhibition describes the process of light-induced inhibition of

photosynthetic electron transport.

Photoinhibition occurs under conditions in which the capacity of the PQ pool to accept

electrons from PSII is exceeded, regardless of whether this is due to excess light intensities

or to less efficient reoxidation of the PQ pool (Gong and Ohad, 1991). The main target of

photoinhibition is PSII, which is inactivated sequentially, followed by degradation of the

D1 protein (Ohad et al., 1984). Photoinactivation of PSII affects primarily, or at least

initially, the acceptor side of PSII, QA (Kyle et al., 1984; Ohad et al., 1988). One of the

responses to excessive light intensities is the appearance of so-called QB non-reducing

centres (Chylla and Whitmarsh, 1989). These centres oxidise QA approximately 1000 times

slower than QB reducing centres (Ort and Whitmarsh, 1990) and are essentially non-

productive in oxygen evolution (Melis, 1991). In QB non-reducing centres, a quasi-stable

or long-lived state of reduced QA (QA_) is established that can relax under low exciting

conditions (Ohad et al., 1988). These “closed” centres are known to develop upon

illumination with excessive light intensities and are proposed to constitute an initial,

reversible stage preceding irreversible photoinactivation (Ohad et al., 1988; Etienne and

Kirilovsky, 1992).

Discussion

80

In C. reinhardtii, PSII activity, as measured by oxygen evolution, decreases by 75 % after

24 h of sulphur deprivation (Wykoff et al., 1998; fig 3-3). It was shown that two processes

account for this loss of activity. First, approximately 30 % of PSII centres are irreversibly

damaged within one day (Wykoff et al., 1998; Melis et al., 2000). Second, an increased

number of the remaining PSII centres is converted to QB non-reducing centres (Wykoff et

al., 1998).

The damage to PSII centres can be followed by quantifying the amount of QA (the primary

electron acceptor of P680) that can be reduced. After the initial 30 % decrease of functional

PSII centres, this degradation process continues steadily in the time course of sulphur

starvation (Melis et al., 2000). In the present study the damage of PSII centres was

followed by PAM fluorescence measurements which allowed determination of the FV/Fm

ratio. This ratio is around 0,8 in healthy plant cells and a decrease in the value of FV/Fm

indicates that some PSII centres are damaged (Krause and Weis, 1992). The determination

of the FV/Fm ratio is a relative measurement that does not provide information about the

absolute number of PSII centres. However, the graph of the values of FV/Fm (fig 3-17 A)

shows a good correlation with the values obtained by QA quantification published

previously (Melis et al., 2000), suggesting that the FV/Fm ratio reflects the conditions in the

cells with good approximation.

It is not yet fully understood why sulphur deprivation causes such a rapid damage

preferentially of PSII centres. One major reason is probably the high turnover rate of the

PSII core protein D1 (PsbA), which is frequently damaged even upon moderate

illumination (Aro et al., 1993). Under normal growth conditions, de novo protein synthesis

and reassembly of PSII centres compensate for photoinactivation and D1 degradation

(Ohad et al., 1984). It seems likely that under sulphur-limiting conditions, the de novo

synthesis of D1 is impaired due to the lack of essential amino acids. PSI is less affected by

photoinhibition, but it is not immune to the effects of excessive light intensities (Sonoike,

1995, 1996). It was shown that the amount of PSI, as measured by quantification of P700,

decreases by approximately 50 % after 100 h of sulphur starvation (Melis et al., 2000).

It might be suggested that two parameters specific to sulphur limitation might have a

dramatic impact on photosynthesis, too. First, PSII activity and heat stability in

C. reinhardtii was shown to depend on sulfoquinovosyl diacylglycerols (SQDG) (Sato et

Discussion

81

al., 1995; Minoda et al., 2002; Sato et al., 2003). SQDGs are anionic sulpholipids that are

almost exclusively found in the thylakoid membranes. When starved for sulphur,

C. reinhardtii cells degrade SQDGs and replace them with phosphatidylglycerols (Sato et

al., 2000). Therefore, PSII of sulphur-deprived C. reinhardtii might exhibit an enhanced

sensitivity towards light stress due to a loss of SQDGs. Secondly, the glutathione content

of sulphur-starved algae might decrease. Glutathione is quite abundant in plant

chloroplasts. It is an important antioxidant (Alscher, 1989), helps in detoxification of

heavy metals (Grill et al., 1989) and is also a large reservoir for reduced sulphur in the

form of cysteine (Kunert and Foyer, 1993). It was also shown to be a mediator in redox-

controlled gene expression (Irihimovitch and Shapira, 2000). An altered ratio of reduced

over oxidised glutathione or a lower glutathione level could be further parameters affecting

PSII integrity and the overall status of the photosynthetic apparatus in sulphur-deprived

green algae, since reactive oxygen species (ROS) might be less efficiently neutralised.

The development of QB non-reducing centres precedes irreversible photodamage of PSII.

The appearance of these “closed” centres can be followed by measuring the kinetics of the

increase in fluorescence, which provides information about the reduction state of QA

(Melis, 1991). Maximal fluorescence is achieved when QA is fully reduced, i.e. in the

presence of DCMU, which blocks electron transfer between QA and QB.

In the initial stages of sulphur deprivation, the maximal fluorescence obtained in

C. reinhardtii wild type samples is significantly lower in the absence than in the presence

of DCMU (fig 3-16 A and B), indicating that QA is only partially reduced in the absence of

DCMU. Furthermore, a particular increase in the fluorescence curve indicates that some

QA is oxidised during the few seconds of darkness that have to be applied before the proper

experiment is initiated (the opening of the shutter which starts illumination) and is reduced

upon the onset of illumination. In the subsequent measurements that were performed daily

in sulphur-deprived C. reinhardtii cells, the fluorescence level was more or less the same

in the absence or in the presence of DCMU, indicating that even in the absence of DCMU,

QA is fully reduced. It should be noted that in whole cells, the analysis of fluorescence

induction curves can not differentiate between quasi-stable reduced QA and a reduced state

of QA due to reduction of the PQ pool by cellular metabolites. Especially under anaerobic

conditions the PQ pool was shown to be strongly reduced (Bulté et al., 1990). From

previous studies it can be assumed that a part of the reduced QA is due to the existence of

QB non-reducing centres (Wykoff et al., 1998). Parallel experiments with dark-adapted

Discussion

82

cells showed that QA can be partially oxidised in the dark, as indicated by a higher

difference in fluorescence between DCMU-treated and –untreated, dark-adapted cells (data

not shown).

“Closure” and irreversible damage of PSII centres are the major factors causing a

decreased oxygen evolution activity in sulphur-deprived C. reinhardtii cells. However, a

further process that diminishes oxygen evolution rates in the sulphur-deprived algae is state

transition. A transition from mainly state 1 (exciting PSII) towards state 2 (exciting PSI)

has been observed (Wykoff et al., 1998). In the present study, 77K measurements were

conducted to analyse the state of the mobile LHCII antennae in sulphur-deprived C.

reinhardtii cultures during 96 h of sulphur starvation. The measurements of the wild type

confirmed and extended previous studies. After 6 h of sulphur deprivation, the F686

(PSII)/F715 (PSI) ratio of the wild type is nearly 2. It was shown previously that

C. reinhardtii cells that are illuminated with far-red “PSI-light” (λ > 714 nm), which is

known to induce state 1 conditions, exhibit a F686/F715 ratio of 2 (Wykoff et al., 1998).

Obviously, algae that are deprived of sulphur for 6 h are in nearly optimal state 1

conditions. Since the state of the mobile LHCII antennae is always directly related to the

reduction state of the PQ pool (Keren and Ohad, 1998), this result demonstrates that the

PQ pool of the cells is more or less oxidised at this point in time.

This, in turn, correlates with the photosynthetic activity of the algae measured at the onset

of sulphur deprivation. Here, the cells show extensive carbon dioxide fixation activity and

starch accumulation, indicating an efficient removal of electrons for reductive biosynthetic

processes.

After a further 24 h of sulphur starvation, the F686/F715 ratio in C. reinhardtii wild type cells

decreases to values around 1,5 and stays at this level throughout the experiment. This

demonstrates a transition towards state 2 as already shown before (Wykoff et al., 1998),

which in turn reflects an increasing reduction state of the PQ pool. This is probably due to

a less efficient removal of electrons by assimilatory processes. Upon transition to state 2,

some light energy is directed away from PSII, resulting in a further decrease of PSII

activity.

In addition to all these processes leading to reduced PSII activity, a further and rapid

deactivation of PSII is observed upon the onset of anaerobiosis. This is shown by the

development of FV’/Fm’ which reflects the efficiency of open PSII centres in the light. In

Discussion

83

sulphur-starved C. reinhardtii wild type cultures, a strong and rapid drop of FV’/Fm’ is

observed upon the transition to anaerobic conditions (fig 3-17 B). A recent study showed

that this rapid decrease of the relative number of open PSII centres is not the trigger for,

but the consequence of anaerobiosis because the PQ pool becomes strongly reduced in the

absence of oxygen (Bulté et al., 1990; Antal et al., 2003).

The progression of PSII activity and the establishment of anaerobic conditions in the

sulphur-deprived C. reinhardtii wild type can be summarised as follows: The degradation

and inactivation of PSII, which is caused by excess light energy, results in a gradual

decrease of photosynthetic activity in an initial phase of sulphur deprivation. At a certain

point of this process, oxygen evolution is exceeded by respiratory oxygen consumption,

resulting in a net removal of oxygen. As soon as anaerobiosis is reached, a further and

strong inactivation of PSII is evoked by the over-reduction of the PQ pool.

In the COX-deficient C. reinhardtii strain M.90, the phase of active oxygen evolution by

PSII is significantly extended (fig 3-11). In contrast to what is expected in a mutant strain

impaired in mitochondrial respiration, the reduced rate of oxygen removal is not due to

significantly lower respiratory rates. Rather, C. reinhardtii M.90 exhibits prolonged

photosynthetic oxygen evolution rates (fig 3-12 A). This demonstrates that the loss of

active and open PSII centres is less marked in strain M.90 than in the wild type.

Several major differences between photosynthetic activity in the wild type and M.90 can

be observed:

C. reinhardtii strain M.90 is predominantly in state 2 conditions throughout the

experimental period (96 h). The F686/F715 ratio always lies around 1. When compared to the

F686/F715 ratio obtained by cells in state 1, which lies around a value of 2, mutant strain

M.90 is in marked state 2 conditions.

In C. reinhardtii, a transition to state 2 (focussing light energy on PSI) can be observed

when oxidative phosphorylation is inhibited by anaerobiosis, uncouplers or inhibitors of

respiratory electron transport (Gans and Rébeillé, 1990). This phenomenon was shown to

be due to a rapid drop in the ATP content which causes a stimulation of glycolysis and an

increase in the NAD(P)H level, which in turn results in non-photochemical reduction of the

PQ pool (Rébeillé and Gans, 1988; Bulté et al., 1990).

Discussion

84

In COX-deficient strains, two proton-pumping complexes (complex III / cytochrome bc1

complex and complex IV / COX) are bypassed, resulting in a lower ATP yield. A recent

study that analysed photochemistry in several mitochondrial mutant strains confirmed that

in mutants impaired in complex IV (COX), the ATP level is lowered by 25 % when

compared to that of the wild type (Cardol et al., 2003). The same study showed that the C.

reinhardtii mutant dum18, which contains a frame-shift in the cox1 gene and is impaired in

complex IV activity (Colin et al., 1995), is mainly in state 2, and cannot be forced to pass

into state 1 by PSI light, which usually results in an oxidation of the PQ pool and transition

to state 1. The authors suggest that in this respiratory mutant, non-photochemical PQ

reduction must be too high to allow efficient re-oxidation of the PQ pool, even if PSI is

optimally excited.

The FV/Fm ratio decreases significantly more slowly in C. reinhardtii strain M.90 than in

the wild type (fig 3-17 A), indicating that the damage to PSII centres is decelerated. This

observation is further supported by Western Blot analyses with anti-D1 antibodies (fig 3-

14). In the COX-deficient strain, a significant decrease in the amounts of D1 is observed

one day later than in the wild type. Only after the “shut-down” of PSII centres due to

anaerobiosis, the FV/Fm ratio decreases rapidly (see below).

In C. reinhardtii strain M.90, PSII centres stay open. Determination of FV’/Fm’ by PAM

measurements indeed showed that in C. reinhardtii strain M.90, the relative efficiency of

open PSII centres in the light remains high during 60 h of incubation in sulphur-free

medium. Only then is a rapid drop in FV’/Fm’ observed (fig 3-17 B). An equivalent

decrease in the relative amount of open PSII centres is observed in the C. reinhardtii wild

type after 24 h of sulphur deprivation, and it follows the establishment of anaerobic

conditions. Also, in strain M.90, the loss of open PSII centres coincides with the onset of

anaerobiosis, suggesting that the same mechanism (strong PQ reduction) is responsible for

this drop of PSII activity in strain M.90 and the wild type. It was shown for the wild type

that the decline of FV’/Fm’ is fully reversible in the first hours after the removal of oxygen

(Antal et al., 2003). Aeration of the cells restored the efficiency of open PSII centres. But

after further 50 h of anaerobic incubation, FV’/Fm’ could only partially be restored by

aeration, indicating that more PSII centres were irreversibly damaged (Antal et al., 2003).

In the COX-deficient strain M.90, the decline of FV’/Fm’ (effective PSII activity) is

followed by a somewhat slower decrease in FV/Fm (maximal PSII activity), which does not

Discussion

85

significantly change before this time point. This suggests that during anaerobiosis, the

inactivation of PSII is favoured.

The “open” status of PSII centres in sulphur-starved C. reinhardtii strain M.90 is also

demonstrated by fluorescence induction curves, which show that in this strain, QA is not

fully reduced. A significant rise in fluorescence after the onset of light reflects that QA is

efficiently oxidised in the dark and only gradually reduced during illumination. The more

oxidised state of QA in the COX mutant is further evinced by the significant differences

between the heights of the fluorescence signals obtained in DCMU-treated or -untreated

cells, which can be observed in all samples from 6 to 72 h of sulphur starvation (fig 3-16).

This difference indicates that QA is not fully reduced in the absence of DCMU.

More efficient PQ oxidation, which can be suggested for C. reinhardtii strain M.90, can

either be performed by PSI activity or by non-photochemical PQ oxidation via PTOX. In

the COX-deficient strain M.90, the capacity for PTOX-mediated oxygen reduction seems

to be some higher than in the wild type (see discussion of respiration below). Furthermore,

this strain is predominantly in state 2, in which PSI is intensely excited (fig 3-15), allowing

an efficient oxidation of the PQ pool by PSI activity.

This, however, requires that PSI has an appropriate electron sink. In strain M.90, carbon

dioxide fixation activity is maintained for longer than in the wild type, and starch

accumulation continues for one more day, indicating that Rubisco is not degraded as fast as

in the wild type. It is not yet clear why Rubisco is degraded as rapidly in sulphur-depleted

C. reinhardtii wild type (Zhang et al., 2002). It was shown for duckweed plants (Lemna

minor L.) that sulphur starvation is the only environmental condition that causes a strong

degradation of Rubisco protein without leading to plant death (Ferreira and Teixeira,

1992). It was proposed that Rubisco, which is rich in sulphurous amino acids, could serve

as an intracellular reservoir of sulphur. In sulphur-starved Dunaliella salina, degradation of

Rubisco enzymes and an increase in intracellular NH3 levels were observed, suggesting

that carbon was directed away from protein synthesis but channelled into sugar metabolism

(Giordano et al., 2000). The same mechanism seems to be active in C. reinhardtii, since

the alga accumulates large amounts of starch in the first one or two days of sulphur

starvation (Melis et al., 2000; fig 3-18).

Discussion

86

It is likely that the same regulatory processes that are active as a response to sulphur

limitations in the C. reinhardtii wild type are present in the COX-deficient strain M.90.

Therefore, the residual Rubisco activity must be due to another mechanism.

It is less commonly known that the Rubisco protein is also sensitive to oxidative stress

(Mehta et al., 1992; Desimone et al., 1996; Knopf and Shapira, 2005), and synthesis of the

large subunit of Rubisco is at least partially regulated by the concentration of reactive

oxygen species (ROS) and the redox state of the glutathione pool, respectively

(Irihimovitch and Shapira, 2000). It is possible that in sulphur-deprived C. reinhardtii

cells, the production of ROS is at least partially responsible for the degradation of Rubisco

protein. Again, as was discussed for PSII D1 protein, a de novo synthesis might not be

feasible or not favourable because of amino acid limitation. Therefore it seems probable

that in C. reinhardtii strain M.90, which is mainly in state 2 and shows less PSII damage

upon sulphur starvation, the production of ROS is less pronounced. This in turn would

allow a prolonged Rubisco activity.

Summarising these data it can be concluded that inactivation, irreversible damage and

degradation of PSII in sulphur-starved C. reinhardtii cultures are slowed in the COX-

deficient strain M.90. In the wild type, at least part of the reduction in PSII activity is

regulated by the sac1 gene product, which was shown to be critical for the acclimation of

C. reinhardtii to sulphur depletion (Davies et al., 1996). It is likely that the specific

responses to limitations in sulphur are regulated in strain M.90 in a similar way as in the

wild type. Therefore, the maintenance of PSII activity is probably a consequence of the

physiological characteristics of this mitochondrial mutant.

In strain M.90, the PQ pool is reduced mainly due to the low ATP and high NAD(P)H

content of the cells. Because of the highly reduced PQ pool, C. reinhardtii strain M.90

cells are mainly in state 2 during 96 h of sulphur depletion. It was shown that in full state 2

conditions, oxygen evolution ceases (Finazzi et al., 1999). Therefore, the relatively high

oxygen evolution rates of the C. reinhardtii mutant strain M.90 show that even these cells

are not under optimal state 2 conditions. In the COX-deficient strain, light energy is

nevertheless directed away from PSII, diminishing charge separation events and thereby

light-induced damage (see a schematic overview in fig 4-1). These considerations are

confirmed by recent results. It was shown that the irreversible damage of PSII that is

caused by high light intensities is more pronounced in state 1 (Finazzi et al. 2001).

Discussion

87

Fig 4-1: Schematic and simplified comparison of the events that lead to PSII inactivation in sulphur-deprived C. reinhardtii wild type (left) and to a significant lesser extent in the COX-deficient strain M.90 (right).

The initial aerobic phase sets the course for subsequent anaerobic survival. Here,

starch is accumulated by photosynthetic activity. This organic reserve is an important

electron donor for subsequent hydrogen production.

Because hydrogen evolution by C. reinhardtii is regarded as a promising biotechnological

approach for the production of hydrogen as an energy source (Melis and Happe, 2001,

2004), the question regarding what is the best electron source for hydrogen production has

attracted much interest. Detailed knowledge, therefore, of these electron pathways would

probably allow manipulation and thus enhancement of the electron supply.

Several results obtained in this work show that hydrogen evolution is both dependent on

residual PSII activity and non-photochemical PQ reduction. The latter, however, depends

on photosynthetic activity during the initial phase of sulphur deprivation.

A PSII-deficient strain (FuD7) and the C. reinhardtii wild type treated with the PSII

inhibitor DCMU pass into anoxia immediately after the closure of the incubation vessel.

Despite the fact that anaerobiosis, one of the most important preconditions for hydrogen

evolution, is provided, and large amounts of active hydrogenase enzyme are synthesised

(tab 3-1), no significant hydrogen evolution can be detected (fig 3-5). These results could

be interpreted in terms of PSII being the major electron donor for hydrogen production, as

Discussion

88

has been proposed by others (Kosourov et al., 2003; Antal et al., 2003). However, the

addition of DCMU to C. reinhardtii cultures that have already passed to the hydrogen

production phase has a much lower effect on hydrogen accumulation (fig 3-6). This

correlates with the real-time effect of DCMU on cell samples as measured by mass

spectrometry. The addition of DCMU to hydrogen-producing algae results in an inhibition

of hydrogen evolution by 0 to 60 % (see fig 3-7), showing that 40 to 100 % of in vivo

hydrogen production are independent of PSII activity.

These results indicate that PSII activity is important in the initial aerobic phase of sulphur

starvation. It was reasoned that the necessity for PSII is due to its contribution in starch

accumulation. Efficient starch biosynthesis depends on the electrons extracted from water

to be used for carbon dioxide fixation in the Calvin cycle. These considerations led to a

detailed analysis of the correlation of starch accumulation and hydrogen production with

the time point of DCMU addition (Fouchard et al., 2005). According to predictions it was

shown that the addition of DCMU prevented starch accumulation in sulphur-deprived

C. reinhardtii, but if DCMU was added to a culture after 24 or 48 h, starch did accumulate.

The amount of hydrogen produced by sulphur-deprived C. reinhardtii cultures correlates

strongly with the amount of starch that can be synthesised before the addition of DCMU

(Fouchard et al., 2005). The importance of starch for hydrogen production is also shown

by the impaired hydrogen evolution rates of C. reinhardtii mutant strains deficient for

isoamlyase or ADP-glucose pyrophosporylase (Posewitz et al., 2004). If these enzymes,

which are involved in starch hydrolysis and biosynthesis, respectively, are absent, the

initial rate of hydrogen evolution in anaerobically adapted C. reinhardtii was shown to be

significantly lower than in the wild type.

During several experiments to determine the direct real-time effect of DCMU on hydrogen

evolution rates it turned out that the inhibitory effect of DCMU varies strongly. The rough

trend that could be observed indicates that DCMU inhibition of hydrogen production is

strongest in cultures with a high chlorophyll content (not shown). This can be explained by

the reduced accumulation of starch in cells due to a less efficient quantum yield of

photosynthesis in self-shadowing dense cultures. This phenomenon has yet to be analysed

in more detail.

It can be concluded from the phenomena described here that C. reinhardtii cells

accumulate starch in the first hours of sulphur deprivation with reductant provided by PSII

activity. In the hydrogen-producing phase, starch is degraded and the electrons are

Discussion

89

transferred to the photosynthetic chain by non-photochemical PQ reduction. It was shown

formerly that hydrogen evolution in C. reinhardtii is dependent on PSI activity, but

electrons can be provided by sources other than PSII (Stuart and Gaffron, 1972; Gfeller

and Gibbs, 1985; Mus et al., 2005).

The data presented in this study disagree with previous studies which considered residual

PSII activity as the major electron source for hydrogen production (Kosourov et al., 2003).

In fact, these data provide the model that was proposed when the hydrogen metabolism of

sulphur-deprived C. reinhardtii was first published with detailed evidence. The special

metabolism of the alga was called the “two stage hydrogen production process”, in which

initial (aerobic) photosynthetic assimilation of organic reserves (stage 1) is temporarily

separated from (anaerobic) hydrogen production at the expense of previously synthesised

cellular substrates (stage 2) (Melis et al., 2000) (fig 4-2).

Fig 4-2: Simplified diagram of the two stages of sulphur-deprived C. reinhardtii cells. In the aerobic phase, electrons provided by PSII are used to build up starch by Rubisco (Rub). In the anaerobic phase, starch is degraded. Electrons originating from glucose oxidation are transferred to the PQ pool and finally to the hydrogenase (HydA1). In this phase, PSII plays a minor role as electron donor.

In vitro hydrogenase activity in sulphur-depleted C. reinhardtii cultures is highest

under marked anaerobic conditions. However, the presence of large amounts of

active hydrogenase enzyme does not necessarily result in high in vivo hydrogen

evolution rates.

Two interesting features of hydrogenase activity and hydrogen production can be

concluded from the experiments with cells in which PSII activity is abolished (by a

mutation or by DCMU). Sulphur-starved algal cultures with impaired PSII activity exhibit

a much higher in vitro hydrogenase activity than wild type cultures (tab 3-1). The

determination of in vitro hydrogenase activity occurs in a reaction mixture that disrupts the

cells and contains saturating amounts of the reduced artificial electron donor methyl

PSII PQ

HydA

Starch

e- + CO2

Rub

e- + CO2

Ndh

PQ

aerobic (stage 1)

anaerobic (stage 2)

H2O ½ O2

H2O ½ O2

PSIIH2

2 H+

Discussion

90

viologen. Therefore, hydrogenase activity measured by this technique does not reflect

cellular hydrogen evolution, but simply the presence of active hydrogenase in the

respective sample. Coincident measurements of in vivo hydrogen accumulation showed

that DCMU-treated or PSII-deficient C. reinhardtii cells hardly produce any hydrogen.

Therefore, the presence of active hydrogenase does not necessarily result in hydrogen

production. Furthermore, it was shown previously that in untreated wild type C. reinhardtii

cultures, in vitro hydrogenase activity can attain levels that are up to 30 times higher than

the respective in vivo hydrogen evolution rate (Hemschemeier, 2002).

These results show that hydrogen evolution is not limited by the amount of hydrogenase

enzyme. Even in the presence of large amounts of hydrogenase, the real hydrogen output

can be close to zero, as shown by PSII-deficient / inhibited cultures. This demonstrates that

the electron supply is limiting hydrogen production.

It was shown previously that the expression of the hydA1 gene is dependent on the oxygen

concentration (Happe et al., 1994; Happe and Kaminski, 2002; Stirnberg and Happe,

2004). However, it was not analysed in detail if in DCMU-treated or PSII-deficient cells

the expression of hydA1 is stronger. Northern Blot analyses, which allow only a rough

quantification of the respective mRNA, indicate that the expression of hydA1 is not

significantly higher in these cells (data not shown). Therefore, it can be suggested that the

higher in vitro hydrogenase activity in PSII-impaired cells is not due to higher expression,

but to less inhibition of the produced enzymes by oxygen.

Sulphur-deprived C. reinhardtii cultures grown autotrophically or in the presence of

5 mM acetate cannot efficiently remove oxygen from the medium and show no

hydrogen production. Additionally, if acetate is supplied as sole carbon source it is

essential for the accumulation of starch.

It was shown previously that in sulphur-deprived C. reinhardtii cultures, which are not

supplemented with acetate, the removal of oxygen is strongly impaired (Hemschemeier,

2002). In this study, the effect of different acetate concentrations on the transition from

aerobic to anaerobic phase was examined. It turned out that in cultures with acetate

concentrations of 10 mM or higher, sulphur-deprived C. reinhardtii cells behave as if they

were incubated in the standard medium (i.e. 20 mM acetate). However, in the presence of

only 5 mM acetate, a culture develops which appears not to sense the presence of acetate

and remains aerobic (fig 3-8). The positive effect of acetate on oxygen removal can be

explained by the stimulatory effect of acetate on respiration (Fett and Coleman, 1994;

Discussion

91

Endo and Asada, 1996, Hemschemeier, 2002) and the inhibitory effect of acetate on

photosynthesis and the expression of genes involved in photosynthesis (Goldschmidt-

Clermont, 1986; Kindle, 1987; Endo and Asada, 1996, Heifetz et al., 2000). Acetate as the

sole carbon source causes limitations in the efficiency and capacity of photosynthesis,

shown by a lower quantum yield and a lower light-saturated rate of photosynthesis (Polle

et al., 2000).

Clearly, a certain level of acetate is needed, which is undershot in medium containing 5

mM and attained or exceeded in medium containing 10 mM acetate. A rough calculation

might explain this threshold. A 300 ml algal culture (~ 18 µg Chl x ml-1) produces roughly

3,2 mmoles of oxygen in the first 10 h of sulphur starvation. To reduce this amount of

oxygen, 12,8 mmoles of electrons in the form of NAD(P)H or FADH2 are needed. This

amount would be reached if 3,2 mmoles of acetate are converted to acetylCoA and

oxidised in the citric acid cycle. Of course, this calculation is very rough and does not

account for oxygen uptake by chloroplast respiration, acetate that is assimilated or the

oxidation of organic matter that is already present in the algae and that would yield further

reducing equivalents. Nevertheless, this might explain the threshold of 10 mM acetate that

is necessary for the removal of oxygen from sulphur-depleted C. reinhardtii cultures.

Above an acetate concentration of 10 mM, the development of algal cultures supplemented

with different amounts of acetate shows no significant deviations. This indicates that the

algae need a certain amount of acetate, not more and not less, for the establishment of

anaerobiosis and hydrogen metabolism. Cultures supplemented with 20 mM acetate

consume approximately half of the acetate in the first 24 h. Subsequently, the level of

acetate in the medium stays more or less constant. These two observations – the minimal

amount for the establishment of anaerobiosis and the initial consumption of acetate –

clearly demonstrate that approximately a 10 mM acetate concentration is necessary and

sufficient for the induction of hydrogen metabolism.

It is remarkable that sulphur-depleted C. reinhardtii cells do not produce significant

amounts of hydrogen in the absence of acetate, even when oxygen is removed from the

culture by argon-flushing when photosynthetic oxygen evolution rates are already low (fig

3-10). This demonstrates a further role of acetate besides being a respiratory substrate

and/or a factor that diminishes the ratio of photosynthetic oxygen evolution over

Discussion

92

respiratory oxygen uptake. It was shown that sulphur-deprived C. reinhardtii cells do not

accumulate starch when they are incubated without any additional carbon source (extra

carbon dioxide or acetate) (Fouchard et al., 2005). Obviously, acetate is also an essential

carbon source for the accumulation of starch.

Cultures with different acetate contents (20 or 40 mM) produce the same amounts of

hydrogen, further demonstrating that the algae can consume only a certain amount of

acetate. The addition of acetate to cells already producing hydrogen has no visible effect.

Obviously, acetate cannot be converted into a form that can be used for hydrogen

production in this phase. This is in accordance with the constant medium level of acetate

after the first 24 h of sulphur-deprivation, which indicates that in this phase, C. reinhardtii

cells no longer consume acetate.

Summarising these data it can be stated that sulphur-deprived C. reinhardtii cells need

acetate, both for the establishment of anaerobic conditions and the biosynthesis of starch.

Since the accumulation of starch was shown to rely on PSII activity (see above) and hardly

occurs in the absence of Rubisco protein (see below), it can be assumed that acetate is

oxidised during respiratory processes in the first hours of sulphur starvation, and that the

resulting carbon dioxide is used for starch accumulation.

In the time course of sulphur starvation, C. reinhardtii cells can efficiently switch

between several respiratory pathways. The capacity of PTOX-catalysed plastid

respiration is enhanced.

The removal of oxygen from a sulphur-starved C. reinhardtii culture is due to respiratory

activity. It was shown that the extent of this activity depends on acetate (see above). It was

interesting to analyse the contribution of each major respiratory pathway via alternative

oxidase (AOX), COX or the plastid terminal oxidase (PTOX) to respiratory oxygen uptake,

especially in the C. reinhardtii COX-deficient strain M.90.

It should be noted first that inhibitor studies always mean a manipulation of cell

physiology, thereby changing the real metabolic processes. The respective inhibitor might

a) also inhibit enzymes or processes other than those in the target pathway and b) might

cause a rapid cellular response that circumvents the inhibited reaction. To test the latter, in

preliminary experiments the respiratory inhibitors used in this work were applied in

different orders to see if their effect would change. This was indeed the case. When applied

to C. reinhardtii cultures that had been incubated in sulphur-free medium for 24 h or

Discussion

93

longer, neither myxothiazol (blocks the COX pathway) nor SHAM (affects AOX) had a

significant effect when added alone. Only when both inhibitors were present did

respiratory activity decrease markedly. This shows that the algae can rapidly switch to use

the non-inhibited respiratory pathway. Therefore, interpretating the results presented in fig

3-13, which show the potential percentage of AOX activity, one should consider the fact

that in untreated cells, this part of respiration could also be due to the COX pathway or to a

mixture of both pathways. Myxothiazol causes a 56 % decrease of respiration when it is

the only present inhibitor, but only in the first hours of sulphur deprivation. This suggests

that under these conditions, the AOX pathway is less efficient compared with sulphur-

depleted cells, even if AOX activity is proposed to be constitutively high in C. reinhardtii

(Goyal and Tolbert, 1989). However, AOX capacity increases during the acclimation of

C. reinhardtii to sulphur limitations. In C. reinhardtii, two aox genes are present, aox1 and

aox2 (Dinant et al., 2001). Previously it was shown that the transcript of aox1 decreases

dramatically upon sulphur starvation (Hemschemeier, 2002). This result was confirmed by

microarray studies (Zhang et al., 2004). The promoter of aox1 is rather unresponsive to

stimuli that are known to induce AOX activity in higher plants, such as stress, respiratory

inhibitors or metabolites (Moller, 2001), but it is stimulated by the absence of ammonium

and the presence of nitrate (Baurain et al., 2003). Nothing is known about the role and

expression of aox2.

C. reinhardtii mutant strain M.90, which is deficient in COX activity, exhibits a lower

respiratory activity than the wild type in the first 24 h of sulphur starvation (fig 3-12 B).

This was expected since C. reinhardtii M.90 lacks one of the three major oxygen-

consuming activities that can be found in plant cells. It was shown for other COX mutants

that they exhibit lower respiratory rates (Cardol et al., 2003). This lower respiratory

oxygen uptake could be one more reason for the prolonged aerobic phase of sulphur-

deprived cultures of strain M.90, even if the comparatively high photosynthetic activity of

this strain seems to be the main reason for this phenomenon (see above). In strain M.90, no

inhibition of respiratory oxygen uptake could be observed in the presence of myxothiazol,

as was expected for a COX-deficient strain. Thus, in C. reinhardtii M.90, the alternative

pathway via AOX is primarily responsible for oxygen uptake (fig 3-13).

Residual respiratory oxygen uptake that still occurs after the addition of myxothiazol and

SHAM and which can be inhibited by propyl gallate is due to plastidic respiration via

PTOX. Strain M.90 shows a higher capacity for plastidic respiration via PTOX than the

Discussion

94

wild type. This might be one reason for the efficient re-oxidation of the PQ pool that can

be observed in this strain.

C. reinhardtii strain M.90 exhibits slightly increased oxygen consumption rates upon

sulphur starvation (fig 3-12 B), whereas in the wild type, respiratory activity decreases. In

the wild type, anaerobic conditions are established after one or two days of sulphur

depletion. The low oxygen content of medium and cells could be an explanation for

reduced respiratory capacity.

It should be noted that a recent study showed different results regarding respiratory activity

than are presented in this work. In that study it was shown that a) KCN (a COX inhibitor)

had a rather strong effect when added alone (30 to 60 % inhibition of respiratory activity),

b) also SHAM alone had a significant inhibitory effect (0,3 to 43 %), and c) KCN and

SHAM together inhibited respiration by 89 to 98 %, indicating that the capacity of the

PTOX pathway accounts for not more than 11 % (Antal et al., 2003). Since inhibitory

studies have to be analysed with caution, the difference between both results is not

evaluated here. It should be noted, however, that those authors used KCN, which is a rather

non-specific inhibitor. Additionally, SHAM was applied in quite high concentrations (4

mM). PTOX is homologous to AOX, which is sensitive to SHAM. Plant PTOX was

reported to be affected by high amounts of SHAM (mentioned in Josse et al., 2003), while

in C. reinhardtii PTOX was described to be unaffected by 4 mM SHAM (Cournac et al.,

2000).

The C. reinhardtii Rubisco-deficient strain CC-2803 produces hydrogen in the

presence of sulphur. In strain CC-2803, hydrogen production is almost completely

dependent on electron supply by PSII and it is the only efficient electron sink for this

strain. The ability to produce hydrogen partially rescues strain CC-2803 from its

light-sensitive phenotype.

C. reinhardtii strain CC-2803 produces hydrogen not only in the absence, but also, and at

higher levels, in the presence of sulphur (fig 3-19). The precondition for this process to

occur is the establishment of anaerobiosis.

In the absence of Rubisco protein, one of the most important electron sinks of

photosynthesis is abolished. Linear photosynthetic electron transport produces reduced

ferredoxin, which can pass the electrons to several further acceptors. Most of the electrons

are used to reduce NADP+ to NADPH by ferredoxin-NADPH reductase (FNR). The

majority of the NADPH is consumed by reductive carbon dioxide assimilation in the

Discussion

95

Calvin cycle via the Rubisco enzyme. It was shown that the absence of FNR or Rubisco

leads to significantly lower photochemical quenching and a decrease in overall

photosynthetic electron transport (Hajirezaei et al., 2002; Allahverdiyeva et al., 2005).

Furthermore, the removal of the most important electron sink leads to increased

photoinactivation, a decreased recovery from photoinhibition and overall oxidative stress

(Palatnik et al., 2003; Takahashi and Murata, 2005). The C. reinhardtii Rubisco mutant

strain CC-2803 that was analysed in this study shows very low PSII fluorescence (data not

shown) under each growth condition. In vivo measurements of the oxygen exchange rate in

CC-2803 reveal that PSII activity of this strain is very low, even in sulphur-replete cultures

(fig 3-22), so that oxygen evolution is always exceeded by oxygen consumption activities.

Therefore, cultures of this Rubisco-deficient strain become anaerobic as soon as they are

cut off from the air (i.e., oxygen) supply (fig 3-21).

Moreover, the high ratio of respiratory oxygen uptake rates over oxygen production rates

explains why C. reinhardtii strain CC-2803 is less dependent on acetate to pass into

anoxia. It was shown that the C. reinhardtii wild type cannot establish anaerobic

conditions in the absence of acetate. If strain CC-2803 is cultivated autotrophically,

removal of oxygen is somewhat slower, but the culture becomes anaerobic within one day.

Hydrogen production by strain CC-2803 is nevertheless lower if the cells are not supplied

with acetate. Clearly, however, there is another reason for the dependence on acetate in the

Rubisco-deficient strain compared with the wild type, since hydrogen production in strain

CC-2803 is not supplied by electrons from oxidative starch degradation (see below).

Indeed, lower hydrogen accumulation in autotrophically grown cultures of strain CC-2803

could be due either to less efficient oxygen removal within the cells or to yet lower

photosynthetic activity. It was suggested that photoinactivation of PSII, as a consequence

of an interruption of the Calvin cycle, is caused primarily by the inhibition of protein

synthesis in the chloroplast, which is essential for the repair of PSII (Takahashi and

Murata, 2005). In the Rubisco-deficient C. reinhardtii strain, the assimilation of acetate

might compensate to a certain degree for the inability to reduce carbon dioxide. In the

absence of acetate, strain CC-2803 has no carbon source and this could be responsible for

an even faster degradation of PSII.

Hydrogenase activity and hydrogen accumulation develop differently in C. reinhardtii

strain CC-2803 and the wild type. In strain CC-2803, in vivo hydrogen evolution can be

Discussion

96

detected only a few hours after the culture has been sealed. Hydrogen production rates

increase for maximally one day and decrease gradually afterwards. It is conspicuous that

both in the presence and in the absence of sulphur, in vivo hydrogenase activities parallel

oxygen evolution rates (compare fig 3-20 and 3-22). In the presence of sulphur,

photosynthetic oxygen production is higher and lasts longer. Accordingly, hydrogen

production in sulphur-containing medium is higher.

The parallel development of PSII activity and hydrogen evolution rates already suggests a

close link between both processes in the C. reinhardtii Rubisco-deficient strain CC-2803.

Furthermore, the inability of this strain to accumulate starch in a large scale indicates that

this organic reserve cannot be a major source of reductant for the hydrogenase pathway.

Consequently, the PSII inhibitor DCMU inhibits hydrogen evolution in Rubisco-deficient

strain CC-2803 almost completely (fig 3-23). The link between hydrogen evolution rates

and PSII activity also explains the different behaviour of sulphur-deprived and sulphur

replete cultures. In the absence of sulphur, the same, and probably even enhanced

influences, as have been observed in the C. reinhardtii wild type, act on PSII activity in

strain CC-2803. A Rubisco mutant is much more light-sensitive (Takahashi and Murata,

2005) even under normal growth conditions. Therefore, PSII activity decreases more

rapidly in the absence of sulphur than in its presence, explaining the lower and more

rapidly decreasing hydrogen evolution rates in sulphur-deprived strain CC-2803.

Summarising these data, it becomes clear that C. reinhardtii Rubisco-deficient strain

CC-2803 has a different hydrogen metabolism than the wild type. Whereas the wild type

only produces hydrogen in significant amounts when it is deprived of sulphur, CC-2803

engages the hydrogenase pathway in sulphur replete medium as soon as it is able to

establish anaerobic conditions. Obviously, the hydrogenase pathway partially rescues

C. reinhardtii strain CC-2803 from its extremely light-sensitive phenotype. In contrast to

sealed cultures that are able to establish anaerobic conditions and dissipate excess electrons

by the hydrogenase, aerated cultures of strain CC-2803 become chlorotic after a few days

(fig 4-3).

Discussion

97

Fig 4-3: Photograph of C. reinhardtii CC-2803

cultures after 5 days of anaerobic (left) or aerobic (right) incubation. The aerated culture shows significant chlorosis.

The behaviour of the Rubisco-deficient C. reinhardtii strain CC-2803 indicates that the

lack of efficient electron sinks is one of the parameters that initiate the drastic

physiological changes that occur in sulphur-deprived C. reinhardtii wild type cultures.

It was proposed that the inactivation of PSII in sulphur-deprived C. reinhardtii cells is not

a consequence of an electron ”tailback” due to a decreased activity of the Calvin cycle,

since it could also be demonstrated in isolated thylakoid membranes in the presence of

artificial electron acceptors (Wykoff et al., 1998). The observed phenomenon could

nevertheless be an irreversible inactivation of PSII caused by less efficient oxidation of the

PQ pool due to replenished electron sinks. The accumulation of starch as a reaction to

nutrient deprivation was described previously (Ball et al., 1990). It is suggested to be a tool

to direct carbon away from biosynthetic pathways that are not functional in the absence of

important nutrients like nitrogen or sulphur (Grossman, 2000). It might be speculated that a

certain threshold of starch is one of the triggers that causes the down-regulation of

photosynthetic activity, since it would signal that the electron and carbon reservoirs of the

cells are filled.

The exceptional behaviour of the Rubisco-deficient C. reinhardtii strain offers further

clues to the advantages of the hydrogenase pathway for the wild type.

This anaerobic plastidic electron “valve” replaces biosynthetic electron sinks and allows

the disposal of electrons. This has several advantages for the algae. The electron potential

within the photosynthetic chain can be relaxed, preventing oxidative and radical damage.

The hydrogenase facilitates continuous photosynthetic electron transport, allowing the

synthesis of ATP. Sulphur-depleted C. reinhardtii cells are able to degrade accumulated

starch and transfer the electrons to the hydrogenase pathway. Thereby, the cells can

Discussion

98

degrade excess carbon reserves and benefit from a further means of generating ATP. The

residual electron transport processes at PSII might also be an instrument to keep some PSII

centres active, so that the cells can generate reductant as soon as they are transferred to

sulphur replete medium again.

4.2 What kind of fermentative system is active in C. reinhardtii?

It was described over 20 years ago that the chlorophyte alga C. reinhardtii exhibits a

complex fermentative metabolism, which is marked by the production of acetate, ethanol

and formate (Gfeller and Gibbs, 1984). The characteristic pattern of end products was

ascribed to the activity of pyruvate formate-lyase (PFL), an enzyme, which was previously

thought to be restricted to prokaryotes (Kreuzberg, 1984). Sulphur-deprived C. reinhardtii

cultures establish a sustained anaerobic metabolism in which the generation of hydrogen

obviously is the major electron sink. However, it was shown that sulphur-deprived green

algae also produce ethanol and formate in significant amounts (Hemschemeier, 2002;

Winkler et al., 2002b). The coexistence of two pathways that are quite unusual for aerobic

(photosynthetic) eukaryotes, a hydrogen pathway catalysed by a [Fe]-hydrogenase and the

fermentation pathway catalysed by PFL, poses interesting questions on the evolutionary

origin of this green alga. This study aimed to characterise the bacterial-type fermentation

system of C. reinhardtii in physiological, genetic and biochemical detail.

The mechanism-based PFL inhibitor hypophosphite prevents the production of

formate in sulphur-deprived C. reinhardtii, but ethanol is still accumulated. Clearly,

further fermentative pathways can replace the PFL system.

PFL cleaves pyruvate non-oxidatively into acetylCoA and formate. While formate is

secreted from C. reinhardtii cells, acetylCoA can be further converted into ethanol and/or

acetate. In dark fermentation, formate, acetate and ethanol are produced in a ratio of 2:1:1,

a typical pattern of PFL fermentation (Gfeller and Gibbs, 1984; Kreuzberg, 1984). In

sulphur-deprived green algae, the ratio of formate to ethanol is roughly 1:1. This could

indicate that all acetylCoA is converted to ethanol. Since the algae are incubated in acetate-

containing medium, it is difficult to differentiate between acetate that was already present

in the medium (roughly 0,4 g x l-1 after 24 h of sulphur starvation) and newly synthesised

acetate, which should make up about 0,03 grams per litre when compared with the amount

of formate (~ 0,06 g x l-1). Previous studies showed a slight increase in acetate

Discussion

99

concentration after approximately 100 h of sulphur starvation (Melis et al., 2000;

Tsygankov et al., 2002). This was not observed in the experiments conducted in this study.

However, if some acetate were to be produced, a part of the accumulated ethanol must

originate from sources other than PFL fermentation.

The first approach to identify PFL as the responsible enzyme for formate production in

sulphur-deprived green algae was to apply hypophosphite, which is a formate analogue and

thereby a mechanism-based inhibitor of PFL (Knappe et al., 1984; Kreuzberg, 1984). If

sulphur-depleted C. reinhardtii cultures are treated with hypophosphite, formate

accumulation is suppressed, indicating that PFL is indeed responsible for formate

production. Interestingly, ethanol accumulation is not significantly influenced by the

addition of hypophosphite. Obviously, another pathway accounts for ethanol production.

Two common fermentative processes that lead to ethanol production are known. The best

characterised of these is ethanol production by yeast. These organisms decarboxylate

pyruvate to acetaldehyde by pyruvate decarboxylase (PDC). Carbon dioxide is released,

while acetaldehyde is further reduced to ethanol by alcohol dehydrogenase.

A second pathway might be of special interest in this context, since it is common for

hydrogen-producing prokaryotes (Pieulle et al., 1997; Stal and Moezelaar, 1997). A

pyruvate ferredoxin-oxidoreductase (PFO) cleaves pyruvate into carbon dioxide and

acetylCoA and reduces ferredoxin during this reaction. Reduced ferredoxin can act as an

electron donor for several hydrogenases. The advantage of this reaction lies in the

production of acetylCoA, which can be converted to acetate to yield ATP. It is also

noteworthy that an enzyme with high similarity to pyruvate ferredoxin-oxidoreductase, the

pyruvate flavodoxin-oxidoreductase, is supposed to be involved in providing reduced

flavodoxin for PFL activation in E. coli (Blaschkowski et al., 1982).

C. reinhardtii possesses a gene encoding a pyruvate decarboxylase (PubMed E15259)

which was shown to be expressed upon sulphur starvation (fig 3-36). Additionally, a

sequence that codes for a protein with high similarity to PFO enzymes was found in the

genome sequence of C. reinhardtii (tab 3-6; appendix 8.3, tab 8-1). However, if PFO is

responsible for ethanol fermentation in sulphur-deprived and hypophosphite-treated algae,

acetylCoA, which is one product of its reaction must almost completely be converted to

ethanol. To test if one of these pathways replaces PFL in hypophosphite-treated

C. reinhardtii cultures, carbon dioxide accumulation was determined in the presence or the

absence of hypophosphite. Indeed, hypophosphite-treated cells produce roughly double the

Discussion

100

amount of carbon dioxide, indicating the occurrence of pyruvate decarboxylation (fig

3-28). It was also shown that during dark fermentation, formate production by

C. reinhardtii can be inhibited by hypophosphite, whereas ethanol and carbon dioxide

production increase (Kreuzberg, 1984).

At the moment, it is not possible to distinguish between PDC or PFO catalysis. The pdc

gene was shown to be transcribed; however, the expression of the putative pfo gene was

not analysed in this study. If PFO is responsible for some ethanol production in

C. reinhardtii, electrons that are transferred to ferredoxin might be a source for hydrogen

production. However, hypophosphite treatment of sulphur-starved C. reinhardtii cultures

does not result in a higher hydrogen yield. Hydrogen accumulation in the absence or the

presence of hypophosphite is nearly the same. Often, a decrease of hydrogen yields could

be observed in hypophosphite-treated cultures (data not shown).

At this point it should be noted that C. reinhardtii was also reported to produce glycerol

and traces of D-lactate (Gfeller and Gibbs, 1984; Kreuzberg, 1984). Low amounts of

glycerol could also be detected in sulphur-deprived C. reinhardtii cells (data not shown).

Genes encoding the enzymes involved in glycerol production, glycerol-3-phosphate

dehydrogenase and glycerol-3-phosphatase, were found in the genomic sequence of

C. reinhardtii (appendix 8.3, tab 8-1). In view of all the potential pathways of fermentation

it can be stated that this green alga has a remarkable fermentative flexibility, which is a

mixture of bacterial-, plant- and yeast-type fermentation (tab 4-1).

end product(s) enzyme(s) most common to

formate, ehanol, acetate PFL (PflA), AdhE, ACK, PTA bacteria, chytrid fungi

ethanol, carbon dioxide PFO, AdhE bacteria, chytrid fungi

ethanol, carbon dioxide PDC, ADH plants, yeast1

glycerol GPD, GPP yeast

Tab 4-1: List of fermentative end products that can be detected in anaerobic C. reinhardtii cultures, the

enzymes responsible for their production and the most common occurrence of the respective pathway.

(PFL (PflA) = pyruvate formate lyase (PFL activase), AdhE = multifunctional acetaldehyde and alcohol dehydrogenase, ACK = acetate kinase, PTA = phosphotransacetylase, PFO = pyruvate ferredoxin oxidoreductase, PDC = pyruvate decarboxylase, ADH = alcohol dehydrogenase, GPD = glycerol-3-phosphate dehydrogenase, GPP = glycerol-3-phosphate phosphatase). 1 PDC is present is very few bacterial species, e.g. Zymomonas mobilis.

Discussion

101

The PFL pathway is typical for bacteria, such as enterobacteria (Kessler and Knappe,

1996) or clostridia (Weidner and Sawers, 1996). Also PFO is typically found in bacteria

(Pieulle et al., 1997; Stal and Moezelaar, 1997). PDC is common in plants (e.g. Kürsteiner

et al., 2003) and yeast (reviewed by Pronk et al., 1996). Glycerol generation is for example

found in yeast, where it is a reaction towards osmotic, anaerobic and oxidative stress

(Påhlman et al., 2001; Remize et al., 2003). A schematic overview of the major (known)

fermentative pathways that, according to the presence of corresponding genes, could be

active in C. reinhardtii is given in fig 4-4.

Fig 4-4: Schematic overview of the possible pathways of glucose fermentation in C. reinhardtii. Glucose is glycolytically converted to pyruvate. One of the intermediates, dihydroxyacetone phosphate (DHAP), can be converted to glycerol-3-phosphate by glycerol-3-phosphate dehydrogenase (GPD) and further dephosphorylated to glycerol (not shown). Pyruvate can be degraded via three possible pathways. Pyruvate decarboxylase (PDC) decarboxylises pyruvate to acet-aldehyde. The latter is reduced to ethanol by an alcohol dehydrogenase (ADH). Pyruvate formate-lyase (PFL) cleaves pyruvate into formate, which is secreted, and acetylCoA. The latter can be reduced to ethanol by AdhE, combining acetaldehyde- and alcohol dehydrogenase, or converted to acetate by the sequential action of phosphotransacetylase (PTA) and acetate kinase (ACK). This reaction yields one ATP per molecule of acetylCoA. Pyruvate ferredoxin/flavodoxin-oxidoreductase (PFO) decarboxylises pyruvate to acetylCoA and reduces ferredoxin (Fd) at the same time. AcetylCoA could be converted in the same way as is described for the PFL pathway (indicated by a dotted line and a question mark).

pyruvate

DHAP

formate acetylCoA

ethanol

acetyl-P

acetate

Pi

CoAPTA

ADP

ATPACK

2 NADH

CoA + 2 NAD+

PFL

CO2

acetylCoA

?

Fdox Fdred

PFO

acetaldehyde

CO2

PDC

ethanol

glycerol-3-phosphate

NADHNAD+

GPD

PflANADH

NAD+ADH

glucose

CoA

AdhE

Discussion

102

The accumulation of fermentative products is dependent on starch. The synthesis of

PFL and AdhE is not correlated with effective fermentative activity.

It is conspicuous that neither the C. reinhardtii PSII mutant FuD7 nor the Rubisco-

deficient strain CC-2803 produces significant amounts of formate and ethanol when

deprived of sulphur (fig 3-29). Since both strains become anaerobic very soon after the

transfer to a sealed culture vessel, the low amount of fermentative products is obviously

not due to the presence of oxygen. It was shown previously that classic fermentation of

dark-adapted C. reinhardtii cultures is accompanied by accelerated starch breakdown,

indicating that starch is the major source of fermentable glucose (Gfeller and Gibbs, 1984).

Neither strain FuD7 (data not shown) nor strain CC-2803 contains significant amounts of

starch. This probably explains the low extent of classic fermentation in these two strains.

Nevertheless, synthesis of PFL and AdhE protein in the Rubisco-deficient C. reinhardtii

strain is not impaired. Both proteins can be detected in significant amounts by Western

Blot analyses. This indicates that the synthesis of these proteins is not correlated with the

effective fermentative activity. In E. coli, the expression of the pfl gene is strongly

enhanced as a response to anaerobiosis (Sawers and Böck, 1988, 1989; Sawers and

Suppman, 1992). Expression of the adhE gene is also enhanced in the absence of oxygen.

Obviously, the NADH/NAD+ ratio plays a significant role in AdhE synthesis: the higher

the ratio, the more AdhE protein is produced (Leonardo et al., 1993).

Both the absence of oxygen and a high NAD(P)H/NAD(P)+ ratio are given in Rubisco-

deficient C. reinhardtii strain CC-2803. Therefore, the production of PFL and AdhE in

C. reinhardtii might be regulated in a similar way as in E. coli inasmuch as the expression

of the pfl and adhE genes is not correlated with the metabolite status of fermentation.

The polypeptide sequences deduced from C. reinhardtii genes encoding PFL and its

activating enzyme, PFL activase (PflA), are significantly homologous to other known

PFL and PflA proteins. A localisation of PFL in an organelle can be suggested.

PFL is an enzyme, which harbours a glycyl radical in its active form (Knappe et al., 1984;

Wagner et al., 1992). The glycyl radical is post-translationally inserted into the PFL

polypeptide by PFL activase (PflA). In E. coli, PflA needs reduced flavodoxin and

S-adenosylmethionine for PFL activation (Knappe et al., 1984). The genes for pfl and pflA

were isolated and characterised in this study. An annotated pfl sequence was found in the

published genome of C. reinhardtii (JGI; C_1330014) and amplified with oligonucleotides

Discussion

103

that were derived from this sequence. In the case of pflA, only a fragment was found in

PubMed (BE025007), making RACE PCR experiments necessary.

The polypeptides that were deduced from the pfl and pflA cDNA sequences were compared

with other known PFL and PflA proteins. The C. reinhardtii PFL protein shows high

similarity to other PFL proteins. E. coli and C. reinhardtii PFL share 52 % identical amino

acids (see an alignment of several PFL polypeptides in appendix 8.2, fig 8-3). The

conserved regions around the catalytic cysteine residues (IACCVS) and the glycyl radical

(RVSGYAV) are identical in both enzymes (fig 3-30). However, the alignments reveal that

the algal protein includes an N-terminal extension (70 amino acids), which shows no

homology to any of the analysed PFL proteins. If this sequence is removed in silico, the

identity between E. coli and C. reinhardtii PFL accounts for 56 %.

The polypeptide deduced from the C. reinhardtii pflA cDNA sequence is identified as PFL

activase by homology searches, but it shows a rather low degree of identity to other PflA

proteins. E. coli and C. reinhardtii PflA exhibit 27 % identity. The highest similarity was

found between C. reinhardtii PflA and the PflA proteins of N. frontalis (34 %) and

Rhodospirillum rubrum (34 %). The PflA enzyme also includes an N-terminal extension

(90 amino acids) with no significant similarity to any proteins or peptides in the data base.

The N-terminal sequences of the algal enzymes could represent organelle-targeting signal

peptides. It was shown previously that PFL activity in C. reinhardtii is located within the

chloroplast and the mitochondrion (Kreuzberg, 1984). The predicted localisation in the

chloroplast has to be taken with caution because the preparation of mitochondrion-free

chloroplasts from C. reinhardtii is quite difficult. However, the results strongly suggest

that PFL is localised in one of the organelles or in both. A dual function of a transit peptide

is rather unusual, but it was described for pea glutathione reductase (Creissen et al., 1995;

Chew et al., 2003).

When analysed with different prediction programs that are available on the ExPASy

Proteomics tools web page (http://au.expasy.org/tools/), both C. reinhardtii PFL and PflA

are predicted to be targeted to organelles. Frequently, the putative signal peptide is

designated to be mitochondrial. Of course these results have to be considered with some

caution, since the prediction programs have not been designed for C. reinhardtii, whose

signal peptides differ from those of higher plants (Franzén et al., 1990).

Discussion

104

It is noteworthy that all enzymes that are involved in the PFL pathway (AdhE, acetate

kinase (ACK) and phosphotransacetylase (PTA)) occur in two copies in the C. reinhardtii

genome. For both putative AdhE enzymes and for ACK1, a putative organellar transit

peptide was annotated (see appendix 8.3, tab 8-1).

In this context a possible interaction of fermentation via PFL and hydrogen production

should be discussed. By engaging the hydrogenase pathway, C. reinhardtii has a very

efficient way to remove reducing equivalents. It was shown, additionally, that the

hydrogenase enzymes present do not operate at full capacity. Therefore an additional

pathway for recycling NAD(P)H in the chloroplast seems redundant. The algae engage the

PFL pathway nevertheless. It might be speculated that the PFL pathway serves a more

local disposal of excess reducing equivalents.

The C. reinhardtii PFL protein is functionally expressed in E. coli mutants, which are

deficient in their own PFL. This demonstrates that C. reinhardtii PFL is recognised by

E. coli PflA. On the other hand, algal PflA fails to activate E. coli PFL.

C. reinhardtii PFL and PflA were analysed for their functionality in E. coli. Mutant strains

of E. coli that are deficient in PFL and PflA, respectively, were transformed with plasmids

that harbour the algal pfl and pflA cDNAs under the control of bacteriophage T7 RNA-

polymerase promoter. The analysis of the respective E. coli strains revealed two interesting

features of E. coli, which will not be discussed in detail since they are not relevant for the

topic of this study. They should be noted nevertheless. Usually, PFL fermentation in E. coli

produces formate, which is secreted from the cells but is re-imported when a certain

threshold of medium pH is reached (Mnatsakanyan et al., 2002; Sawers, 2005). Inside the

cells, formate triggers the induction of the formate hydrogen lyase (FHL) complex that

disproportionates formate into carbon dioxide and hydrogen. This well studied behaviour

could not be observed in the BL21 E. coli strains that were used in this study, indicating

that an important factor for formate degradation is missing in this constructed strain.

Therefore, active PFL enzyme in BL21 derivatives was concluded from formate

accumulation, since the analysed strains did not re-import formate.

A second, maybe even more unusual behaviour was observed in derivatives of E. coli

strain 234M11 (which, in turn, is a derivative of E. coli MC4100; Sauter and Sawers,

1990). Since pflA cDNA is present in pET9a under the control of T7 promoter, an extra

Discussion

105

source of T7 polymerase has to be introduced into this strain, which has no genomic copy

of T7 polymerase. Unfortunately, co-expression with two vectors – one of the pET9a

derivatives and pUPI-2 that contains a gene for T7 polymerase – failed. However, it was

observed that hydrogen production of the control strain (234M11 which was transformed

with a plasmid harbouring the E. coli pflA gene) was restored upon introduction of the pflA

gene present in pET9a.

No reference describing the ability of E. coli RNA polymerases to recognize T7 promoter

could be found. However, transcription from other viral promoters such as SV40 or CMV

was described (Lewin et al., 2005). Furthermore, the leakiness of T7 promoter systems is a

well known problem. Therefore, a weak but significant transcription of the pflA gene by

E. coli RNA polymerase seems possible, especially if the presence of the respective gene

product is an advantage for the cells. In the analysed system, low expression of pflA is

probably sufficient to activate enough PFL proteins for efficient fermentative growth.

Generally, the complementation of E. coli strains BL21∆pfl and 234M11 (∆pflA) with

E. coli pfl and pflA genes, respectively, shows that the chosen system, even if rather

designed for protein over-production, is a suitable tool to test the activity of the algal

proteins in vivo. Strain BL21.36 (BL21∆pfl in which E. coli pfl was re-introduced) is

clearly rescued from the phenotype of pfl knockout. Whereas BL21∆pfl does not excrete

formate and grows poorly under anaerobic conditions, BL21.36 shows both formate

production and growth which are even higher than in the “wild type” (BL21(DE3)pLys).

This “over-complementation” is probably due to the enhanced PFL synthesis in BL21.36

that was shown by Western Blot analyses (fig 3-34). However, both growth and formate

production in BL21.36 started later than in the BL21 “wild type”. This might be due to

some regulatory events that occur differently in “wild type” and complemented mutant

strains. In E. coli wild type, PFL expression is enhanced 10 to 15 times upon anaerobiosis.

In the constructed strains analysed in this study, PFL protein is constitutively produced in

high amounts due to the leakiness of the T7 system. It is possible that the relatively high

amounts of inactive PFL protein interfere with efficient induction of the fermentative

machinery.

Strains BL21.28 (transformed with complete C. reinhardtii pfl) and BL21.32 (transformed

with truncated C. reinhardtii pfl) show significant formate production, which demonstrates

that the algal PFL proteins are functionally synthesised in E. coli. Both forms of the

Discussion

106

C. reinhardtii PFL (with and without putative transit peptide) are active in formate

production, indicating that the N-terminal extension sequence is not required for catalytic

activity of PFL in E. coli. This result also shows that the algal PFL enzyme is recognised

and activated by E. coli PflA. It was proposed that PflA recognises a heptapeptide

sequence, which surrounds the radical harbouring glycine residue (Frey et al., 1994). In

E. coli PFL, this sequence is RVSGYAV, and it is exactly the same sequence in C.

reinhardtii PFL.

Although E. coli pfl mutants complemented with algal PFL produce formate, they are not

phenotypically rescued by the algal proteins, since their growth is comparable to that of

BL21∆pfl. This might be due to less efficient pyruvate degradation in BL21.28 and

BL21.32. It was shown in parallel experiments that strep-tagged C. reinhardtii PFL is not

produced in a soluble form in E. coli strains incubated at 37°C or at ambient temperatures.

Only when the cells were incubated at 4°C did expression of C. reinhardtii PFL in E. coli

yield soluble protein that was active in in vitro pyruvate degradation (Jacobs, 2005). The

complementation assays performed in this study were conducted at room temperature to be

able to observe physiological reactions which would have been too slow at 4°C. Therefore

it can be assumed that only a small fraction of the algal PFL protein was soluble in the

BL21∆pfl derivatives.

On the other hand, the BL21∆pfl strains complemented with C. reinhardtii proteins show a

similar formate production as the control strains when related to the optical density. This

suggests that the relative efficiency of the algal proteins in E. coli should be high enough to

allow anaerobic growth. It is conceivable that the expression of the algal PFL protein

depletes the pools of rare tRNAs in E. coli because of the different codon usage of both

species. The average GC content of the E. coli and C. reinhardtii genomes are 50 % and

66 %, respectively. The comparison of the C. reinhardtii pfl sequence with the E. coli

codon usage table using the internet programme graphical codon usage analyser

(http://gcua.schoedl.de/) showed that there are more than 100 codons in the C. reinhardtii

pfl sequence that require tRNAs, which are rare in E. coli. Some of these critical codons

are directly adjacent or close to each other (data not shown). Therefore, the enforced

expression of the algal PFL protein could deplete the E. coli tRNA pools and therewith

interfere with its physiological status.

Whereas C. reinhardtii PFL is produced in an active form in E. coli, the algal PFL activase

(PflA) is not functional in activating the E. coli PFL protein. E. coli 234 M11 strains that

were transformed with the complete or a truncated C. reinhardtii pflA cDNA sequence did

Discussion

107

not exhibit growth, formate production or hydrogen accumulation under anaerobic

conditions. However, a 234M11 strain in which the E. coli innate pflA gene was re-

introduced was clearly rescued from the phenotype of 234M11, demonstrating that the

chosen system is functional in principle.

The fact that the algal PFL is active in E. coli, but PflA is not, seems paradoxical. To

activate a PFL enzyme, the respective PflA protein has to recognise and bind the

polypeptide substrate. Therefore, if the algal PFL is recognised by E. coli PflA, the

interacting three-dimensional structures should be highly similar. It was suggested that in

E. coli, PflA recognises a short domain of seven amino acids around the glycyl radical

(Frey et al., 1994). In the C. reinhardtii PFL sequence, these seven amino acids are

completely identical to those in the E. coli polypeptide. Consequently, it could be assumed

that the recognition sites in both C. reinhardtii and E. coli PflA enzymes are highly similar.

Two reasons could account for E. coli PFL being not activated in the presence of algal

pflA-genes. First, the C. reinhardtii PflA enzyme might not be produced in its active form.

It has no unusual prosthetic groups which might be difficult for E. coli to synthesise. In

fact, the polypeptide sequence of C. reinhardtii PflA suggests that it has the same type of

FeS cluster that it is usually present in PflA proteins (Külzer et al., 1998; Sofia et al.,

2001). Again, the different codon usage of C. reinhardtii and E. coli might be the reason

why the algal PflA is not functionally produced. The C. reinhardtii pflA sequence has a GC

content of 60 %.

Second, it is possible that E. coli PflA has a broader substrate specificity than the algal

enzyme. According to the current version of the C. reinhardtii genomic sequence, this

organism has only one PFL protein and no further PFL-like enzymes. In E. coli, however,

three more genes that encode proteins with amino acid sequence similarity to PFL have

been identified (Gelius-Dietrich and Henze, 2004). At least one of them, the 2-ketobutyrate

formate-lyase TdcE, exhibits PFL activity and can be activated by PflA (Heßlinger et al.,

1998). It is conceivable that the algal PflA enzyme is more specific for its PFL enzyme

than the E. coli PflA, which might be needed for the activation of several PFL-like

proteins. It should also be noted again that the C. reinhardtii and E. coli PflA polypeptides

share only 27 % amino acid identity. Even if the putative transit peptide of the algal PflA is

cleaved off in silico, the identity between both sequences is still only at 36 %.

Discussion

108

C. reinhardtii is one of the few eukaryotic organisms that possesses a PFL system.

Moreover, this alga is the only eukaryote with both an oxygenic photosynthesis and a

marked anaerobic metabolism.

Summarising all the data presented in this work it can be stated that C. reinhardtii

possesses a bacterial-type fermentative system via PFL. There are only two eukaryotic

lineages described to date that carry out PFL fermentation, some chlorophyte algae

(Kreuzberg, 1984) and some obligate anaerobic chytrid fungi (Marvin-Sikkema et al.,

1993; Akhmanova et al., 1999; Boxma et al., 2004). A monophyletic origin of the

eukaryotic PFL proteins was suggested (Gelius-Dietrich and Henze, 2004).

In the chytridiomycetes Neocallimastix frontalis and Piromyces sp. E2 that live in the

gastrointestinal tracts of ruminants, the PFL pathway is involved in anaerobic energy

production in hydrogenosomes. These organelles are present in several strictly anaerobic

protists and they are proposed to be relatives of mitochondria (Akhmanova et al., 1998;

Embley et al., 2002). Interestingly, hydrogenosome-containing protists are, besides

chlorophyte algae, also the only other known eukaryotes in which a [Fe]-hydrogenase has

been characterised (Horner et al., 2000).

At first glance, the occurrence of a complex fermentative metabolism characterised by PFL

and a [Fe]-hydrogenase in green algae seems to be a biological curiosity, since both

enzymes are exquisitely sensitive towards oxygen, while green algae are organisms with an

oxygenic photosynthesis. However, recent publications and the presented study show that

both anaerobic pathways are extensively engaged in C. reinhardtii, especially under

anaerobic conditions, which are internally induced by the absence of sulphur.

This metabolism is distinguished by the presence of fermentative pathways catalysed by

oxygen-sensitive proteins in illuminated green algae that usually carry out oxygenic

photosynthesis. Because of this apparently paradoxical behaviour of sulphur-deprived

C. reinhardtii, the term “photofermentation” was proposed (Winkler et al., 2002b).

Summary

109

5 Summary

The unicellular green alga C. reinhardtii is a plant organism that shares many features with

its younger relatives, the higher plants. However, this chlorophyte alga shows a special

type of anaerobic metabolism that is quite unusual for plants or eukaryotes at all. In the

absence of sulphur, a sustained anaerobic metabolism is established in illuminated

C. reinhardtii cultures. This metabolism is characterised by the production of hydrogen,

formate and ethanol.

This study aimed to get deeper insights into the hydrogen and fermentative metabolism of

C. reinhardtii and has come to several new conclusions regarding metabolic adjustments

that occur in the initial aerobic phase, the electron source for hydrogen production and the

vital role of the hydrogenase pathway. Furthermore, the pyruvate fermentation system of

C. reinhardtii could be characterised in detail, demonstrating a remarkable efficiency and

flexibility of the anaerobic metabolism of this green alga.

After the transfer to sulphur-free medium, C. reinhardtii cultures first pass an aerobic

phase which is characterised by active photosynthetic oxygen evolution and carbon dioxide

fixation. In this phase, the algae accumulate large amounts of starch. It could be shown that

in the subsequent anaerobic phase, electrons both from this organic reserve and from

residual PSII activity enter the hydrogenase pathway and contribute to hydrogen

production.

One of the most remarkable reactions to sulphur deprivation is the strong decrease of PSII

activity that was proposed to be a regulated process and a specific response to limitations

in sulphur. This work demonstrates that the decline of photosynthetic oxygen evolution is

mainly correlated with the light intensity that is centred on PSII. Sulphur deprivation seems

to be a condition in which photoinhibitory processes cannot be compensated any more,

leading to a sequential loss of active PSII centres.

The establishment of anaerobic conditions, which goes hand in hand with the decrease of

PSII activity, can only be observed in the presence of acetate, which is furthermore a

carbon source for starch accumulation.

A very special behaviour was observed in the Rubisco-deficient C. reinhardtii strain

CC-2803. This strain produces hydrogen in the absence, but also in the presence of

sulphur. This is the first proof that algae can also establish a sustained hydrogen

metabolism in the presence of sulphur, which might provide new possibilities for

biotechnological approaches.

Summary

110

The only significant electron source for hydrogen generation in strain CC-2803 is PSII.

The possibility to transfer electrons to the hydrogenase is the major electron sink for

Rubisco-deficient algae and partially rescues the light-sensitive phenotype of strain

CC-2803.

The sustained anaerobic metabolism of sulphur-deprived C. reinhardtii cultures is not only

characterised by hydrogen accumulation, but also by the accumulation of formate and

ethanol. Usually, formate production is due to pyruvate formate-lyase (PFL). The PFL

system of C. reinhardtii was analysed in genetic and biochemical details. The genes

encoding PFL and its activating enzyme, PFL activase (PflA), were isolated. The deduced

polypeptide sequences show significant homologies to other known PFL and PflA proteins.

The algal PFL protein and AdhE, a multifunctional protein complex acting in downstream

reactions of the PFL pathway, cross-react with antibodies raised against the respective

E. coli proteins, indicating a high similarity between the algal and the bacterial enzymes.

The algal pfl gene is functionally expressed in E. coli as shown by restored formate

production of E. coli ∆pfl strains that were transformed with the C. reinhardtii pfl gene.

C. reinhardtii PflA, however, fails to activate E. coli PFL.

The bacterial-type PFL system is not the only fermentative pathway in C. reinhardtii.

Ethanol production can also be performed by pyruvate decarboxylase or pyruvate

ferredoxin-oxidoreductase. Additionally, the fermentative production of glycerol is

observed. Obviously, this alga disposes of a highly complex and flexible fermentation

apparatus, in which aspects of bacterial-, plant- and yeast-type fermentation are mixed.

The establishment of a marked anaerobic metabolism in illuminated cultures of

C. reinhardtii, which is an organism with oxygenic photosynthesis, is a particularity. Since

hydrogen production in algae is also light-dependent, we have termed this special

metabolism of sulphur-starved green algae “photofermentation”.

Zusammenfassung

111

6 Zusammenfassung

Die einzellige Grünalge C. reinhardtii ist ein pflanzlicher Organismus, der seinen jüngeren

Verwandten, den höheren Pflanzen, in vielerlei Hinsicht ähnelt. C. reinhardtii verfügt

jedoch über einige Eigenschaften, die für Pflanzen, oder Eukaryonten generell, sehr

ungewöhnlich sind. Unter Schwefelmangel bilden belichtete C. reinhardtii-Kulturen einen

lang anhaltenden anaeroben Stoffwechsel aus, der durch die Bildung von Wasserstoff,

Formiat und Ethanol gekennzeichnet ist.

Ziel dieser Arbeit war es, tiefere Einsichten in den Wasserstoff- und Gärungsstoffwechsel

von C. reinhardtii zu gewinnen. Es wurden einige neue Erkenntnisse bezüglich der

Stoffwechselanpassungen in der anfänglichen aeroben Phase, der Elektronenquelle der

Wasserstoffbildung und der essentiellen Bedeutung der Hydrogenase gewonnen.

Weiterhin konnte durch eine detaillierte Charakterisierung der Pyruvatgärung in

C. reinhardtii eine außergewöhnliche Effizienz und Flexibilität des anaeroben

Stoffwechsels dieser Alge gezeigt werden.

Nach der Überführung in schwefelfreies Medium durchlaufen C. reinhardtii-Kulturen

zunächst eine aerobe Phase, die durch aktive photosynthetische Sauerstoffproduktion und

Kohlendioxidfixierung geprägt ist. In dieser Phase akkumulieren die Algen große Mengen

an Stärke. In der vorliegenden Arbeit konnte gezeigt werden, dass sowohl diese organische

Kohlenstoffreserve als auch die noch vorhandene PSII-Aktivität Elektronenquellen für die

in der anschließenden anaeroben Phase erfolgende Wasserstoffproduktion sind.

Eine der auffälligsten Folgen des Schwefelmangels ist die deutliche Abnahme der PSII-

Aktivität. Es wurde vermutet, dass dieser Prozess reguliert und eine spezifische Reaktion

auf das Fehlen von Schwefel ist. Diese Arbeit zeigt, dass die Abnahme der PSII-Aktivität

weitgehend auf lichtabhängige Photoinaktivierung zurückzuführen ist. Schwefelmangel

scheint eine Bedingung zu sein, in der photoinhibitorische Prozesse nicht mehr

kompensiert werden können, was zu einer stetigen Abnahme an funktionalen PSII-Zentren

führt. Die Entstehung anaerober Bedingungen, die einhergeht mit der Abnahme an PSII-

Aktivität, kann nur beobachtet werden, wenn den Zellen Acetat angeboten wird. Acetat ist

außerdem eine Kohlenstoffquelle für den Aufbau von Stärke.

Ein sehr auffälliges Verhalten konnte bei Untersuchungen des Rubisco-defizienten

C. reinhardtii-Stammes CC-2803 beobachtet werden. Dieser Stamm bildet zwar auch

Wasserstoff in Schwefelmangelmedium, jedoch ebenfalls und in größerer Menge in

schwefelhaltigem Medium. Damit wurde erstmalig gezeigt, dass Algen auch in

Zusammenfassung

112

Anwesenheit von Schwefel einen ausgeprägten Wasserstoffmetabolismus entwickeln

können. Dies zeigt ganz neue Möglichkeiten für die biotechnologische Anwendung auf.

In Stamm CC-2803 ist die Wasserspaltung an PSII die vorwiegende Elektronenquelle für

die Wasserstoffbildung. Die Möglichkeit, Elektronen auf die Hydrogenase zu übertragen

ist die einzige bedeutsame Elektronensenke für die Rubisco-defizienten Algen und lindert

die extreme Lichtempfindlichkeit der Zellen.

Der lang anhaltende anaerobe Stoffwechsel von C. reinhardtii-Kulturen in

Schwefelmangelmedium ist nicht nur durch die Bildung von Wasserstoff, sondern auch

durch die Akkumulation von Formiat und Ethanol gekennzeichnet. Im Allgemeinen wird

die fermentative Bildung von Formiat durch Pyruvat-Formiat-Lyase (PFL) katalysiert. Das

PFL-System von C. reinhardtii wurde in der vorliegenden Arbeit in genetischen und

biochemischen Details charakterisiert.

Die Gene, welche die PFL und das PFL-aktivierende Enzym, PFL-Aktivase (PflA)

kodieren, wurden isoliert. Die abgeleiteten Polypeptidsequenzen zeigen deutliche

Homologien zu bereits bekannten PFL- and PflA-Proteinen. Sowohl PFL als auch AdhE,

ein multifunktionaler Enzymkomplex, der ebenfalls am PFL-Weg beteiligt ist, reagieren

mit Antikörpern, die gegen die entsprechenden E. coli-Proteine gerichtet sind. Dies deutet

auf eine starke Ähnlichkeit zwischen den jeweiligen Proteinen aus C. reinhardtii und

E. coli hin.

Das pfl-Gen aus C. reinhardtii wird in E. coli funktionell exprimiert. Dies zeigte sich

anhand der wiederhergestellten Fähigkeit zur Formiat-Bildung in E. coli-Stämmen, deren

eigenes pfl-Gen inaktiv ist, und die mit dem pfl-Gen aus C. reinhardtii transformiert

wurden. Das PflA-Enzym aus der Alge dagegen kann die PFL aus E. coli nicht aktivieren.

Das PFL-System ist nicht der einzige Gärungsweg in C. reinhardtii. Ethanol-Bildung kann

auch durch Pyruvat-Decarboxylase oder Pyruvat-Ferredoxin-Oxidoreduktase erfolgen.

Zusätzlich ist Glyceringärung möglich. Offensichtlich verfügt diese Alge über einen hoch

komplexen und flexiblen Gärungsapparat, welcher Aspekte bakterieller, pflanzlicher und

Hefe-Gärung beinhaltet.

Die Besonderheit des Gärungsstoffwechsels in Schwefelmangelkulturen von C. reinhardtii

ist die Tatsache, dass er in einem Organismus mit grundsätzlich oxygener Photosynthese

unter normalen Lichtbedingungen stattfindet. Wegen dieser Einzigartigkeit und der

Tatsache, dass die Wasserstoffbildung in C. reinhardtii lichtabhängig ist, haben wir diesen

speziellen Stoffwechsel als Photofermentation bezeichnet.

.

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Appendix

125

8 Appendix

8.1. Assembly of the pflA-cDNA and deduced oligonucleotides

Fig 8-1: Schematic overview of oligonucleotides that were used for the isolation of the complete pflA cDNA by RACE-PCR.

Fig 8-2: Schematic overview of the assembly of the pflA-sequence and the oligonucleotides that were used for the amplification of the whole cDNA and the amplification of the open reading frame for subsequent cloning into pASK-IBA-7 and pET9a.

0,1 kb

PFL-Act-1110

PFL-Act-2 425

PFL-Act-380

PFL-Act-4 360

PFL-Act-5 330

Putative pflA-fragment (PubMed BE025007), 559 bps

PFL-Act-6 314

0,1 kb

5’-RACE fragment, 540 bps3’-RACE fragment, 1683 bps

ORF (1011 bps)2022 bps

Putative pflA-fragment (PubMedBE025007), 559 bps

Act-total-31819

Act-total-586

Act-total-3-nest1722

Act-total-5-nest90

Act-IBA7-3 + Act-pET-Bpu-21115

Act-IBA7-5 + Act-pET-Nde105

Act-pET-ohneSP375

Appendix

126

8.2. Alignments of PFL and PflA polypeptides

Fig 8-3: Alignment of selected PFL polypeptide sequences. 1 = C. reinhardtii, 2 = Piromyces sp. E2 (Akhmanova et al., 1999), 3 = E. coli (Blattner et al., 1997), 4 = Clostridium pasteurianum (Weidner and Sawers, 1996), 5 = Bacillus anthracis (Read et al., 2003), 6 = Thermosynechococcus elongates (Nakamura et

al., 2002). The alignment was performed with the programme SECentral. The black frames indicate the two active site cysteine residues and the conserved region around thy glycyl radical.

Appendix

127

Fig 8-4: Alignment of selected PflA polypeptide sequences. 1 = C. reinhardtii, 2 = Neocallimastix frontalis (Marvin-Sikkema et al., 1993), 3 = E. coli (Blattner et al., 1997), 4 = C. pasteurianum (Weidner and Sawers, 1996), 5 = Streptococcus thermophilus (Bolotin et al., 2004), 6 = T. elongates (Nakamura et al., 2002). The Alignment was done in SECentral. The black frames indicate the typical Cys-x-x-x-Cys-x-x-Cys motif (with x = any amino acid), which coordinates the FeS-cluster of PflA, and the glycine rich region.

8.3. Annotated sequences encoding fermentative enzymes in C. reinhardtii

enzyme number name in JGI annotation (user) (size)

PFL 1 C_1330014 pyruvate formate-lyase; may contain organelle targeting peptide

(Laurens Mets) (830 aas, ~91 kDa)

C_840029

(complete)

Adh1; dual function alcohol dehydrogenase / acetaldehyde

dehydrogenase [EC:1.1.1.1 /1.2.1.10], probably mitochondrial

(PMID: 14756315) (Olivier Vallon) (954 aas, ~102kDa)

AdhE 2 C_920036,

C_5260001

(fragments)

Adh2; C_920036: alcohol dehydrogenase, class IV (iron-

containing), probably mitochondrial (PMID: 14756315) (Olivier

Vallon); C_5260001: acetaldehyde dehydrogenase; could be the

N-terminal part of C_920036 (Olivier Vallon)

FDH2;

C_70224

formaldehyde dehydrogenase, glutathione-dependent, Zinc-

containing alcohol dehydrogenase class III (Laurens Mets)

(377 aas, ~40 kDa)

FDH1;

C_70223,

formaldehyde dehydrogenase, glutathione-dependent, Zinc-

containing alcohol dehydrogenase class III (Laurens Mets)

(419 aas, ~55 kDa)

C_10090

Zinc-containing alcohol dehydrogenase, KOG1 description:

alcohol dehydrogenase, class V (395 aas, ~42 kDa)

ADH 5

C_130168, Zinc-containing alcohol dehydrogenase, KOG1 description:

sorbitol dehydrogenase (424 aas, ~44 kDa)

Appendix

128

C_570031

(fragment)

Zinc-containing alcohol dehydrogenase, KOG1 description:

sorbitol dehydrogenase

ACK1;

C_330070

acetate kinase; acetokinase (Martin H. Spalding); potential N-

terminal organelle targeting sequence (Laurens Mets)

(431 aas, ~45,5 kDa) ACK 2

ACK2;

C_170112

acetate kinase; acetokinase (Martin H. Spalding)

(408 aas, ~ 43 kDa)

PTA1;

C_870001

related to phosphate acetyltransferase (phosphotransacetylase)

(Sabeeha Merchant) (581 aas, ~ 60 kDa,) PTA 2

PTA2;

C_330071

related to phosphate acetyltransferase (phosphotransacetylase)

(Sabeeha Merchant) (792 aas, ~85 kDa)

PDC 1 C_340129 pyruvate decarboxylase [EC:4.1.1.1] (497 aas, ~53 kDa)

PFO 1 PFR1,

C_140055

pyruvate ferredoxin oxidoreductase (= NifJ) (Sabeeha Merchant)

(1303 aas, ~140 kDa)

C_180128 KOG1 description: Glycerol-3-phosphate dehydrogenase /

dihydroxyacetone 3-phosphate reductase (702 aas, ~75 kDa)2

C_180126 KOG1 description: glycerol-3-phosphate dehydrogenase /

dihydroxyacetone 3-phosphate reductase (670 aas, ~73 kDa)2

GPD1;

C_490040

Interconverts dihydroxyacetone phosphate and glycerol-3-

phosphate. Possibly involved in G-3-P production for glycerolipid

biosynthesis, by homology to an uncharacterized Arabidopsis

protein (Wayne R. Riekhof) (217 aas, ~23 kDa)


dihydroxyacetone 3-phosphate reductase (217 aas, ~23 kDa)


dihydroxyacetone 3-phosphate reductase (221 aas, ~23 kDa)

GPD 6

C_18990001;

fragment KOG1 description: glycerol-3-phosphate dehydrogenase

GPP 1 C_590009,

fragment

KOG1 description: predicted haloacid-halidohydrolase and related

hydrolases3

Tab 8-1: Detailed list of annotated sequences that were found in the C. reinhardtii genomic sequence on

JGI, which encode enzymes that are involved in one of the fermentative pathways of C. reinhardtii. PFL = pyruvate formate-lyase, AdhE = acetaldehyde / alcohol dehydrogenase and PFL-deactivase, ADH = Zn-containing alcohol dehydrogenase, ACK = acetate kinase, PTA = phosphotransacetylase, PDC = pyruvate decarboxylase, PFO = pyruvate ferredoxin/flavodoxin oxidoreductase, GPD = glycerol-3-phosphate dehydrogenase, GPP = glycerol-3-phosphate phosphatase. 1) KOG = clusters of euKaryotic Orthologous Groups; part of the Conserved Domain Database (CDD) (Marchler-Bauer et al., 2003). 2) C_180128 and C_180126 might have dual functions since, according to an NCBI Conserved Domain Search, they contain an N-terminal hydrolase- and a C-terminal GPD-domain. 3) Detected by blasting the Aspergillus fumigatus glycerol-3-phosphate phosphatase GppA (PubMed XP_752423) against the C. reinhardtii genome.

Curriculum vitae

129

9 Curriculum vitae

Personal Data

Name: Anja Christine Hemschemeier

Date of birth: June 29th, 1976

Place of birth: Engelskirchen

Citizenship: German

Marital status: single

School attendance

1982 - 1986 Basic primary school in Wegescheid, Gummersbach

1986 - 1996 Comprehensive secondary school; Gymnasium Grotenbach in

Gummersbach

1993 Stay abroad in Santiago de Chile, attendance at the Deutsche Schule

June 1996 Acquirement of the general qualification for university (Abitur), with

average grade 1,7

Professional training

1997 - 2001 Study of biology at the Rheinische Friedrich-Wilhelms-University in Bonn

June 2002 Acquirement of the diploma-grade of biology at the institute for botany of

the University in Bonn (grade 1,0 with commendation)

Title of the diploma thesis (translated): The H2-metabolism of

Chlamydomonas reinhardtii in the absence of sulphur: physiological and

molecular biological studies. Tutor: Prof. Dr. W. Hachtel

July 2002 Begin of the dissertation in the department for molecular biochemistry at

the University of Bonn. Tutor: PD Dr. T. Happe

Title of the thesis: “The anaerobic life of the photosynthetic alga

Chlamydomonas reinhardtii. Photofermentation and hydrogen production

upon sulphur deprivation.” Since October, 2002, this thesis is promoted by

the Studienstiftung des Deutschen Volkes.

July / August 2002 Stay abroad for research in the Food and Bioprocess Engineering Group at

the University of Wageningen (the Netherlands) (promoted by COST

Action 841).

October -

December 2002 Stay abroad for research in the Laboratoire d’Ecophysiologie de la

Photosynthése of Prof. Gilles Peltier in CEA Cadarache (France)

(promoted by the European Molecular Biology Organization, EMBO).

Curriculum vitae

130

April 2003 Change to the Ruhr-University Bochum, continuation of the PhD thesis in

the working group Photobiotechnology of Prof. Dr. T. Happe

May 2003 Affiliation into the European Graduate College 795 (Regulatory Circuits in

Cellular Systems: Fundamentals and Biotechnological Applications)

July 2004 Stay abroad for research in the Laboratoire d’Ecophysiologie de la

Photosynthése of Prof. Gilles Peltier in CEA Cadarache (France)

(promoted by the DAAD, Project Based Personnel Exchange Programme

with France - PROCOPE)

Publications

Happe T., Hemschemeier A., Winkler M. and Kaminski A. (2002) Hydrogenases in green algae: Do they save the algae´s life and solve our energy problems? Trends Plant Sci 7, 246-250

Winkler M., Hemschemeier A., Gotor C., Melis A. and Happe T. (2002) [Fe]-hydrogenases in green algae: Photo-fermentation and hydrogen evolution under sulfur deprivation. Int J Hydrogen Energy 27, 1431-1439

Winkler M., Maeurer C., Hemschemeier A. and Happe T. (2002) The isolation of green algal strains with outstanding H2-productivity. Int J Hydrogen Energy 27, 1431-1439

Hemschemeier A. and Happe T. (2005) The exceptional photofermentative hydrogen metabolism of the green alga Chlamydomonas reinhardtii. Biochem Soc Trans. 33:39-41 Fouchard S., Hemschemeier A., Caruana A., Pruvost J., Legrand J., Happe T., Peltier G. and Cournac L. (2005) Autotrophic and mixotrophic hydrogen photoproduction in sulfur-deprived Chlamydomonas cells. Appl Environ Microbiol, in press

Contributions to conferences

Schwarzer S., Kaminski A., Florin L., Hemschemeier A., Winkler M., Heil B. and Happe T. (2001) Hydrogen production in green algae – A novel type of iron hydrogenase is linked to the photosynthetic electron transport chain. Workshop of Biodiversity of Hydrogenases. Reading, UK. Poster

Schwarzer S., Hemschemeier A., Weber A., Winkler M., Heil B. and Happe T. (2001) Light driven hydrogen production by microalgae. Working shop and expert meeting of Photobiological Hydrogen

Production. Szeged, Hungary. Poster

Hemschemeier A. and Winkler M. (2001) Iron hydrogenases in unicellular green algae: structure and physiology. NEDO Start-up meeting. Bonn, Germany. Oral presentation Hemschemeier A., Weber A., Radix M., Smolny S., Winkler M. and Happe T. (2002) Hydrogen production by green algae in the absence of sulphur: a specific response or just a consequence of anaerobiosis? Biohydrogen 2002. Ede, The Netherlands. Poster and oral presentation

Hemschemeier A., Winkler, M., Weber, A., Radix M., Kaminski, A. and Happe T. (2002) Hydrogen metabolism in green algae - New insights coming from the genes. The 10

th International Conference on the

Cell and Molecular Biology of Chlamydomonas. Vancouver, Canada. Poster

Hemschemeier A., Weber, A., Radix M. and Happe T. (2002) Structure and function of a new type of [Fe]-hydrogenase in green algae. International meeting on Iron-sulphur proteins: Biogenesis, Structure, Function,

Pathogenesis and Evolution. Marburg, Germany. Poster

Curriculum vitae

131

Hemschemeier A., Kästle, S. Winkler, M. and Happe T. (2002) What happens with Chlamydomonas

reinhardtii upon sulphur-deprivation? Workshop COST 841, Biosynthesis and Regulation of Hydrogenases.

Madrid, Spain. Poster

Happe T., Mäurer, C. and Hemschemeier A. (2002) Identification of Chlamydomonas reinhardtii mutants for hydrogen production. NEDO Meeting: Development of molecular devices for hydrogen production.

Kyoto, Japan. Poster

Hemschemeier A., Winkler M., Weber A., Stirnberg M. and Happe T. (2003) S-deprivation induces a photofermentative metabolism in the unicellular green alga Chlamydomonas reinhardtii. Working Group 2

Workshop „Active Centres of Hydrogenases“. Mülheim/Ruhr, Germany. Oral presentation Hemschemeier A., Weber A., Stirnberg M. and Happe T. (2004) New insights into the photofermentative hydrogen metabolism of Chlamydomonas reinhardtii. 11th International Conference on the Cell and

Molecular Biology of Chlamydomonas. Kobe, Japan. Poster Hemschemeier A., Jacobs J. and Happe T. (2005) [Fe]-hydrogenases of green algae: powerful enzymes in (photobio)technological hydrogen production. SolarH workshop (EU-Projekt 516510). Paris, France. Oral

presentation Hemschemeier A., Winkler M., Weber A., von Abendroth G., Müllner K. and Happe T. (2005) [Fe]-hydrogenases of green algae: powerful enzymes in photobiological or semiartificial hydrogen production. International Hydrogen and Energy Congress & Exhibition. Istanbul, Turkey. Oral presentation

Danksagungen

Diese Dissertation begann ich im Juli 2002 an der Rheinischen Friedrich-Wilhelms-Universität in Bonn und schließe sie nun, im August 2005, an der Ruhr-Universtät Bochum ab. Während dieser Zeit sind mir natürlich viele liebe Menschen begegnet, die mich auf die eine oder andere Weise bei meiner Arbeit unterstützt haben, und denen ich hiermit danken möchte. Meinem Betreuer, Prof. Dr. Thomas Happe möchte ich aus vielen Gründen danken. Zum einen hat er die Betreuung dieser Arbeit übernommen und mir ein faszinierendes Forschungsthema überlassen. Dabei durfte ich frei schalten und walten, aber immer im genau richtigen Augenblick brachte mein Chef die ein oder andere richtungweisende Idee ein. Ich bin außerdem stolz darauf, eine der Pionierinnen der AG Photobiotechnologie sein zu können. Herr Happe hat schon meine allerersten Schritte in die Welt der Forschung betreut und begleitet und mir mit sanftem Druck immer wieder Aufgaben anvertraut, die mich zunächst erschreckten, die mich aber stets ein kleines Stückchen haben wachsen lassen. Bei Herrn Prof. Dr. Franz Narberhaus vom Lehrstuhl für Biologie der Mikoorganismen möchte ich mich sehr herzlich für die bereitwillige Übernahme des Zweitgutachtens bedanken. Ich danke Prof. Dr. Matthias Rögner dafür, dass ich die vielfältige technische Ausstattung seines Lehrstuhls nutzen durfte, und für das harmonische Arbeitsklima. Ich danke ihm ebenfalls als dem Sprecher des Europäischen Graduiertenkollegs 795, in das ich als Kollegiatin aufgenommen wurde. Die Teilnahme an diesem Kolleg hat mir nicht nur wertvolle wissenschaftliche Erkenntnisse, sondern auch das Kennenlernen vieler anderer netter ForscherInnen ermöglicht. In diesem Zusammenhang möchte ich auch Dr. Petra Schrey und Dr. Christiane Wüllner danken, die als Koordinatorinnen des EGC immer ein offenes Ohr für Fragen und Probleme hatten. Petra Schrey möchte ich auch in ihrer neuen Funktion im Dekanat danken. Es gibt nichts, wobei sie einem nicht helfen kann. Ebenso herzlich danken möchte ich Skadi Heinzelmann, die ich in ihrer Funktion als Promotions-Beraterin besonders in den letzten Wochen mit Fragen gelöchert habe. Die Antwort kam immer prompt, freundlich und mit sofortigem Hilfsfaktor. Einen beträchtlichen Teil meiner Arbeit habe ich in Kooperation mit Dr. Laurent Cournac und Prof. Dr. Gilles Peltier vom Laboratoire d’Ecophysiologie de la Photosynthése im CEA Cadarache (Frankreich) anfertigen können. Das dortige technische Equipment sowie die unerschöpfliche Diskutierbereitschaft von Laurent (und vielleicht ebenso die faszinierende Landschaft der Provence) haben mich große Schritte vorangebracht. Laurent möchte ich außerdem besonders dafür danken, dass er mich zwei Wochen lang ins Haus seiner Familie aufgenommen hat. Einen weiteren, nicht minder bedeutsamen Teil meiner Arbeit habe ich nur mit tatkräftiger Unterstützung von Dr. Gary Sawers vom Department of Molecular Microbiology, John Innes Centre, Norwich, Großbritannien, anfertigen können. Er hat mir E. coli-Stämme, Antikörper und tausende von Antworten geschenkt. Besonders gefreut hat mich, dass er sich ohne zu zögern bereiterklärte, einige Teile meiner englischsprachigen Arbeit in englischwürdiges Englisch zu korrigieren. Daher gebührt ihm ein ganz besonders herzliches Dankeschön!

Dr. Marcel Janssen und Prof. Dr. René Wijffels von der Food and Bioprocess Engineering Group, Department of Agrotechnology and Food Sciences, Wageningen University, Niederlande, danke ich für vier Wochen gemeinsames Forschen an Photobioreaktoren. Auch sehr herzlich zu danken habe ich Dr. Wolfgang Schiefer, der schon in Bonn ein stets hilfsbereiter Kollege war, mit dem ich immer so schön lachen konnte (auch wenn er immer RNasen auf meinem Platz verteilt hat). Hier in Bochum hat er mich in die hohe Kunst der Fluoreszenzmessungen eingeführt, und mir geduldig immer wieder diese seltsamen Apparaturen erklärt. Der gesamten Arbeitsgruppe Happe schulde ich großen Dank für all die kleinen und großen Hilfeleistungen, für schöne Feiern, nette Gespräche und ein insgesamt angenehmes Arbeitsklima. Besonderer Dank gebührt Astrid Weber als unserer „Oberschwester“, und Martin Winkler, mit dem man immer wissenschaftliche oder private Diskussionen führen kann. Dr. Kathrin Happe, die nun leider nicht mehr bei uns arbeitet, danke ich für die schöne und harmonische Zeit unserer Zusammenarbeit. Brigitte Depka, die erst vor kurzem zu uns gekommen ist und sich erst an diesen bunten Haufen gewöhnen musste, danke ich ganz besonders, denn sie hat eine ganze Menge wunderschöner Western Blots für mich fabriziert. Auch der gesamten Lehrstuhlbelegschaft möchte ich danken für die nette Aufnahme der Bonner Exilanten. Dabei ist besonders Gertrud Lideka zu nennen, die uns Neulingen immer mit Rat und Tat zur Seite steht. Ganz besonders herzlich möchte ich Thomas Schröder danken. Ohne sein technisches Geschick wäre so manche Forschung nicht möglich gewesen. Aber noch viel mehr danke ich ihm für die geistige Unterstützung, die immer wieder arbeitsbedingte Wut- und Frustausbrüche aufgefangen hat, und für den häufigen Lachmuskelkater. Für die Förderung meiner Arbeit habe ich der Studienstiftung des deutschen Volkes besonderen Dank auszusprechen. Jedoch ging ihre Förderung meiner Dissertation über das rein Finanzielle hinaus. Die gemeinsamen Treffen mit anderen Studienstiftlern und meinem Vertrauensdozenten Prof. Dr. Franz Lebsanft haben mir Einblicke in ganz andere Forschungsgebiete und Denkweisen ermöglicht. Außerdem fühlte ich mich durch stete Korrespondenz und den Zutritt zum Intranet in einer großen Gemeinschaft aufgehoben. Herrn Lebsanft danke ich besonders als meinem Vertrauensdozenten. Ich möchte mich auch recht herzlich bei der European Molecular Biology Organization

(EMBO), der COST Action 841 und dem deutschen akademischen Austauschdienst (DAAD)

für die Förderung meiner Auslandsaufenthalte, sowie dem Frauenförderungsprogramm der Fakultät für Biologie der Ruhr-Universität Bochum für die großzügigen Reisekostenzuschüsse für Konferenzen in Japan und der Türkei bedanken. Last, aber sicher nicht least, danke ich über alles meinen Eltern und meiner Schwester, die schon so einiges mit mir mitgemacht haben und immer für mich da waren. Sie haben das ständige Auf und Ab der Forschung sowie das in den letzten Wochen konzentrierte Gejammere stoisch über sich ergehen lassen und waren immer mit aufmunternden Worten parat. Ich danke Euch so sehr für alles!

diss total korrigiert - mit - massachusetts institute of...

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