Regulation of glutamate dehydrogenase in Corynebacterium ......2.1 Corynebacterium glutamicum During...

Regulation of glutamate dehydrogenase in Corynebacterium glutamicum

Der Naturwissenschaftlichen Fakultät

der Friedrich-Alexander-Universität Erlangen-Nürnberg

zur

Erlangung des Doktorgrades

vorgelegt von

Eva Hänßler

aus Aachen

Als Dissertation genehmigt von der Naturwissen-

schaftlichen Fakultät der Universität Erlangen-Nürnberg

Tag der mündlichen Prüfung: 15.02.2008

Vorsitzender der Promotionskommission: Prof. Dr. Eberhard Bänsch

Erstberichterstatter: Prof. Dr. Andreas Burkovski

Zweitberichterstatter: Prof. Dr. Reinhard Krämer

Content

Content

1 Zusammenfassung/Summary........................................................1

2 Introduction.....................................................................................3

2.1 Corynebacterium glutamicum.............................................................................3 2.2 Uptake and assimilation of nitrogen sources....................................................4 2.3 Nitrogen-dependent regulation...........................................................................6 2.4 GDH at the interface between nitrogen and carbon metabolism...................11

2.4.1 Glutamate dehydrogenase of E. coli ................................................................12 2.4.2 Glutamate dehydrogenase of B. subtilis...........................................................13 2.4.3 Glutamate dehydrogenase of C. glutamicum ...................................................15

2.5 Objectives ...........................................................................................................17

3 Materials and methods .................................................................19

3.1 Bacterial strains and plasmids .........................................................................19 3.2 Cultivation of bacteria........................................................................................22

3.2.1 Culture medium for E. coli ................................................................................22 3.2.2 Culture media for corynebacteria .....................................................................22 3.2.3 Antibiotics .........................................................................................................23 3.2.4 Growth conditions.............................................................................................24

3.3 Genetic manipulation of bacteria......................................................................25 3.3.1 Preparation of competent E. coli cells and transformation ...............................25 3.3.2 Preparation of competent C. glutamicum cells and transformation..................26

3.4 Working with DNA ..............................................................................................26 3.4.1 Isolation of plasmid DNA from E. coli ...............................................................26 3.4.2 Gel electrophoresis and extraction of DNA from agarose gels.........................27 3.4.3 Preparation of chromosomal DNA from C. glutamicum....................................27 3.4.4 Purification and enrichment of DNA .................................................................27 3.4.5 Polymerase chain reaction (PCR) ....................................................................28 3.4.6 Two-step PCR ..................................................................................................28 3.4.7 Restriction of DNA............................................................................................29 3.4.8 Ligation of DNA fragments ...............................................................................29 3.4.9 Sequencing of DNA..........................................................................................29

3.5 Working with RNA ..............................................................................................30 3.5.1 Isolation of total RNA and RNA gel electrophoresis .........................................30

Content

3.5.2 Synthesis of digoxigenin-labeled RNA probes .................................................31 3.5.3 Northern blot analysis.......................................................................................31 3.5.4 Dot blot analysis ...............................................................................................33 3.5.5 Reverse transcriptase (RT) PCR......................................................................34 3.5.6 Quantitative real time RT PCR .........................................................................34 3.5.7 Primer extension analysis ................................................................................35

3.6 Working with proteins........................................................................................37 3.6.1 Protein purification............................................................................................37 3.6.2 Quantification of proteins..................................................................................38 3.6.3 SDS polyacrylamide gel electrophoresis (PAGE).............................................38 3.6.4 Staining with Coomassie Brilliant Blue .............................................................39 3.6.5 Western blotting ...............................................................................................39 3.6.6 Determination of enzyme activity .....................................................................41

3.6.6.1 GDH activity measurements.....................................................................41 3.6.6.2 Glutamyltransferase test...........................................................................42

3.6.7 Determination of promoter activity....................................................................43 3.6.8 Gel shift assays and competition assays .........................................................43

4 Results ...........................................................................................45

4.1 Purification and characterization of glutamate dehydrogenase....................45 4.1.1 Purification of GDH...........................................................................................45 4.1.2 Characterization of GDH in lysine-producing strains........................................47 4.1.3 Gradual expression of gdh ...............................................................................49

4.2 Transcriptional regulation of gdh .....................................................................51 4.2.1 Mutational analyses of the gdh promoter region ..............................................51 4.2.2 Determination of the transcription start.............................................................57 4.2.3 Nitrogen-dependent transcription .....................................................................60

4.2.3.1 Function of AmtR......................................................................................61 4.2.3.2 Influence of putative regulators on gdh transcription................................66

4.2.4 Studies on sigma factor-dependent gdh expression ........................................71 4.2.5 Investigation of the putative orf Cg2281 ...........................................................73

4.3 Identification of AmtR and FarR target genes .................................................75 4.3.1 Identification of FarR target genes ...................................................................75

4.3.1.1 Characterization of arginine biosynthesis genes ......................................76 4.3.1.2 Determination of FarR and ArgR binding sites .........................................79 4.3.1.3 Transcriptional regulation of arginine biosynthesis genes........................81

Content

4.3.2 Identification of AmtR target genes ..................................................................83

5 Discussion.....................................................................................89

5.1 Characterization of GDH in the context of systems biology..........................89 5.2 Transcriptional regulation of gdh .....................................................................93 5.3 Identification of FarR and AmtR target genes ...............................................100 5.4 The interface between nitrogen and carbon metabolism.............................106

6 Appendix......................................................................................109

6.1 Regulation of glutamine synthetase in corynebacteria................................109 6.2 Plasmid constructions.....................................................................................112

References ........................................................................................117

Publications ......................................................................................131

Abbreviations and units...................................................................132

Zusammenfassung 1

1 Zusammenfassung

Die Glutamatdehydrogenase (GDH) aus Corynebacterium glutamicum, einem

Actinomyceten mit herausragender biotechnologischer Bedeutung, befindet sich an einer

wichtigen Position innerhalb des Stoffwechsels, da sie Stickstoffassimilation und den

Zentralstoffwechsel verbindet. Unter Überschussbedingungen ist die GDH an der

Ammoniumassimilation beteiligt und über das Substrat α-Ketoglutarat besteht eine direkte

Verknüpfung zum Citrat Zyklus. Aufgrund der NADPH-Abhängigkeit kann weiterhin der pool

an Reduktionsäquivalenten beeinflusst werden. Trotz dieser scheinbar bedeutsamen

Stellung konnte bis auf die mögliche Beteiligung zweier Transkriptionsregulatoren kein

detaillierter Regulationsmechanismus beschrieben werden.

Um die GDH genauer charakterisieren zu können, wurde das Protein gereinigt und in

definierten Lysinproduktionsstämmen untersucht. Hierzu wurde im Rahmen dieser Arbeit

ein Protokoll zur Überexpression in C. glutamicum, gefolgt von einer Kombination aus Ni2+

NTA Affinitätschromatographie und Gelfiltrationschromatographie etabliert. Dadurch wurde

die Grundlage für weiterführende kinetische Messungen gelegt. Weiterhin wurde GDH-

Aktivität, Proteinmenge und Transkriptlevel in Produktionsstämmen bestimmt.

Veränderungen von Stoffwechselflüssen, die auf eine erhöhte Lysinproduktion

zurückzuführen waren, zeigten keinen Einfluss auf die GDH im Vergleich zum Wildtyp.

Als Schwerpunkt der Arbeit wurde, auf Grund der widersprüchlichen und geringen

Informationen, die gdh Transkription näher betrachtet sowie die Regulatoren AmtR und

FarR hinsichtlich neuer Targetgene untersucht. Mit diesem Ansatz konnte durch

Mutagenese die putative -10-Region des gdh Promotors experimentell bestätigt und

Promotoren mit abgestufter Aktivität konstruiert werden. Im Rahmen dieser

Untersuchungen wurden zusätzlich zu dem bekannten Promoter ein zweiter Promotor

sowie ein bisher nicht bekannter Transkriptionsstartpunkt identifiziert. Es wurde erstmalig in

vivo nachgewiesen, dass die gdh Transkription unter Stickstoffmangel von beiden

Promotoren zunimmt und dass dieser Mechanismus AmtR-abhängig ist. Weitere putative

Regulatoren, OxyR und FarR, zeigten keine Effekte, so dass zusätzlich mit

Untersuchungen zur Regulation durch alternative Sigmafaktoren begonnen wurde.

Mit mez (kodiert für das Malat Enzym) und dapD (kodiert für die

Tetrahydrodipicolinatsuccinylase) wurden zwei bisher nicht bekannte AmtR Targetgene

inklusive Bindestellen identifiziert. Für FarR, einen weitern putativen Regulator der gdh

Transkription, wurde eine mögliche Beteiligung an der Regulation der Argininbiosynthese

nachgewiesen.

Summary 2

1 Summary

Glutamate dehydrogenase (GDH) of the industrially highly relevant actinomycete

Corynebacterium glutamicum is located at an important branch-point of metabolism. On the

one hand, it is the enzyme primarily involved in ammonium assimilation under ammonium

surplus. On the other hand, it is connected to the tricarboxylic acid cycle by its substrate 2-

oxoglutarate and influences the intracellular pool of reductive equivalents due to its NADPH

dependency. Despite this crucial position and intense studies, only an incomplete model of

regulation has been proposed including two putative transcriptional regulators.

To characterize the GDH enzyme, within this work studies were performed that included

purification of GDH and investigation of GDH in lysine-producing strains. It was possible to

establish a protocol for overexpression of GDH in C. glutamicum followed by purification by

Ni2+ NTA affinity chromatography and size exclusion chromatography, so that the

foundation for kinetic measurements has been laid. Examination of GDH activity, protein

level, and gdh transcript in lysine-producing strains revealed that altered metabolic fluxes

due to the enhanced production did not lead to changes compared to the wild type.

However, because of contrary results reported and an apparent lack of information, the

main focus was put on the reinvestigation of transcriptional control of gdh, which included

characterization and mutagenesis of the gdh promoter as well as further examination of the

two regulators AmtR and FarR. The predicted -10 region of the gdh promoter was verified

by experimental approaches. In the course of these studies, promoters varying in activity

were constructed. Besides the known promoter, it was possible to identify an additional

promoter including a so far not determined transcriptional start site. Transcription of gdh

was shown to be induced upon nitrogen deficiency from both promoters and for the first

time, AmtR-mediated regulation could be demonstrated in vivo. Studies on other putative

regulators of gdh transcription, OxyR and FarR, did not show any effects so that the focus

was put on transcription by alternative sigma factors as well.

Two novel AmtR target genes, namely, mez encoding malic enzyme and dapD encoding

tetrahydrodipicolinate succinylase, were additionally identified including respective binding

sites. Investigations on the second putative regulator of gdh transcription, FarR, suggested

a role in regulation of arginine biosynthesis.

Introduction 3

2 Introduction

2.1 Corynebacterium glutamicum

During a screening program for glutamate-producing microorganisms, the Gram-positive

bacterium Corynebacterium glutamicum was isolated from soil samples taken at the Ueno

Zoo in Tokyo, Japan (Kinoshita et al., 1957). It is an aerobic, non-sporulating, and immobile

bacterium marked by a high G+C content of the DNA. The characteristic rod-shape (koryne,

Greek word for rod) is eponymous for corynebacteria. Another feature of this genus is the

so-called snapping cell division, during which cells are laterally connected prior to the actual

division (figure 2.1). Due to the complex composition of the cell wall, corynebacteria are

referred to as mycolic acids-containing actinomycetes (Stackebrandt et al., 1997). Besides

nocardia, well-known human pathogens such as Corynebacterium diphtheriae,

Mycobacterium leprae, and Mycobacterium tuberculosis are members of this group

(Pascual et al., 1995). However, C. glutamicum is non-pathogenic and therefore save to

handle. Because of these properties and the availability of a broad range of experimental

techniques, C. glutamicum is suitable as model organism for its pathogenic relatives.

The identification of C. glutamicum as a

glutamate-excreting bacterium by Kinoshita

and coworkers (1957) laid the foundation for

an extended biotechnological application of

this organism. Under conditions of biotin

limitation, treatment with antibiotics or certain

surfactants, C. glutamicum is able to

accumulate high amounts of glutamate in the

surrounding medium (Gutmann et al., 1992).

During the last decades, improved

production strains were created by random

mutagenesis programs, and within the last

years rational design approaches were applied (Sahm et al., 1996; Ohnishi et al., 2002).

Upscaling and optimization of cultivation conditions as well as the use of heat-tolerant

species such as Corynebacterium efficiens contributed to an increase in yield combined

with a reduction of production costs (Fudou et al., 2002; Hermann, 2003). So, today

C. glutamicum is one of the most important organisms used in biotechnological production

processes. This biotechnological relevance of the organism is reflected in recent numbers

on amino acid production by various C. glutamicum strains. Of the flavor enhancer L-

Fig. 2.1: Scanning-electromicrogram of C. glutamicum cells. Snapping cell division as well as the characteristic V-shape of the cells is visible (Forschungszentrum Jülich).

Introduction 4

glutamate 1.5 million tons per year and of the feed additive L-lysine 650,000 tons per year

are produced. In addition to that, other amino acids such as L- alanine, L-isoleucine, and L-

proline as well as vitamins and nucleotides are yielded by fermentation with C. glutamicum

strains (Leuchtenberger et al., 2005).

This rapid increase in industrial importance led to an intense study of this organism.

Whereas first approaches mainly focused on the respective pathways of amino acid

biosynthesis as well as on import and export systems, today more emphasis is put on

global approaches. While the central metabolism has also been studied in detail already,

the publication of the full genome sequence by two different industry-funded groups (Ikeda

& Nakagawa, 2003; Kalinowski et al., 2003) enables a closer look on interacting metabolic

pathways and connected regulatory mechanisms. Besides the characterization of enzymes

on the biochemical and genetic level, currently techniques are available that allow global

investigation of transcriptome (Wendisch, 2003), proteome (Schaffer and Burkovski, 2005),

and metabolome (Strelkov et al., 2004). Within the last decade especially nitrogen

metabolism has been well investigated, leading to new insights into regulatory processes

and signal transduction in response to the environment (for overview, Burkovski, 2003a;

2003b; 2005; 2007). This approach has recently even been extended to related

actinomycetes with published genome sequence, namely, C. diphtheriae, the causative

agent of diphtheria (Cerdeno-Tarraga et al., 2003), Corynebacterium jeikeium (Tauch et al.,

2005) a pathogen found in the human skin flora, and the heat-tolerant amino acid producer

C. efficiens (Fudou et al., 2002). Currently, nitrogen metabolism is in the focus of research

in connection with finding a common theme among related organisms (Walter et al., 2007).

2.2 Uptake and assimilation of nitrogen sources

Nitrogen is an essential macro-element, which is part of important components of the cell

such as nucleotides and amino acids or amino sugars within the bacterial murein sacculus.

In general, bacteria can use a variety of different organic and inorganic nitrogen sources

depending on their genetic repertoire. As a first step of utilization, compounds need to be

taken up by the cell. This can either occur via passive diffusion along a gradient of

concentration or by active transport processes. In C. glutamicum, transport systems and

respective assimilatory enzymes have been characterized biochemically and on the genetic

level for ammonium, creatinine, glutamate, and urea (Börmann et al., 1992; Kronemeyer et

al., 1995; Siewe et al., 1996; Jakoby et al., 1997; Nolden et al., 2000; Beckers et al., 2001;

Meier-Wagner et al., 2001; Nolden et al., 2001a; Schulz et al., 2001; Beckers et al., 2004;

Introduction 5

Bendt et al., 2004). Furthermore, organic nitrogen sources such as alanine, asparagine,

glutamine, serine, threonine, and different peptides can be used, even though uptake and

assimilation in respect of nitrogen supply are less investigated (Burkovski, 2005).

Ammonium serves as an excellent nitrogen source for C. glutamicum. The protonated

ammonium ion (NH4+) is in equilibrium with uncharged ammonia (NH3), which is able to

pass through the cell membrane by diffusion. Under ammonium surplus, as for instance

found in standard minimal medium, diffusion of ammonia is sufficient to promote cell

growth. As soon as passive diffusion of ammonia becomes limiting resulting in a deprivation

of nitrogen, two ammonium transport systems, AmtA and AmtB, are recruited for uptake.

AmtA has been characterized as a high affinity uptake system, which works in a membrane

potential-dependent manner (Siewe et al., 1996; Meier-Wagner et al., 2001; Walter, 2007).

Since AmtA also accepts the radioactively labeled substrate analog [14C]methylammonium,

determination of kinetic parameters such as KM (44.7 µM for methylammonium) and

maximal velocity (20±5 nmol min-1(mgdw)-1) was possible (Siewe et al., 1996; Meier-Wagner

et al., 2001). More recently, it was demonstrated that AmtA-mediated uptake is independent

of assimilatory enzymes excluding the mechanism of so-called metabolic trapping.

Furthermore, ammonium is accumulated by AmtA in a membrane potential-dependent

manner (Walter, 2007). This underlines previous results describing AmtA as an active

(methyl)ammonium carrier. The second transport system, AmtB, seems to be a low affinity

uptake system, which has channel-like properties and facilitates diffusion of ammonia

(Walter, 2007).

Once inside the cell, ammonium can be assimilated via two different pathways yielding

glutamate and glutamine, the major donors of intracellular nitrogen (Merrick & Edwards,

1995) (figure 2.2). Glutamate is the donor of nitrogen in transamination reactions, whereas

glutamine donates nitrogen for the synthesis of nucleotides, arginine, asparagine, histidine,

tryptophane, glucosamine, and p-aminobenzonate (Reitzer, 2003).

When present in high concentrations, ammonium is primarily fixed by glutamate

dehydrogenase (GDH, encoded by gdh; Börmann et al., 1992). This low affinity enzyme

catalyzes the reductive amination of 2-oxoglutarate to L-glutamate in an NADPH-dependent

reaction. Because of the low affinity to ammonium (KM=3.08 mM), at concentrations below

5 mM a second pathway for assimilation is mainly used (Shiio & Ozaki, 1970; Tesch et al.,

1998). This alternative pathway consists of two reactions catalyzed by glutamate synthase

(GOGAT, encoded by gltBD; Beckers et al., 2001; Schulz et al., 2001) and glutamine

synthetase (GS, encoded by glnA; Jakoby et al., 1997). First, ammonium is assimilated by

GS, which has a high substrate affinity compared to GDH. Under conditions of nitrogen

Introduction 6

2-oxoglutarate + NH4+ + NADPH+H+ GDH L-glutamate + NADP+

L-glutamate + NH4+ + ATP GS L-glutamine + ADP

L-glutamine + 2-oxoglutarate + NADPH+H+ GOGAT 2 L-glutamate + NADP+

A: Nitrogen surplus

B: Nitrogen deprivation




A: Nitrogen surplus





A: Nitrogen surplus


surplus, 28 % of ammonium is assimilated by GS to fulfill the cellular requirement for

glutamine (Tesch et al., 1999). This is due to regulation of GS on the level of enzyme

activity by an adenylyltransferase (ATase, encoded by glnE; Nolden et al., 2001a). When

most of ammonium can be assimilated by GDH, GS is present in its less active,

adenylylated form. At decreasing ammonium concentrations, GS is deadenylylated by

ATase, which results in an increased activity (Jakoby et al., 1999; Nolden et al., 2001a). GS

attaches ammonium in an ATP-dependent reaction to glutamate forming glutamine. In the

second reaction, the amide nitrogen is transferred to 2-oxoglutarate in an NADPH-

dependent reaction by GOGAT yielding glutamate (figure 2.2). Despite a higher energy

demand, this is the preferred way of ammonium assimilation under limiting conditions.

Fig. 2.2: Assimilation of ammonium in C. glutamicum. Under ammonium surplus (A) assimilation occurs mainly via the glutamate dehydrogenase (GDH) pathway, while under nitrogen deprivation ATP-dependent glutamine synthetase (GS) and glutamate synthase (GOGAT) are responsible (Tesch et al., 1999).

2.3 Nitrogen-dependent regulation

In order to keep a balance between sufficient nitrogen supply and energy metabolism as

well as to coordinate the use of alternative nitrogen sources, elaborate regulatory

mechanisms need to be present in the bacterial cell. These regulatory networks, which are

responsible for the arrangement of transport and assimilation of nitrogen sources, are

summarized in the term nitrogen control. This term describes sophisticated mechanisms of

Introduction 7

transcriptional regulation as well as fine-tuning of enzyme activity by post-translational

modification.

Intense studies on nitrogen control in C. glutamicum led to the identification of the major

players involved within the respective regulatory network (for overview, Burkovski, 2003a;

2003b; 2005; 2007). In many bacteria, PII type signal transduction proteins play a crucial

role in sensing the current metabolic state and then transmitting the signal. In contrast to

other organisms, only one PII protein, designated GlnK, is present in C. glutamicum

(Nolden et al., 2001b; Ninfa & Jiang, 2005). The glnK gene is organized in an operon with

amtB, encoding an ammonium uptake system, and glnD as it has been described for other

actinomycetes (for review, Arcondéguy et al., 2001; Nolden et al., 2001b). The glnD gene

product, a bifunctional adenylyltransferase, and the AmtR protein are essential components

of nitrogen control (Jakoby et al., 2000; Strösser et al., 2004). AmtR was identified as global

regulator of nitrogen metabolism. Applying a set of bioinformatic tools, transcriptome and

proteome analyses as well as DNA binding studies, the AmtR regulon was closer

investigated. Based on previously identified AmtR binding sites, it was possible to define a

consensus sequence of an AmtR binding motif, which is displayed in figure 2.3 (Beckers et

al., 2005).

Fig. 2.3: Consensus motif of AmtR binding sites. The height of the letters represents the frequency of the respective nucleotide at a particular position (Beckers et al., 2005).

By combination of different global approaches, it was revealed that under conditions of

nitrogen surplus AmtR represses the transcription of at least 36 genes underlining its

function as a global regulator of nitrogen metabolism (Beckers et al., 2005). Genes

controlled by AmtR can be grouped into different categories according to their function. A

broad number of these genes encode transport systems for alternative nitrogen sources.

Besides the genes of the two ammonium transport systems AmtA (amtA, Jakoby et al.,

2000) and AmtB (amtB, Meier-Wagner et al., 2001), genes encoding transport systems for

glutamate (gluABCD, Kronemeyer et al., 1995), urea (urtABCDE, Beckers et al., 2004),

creatinine (crnT, Bendt et al., 2004), and a putative uptake system for oligopeptides

Introduction 8

(NCgl1915-1918, Beckers et al., 2005) are regulated. Even though it was not possible to

prove AmtR binding to the respective upstream region, the gene encoding a putative

protocatechuate transport system (vanK, Merkens et al., 2005) is also part of the AmtR

regulon.

Furthermore, expression of assimilatory enzymes for the utilization of different nitrogen

sources is controlled by AmtR. These AmtR-regulated genes encode enzymes involved in

ammonium assimilation at low concentrations (GS and GOGAT), urease (ureABCEGD,

Nolden et al., 2000; Beckers et al., 2004) catalyzing the formation of ammonium and carbon

dioxide from urea, and creatinine deaminase (codA, Bendt et al., 2004), which is

responsible for the degradation of creatinine leading to the dead-end product

methylhydantoin and ammonium. Interestingly, genes being part of signal transduction

cascade (glnD and glnK, Jakoby et al., 1999; Nolden et al., 2001b), are also among the

AmtR-regulated genes. Additionally, vanillate demethylase (vanAB, Merkens et al., 2005)

and genes of miscellaneous or unknown function were identified as part of nitrogen control

(Beckers et al., 2005). An overview of AmtR-regulated genes is given in figure 2.4.

Introduction 9

Fig. 2.4: Nitrogen control network and AmtR regulon of C. glutamicum. Under ammonium surplus, AmtR represses the transcription of genes encoding transport proteins, assimilatory enzymes, and components of the regulatory network. At decreasing ammonium concentrations, repression is released due to interaction of AmtR with the modified signal transduction protein GlnK. Modification/demodification is carried out by the adenylyltransferase GlnD in response to changes in ammonium availability. A possible sensor of the nitrogen status remains to be identified (Müller, 2005; Beckers et al., 2005). AmtR belongs to the family of TetR/ArcR-type regulators, which are the transcription

regulators occurring with the highest frequency in corynebacteria (Jakoby et al., 2000;

Brune et al., 2005). Typically, TetR-type regulators exhibit DNA binding depending on the

presence of small effector molecules as it has been described for the prototype of these

proteins TetR. Upon addition of Mg2+ and tetracycline, TetR is released from the DNA due

to conformational changes (Ramos et al., 2005). However, this mechanism does not apply

to AmtR. Instead, DNA binding of AmtR is regulated by protein-protein interaction. Under

nitrogen surplus, transcription of genes belonging to the AmtR regulon is repressed. At

decreasing concentrations, AmtR is released from its target DNA as a result of interaction

extracellular intracellular

NH4+

NH4+

glutamate

urea

NCgl1915-1918peptides

creatinine

ureaseurea

creatinine methylhydantoinCodA NH4+ NCgl

1099NCgl1100

NCgl1362

GlnD

AmtR

glutamineglutamate

NH4+

2-oxoglutarateglutamate

GS

GOGAT

GlnK

: transcription repression by AmtR

: metabolism and transport

: protein interaction

GluABCD

AmtA

AmtB

UrtABCDE

CrnT

H2CO3 + NH4+

: transport system

: enzyme

: signal transduction

: protein of unknown function

ATase

extracellular intracellular

NH4+

NH4+

glutamate

urea

NCgl1915-1918peptides

creatinine

ureaseurea

creatinine methylhydantoinCodA NH4+ NCgl

1099NCgl1100

NCgl1362

GlnD

AmtR

glutamineglutamate

NH4+

2-oxoglutarateglutamate

GS

GOGAT

GlnK

: transcription repression by AmtR

: metabolism and transport

: protein interaction

: transcription repression by AmtR: transcription repression by AmtR

: metabolism and transport: metabolism and transport

: protein interaction: protein interaction

GluABCD

AmtA

AmtB

UrtABCDE

CrnT

H2CO3 + NH4+

: transport system

: enzyme



: transport system

: enzyme



ATaseATase

Introduction 10

with the trimeric PII type protein GlnK. As a prerequisite for this, GlnK needs to be modified

at a specific tyrosyl residue at position 51 within the T-loop of the protein (Nolden et al.,

2001b; Beckers et al., 2005). This modification is performed by bifunctional adenylyl-

transferase GlnD (Stösser et al., 2004). Depending on the nitrogen status of the cell, three

AMP residues are attached to the PII protein, which then is able to interact with AmtR

(Beckers et al., 2005). Upon sudden increase in nitrogen supply, GlnD is also involved in

demodification of the signal transduction protein. Unmodified GlnK is not able to bind AmtR,

so that gene expression is repressed again. Furthermore, under these conditions GlnK is

sequestered to the membrane, interacts with the ammonium transporter AmtB and is

rapidly degraded (Strösser et al., 2004). Besides the regulation of gene expression, post-

translational modification is also a part of nitrogen control in C. glutamicum. GS is regulated

on the level of activity by adenylylation in response to the nitrogen status. This modification

carried out by GlnE seems to work independent of the GlnD/GlnK/AmtR cascade

(Burkovski, 2003b).

Whereas in Gram-negative bacteria such as Escherichia coli, Salmonella typhimurium, and

Klebsiella pneumoniae signals for the nitrogen status of the cell and the respective sensors

have been identified and described in detail (Merrick & Edwards, 1995; Ikeda et al., 1996;

Jiang et al., 1998a; Schmitz, 2000), this is not the case for C. glutamicum. Based on the

E. coli model of nitrogen control, GlnD was assumed to be the sensor of the nitrogen status.

But the fact that glnD is part of the AmtR regulon and expressed depending on the

availability of nitrogen, rules out its function as sensor. Overexpression of glnD as well as of

the whole amtB-glnK-glnD operon results in deregulation of nitrogen control. If GlnD was

the sensor, overexpression would not be expected to change the basic properties of

nitrogen control. Therefore, the sensor remains to be identified (Nolden et al., 2001b).

Concerning signals indicating the metabolic state different molecules have been discussed.

In other microorganisms glutamine serves as signal for the cellular nitrogen status (Ikeda et

al., 1996; Brauer et al., 2006). However, this seems not to be true for C. glutamicum. High

internal concentrations of glutamine and glutamate (Krämer & Lambert, 1990; Tesch et al.,

1999; Nolden et al., 2001b) react too slow to be responsible for signaling changes in

nitrogen supply. Instead, they might serve as buffers for nitrogen and/or carbon supply

(Nolden et al., 2001b). Recent studies on the effect of nitrogen availability on different

metabolites contributed significantly to the identification of possible compounds transmitting

the current state of nitrogen metabolism. The expression of AmtR-regulated gltB directly

correlates with the internal ammonium, but not glutamine, concentration indicating that the

nitrogen status might be resembled by the ammonium supply. Most probably, internal

Introduction 11

ammonium concentration rather than the external concentration is responsible. Additionally,

a second marker metabolite seems to be involved. It could be observed that within

30 minutes of nitrogen deprivation the internal concentration of 2-oxoglutarate increased up

to a factor of about 30 and decreased again immediately after the addition of ammonium

(Müller et al., 2006). Generally, PII type signal transduction proteins are referred to as

sensors of 2-oxoglutarate concentration (Ninfa & Jiang, 2005) and therefore integrate

signals from central metabolism and nitrogen metabolism. This has been demonstrated for

PII of E. coli in great detail, which synergistically binds 2-oxoglutarate and ATP, and on the

other hand is modified in response to the internal glutamine concentration (Kamberov et al.,

1995; Ninfa & Jiang, 2005). For GlnK, the only PII type protein in C. glutamicum, binding of

2-oxoglutarate has not been shown so far. However, regulation of nitrogen metabolism in

response to signals indicating nitrogen and carbon status of the cell seems to be conserved

within different organisms (Brauer et al., 2006). Also in C. glutamicum, intermediates of

both, central and nitrogen metabolism seem to exhibit important functions in transferring

signals concerning nitrogen availability into the cell resulting in an appropriate reaction of

the metabolism. This underlines the importance of the interplay between these different

metabolic networks.

2.4 GDH at the interface between nitrogen and carbon metabolism

As already reflected by regulatory mechanisms of nitrogen metabolism, different pathways

interact in assuring an optimal cellular response to environmental stimuli. So, there is more

and more interest in understanding crosstalk of pathways and regulatory networks.

Consequently, mechanisms controlling enzymes connecting these are in the focus of

research.

Besides glutamine synthetase and glutamate synthase, glutamate dehydrogenase acts as

direct link between carbon and nitrogen metabolism. In contrast to eukaryotic enzymes,

bacterial GDH enzymes are characterized by coenzyme specificity, meaning that they

either use NADH or NADPH. Generally, NADH-dependent enzymes catalyze catabolic

reactions, in this case the oxidative deamination of glutamate leading to 2-oxoglutarate. The

reaction contributing to anabolism, the synthesis of glutamate, is performed by NADPH-

dependent enzymes (for review, Minambres et al., 2000). Due to a common structural

feature, namely the molecular weight of respective subunits, GDH enzymes are combined

in families. Based on hierarchical homology grouping, the subfamily of small GDH (α6-50

subfamily) enzymes, which consists of hexameric proteins with a molecular weight of about

Introduction 12

50 kDa per subunit, is further divided into two classes (Minambres et al., 2000). Regarding

the three-dimensional structure as well as catalytic mechanism, the enzymes of Clostridium

symbiosum and E. coli have been well investigated (Baker et al., 1992; Korber et al., 1993;

Stillman et al., 1999). Despite a broad structural knowledge, comparably little is known on

mechanisms involved in regulation of GDH enzymes, even in well-characterized enteric

bacteria (Yan, 2007). So far, regulation by covalent modification as for instance described

for GS, has been excluded for bacterial GDH enzymes (Minambres et al., 2000).

Nevertheless, for the model organisms E. coli and Bacillus subtilis regulatory mechanisms

have been described in dependence of nitrogen and carbon metabolism. The following

overview on these processes underlines the crucial function of GDH in connecting the two

metabolic pathways.

2.4.1 Glutamate dehydrogenase of E. coli

The E. coli GDH (encoded by gdh; Valle et al., 1983) is a α6-50I class enzyme (Minambres

et al., 2000) involved in NADPH-dependent ammonium assimilation under nitrogen surplus

as it has also been described for C. glutamicum. According to its function, gdh transcription

is regulated depending on nitrogen availability leading to a decrease in transcript level upon

nitrogen deprivation. Repression under low ammonium concentrations is mediated by the

Nac protein. It is referred to as the adapter between the nitrogen regulatory (Ntr) system

and primary sigma factor (σ70)-dependent promoters and it also acts as a repressor of

serine and glutamate metabolism (Muse & Bender, 1998; Camarena et al., 1998; Zimmer et

al., 2000; Blauwkamp & Ninfa, 2002). Nac itself is considered a so-called nitrogen-

controlled (Ntr) gene. Transcription of these genes is induced upon nitrogen deprivation.

Nitrogen control in E. coli differs from the C. glutamicum model, even though PII type

proteins (encoded by glnB and glnK) and a bifunctional uridylyltransferase (encoded by

glnD) are also key components in the respective regulatory network (Jiang et al., 1998a).

Nitrogen-dependent transcription is controlled by a two component system consisting of the

histidine kinase NRII (encoded by glnP) controlling the phosphorylation state of NRI

(encoded by glnL). NRII exhibits phosphatase activity when interacting with unmodified PII

resembling sufficient nitrogen supply. At nitrogen deprivation, PII-UMP3 does not interact

with NRII and as a result NRI is phosphorylated by the histidine kinase (Jiang et al., 1998b).

Via interaction with σ54 RNA polymerase, NRI-P activates transcription of Ntr genes leading

to an adaptation to limited nitrogen supply (Kustu et al., 1989; Atkinson et al., 2002). Due to

its low substrate affinity, the regulation of GDH in response to decreasing nitrogen supply

seems to be a reasonable reaction (for overview, Merrick & Edwards, 1995; Reitzer, 2003).

Introduction 13

So far, little knowledge is available about regulation of gdh depending on the cellular carbon

state. But in addition to the Nac-controlled transcription, it was postulated that another

regulatory mechanism is present (Camarena et al., 1998). Central metabolism of E. coli and

regulation by carbon catabolite repression have been in the focus of interest for decades.

The global regulator CAP (catabolite activator protein) activates transcription of more than

100 promoters (Busby & Ebright, 1999). As a prerequisite for DNA binding, the allosteric

effector cAMP needs to be present indicating growth on less preferred non-PTS

(phosphoenolpyruvate-dependent phosphotransferase system) sugars (Brückner &

Titgemeyer, 2002; Mao et al., 2007). Interaction of regulatory networks responding to

carbon source and nitrogen metabolism has been examined. A well-investigated example

of how regulators of nitrogen and carbon metabolism contribute to fine-tuning of expression

is the regulation of glutamine uptake and synthesis by GlnHPQ and GS. The glnA gene and

the glnHPQ operon are both expressed from a σ70-and σ54-dependent promoter. Besides

NRI-P and σ54-dependent transcription, CAP antagonizes this activating effect by

repressing the respective σ54-dependent promoters. Fine-tuning of transcription is achieved,

on the other hand, by CAP-mediated activation of σ70-dependent promoters in response to

changes in nitrogen and carbon supply. Furthermore, the influence of carbon source and

cAMP on the uridylylation state of PII and consequently the Ntr system in the presence of

glutamine as nitrogen source hint at coordinated control of global carbon and nitrogen

signal transduction pathways. (Tian et al., 2001; Maheswaran & Forchhammer 2003; Mao

et al., 2007). Put together, studies on glutamine transport and assimilation provided first

ideas on the complexity and the importance of crosstalk between carbon and nitrogen

metabolism. However, for the regulation of gdh expression and glutamate synthesis in

E. coli details are still missing. Within the gdh promoter region a possible CAP-binding site

has been identified (Valle et al., 1983). Together with the observation that cAMP increases

GDH activity (Prusiner et al., 1972), regulation of gdh in dependence of the carbon source

and status can be implied.

2.4.2 Glutamate dehydrogenase of B. subtilis

In B. subtilis, the interplay of regulatory networks regarding nitrogen and carbon state is

investigated in more detail describing a combination of regulatory proteins as well as

protein-protein interactions. Glutamate dehydrogenase RocG (encoded by rocG; Belitsky &

Sonenshein, 1998) of B. subtilis is an α6-50II class enzyme, which in contrast to the GDH

enzymes described so far accepts NADH as substrate (Minambres et al., 2000). The

B. subtilis protein is not involved in ammonium assimilation, but rather exhibits a catabolic

Introduction 14

rocG

Arginine Glucose

CcpA HPrAhrCσL

function in the degradation of arginine and ornithine (Belitsky & Sonenshein, 1998). This

implies that nitrogen metabolism and its regulation differ significantly from the examples

described so far. In fact, glutamine and not ammonium serves as the preferred nitrogen

source in B. subtilis. Ammonium assimilation is catalyzed by GS and GOGAT (glnA and

gltAB; Deshpande & Kane, 1980; Fisher et al., 1984) exclusively (Belitsky & Sonenshein,

1998). Despite the presence of a PII type protein, mechanisms of sensing and transmission

of the nitrogen status are not yet understood (Wray et al., 1994). However, at least three

regulatory proteins are involved in controlling expression of gene products of nitrogen

metabolism (for overview, Fisher, 1999). In addition to GlnR and CodY, the regulator TnrA

exhibits a dual function in activation and repression of gene expression. Further regulators

are required for fine-tuning and integration of signals from central metabolism. Regulation of

central metabolism has been well investigated, so that detailed information is available on

catabolite repression mediated by the carbon catabolite protein A (CcpA) and the PTS (for

overview, Titgemeyer & Hillen, 2002; Warner & Lolkema, 2003).

The expression of rocG is controlled by different proteins depending for instance on the

availability of glucose, arginine, ornithine, and glutamate (Belitsky & Sonenshein, 1998). In

the presence of glucose, rocG transcription is repressed by CcpA (Belitsky et al., 2004).

Furthermore, rocG is transcribed depending on the σL subunit of RNA polymerase, a

homolog of the E. coli σ54 subunit. In the presence of arginine and ornithine, transcription is

activated by AhrC and RocR (Belitsky & Sonenshein, 1999; Commichau et al., 2007b). This

complex regulation depending on carbon and nitrogen source indicates the crucial and

versatile function of RocG (figure 2.5).

Similar patterns of controlling expression in response to signals of both nitrogen and carbon

metabolism have been reported also for the gltAB operon and citB encoding aconitase

(Faires et al., 1999; Belitsky et al., 2000; Blencke et al., 2006; Commichau et al., 2007a).

Fig. 2.5: Schematic picture showing regulation of rocG expression. RocG is transcribed in a σL-dependent manner. Induction in the presence of arginine depends on AhrC and RocR (not shown). In the presence of glucose, rocG is repressed by the CcpA/HPr complex (Commichau et al. 2007b).

Introduction 15

2.4.3 Glutamate dehydrogenase of C. glutamicum

As mentioned above, GDH of C. glutamicum is an anabolic, strictly NADPH-dependent

enzyme belonging to the α6-50I class of GDH enzymes. Flux measurements could show

that under ammonium surplus 72 % of ammonium assimilation is performed by this enzyme

and that it uses more than 50 % of NADPH available in the cell (Tesch et al., 1999; Marx et

al., 1999). Due to industrial relevance of glutamate production, GDH has been studied

regarding its function already decades ago (Kimura, 1962; Shiio & Ujigawa, 1978), but

compared to other components of nitrogen metabolism, little is known about regulation of

GDH. Moreover, contrary results make conclusions about regulatory mechanisms involved

even harder. Activity measurements suggested that GDH activity is not altered upon

changes in nitrogen supply (Tesch et al., 1998) and later these results were supported by

transcriptome analyses (Beckers et al. 2005; Silberbach et al., 2005a). This observation is

consistent with data on regulation of GDH in other organisms such as S. typhimurium

(Brenchley et al., 1975). However, recent studies on GDH of C. glutamicum showed

astonishing results. Upon nitrogen starvation, gdh transcription was shown to be enhanced,

which was accompanied by an increase in protein level and enzyme activity (Müller, 2005;

L. Nolden, unpublished results). So far, similar results have only been observed for

NADPH-dependent GDH of Ruminococcus flavefaciens FD-1. Interestingly, this enzyme

shares an amino acid identity of 63 % with the enzyme of C. glutamicum and is so far the

closest homolog known (Antonopoulos et al., 2003). The physiological function of this

unusual regulation remains unknown, but recent work on the transcriptional regulation of

gdh could give some insights into possible regulatory mechanisms. The gdh gene is

transcribed monocistronically and the start of transcription has been identified 284 bp

upstream of the start codon (Börmann et al., 1992). Within this relatively large 5’UTR and

upstream of the transcription start four putative regulators seem to bind. Two of them were

investigated in detail concerning binding sites and influence on transcriptional regulation.

AmtR, the global repressor of nitrogen metabolism, has been characterized quite well as

described above, while the second regulator, FarR, had not been identified in

C. glutamicum before. The latter is a HutC/FarR-type regulator of the GntR family (Müller,

2005). As displayed in figure 2.6, there are two AmtR binding sites and one FarR binding

site within the promoter region, which were identified by DNA affinity purification and gel

shift assays. For AmtR, one binding site upstream of the transcription start had been

predicted before using Hidden-Markov models (Beckers et al., 2005).

Introduction 16

Fig. 2.6: Schematic picture of the gdh promoter region from C. glutamicum. The transcription start has been determined 284 bp upstream of the start codon (Börmann et al., 1992). By a combination of gel shift assays and DNA affinity purification, one FarR binding site and two AmtR binding sites were identified (Müller, 2005).

Despite the identification of the binding site for FarR, it was not yet possible to further

elucidate the function of this regulator. The finding that AmtR binds to the gdh promoter

region is astonishing, since RNA hybridization experiments showed that AmR is not

involved in nitrogen-dependent transcription of gdh (Müller, 2005; L. Nolden, unpublished

results). Since gene expression mediated by a Nac-homolog has been excluded (Faust,

2002), mechanisms of nitrogen-dependent regulation remain unknown. However, GDH has

a significant influence on the nitrogen starvation response, since the lack of this enzyme

leads to transcription of AmtR-regulated genes even in the presence of nitrogen. It has

been proposed that this is due to the missing metabolization of 2-oxoglutarate, which might

serve as a signaling metabolite in C. glutamicum (Müller et al., 2006). These data underline

again the key role of GDH for the metabolism.

Also by a so far not described mechanism, gdh transcription is downregulated in the

absence of glucose and as response to utilization of either 2-oxoglutarate, glutamate,

mannose, or fructose as carbon source (Müller, 2005). This, in addition to the observation

that decreased activity of GS results from carbon deprivation (Schulz et al., 2001), suggests

interplay of regulation between nitrogen and carbon metabolism. Unlike in other model

organisms, little is known about global regulatory mechanisms of central metabolism in

C. glutamicum. There is no experimental evidence for the presence of CcpA or CAP

homologs (Gerstmeir et al., 2004). The lack of diauxic growth and the resulting coutilization

of different carbon sources (Cocaign et al., 1993; Dominguez et al., 1993; Wendisch et al.,

2000) indicate substantial differences in carbon metabolism of C. glutamicum compared to

E. coli and B. subtilis. So far, only regulators of single enzymes of the tricarboxylic acid

(TCA) cycle and acetate metabolism have been identified, such as RamA and RamB and a

regulatory network controlling aconitase expression in dependence of iron availability

(Gerstmeir et al., 2004; Krug et al., 2005; Wennerhold et al., 2005; Cramer et al., 2006;

Bott, 2007). Recently, a possible mechanism of interaction between nitrogen assimilation

gdh

-469 -450 -358 -339 -208 -189

FarR AmtR AmtRlow affinity high affinity

start of transcription

-35 -10gdh

-469 -450 -358 -339 -208 -189

FarR AmtR AmtRlow affinity high affinity

start of transcription

-35 -10

Introduction 17

and the TCA cycle has been proposed for the regulation of the 2-oxoglutarate

dehydrogenase complex (ODHC). At this branch point, 2-oxoglutarate is either used as

substrate in the GDH reaction or oxidized in the TCA cycle. Carbon flux is controlled by

regulation of ODHC, which has a significantly higher substrate affinity than GDH (Bott,

2007). By interaction with OdhI, ODHC activity is inhibited. When OdhI is modified by

protein kinase G (PknG), this interaction does not take place and inhibition is released

(Niebisch et al., 2006). Therefore, the presence of an intact OdhI protein is of crucial

importance for glutamate formation. Furthermore, the deletion of the pknG gene, which

leads to a decreased amount of phosphorylated OdhI, showed a positive influence on

glutamate production depending on the cultivation conditions. Interestingly, an intermediate

of nitrogen metabolism has been proposed as signal of this regulatory cascade. It is

assumed that phosphorylation activity of PknG is stimulated by glutamine (Schultz et al.,

2007).

That means, this newly identified regulatory network is an excellent example for the

connection of nitrogen and carbon metabolism. The fact that assimilation of nitrogen

sources depends on the central metabolism as donor of precursors and energy and

therefore exhibits substantial influence has additionally been demonstrated by global

proteome analyses (Schmid et al., 2000). For these reasons, investigating the crosstalk of

possible regulatory mechanisms is of great interest. In this context, GDH and its regulation

are of notable interest and great relevance.

2.5 Objectives

The availability of global analyses techniques allows a detailed investigation of metabolic

networks. Due to the industrial importance of C. glutamicum, the organism has been in the

focus of research for decades. In order to optimize the application of C. glutamicum for

amino acid production, the TCA cycle as donor of energy and precursors is currently

investigated applying a systems biology-based approach. The GDH reaction is included,

since it is the direct link between TCA cycle and ammonium assimilation. Furthermore, due

to its NADPH dependency, GDH was shown to exhibit an influence on generation of

reductive equivalents. These are of importance for anabolic processes such as lysine

production. Because of the position at this crucial branch-point, the aims of this work are to

characterize the enzyme and to focus on transcriptional regulation of gdh.

For determination of kinetic parameters a protocol for overexpression and purification of

GDH needs to be established. By combining techniques to quantify protein and transcript

Introduction 18

level, GDH will be investigated in well-characterized, lysine-producing strains and

compared to the wild type.

Studies regarding transcriptional control of the gdh gene include the characterization of the

presumed promoter region by mutagenesis and examination of nitrogen-dependent

transcriptional control. Data obtained by mutagenesis studies might be used for the

construction of promoters varying in strength, which is of interest considering the

biotechnological relevance of C. glutamicum. Another aim is to elucidate mechanisms

involved in nitrogen-dependent regulation of gdh transcription, because previous studies

presented contrary results. Furthermore, putative regulators of gdh transcription, namely

FarR and AmtR, need to be closer analyzed regarding influence on gdh expression and

further target genes.

Materials and methods 19

3 Materials and methods

3.1 Bacterial strains and plasmids

Bacterial strains and plasmids used in this study are listed in table 3.1 and table 3.2,

respectively.

Tab. 3.1: C. diphtheriae, C. efficiens, C. glutamicum, C. jeikeium, and E. coli strains used. KmR: resistance to kanamycin.

Strain Genotype, phenotype Reference

C. diphtheriae

DSM 44123 designated ATCC 27010, non-pathogenic strain of unknown

origin Hauser et al., 1993

C. efficiens

DSM 44549 designated YS-314, isolated from soil samples Fudou et al., 2002

C. glutamicum

ATCC 13032 wild type Abe et al., 1967

CL1 RES167 with a deletion of sigB Larisch et al., 2007

DM1132 sequenced wild type

Abe et al., 1967

Degussa GmbH,

Halle

DM1799 DM1132 pycP458S Degussa GmbH,

Halle

DM1800 DM1132 pycP458S_lysCT311I Degussa GmbH,

Halle

DM1868 DM1132 lysCT311I Degussa GmbH,

Halle

LNΔGDH ATCC 13032 with a deletion of gdh Müller et al., 2006

MJ6-18 ATCC 13032 with a deletion of amtR Jakoby et al., 2000

RES167 restriction-deficient mutant of ATCC 13032 Δ(cgIIM-cgIIR-

cgIIR) Tauch et al., 2002


Strain Genotype, phenotype Reference

RES ΔsigD RES167 with a deletion of sigD

Larisch & Kalinowski,

University of Bielefeld,

unpublished results

RES ΔsigE RES67 with a deletion of sigE



unpublished results

RES ΔsigM RES167 with a deletion of sigM



unpublished results

RES INTsigH RES167 with an integration in sigH, KmR



unpublished results

TMΔfarR RES167 with a deletion of farR Hänßler et al., 2007

TMΔoxyR RES167 with a deletion of oxyR Müller, 2005

C. jeikeium

K411 nosocomial pathogen of the human skin flora,

isolated from blood cultures Tauch et al., 2005

E. coli

DH5αmcr

endA1 supE44 thi-1 λ- recA1 gyrA96 relA1 deoR

Δ(lacZYA-argF) U169 φ80ΔlacZ ΔM15mcrA

Δ(mmr hsdRMS mcrBC)

Grant et al., 1990

JM109

f`traD36 laclq Δ(lacZ)M15 proA+B+ / e14- (McrA-)

Δ(lac-proAB) thi gyrA96 (NxR) endA1 hsdR17

(r-km-

k) relA1 supE44 recA1

Yanisch-Perron et al.,

1985


Tab. 3.2: Plasmids used in this work. For plasmids, which were constructed as part of this work, a detailed description is given in the appendix. ApR: resistance to ampicillin, KmR: resistance to kanamycin.

Plasmid Description Reference

pEKEx2 C. glutamicum expression vector ptac, lacIq,

KmR, ori C. g., ori E. c. Eikmanns et al., 1991

pEKExgdh C. glutamicum expression vector for his-tagged

GDH, ptac, lacIq, KmR, ori C. g., ori E. c. this work

pEPR1 E. coli-Rhodococcus promoter-probe vector,

promoterless gfpuv as reporter, KmR Knoppová et al., 2007

pEPRP45

E. coli-Rhodococcus promoter-probe vector,

gfpuv under control of the P45 promoter as

positive control, KmR

Pátek, unpublished work

pEPRpgdh527 pEPR1 carrying a 527 bp fragment of the gdh

promoter region this work

pEPRpgdh527_1 pEPR1 carrying a 527 bp fragment of the gdh

promoter region including a mutated -10 region this work

pEPRpgdh527_2

pEPR1 carrying a 527 bp fragment of the gdh

promoter region with mutated extended -10

region

this work

pEPRpgdh527_3 pEPR1 carrying a 527 bp fragment of the gdh

promoter region including a mutated -10 region this work

pEPRpgdh527_4


promoter region with mutated extended -10

region

this work

pEPRpgdh527inv pEPR1 carrying the inverted 527 bp fragment of

the gdh promoter region this work



pEPRpgdh398_1


promoter region with a mutated extended -10

region

this work

pEPRpgdh398inv pEPR1 carrying the inverted 398 bp fragment of

the gdh promoter region this work




Plasmid Description Reference

pEPRpgdh298


promoter region with a mutated extended -10

region

this work





pQE30Xagdh Expression vector for his-tagged GDH from

C. glutamicum, ApR, ori ColE1, pT5 Müller, 2005

pUC18 ApR, lacZα Viera & Messing, 1982

pUCargR Expression vector of argR from C. glutamicum

derived from pUC18, ApR, lacZα this work

3.2 Cultivation of bacteria

3.2.1 Culture medium for E. coli

E. coli strains were routinely grown in Luria-Bertani (LB) medium or on LB plates (table 3.3).

Plates were supplemented with 15 g/l Bacto-Agar (Difco, Detroit, USA).

3.2.2 Culture media for corynebacteria

As complex medium for cultivation of C. diphtheriae, C. efficiens, and C. glutamicum Brain

Heart Infusion (BHI) medium and BHI plates were used. C. jeikeium was cultivated in BYT

medium (Tauch et al., 2005). To the respective plates 15 g/l Bacto-Agar (Difco, Detroit,

USA) were added. As defined minimal medium for C. glutamicum CgC medium was used

(Keilhauer et al., 1993). In order to induce nitrogen and carbon starvation, cells were

transferred to CgCoN (Jakoby et al., 2000) or CgCoC medium. The composition of all

media used is listed in table 3.3.


Tab. 3.3: Media used for cultivation of E. coli, C. diphtheriae, C. efficiens, C. glutamicum, and C. jeikeium strains. The composition is given for 1 liter.

Medium Ingrediens (per l)

LB 10 g tryptone, 5 g yeast extract, 10 g NaCl. Sterilization by autoclavation

at 121°C for 20 minutes

BHI 37 g Brain Heart Infusion (Difco, Detroit, USA and Oxoid, Heidelberg).

Sterilization by autoclavation at 121°C for 20 minutes

BYT

37 g Brain Heart Infusion (Difco, Detroit, USA and Oxoid, Heidelberg),

0.4 % (w/v) yeast extract, 0.2 % (v/v) Tween 80. Sterilization by

autoclavation at 121°C for 20 minutes

CgC

42 g MOPS, 20 g (NH4)2SO4, 5 g urea, 0.5 g KH2PO4, 0.5 g K2HPO4, pH

(NaOH) 7.0. After autoclavation addition of: 10 ml 100 mM CaCl2, 10 ml

1 M MgSO4, 200 µg biotin, 1 ml trace element solution, 80 ml 50 % (w/v)

glucose

CgCoN

42 g MOPS, 0.5 g KH2PO4, 0.5 g K2HPO4, pH (NaOH) 7.0. After

autoclavation addition of: 10 ml 100 mM CaCl2, 10 ml 1 M MgSO4, 200 µg

biotin, 1 ml trace element solution, 80 ml 50 % (w/v) glucose

CgCoC

42 g MOPS, 20 g (NH4)2SO4, 5 g urea, 0.5 g KH2PO4, 0.5 g K2HPO4, pH

(NaOH) 7.0. After autoclavation addition of: 10 ml 100 mM CaCl2, 10 ml

1 M MgSO4, 200 µg biotin, 1 ml trace element solution, 80 ml H2O

CgCoN with glutamine

42 g MOPS, 0.5 g KH2PO4, 0.5 g K2HPO4, pH (KOH) 7.0. After

autoclavation addition of: 10 ml 100 mM CaCl2, 10 ml 1 M MgSO4, 200 µg

biotin, 1 ml trace element solution, 80 ml 50 % (w/v) glucose, and 100 mM

glutamine

Trace element solution

28.5 g FeSO4 x 7H2O, 16.5 g MnSO4 x H2O, 6.4 g ZnSO4 x 7H2O, 764 mg

CuSO4 x 5H2O, 128 mg CoCl2 x 6H2O, 44 mg NiCl2 x 6H2O, 64 mg

Na2MoO4 x 2H2O, 48 mg H3BO3, 50 mg SrCl2, 50 mg BaCl2 x 2H2O,

28 mg KAl(SO4)2 x 12H2O, pH (H2SO4) 1. Sterilization by filtration

3.2.3 Antibiotics

If appropriate, antibiotics were added to the media in the following final concentrations:

100 µg/ml ampicillin, 50 µg/ml carbenicillin, 25 µg/ml kanamycin, or 10 µg/ml kanamycin

(latter only for C. glutamicum strains after electroporation).


3.2.4 Growth conditions

E. coli strains as well as C. diphtheriae, C. efficiens, and C. jeikeium were grown at 37°C,

whereas the temperature for cultivation of C. glutamicum strains was 30°C. In order to

ensure sufficient agitation during growth, baffled flasks were used and the frequency of the

rotary shaker (SM30, Edmund Bühler, Tübingen) was set to 135 and 125 rpm, respectively.

To assure highly reproducible conditions, a standardized inoculation scheme was applied

for cultivation of C. glutamicum: 20 ml BHI medium were inoculated and incubated as

described previously for about 8 hours. This culture was used for inoculation of 50 ml of

CgC medium to an optical density at a wavelength of 600 nm (OD600) of 0.5 as overnight

culture. This procedure leads to an optimal adaptation to the minimal medium. On the next

day, fresh CgC medium was inoculated to an OD600 of 1. This corresponds to approximately

109 cells per ml. Depending on the stress condition to be investigated cells were treated

differently, when exponential growth phase was reached. To cause nitrogen or carbon

starvation, cultures were transferred to CgCoN or CgCoC medium by centrifugation

(4,000 xg, 10 min) and resuspension in the respective medium. Incubation was carried out

for 1 hour. As control, one culture was resuspended in CgC medium again after

centrifugation. Cold shock was induced by transferring cells to fresh CgC medium, followed

by an hour of cultivation at 15°C. The procedure of sample preparation is further described

in the next sections.

For overexpression studies using pEKEx2, C. glutamicum strains were cultivated as just

described and appropriate concentrations of IPTG were used for induction when an OD600

of about 4 was reached.

In order to purify GDH, cells were not transferred to minimal medium. Overnight growth also

occurred in BHI medium and on the next day fresh BHI medium was inoculated to an OD600

of 1. When the exponential growth phase was reached at about an OD600 of 4, IPTG was

added to a final concentration of 0.5 mM and the culture was incubated for 2 hours. The

procedure is further described in section 3.6.1.

For cultivation of C. diphtheriae, C. efficiens, and C. jeikeium, the standardized inoculation

scheme was slightly modified. For reason of better comparison, C. glutamicum was also

cultivated according to this protocol. 100 ml BHI or BYT medium were inoculated and

cultivation was performed overnight at 30°C and 37°C, respectively. On the next day, these

cultures were used to inoculate 100 ml fresh BHI or BYT medium to an OD600 of about 0.2.

After 2-3 hours, exponential growth phase was reached. All cultures were divided into 50 ml

aliquots and centrifuged (4,000 xg, 7 min). One aliquot was used to carry out uptake

measurements and determine enzyme activity under nitrogen surplus, whereas the other


aliquot was resuspended in CgCoN medium. After one hour of incubation without nitrogen

source, samples were taken again for uptake measurements and determination of enzyme

activity.

For long-term storage of bacterial strains permanent cultures were made using Roti®-Store

tubes (Roth, Karlsruhe) as described by the manufacturer and stored at -80°C.

3.3 Genetic manipulation of bacteria

3.3.1 Preparation of competent E. coli cells and transformation

Chemically competent E. coli cells were prepared as described by Inoue et al. (1990). For

this, 5 ml LB medium were inoculated with E. coli cells and cultivated for 8 hours at 37°C.

1 ml of this culture was used to inoculate 250 ml SOB medium for overnight growth at 20°C

in a 2 l flask. On the next day, the culture was chilled on ice for 10 minutes. Cells were

harvested by centrifugation (4,000 xg, 10 min, 4°C). The cell pellet was resuspended in

80 ml ice-cold TB buffer and centrifuged again (4,000 xg, 10 min, 4°C). After resuspending

in 40 ml TB buffer supplemented with 1.4 ml DMSO, cells were incubated for 10 minutes on

ice. Aliquots of 100 µl were transferred to pre-cooled reaction tubes. These were

immediately frozen in liquid nitrogen and stored at -80°C.

For transformation, a 100 µl aliquot of competent E. coli cells was thawed on ice and

plasmid DNA was added. Cells were incubated for 30 minutes on ice. After heat shock at

42°C for 30 seconds, 400 µl SOC medium were added. The cell suspension was cultivated

for 1 hour at 37°C. After that, 200 µl of the cell suspension were plated on LB plates

containing the appropriate antibiotic.

TB buffer:

10 mM Pipes, 15 mM CaCl2, 250 mM KCl, pH (KOH) 6.7, after adjustment of pH, addition of

55 mM MnCl2; sterilization by filtration

SOB medium:

20 g/l tryptone, 5 g/l yeast extract, 0.5 g/l NaCl, 2.5 mM KCl, after autoclavation, addition of

5 ml 2 M MgCl2


SOC medium:

10 g/l tryptone, 5 g/l yeast extract, 5 g/l NaCl, 3.6 g/l glucose, after autoclavation, addition of

5 ml 2 M MgCl2

3.3.2 Preparation of competent C. glutamicum cells and transformation

To prepare competent C. glutamicum cells, 20 ml LB medium with 2 % (w/v) glucose were

inoculated and cultivated for 8 hours at 30°C. Subsequently, this culture was used to

inoculate 200 ml LB medium with growth inhibitors to an OD600 of about 0.2-0.3. Cells were

cultivated at 20°C in a 2 l flask overnight. On the next day, cells were harvested by

centrifugation (4,000 xg, 5 min, 4°C). The cell pellet was washed five times with ice-cold

10 % (v/v) glycerol. Finally, the cell pellet was resuspended in 1 ml ice-cold 10 % (v/v)

glycerol. Aliquots of 50 µl were transferred to pre-cooled reaction tubes. These were

immediately frozen in liquid nitrogen and stored at -80°C (Liebl et al., 1989).

For transformation, a 50 µl aliquot of competent C. glutamicum cells was thawed on ice and

transferred to a pre-cooled electroporation cuvette with 2 mm electrode clearance (peqlab

Biotechnologie GmbH, Erlangen). 0.3-1 µl plasmid DNA were added and electroporation

was performed with a Gene-Pulser (Biorad, Munich) at 2.5 kV, 600 Ω, and 25 µF. Right

after that, 1 ml BHIS medium was added and heat shock for 6 minutes at 46°C was

performed. Cells were cultivated for 2 hours at 30°C and 500 µl of the cell suspension were

plated on a BHI plate containing the appropriate antibiotic.

LB medium with growth inhibitors:

10 g/l tryptone, 5 g/l yeast extract, 5 g/l NaCl, 4 g/l isonicotine acid hydrazide, 25 g/l glycine,

0.1 % (v/v) Tween 80, sterilization by filtration

BHIS medium:

37 g/l Brain Heart Infusion, 91 g/l sorbitol, sterilization by filtration

3.4 Working with DNA

3.4.1 Isolation of plasmid DNA from E. coli

For plasmid preparation, E. coli colonies were grown overnight in 4 ml LB medium

containing an appropriate antibiotic. To isolate plasmid DNA from E. coli, the NucleoSpin®

Plasmid kit and the NucleoSpin® QuickPure kit (Macherey-Nagel, Düren) were used as


recommended by the supplier followed by agarose gel electrophoresis to check the

resulting plasmids.

3.4.2 Gel electrophoresis and extraction of DNA from agarose gels

Gel electrophoresis of DNA was performed in 0.8-2 % agarose gels in 1 x TAE buffer as

described by Sambrook et al. (1989). Samples were mixed with 6 x loading dye.

Electrophoresis was performed at 10 V/cm of gel length. After electrophoresis, DNA was

stained with ethidium bromide solution. To be able to determine the size of DNA fragments,

DNA marker (Lambda-DNA/Eco911 (BstE(II) cut), MBI Fermentas, Wilna and peqGOLD

ladder, peqlab Biotechnologie GmbH, Erlangen) was applied. For detection of stained DNA,

the Doc Print 1000 Gel documentation (peqlab Biotechnologie GmbH, Erlangen) was used.

DNA was isolated from agarose gels using the NucleoSpin® Extract kit (Macherey-Nagel,

Düren) as recommended by the supplier.

1 x TAE buffer:

40 mM Tris, 0.5 mM EDTA, pH (acetic acid) 7.5

6 x loading dye:

0.25 % (w/v) bromphenol blue, 0.25 % (w/v) xylene cyanol FF, 40 % (w/v) sucrose

3.4.3 Preparation of chromosomal DNA from C. glutamicum

Preparation of chromosomal DNA was performed using phenol chloroform extraction.

C. glutamicum strains were cultivated in 5 ml BHI medium overnight. On the next day,

cultures were centrifuged (4,000 xg, 10 min, 4°C) and the pellet was resuspended in 200 µl

H2O. 200 µl phenol were added and this mixture was incubated at 65°C for 10 minutes.

Prior to the addition of 200 µl chloroform, the mixture was put on ice to cool down. After

centrifugation at 13,000 xg and 4°C for 5 minutes, the supernatant was transferred into a

fresh reaction tube. To this, again 200 µl chloroform were added and the mixture was

centrifuged as described above. The supernatant containing the DNA was separated from

the organic phase and the DNA was diluted 1:10 in H2O. For PCR experiments (3.4.5) 1 µl

of chromosomal DNA was used as template in a 25 µl reaction volume.

3.4.4 Purification and enrichment of DNA

In order to purify DNA and to increase the concentration, ethanol precipitation was carried

out. To the reaction mixture 1/10 volume 3 M sodium acetate pH 4.8 and 2.5-fold volume


100 % ethanol were added. After incubation for at least one hour at -20°C, the precipitated

nucleic acids were centrifuged (16,060 xg, 30 min, 4°C), the pellet was washed with

70 % (v/v) ethanol (16,060 xg, 5 min, 4°C) and air dried. Finally, the pellet was suspended

in H2O or TE buffer.

TE buffer:

10 mM Tris, 1 mM EDTA, pH (HCl) 7.5

3.4.5 Polymerase chain reaction (PCR)

The selective amplification of specific DNA fragments was performed by polymerase chain

reaction (PCR) (Mullis et al., 1986). As template either genomic DNA or a cell suspension,

which had previously been incubated at 95°C for 5 minutes, was used. Primers were

supplied by Operon (Cologne), Eurogentech (Seraing, Belgium), and MWG (Ebersberg)

and diluted in H2O to a final concentration of 100 pmol/µl. Depending on the primer

sequence the annealing temperature was calculated. For breaking of hydrogen bonds

between guanine and cytosine 4°C were estimated and for adenine and thymine 2°C. Since

only DNA fragments smaller than 2 kb were amplified, the Master-Mix Kit (Qiagen, Hilden),

Taq Polymerase (NEB, Frankfurt/Main) and PuRe Taq Ready-To-Go PCR beads (GE

Healthcare, Munich) were used as recommended by the suppliers. The PCR was

performed with a Mastercycler® personal or Mastercycler® gradient (Eppendorff, Hamburg).

First of all, template DNA was denatured by an initial incubation at 95°C for 5 minutes. After

that, the following steps were repeated 30 times: double-stranded DNA was denatured for

15 seconds at 95°C. Depending on the sequence of the primers used, annealing was

performed for 15 seconds at 57°C to 60°C. At a temperature of 72°C double-stranded DNA

was synthesized by the polymerase. This step was carried out for 1 minute per kb. By final

incubation at 72°C for 10 minutes, it was ensured that the synthesis of fragments was

completed. The resulting PCR product was analyzed by agarose gel electrophoresis and

subsequently stored at 4°C. If appropriate, it was purified either with the NucleoSpin®

Extract Kit (Macherey-Nagel, Düren) as recommended by the supplier or by gel

electrophoresis as described in section 3.4.2.

3.4.6 Two-step PCR

In order to create specific mutations in the gdh promoter region, two-step PCR was carried

out. In a first PCR performed as described in section 3.4.5 a primer carrying the desired

mutation was used. Subsequently, in a second PCR 2 µl of the purified, first PCR product


were used instead of the forward primer in a 25 µl reaction. The resulting PCR product

carrying point mutations was then purified as described in 3.4.2 and cloned into the plasmid

pEPR1.

3.4.7 Restriction of DNA

For restriction of DNA, restriction enzymes were used as recommended by the suppliers

(NEB, Frankfurt/Main; MBI Fermentas, Wilna). If appropriate, 1 µl shrimp alkaline

phosphatase or 1 µl antarctic phosphatase was added to dephosphorylate 5’ ends of

plasmid DNA (NEB, Frankfurt/Main). After restriction and dephosphorylation, DNA was

purified by gel electrophoresis with the NucleoSpin® Extract kit (Macherey-Nagel, Düren)

following the supplier’s protocol or as described in section 3.4.2.

3.4.8 Ligation of DNA fragments

For the ligation of DNA fragments to restricted vectors, the Rapid DNA Ligation kit (MBI

Fermentas, Wilna) or T4 DNA ligase (NEB, Frankfurt/Main) were used as recommended.

After ligation, 5 µl of the reaction mix were used to transform 100 µl competent E. coli cells

as described in section 3.3.1.

3.4.9 Sequencing of DNA

Sequencing of DNA was performed by using the chain termination method described by

Sanger et al. (1977). As chain terminators didesoxynucleotides labeled with fluorescent dye

were used (Big Dye Terminator Mix, Perkin Elmer, Weiterstadt). Sequencing reactions were

carried out as recommended by the supplier using the following PCR program:

96°C 19 s

25 x 60°C 5 s

60°C 4 min

The resulting fragments were analyzed using the ABI PRISM Genetic Analyser 310

(Applied Biosystems, Freiburg) and sequence data were evaluated with the Seqman

software (DNAstar, Madison, USA).

Further sequence analyses were carried out by the bioanalytics service unit at the Center

for Molecular Medicine Cologne (ZMMK). Samples contained between 300-500 ng of

plasmid DNA and 1 µl primer (10 µM). For the sequencing reaction, the Taq FS BigDye-

terminator cycle sequencing method and a Prism 377 DNA Sequencer (Applied


Biosystems, Freiburg) were used. Data were analyzed using Sci Ed Central for Windows 95

(Scientific & Educational Software, Cary, USA) and Chromas Lite Version 2.0

(Technelysium, Tewantin, Australia) as well as the Seqman software (DNAstar, Madison,

USA).

3.5 Working with RNA

3.5.1 Isolation of total RNA and RNA gel electrophoresis

1 ml of a C. glutamicum culture with an OD600 of 4-5 was centrifuged (16,060 xg, 30 s) and

the cell pellet was immediately frozen in liquid nitrogen and stored at -80°C. The cell pellet

was thawed on ice and resuspended in RA1 buffer containing 1 % (v/v) 2-mercaptoethanol

(NucleoSpin® RNA II Kit, Macherey-Nagel, Düren). Immediately after resuspension of the

cell pellet, the suspension was transferred to a tube containing glass beads and cells were

disrupted by vigorous shaking at 6.5 m s-1 for 30 seconds using a FastPrep FP120

instrument (Q-BIOgene, Heidelberg). After another cycle of disruption, cell debris and glass

beads were removed by centrifugation (16,060 xg, 3 min, 4°C) and the supernatant was

mixed with 350 µl 70 % (v/v) ethanol. In the following, total RNA was isolated using the

NucleoSpin® RNA II kit (Macherey-Nagel, Düren) as recommended by the supplier. The

purity of total RNA isolated by this approach was sufficient for Northern blot and Dot blot

hybridization experiments. However, for reverse transcriptase (RT) PCR and quantitative

real time RT PCR experiments, total RNA was additionally treated with TURBO-DNase

(Ambion, Austin, USA) and subsequently purified again with the NucleoSpin® RNA II kit

(Macherey-Nagel, Düren) as recommended by the supplier. Alternatively, the RNeasy Kit

(Qiagen, Hilden) was used in combination with on column DNaseI treatment (Qiagen,

Hilden). For this, sample volumes up to 20 ml were used.

The integrity of isolated total RNA was analyzed by gel electrophoresis. 3 µl of purified RNA

were mixed with 5 µl RNA loading buffer and incubated for 10 minutes at 65°C. After that,

the sample was placed on ice for 5 minutes. In the following, gel electrophoresis and

detection of RNA was performed in accordance to the analysis of DNA described in section

3.4.2. The concentration of RNA samples was measured at a wavelength of 260 nm

(ε= 5 cm2/mg) using the Nanodrop spectrophotometer ND-100 (peqlab Biotechnologie

GmbH, Erlangen). Purity was analyzed by determination of the quotient of A260 / A280, which

is 2.0 for pure RNA.


RNA loading buffer:

0.2 % (w/v) bromphenol blue, 0.2 % (w/v) xylene cyanol, 65 % (v/v) formamide, 12 % (v/v)

formaldehyde, 2 x MOPS, 2 % (w/v) sucrose

10 x MOPS buffer:

200 mM MOPS, 50 mM sodium acetate, 10 mM EDTA, pH (NaOH) 7.0

3.5.2 Synthesis of digoxigenin-labeled RNA probes

Digoxigenin-labeled RNA probes were synthesized by in vitro transcription. The probe was

labeled by the use of digoxigenin-11-dUTP instead of dUTP. The reaction mixture for the in

vitro transcription was composed of:

14 µl template DNA (purified PCR product)

2 µl DIG RNA Labeling Mix (Roche Diagnostics, Mannheim)

2 µl 10 x RNA Polymerase Buffer (NEB, Frankfurt/Main)

1 µl T7 RNA Polymerase (NEB, Frankfurt/Main)

The reaction mixture was incubated for 2 hours at 37°C. After that, 1 µl RNase-free

peqGOLD DNaseI (peqlab Biotechnologie GmbH, Erlangen) was added and the mixture

was incubated for another 20 minutes at 37°C. The resulting digoxigenin-labeled RNA

probe was stored at -80°C.

3.5.3 Northern blot analysis

In order to determine the size of specific mRNA, Northern blot experiments were carried

out. For these, 5 µg of total RNA were mixed with RNA loading buffer and incubated at

65°C for 10 minutes. Samples were then loaded onto a denaturing MOPS-buffered RNA

gel. To be able to determine the fragment size, 3 µl RNA ladder (high range, Fermentas,

Wilna) were applied as well. Electrophoresis was performed at 80 V for about 2 hours in 1 x

MOPS. RNA was transferred onto a positively charged nylon membrane (Roche

Diagnostics, Mannheim) by using a vacuum blot apparatus (Amersham Pharmacia,

Freiburg) at 50 mbar. The gel was constantly covered with liquid, first of all for 7 minutes

with denaturing buffer followed by 10 minutes with neutralizing buffer. During the remaining

time it was covered with 20 x SSC. After this transfer, RNA was crosslinked twice to the

membrane by UV radiation at 1200 kJ (UV Stratalinker® 1800, Stratagene, LaJolla, USA).

The denaturing gel was stained with ethidium bromide solution to check the integrity of the


RNA and the RNA concentration. Meanwhile, the membrane was incubated at 68°C for

2 hours in hybridization solution and for overnight hybridization the digoxigenin-labeled

probe was added in a 1:10,000 dilution. On the next day, two washing steps with washing

buffer 1 were carried out for 5 minutes at room temperature. The two following washing

steps were performed at 68°C with washing buffer 2 for 25 minutes. After 1 minute in maleic

acid buffer, 1 x blocking buffer was added and after 30 minutes alkaline phosphatase

conjugated anti-digoxigenin Fab fragments (Roche Diagnostics, Mannheim) were applied in

a 1:10,000 dilution. Incubation with the antibody lasted 30 minutes. Then, the membrane

was washed twice in maleic acid buffer for 15 minutes. For equilibration, the membrane

was put into buffer 3 for 5 minutes. All washing steps were carried out under gentle

agitation at room temperature. For detection, the membrane was covered with CDP* ready

to use reagent (GE Healthcare, Munich) and incubated in the dark for 5 minutes. Signals

were detected using Amersham HyperfilmTM MP (GE Healthcare, Munich).

Denaturing RNA gel:

0.6 g agarose, 5 ml 10 x MOPS, 40 ml H2O, after heating and incubation at 68°C for

30 minutes, addition of 4.25 ml formaldehyde (37 %)

Denaturing buffer:

50 mM NaOH, 10 mM NaCl

Neutralizing buffer:

0.1 M Tris HCl pH 7.4

20 x SSC:

3 M NaCl, 0.3 M sodium citrate, pH (HCl) 7.0

Hybridization mix:

50 ml formamide, 20 ml 10 x blocking buffer, 25 ml 20 x SSC, 1 ml 10 % (w/v) sodium

lauroyl sarcosinate, 0.2 ml 10 % (w/v) SDS, 3.8 ml H2O

Washing buffer 1:

2 x SSC, 0.1 % (w/v) SDS


Washing buffer 2:

0.2 x SSC, 0.1 % (w/v) SDS

10 x blocking buffer:

10 % blocking reagent (Roche Diagnostics, Mannheim) in maleic acid buffer; 10 x blocking

buffer was diluted 1:10 in maleic acid buffer to obtain 1 x blocking buffer.

Maleic acid buffer:

0.1 M maleic acid, 0.15 M NaCl, pH (NaOH) 7.5

Buffer 3:

0.1 M diethanolamine

3.5.4 Dot blot analysis

For the preparation of RNA Dot blots, 1-2.5 µg of total RNA were mixed with 100 µl 10 x

SSC and spotted onto a positively charged nylon membrane (Roche Diagnostics,

Mannheim) previously equilibrated in 10 x SSC using the Minifold I Dot Blotter (Schleicher

& Schuell, Dassel). Subsequently, the nylon membrane was dried and RNA was

crosslinked by UV radiation (125 mJ/cm2) using a Bio-Link instrument (LFT-Labortechnik,

Wasserburg). The membrane was incubated for 1 hour in 12 ml hybridization mix.

Subsequently, 1.2 µl digoxigenin-labeled RNA probe (for preparation refer to section 3.5.2.)

were added and hybridization was performed at 68°C overnight. The following washing

steps were performed similar as described for Northern blot experiments in section 3.5.3.

On the next day, the membrane was washed twice with 20 ml washing buffer 1 for

15 minutes at room temperature. Then, the membrane was washed twice with 20 ml

washing buffer 2 for 15 minutes at 68°C and subsequently, with 20 ml washing buffer 3 for

1 minute. After that, the membrane was incubated for 30 minutes in 12 ml 1 x blocking

buffer. Then, 1.2 µl alkaline phosphatase conjugated anti-digoxigenin Fab fragments

(Roche Diagnostics, Mannheim) were added and the membrane was incubated for another

30 minutes. After three washing steps with washing buffer 3 for 30 minutes at room

temperature, the membrane was equilibrated in detection buffer for 3 minutes. 10 µl CSPD

reagent (Roche Diagnostics, Mannheim) were diluted in 990 µl detection buffer. This

solution was used to moisten the membrane, which was subsequently covered. After an

incubation of 15 minutes at 37°C, light emission was detected using Amersham HyperfilmTM

MP (GE Healthcare, Munich).


Washing buffer 3:

0.3 % (v/v) Tween 20 in maleic acid buffer

Detection buffer:

0.1 M Tris, 0.1 M NaCl, pH (NaOH) 9.0

3.5.5 Reverse transcriptase (RT) PCR

Reverse transcriptase PCR was carried out using the OneStep RT PCR kit (Qiagen, Hilden)

as recommended by the supplier. Of total, Turbo-DNase treated RNA (3.5.1) 0.5 µg were

used in the reaction, which was performed according to the following program: for reverse

transcription of mRNA into cDNA, the reaction mix was incubated for 30 minutes at 50°C.

The resulting cDNA was amplified in a PCR with an initial activation of HotStarTaq

polymerase for 15 minutes at 94°C. The PCR included the following 30 cycles: 45 seconds

denaturation at 94°C, 45 seconds at 58°C for primer annealing and elongation at 72°C

between 45 seconds and 90 seconds depending on the fragment size. To complete the

synthesis of all DNA fragments, final extension was performed for 10 minutes at 72°C. To

check the size of the PCR products, agarose gel electrophoresis was carried out as

described in 3.4.2. As negative control for each reaction, one mixture was carried along that

was not incubated at 50°C, so that reverse transcription could not take place.

3.5.6 Quantitative real time RT PCR

Quantitative real time RT PCR was used to detect the amount of a specific mRNA present.

For this purpose, total RNA was isolated from C. glutamicum cells as described in section

3.5.1. Real time RT PCR experiments were carried out in a MyiQ Single-Color Real Time

PCR Detection system (Biorad, Munich). In order to quantify mRNA levels, RNA was

transcribed into cDNA in a reverse transcription reaction. The increase in product during the

following PCR reaction was determined using the fluorescent dye SYBR Green 1, which

binds to double-stranded DNA emitting a fluorescent signal. In the real time RT PCR,

100 ng DNA-free total RNA were used as template. According to the instruction of the

manufacturer, the QuantiTectTM SYBR® Green RT PCR KIT (Qiagen, Hilden) was used.

Primers were supplied by MWG (Ebersberg) and dissolved in H2O to a final concentration

of 100 pmol/µl. In order to guarantee a constant efficiency of the polymerase, the size of the

amplified fragment was always set to 150 bp.


A 20 µl real time RT PCR mixture contained:

10 µl 2 x QuantiTectTM SYBR® Green RT PCR Master Mix (Qiagen, Hilden)

1 µl primer forward [1 µM]

1 µl primer reverse [1 µM]

0.2 µl QuantiTectTM RT Mix (Qiagen, Hilden)

2 µl fluorescein [100 nM] (Biorad, Munich)

100 ng total RNA

RNase-free H2O ad 20 µl

The addition of fluorescein to the reaction mix allows normalization of fluctuations resulting

from pipetting errors. Real time RT PCR was performed using the following program:

During the first step, mRNA was transcribed into cDNA at 50°C, followed by denaturing of

the DNA/RNA hybrid at 95°C. cDNA was amplified during the following 40 cycles. After

each cycle, the rising amount of PCR product was monitored by measuring the increase in

fluorescence due to the binding of SYBR Green dye to the DNA. In order to check the purity

and specificity of the product, a melt curve was created after the PCR. Of each sample, at

least three replicates were used for determination of transcript level. To assure the quality

of the experiment, for each set of primers a “no template” control was added. For evaluation

the software provided by the supplier was used, so that differences in gene expression

could be quantified using the application “Gene Expression Analysis”.

3.5.7 Primer extension analysis

Non-radioactive primer extension analysis was performed as described by Engels et al.

(2004). 15 µg of total RNA were combined with 2 pmol IRD800-labeled oligonucleotide

(MWG, Ebersberg) and 2 µl annealing buffer in a total volume of 10 µl. This reaction was

heated to 65°C for 5 minutes and slowly cooled down to 42°C (0.5°C every 2 minutes).

Reverse transcription was then started by the addition of 10 µl 5 x RT buffer, 23 µl H2O,

30 min 50°C

15 min 95°C

15 s 94°C

15 s 60°C

15 s 72°C

Melt curve 55°C-100°C

40 cycles


0.5 µl actinomycin D (5 mg ml-1 in ethanol), 1 µl desoxyribonucleotides (25 mM ATP, CTP,

GTP, and TTP), 5 µl DTT and, 1 µl of SuperScript II RNase H- reverse transcriptase

(200 units µl-1, Invitrogen, Karlsruhe). After 1 hour of incubation at 42°C, the reaction was

stopped by the addition of 120 µl RNase A reaction mix and incubated for another hour at

37°C. DNA was precipitated at -20°C overnight by adding 17 µl sodium acetate (pH 5.2)

and 380 µl ice-cold ethanol. The resulting pellet (16,000 xg, 30 min, 4°C) was washed with

500 µl 70 % (v/v) ethanol and dissolved in 2 µl H2O. Before applying samples onto a

denaturing 4.6 % (w/v) Long Ranger (Cambrex, Bedford, USA) sequencing gel, 2 µl

formamide loading dye (Thermo Sequenase Primer Cycle, Amersham Biosciences,

Freiburg) were added followed by incubation at 70°C for 5 minutes. 1.5 µl sample were

applied and separated (length 41 cm) using 1 x TBE as running buffer in a Long Red IR

DNA sequencer (Licor, Bad Homburg). In order to determine the length of the product and

the exact transcription start, four lanes of sequencing reaction were also loaded onto the

gel (Thermo Sequenase Primer Cycle, Amersham Biosciences, Freiburg). For these

reactions, the same oligonucleotides as for reverse transcription were used. In total, five

different primers were chosen to verify the results.

Annealing buffer:

50 mM Tris HCl pH 7.9, 1.25 M KCl

5 x RT buffer:

250 mM Tris HCl pH 8.3, 125 mM KCl, 15 mM MgCl2

RNaseA reaction mix:

10 mM Tris HCl pH 7.5, 1 mM EDTA, 100 mM NaCl, 100 µg ml-1 salmon sperm

(sonicated), 200 µg ml-1 RNase A (Qiagen, Hilden)

10 x TBE buffer:

890 mM Tris, 890 mM boric acid, 20 mM EDTA, pH 8.3


Long Ranger gel: 2.85 ml Long Ranger solution (50 %) (Cambrex, Bedford, USA)

13 g urea

3.1 ml 10 x TBE

310 µl DMSO

31 µl TEMED

217 µl 10 % (w/v) APS

3.6 Working with proteins

3.6.1 Protein purification

The GDH protein was purified after overexpression in C. glutamicum followed by a

combination of Ni2+ NTA affinity purification and size exclusion chromatography. The

C. glutamicum strain LNΔGDH carrying the expression vector pEKExgdh was cultivated as

described in section 3.2.4. After 2 hours of induction with 0.5 mM IPTG, 600 ml of culture

were harvested by centrifugation (4,000 xg, 10 min, 4°C). The pellets were solved in 3 ml

HisA buffer and transferred to a tube containing glass beads. Cells were disrupted by

vigorous shaking at 6.5 m s-1 for 30 seconds using a FastPrep FP120 instrument (Q-

BIOgene, Heidelberg). After two more cycles of disruption, cell debris and glass beads were

removed by centrifugation (10,000 xg, 30 min, 4°C). To assure that the resulting crude

extract was free of particulate material, it was filtered using Filtropur S 0.2 filter (Sarstedt

AG, Nümbrecht).

His-tagged GDH was subsequently purified using an Äkta prime with a 1 ml HighTrap

affinity column (GE Healthcare, Munich) according to the description of the manufacturer. In

order to achieve a rising imidazole concentration, HisB buffer was added gradually. Finally,

fractions that showed the highest protein concentration were applied onto an SDS gel

(3.6.3) and combined for size exclusion chromatography on Superdex G75 in 0.1 M Tris

HCl pH 8 using an Äkta FPLC system (Pharmacia Biotech, Munich).

HisA buffer pH (NaOH) 7.5:

300 mM NaCl, 50 mM NaH2PO4, 10 mM imidazole

HisB buffer pH (H3PO4) 7.5:

300 mM NaCl, 50 mM NaH2PO4, 500 mM imidazole


3.6.2 Quantification of proteins

Protein concentration was determined using the method described by Lowry (modified by

Dulley & Grieve, 1975). For a standard curve 2, 4, 12, 16, 20, 30, and 40 µg bovine serum

albumine (BSA) were used. To the protein sample 20 µl 100 mM SDS were added. After

addition of 1 ml fresh coloring reagent, samples were incubated for 5 minutes at 37°C.

100 µl of folin-ciocalteus phenol reagent (Merck, Darmstadt) were applied, samples were

mixed and incubated for another 20 minutes at 37°C. Subsequently, the absorbance at

650 nm was measured, so the protein concentration could be determined using the

standard curve.

Coloring reagent:

100 ml solution A (2 g Na2CO3, 0.4 g NaOH/100 ml)

+ 2 ml solution B (2 % (w/v) Na+K+ tartrate)

+ 2 ml solution C (1 % (w/v) CuSO4)

3.6.3 SDS polyacrylamide gel electrophoresis (PAGE)

The electrophoretic separation of proteins was performed in 14 % SDS polyacrylamide gels

(Schägger & von Jagow, 1987).

A separation gel was composed of:

10 ml acrylamide/bisacrylamide (37.5 %/1 %)

10 ml gel buffer

10.8 g urea

10 µl TEMED

100 µl 10 % (w/v) APS

The stacking gel was composed of:

1 ml acrylamide/bisacrylamide (37.5 %/1 %)

3.1 ml gel buffer

8 ml water

10 µl TEMED

100 µl 10 % (w/v) APS


Protein samples were mixed with loading buffer, incubated for 5 minutes at 65°C, and

loaded onto the gel together with protein marker (Prestained Protein Marker Broad Range,

10-180 kDa, MBI Fermentas, Wilna and peqGOLD Prestained Protein-Marker IV, peqlab

Biotechnologie GmbH, Erlangen). Gel electrophoresis was performed using a BlueVertical

101 apparatus (Serva Electrophoresis GmbH, Heidelberg) at 50 V for 30 min and

subsequently at 120 V. As buffers cathode and anode buffers were used.

Gel buffer:

3 M Tris, 1 M HCl, 0.3 % (w/v) SDS

5 x loading buffer:

20 % (w/v) SDS, 60 % (w/v) glycerol, 250 mM Tris, 10 % (v/v) mercaptoethanol, 0.01 %

(w/v) serva blue G, pH (HCl) 6.8

Cathode buffer:

0.1 M Tris, 0.1 M tricine, 0.1 % (w/v) SDS

Anode buffer:

0.2 M Tris, pH (HCl) 8.9

3.6.4 Staining with Coomassie Brilliant Blue

For staining with Coomassie Brilliant Blue, the polyacrylamide gel was first incubated in

staining solution at room temperature for at least 1 hour. After that, it was incubated

overnight in 10 % (v/v) acetic acid.

Staining solution:

45 % (v/v) methanol, 10 % (v/v) acetic acid, 0.1 % (w/v) Coomassie Brilliant Blue G-250

3.6.5 Western blotting

After SDS PAGE carried out as described in section 3.6.3, the gel-separated proteins were

transferred onto a polyvinylidene difluoride membrane (Millipore Immobilon P, Roth,

Karlsruhe) by electroblotting. For this, the membrane was first incubated in 60 % (v/v)

methanol and then in transfer buffer. It was put on filter paper soaked in transfer buffer. On

this, the SDS gel and another stack of filter paper were applied. Blotting was performed for

1 hour at 0.8 mA per cm2 of membrane. When transfer was completed, the membrane was


incubated in blocking buffer for at least 1 hour. GDH-specific anti-serum generated in

rabbits was added in a 1:10,000 dilution. After another hour, 3 washing steps, 20 minutes

each, were performed and anti-antibody coupled to alkaline phosphatase (Sigma-Aldrich,

Steinheim) was added diluted in blocking buffer (1:10,000). Washing steps were repeated

and detection was carried out using the BCIP/NBT alkaline phosphatase substrate (Roth,

Karlsruhe). For this, to 10 ml of detection buffer 65 µl of BCIP and NBT stock solution were

added, respectively. The reaction was stopped by the addition of H2O.

As a more sensitive method the ECL Western Blot plus kit (GE Healthcare, Munich) was

used as recommended by the supplier. The signal was detected on Amersham HyperfilmTM

MP (GE Healthcare, Munich).

Transfer buffer:

10 mM CAPS, 1.5 M NaCl, 10 % (v/v) methanol, pH (NaOH) 11

Washing buffer:

50 mM Tris, 0.15 M NaCl, pH (HCl) 7.5

Blocking buffer:

5 % (w/v) skim milk powder in washing buffer

NBT stock solution:

0.5 g 4-nitro blue tetrazolium chloride (NBT) were dissolved in 10 ml 70 % (v/v) dimethyl-

formamide and stored at -20°C.

BCIP stock solution:

0.5 g 5-bromo-4-chloro-3-indolyl phosphate toluidine salt (BCIP) were dissolved in 20 ml

100 % dimethylformamide and stored at -20 °C.

Detection buffer:

100 mM NaCl, 5 mM MgCl2, 100 mM Tris, pH (NaOH) 9.5


3.6.6 Determination of enzyme activity

3.6.6.1 GDH activity measurements

To measure GDH activity as described by Meers and Tempest (1970), a C. glutamicum

culture (OD600 = 4-5) was centrifuged (4,000 xg, 6 min, 4°C). The cell pellet was washed

with ice-cold potassium phosphate buffer (200 mM, pH 7.0). Subsequently, the cell pellet

was resuspended in 1.5 ml ice-cold potassium phosphate buffer (200 mM, pH 7.0). The cell

suspension was transferred to a tube containing glass beads and cells were disrupted by

vigorous shaking at 6.5 m s-1 for 30 seconds using a FastPrep FP120 instrument (Q-

BIOgene, Heidelberg). After repeating this procedure twice, cell debris and glass beads

were removed by centrifugation (10,000 xg, 30 min, 4°C). To measure the GDH activity of

this cell extract, the following reaction mixture was added to an UV-cuvette:

100 µl 1 M Tris HCl, pH 8.0

100 µl 2.5 mM NADPH

100 µl 200 mM NH4Cl

10-100 µl cell extract

ad 900 µl water

The reaction was started by the addition of 100 µl 100 mM 2-oxoglutarate and the initial

rate of absorbance decrease at 340 nm was measured using the V-560 UV/VIS-spectro-

photometer (JASCO, Gross-Umstadt).

In order to determine the maximal velocity of the deamination of glutamate by GDH, the

following reaction mixture was used as described by Shiio and Ozaki (1970):

100 µl 1 M Tris HCl, pH 8.8

100 µl 1.5 mM NADP+

10-100 µl cell extract

ad 900 µl water

This reaction was started by the addition of 100 µl 1 M glutamate and the initial increase of

absorbance at 340 nm was measured as described above.

In addition to that, the concentration of total protein in the cell extract was determined as

described in section 3.6.2. From the resulting data, the specific GDH activity was calculated

using the following equation:


dmtVAactivity⋅⋅⋅

⋅Δ=

ε

activity: specific GDH activity

ΔA: change of absorbance at 340 nm

V: total volume

ε: extinction coefficient (6.3 cm2 µmol-1)

t: reaction time

m: mass of total protein in the reaction mixture

d: layer thickness

3.6.6.2 Glutamyltransferase test

Besides its physiological reaction, the ATP-dependent synthesis of L-glutamine from

glutamate and ammonium, the glutamine synthetase can also catalyze the ADP-dependent

formation of glutamylhydroxamate from glutamine and hydroxylamine in the presence of

arsenic. In combination with Fe3+ ions, glutamylhydroxamate forms a brown-colored

complex, which can be detected spectrometrically. Under these conditions, the

physiological reaction is completely inhibited (Shapiro & Stadtman, 1970).

Cultivation was performed as described in section 3.2.4. 20 ml to 100 ml of culture were

harvested by centrifugation (4,000 xg, 5 min, 4°C), washed in disruption buffer and

resuspended in 700 µl disruption buffer. Cells were transferred into 2 ml tubes containing

approximately 300 mg glass beads (diameter 0.2-0.3 mm) and disrupted by vigorous

agitation (4 x 30 s, 6.5 m s-1) in a FASTPREP (FP120, QBIOGENE, Heidelberg). In

between, samples were chilled on ice for 3 minutes. After centrifugation (16,060 xg, 30 min,

4°C), the cell-free supernatant was used for determination of enzyme activity. For this,

380 µl test buffer, 20 µl 8 mM ADP solution and between 10 µl and 100 µl crude extract

were mixed and incubated at 30°C for 5 minutes. By adding 20 µl hydroxylamine solution

[360 mM], the reaction was started. After 10 minutes of incubation at 30°C, the reaction

was stopped with 1 ml stop solution. For each dilution of crude extract another sample as

reference was used, in which hydroxylamine was substituted by H2O. Measurements were

ADP

glutamine + NH2OH 3-glutamylhydroxamate + H2O arsenic


carried out at a wave length of 540 nm. The amount of the iron-γ-glutamylhydroxamate

complex formed was determined using a standard curve with 0 to 3 mM γ-

glutamylhydroxamate.

Disruption buffer:

2.5 mM MnCl2, 20 mM imidazole, pH (HCl) 7.2

Test buffer:

20 mM glutamine, 25 mM KHAsO4, 0.27 mM MnCl2, 135 mM imidazole, pH (HCl) 7.2

Stop solution:

11 g FeCl3 x 6H2O, 4 g trichloroacetic acid, 4.2 ml 37 % HCl in 200 ml

3.6.7 Determination of promoter activity

As a reporter system for quantification of promoter activity, the green fluorescent protein

(GFP) was used. Strains carrying plasmids expressing a UV-optimized GFP variant under

the control of the gdh promoter were cultivated as described in section 3.2.4. During

exponential growth (OD600= 4-5), a 10 ml sample was centrifuged (4,000 xg, 7 min, 4°C)

and resuspended in 1 x PBS. In order to test the influence of nitrogen starvation on the gdh

promoter activity, CgCoN medium containing 100 mM glutamine was inoculated with the

overnight culture to an OD600 of 1 and samples were also taken at about OD600 of 4. After

excitation at a wavelength of 395 nm, the emission between 490 nm and 520 nm was

detected. These measurements were carried out using the fluorimeter (Fluorolog-3

Instruments S.A., Inc, Edison, USA). Promoter strength was calculated by putting fluores-

cence in relation to the dry weight per ml. An OD600 of 1 corresponds to 0.36 mgdry weight per

ml (Weinand, 2004).

1 x PBS:

8 g/l NaCl, 3.58 g/l Na2HPO4 x 12H2O, 0.2 g/l KCl, 0.24 g/l K2HPO4, pH (HCl) 7.4

3.6.8 Gel shift assays and competition assays

By gel shift assays, binding of a transcriptional regulator to a distinct DNA fragment can be

analyzed. Unless provided, protein extract of E. coli heterologoulsy expressing the

transcriptional regulator to be investigated was prepared as follows. Respective E. coli

strains were cultivated in 250 ml LB medium at 37°C overnight. On the next day, cells were


harvested by centrifugation (4,000 xg, 15 min, 4°C). The cell pellet was resuspended in

TEN buffer (2 ml per g cell pellet weight). The cell suspension was transferred to tubes

containing glass beads and cells were disrupted by vigorous shaking twice at 6.5 m s-1 for

30 seconds using a FastPrep FP120 instrument (Q-BIOgene, Heidelberg). Cell debris and

glass beads were removed by centrifugation (13,000 xg, 30 min, 4°C). The protein extract

was stored at -80°C. The 200 bp target DNA for the gel shift assays was synthesized by

PCR (section 3.4.5) and purified by gel electrophoresis (section 3.4.2). For labeling of the

DNA and the setup of the reaction mixture for the gel shift assays, the DIG Gel Shift Kit

(Roche Diagnostics, Mannheim) was used following the supplier’s protocol. Separation by

gel electrophoresis was performed in native 6 % polyacrylamide gels (Anamed

Electrophorese GmbH, Darmstadt) using 0.5 x TBE buffer as running buffer. Subsequently,

the labeled DNA was blotted on a positively charged nylon membrane (Roche Diagnostics,

Mannheim) by electroblotting as described in the protocol of the DIG Gel Shift Kit (Roche

Diagnostics, Mannheim). Detection of the labeled DNA was performed as described in

section 3.5.4.

Competition assays were performed in accordance to the protocol of gel shift assays. To

each reaction, unlabeled, overlapping 50 bp DNA fragments were added, in order to

analyze their ability to inhibit a shift of the labeled target DNA.

TEN buffer:

10 mM Tris, 1 mM EDTA, 0.1 M NaCl, pH (HCl) 8.0

10 x TBE buffer:

890 mM Tris, 890 mM boric acid, 20 mM EDTA, pH (HCl) 8.0

Results 45

4 Results

GDH is the enzyme primarily responsible for assimilation of ammonium in C. glutamicum.

Especially, due to its function of providing glutamate, it was investigated concerning kinetic

parameters and substrate specificity decades ago (Kimura, 1962; Oshima et al., 1964; Shiio

& Ozaki, 1970; Shiio & Ujigawa, 1978). After the identification of the gdh gene, first studies

on transcriptional regulation were performed (Börmann et al., 1992). Also, nitrogen

metabolism in general was examined intensely (for overview, Burkovski, 2003a; 2003b;

2005; 2007). In addition to the TCA cycle as donor of carbon backbones, nitrogen

metabolism is of great importance as donor of amino groups for amino acid biosynthesis. In

the course of studying nitrogen control, contrary results about the influence of nitrogen

availability on regulation of GDH were obtained. More recent work on GDH and its impact

on nitrogen control could provide a vast amount of new details, but still failed to give a

coherent model describing regulatory mechanisms involved (Müller, 2005). Therefore,

additional data on this are needed. Furthermore, the connection of GDH not only to nitrogen

metabolism, but also to the central metabolism is in the focus of interest.

4.1 Purification and characterization of glutamate dehydrogenase

GDH is positioned at an important branch-point of metabolism connecting ammonium

assimilation and the TCA cycle. Furthermore, the reaction consumes more than 50 % of

NADPH available. The presence of this NADPH-dependent enzyme has a crucial influence

on the cellular generation of NADPH (Marx et al., 1999). Reductive equivalents next to

respective carbon backbones derived from the TCA cycle are important cofactors for lysine

biosynthesis. For these reasons, there is a strong interest in not only characterizing

enzymes directly involved in the TCA cycle, but also GDH. In the light of this, closer

investigation of the GDH enzyme is one aim so that derived data can be seen in a more

global context leaving behind studies that focused only on analyses of the gene/protein

itself.

4.1.1 Purification of GDH

In addition to established methods for determination of enzyme activity in crude extract,

Western blots are another method for quantification of enzyme level. In order to be able to

precisely quantify the amount of a certain protein present, the purified enzyme is used as

standard. As a prerequisite for the performance of quantitative Western blots a protocol

Results 46

needed to be established for purification of GDH. This purified enzyme can also be used for

biochemical characterization and studies on the impact of effector molecules on enzyme

kinetics. One of the most common approaches for protein purification, Ni2+ NTA affinity

purification using a commercially available vector expressing an N-terminal histidine (his)-

tag (Qiagen, Hilden), was chosen. By applying standard methods described for purification

after overexpression in E. coli, it was not possible to obtain purified and active GDH

enzyme. Even commonly performed changes in cultivation conditions such as variation in

temperature, IPTG concentration, induction time, and the use of different E. coli strains did

not bring about the desired results (Müller, 2005; data not shown). As a next approach,

C. glutamicum was chosen as host organism for the overexpression of his-tagged GDH.

The respective expression vector pEKExgdh was transformed into the gdh deletion strain

LNΔGDH and cultivation was performed as described in 3.2.4. In order to obtain protein of

high purity and to be able to choose buffer condition suitable for subsequent activity

measurements, a combination of Ni2+ NTA affinity purification and size exclusion

chromatography was applied. Crude extract was obtained from 600 ml of culture after

2 hours of induction with 0.5 mM IPTG. After Ni2+ NTA affinity purification, fractions

containing the highest protein concentration (figure 4.1 A) were combined for a second

round of size exclusion chromatography (figure 4.1 B).

Fig 4.1: Results of GDH purification after overexpression in C. glutamicum. 16 µl of fractions containing the highest protein amount after Ni2+ NTA affinity purification were applied (A). After size exclusion chromatography, the same volume of fractions with the highest protein concentration (lanes 2-8) was used (B). For comparison, lane 1 contained protein after Ni2+ NTA affinity purification.

By combining both purification methods, it was possible to obtain pure GDH protein.

Furthermore, activity measurements confirmed that GDH yielded by this approach was still

active. In the course of the measurements, it was also demonstrated that GDH had a high

substrate specificity and did not use diaminopimelic acid as substrate (data not shown) as

proposed by Oshima and coworkers (1964). After upscaling, this method of purification can

453525

15

55

70130170

130

45

3525

55

15

70A B1 2 3 4 1 2 3 4 5 6 7 8

453525

15

55

70130170

130

45

3525

55

15

70A B

453525

15

55

70130170

453525

15

55

70130170

453525

15

5555

7070130130170170

130130

45

3525

55

15

70

45

3525

5555

1515

7070A B1 2 3 4 1 2 3 4 5 6 7 8

Results 47

be used to obtain GDH in sufficient amounts for creating standard curves for quantitative

Western blots. Figure 4.2 shows different dilutions of purified GDH detected using the

highly sensitive ECL Western Blot plus kit (GE Healthcare, Munich).

By moving away from the standard organism for heterologous expression of proteins and

successfully applying a combination of two purification steps, active GDH protein was

yielded. First attempts of performing Western blots indicated that appropriate dilutions

resulted in a pattern of a well visible decrease in protein amount. Starting out from this

point, the foundation for future work of establishing an additional method for quantification

of GDH has been laid.

4.1.2 Characterization of GDH in lysine-producing strains

As described above, one approach to collect data on enzymes is purification in combination

with subsequent determination of kinetic parameters. On the other hand, enzymes can also

be analyzed in the crude extract of different strains under selected cultivation conditions.

Besides the wild type, genetically manipulated strains, as for instance well-defined lysine-

producing strains, are suitable for this approach. In addition to the biotechnological

relevance of these strains, another advantage is the detailed investigation concerning

metabolic fluxes. Comparison to the C. glutamicum wild type might lead to new insights into

metabolic changes in response to the altered flux towards lysine. Within this work, GDH

activity, protein level, and gdh transcription were determined in strains with mutated

aspartatokinase (encoded by lysC) and pyruvate carboxylase (encoded by pyc).

Aspartatokinase is the key enzyme within the biosynthetic pathways for lysine, threonine,

and isoleucine and therefore its regulation has been investigated for decades (Shiio &

Miyajima, 1969; Kalinowski et al., 1990). Pyruvate carboxylase reaction provides

oxaloacetate as precursor for the synthesis of amino acids derived from aspartate by

catalyzing the carboxylation of pyruvate (Peters-Wendisch et al., 1998). In the following

experiments, strains with single mutations and a combination of mutations were used. The

strain DM1868 expressed a deregulated aspartatokinase (lysCT311I), in which lysine- and

threonine-mediated feedback inhibition was released up to concentrations of 10 mM

1 2 3 4 51 2 3 4 5 Fig. 4.2: Western blot with different dilutions of purified GDH. 8 µl of the purified protein (1) and the following dilutions were applied: 1:2 (2), 1:5 (3) and 1:10 (4), and 1:100 (5). Detection was performed using the ECL Western Blot plus kit (GE Healthcare, Munich).

Results 48

(Ohnishi et al., 2002). DM 1799 had a mutated pyruvate carboxylase (pycP458S), which

probably showed an improved catalytic ability due to a more flexible conformation caused

by the exchange of proline to serine (Ohnishi et al., 2002). Strain DM1800 carried both

mutations. For DM1868 and DM1800 enhanced lysine formation was reported (Degussa

GmbH, Halle). In the next figure and table, the results concerning the investigation of GDH

are displayed.

Tab. 4.1: Determination of GDH activity. Enzyme activity was determined in the wild type and lysine-producing strains by monitoring changes in NADPH concentration at 340 nm. Results are from two independent experiments.

Strain Activity

[µmol mgProtein-1 min-1]

ATCC13032 2.61

DM1132 2.66

DM1799 2.25

DM1800 2.75

DM1868 2.35

Fig. 4.3: Detection of GDH protein and gdh transcript. The amount of GDH was determined by Western blots using cell extract obtained from the wild type and lysine-producing strains. 10 µg of total protein were applied (A). To determine the amount of gdh transcript in the respective strains, 1 µg total RNA was used for Dot blot experiments (B).

The results of the activity measurements showed that there were basically no differences in

GDH activity in the lysine-producing strains and the wild type. Western blot experiments

supported these data, since the amount of GDH was comparable in all strains. Also, gdh

transcription was not altered considerably compared to the wild type. Put together,

ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

72

56

43

3425

ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

A B

ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

72

56

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ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

72

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ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

A B

ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

ATC

C13

032

DM

1132

DM

1799

DM

1800

DM

1868

A B

Results 49

metabolic perturbations due to enhanced lysine formation did not show any great influence

on the GDH activity under the tested conditions.

4.1.3 Gradual expression of gdh

Even though it has been demonstrated that alterations of metabolism resulting in lysine

formation did not significantly influence the GDH reaction, it might be the other way around,

meaning that different levels of GDH activity lead to metabolic changes in favor of or

against lysine formation. Hints going in this direction were obtained by Marx and coworkers

(1999). After deletion of the gdh gene in the C. glutamicum lysine-producing strain MH20-

22B, plasmid-encoded NADPH-dependent GDH from C. glutamicum, as well as an NADH-

dependent GDH from Peptostreptococcus asaccharolyticus, was expressed. The exchange

of the NADPH-consuming enzyme led to a decrease in cellular NADPH demand, which

then resulted in a drastically reduced generation of the reductive equivalent. These results

indicated the flexibility of the cellular responses to metabolic changes.

In order to be able to monitor the effect of slight alterations of enzyme activity on the

metabolism, one goal was to establish a method that would allow differential expression of

a respective enzyme, in this case GDH. The advantage compared to a deletion or an

exchange would be the possibility of fine-tuning expression and/or enzyme activity to

monitor putative effects on the metabolism. The easiest way of achieving different levels of

enzyme activity is to change the expression level in order to get different amounts of

protein. This can either be done by the more complex approach of manipulating the

promoter region or by using a plasmid encoding the desired protein. Since pEKExgdh had

been constructed for purification, it was tested, whether it would be possible to use the

vector for expressing GDH at different levels. For this, the strain LNΔGDH carrying the

respective plasmid was grown in CgC medium and when an OD600 of about 4 was reached,

induction using different concentrations of IPTG was performed. After one and two hours,

samples were taken for determination of enzyme activity and Western blots.

Results 50

Tab. 4.2: Determination of GDH activity after gradual expression. Expression in LNΔGDH carrying pEKExgdh was induced with IPTG concentrations described below. The wild type was used as control. Induction was performed for one (A) and two hours (B) and GDH activity was determined by monitoring the decrease in NADPH concentration at 340 nm.

Strain IPTG-concentration

[mM]

Activity [µmol mgProtein

-1 min-1]after 1 h

Activity [µmol mgProtein

-1 min-1]after 2 h

Wild type 0 2.32 1.90

LNΔGDH pEKEx2gdh 0 0.19 0.17

LNΔGDH pEKEx2gdh 0.005 0.34 0.48





Fig. 4.4: Expression of gdh at different levels. Expression in LNΔGDH carrying pEKExgdh was induced with IPTG concentrations described above. The wild type was used as control. Induction was performed for one (A) and two hours (B) and 20 µg of total protein were used. The results of Western blots and activity measurements showed that pEKExgdh was

suitable for expressing GDH at different levels. Within the course of these experiments, also

first hints were obtained suggesting a connection between the presence of GDH and lysine

formation. When comparing the external concentration of lysine secreted by the wild type,

LNΔGDH pEKExgdh overexpressing gdh, and LNΔGDH carrying pEKEx, significantly less

lysine was detected in the strain lacking a functional GDH (data kindly provided by A. Wirtz,

University of Cologne). Of course, this interesting effect needs to be further investigated,

but it confirmed that studies on the expression level of gdh might be a useful tool to explore

72

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Wild

type

0 m

M

0.00

5 m

M

0.01

mM

0.05

mM

0.1

mM

0.5

mM

IPTG

72

56

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Wild

type

0 m

M

0.00

5 m

M

0.01

mM

0.05

mM

0.1

mM

0.5

mM

IPTG

A B

72

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Wild

type

0 m

M

0.00

5 m

M

0.01

mM

0.05

mM

0.1

mM

0.5

mM

IPTG

72

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type

0 m

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5 m

M

0.01

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0.05

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0.1

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0.5

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IPTG

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0 m

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5 m

M

0.01

mM

0.05

mM

0.1

mM

0.5

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IPTG

72

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0 m

M

0.00

5 m

M

0.01

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0.05

mM

0.1

mM

0.5

mM

IPTG

A B

Results 51

a putative influence on the metabolism. Using pEKExgdh differential expression was

achieved successfully. However, it seemed that the tac promoter of pEKEx was leaky and

therefore not fully repressed in the absence of IPTG. This led to a very low basal enzyme

activity as well as to a weak signal in Western blot experiments even in the absence of

IPTG. Another disadvantage of expressing a plasmid-encoded protein at different levels is

the fact that cultures carrying the respective plasmid are never homogenous. Therefore,

only the overall effect of a whole culture can be detected. In order to avoid this problem, the

focus has to be put on the respective promoter region. By bringing in mutations into the

genome directly, it is possible to circumvent this problem. As a prerequisite, it is necessary

to test the effect of respective mutations in reporter gene assays with promoter fusions.

4.2 Transcriptional regulation of gdh About regulation of gdh, controversial data have been presented in the past. While some

studies reported constant level of activity and transcript irrespective of the nitrogen supply

(Tesch et al., 1999; Beckers et al., 2005; Silberbach et al., 2005a), others suggested an

enhanced activity, protein level, and transcript level under nitrogen deficiency (Müller, 2005;

L. Nolden, unpublished results). So far, regulation of bacterial GDH enzymes is believed to

be on the level of transcription, since post-translational modification has not been reported

(Minambres et al., 2000). Identification of putative transcriptional regulators of

C. glutamicum gdh did not contribute to solve the controversy about mechanisms

controlling expression (Müller, 2005). Therefore, emphasis was put on the direct

characterization of the gdh promoter region, hoping to get further information on regulation

of transcription that would contribute to explain processes involved. Information on the

promoter structure and components of this region might be used, as described above, for

the construction of strains expressing gdh at different levels.

4.2.1 Mutational analyses of the gdh promoter region

Within the last decade, C. glutamicum promoter structures have been increasingly in the

focus of attention. Based on the sequence of 53 experimentally determined promoters,

consensus sequences for -35 and an extended -10 region of the σA-dependent house-

keeping promoter have been identified. The -35 region in C. glutamicum is conserved at a

very low level and it seems that especially in these cases the extended -10 region plays a

more important role in interacting with RNA polymerase. The consensus sequence of the

extended -10 region is as follows: TgtG(c/g)TAtAATGG. Capital letters indicate a

Results 52

GACACTGCCATAATTGAACGTGAGCA-10

Wild type:

GACACCCCCATAATTGAACGTGAGCAMutant 3:

*

Mutant 1: GACACTGCTATAATTGAACGTGAGCA*

GACACTGCCATAAATGAAAGTGAGCAMutant 2: *

*

GACACCCCCATAAATGAACGTGAGCAMutant 4: *

GACACTGCCATAATTGAACGTGAGCA-10

Wild type:

GACACCCCCATAATTGAACGTGAGCAMutant 3:

*

Mutant 1: GACACTGCTATAATTGAACGTGAGCA*

GACACTGCCATAAATGAAAGTGAGCAMutant 2: *

*

GACACCCCCATAAATGAACGTGAGCAMutant 4: *

conservation of more than 40 % (Pátek, 2005). An additional component in some promoters

is the AT-rich region, which is located upstream of the -35 region and might induce

activation by DNA bending or act as an UP element as described for E. coli promoters

(Pátek et al., 2003; Browning & Busby, 2004).

The gdh promoter has been investigated previously with respect to transcription start and

putative core promoter regions (Börmann et al., 1992). Transcription initiates 284 bp

upstream of the start codon, which results in a long 5’ UTR. Putative -10 and -35 regions

have been determined by sequence analyses due to a high similarity to the consensus

sequence of housekeeping promoters, but have not been verified by experimental

approaches. Consequently, the first experimental set-up for the characterization of the gdh

promoter was to confirm these predictions by introducing specific mutations into the

promoter region. Information gained from these experiments might be suitable for creating

promoters of different strength to make gradual expression of gdh possible. Based on the

acquired knowledge on differently conserved nucleotides, mutations were introduced in the

gdh promoter region that would putatively lead to an increase or decrease in promoter

strength. The effect of the respective mutations was determined in reporter gene assays

using the plasmid pEPR1, which harbors a promoterless gfpuv gene (Knoppová et al.,

2007; kindly provided by M. Pátek, Academy of Sciences of the Czech Republic, Prague). A

527 bp fragment spanning the gdh upstream region including the wild type promoter was

cloned upstream of the gfpuv gene in pEPR1. The same was done for 527 bp fragments

carrying mutated promoters. In figure 4.5, the sequence of the gdh wild type core promoter

is displayed. Based on this sequence the respective mutations marked in red were

introduced.

Fig. 4.5: Sequence of the gdh core promoter region. Transcription initiates 284 bp upstream of the start codon (*). The extended -10 region (bold) was identified by sequence comparison (Börmann et al., 1992). Based on the wild type promoter, mutations marked in red were introduced in the promoter.

Results 53

0

200x103

800x103

1x106

control mutant1

wt mutant3

mutant4

mutant2

Cps

(mg d

w/m

l)-1

0

200x103

800x103

1x106

control mutant1

wt mutant3

mutant4

mutant2

0

200x103

800x103

1x106

control mutant1

wt mutant3

mutant4

mutant2

Cps

(mg d

w/m

l)-1

Examination of the respective sequence showed that the gdh promoter has a high similarity

with the consensus sequence. However, to possibly increase promoter strength the

cytosine (C) at the first position of the -10 region was exchanged to thymine (T), so that the

-10 region matched the consensus sequence (mutant 1). This T at the first position of the

-10 region was shown to be conserved in 80 % of 44 promoters examined (Vašicová et al.,

1999). In the other three mutants, conserved nucleotides were substituted to presumably

decrease promoter activity. The highly conserved T at the end of the putative -10 region,

which was shown to be conserved in 91 % of 44 promoters (Vašicová et al., 1999), was

exchanged. Also, one C located closer to the transcription start was mutated to adenosine

(A) (mutant 2). In mutant 3, T and guanine (G) upstream of the -10 region were both

substituted to C. A combination of these selected mutations was introduced in the promoter

region designated mutant 4. These were expected to inactivate the promoter and therefore

confirm the predicted -10 region (M. Pátek, personal communication). Wild type cultures

carrying the plasmids harboring the GFP promoter fusions described above were grown in

minimal medium until exponential phase (OD600 4 to 5). Samples were taken and after

excitation at 395 nm the emission spectrum was measured. Maximal fluorescence was then

put into relation to the dry weight per ml of the cultures. The next figure summarizes results

of determination of gdh promoter strength and the impact of the respective mutations on

activity of the gdh promoter. As negative control cultures carrying pEPR1 without promoter

were used.

Fig. 4.6: Determination of gdh promoter activity. Activity of the wild type promoter (wt) was compared to the activity of mutated promoter regions. Sequences of the respective promoters are displayed in figure 4.5. C. glutamicum wild type carrying pEPR1 without promoter (control) and fragments of the gdh promoter were cultivated until exponential phase. After excitation at 395 nm an emission spectrum ranging from 490 nm to 520 nm was monitored. Maximal fluorescence was put into relation to mgdry weight per ml.

Results 54

Data obtained from measurements of fluorescence indicated that the putative -10 region

indeed resembles the actual -10 region. The exchange of nucleotides led to drastic

differences in promoter activity. As shown for mutant 1, a sequence identical to the

consensus sequence of the C. glutamicum -10 region resulted in a significant increase in

promoter strength. On the other hand, promoter activity was decreased by exchange of

conserved nucleotides within the extended -10 region. Exchange of T to A and C to A in

mutant 2 led to significantly less promoter activity. Similar decrease in activity was observed

after exchange of the 5’ extension of the -10 region to CC in mutant 3. The combination of

mutations (mutant 4) did cause a lower activity, but residual activity of 38 % compared to

the wild type promoter was still detectable. Besides verification of the -10 region, this

approach could prove that mutagenesis of the promoter region is a useful tool to achieve

gradual expression of a respective gene. It was possible to enhance promoter activity as

well as to reduce the level of transcription from the gdh promoter. Integration of these kinds

of mutations into the genome would make expression at different levels easy to handle

compared to plasmid-encoded expression described in chapter 4.1.3.

However, the question arising from this result was why the exchange of nucleotides in

mutant 4 only resulted in a partial decrease of promoter activity instead of complete

inactivation. Transcription despite a mutated -10 region has been observed before for other

promoters. As reason for the residual promoter activity DNA bending due to the flexibility of

the AT-rich region or interaction of the so-called UP element with RNA polymerase has

been proposed. Residual activity might also have been due to interaction of -35 region with

RNA polymerase or the presence of an additional less optimal -10 region. Another

possibility was the presence of a second start of transcription. The chance that RNA

polymerase recognized the mutated -10 region was almost negligible. However, it is hard to

predict which element of the promoter actually contributes to which extend to the activity

(Browning & Busby, 2004). In order to find out about the mechanisms involved in

transcription, despite a presumably inactive core promoter, the same combination of

mutations was introduced into a 398 bp and a 298 bp fragment of the gdh promoter. While

the 398 bp fragment still included the region upstream of the -35 region, the 298 fragment

only contained the mutated core promoter. As control the 527 bp and the 398 bp fragment

without mutations were included in the experimental set-up.

Results 55

Fig. 4.7: Determination of residual promoter activity. Activity of the wild type promoter (527 bp and 398 bp fragments, respectively) was compared to activity of promoters with a mutated (*) extended -10 region (GACACCCCCATAAATGAACGTGAGCA, respective mutations are underlined, -10 region and transcription start are marked in bold). Fragments including the mutation spanned 527 bp, 398 bp, and 298 bp of the gdh upstream region. C. glutamicum wild type carrying pEPR1 without promoter (control) and pEPR1 including fragments of the gdh promoter (527 and 398) were cultivated until exponential phase. After excitation at 395 nm, an emission spectrum ranging from 490 nm to 520 nm was monitored. Maximal fluorescence was put into relation to mgdry weight per ml.

Fragment size did not seem to have an impact on residual promoter activity as can be seen

in figure 4.7. The promoter activity was at a comparable level for all three fragments

including the mutations. The 298 bp fragment only contained the core promoter. For this

reason, transcription as a result of interaction of RNA polymerase with a putative UP

element or DNA bending was unlikely. Therefore, other mechanisms causing residual

activity had to be taken into consideration. However, some seemed to be more likely than

others. Recognition of the mutated promoter resulting in such a high level of activity was a

doubtful explanation. Interaction of RNA polymerase with the -35 region might have been

possible, but also not very likely since the -35 region is less conserved and probably

marginal in C. glutamicum (Pátek et al., 2003). Therefore, the presence of at least one

additional promoter had to be taken into account. This not yet identified promoter was

thought to be located in the vicinity of the known promoter or downstream of the

transcriptional start site identified by Börmann and coworkers (1992). In order to prove this

hypothesis, fragments of the promoter region ranging between 527 bp and 172 bp were

used for the construction of additional GFP promoter fusions to determine promoter activity

by measuring fluorescence. The following figure gives a schematic overview of the

fragments used in the subsequent measurements.

0

50x103

100x103

150x103

200x103

250x103

300x103

control 527 527* 398 398* 298*

Cps

(mg d

w/m

l)-1

0

50x103

100x103

150x103

200x103

250x103

300x103

control 527 527* 398 398* 298*0

50x103

100x103

150x103

200x103

250x103

300x103

control 527 527* 398 398* 298*

Cps

(mg d

w/m

l)-1

Results 56

Fig. 4.8: Schematic presentation of the fragments used in reporter gene assays harboring the gdh promoter region. Fragments ranging between 527 bp and 172 bp were cloned in pEPR1 to determine the influence of fragment size on the promoter strength. The ribosomal binding site was not included in any fragment.

As mentioned above, the longest fragment spanned 527 bp of the promoter region. It

included all regulator binding sites identified within the gdh upstream region, namely a FarR

binding site as well as both AmtR binding sites (Müller, 2005). The 398 bp fragment lacked

the FarR binding site. The 313 bp fragment began just upstream of the -35 region. That

means, it did neither contain the AT-rich upstream region nor the low affinity AmtR binding

site. The 276 bp fragment did not include the known promoter and transcriptional start site,

but harbored the high affinity AmtR binding site. The smallest, 172 bp fragment, did neither

contain any of the identified regulator binding sites nor the transcriptional start site.

Determination of promoter activity was carried out in the wild type as described previously.

This time, as positive control, gfpuv under the control of the strong p45 promoter (Pátek et

al., 1996) was additionally included.

527 bp

gdh

FarR AmtR AmtR

-35 -10

gdh

AmtR AmtR

-35 -10

gdh

AmtR

-35 -10

gdh

398 bp

313 bp

172 bp

gdh

AmtR

276 bp

527 bp

gdh

FarR AmtR AmtR

-35 -10

gdh

AmtR AmtR

-35 -10

gdh

AmtR

-35 -10

gdh

398 bp

313 bp

172 bp

gdh

AmtR

276 bp

Results 57

0

100x103

200x103

2x106

control p45 527 398 313 276 172

Cps

(mg d

w/m

l)-1

0

100x103

200x103

2x106

control p45 527 398 313 276 172control p45 527 398 313 276 172

Cps

(mg d

w/m

l)-1

Fig. 4.9: Determination of gdh promoter strength. Activity of the wild type promoter was determined depending on the length of the respective fragments. For a detailed description refer to figure 4.8. C. glutamicum wild type carrying pEPR1 without promoter (control) and different fragments of the gdh promoter were cultivated until exponential phase. As positive control pEPR1 carrying the p45 promoter was used. After excitation at 395 nm, an emission spectrum ranging from 490 nm to 520 nm was monitored. Maximal fluorescence was put into relation to mgdry weight per ml.

For promoter fusions with fragments of the gdh upstream region ranging between 527 bp

and 172 bp, a correlation between promoter activity and fragment size became obvious.

While promoter activity of the three longest fragments was at a comparable level, it

decreased significantly for the two shortest fragments. These results indicated that the

region upstream of the core promoter did not contribute significantly to the activity of the

gdh promoter. However, the residual activity observed for the 276 bp fragment was only

reduced to 44 % compared to the activity determined for the 527 pb fragment and lay within

the same order of magnitude as measured for the fragments with the mutated extended -10

region (figure 4.7). This result excluded the possibility of residual activity solely due to

interaction of RNA polymerase with the -35 region or the mutated -10 region. Instead, these

facts led to the assumption that the relatively high residual activity was the result of a

second promoter downstream of the identified transcription start within the 276 bp fragment.

4.2.2 Determination of the transcription start

Since results on the characterization of the gdh promoter indicated the presence of an

additional, so far not identified transcription start, Northern blot experiments were carried

out to determine size and number of gdh transcripts. Previous results demonstrated that

one transcript with a size of 1.65 kb was present, which is consistent with the transcriptional

start site located 284 bp upstream of the start codon (Börmann et al., 1992). The presence

Results 58

of one transcript could be confirmed by repeating Northern blot experiments (data not

shown). However, due to the experimental limits of this method, only transcripts significantly

different in size can be detected. Minor variations in fragment size cannot be resolved. To

be able to precisely determine the gdh transcription start, primer extension experiments

were carried out. For these, RNA obtained after cultivation under the following conditions

was used: standard conditions, 1 hour of nitrogen starvation, 1 hour of carbon starvation,

1 hour of cold shock at 15°C, and after 8 hours of cultivation. For most of these conditions,

changes in gdh transcription were reported (Müller, 2005). The reason for the application of

RNA obtained after different cultivation conditions was the possibility that transcription from

a distinct promoter might be only induced upon a certain stress conditions. That means, the

resulting transcripts would not be detectable under standard conditions.

Fig. 4.10: Expression analysis and determination of gdh transcription start. For primer extension experiments, 15 µg total RNA were used obtained from cultures grown under ammonium surplus (+N), 1 hour of ammonium deprivation (-N), 1 hour of carbon deprivation (-C), cold shock at 15°C for 1 hour, and at the late exponential phase (8 h). The transcriptional start sites, labeled by asterisks, were determined using IRD800-labeled oligonucleotides. Those were also used for sequencing reactions of the PCR product spanning the investigated region upstream of gdh.

Primer extension experiments shown in figure 4.10 were carried out using different primers

which bound in the upstream region of the gdh transcript. It was possible to verify the

transcriptional start site determined by Börmann and coworkers (1992). Based on the

results of the recent sequencing reactions, transcription started at G 285 bp upstream of the

start codon. As can be seen in figure 4.10 A, there might also be a second start of

transcription at T 286 bp upstream of the translational start. However, this result needs to

be verified using additional primers. Overlapping core promoter regions and transcripts

starting from nucleotides located closely to each other might indicate transcription initiating

from promoters recognized by different sigma factors in addition to the housekeeping sigma

factor σA (for overview, Pátek & Nešvera, 2008). Whether in the case of the gdh promoter

two adjacent transcriptional start sites are present or another promoter besides the

A T C G

CTTGCACTCGT

*

+N -N -C 15°C 8 h A C G T

AATTGGCAATG

*

+N -N -C 15°C 8 h

A B

A T C G

CTTGCACTCGT

*

+N -N -C 15°C 8 h A C G T

AATTGGCAATG

*

+N -N -C 15°C 8 hA C G T

AATTGGCAATG

*

+N -N -C 15°C 8 h

A B

Results 59

GTGGCCAGGT TATATAACCA GTCAGTCAAC TGGTCTCATT CGCTGGTCGG

ATGAATTTAA TTAAAGAAGA GACTTCATGC AGTTACCGCG CGTTTTGGCG

ATACAAAATT GATAAACCTA AAGAAATTTT CAAACAATTT TAATTCTTTG

TGGTCATATC TGTGCGACAC TGCCATAATT GAACGTGAGC ATTTACCAGC

CTAAATGCCC GCAGTGAGTT AAGTCTCAAA GCAAGAAGTT GCTCTTTAGG

GCATCCGTAG TTTAAAACTA TTAACCGTTA GGTATGACAA GCCGGTTGAT

-N120-TACAC ACTTTAAAAT CGTGCGCGCA TGCAGCCGAG ATGGGAACGA

GGAAATCATG

AmtR

AmtR

FarR

*

*-10-35

+1

1

61

121

181

241

301

361

526

-35








GGAAATCATG

AmtR

AmtR

FarR

*

*-10-35

+1

1

61

121

181

241

301

361

526

-35

housekeeping promoter was used, needs to be further elucidated and will be addressed in

one of the following sections.

Downstream of the known transcription start another transcriptional start site was identified

at G 195 bp upstream of the gdh gene. Interestingly, it is positioned within the high affinity

AmtR binding site 189 bp to 208 bp upstream of the gdh gene. An overview of the gdh

upstream region showing the identified transcriptional start sites is given in figure 4.11. The

promoter regions with the putative -35 regions and the experimentally proven -10 region as

well as the determined FarR and AmtR binding sites are included.

Fig. 4.11: Overview of the gdh promoter region. Transcriptional start sites indicated by asterisks are located 285 bp and 195 bp upstream of the start codon (284 bp and 194 bp, respectively, according to the sequence published by Kalinowski et al., 2003). -10 region and putative -35 regions are marked in bold letters and regulator binding sites are indicated as well. The start codon of the gdh gene is underlined.

While sequence analyses allowed the determination of the core promoter located at the

upstream transcriptional start site, it was complicated to predict promoter elements in the

vicinity of the newly identified downstream transcriptional start site. A putative -35 region

with the sequence TTGCTC matched the low conserved consensus sequence determined

for C. glutamicum (Pátek et al., 2003; Pátek, 2005). Prediction of a putative -10 region of

the housekeeping promoter solely based on the sequence was difficult to make in this

region. The most probable, but very weak hexamers would be the sequences AAAACT and

TAAAAC (M. Pátek, personal communication). The first hexamer showed little agreement

with the published consensus sequence, but the T at the last position tends to be highly

Results 60

conserved within the C. glutamicum -10 region (Vašicová et al., 1999). The second putative

hexamer shared four conserved positions with the consensus sequence. Based on

sequence analyses, no conclusions could be drawn about a putative housekeeping

promoter upstream of the newly identified transcription start, even though transcription

initiated there under standard conditions during exponential growth as expected for σA-

dependent promoters.

The results of the primer extension experiments also displayed the known changes in

transcript level in response to different environmental conditions. Transcription of gdh was

induced upon nitrogen starvation and repressed as a result of carbon deprivation. This

transcription pattern was not that obvious for transcripts starting at G285. On the other

hand, drastic changes in response to nitrogen and carbon deficiency could be observed for

the amount of transcript starting at G195.

Regulation of gdh expression in response to different environmental stimuli is necessarily

not only controlled by binding of transcriptional regulators. Also, alternative sigma factors

could be involved. Especially, the lack of an obvious σA-dependent housekeeping promoter

gave rise to this assumption. However, transcriptional regulation by sigma factors will be a

topic of one of the following sections. The fact that the transcriptional start site G195 is

located within an AmtR binding site led to the reinvestigation of the controversially

discussed topic of nitrogen control and its impact on gdh transcription.

4.2.3 Nitrogen-dependent transcription

For AmtR-regulated genes with a determined transcription start, namely amtA and the

gltBD operon, transcriptional start sites have been identified within AmtR binding motifs

(Jakoby et al., 2000; Beckers et al., 2001; Beckers et al., 2005). It seems that this

organization is somehow characteristic for AmtR-regulated genes. The gdh gene is not

regarded as part of the AmtR regulon. Despite an obvious upregulation of transcription

under nitrogen deprivation shown in Dot blot experiments, results could not be verified by

DNA microarrays (Müller, 2005; Beckers et al., 2005; Silberbach et al., 2005a) leaving the

question on regulatory mechanisms of gdh transcription unanswered. The discovery of an

apparently common organization of nitrogen-dependent promoters in C. glutamicum led to

the reinvestigation of nitrogen-dependent gdh transcription.

Results 61

4.2.3.1 Function of AmtR

Applying a novel experimental set-up, gdh transcription depending on the nitrogen

availability was monitored by GFP promoter fusions including different fragments of the gdh

promoter region. The longest fragment used in this set-up (398 bp) harbored both AmtR

binding sites as well as both promoters. The 276 bp fragment only contained the high

affinity AmtR binding site and the transcription start located closer to the gdh gene (G195).

For detailed information on the fragments used, refer to figure 4.8. Wild type cultures

carrying the respective plasmids were cultivated as described in section 3.2.4. After

overnight growth, fresh CgC medium was inoculated to an OD600 of 1. In order to induce

nitrogen starvation, CgCoN medium was used supplemented with 100 mM glutamine as

sole nitrogen source. Samples for measurements of promoter activity were taken during

exponential growth at OD600 of about 4. To check the integrity of the nitrogen starvation

response in the presence of the pEPR1 plasmids, Dot blots with an amtA probe were

carried out in parallel. In the course of the experiments on nitrogen-dependent gdh

transcription, a possible function of AmtR in the regulatory process was reinvestigated as

well. For this, pEPR1 harboring a 527 bp fragment of the gdh promoter region was

transformed into the amtR deletion strain MJ6-18 and cultivation was carried out as

described above. Results of the measurements are summarized in the following figure.

Results 62

0

100x103

200x103

300x103

400x103

500x103

600x103

control 398 276 172

Cps

(mg d

w/m

l)-1

amtA

+N -N +N -N +N -N +N -N

0

100x103

200x103

300x103

400x103

500x103

600x103

control 398 276 172

Cps

(mg d

w/m

l)-1

0

100x103

200x103

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control 398 276 172

Cps

(mg d

w/m

l)-1

amtA

+N -N+N -N +N -N+N -N +N -N+N -N +N -N+N -N

Fig. 4.12: Investigation of nitrogen-dependent gdh transcription in the wild type and the amtR deletion strain MJ6-18. Activity of the promoter was determined depending on the nitrogen source. The C. glutamicum wild type (A) and the amtR deletion strain MJ6-18 (B) carrying pEPR1 without promoter (control) and different fragments of the gdh promoter were cultivated until exponential phase. For a detailed description of the fragments refer to figure 4.8. Either ammonium (black bars) or glutamine (100 mM, grey bars) was used as nitrogen source. After excitation at 395 nm, an emission spectrum ranging from 490 nm to 520 nm was monitored. Maximal fluorescence was put into relation to mgdry weight per ml. In addition to that, Dot blots with an amtA probe containing 2.5 µg total RNA were performed to check the functionality of the nitrogen starvation response.

leer 5270

50x103

400x103

500x103

control 527

Cps

(mg d

w/m

l)-1

+N -N +N -N

amtA

leer 5270

50x103

400x103

500x103

control 527

Cps

(mg d

w/m

l)-1

+N -N +N -N

leer 5270

50x103

400x103

500x103

control 527

Cps

(mg d

w/m

l)-1

+N -N+N -N +N -N+N -N

amtA

A

B

Results 63

Dot blot experiments shown in figure 4.12 indicated that wild type cultures grown in the

presence of glutamine as sole nitrogen source turned on the nitrogen starvation response.

Accompanied by enhanced expression of the AmtR-regulated amtA gene, gdh transcription

was upregulated under these conditions as well. Furthermore, Dot blots proved that AmtR

was not titrated out by the additional, plasmid-encoded binding sites located in the gdh

promoter region. If that had been the case, amtA transcription would have been detectable

in the wild type even under sufficient nitrogen supply. Determination of gdh promoter

activity in a wild type background therefore clearly underlined results on nitrogen-dependent

transcription obtained by Dot blot experiments (Müller, 2005). The 172 bp fragment without

AmtR binding sites only showed little promoter activity and almost no increase in activity

upon nitrogen deprivation. For the 398 bp fragment including two AmtR binding sites, a 2.4-

fold enhancement of transcription was observed upon nitrogen starvation. A 2.5-fold

increase in promoter activity was detected for the 276 bp fragment indicating upregulation

even in the presence of only one AmtR binding site. The expression of gdh in the amtR

deletion strain MJ6-18 was monitored as well. It allowed drawing conclusions about a

possible participation of AmtR in regulating gdh transcription. As expected, Dot blot

confirmed that in the absence of AmtR, amtA was transcribed even under nitrogen surplus.

In accordance to this, gdh transcription was at a higher level even in the presence of

nitrogen and did not increase significantly upon nitrogen deficiency.

Put together, the results in figure 4.12 confirm the nitrogen-dependent regulation of gdh

transcription, which in the past has been reported contradictorily. For the first time, it was

possible to show that AmtR is necessary for repression of gdh transcription under nitrogen

surplus. To support the data obtained and investigate the influence of AmtR on gdh

expression, further measurements were performed with GFP promoter fusions. The

constructs spanning 527 bp and 276 bp of the gdh upstream region were used for

measurements in a wild type background and background of MJ6-18 under standard

conditions in CgC medium. By comparing promoter activity in the two strains, the general

capacity of AmtR to repress gdh transcription in the presence of sufficient nitrogen could be

analyzed.

Results 64

0

100x103

200x103

300x103

400x103

500x103

control 527 276

Cps

(mg d

w/m

l)-1

0

100x103

200x103

300x103

400x103

500x103

control 527 276

Cps

(mg d

w/m

l)-1

Fig. 4.13: Impact of AmtR on gdh transcription. Activity of the gdh promoter was determined in the C. glutamicum wild type (black bars) and the amtR deletion strain MJ6-18 (grey bars) carrying pEPR1 without promoter (control) and two different fragments of the promoter region. After excitation at 395 nm, an emission spectrum ranging from 490 nm to 520 nm was monitored. Maximal fluorescence was put into relation to mgdry weight per ml.

The comparison of promoter activity in the wild type and MJ6-18 revealed that expression in

the deletion mutant was significantly higher for both fragments used. This indicated that

AmtR repressed gdh transcription under nitrogen surplus and that this AmtR-mediated

repression of gdh was released even in the presence of sufficient nitrogen in the deletion

mutant. Interestingly, the promoter activity in the deletion mutant was at the same level

irrespective of the number of transcriptional start sites and AmtR binding sites. For the

527 bp fragment harboring the FarR and both AmtR binding sites, a 1.9-fold increase was

measured compared to the wild type. For the shorter fragment, including only the high

affinity AmtR binding site and the transcription start located 195 bp upstream of the gdh

gene, a 4.1-fold increase was determined. Therefore, it is very likely that despite contrary

results from DNA microarrays and earlier studies on GDH activity (Tesch et al., 1999;

Beckers et al., 2005; Silberbach et al., 2005a), gdh is regulated depending on the nitrogen

availability. Furthermore, AmtR might be involved in this regulation as has been proposed

using a bioinformatic approach as well gel shift assays and DNA affinity purification, which

could prove AmtR binding to the gdh upstream region in vitro (Müller, 2005; Beckers et al.,

2005). However, this is the first time the AmtR-mediated repression of gdh transcription was

shown in vivo.

Since AmtR binding to the gdh upstream region has been clearly demonstrated in previous

work as well as in this work, the binding behavior of AmtR was closer investigated. AmtR is

a TetR/AcnR type regulator. In general, these kinds of regulators bind small effector

Results 65

free

DN

A

0.5 5 50 ng AmtR+ + + 5 mM AMPfre

eD

NA

0.5 5 50 ng AmtR+ + + 5 mM AMP

molecules that lead to conformational changes and thereby influence interaction with DNA

(Ramos et al., 2005). For AmtR, release from its target DNA has been shown to be the

result of protein-protein interaction with the adenylylated signal transduction protein GlnK

(Beckers et al., 2005). In connection with the detailed characterization of the gdh promoter,

it was also investigated, whether AmtR binding is influenced not only by the modified PII

signal transduction protein, but also by AMP itself. This was tested by gel retardation

experiments as shown below. In these, purified AmtR protein (kindly provided by J. Reihlen,

University of Cologne) and 5 mM AMP were added to the binding reaction.

Fig. 4.14: Binding of AmtR to the gdh promoter in the presence of AMP. In gel shift assays with a digoxigenin-labeled fragment of the gdh upstream region, rising amount of purified AmtR protein were applied. In addition to that, 5 mM AMP were added. As control, one reaction did not contain AmtR (free DNA).

As reported before, gel retardation assays confirmed AmtR binding to the gdh promoter

region. However, an influence of AMP on AmtR binding was not shown. Therefore, it seems

that GlnK-AMP3 is necessary to release AmtR from its target DNA as has been proposed by

Beckers and coworkers (2005).

Results 66

4.2.3.2 Influence of putative regulators on gdh transcription

Besides AmtR, DNA affinity purification led to the identification of other putative regulators

of gdh transcription. These were FarR and OxyR, which both exhibited nitrogen-dependent

binding to the gdh promoter (Müller, 2005). Within this section, influence of these regulators

on gdh transcription was investigated.

For OxyR, sequence alignments suggested similarity to a transcriptional regulator of

M. leprae (Müller, 2005). In other organisms, for instance enteric bacteria, a regulatory

function of OxyR in response to oxidative stress has been reported (Christman et al., 1989).

During DNA affinity purification, OxyR was isolated from C. glutamicum cell extract after

cultivation under nitrogen limitation. The respective binding region was narrowed down to

173 bp to 376 bp upstream of the start codon. Since fluorescence measurements have

been proven to be a suitable method to determine the effect of putative regulators on the

promoter activity, the method was used to investigate the influence of OxyR on nitrogen-

dependent transcription of gdh. In Dot blot experiments, the deletion of oxyR did not alter

the nitrogen-dependent transcription pattern of gdh. This lacking effect of the deletion on

nitrogen-dependent gdh transcription in Dot blots was also observed for AmtR (Müller,

2005). Eventually, it was shown not to be true and therefore this alternative method to

determine transcription level was used for investigation of OxyR as well. pEPR1 carrying a

398 bp and 313 bp fragment, respectively, was transformed into TMΔoxyR and the parental

strain RES167. The longer construct included the whole fragment that presumably harbored

the OxyR binding site, while the 313 bp fragment contained only 149 bp of the 204 bp

region, to which OxyR bound in the DNA affinity purification. A possible effect on gdh

transcription due to the absence of a functional oxyR gene, would additionally allow

narrowing down the putative binding region of the OxyR protein. Samples were taken as

described before during exponential growth, under good nitrogen supply and with glutamine

as sole nitrogen source. Again, Dot blot experiments were carried out in parallel to confirm

the functionality of the nitrogen starvation response in the respective strains.

Results 67

0

100x10 3

200x10 3

300x10 3

400x10 3

500x10 3

600x10 3

700x10 3C

ps(m

g dw/m

l)-1

+N -N +N -N +N -N +N -N +N -N +N -N

amtA

0

100x10 3

200x10 3

300x10 3

400x10 3

500x10 3

600x10 3

700x10 3C

ps(m

g dw/m

l)-1

+N -N+N -N +N -N+N -N +N -N+N -N +N -N+N -N +N -N+N -N +N -N+N -N

amtA

Fig. 4.15: Investigation of the impact of OxyR on nitrogen-dependent gdh transcription. Activity of the promoter was determined depending on the nitrogen source. C. glutamicum RES167 and TMΔoxyR carrying pEPR1 without promoter (control) and different fragments of the gdh promoter were cultivated until exponential phase. For a detailed description of the fragments refer to figure 4.8. Either ammonium (black bars) or glutamine (100 mM, grey bars) was used as nitrogen source. After excitation at 395 nm, an emission spectrum ranging from 490 nm to 520 nm was monitored. Maximal fluorescence was put into relation to mgdry weight per ml. In addition to that, Dot blots with an amtA probe containing 2.5 µg total RNA were performed to check the functionality of the nitrogen starvation response.

In the examined strains nitrogen control was functional as it was proven by Dot blot

experiments with an amtA probe. As shown before, the presence of pEPR1 did not exhibit

any influence on nitrogen control. In addition to that it was demonstrated that the deletion of

oxyR did not have an impact on the expression of the AmtR-regulated amtA gene. This had

been shown for gdh already (Müller, 2005), but now it was also obvious for a strongly

regulated gene of the nitrogen starvation response. Basically, this excludes a function of

OxyR in regulation of nitrogen-controlled genes. In accordance with the detected nitrogen

starvation response, in the parental strain RES167 as well as in the deletion mutant gdh

expression was enhanced upon nitrogen deprivation. In both strains, the expression level

under good ammonium supply and under limiting conditions was at the same level as in the

wild type. That means that first of all, regulation in RES167 was comparable to the wild type

and as reported before (Müller, 2005), the deletion of oxyR did not show an influence on the

TMΔoxyR control

TMΔoxyR 398

TMΔoxyR 313

RES167 control

RES167 398

RES167 313

Results 68

nitrogen-dependent gdh expression. The function of OxyR and the reason for nitrogen-

dependent binding during DNA affinity purification therefore remain unknown.

FarR is a HutC/FarR-type regulator of the GntR family. During DNA affinity purification, it

was isolated from C. glutamicum cell extract after cultivation under good nitrogen supply. In

vitro binding to the promoter region was verified in gel retardation assays. Furthermore, the

principal capacity to repress transcription starting from the gdh promoter in vivo was

demonstrated in reporter gene assays after heterologous expression in E. coli (Müller,

2005). But still, the function of FarR in C. glutamicum, especially in connection with

regulating gdh expression, remained to be elucidated. In E. coli, FarR has been described

controversially as fatty acyl responsive regulator of TCA cycle genes or as a regulator of

genes encoding a 2-O-α-mannosyl-D-glycerate transport and metabolism system (Quail et

al., 1994; Sampaio et al., 2004). One approach to solve the question concerning the

function of FarR in regulation of gdh transcription was to perform reporter gene assays with

the wild type carrying two different pEPR1 constructs. The 527 bp fragment spanned the

whole promoter region including the FarR binding site 450 bp to 469 bp upstream of the

start codon. The 398 bp fragment only harbored the two AmtR binding sites, but did not

include the FarR binding site. During cultivation under nitrogen surplus, FarR was shown to

bind to the gdh promoter region (Müller, 2005). Therefore, if there occurred FarR-mediated

regulation under the tested conditions, differences in promoter activity would be expected

for the 527 bp and 398 bp fragments during cultivation in CgC medium. In addition, pEPR1

constructs were used that contained the two fragments in an inverted direction to be able to

monitor transcription from a possibly divergently orientated promoter.

Results 69

0

100x103

200x103

300x103

control 527 398 398*527*0

100x103

200x103

300x103

control 527 398 398*527*

Fig. 4.16: Investigation of FarR-dependent transcription of gdh and detection of a divergently orientated promoter. C. glutamicum wild type carrying one fragment of the gdh promoter including a FarR binding site (527) and one fragment without FarR binding site (398) were cultivated until exponential growth phase. For a detailed description of the fragments refer to figure 4.8. As negative control cultures carrying pEPR1 without promoter were used. Additionally, wild type cultures with plasmids harboring inverted fragments (*) were cultivated as described above. After excitation at 395 nm, an emission spectrum ranging from 490 nm to 520 nm was monitored. Maximal fluorescence was put into relation to mgdry weight per ml.

From the results of the reporter gene assays, it could be concluded that FarR was not

responsible for regulation gdh expression under standard cultivation and during exponential

growth. There was no difference in promoter activity detectable between the fragment with

and without FarR binding site showing that FarR did not play a crucial role under the

conditions tested. Despite previous results indicating binding to gdh promoter, the function

of this putative regulator remaines unknown. By including two inverted fragments of the gdh

promoter in this approach, it was also possible to show that there is no promoter orientated

in an opposite direction in the vicinity of the gdh promoter. The level of activity for the

inverted fragments was comparable to the culture containing the empty plasmid. These

results showed that there is no transcriptional start site and consequently no transcription of

for instance antisense RNA. Furthermore, it proved that the promoter of the divergently

orientated glxK gene located upstream of gdh is obviously not positioned within the gdh

promoter region. Therefore, it was concluded that the identified regulator binding sites lie

within the gdh promoter region and the respective regulators are responsible for controlling

gdh expression.

Results 70

Despite the results that FarR did not exhibit an influence on gdh transcription under

standard conditions, nitrogen-dependent expression was investigated in the farR deletion

strain TMΔfarR and the parental strain RES167. To do so, Dot blot experiments and the

more sensitive method of quantitative real time RT PCR were performed. Real time RT

PCR was carried out using two different sets of primers in order to circumvent primer

specific effects such as dimerization and formation of secondary structures. Primers used

amplified different fragments of the gdh transcript. Therefore, an impact on the results

caused by possible secondary structures of the PCR product could also be excluded.

Fig. 4.17: Analysis of gdh expression in RES167 and TMΔfarR depending on nitrogen availability. RNA for Dot blot experiments was obtained from both strains after cultivation under nitrogen surplus (+N) and after 1 hour without nitrogen source (-N). From each sample 1 µg of total RNA was applied. Tab. 4.3: Analysis of gdh expression in RES167 and TMΔfarR. Quantitative real time RT PCR was performed with RNA from cultures grown under ammonium surplus (+N) and 1 hour without nitrogen source (-N). Three replicates were used and standard deviation was calculated. Expression of gdh in RES167 under nitrogen surplus was set to 1. In order to verify results, two sets of primers were used.

Strain Primer 1 Primer 2 RES167 +N 1.00±0.16 1.00±0.10

RES167 -N 2.61±0.37 2.34±0.13

TMΔfarR +N 1.05±0.23 1.05±0.14

TMΔfarR -N 2.25±0.46 2.42±0.39

Dot blot experiments revealed the expected transcription pattern (Müller, 2005). The gdh

gene was transcribed at a low level under good ammonium supply and after cultivation

without nitrogen source, transcription was significantly enhanced. This was observed in

RES167 as well as in TMΔfarR, showing that FarR did not alter nitrogen-dependent gdh

transcription. Quantitative real time RT PCR experiments underlined the results. First of all,

both primer sets showed comparable results indicating that neither the chosen primers nor

the amplified fragment had an effect on the outcome of the experiment. The level of gdh

TMΔfarR

RES167

+N -N

Results 71

transcript rose when nitrogen supply declined. This was the case in RES167 as well in the

deletion mutant. Therefore, an exclusively FarR-mediated regulation of nitrogen-dependent

transcription could be excluded and the function of FarR in regulating gdh transcription

remains unknown.

4.2.4 Studies on sigma factor-dependent gdh expression

In addition to transcriptional control mediated by regulatory proteins so far mainly covered

within this work, gene expression is also controlled by interaction of the core promoter with

respective σ subunits of the RNA polymerase (sigma factors). In C. glutamicum, seven

sigma factors have been described based on analysis of the genome sequence. The

essential primary sigma factor SigA (σA) is responsible for expression of most genes during

exponential growth under optimal growth conditions (Halgasova et al., 2001). Alternative

sigma factors are named according to the respective homologs in M. tuberculosis, namely

SigB, SigC, SigD, SigE, SigH, and SigM (for review, Nešvera & Pátek, 2008). SigB is a

primary-like sigma factor and is involved in regulating gene expression during certain stress

conditions, the transition between exponential growth and stationary phase, and during

stationary phase (Halgasova et al., 2001; Larisch et al., 2007). The remaining sigma factors

are so-called extracytoplasmic function (ECF) factors that exhibit functions during various

stress conditions depending on different environmental stimuli. SigH for instance was

shown to be involved in regulating the expression of genes of the heat stress response

(Engels et al., 2004; for review, Nešvera & Pátek, 2008).

Transcription of gdh has been reported to be regulated depending on a variety of

environmental conditions such as variation of nitrogen and carbon source, chill stress, and

oxygen limitation (Müller, 2005). Since more and more knowledge is available on possible

functions of sigma factors as well as on respective promoter structures, a first approach

was carried out to investigate the influence of certain sigma factors on regulation of gdh

expression. Since in the vicinity of the newly identified gdh transcriptional start no clear

housekeeping promoter is present, there is a chance that alternative sigma factors

recognize this promoter using a different recognition site. To gain further knowledge on the

involvement of sigma factors, gdh expression was analyzed in the respective mutants using

real time RT PCR. Growth of the parental strain RES167 and the sigB, sigD, sigE, and sigM

deletion mutants as well as of a sigH insertion mutant was monitored. Cultures were grown

as described in section 3.2.4. Increase in cell density was determined by hourly

measurement of OD600.

Results 72

Fig. 4.18: Growth of C. glutamicum RES167 and mutants lacking functional sigma factors. Strains were cultivated in CgC medium and growth of RES167 (+), CL1 (+), RES ΔsigD (+), RES ΔsigE (+), RES ΔsigM (+), and RES INTsigH (+) was monitored by hourly measurements of the optical density at 600 nm (OD600).

Growth of all strains examined did not differ during cultivation in CgC medium as can be

seen in figure 4.18. This result was consistent with previous data obtained by Ley (2005).

Samples for real time RT PCR were taken during exponential growth, five hours after

inoculation, and after ten hours at the transition between exponential and stationary phase.

Expression of gdh in the mutant strains and RES167 is summarized in table 4.4.

Tab. 4.4: Analysis of gdh expression in RES167 and mutants lacking functional sigma factors. Quantitative real time RT PCR was performed with RNA from cultures grown in CgC medium. Samples were taken 5 and 10 hours after inoculation. Three replicates were used and standard deviation was calculated. Expression of gdh in RES167 during exponential growth was set to 1.

Strain Expression during

exponential growth (5 h) Expression during the transition phase (10 h)

RES167 1.00±0.12 1.00±0.05

CL 1 1.91±0.31 1.01±0.07

RES ΔsigD 1.72±0.27 0.88±0.05

RES ΔsigE 1.07±0.07 0.80±0.07

RES ΔsigM 1.18±0.09 0.72±0.08

RES INTsigH 2.05±0.16 0.90±0.04

0.1

1

10

100

0 5 10 15 20 25time [h]

OD

600

Results 73

Data from quantitative real time RT PCR experiments showed first of all that gdh was

expressed at the same level during exponential growth and during the transition between

exponential and stationary phase. Furthermore, the missing sigma factors did not have a

crucial influence on gdh expression during exponential growth and the transition phase.

Expression of gdh in the respective mutants did not change drastically compared to

RES167. Due to these marginal changes in transcript level, this first approach taking into

account regulation of gdh expression by sigma factors did not provide any further results.

However, cultivation was only performed under standard condition using CgC medium

without causing a particular stress situation, which might be necessary for the expression of

certain sigma factors as shown for instance for sigB (Larisch et al., 2007). Therefore,

starting out from this approach, there is a wide range of possibilities to examine sigma

factor-dependent gdh expression.

4.2.5 Investigation of the putative orf Cg2281

Another feature of the gdh upstream region, so far not mentioned, is the putative open

reading frame (orf) Cg2281. It was revealed by sequence analyses and spans 93 bp

(GenDB, CeBiTec, University of Bielefeld). Since it was only predicted by a bioinformatic

approach and no function was suggested, it was investigated in the course of the

experiments, whether there is a promoter present that might be responsible for transcription

of Cg2281. The putative start codon of Cg2281 is located 422 bp upstream of the gdh gene.

In the course of primer extension experiments and investigation of the promoter region

applying reporter gene assays one aim was to investigate, whether there is an additional

transcriptional start site located upstream of the one identified by Börmann and coworkers

(1992). This region would harbor the promoter of the putative orf Cg2281.

Results 74








GGAAATCATG

*

*-10-35

+1

1

61

121

181

241

301

361

526

Cg2281

-35








GGAAATCATG

*

*-10-35

+1

1

61

121

181

241

301

361

526

Cg2281

-35

Fig. 4.19: Overview of the gdh promoter region. Transcriptional start sites indicated by asterisks are located 285 bp and 195 bp upstream of the start codon (284 bp and 194 bp, respectively, using the sequence published by Kalinowski et al., 2003). -10 region and putative -35 regions are marked in bold letters. The putative orf Cg2281 is underlined as well as the gdh start codon.

In the primer extension experiments, the intergenic region located upstream of the gdh

gene was investigated. By this approach, up to 460 bp and 530 bp upstream of the gdh

start codon were covered. This region included about 100 bp of the Cg2281 upstream

region. Within this region it was not possible to identify a transcriptional start site under the

tested cultivation conditions that would correspond to Cg2281. Also, the construction of

GFP promoter fusions did not give any hints which would suggest transcription of Cg2281.

The 527 bp fragment used in the measurements included about 110 bps of the Cg2281

upstream region. If there had been an additional promoter located upstream of Cg2281, this

would have led to an enhanced promoter activity of the 527 bp fragment compared to the

398 bp and 313 bp fragments. Since there was no significant difference in promoter

strength of the three fragments (figure 4.9), it was assumed that there is no additional

housekeeping promoter located in the direct upstream region of Cg2281. Whether there is

transcript and promoter activity detectable under certain environmental stress conditions or

a transcriptional start site located further upstream than 100 bp can of course not be

excluded.

Results 75

4.3 Identification of AmtR and FarR target genes For AmtR and FarR, participation in regulating gdh transcription has been suggested based

on the results of DNA affinity purification and gel retardation experiments (Müller, 2005).

Generally, the repressor AmtR is well characterized in C. glutamicum regarding the

respective regulon and binding motif (Beckers et al., 2005). Nevertheless, recent studies

imply that further target genes might have been left out in studies of the AmtR regulon

(S. Hans, unpublished data).

As already mentioned above, comparably little is known about function and stimulus of

FarR-dependent regulation of gene expression. For these reasons, in the following section

the focus lay on the identification of additional target genes for both regulators.

4.3.1 Identification of FarR target genes

Different experimental set-ups have already been used to find FarR target genes other than

gdh. Based on a putative FarR binding motif, the genome was screened for further target

genes. Binding was then investigated by competitive gel retardation assays. A more global

approach was also chosen by performing DNA microarrays with RNA of the respective

deletion mutant and RES167. While DNA microarrays led to the identification of a variety of

putative FarR targets, only one target gene was identified by the screening approach in

combination with competitive gel shift assays (Müller, 2005). For this target gene, dtsR2

encoding a detergent sensitive rescuer protein, FarR binding to the upstream region was

verified by gel retardation experiments (data not shown). As a result of the DNA

microarrays, a broad number of genes possibly regulated by FarR was identified. However,

many of them had not been characterized before and a possible function was only

annotated. Only genes that were expressed differently with a factor of at least three in DNA

microarrays were taken into account for further investigation within this work. The

respective genes are listed in table 4.5.

Results 76

Tab. 4.5: Genes expressed differently in TMΔfarR compared to RES167 in DNA microarrays. Listed are genes that were regulated at least by a factor of 3. A negative value indicates stronger expression in RES167 (Müller, 2005).

NCgl Gene Annotation RES167 vs. TMΔfarR

0689 Putative short-chain alcohol dehydrogenase -3.36 1331 ugpB ABC sugar transport system 3.42 1341 argJ Ornithine acetyltransferase 3.57 1342 argB Acetylglutamate kinase 4.19 1343 argD Ornithine/Acetylornithine aminotransferase 3.58 1466 Phospholipid binding protein 3.14 1934 Response regulator 3.09 2716 cysD 3'-phosphoadenosine 5'-phosphosulfate synthetase 3.01 2717 cysH 3'-phosphoadenosine 5'-phosphosulfate synthetase 3.35 2718 cysI Sulfite reductase 3.88 2787 Putative flavoprotein involved in K+ transport -9.32

Genes of different metabolic pathways are apparently regulated by FarR. The cys genes

are organized in an operon and are involved in the assimilatory reduction of inorganic

sulphur compounds (Rückert et al., 2005). Except for the cys operon and the arg genes,

function of the identified genes is unknown. FarR-mediated regulation of gene expression of

the arg genes, cys operon, and NCgl2787 as suggested by DNA microarrays was

additionally investigated using gel retardation assays and real time RT PCR (data not

shown). Since it was only possible to verify results for the arg genes, the focus of further

experiments was put on the regulation of arginine biosynthesis.

4.3.1.1 Characterization of arginine biosynthesis genes

Arginine is synthesized from the precursor glutamate in eight enzymatic steps via a so-

called cyclic pathway (Sakanyan et al., 1996). The respective biosynthesis genes in

C. glutamicum are organized in a cluster as shown in figure 4. 20 A. Single enzymes have

been characterized biochemically and on the genetic level, such as ornithine

carbamoyltransferase (encoded by argF; Chun & Lee, 1999), argininosuccinate synthetase

(encoded by argG; Soon-Young et al., 2003), and argininosuccinate lyase (encoded by

argH; Yim & Lee, 2005).

Results 77

Fig. 4.20: Arginine biosynthesis. Arginine biosynthesis genes of C. glutamicum are organized in a cluster as shown in A. The respective gene products catalyze the formation of arginine from glutamate in a cyclic pathway, in which transacetylation between glutamate and ornithine is mediated by ornithine acetyltransferase (encoded by argJ). The successive enzymatic reactions are performed by acetylglutamate kinase (encoded by argB), acetylglutamate semialdehyde dehydrogenase (encoded by argC), ornithine/acetylornithine aminotransferase (encoded by argD), ornithine carbamoyltransferase (encoded by argF), argininosuccinate synthetase (encoded by argG), and argininosuccinate lyase (encoded by argH). Carbamoylphosphate is synthesized from glutamine by carbamoylphosphate synthase (encoded by carAB). The argR gene is annotated as encoding the arginine repressor. Compared to Gram-negative organisms, regulation of arginine biosynthesis is less

investigated in C. glutamicum (Maas, 1994; Lu et al., 2004) especially regarding regulatory

mechanisms on the level of transcription. Due to the chromosomal organization of the

arginine biosynthesis genes, an operon structure was assumed. Probably because of the

transcript length, Northern blot experiments did not show clear results (data not shown), so

that RT PCR experiments were carried out using overlapping sets of primers as shown in

figure 4.20 A. For the RT PCR experiments, RNA prepared from samples after cultivation

with good nitrogen supply and after 30 minutes without nitrogen source was used.

argF

aspartate

1074 1167 954 1176 516 1206 1434

argC argJ argB argD argF argR argG argH

960

[83] [45] [0] [13] [3] [167] [60]

A

glutamateargC argJargD argB

N-acetylornithine

argJ

ornithine

citrulline

L-argininosuccinate

arginine

argGargH

fumarateUrea cycle

B

glutamine

glutamate

carbamoyl-PcarAB

+

argF

aspartate

1074 1167 954 1176 516 1206 1434


960

[83] [45] [0] [13] [3] [167] [60]

1074 1167 954 1176 516 1206 1434


9601074 1167 954 1176 516 1206 1434


960

[83] [45] [0] [13] [3] [167] [60]

A

glutamateargC argJargD argB

N-acetylornithine

argJ

ornithine

citrulline

L-argininosuccinate

arginine

argGargH

fumarateUrea cycle

B

glutamine

glutamate

carbamoyl-PcarAB

+

Results 78

Fig. 4.21: Characterization of the chromosomal organization of arginine biosynthesis genes. Arginine biosynthesis genes were analyzed by RT PCR using a combination of different primer sets as displayed in figure 4.20 A. As negative control for each sample a set-up was carried along without performance of RT reaction (-). RNA was prepared from samples taken under good nitrogen supply (+N) and after 30 minutes of nitrogen deprivation (-N). The following sets of primers were used: A: amplification of argC, argJ, and argB B: amplification of argF and argR C: amplification of argR, argG, and argH D: amplification of argB, argD, and argF (1 and 4); argG and argH (2 and 5); argF, argR, argG, and argH (3 and 6)

Results of the RT PCR experiments led to the assumption that arginine biosynthesis genes

were transcribed at a higher level during nitrogen starvation. But since RT PCR can only be

used for quantification of transcripts under some restraints, these hints needed to be

confirmed using a more suitable method (4.3.1.3). By combining the results derived from

the RT PCR experiments shown in figure 4.21, it was concluded that the first six genes of

the cluster, argC, argJ, argB, argD, argF, and argR, form a common transcript. The last two

genes argG and argH were also cotranscribed. There is a high probability, that even all

eight genes form one operon. Because of an apparently enhanced transcription under

nitrogen starvation, it was possible to detect a transcript starting in argF ending in argG

under these conditions. This hypothesis was underlined by the fact that a terminator

M + - + - M +N -N

M + - + - M +N -N

M + - + - M +N -N

M + - + - M +N -N

M 1 - 2 - 3 - 4 - 5 - 6 - M +N -N

A B C

D

Results 79

A

free

DN

Aco

ntro

l

FarR

free

DN

Aco

ntro

l

FarR

free

DN

Aco

ntro

lFa

rR

free

DN

Aco

ntro

lFa

rR

free

DN

Aco

ntro

lFa

rRB

structure is only located downstream of the argH gene (GenDB, CeBiTec, University of

Bielefeld).

4.3.1.2 Determination of FarR and ArgR binding sites

Genes of the arg operon are presumably regulated by FarR. While real time RT PCR

experiments failed to verify the results of DNA microarrays (data not shown), binding of the

putative regulator to the upstream regions of argC and argG was investigated. To prove in

vitro binding of FarR, gel retardation experiments were performed using cell extract of

E. coli heterologously expressing FarR of C. glutamicum (kindly provided by T. Müller,

University of Cologne). For this approach, a 200 bp fragment of the upstream region of the

first gene in the arg operon was chosen. Because of the long intergenic region that might

harbor a regulator binding site, binding to the argG upstream region was also examined.

Fig. 4.22: Determination of FarR binding in the upstream regions of arginine biosynthesis genes. Gel retardation assays with digoxigenin-labeled fragments of the argC (A) and argG (B) upstream regions were performed with 1.5 µg cell extract of E. coli DH5αmcr heterologously expressing FarR of C. glutamicum. For a more precise determination of the FarR binding site upstream of argG (C) competition assays using the indicated unlabeled 50 bp-fragments (D) in a 1500-fold molar excess were performed. Control: cell extract of E. coli containing pUC18. Free DNA: without any cell extract.

1 2 3 4 5 6 7C

1 2 3 4 5 6 7C

D

argGargR

-100

71

2

3

4

5

6

-176 +23

+1

-176 +24

argGargR

-100

71

2

3

4

5

6

-176 +23

+1

-176 +24

Results 80

A

BGCCAGGTTA--TATAACCAGTC gdhAACCGGTTAGCGAAACGATTAG dtsR2GGCAGGTAAGGTATAACCCGAG argG

A

BGCCAGGTTA--TATAACCAGTC gdhAACCGGTTAGCGAAACGATTAG dtsR2GGCAGGTAAGGTATAACCCGAG argG

Gel shift assays proved that FarR bound in vitro to the upstream regions of argC and argG.

By this, transcription of arginine biosynthesis genes is probably repressed. In the case of

argG, it was furthermore possible to narrow down the putative FarR binding site. In addition

to the labeled fragment of the argG upstream region, unlabeled 50 bp DNA fragments as

indicated in figure 4.22 D, were applied. Fragments 2 and 3 inhibited the shift. That means

the binding site was located in the overlapping region of the two fragments, which would be

77 bp to 151 bp upstream of the argG start codon. There is only little knowledge available

on a consensus binding motif of FarR. A consensus sequence for the FarR binding motif

based on two putative FarR binding sites has been proposed by Müller (2005). The

respective consensus sequence as well as putative FarR binding sites identified by gel shift

assays is shown in figure 4.23, indicating a rather low level of conservation.

Fig. 4.23: Comparison of deduced FarR binding motifs. Known putative FarR binding motifs upstream of gdh, dtsR2, and argG are listed. Nucleotides conserved in all three sequences are marked in bold letters. All three were proven by gel retardation experiments.

In the course of the experiments dealing with transcriptional regulation of arginine

biosynthesis, the function of the annotated arginine repressor ArgR (encoded by argR) was

investigated as well. The C. glutamicum ArgR protein shares 33 % and 31 % amino acid

identity with the respective regulators of B. subtilis and E. coli (J. Amon, personal

communication). In Gram-negative organisms, such as E. coli and Pseudomonas

aeruginosa as well as in B. subtilis, regulation of arginine metabolism is understood in quite

good detail (Czaplewski et al., 1992; Maas, 1994; Dennis et al., 2002; Lu et al., 2004). In

E. coli, ArgR represses the transcription of arginine biosynthesis genes in the presence of

arginine. About mechanisms involved and ArgR binding sites broad structural knowledge is

available (for overview, Maas, 1994; Makarova et al., 2001). About ArgR-mediated

Results 81

transcriptional regulation of arginine biosynthesis genes, nothing is known in

C. glutamicum. For this reason, the function of ArgR was closer investigated.

Using gel retardation experiments, in vitro binding of ArgR to the upstream regions of

arginine biosynthesis genes was tested. For these, cell extract of E. coli heterologously

expressing ArgR of C. glutamicum was used. However, binding to the argC, argR, and

argG upstream regions could not be demonstrated (data not shown). Therefore, by using

this approach it was not possible to gain information about a putative ArgR-mediated

regulation of the arg operon. It was also tested, whether FarR bound to the argR upstream

region directly exhibiting a regulatory function on the expression of the regulator. If that was

the case, a regulatory mechanism including a set of two regulators would be present. But

under the tested conditions, binding of FarR to the amtR upstream region was not shown

irrespective of the presence of arginine.

However, with FarR, a putative, novel regulator involved in controlling transcription of

arginine biosynthesis genes was identified. Data from transcriptome analyses and binding

studies hint at a FarR-meditated repression of arg operon. Interestingly, the same protein

might be involved in transcriptional regulation of gdh (Müller, 2005). This reaction provides

the precursor for arginine synthesis. The function of the second regulator presumably

involved in controlling arginine metabolism, ArgR, was so far not clarified.

4.3.1.3 Transcriptional regulation of arginine biosynthesis genes

Now, that it was possible to identify a putative regulator of the arg operon, transcription of

the respective genes in response to different cultivation conditions was investigated. As it is

the case for the regulator ArgR, in respect of environmental stimuli comparably little is

known about transcriptional regulation of arginine biosynthesis in C. glutamicum. Because

of hints obtained from RT PCR experiments, nitrogen-dependent transcription of three arg

genes was analyzed in Dot blot experiments. For these, RNA was prepared from samples

taken during cultivation in CgC medium, after 30 minutes without nitrogen source and

30 minutes after a pulse with 100 mM NH4(SO4)2. Also, transcription depending on the

presence of arginine was of interest. To detect a possible influence of the end product of

the biosynthetic pathway, real time RT PCR was carried out. RNA was obtained from

samples after cultivation in CgC medium and 1 hour after the addition of 10 mM arginine.

As the first gene of the arg operon argC was chosen, in addition to that argR and argH

transcription was examined. Results of Dot blot and realtime RT PCR experiments are

summarized in figure 4.24.

Results 82

0.0

0.5

1.0

1.5

argC

argR

+N –N pulse

argH

argC argR argH

A B

Rel

ativ

e fo

ldex

pres

sion

0.0

0.5

1.0

1.5

argC

argR

+N –N pulse

argH

argC argR argH

A B

Rel

ativ

e fo

ldex

pres

sion

Fig. 4.24: Expression analyses of arginine biosynthesis genes. Using Dot blot experiments (A) expression of argC, argR, and argH was investigated under nitrogen surplus (+N), 30 minutes of nitrogen starvation (-N), and after an ammonium pulse (100 mM NH4(SO4)2, 30 minutes). 1 µg total RNA was applied. By quantitative real time RT PCR, expression of argC, argR, and argH was analyzed depending on the presence of arginine (B). Black bars indicate transcript level without arginine and grey bars show transcript level after cultivation with 10 mM arginine for 1 hour. Three replicates were used and standard deviation was calculated. Expression without arginine was set to 1. As already assumed from the results of the RT PCR, transcription of argC and argR was

induced upon nitrogen starvation and reduced again upon addition of ammonium. The

results concerning nitrogen-dependent argH transcription were less clear. In order to test

whether AmtR might be involved in regulating nitrogen-dependent transcription of the arg

operon, gel retardation assays with E. coli cell extract heterologously expressing AmtR of

C. glutamicum were performed as well (kindly provided by T. Müller, University of Cologne).

However, by this approach, binding to the argC, argR, and argG upstream region could not

be shown under the tested conditions (data not shown). Therefore, the mechanism of

transcriptional regulation of the arg operon in dependence of the nitrogen supply remains to

be investigated.

This is also the case for the transcriptional regulation depending on the presence of

arginine. As expected for genes of the biosynthesis pathway, transcription of argC was

significantly reduced upon addition of arginine. The same was true for argR transcription.

On the other hand, argH transcription was only slightly altered in the presence of arginine.

Real time RT PCR experiments with RNA of a farR deletion mutant led to the same results

indicating that arginine-dependent regulation was not mediated by FarR (data not shown).

Based on these observations, a regulatory mechanism controlling expression of

biosynthesis genes in response to the presence of the respective end product was

detectable, whereas genes of the same operon were expressed at different levels. Despite

Results 83

the involvement in regulating arginine metabolism in other organisms, the function of ArgR

in this context remains to be elucidated in C. glutamicum, especially since binding to the

respective upstream regions of arginine biosynthesis genes could not be demonstrated.

4.3.2 Identification of AmtR target genes

The AmtR regulon has been investigated to great extend applying global approaches such

as DNA microarrays, two dimensional gel electrophoresis, and bioinformatic tools (Beckers

et al., 2005). In total, 36 genes are regulated by AmtR directly and indirectly. Knowledge on

AmtR binding sites, acquired by the examination of promoters of different nitrogen-

regulated genes, allowed the identification of a consensus sequence for the AmtR binding

motif (Beckers et al., 2005). Binding studies and mutational analysis of respective promoter

regions led to the collection of a large set of detailed information (for review, Hänßler &

Burkovski, 2008). Nevertheless, with optimization of experimental techniques new putative

target genes of the global repressor are presumed. As for instance shown by the example

of gdh, the combination of different methods might give better insight into regulatory

mechanisms. Based on recently performed DNA microarrays the focus was put onto two

new putative AmtR target genes. These genes were dapD encoding tetrahydrodipiconlinate

succinylase and mez encoding malic enzyme (S. Hans, unpublished data).

Tetrahydrodipiconlinate succinylase is the first enzyme within the so-called succinylase

variant of diaminopimelate synthesis. Diaminopimelate is the precursor of the cell wall

component peptidoglycan and lysine, an amino acid of outstanding industrial importance.

The succinylase variant of diaminopimilate synthesis is comprised of four enzymatic steps

yielding diaminopimelate from 2,3,4,5-tetrahydrodipicolinate. There is a parallel pathway for

synthesis of diaminopimelate in C. glutamicum. In an NADPH-dependent reaction, 2,3,4,5-

tetrahydrodipicolinate and ammonium form diaminopimelate catalyzed by diaminopimelate

dehydrogenase (figure 4.25). It has been shown that there is a difference between the two

pathways regarding activity depending on the nitrogen source. The succinylase variant is

necessary for the use of organic nitrogen compounds. Deletion of dapD showed that the

dehydrogenase variant could only compensate the loss of the succinylase variant at high

ammonium concentrations, but not in the presence of glutamate (Wehrmann et al., 1998).

Considering this background information, a possible involvement of AmtR is not only

reasonable, but also very interesting due to the connection of lysine production.

Results 84

L-Aspartate

L-Aspartate semialdehyde

2,3,4,5-Tetrahydrodipicolinate

Succinyl-2-amino-6-ketopimelate

Succinyl-2,6,L,L-diaminopimelate

L,L-Diaminopimelate

D,L-Diaminopimelate

Peptidoglycan L-Lysine

SuccinylCoA

CoA

Glutamate

2-Oxoglutarate

Succinate

NH4+ + NADPH

NADP

dapD

dapE

dapC

dapF

murE

ddh

lysA

L-Aspartate

L-Aspartate semialdehyde

2,3,4,5-Tetrahydrodipicolinate

Succinyl-2-amino-6-ketopimelate

Succinyl-2,6,L,L-diaminopimelate

L,L-Diaminopimelate

D,L-Diaminopimelate

Peptidoglycan L-Lysine

SuccinylCoA

CoA

Glutamate

2-Oxoglutarate

Succinate

NH4+ + NADPH

NADP

dapD

dapE

dapC

dapF

murE

ddh

lysA

Fig. 4.25: Synthesis of diaminopimelate in C. glutamicum. The succinylase variant and the dehydrogenase variant of diaminopimelate formation are presented. The succinylase variant is comprised of four enzymatic steps catalyzed by tetrahyrdodipicolinate succinylase (dapD), succinyl-aminoketopimelate transaminase (dapC), succinyldiaminopimelate desuccinylase (dapE) diaminopimelate epimerase (dapF). Diaminopimelate dehydrogenase (ddh) catalyzes the formation of diaminopimelate from tetrahydrodipicolinate. This serves as precursor for the synthesis of peptidoglycyan and lysine. The respective reactions are catalyzed by the diaminopimelate adding enzyme (murE) and diaminopimelate decarboxylase (lysA) (Wehrmann et al., 1998).

DNA microarrays with wild type RNA and RNA of an amtR deletion mutant indicated a 2.87-

fold induction of dapD expression in the deletion mutant (S. Hans, unpublished data). Using

quantitative real time RT PCR to verify this result, induction in the same order of magnitude

was observed, namely 2.52 ±0.09-fold. In addition to the analysis of dapD transcription in

dependence of AmtR, a putative AmtR binding motif was identified upstream of the dapD

gene in an approach of screening the genome using PREDetector software (Hiard et al.,

2007; J. Amon, personal communication). Therefore, by two independent approaches,

Results 85

AmtR-mediated regulation of dapD expression was demonstrated. The next step was to

investigate AmtR binding to the respective promoter region and to prove the predicted

binding motif. To do so, gel retardation assays were carried out with a 210 bp fragment of

the dapD upstream region. Since the aroP gene encoding an aromatic amino acid transport

protein ends only 29 bp upstream of the dapD start codon, the 3’-end of this gene was

included. In order to investigate the binding behavior, rising amounts of purified AmtR

protein were applied (kindly provided by J. Reihlen, University of Cologne).

Fig. 4.26: Investigation of AmtR binding to the dapD upstream region. Gel retardation experiments with digoxigenin-labeled fragments of the dapD upstream region were performed by adding rising amounts of purified AmtR protein (5 ng, 50 ng, and 100 ng) (A). For a more precise determination of the AmtR binding site upstream of dapD, competition assays with unlabeled 50 bp fragments (1-8) in a 1900-fold excess were performed. As control, H2O was added to one reaction (B). Additionally, one set-up did not contain AmtR protein (free DNA).

Binding of purified AmtR to the labeled fragment of the dapD upstream region was proven

as can be seen in figure 4.26. In order to identify the AmtR binding site and to find out,

whether it is consistent with the data obtained by application of PREDetector software,

competition assays were performed. To the labeled fragment, unlabeled, overlapping 50 bp

DNA fragments were applied in a 1900-fold excess. Addition of fragment 7 led to an

inhibition of the shift, indicating that the AmtR binding site was located within the region 48

bp upstream of the start codon. Indeed, with this approach it was possible to confirm the

predicted binding site. The AmtR binding site within the dapD promoter was shown to be

located 11 bp to 38 bp upstream of the start codon overlapping with the 3’-end of the aroP

gene.

free

DN

A

AmtR free

DN

AH

2O

1 2 3 4 5 6 7 8A B

free

DN

A

AmtRfree

DN

A

AmtR free

DN

AH

2O

1 2 3 4 5 6 7 8A B

Results 86

Fig. 4.27: Determination of the AmtR binding site in the dapD upstream region. Based on the results of gel shift assays, it was possible to confirm the predicted AmtR binding site in the dapD upstream region. Nucleotides marked in bold are highly conserved and match with the consensus sequence published by Beckers and coworkers (2005). This location within a coding region was the reason that this binding motif, despite high

similarity to the consensus sequence, had not been identified before. All together, it was

possible to identify dapD as a new AmtR target gene. Besides the identification of the

binding site, AmtR-mediated repression under nitrogen surplus could be demonstrated as

well. Due to the apparently nitrogen-dependent regulation of diaminopimelate synthesis

(Wehrmann et al., 1998), a participation of AmtR in this regulatory process seems

reasonable.

The other putative AmtR target gene mez encodes malic enzyme. This enzyme catalyzes

the NADP+-dependent decarboxylation of malate resulting in pyruvate. Malic enzyme has

been biochemically characterized and its impact on lactate metabolism has been studied

(Gourdon et al., 2000). As for dapD, only recent studies indicated involvement of AmtR in

regulating mez transcription. DNA microarrays showed a 3.46-fold enhanced transcription

in the amtR deletion mutant compared to the wild type (S. Hans, unpublished data). To find

transcriptional regulators of the mez gene, DNA affinity purification was performed and led

to the identification of AmtR as a putative regulator of mez transcription (V. Wendisch,

personal communication). But since there is no obvious connection to nitrogen metabolism

and the presence of an AmtR binding site in the mez upstream region has never been

proposed, direct regulation of mez by AmtR was investigated. To verify the results of DNA

affinity purification and to identify a possible binding site of AmtR, gel retardation

experiments were performed. For these, the same 291 bp fragment as for DNA affinity

purification as well as purified AmtR protein was used.

CACCCCTTCC GACTTGAACT GATAGGCCGA TAGAAATTAT TCTGGACGTC ATGACTACTG

ATAATTTCTATCGGCCTATCAGTTCAAG

+1-11-38dapDaroP

CACCCCTTCC GACTTGAACT GATAGGCCGA TAGAAATTAT TCTGGACGTC ATGACTACTG


+1-11-38dapDaroP


+1-11-38dapDaroP

Results 87

ACATTGCGAA ATTTTTGTTG AGCTACAGAT TTAGCTAGTG TTTTTGTTCC AGAACCCTAA

ATGAGGTTCT ACCCTTAACA GAGCTTCCCG CAAAAACACC GATTAACAAG GCTAAATGAT

ATGACCATCG+1

GTTGAGCTACAGATTTAGCTAGTGTT

-79-104

mez

ACATTGCGAA ATTTTTGTTG AGCTACAGAT TTAGCTAGTG TTTTTGTTCC AGAACCCTAA

ATGAGGTTCT ACCCTTAACA GAGCTTCCCG CAAAAACACC GATTAACAAG GCTAAATGAT

ATGACCATCG+1

GTTGAGCTACAGATTTAGCTAGTGTT

-79-104

mez

Fig. 4.28: Investigation of AmtR binding to the mez upstream region. Gel retardation experiments with digoxigenin-labeled fragments of the mez upstream region were performed by adding rising amounts of purified AmtR protein (5 ng, 50 ng, and 100 ng) (A). For a more precise determination of the AmtR binding site upstream of mez, competition assays with unlabeled 50 bp fragments (1-12) in a 1900-fold excess were performed. As control, H2O was added to one reaction (B). Additionally, one set-up did not contain AmtR protein (free DNA).

In gel retardation assays, binding of AmtR to the mez upstream region was confirmed. In

order to localize the exact binding site, competition assays with overlapping, unlabeled 50

bp DNA fragments were carried out. Fragment 7 located 71 bp to 120 bp upstream of the

start codon caused an inhibition of the shift, indicating that the presumed binding site lay in

this region.

Fig. 4.29: Determination of the AmtR binding site in the mez upstream region. Based on the results of gel shift assays an AmtR binding site was determined in the mez upstream region. Nucleotides marked in bold are highly conserved and match with the consensus sequence published by Beckers and coworkers (2005).

Within fragment 7, a sequence similar to the consensus motif of the AmtR binding site was

found located 79 bp to 104 bp upstream of the mez gene. Using different bioinformatical

approaches, namely Hidden-Markov models by Beckers and coworkers (2005) and

PREDetector software (J. Amon, personal communication), it was not possible to identify an

AmtR binding site upstream of the mez gene. However, by the gel retardation experiments

free

DN

AH

2O

1 2 3 4 5 6 7 8 9 10 11 12free

DN

AAmtR

A B

free

DN

AH

2O

1 2 3 4 5 6 7 8 9 10 11 12free

DN

AH

2O

1 2 3 4 5 6 7 8 9 10 11 12free

DN

AAmtR

A B

Results 88

results of the DNA affinity purification were verified. It was demonstrated that the changes

in transcript level observed in DNA microarrays were probably due to direct regulation of

mez transcription by AmtR and not the result of secondary effects.

By reinvestigating the AmtR regulon using DNA microarrays and subsequently investigating

the regulation of selected genes, it was possible to approve AmtR-mediated regulation of

two novel target genes. While for the regulation of dapD the physiological role of AmtR

regulation is obvious due to the nitrogen dependency of the split pathway for

diaminopimelate synthesis, this is not the case for mez.

Discussion 89

5 Discussion

GDH of the biotechnologically highly relevant actinomycete C. glutamicum was already the

subject of biochemical investigation decades ago (Kimura, 1962; Oshima et al., 1964; Shiio

& Ozaki, 1970; Shiio & Ujigawa, 1978). These studies were driven by the commercial

interest in the production of glutamate as a flavor enhancer. But despite this constant

attention, comparably little is known about regulation of GDH, besides data on the

biochemical properties. This goes together with a lack of information on the influence of this

central enzyme on carbon and nitrogen metabolism as well as on redox fluxes. Therefore,

within this work several approaches were chosen to gather information on GDH. With

respect to its position at the interface between nitrogen and carbon metabolism, the GDH

enzyme was closer characterized bearing in mind the biotechnological importance of

C. glutamicum. Furthermore, contradictory results on a possibly nitrogen-dependent

regulation of gdh expression led to the reinvestigation of gdh transcription accompanied by

a detailed examination of putative regulators involved.

5.1 Characterization of GDH in the context of systems biology In order to gain information concerning the role of GDH moving away from plain studies on

the single enzyme, the GDH reaction is included in a systems biology-based approach on

modeling the TCA cycle in C. glutamicum. The TCA cycle is the major cellular donor of

carbon backbones and reductive equivalents and therefore of crucial importance for the

synthesis of amino acids. Because of its outstanding position, currently regulation of this

metabolic pathway is investigated in respect of transcriptional control, metabolome and on

protein level. Subsequent modeling approaches can be considered fundamental work for

applying systems biology-based analyses later to the whole metabolism. The emerging field

of systems biology describes the investigation of a whole cell concerning structure as well

as dynamics of the metabolism moving away from focusing only on the response of single

pathways to metabolic changes (for review, Kitano, 2002). In order to establish analytical

methods, this very complex set-up is so far limited to the TCA cycle of C. glutamicum.

Generally, modeling the complex regulatory network of the cell is a challenge which still

needs to be overcome, so that currently simplification of mathematical modeling is priority

(Bornholdt, 2005). However, the perspective of integrating global analyses techniques of

systems biology approaches and strain improvement (Wang et al., 2006), for instance by

metabolic engineering of biotechnologically relevant organisms, is promising.

Discussion 90

The GDH reaction is included in these studies on the TCA cycle in C. glutamicum, because

it is located at an important branch-point of metabolism. The substrate 2-oxoglutarate can

either be oxidized in the TCA cycle by the oxoglutarate dehydrogenase complex or is

aminated to form glutamate. Furthermore, the GDH reaction uses more than 50 % of

NADPH available in the cell (Marx et al., 1999). Taking into account this crucial function

within this work different experimental set-ups were chosen to collect data. These included

characterization of the purified enzyme and examination of GDH in different strains to

detect possible changes in transcript level, protein amount, or enzyme activity. Also,

preliminary work to investigate the influence of differential GDH activity on the metabolism

was performed. The purification of GDH after overexpression in C. glutamicum is the

prerequisite for kinetic measurements as well as quantification of GDH using Western blots.

The latter technique can be applied in addition to the established method of activity

determination by spectrometrically measuring changes in NADPH concentration (Meers &

Tempest, 1970). Despite the challenging procedure of overexpression and subsequent

purification described within this work, the protocol to obtain active, purified GDH was

successfully established, so that the foundation for future projects has been laid.

Furthermore, GDH was investigated using well-established techniques such as the activity

measurements just mentioned, Dot blot analyses and Western blots. For these approaches,

lysine-producing strains and the wild type were compared. These kinds of studies on the

influence of metabolic changes due to defined mutations are a useful basis for monitoring

system dynamics. It was demonstrated that mutations leading to enhanced lysine

production and therefore resulting in metabolic changes did not have an effect on the

activity, protein, and transcript level of gdh. However, changes in cellular fluxes due to

differences in the genetic repertoire do not always exhibit an influence on transcriptome and

proteome, but may lead to variation in concentrations of numerous metabolites (Wang et

al., 2006). Studies on the relation between gene expression levels and metabolic fluxes

during lysine production indicated that there was not necessarily a correlation occurring

(Krömer et al., 2004). This demonstrates the importance of combining studies on all levels

to get a coherent model of cellular processes.

As another possibility to gain knowledge on the connection between GDH, the TCA cycle

and maybe amino acid production, gradual expression of the enzyme was considered. Due

to its position at the interface between nitrogen metabolism and the TCA cycle and because

of the extensive use of NADPH in glutamate synthesis, an impact on the metabolism can be

implied. This assumption was underlined by flux measurements in the lysine-producing

strain MH20-22B (Schrumpf et al., 1992; Marx et al., 1999). After deletion of the gdh gene,

Discussion 91

plasmid-encoded NADPH-dependent GDH of C. glutamicum and an NADH-dependent

GDH of P. asaccharolyticus, were expressed, respectively. One expectation was that this

exchange would potentially lead to an increased NADPH availability for lysine production.

As a result of exchanging the NADPH-consuming enzyme, the cellular NADPH demand

decreased. However, this was accompanied by a drastically reduced generation of the

reductive equivalent, which suggested that lysine production is not limited by the availability

of the reductive equivalent and indicated the flexibility of the cellular responses to metabolic

changes (Marx et al., 1999).

Since the exchange or deletion of respective genes leads to severe alterations of the

metabolism, within this work an expression system for gradual expression of GDH was

established. First attempts were carried out with plasmid encoded gdh under the control of

the tac promoter. Activity measurements and Western blots confirmed the principal

functionality, but due to several drawbacks of the method it remains to be preliminary work.

The tac promoter was shown to be leaky that means expression cannot be turned off

completely. Furthermore, cultures carrying plasmids expressing gdh could not be

considered homogenous. So far, no expression vectors with strong, non-leaky promoters

are available for C. glutamicum. Despite its frequent use, IPTG inducible expression

systems were shown to be disadvantageous due to the low permeability for IPTG observed

in most C. glutamicum strains (Pátek et al., 2003; Nešvera & Pátek, 2008). Bearing in mind

the industrial relevance of C. glutamicum, applying lab scale techniques to industrial

processes needs to be considered. In this context, plasmid-carrying strains might be non-

economical and serve as unstable producers. Risks of spreading resistance genes by

horizontal gene transfer have to be taken into account (Nešvera & Pátek, 2008). To

circumvent these problems, chromosomal integration of vectors as well as the use of

strains expressing enzymes of interest at different levels due to manipulation of the

respective promoter regions is more favorable. Successful application of this approach was

impressively demonstrated on the example of valine production. By combining

chromosomally integrated promoter mutations, overall flux towards valine was increased

significantly without worrying about the disadvantages of plasmid-carrying strains (Nešvera

& Pátek, 2008).

Even though biotechnological use of plasmids expressing heterologous and homologous

genes has been described, working on integration of mutations into the genome seems

more promising. Approaches chosen include introduction of additional copies of a desired

gene into the genome or fine-tuning of expression by mutagenesis of the promoter region.

Promoters are excellent targets for optimizing fluxes towards a desired product by changing

Discussion 92

transcription levels. It has been reported that due to the interactions of various structural

features of the promoter with RNA polymerase and respective regulators, frequency of

transcription can range over four orders of magnitude (Nešvera & Pátek, 2008).

It would be advantageous to construct strains that express gdh at different levels due to

differences in promoter strength. Fundamental experiments contributing to this were also

part of the work presented and will be addressed in the following section. Successful

application of gradual expression due to chromosomally integrated mutations has been

shown on the example of the gdh promoter already. In order to increase the yield of

glutamate, genetically altered C. glutamicum strains were cultivated under overproduction-

inducing conditions and metabolic flux was monitored (Asakura et al., 2007). Results of this

and other studies revealed the importance of this branch-point of glutamate formation and

TCA cycle (Schultz et al., 2007). Enhanced glutamate formation correlates with less activity

of the oxoglutarate dehydrogenase complex. Consequently, the deletion of the odhA gene,

encoding the E1 subunit, led to an enhanced glutamate formation, which was accompanied

by intracellular accumulation of 2-oxoglutarate. To restore the metabolic flux between TCA

cycle and glutamate formation, strains with higher GDH activity were constructed by

mutagenesis of the promoter leading to an increased yield of glutamate. Astonishingly, it

was shown in the same study that a very high GDH activity caused the opposite effect,

namely a reduction of glutamate production. This was probably due to changes in the

metabolic balance (Asakura et al., 2007). The effect of imbalance of metabolic fluxes due to

significant changes in enzyme activity leading to a severe growth defect has also been

reported for Klebsiella aerogenes after overexpression of gdhA encoding an NADPH-

dependent GDH. A single nucleotide exchange led to the creation of a stronger promoter

and enhanced GDH activity. This presumably resulted in succinylCoA limitation within the

TCA cycle, because of a higher demand of 2-oxoglutarate in the GDH reaction (Janes et

al., 2001).

By showing the drastic metabolic effects occurring from changes in protein levels, these

results call for an expression system that allows slight variations in gene expression instead

of knock out or strong induction. Besides insights into systems dynamics, this might

eventually allow fine-tuning of enzymes to possibly increase production rates. Predictions

about reasonable interventions could be made based on data collected by systems biology-

based approaches resulting in the construction of improved production strains. Therefore,

currently fundamental work is being done in this field, so that acquired knowledge might be

used in the future.

Discussion 93

5.2 Transcriptional regulation of gdh Transcriptional control was not only analyzed in order to gain knowledge eventually

applicable for strain construction, due to contrary results reported in the past, nitrogen-

dependent gdh transcription was reinvestigated as well. Activity measurements and

transcriptome analyses suggested constant activity and transcript level irrespective of the

nitrogen availability (Tesch et al., 1999; Beckers et al., 2005; Silberbach et al., 2005a).

Other studies reported increase in activity, protein level as well as transcript level upon

nitrogen deficiency (L. Nolden, unpublished results; Müller, 2005). In the course of the latter

studies, putative transcriptional regulators were found. Since intense investigations did not

lead to the identification of the related stress conditions and therefore did not succeed in

creating a coherent model of transcriptional regulation of gdh, this work mainly focused on

the gdh promoter itself.

Promoter studies in C. glutamicum have become more and more common within the last

decade. Based on a collection of over 50 housekeeping promoters, consensus motifs for

the -35 and -10 region of the σA-dependent promoter were identified (for review, Pátek et

al., 2003; Pátek, 2005; Nešvera & Pátek, 2008). While the -10 region was shown to be

highly conserved, the -35 region was considered to be marginal. This lack of conservation

often seems to be compensated by a 5’-TG-3’ extension of the hexameric -10 region, which

is then additionally recognized by domain 3 of the σ subunit of RNA polymerase (Browning

& Busby, 2004). For the C. glutamicum extended -10 region, the following consensus

sequence was identified: TgtG(c/g)TAtAATGG. Capital letters indicate a conservation

of more than 40 % (Pátek, 2005). On the example of E. coli promoters, occurrence and

function of the 5’ extension of the -10 hexamer, which apparently was found in 20 % out of

554 analyzed promoters, have been studied intensely. Here, it was also suggested that lack

of conservation within the core promoter is compensated by this conserved region. Indeed it

was proven that a promoter with an extended -10 region, but lacking -35 region, nearly

contained the same amount of information as one consisting of -10 and -35 region (Mitchell

et al., 2003; Shultzaberger et al., 2007). The detailed knowledge acquired from studies on

E. coli promoters was applied to explain mechanisms of transcription and its regulation in

C. glutamicum. Therefore, for this organism a rising amount of knowledge on promoters

and their structures has been collected as well (for review, Pátek et al., 2003; Pátek, 2005;

Nešvera & Pátek, 2008). Obtained information was applied in this study to characterize the

gdh promoter. It was possible to experimentally prove the predicted -10 region of the gdh

promoter by mutagenesis and demonstrate the significance of the 5’ extension for promoter

activity. Similar experiments of exchanging 5’-TG-3’ to 5’-CC-3’ have been performed with

Discussion 94

different E. coli promoters leading to a significant decrease in promoter activity as well

(Mitchell et al., 2003). That means this important feature is conserved in the core promoters

of different organisms.

Furthermore, it was possible to use the technique of site-directed mutagenesis to construct

promoters varying in activity. Comparison of C. glutamicum promoters allowed determining

the grade of conservation of different nucleotides within the -10 region. Based on this,

mutagenesis of the dapA promoter had been performed before proving the success of this

approach (Vašiková et al., 1999). By exchanging the C at the first position of the -10 region

of the gdh promoter to T, which was shown to be conserved in 80 % of 44 promoters

(Vašiková et al., 1999), the gdh promoter resembled the deduced consensus sequence and

activity was strongly enhanced. Exchange of the 5’ extension and of the T at the last

position, which was shown to be conserved in over 90 % of 44 promoters (Vašicová et al.,

1999), resulted in less promoter activity. So put together, with this approach fundamental

work for integration of mutations changing promoter activity into the genome has been

carried out successfully. Using knowledge acquired by reporter gene assays, it will be

possible to create strains expressing gdh at different levels avoiding the drawbacks of

plasmid-encoded expression mentioned above.

The application of this kind of genetic manipulation in connection with improvement of

amino acid production has been shown in detail by Asakura and coworkers (2007). As

mentioned above, in this study GDH activity was enhanced by introducing a stronger gdh

promoter into the genome. It was demonstrated in the same study that changes in enzyme

level and activity may lead to metabolic perturbations which are difficult to predict (Asakura

et al., 2007). This fact points out the importance of the systems biology approach

mentioned before. Detailed collection of data might allow prediction of this kind of

phenomenon. Furthermore, strains expressing gdh and additional genes encoding enzymes

of central metabolism differently might be used to understand metabolic fluxes especially

with respect to production of amino acids other than glutamate. Another interesting fact was

brought up by these mutagenesis studies of the gdh promoter (Asakura et al., 2007).

Despite the assumption that the -35 region plays a less important role in transcription

initiation in C. glutamicum, it was proven that mutagenesis leading to sequences

resembling the derived consensus motif ttGc/gca resulted in a significant increase in

promoter activity (Asakura et al., 2007). In the case of the gdh promoter, the -35 region

seems to be involved in interaction with RNA polymerase. Thereby, the conclusion that the

relative contribution of the respective promoter elements differs from promoter to promoter

was shown to be true (Browning & Busby, 2004). It points out the difficulty of making

Discussion 95

predictions solely based on sequence analyses. The dimension of this statement became

obvious during mutational analyses of the gdh promoter within this work as well. Despite an

expected inactivation of the promoter as it has been shown in E. coli or on the example of

the C. glutamicum dapA promoter (Vašiková et al., 1999; Mitchell et al., 2003) residual

activity of 38 % was retained after introduction of the respective mutations. By subsequent

elimination of possible reasons, this question was eventually answered. Interaction of RNA

polymerase with a putative UP element and activation of the promoter by DNA bending

were excluded. Consequently, interaction with the mutated extended -10 region, the -35

region or additional promoter were taken into account. By construction of further promoter

fusions the first two possibilities were ruled out, so that subsequent studies focused on the

identification of an additional transcriptional start site downstream of G284 (Börmann et al.,

1992). Localization upstream of the known promoter could be excluded. By performing

primer extension experiments, it was indeed possible to identify a transcriptional start site at

G195 in addition to the known start point. Therefore, the residual activity of the intentionally

inactivated gdh promoter could easily be explained. Despite the presence of the newly

identified transcriptional start site, attempts to define respective elements of the σA-

dependent core promoter by sequence analyses failed. Only a sequence resembling the

consensus motif for a -35 region was found. The spacing with 19 nts instead of 17 nts to the

possible -10 hexamer is quite unusual (M. Pátek, personal communication; Nešvera &

Pátek, 2008). The possible impact of optimal spacing of -10 and -35 region on promoter

activity became obvious during studies on gdh promoters in S. typhimurium and

K. aerogenes. Mutations that led to a spacer length closer to the 17 nts of the consensus

housekeeping promoter were thought to be responsible for a significant increase in GDH

activity (Janes et al., 2001; Yan, 2007). Therefore, especially when examining promoters

which do not contain an extended -10 region the correct spacing has to be taken serious.

The putative -10 hexamer in the vicinity of G195 has a weak degree of conservation

compared to the consensus motif of the housekeeping promoter, so that mutational

analyses are indispensable to confirm the -10 region. Another possibility, which will be

addressed later on, is transcription starting at G195 from a not σA-dependent promoter

involving alternative sigma factors.

Interestingly, the newly identified transcription start is located within the high affinity AmtR

binding site identified by Müller (2005). Comparison of the relationship between AmtR

binding sites and respective transcriptional start sites of nitrogen-dependent genes revealed

that this organization seems to be a common feature (for review, Hänßler & Burkovski,

2008). For amtA and the gltBD operon, it was shown that the transcriptional start site is

Discussion 96

located within the AmtR binding site. Promoters of other nitrogen-controlled genes are

located closely downstream of the respective AmtR binding sites. As mentioned above, in

the case of gdh, nitrogen-dependent transcription has been reported controversially.

However, AmtR binding sites were determined by two independent approaches and in vitro

binding to the gdh promoter region was shown as well (Müller, 2005; Beckers et al., 2005).

Generally, the apparent upregulation of gdh in response to nitrogen deficiency is

astonishing and has so far only been reported for the enzyme of R. flavefaciens, without

providing an explanation for the physiological benefit (Antonopoulos et al., 2003). One

explanation could be a minor contribution to maintain a sufficient glutamate pool. The

importance of keeping the glutamate to glutamine ratio in balance has recently been

pointed out in a study on S. typhimurium (Yan, 2007).

The inconsistency of previous results with the finding that organization of both transcription

start points and respective AmtR binding sites is conserved within the gdh promoter led to

the reinvestigation of nitrogen-dependent transcription. Within this work, reporter gene

assays with an UV-optimized variant of the gfp gene under the control of the gdh promoter

were shown to be a useful tool and were therefore selected as a novel experimental

approach. Using glutamine as nitrogen source, the nitrogen-starvation response was turned

on as could be proven by analyzing transcription of the AmtR-regulated amtA gene. As

observed in real time RT PCR experiments, in reporter gene assays gdh transcription was

enhanced in the absence of ammonium by a factor higher than 2. This was true for a

fragment of the promoter region including both transcriptional start sites and consequently

both AmtR binding sites (398 bp) as well as for a shorter fragment harboring the newly

identified transcriptional start site and one AmtR binding site (276 bp). Therefore, it can be

concluded that gdh is transcribed at a higher level during nitrogen deficiency in

C. glutamicum, despite contrary results mentioned before. Discrepancy between results

from microarray data and reporter gene assays was also described for the hkm gene

encoding a putative histidine kinase. Chloramphenicol acetylase assays demonstrated a

moderate induction of transcription under nitrogen deprivation (Schulz et al., 2001).

Additionally, two AmtR binding sites where identified by bioinformatic sequence analyses

(Beckers et al., 2005). Nevertheless, DNA microarrays did neither confirm hkm as nitrogen-

regulated nor as part of the AmtR regulon. There is a chance that slight alterations of

expression level in response to changes in nitrogen supply are below the detection limit of

the microarrays (Beckers et al., 2005) indicating constrains of this method.

Within this work, it was demonstrated by reporter gene assays with an amtR deletion

mutant that regulation of gdh transcription in the response to the nitrogen supply occurs in

Discussion 97

an AmtR-dependent manner. AmtR-mediated regulation of gdh transcription depending on

the nitrogen availability was shown for the first time in vivo.

The other putative regulator of gdh expression FarR might influence transcription in the

background of the amtR deletion. Transcription from the gdh promoter was monitored in the

wild type and in the amtR deletion mutant under nitrogen surplus. Comparison was drawn

between a 527 bp fragment with both AmtR binding sites and a FarR binding site and a

shorter fragment including one AmtR binding site. In the wild type, promoter activity was

about 2-fold higher when both transcriptional start sites were present, while in the deletion

mutant, promoter activity was at the same level irrespective of the fragment size. Resulting

from this, transcription from the longer fragment was induced only about 2-fold in the amtR

deletion mutant compared to the wild type. While a 4-fold induction was measured for the

shorter fragment. Since it has been demonstrated that FarR posses the capacity to repress

transcription from the gdh promoter (Müller, 2005), it can be assumed that the constant

level of promoter activity in the amtR deletion mutant was due to binding of FarR to the 527

bp fragment of the gdh promoter under ammonium surplus. However, this function was only

observed in an amtR deletion mutant. Experiments in a wild type background could not

verify an effect of FarR on gdh transcription. The most plausible explanation for this

observation might be alterations in metabolite concentrations due to the amtR deletion. For

GntR type regulators ligand-influenced DNA binding has been reported (Aravind &

Anatharaman, 2003), so that changes in the intracellular concentration of metabolites might

exhibit substantial influence on the FarR binding behavior. Metabolome analyses indeed

suggested alterations in response to the amtR deletion (S. Buchinger & J. Reihlen,

unpublished results). In order to confirm this assumption, respective GFP measurements

with shorter fragments of the gdh promoter in an amtR deletion mutant could be performed.

Alternatively, distinct mutations could be introduced into the FarR binding site preventing

binding of the regulator. The results presented suggest that FarR might play a role in

regulation of gdh transcription, whereas the respective stress conditions still need to be

determined.

In the course of the studies on AmtR-mediated regulation of gdh transcription hints were

obtained that the affinity of the binding site located downstream is higher compared to the

second AmtR binding site (Müller, 2005). For the 276 bp fragment harboring only the high

affinity binding site a discrepancy in promoter activity was observed between nitrogen

deprivation and activity in an amtR deletion mutant. Transcription from the gdh promoter

was enhanced in the complete absence of AmtR compared to cultivation with glutamine.

This indicates that due to the higher affinity, AmtR-mediated repression was not fully

Discussion 98

released when glutamine served as nitrogen source compared to the deletion mutant. It

was proposed that the presence of two AmtR binding sites in the gdh promoter region is

primarily beneficial for increasing the local concentration of repressor (Müller, 2005; Müller-

Hill, 2006). However, since an additional transcriptional start site was identified this seems

not to be the main function.

In the course of the studies on the gdh promoter and transcriptional regulation, the function

of OxyR was reinvestigated. During DNA affinity purification this regulator was isolated from

C. glutamicum cell extract after cultivation under nitrogen starvation (Müller, 2005). Despite

the lack of information concerning a respective binding site, nitrogen-dependent

transcription of gdh in the absence of OxyR was investigated. Additionally, amtA

transcription was analyzed in an oxyR deletion mutant. From the results obtained, it can be

concluded that OxyR does not play an important role in regulating nitrogen-dependent gene

expression. This is the case for gdh as well as for the AmtR-regulated amtA gene. Reasons

for nitrogen-dependent binding therefore remain unknown. The possibility that FarR and

OxyR might regulate expression of the glxK gene located in a divergent orientation

upstream of the gdh gene was ruled out based on experiments performed as part of this

work. In measurements with inverted fragments of the gdh promoter, no activity was

detectable. Therefore, the glxK promoter seems not to be located within the investigated

fragments. Furthermore, this generally applied control experiment excluded the presence of

antisense RNA upstream of the gdh gene. Antisense RNAs have mainly been studied in

enteric bacteria and have been shown to be involved in post-transcriptional control

(Udekwu & Wagner, 2007). However, it seems that predictions are hard to make in the first

place, since chromosomally encoded small regulatory RNAs often display limited

complementarity and are non-contiguous (Udekwu & Wagner, 2007).

Besides transcriptional control mediated by regulator proteins, there are further

mechanisms that are able to influence gene expression. Very well investigated, especially

in the model organism E. coli, is transcription starting from promoters which are recognized

by different sigma (σ) subunits of RNA polymerase. Generally, it is discriminated between

housekeeping sigma factors responsible for recognition of promoters that are expressed

during exponential growth and alternative sigma factors required for expression of distinct

genes in response to a certain stress condition. An example of a well characterized stress

response is the σ54-dependent expression of ntr genes in E. coli, even though structural

features and the desperate need for an activator make σ54 an exception. However,

Discussion 99

homologs of σ54 are found in many microorganisms (for review, Kustu et al., 1989;

Browning & Busby, 2004).

In C. glutamicum, seven sigma factors are present named after the respective homologs in

M. tuberculosis. The primary sigma factor SigA was shown to be essential and responsible

for transcription from housekeeping promoters during exponential growth (Halgasova et al.,

2001). SigB is described as primary-like sigma factor involved in gene expression in

response to several stress conditions and the transition form exponential to stationary

phase (Halgasova et al., 2001; Larisch et al., 2007). The remaining sigma factors SigC,

SigD, SigE, SigH, and SigM are so-called extracytoplasmic-function (ECF) factors. Besides

housekeeping promoters in C. glutamicum, so far only promoters recognized by SigB, SigH,

and SigM have been described (for review, Nešvera & Pátek, 2008). The respective genes

belong for instance to the regulon of heat shock response, disulphide stress response or

are expressed during transition and stationary phase (Engels et al., 2004; Larisch et al.,

2007; Nakunst et al., 2007). As for instance described for genes encoding Clp protease

subunits, housekeeping promoter and the promoter recognized by an alternative σ factor

are located adjacent to each other and elements of the core promoter overlap (Engels et

al., 2004). Similar promoter organization has been observed for SigM controlling expression

of genes involved in disulphide stress (Nakunst et al., 2007).

In C. glutamicum σ factor-dependent regulation of nitrogen control has so far been

excluded, but based on results obtained in this work should be taken into consideration with

respect to gdh transcription. Primer extension experiments suggested adjacent start sites at

G285 and T286 which might be the result of overlapping promoters. Furthermore, the lack

of a clear -10 region next to the newly identified transcriptional start site at G195 underlines

the assumption that σ factors other than SigA might be involved in transcription initiation. To

get some first ideas, gdh transcription in mutants lacking functional sigma factors was

investigated. However, under the tested conditions no influence was observed. One reason

could be that alternative σ factors are generally more abundant in response to changes in

environmental conditions as for instance described for SigB (Larisch et al., 2007).

Therefore, under the investigated conditions respective σ factors might not exhibit a

regulatory function. A low basal level of transcription of σ factors, might explain the

transcript detected in the primer extension experiments though (for review, Nešvera &

Pátek, 2008). In order to get clear results about a possible participation of a certain sigma

factor in regulation of gdh transcription, further experiments need to be performed as for

instance reporter gene assays in mutants lacking functional σ factors.

Discussion 100

Also, when talking about mechanisms involved in transcriptional regulation, stringent

response and the alarmone (p)ppGpp should not be ignored. In response to amino acid

starvation, (p)ppGpp is synthesized and by interaction with RNA polymerase is responsible

for gene expression according to the changing environmental conditions. C. glutamicum

posses a bifunctional guanosine pentaphosphate synthetase (Rel), which is essential for

(p)ppGpp synthesis (Wehmeier et al., 1998). Transcriptome analyses of a rel deletion

mutant revealed that genes of nitrogen metabolism were regulated in a negative Rel-

dependent fashion. Besides genes of the AmtR regulon such as amtA and glnK, gdh was

also under Rel-dependent stringent control. It remains to be elucidated whether this

regulation can be ascribed to a direct influence or interaction of AmtR with (p)ppGpp or if

nitrogen-dependent promoters in general are sensitive to (p)ppGpp-dependent repression

(Brockmann-Gretza & Kalinowski, 2006). However, this influence on nitrogen-dependent

genes mediated by another global regulatory network, points out that some aspects of well-

investigated nitrogen control are still left out. Especially, with respect to open questions

concerning gdh transcription, regulation by stringent control should be kept in mind. In

connection with control of GDH in E. coli, participation of SpoT-regulated (p)ppGpp

synthesis was suggested (Saroja & Gowrishankar, 1996). However, regulation of GDH as

well as stringent response differs substantially in enteric bacteria and C. glutamicum.

Within this work it was possible to gain further information on the gdh promoter. In addition

to the determination of a novel transcriptional start site, AmtR-regulated, nitrogen-

dependent transcription of gdh could be demonstrated for the first time in vivo. Investigation

of the promoter region laid the foundation for further studies on mechanisms controlling gdh

expression as for instance alternative sigma factors. Considering future projects on

constructing promoters varying in strength, preliminary work on gradual expression has

been carried out successfully. This approach would allow great flexibility and therefore in

general can be considered a powerful tool for changing expression for a possible

biotechnological application.

5.3 Identification of FarR and AmtR target genes In addition to studies on regulation of gdh, the transcriptional regulators apparently involved

in controlling gdh expression were closer analyzed. For the global regulator of nitrogen

metabolism AmtR as well as for the less investigated regulator FarR, novel target genes

were identified. As mentioned before, the AmtR regulon has been studied in great detail,

which led to the identification of at least 36 regulated genes mainly involved in nitrogen

Discussion 101

metabolism (Beckers et al., 2005). For FarR, the second putative regulator of gdh

transcription, no detailed knowledge was available. It is a HutC/FarR-type regulator of the

GntR family that has not been characterized in C. glutamicum. DNA microarrays and

sequence analyses suggested additional FarR target genes besides gdh (Müller, 2005),

which were closer examined in the course of this work.

As putative FarR targets, arginine biosynthesis genes were suspected. Arginine is

synthesized from its precursor glutamate in eight enzymatic steps. It was demonstrated that

genes encoding the respective enzymes form an operon, which additionally includes the

argR gene encoding a putative regulator of arginine biosynthesis. By in vitro binding studies

interaction of FarR with the upstream region of the first gene in the operon argC and of the

argG gene was shown. In the case of argG, it was possible to more precisely determine the

FarR binding site. These results indicated that the changes of expression observed in DNA

mircroarrays (Müller, 2005) were due to direct interaction with the respective upstream

regions and a secondary effect can be excluded. However, determination of the exact

binding site was complicated because of varying distances of conserved nucleotides within

the binding site. The search was performed based on two putative binding sites determined

in the gdh and dtsR2 upstream region (Müller, 2005). Repeated bioinformatic screening

with the two previously identified motifs and the newly identified motif upstream of argG did

not lead to coherent results (J. Kalinowski, personal communication). Therefore, further

target genes besides gdh, dtsR2 encoding a detergent sensitive rescuer protein, and the

arg operon were not found. The physiological reason for regulation of dtsR2 in this context

remains unknown. It was proposed that the respective gene product might play a role in

glutamate production due to its influence on the fatty acid composition of the cell membrane

(Kimura, 2002; Müller, 2005). However, this was shown to be unlikely using transcriptome

analyses to investigate the molecular mechanisms of glutamate overproduction (Kataoka et

al., 2006).

Since the stimulus for FarR-mediated regulation was not known, it was tested which

conditions would lead to alterations in expression of the arg operon. Upregulation under

nitrogen-starvation was demonstrated and an involvement of the global regulator AmtR was

excluded due to results of different experimental approaches. Gel retardation experiments

could not confirm AmtR binding and previous studies including bioinformatic screening,

DNA microarrays or proteome analyses neither gave rise to the assumption that the arg

operon might be transcribed at higher levels under nitrogen deficiency nor indicated the

presence of a respective AmtR binding site (Beckers et al., 2005; Silberbach et al., 2005a).

Discussion 102

Generally, genes of biosynthetic pathways are regulated depending on the presence of the

final product. This was also shown as part of this work for genes of the arg operon in the

presence of arginine. Arginine metabolism has been investigated in great detail for a variety

of different organisms. Its regulation seems to be conserved in a wide range of

microorganisms and in all of them the regulator ArgR/AhrC and the respective DNA binding

motifs designated ARG boxes plays a key role in transcriptional control. In E. coli,

transcription of arginine biosynthesis genes is repressed by ArgR in the presence of

arginine. Furthermore, transcription in dependence of the alternative stationary phase

sigma factor RpoS was shown, giving an example of interacting regulatory networks (Maas,

1994; Weerasinghe et al., 2006). The arg cluster of B. subtilis is regulated from an arginine-

dependent promoter by the AhrC protein, which has been shown to exhibit activating as

well as repressing activity (Czaplewski et al., 1992). A mechanism of transcriptional control

involving activation as well as repression by two regulators seems to be widely distributed

in low G+C Gram-positves. On the example of Lactococcus lactis regulation by the two

homologs ArgR and AhrC has been described (Larsen et al., 2005).

The argR gene located within the C. glutamicum arg operon is annotated as arginine

repressor. The respective protein shares more than 30 % amino acid identity with the

respective regulators of E. coli and B. subtilis (J. Amon, personal communication).

Concerning the function of ArgR in regulating expression of the arg operon, binding studies

were performed with and without addition of arginine. Under the tested conditions

interaction with the respective upstream regions could not be observed. It was taken into

consideration that maybe ArgR does not exhibit a regulatory function due to the additional

participation of FarR. On the example of the ure operon encoding urease, it was

demonstrated that an adjacently located regulator (UreR) does not necessarily exhibit a

regulatory function (Beckers et al., 2004). So this might be a common feature in

C. glutamicum. However, recent data from transcriptome analyses of an argR deletion

strain disagree with this hypothesis. It was clearly shown that the deletion of the argR gene

was followed by an enhanced transcript level of the arg genes. For argC, argF, argG, and

carAB results were verified by real time RT PCR. Therefore, it seems that genes encoding

enzymes responsible for the arginine biosynthesis are repressed by argR. Repression was

shown to be even stronger in the presence of arginine (A. Hüser, personal communication).

Therefore, together with data from this work, it can be assumed that in C. glutamicum

transcription of the arg operon is directly regulated by ArgR, especially in the presence of

arginine. However, a second regulator, FarR represses transcription of the operon in

response to a yet unknown stimulus. Furthermore, the principal capacity to repress gdh

Discussion 103

providing the precursor for arginine synthesis was shown in previous work as well as a

possible repression of carB encoding carbamoylphosphate synthase (Müller, 2005). Put

together, arginine biosynthesis in C. glutamicum is controlled by at least two regulators.

Studies on hierarchical order of regulatory processes still have to be performed and will be

simplified after the identification of the stimulus for FarR-mediated control. In

C. glutamicum, a wide range of regulatory networks have been described starting with

relatively simple feed-forward loops as for instance SigH-dependent regulation of the clp

genes up to multi-input motifs. The latter have been described for instance for regulation of

TCA cycle genes (Engels et al., 2004; Bott, 2007; Brinkrolf et al., 2007). So it might be that

transcriptional regulation of arginine biosynthesis is quite complex.

In addition, the presence of regulatory mechanisms besides transcriptional control by FarR

has been suggested based on data indicating that the internal arginine concentration was

not significantly influenced by the farR deletion (Müller, 2005). Besides transcriptional

control, regulation by feedback inhibition of acetylglutamate kinase (encoded by argB) was

shown to be involved (Udaka, 1966; Sakanyan et al., 1996). The presence of a putative

second promoter upstream of argB located within the operon gives rise to the assumption

that genes of the operon are regulated independently as well (Sakanyan et al., 1996). A

similar organization of an operon structure additionally including internal promoters has

been described for the gap-pgk-tpi-ppc gene cluster (Schwinde et al., 1993). Different

transcript levels observed for arg genes during studies on transcriptional control might be a

result of this.

Even though arginine biosynthesis has been examined to little extend so far in

C. glutamicum, this work could provide some first information on a putative regulator

involved as well as on experimental conditions that lead to changes in gene expression. In

addition to the putative regulation of gdh transcription (Müller, 2005), information on the

possible function of the so far not characterized regulator FarR was gathered.

Compared to the minimal amount of information available on FarR, the regulon of AmtR has

been investigated in great detail using different global approaches (Beckers et al., 2005). A

repeated comparison of the transcriptome of an amtR deletion mutant and the wild type led

to identification of two new putative AmtR target genes. Hints obtained by independent

approaches, namely DNA affinity purification and sequence analyses, underlined these

assumptions. For one of the genes dapD, encoding tetrahydrodipicolinate succinylase,

nitrogen-dependent regulation has been observed before (Silberbach et al., 2005b),

whereas this was not the case for mez encoding malic enzyme.

Discussion 104

The dapD gene product catalyzes the first of four steps of the so-called succinylase variant

of diaminopimelate synthesis. In a parallel pathway, diaminopimelate is formed from

tetrahydrodipicolinate and ammonium by the diaminopimelate dehydrogenase (encoded by

ddh) variant. Between these two split pathways differences concerning the activity

depending on the nitrogen source were observed. The succinylase variant is necessary for

the use of organic nitrogen compounds. The dehydrogenase variant seems to be the low

affinity branch, since the deletion of the dapD gene could only be compensated by

diaminopimelate dehydrogenase under high ammonium concentrations and not in the

presence of glutamate as nitrogen source (Wehrmann et al., 1998). In accordance with this,

transcriptome analyses after cultivation in a chemostat performed by Silberbach and

coworkers (2005b) showed that expression of dapD and ddh was regulated depending on

the ammonium availability. Under ammonium limitation, expression of ddh was repressed

whereas dapD transcript level and protein level increased.

Within this work, the assumption that AmtR might be involved in regulating dapD

expression was confirmed. Real time RT experiments verified results obtained by DNA

microarrays. The predicted AmtR binding site in the dapD upstream region was

experimentally proven as well. Therefore, by reinvestigation and application of a

combination of different methods a new AmtR target gene was identified. By results

presented in this work, the mechanism of nitrogen-dependent diaminopimelate synthesis

can be explained. Under ammonium surplus mainly the dehydrogenase variant is active,

while decrease in ammonium concentrations leads to a release of AmtR-mediated

repression of dapD and consequently synthesis of diaminopimelate via the succinylase

variant. When monitoring flux distribution of lysine production during fermentation, it

became obvious that there was a variation of pathway use correlating with cultivation time

and ammonium content. While in the beginning most of the lysine was made by the

dehydrogenase variant, later on the high affinity succinylase variant was used. In total 66 %

of lysine was formed by the latter pathway (Sahm et al., 2000). The involvement of AmtR in

regulation of diaminopimelate synthesis, explains also the increased lysine concentration

observed in an amtR deletion mutant (S. Buchinger & J. Reihlen, unpublished data). That

means with dapD not only a new, but also very interesting AmtR target gene was identified.

Less is known about the reasons for AmtR-mediated regulation of mez expression. The

mez gene encodes malic enzyme, which catalyzes the decarboxylation of malate to

pyruvate accompanied by NADPH generation. However, its role in the metabolism of

C. glutamicum is not yet fully understood (Georgi et al., 2005). Its function has been

investigated regarding the utilization of different carbon sources and in respect of lysine

Discussion 105

production (Gourdon et al., 2000; Georgi et al., 2005). It was proposed that due to its

function in NADPH generation, overexpression would be beneficial for lysine production

when using fructose or a glucose/fructose mix as carbon source (Dominguez et al., 1998).

This could not be confirmed by overexpression studies in lysine-producing strains (Georgi

et al., 2005). As in the case of dapD, reinvestigation of the transcriptome of an amtR

deletion mutant revealed an enhanced level of mez transcript in the absence of AmtR. In

accordance to this, the regulator was isolated by DNA affinity purification with a fragment of

the mez upstream region (V. Wendisch, personal communication). In this work, it was

verified that the changes in expression level are the result of direct interaction between

AmtR and the mez upstream region. By this independent approach, the localization of the

AmtR binding site could be narrowed down to 50 bp. Based on sequence analyses a

putative binding motif was determined. Probably due to a low similarity to the consensus

motif (Beckers et al., 2005), it had not been identified before. However, to confirm this

binding site mutational analyses and binding studies as described for the binding sites

upstream of amtA (Jakoby et al., 2000) should be performed. The putative AmtR binding

site is located 79 bp to 104 bp upstream of the mez start codon right in the vicinity of the

transcription start site located 70 bp upstream of the start codon (V. Wendisch & J-W.

Youn, personal communication). That means, the binding site overlaps with key structures

of the core promoter and by doing this probably interferes with RNA polymerase

recognition. This organization seems to be conserved for AmtR binding sites, because they

tend to be located upstream of the respective transcription start or in the close

neighborhood (for review, Hänßler & Burkovski, 2008). Even though direct regulation of

mez expression by AmtR could be proven within this work, the physiological role of AmtR-

mediated regulation remains unknown.

In contrast to previously performed DNA microarrays (Beckers et al., 2005) recent

experiments were of a better quality marked by a higher sensitivity and an increase in

dynamic range (S. Hans, unpublished data). Even in these improved experiments, the

regulation factors of dapD and mez of only 2.9 and 3.5, respectively, were quite low

compared to the 137-fold induction of amtB transcription in the absence of AmtR. This

minor level of regulation seems to be one reason for the later discovery of these novel

target genes. In the case of mez the moderate regulation might be due to a low level of

conservation of the respective binding site, but since the AmtR box upstream of dapD is

well conserved, there need to be other features which correlate with the degree of

repression. Localization with respect to the transcriptional start point, number, and length of

the binding sites are discussed as decisive criteria. However, detailed studies on AmtR and

Discussion 106

its interaction with DNA need to be performed to gain detailed understanding. Different

expression levels of nitrogen-regulated genes have also been reported for NRI-P-

dependent transcription in E. coli. While glnA transcript was shown to be present already at

low levels of NRI-P, for the transcription starting form the glnK and nac promoters higher

concentrations of the activator were required. This mechanism of regulation led to

significant differences in the level of transcription as well as in the chronology of expression

(Atkinson et al., 2002). Similar increments in AmtR-mediated expression are imaginable

due to varying affinities observed in gel shift assays as well as differential regulation shown

in transcriptome analyses and real time RT PCR experiments (Müller, 2005; Beckers et al.,

2005).

In summary, it was possible to successfully verify results from DNA microarrays and

thereby prove direct regulation of two novel target genes of the global repressor of nitrogen

metabolism. Despite intense studies on AmtR-mediated nitrogen control (for review,

Hänßler & Burkovski, 2008), by applying a combination of methods dapD and mez were

just recently confirmed as novel members of the AmtR regulon.

5.4 The interface between nitrogen and carbon metabolism Data on transcriptional regulation of gdh and respective regulators involved provide a solid

foundation for future work dealing with detailed investigation on the interplay between

nitrogen and carbon metabolism. While in other organisms such as E. coli or B. subtilis

comparably detailed knowledge is available on interacting regulatory networks of respective

pathways, experimental data from C. glutamicum are rather rudimental. However, emerging

projects on the characterization of TCA cycle enzymes in connection with the GDH reaction

are promising approaches.

So far, in C. glutamicum analyses of the proteome and transcriptome gave hints about the

connection of central metabolism and nitrogen control. It was demonstrated that nitrogen

limitation during cultivation in a chemostat resulted in an enhanced transcription of genes

involved in central metabolism to deal with the increased demand of energy (Silberbach et

al., 2005b). Similar conclusions were drawn from proteome analyses after cultivation

without nitrogen source, which indicated an increased amount of proteins of energy

metabolism in response to nitrogen starvation. This was accompanied by a higher oxygen

consumption rate of the starved cells (Schmid et al., 2000). These results clearly suggest

an interconnection between nitrogen and carbon metabolism. Also, enzymes involved in

nitrogen assimilation, namely GS and GDH have been shown to be regulated not only in

Discussion 107

response to the nitrogen status, but also depending on carbon source and availability

(Schulz et al., 2001; Müller, 2005). In the case of GDH, the influence of intermediates of the

TCA cycle on enzyme activity has been investigated and a slight activation of glutamate

synthesis in the presence of citrate, cis-aconitate and further metabolites was demonstrated

(Shiio & Ozaki, 1970).

Nevertheless, concerning details on the molecular level, basically nothing is known on

regulation of gdh of C. glutamicum in connection with carbon metabolism. This is almost the

same for the enzyme of E. coli. However, in this model organism details on transcription

initiating from σ70- and σ54-dependent promoters have been described for genes encoding

GS and a glutamine transport system (Tian et al., 2001; Mao et al., 2007). Another

prominent example of transcriptional control by global carbon and nitrogen regulatory

mechanisms in cooperation with local regulators is for instance the gltBDF operon.

Participation of Lrp, CAP, and ArgR indicates a crucial function of the respective gene

products linking ammonium assimilation and carbon metabolism (Paul et al., 2007). Even

though glutamate dehydrogenase (RocG) of B. subtilis fulfills a completely different

function, regulatory mechanisms responding to signals of central metabolism as well as to

the availability of amino acids such as arginine have been described. RocG executes a

regulatory function itself by interacting with GltC, the activator of the gltAB operon

(Commichau et al., 2007a; 2007b).

Data collected on GDH of C. glutamicum so far are not sufficient to make proposals on

regulatory mechanisms in response to central metabolism. This work contributed to verify

previous data on nitrogen-dependent regulation and confirmed involvement of the global

repressor of nitrogen metabolism AmtR. Conclusion on the physiological function of this

nitrogen-dependent regulation cannot be drawn at this point. Of great interest in this context

is the apparent participation of AmtR in controlling not only expression of genes directly

linked to nitrogen metabolism. Indirect regulation of vanAB and vanK encoding vanillate

demethylase and a putative protocatechuate transporter has been reported previously

(Merkens et al., 2005) and as part of this work, a direct influence on mez transcription could

be proven. Put together, these results give first indications on possible connections

between regulatory networks of central and nitrogen metabolism. However, the complexity

of these processes has been shown clearly by investigation of ammonium assimilation in

the well-characterized S. typhimurium. In order to reduce complexity of the system to be

examined, only GDH, GS, and GOGAT were included. By this approach remarkable results

concerning regulation of homeostasis of glutamine and glutamate pools were obtained

Discussion 108

(Yan, 2007). Studies limited to a small system provide information for future work on more

complex systems and other microorganisms.

To be able to apply this kind of approach to the metabolism of C. glutamicum, this work

provided data on nitrogen-dependent regulation of GDH as well as information on further

putative regulatory mechanisms involved. Also preliminary work for future application of

techniques to investigate GDH and its possible impact on the metabolism was successfully

performed.

Appendix 109

6 Appendix

6.1 Regulation of glutamine synthetase in corynebacteria

Recent approaches focused on finding a common theme of nitrogen metabolism among

related microorganisms. Within these studies, nitrogen control in corynebacteria with a

published genome sequence was analyzed regarding the presence of genes encoding the

key components involved in regulation, assimilation, and transport processes (Walter et al.,

2007). Besides well-investigated C. glutamicum (Ikeda & Nakagawa, 2003; Kalinowski et

al., 2003), the genetic repertoire of C. diphtheriae (Cerdeno-Tarraga et al., 2003), C.

efficiens (Fudou et al., 2002), and C. jeikeium (Tauch et al., 2005) was compared to find out

about differences in the cellular response to changes in nitrogen availability in the soil-living

amino acid producers and the two human pathogens. Besides transcriptional regulation of

nitrogen-dependent genes, uptake measurements, and search for common AmtR binding

motifs, activity of glutamine synthetase was determined in respect of nitrogen dependency

(Walter et al., 2007).

In each organism, except for C. diphtheriae, a glnA gene encoding the GSI-β type enzyme

responsible for glutamine synthesis and ammonium assimilation is present. The C. efficiens

genome contains even a second copy of this gene. In addition, glnA2 encoding the GSI-α

type enzyme is present in all four genomes. Generally, C. glutamicum and C. efficiens

genomes show a high degree of similarity regarding the presence of genes encoding

components of nitrogen metabolism, while the number of genes involved in nitrogen

metabolism in the two pathogenic organisms is significantly reduced, especially in terms of

transport proteins involved in uptake of alternative nitrogen sources.

In order to get a complete picture of nitrogen control in addition to transcriptional regulation

of nitrogen-dependent genes and ammonium uptake, regulation of glutamine synthetase

was investigated by determination of the glutamyltransferase activity. For this, all organisms

were first cultivated in complex medium. To induce nitrogen deprivation, cultures were

divided and one aliquot was transferred to CgCoN medium. The results of the activity

measurements are summarized in figure 6.1.

Appendix 110

Fig. 6.1: Nitrogen-dependent regulation of glutamine synthetase in corynebacteria. Activity of glutamine synthetase was determined in C. diphtheriae, C. efficiens, C. glutamicum, and C. jeikeium under nitrogen surplus (white bars) and nitrogen deficiency (black bars). For this, glutamyltransferase activity was measured.

The results of the determination of glutamine synthetase activity correlated with the results

derived from the comparison of the respective genome sequences. In crude extract of

C. diphtheriae, glutamyltransferase activity was not detectable. This could easily be

explained by the lack of the glnA gene. The same results were obtained for a

C. glutamicum glnA deletion strain containing only the glnA2 gene (Nolden et al., 2001a;

Ansorge, 2005). Either the applied glutamyltransferase test is not suitable for determination

of the activity of the GSI-α type enzyme in general or an active glnA2 gene product was not

present under the conditions tested. In the related human pathogen M. tuberculosis, the

absence of the glnA1 gene, encoding a GS type I enzyme, was also shown to result in loss

of GS activity despite the presence of three additional genes encoding putative glutamine

synthetases (Tullius et al., 2003). This phenomenon was explained in a subsequent study

which indicated that of the three respective genes transcript was present, but translation did

not occur (Harth et al., 2005). Whether this kind of mechanism is also present in

C. diphtheriae, remains to be investigated.

In C. efficiens and C. glutamicum, an active glutamine synthetase was present. In response

to nitrogen starvation activity increased. Both organisms contain the glnE gene encoding an

adenylyltransferase (ATase) as well as amtR encoding the global repressor of nitrogen

metabolism. That means, a functional nitrogen control as reported for C. glutamicum can be

assumed for C. efficiens as well. Under ammonium surplus, glnA transcription is partially

0 .0 0 0

0 .2 0 0

0 .4 0 0

0 .6 0 0

0 .8 0 0

1 .0 0 0

1 .2 0 0

GS

activ

ity [µ

mol

(mg

prot

ein)

-1m

in-1

]

C. diphteriae C. glutamicumC. efficiens C. jeikeium0 .0 0 0

0 .2 0 0

0 .4 0 0

0 .6 0 0

0 .8 0 0

1 .0 0 0

1 .2 0 0

GS

activ

ity [µ

mol

(mg

prot

ein)

-1m

in-1

]

C. diphteriae C. glutamicumC. efficiens C. jeikeium

Appendix 111

repressed by AmtR and this repression is released as response to the lack of nitrogen

source. Demodification of GS by ATase leads additionally to an increase in activity under

these conditions (Nolden et al., 2001a). The presence of two copies of the glnA gene in

C. efficiens might explain the generally higher level of activity compared to C. glutamicum.

In C. jeikeium, enzyme activity was detectable as well. However, the level of activity

remained unaltered irrespective of the nitrogen availability. It was in the same order of

magnitude as determined for C. glutamicum. The constant level of activity might have been

due to the lack of AmtR in C. jeikeium. However, the presence of the glnE gene suggests

ATase-mediated regulation on the level of activity. Under the conditions tested upregulation

of GS due to nitrogen deprivation as shown for C. glutamicum (Jakoby et al., 1999) was not

detectable, so that the involvement of ATase in post-translational regulation of GS activity in

C. jeikeium is not clear.

By comparing regulation of GS in the four different organisms, it was demonstrated that the

response to changes in nitrogen availability and connected regulatory mechanisms are

conserved among the species. Furthermore, the known phenomenon of gene decay for

pathogenic organisms became obvious, since C. diphtheriae lacks the glnA gene and

C. jeikeium lacks the amtR gene (Walter et al., 2007). By combining bioinformatic

approaches and transcriptional analyses with determination of enzyme activity and uptake

measurements, it was possible to draw a coherent picture of regulation of nitrogen

metabolism in different corynebacteria. It was shown that key components are conserved,

whereas different habitats call for adaptation to environmental conditions going together

with modification of the genetic equipment.

Appendix 112

6.2 Plasmid constructions

pEKExgdh: This plasmid is an expression vector of his-tagged gdh of C. glutamicum. The plasmid

pQE30Xagdh (Müller, 2005) was restricted with EcoRI and PvuII. The resulting insert

including the full-length gdh gene of C. glutamicum and the sequence of an N-terminal

histag was ligated to the EcoRI- and DraI-restricted vector pEKEx2 (Eikmanns et al., 1991)

leading to pEKExgdh.

pEPRpgdh527 This plasmid harbors a fusion of the gdh promoter region to the gfpuv gene and can

therefore be used for reporter gene assays. For subsequent cloning steps, SmaI and XbaI

restriction sites (shown in bold) were introduced into the primer sequence (5’-gcg cgc ccc ggg ggt caa att tct ccg atg tag-3’/5’-gct cta gag ttc cca tct cgg ctg cat g-3’). A 527 bp

fragment of the gdh promoter region was amplified by PCR and ligated to the SmaI-XbaI-

restricted vector pEPR1 (Knoppová et al., 2007) leading to pEPRpgdh527. The cloned

fragment of the promoter region was sequenced.

pEPRpgdh527_1 This plasmid harbors a fusion of the gdh promoter region to the gfpuv gene and can

therefore be used for reporter gene assays. This plasmid was derived from pEPRpgdh527

and carries a modified promoter region. The respective mutation is underlined in the primer

sequence. Mutations were created by two-step PCR using the following primers in a first

reaction: 5’-gcg cgc ccc ggg ggt caa att tct ccg atg tag-3’/5’-ttc aat tat agc agt gtc gc-3’.

The resulting PCR product was used as primer in a second PCR together with the primer

having the following sequence: 5’-gct cta gag ttc cca tct cgg ctg cat g-3’. For subsequent

cloning steps, SmaI and XbaI restriction sites (shown in bold) were introduced into the

primer sequence. The 527 bp fragment of the gdh promoter region including the mutation

was ligated to the SmaI-XbaI-restricted vector pEPR1 (Knoppová et al., 2007) leading to

pEPRpgdh527_1. The cloned fragment of the mutated promoter region was sequenced.




Appendix 113


reaction: 5’-gcg cgc ccc ggg ggt caa att tct ccg atg tag-3’/5’-ttc aat tat ggg ggt gtc gc-3’.











reaction: 5’-gcg cgc ccc ggg ggt caa att tct ccg atg tag-3’/5’-ttc att tat ggc agt gtc gc-3’. The

resulting PCR product was used as primer in a second PCR together with the primer having

the following sequence: 5’-gct cta gag ttc cca tct cgg ctg cat g-3’. For subsequent cloning

steps, SmaI and XbaI restriction sites (shown in bold) were introduced into the primer

sequence. The 527 bp fragment of the gdh promoter region including the mutation was

ligated to the SmaI-XbaI-restricted vector pEPR1 (Knoppová et al., 2007) leading to






reaction: 5’-gcg cgc ccc ggg ggt caa att tct ccg atg tag-3’/5’-ttc att tat ggg ggt gtc gc-3’. The

resulting PCR product was used as primer in a second PCR together with the primer having

the following sequence: 5’-gct cta gag ttc cca tct cgg ctg cat g-3’. For subsequent cloning

steps, SmaI and XbaI restriction sites (shown in bold) were introduced into the primer

sequence. The 527 bp fragment of the gdh promoter region including the mutation was

ligated to the SmaI-XbaI-restricted vector pEPR1 (Knoppová et al., 2007) leading to


Appendix 114

pEPRpgdh527inv This plasmid harbors a fusion of the gdh promoter region to the gfpuv gene and can


restriction sites (shown in bold) were introduced into the primer sequence (5’-gcg cgc tct aga ggt caa att tct ccg atg tag-3’/5’-gcg cgc ccc ggg gtt ccc atc tcg gct gca tg-3’). A 527 bp


restricted vector pEPR1 (Knoppová et al., 2007) in the inverted direction leading to

pEPRpgdh527inv. The cloned fragment of the promoter region was sequenced.



restriction sites (shown in bold) were introduced into the primer sequence (5’-gcg cgc ccc

ggg gaa gag act tca tgc agt tac-3’/5’-gct cta gag ttc cca tct cgg ctg cat g-3’). A 398 bp








reaction: 5’-gcg cgc ccc ggg gaa gag act tca tgc agt tac-3’/5’-ttc att tat ggg ggt gtc gc-3’.







pEPRpgdh398inv This plasmid harbors a fusion of the gdh promoter region to the gfpuv gene and can


restriction sites (shown in bold) were introduced into the primer sequence (5’-gct cta gag

Appendix 115

aag aga ctt cat gca gtt acc-3’/5’-gcg cgc ccc ggg gtt ccc atc tcg gct gca tg-3’). A 398 bp


restricted vector pEPR1 (Knoppová et al., 2007) in the inverted direction leading to

pEPRpgdh398inv. The cloned fragment of the promoter region was sequenced.




ggg tgg tca tat ctg tgc gac ac-3’/5’-gct cta gag ttc cca tct cgg ctg cat g-3’). A 313 bp





therefore be used for reporter gene assays. This plasmid carries a modified promoter

region. The respective mutation is underlined in the primer sequence. Mutations were

created by using a primer containing the desired base exchange (5’- gcg cgc ccc ggg gac

acc ccc ata aat gaa cg -3’/5’-gct cta gag ttc cca tct cgg ctg cat g-3’). The respective

mutations are underlined in the primer sequence. For subsequent cloning steps, SmaI and

XbaI restriction sites (shown in bold) were introduced into the primer sequence. The 298 bp

fragment of the gdh promoter region including the mutations was ligated to the SmaI-XbaI-


fragment of the mutated promoter region was sequenced.




ggg agc att tac cag cct aaa tgc c-3’/5’-gct cta gag ttc cca tct cgg ctg cat g-3’). A 276 bp




Appendix 116




ggg ccg gtt gat gtg aac gca g-3’/5’-gct cta gag ttc cca tct cgg ctg cat g-3’). A 172 bp



fragment of the promoter region was sequenced. pUCargR This plasmid is an expression vector of argR of C. glutamicum. The full-length argR gene of

C. glutamicum was amplified by PCR. For subsequent cloning steps, KpnI and PstI

restriction sites (shown in bold) were introduced in the primer sequences (5’- cgg ggt acc

aga cat gtc cct tgg ctc aac c-3’/5’- gcg cct gca gtt aag tgg tgc gcc cgc tga gt-3’). After

restriction of the PCR product and the expression vector pUC18 (Viera & Messing, 1982),

the argR gene was ligated to pUC18 leading to plasmid pUCargR. The resulting plasmid

was sequenced.

References 117

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Publications 131

Publications

Hänßler, E. & Burkovski, A. (2008). Molecular mechanisms of nitrogen control in

corynebacteria. In: Burkovski, A. (ed.) Corynebacteria: genomics and molecular biology.

Horizon Scientific Press, Norwich, UK, in press.

Hänßler, E., Müller, T., Jeßberger, N., Völzke, A., Plassmeier, J., Kalinowski, J., Krämer, R. & Burkovski, A. (2007). FarR, a putative regulator of amino acid metabolism in

Corynebacterium glutamicum. Appl. Microbiol. Biotechnol. 76, 625-632.

Walter, B., Hänssler, E., Kalinowski, J. & Burkovski, A. (2007). Nitrogen metabolism in

corynebacteria: variations of a common theme. J. Mol. Microbiol. Biotechnol. 12, 131-138.

Abbreviations and units 132

Abbreviations and units

A adenine

ADP adenosine diphosphate

ApR resistant to ampicillin

APS ammonium peroxodisulfate

ATP adenosine triphosphate

ATCC American type culture collection

BCIP 5-bromo-4-chloro-3-indolyl phosphate toluidine salt

BSA bovine serum albumine

C cytosine

CAPS 3-(Cyclohexylamino)-1-propansulfonic acid

CSPD disodium 3-(4-methoxyspiro {1,2-dioxetane-3,2´-(5´-chloro)tricyclo

[3.3.1.13´7]decan}-4-yl)phenylphosphate

DIG digoxigenin

DMSO dimethyl sulfoxide

DTT dithiothreitol

DW dry weight

EDTA ethylendiaminetetraacetic acid

et al. et alii

Fig. figure

G guanine

g 9.81 m s-2

GFP green fluorescent protein

IPTG isopropyl β-D-1-thiogalactopyranoside

KmR resistant to kanamycin

MOPS 3-[N-morpholino]propansufonic acid

NAD(P)H nicotinamide adenine (phosphate) dinucleotide

NBT 4-nitro blue tetrazolium chloride

nt nucleotide

OD600 optical density at 600 nm

orf open reading frame

PAGE polyacrylamide gel electrophoresis

(p)ppGpp guanosine hepta (penta) phosphate

PCR polymerase chain reaction

RT reverse transcriptase

SDS sodium dodecylsulphate

T thymine

Abbreviations and units 133

Tab. table

TAE tris acetate EDTA

TCA tricarboxylic acid

TE Tris EDTA

TEMED N,N,N’,N’-tetramethyl-ethylendiamine

Tris 2-amino-hydroxymethylpropane-1,3-diol

5’UTR 5’ untranslated region

v/v volume per volume

w/v weight per volume

Units Prefixes bp base pairs k kilo

°C degree Celsius m milli

Cps counts per second µ micro

Da Dalton n nano

g gram

kb kilo base pairs

l liter

m meter

M molar

min minutes

s seconds

U unit

V volt

Amino acid nomenclature (IUPAC-IUB,1969) A alanine M methionine

C cysteine N asparagine

D aspartate P proline

E glutamate Q glutamine

F phenylalanine R arginine

G glycine S serine

H histidine T threonine

I isoleucine V valine

K lysine W tryptophane

L leucine Y tyrosine

Regulation of glutamate dehydrogenase in Corynebacterium ......2.1 Corynebacterium glutamicum During...

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