Cloning, characterization and transcriptional analysis of twophosphate acetyltransferase isoforms from Azotobacter vinelandii

Maria Dimou • Anastasia Venieraki •

Georgios Liakopoulos • Panagiotis Katinakis

Received: 30 June 2010 / Accepted: 9 November 2010 / Published online: 21 November 2010

� Springer Science+Business Media B.V. 2010

Abstract Acetate is abundant in soil contributing to a

great extent on carbon cycling in nature. Phosphate ace-

tyltransferase (Pta, EC 2.3.1.8) catalyzes the reversible

transfer of the acetyl group from acetyl-P to CoA forming

acetyl-CoA and inorganic phosphate, participating to ace-

tate assimilation/dissimilation reactions. In the present

study, we demonstrate that Azotobacter vinelandii, a

nitrogen-fixing, free-living, soil bacterium, possesses two

class II phosphate acetyltransferase isoforms, AvPTA-1 and

AvPTA-2, with different kinetic properties. At the acetyl-

CoA forming direction, AvPTA-1 has lower affinity for

acetyl-P and higher affinity for CoA than AvPTA-2 while at

the acetyl-P forming direction; activity was measured only

for AvPTA-1. Quantification of their expression patterns by

RT-qPCR indicated that both genes are expressed during

exponential growth on glucose or acetate and are down-

regulated in the stationary phase. The ammonium avail-

ability during acetate growth resulted in up-regulation of

Avpta-2 expression only. Further, the gene expression

patterns of other related gene transcripts were also inves-

tigated, in order to understand the influence of each path-

way in the assimilation/dissimilation of acetate.

Keywords Acetate � Azotobacter vinelandii � Phosphate

acetyltransferase � RT-qPCR

Abbreviations

P Phosphate

CoA Coenzyme A

RT-qPCR Real-time quantitative polymerase chain

reaction

Acs-A Acetyl-CoA synthetase

Ack-A Acetate kinase

Pta Phosphate acetyltransferase

PoxB Pyruvate oxidase

Introduction

Acetate, a short-chain fatty acid, is abundant in soil and

other environments since it is a product of many fer-

mentative processes [1] and an important intermediate in

the anaerobic degradation of organic matter [2, 3]. Many

bacteria, including the non-symbiotically nitrogen-fixing

A. vinelandii, are able to use acetate as a source of carbon

and energy [4] while it represents the growth substrate for

methane producing archaea as well, contributing greatly

on carbon cycling in nature [5]. The first step in the uti-

lization of acetate is its activation into acetyl-CoA and

subsequently the operation of the glyoxylate cycle as an

anaplerotic pathway [6]. Once acetyl-CoA is made, it is

used to synthesize lipids, can enter the tricarboxylic acid

cycle to generate precursors of amino acids, or can be

oxidized to generate reducing power via diverse pathways

[7–10].

Two pathways are central to acetate catabolism in pro-

karyotes. Both of these pathways activate acetate into

acetyl-CoA, but they do it via different intermediates and in

response to different concentrations of acetate in the envi-

ronment. The first pathway is catalyzed by the acetyl-CoA

synthetase (Acs-A, EC 6.2.1.1) [11] while the second path-

way is comprised of the acetate kinase (Ack-A, EC 2.7.2.1)

M. Dimou � A. Venieraki � G. Liakopoulos � P. Katinakis (&)

Department of Agricultural Biotechnology, Agricultural

University of Athens, Iera Odos 75, 11855 Botanikos,

Athens, Greece

e-mail: bmbi2kap@aua.gr

123

Mol Biol Rep (2011) 38:3653–3663

DOI 10.1007/s11033-010-0478-3

and the phosphate acetyltransferase (Pta, EC 2.3.1.8) [12].

Although both pathways are reversible, hydrolysis of pyro-

phosphate generated by Acs-A prevents the Acs-A from

converting acetyl-CoA to acetate. The role of these path-

ways is to carefully maintain a physiological balance of

acetyl-CoA and free CoA [13–15]. On the contrary, acetate

formation occurs anaerobically during mixed acid fermen-

tation or aerobically when growth on excess glucose inhibits

respiration [15]. Two pathways are known to contribute to

acetate production: the first, assumed to be constitutive, is

the conversion of acetyl-CoA through phosphate acetyl-

transferase and acetate kinase [16, 17] while the second is

the conversion of pyruvate directly into acetate via pyruvate

oxidase (poxB, E.C 1.2.2.2) [18, 19].

Acetate/acetyl-CoA homeostasis is of great importance

to all cells and phosphate acetyltransferase which rapidly

converts acetyl-CoA via acetyl-P to acetate as an overflow

pathway or converts exogenous acetate back to acetyl-CoA

[16, 20], probably plays a central role in maintaining a

balance flux between biosynthesis and energy generation

[14]. Microbial genomes contain two main classes of

phosphate acetyltransferase enzymes [21] while an addi-

tional evolutionary distinct class also exists [22]. Class I

enzymes, which are *350 amino acids in length and are

not allosterically regulated, include enzymes from Clos-

tridium kluyverii [23], Veillonella alcalescens [24], Ther-

motoga maritime [25], Methanosarcina thermophila [26]

and Bacillus subtilis [27]. Class II enzymes, which are

*700 amino acids and allosterically regulated, with the

exception of enzymes from the enterobacteria Escherichia

coli [28, 29] and Salmonella enterica [21], remain less

investigated.

In the present study we present the enzymatic charac-

terization of two phosphate acetyltransferase class II iso-

forms from A. vinelandii. In addition, we examine the

transcript accumulation of acetate metabolism genes of

A. vinelandii under aerobic growth on glucose or acetate

minimal media. Since A. vinelandii is a nitrogen-fixing,

soil bacterium, we also investigate the influence of

ammonium source on the transcript accumulation of the

tested genes.

Materials and methods

Bacterial strains and growth conditions

E. coli XL-Blue1 strain (Invitrogen) was used for the

propagation of recombinant forms of the plasmid pET28a.

E. coli strain BL21 (DE3) (Novagen) was used for the

expression of recombinant proteins. All E. coli strains were

grown in LB medium supplemented with kanamycin.

A. vinelandii was grown at 30�C in LB medium or Burk’s

nitrogen-free salts [30], supplemented with either glucose

at 1% (BG) or sodium acetate at 0.3% (BA) and in Burk’s

salts supplemented with ammonium chloride at 0.1% and

either glucose at 1% (BNG) or sodium acetate at 0.3%

(BNA).

Heterologous expression of AvPTA-1 and AvPTA-2

in E. coli and purification of recombinant proteins

The coding sequences of Avpta-1 (YP_002800579) and

Avpta-2 (YP_002801221) were PCR-amplified from

A. vinelandii genomic DNA. The primers used were

AvPTA.1-F: 50-CCCCCATGGACACTCTATTTCTCGCC

CCCAC-30 with AvPTA.1-R: 50-AAACTCGAGGGCTCC

GACCGGCTGCCCCTG-30 and AvPTA.2-F: 50-CCCTCA

TGAAGACCTTCCTGATCGCCCCCAG-30 with AvPTA.

2-R: 50-AAAAAGCTTGCCGGCGGCGGTTTCGGCCT

G-30, carrying restriction sites for ligation to the pET28a

expression vector. The PCR-fragments were digested with

the appropriate restriction enzymes and cloned to the cor-

responding sites of pET28a, resulting in pET28a-AvPTA-1

and pET28a-AvPTA-2, respectively. The resulting con-

structs were confirmed by sequencing. Synthesis of

recombinant proteins was initiated by addition of 0.5 mM

isopropyl 1-thio-b-D-galactopyranoside when the cultures

reached A600 of 0.6 and continued cultivation for an

additional 4 h at 30�C. Cells were harvested by centrifu-

gation and were disrupted by sonication in Lysis buffer

(50 mM NaH2PO4, 300 mM NaCl, 10 mM imidazole

supplemented with 1 mg/ml lysozyme). Cellular lysates

were centrifuged and the supernatants were used for pro-

tein purification. Recombinant proteins were purified with

Ni–NTA chromatography (Ni2?-nitrilotriacetate, Qiagen)

according to the manufacturer’s instructions. To remove

any imidazole and salts in the collected fractions, fractions

were dialysed against 50 mM Tris–HCl buffer (pH 7.2), for

12 h. The purity of the purified proteins was analyzed by

15% SDS-PAGE electrophoresis.

Phosphate acetyltransferase activity assay

The rate for both the forward (acetyl-CoA forming) and

reverse (acetyl-P forming) directions of the reaction cata-

lyzed by pta were measured at 35�C by monitoring the

change in absorbance at 233 nm concomitant with the

formation or the hydrolysis of the thioester bond of acetyl-

CoA (e = 4.36 M-1), using a 1 cm path-length quartz

cuvette in a HITACHI U-2800 spectrophotometer. The

standard reaction mixture (800 ll) for the forward direc-

tion contained 50 mM Tris–HCl (pH 7.2), 20 mM NH4Cl,

20 mM KCl, the appropriate substrate and a concentration

of enzyme sufficient to yield a linear rate over at least

2 min (approximately 3 lg). The standard reaction mixture

3654 Mol Biol Rep (2011) 38:3653–3663

123

(800 ll) for the reverse direction contained 50 mM Tris–

HCl (pH 7.2), 20 mM MgCl2, the appropriate substrate and

a concentration of enzyme sufficient to yield a linear rate

over at least 2 min (approximately 5 lg). Reactions were

initiated by addition of the second substrate as described

[31]. Kinetic parameters were estimated from non-linear

curve fitting using GraphPad Prism software (version 5 for

Windows, GraphPad Software). The kinetic values were

calculated by Michaelis–Menten or Substrate Inhibition

models. In the latter case, the equation used was:

Y = Vmax 9 X/[Km ? X 9 (1 ? X/Ki)], where Vmax is the

maximum enzyme velocity, if the substrate didn’t also

inhibit enzyme activity, Km is the Michaelis–Menten con-

stant, and Ki is the dissociation constant for substrate

binding in such a way that two substrates can bind to an

enzyme.

Size exclusion chromatography

Size exclusion chromatography was performed on a

Prominence liquid chromatography system (Shimadzu Co.

Tokyo, Japan) equipped with a ZORBAX GF-250 4 lm

column (4.6 9 250 mm, Agilent Technologies, Palo Alto,

CA, USA) and a SPD 20A diode array detector. Aliquots of

20 ll of protein samples were injected and eluted isocrat-

ically with 20 mM Na2HPO4 pH 7.0, 130 mM NaCl at a

flow rate of 1 ml/min. Molecular weight calibration was

performed by using bovine thyroglobulin (670 kDa),

bovine c-globulin (158 kDa), chicken ovalbumin (44 kDa),

horse myoglobin (17 kDa) and vitamin B12 (1.35 kDa)

mixture, all components of the Bio-Rad gel filtration

standard kit. Chromatograms were analyzed at 280 nm.

RNA isolation

Total RNA was isolated by the hot SDS/hot phenol method

[32]. Briefly, bacterial cultures were added to a 1/10 vol-

ume of 95% ethanol plus 5% saturated phenol to stabilize

cellular RNA and the cells were then harvested by centri-

fugation at 6,000 rpm for 5 min at 4�C. The supernatant

was aspirated and the pellets were kept at -80�C for no

longer than 1 week. Pellets were resuspended in 800 ll of

lysis buffer (10 mM Tris–HCl pH 8.0, 1 mM EDTA sup-

plemented with 3 mg/ml lysozyme) and 80 ll of a 10%

SDS solution was added to the lysate and incubated at 64�C

for 2 min. Following incubation, 88 ll of 1 M NaOAc (pH

5.2) was added to the lysate and an equal volume of water-

saturated phenol was added and incubated at 64�C for

6 min, inverting the tubes six times every 40 s. The sam-

ples were chilled on ice and centrifuged at 13,000 rpm for

10 min at 4�C. The aqueous layer was transferred to a tube

with equal volume of chloroform and centrifuged at

13,000 rpm for 5 min at 4�C. Subsequently, the aqueous

layer was ethanol precipitated by adding 1/10 volume of

3 M NaOAc (pH 5.2), 1 mM EDTA and two volumes cold

ethanol. The samples were incubated at -80�C overnight.

The RNA was pelleted by centrifugation at 13,000 rpm for

25 min at 4�C. Pellets were washed with ice cold 80%

ethanol and centrifuged at 13,000 rpm for 5 min at 4�C.

The pellets were resuspended in a total of 100 ll of RNase-

free water.

Total RNA was quantified using micro-spectrophotom-

etry (NanoDrop Technologies, Inc.) and its quality was

assessed by agarose gel electrophoresis. DNA was removed

with DNase I (Promega). Removal of DNA from the RNA

samples was confirmed by performing real-time PCR on

100 ng of total RNA using the Avin_38560-qPCR primer

set (Table 1), but without a reverse transcriptase step.

Those RNA samples found to yield Ct values larger than 35

were judged to be sufficiently free of contaminating DNA

for further analysis. Once more, total RNA was quantified

using micro-spectrophotometry (NanoDrop Technologies,

Inc.) while the A260/A280 ratio was always above two.

Purified RNA was either converted to cDNA immediately

or stored frozen at -80�C.

cDNA synthesis

First-strand cDNA synthesis was performed using Super-

script II RT (Invitrogen). Briefly, 1.5 lg of each RNA

sample was mixed with 1 ll of 250 ng/ll random hexamer

primers, 1 ll of 10 mM dNTP mix (Finnzymes), and

RNase-free water to make 12 ll. The mixture was

Table 1 Primers used in RT-qPCR reactions

Primer name Nucleotide sequence

Avin_34560-qPCR-F 50-AACTGGGCGCTGGTGCGTAT-30

Avin_34560-qPCR-R 50-TCCATGGCGGCGAGCATTTCA-30

Avin_41120-qPCR-F 50-TGGGCTGGAGCCTCACGGAAAT-30

Avin_41120-qPCR-R 50-CCAGGTTGCGCATGTCGTTGGA-30

Avin_34550-qPCR-F 50-CCGATACTGCTGGATCAGGT-30

Avin_34550-qPCR-R 50-CTGATAGCGGCTGACCACTT-30

Avin_41130-qPCR-F 50-GACTACTGTGCCAGTCACC-30

Avin_41130-qPCR-R 50-CCCGCTTTACCAGGTTGTAG-30

Avin_36780-qPCR-F 50-ATCCCTGCAGGCCCACCTCC-30

Avin_36780-qPCR-R 50-TGGCTATGGCGTGGTCCCGA-30

Avin_34530-qPCR-F 50-ATGTCGCGATGCGAGTGCCA-30

Avin_34530-qPCR-R 50-GCGTTGACCAATCCGGACACCT-30

Avin_42750-qPCR-F 50-GCAGCATCTACGGCGACCATCA-30

Avin_42750-qPCR-R 50-TGCGCCATCGCCGGTGAAAT-30

Avin_08480-qPCR-F 50-GGCCGGTGCTGATCGAGTTCAA-30

Avin_08480-qPCR-R 50-ACAGCAAGGTGCCGGCGAAT-30

Avin_38560-qPCR-F 50-TCCGGGTTGCCGAACATCACA-30

Avin_38560-qPCR-R 50-TGCGCAAGATCACCGGCAACA-30

Mol Biol Rep (2011) 38:3653–3663 3655

123

denatured at 65�C for 5 min and immediately chilled. 4 ll

of 59 first-strand buffer, 2 ll of 0.1 M DTT, 1 ll of

40 units/ll RNaseOUT (Invitrogen) and 1 ll of 200 units/

ll SuperScript II (Invitrogen) were then added and the

volume was adjusted to 20 ll. The mixture was incubated

at 25�C for 10 min, 42�C for 50 min, and finally 70�C for

15 min to inactivate the reaction.

RT-qPCR

The oligonucleotide primers used for RT-qPCR are listed

in Table 1. All primers were supplied by Invitrogen. The

primers were designed using Primer-BLAST (http://blast.

ncbi.nlm.nih.gov/Blast.cgi) and optimised to an equal

annealing temperature of 60�C. Each primer pair further

assessed for specificity with melting curve analysis and gel

electrophoresis of amplification products.

One microliter of diluted cDNA was added to the primer

pair mix and SYBR Green Master Mix (Applied Biosys-

tems) in each well. RT-qPCR was conducted on an MxPro

Mx3005P PCR system and analyzed with MxPro v4.01

software (Stratagene). The PCR cycling conditions were

95�C for 10 min, 40 cycles of 95�C for 30 s and 60�C for

1 min, 95�C for 1 min, 60�C for 30 s and 95�C for 30 s.

Plates were run for 40 cycles and fluorescence intensity

measured after every cycle. For each target sequence the

average cycle number at which fluorescence was detected

above background, in the exponential phase of amplifica-

tion, was obtained. No-reverse transcription controls were

included. The efficiency of each RT-qPCR reaction was

calculated using the LinRegPCR software [33].

Relative expression of the target gene in the various

Burk’s minimal media versus LB full medium was calcu-

lated using the following equation, described by Pfaffl [34].

Relative expression ratio per gene: R = EGOI^ (Ct, CONTROL

- Ct, SAMPLE) GOI/EREF^ (Ct, CONTROL-Ct, SAMPLE) REF. recA

(Avin_38560) was used as internal control. Statistical sig-

nificance was obtained using a pair-wise fixed reallocation

randomization test using the REST software [35].

Results

Avpta-1 and Avpta-2 encode putative phosphate

acetyltransferases

We identified three putative phosphate acetyltransferases

from A. vinelandii, AvPTA-1 (ACO79604), AvPTA-2

(ACO80246) and AvPTA-3 (ACO80523), all belonging to

the phosphate acetyl/butaryl transferase superfamily by

sequence homology searches using BLAST [36]. Microbial

genome sequencing has revealed two main classes of

phosphate acetyltransferase enzymes. Class I enzymes

are *350 amino acids in length, whereas class II enzymes

are *700 amino acids. Class I enzymes share end-to-end

homology with the C-terminal domain of class II enzymes;

hence it is inferred that the active site of class II enzymes is

located within their C-terminal domain [21]. AvPTA-1 (712

amino acids) and AvPTA-2 (691 amino acids) show 61.2%

identity between each other and 50.2 and 53% identity with

E. coli pta, respectively, all belonging to the Class II

enzymes. The N-terminal region of AvPTA-1 and AvPTA-2

contains a Bio-D and a DRTGG domain, as revealed by

CDD searches [37]. AvPTA-3 (317 amino acids), on the

other hand, has no additional domains, shows approxi-

mately 21% identity with E. coli eutD and only 8.9%

identity with E. coli pta, probably representing another

class of phosphate acetyl/butaryl transferases superfamily.

Figure 1 shows a ClustalW multiple protein alignment

among A. vinelandii AvPTA-1, AvPTA-2, AvPTA-3 and

other characterized phosphate acetyltransferases. Catalyti-

cally important Ser309, Arg310 and Asp316 residues (Met-

hanosarcina thermophila numbering) are all conserved in

AvPTA-1 and AvPTA-2 but not in AvPTA-3.

AvPTA-1 and AvPTA-2 exhibit phosphate

acetyltransferase activity

Using purified recombinant proteins, the catalytic proper-

ties of both AvPTA-1 and AvPTA-2 were investigated.

Both genes were PCR-amplified from A. vinelandii and

cloned to an expression vector to obtain the corresponding

His-tagged proteins, which were overproduced in E. coli

BL21 (DE3) cells and purified using Ni–NTA chroma-

tography. Analysis of the purified recombinant proteins by

SDS-PAGE and Coomassie blue staining, revealed single

protein bands, close to the theoretical molecular masses of

both proteins (77 and 76 kDa, respectively) (Fig. 2a, b). To

determine the biochemical characteristics of AvPTA-1 and

AvPTA-2, the kinetic constants were determined for both

directions of the reaction: acetyl-P ? CoA $ acetyl-

CoA ? Pi. The apparent kinetic constants of the substrates

were determined by varying the concentration of one

substrate around its Km value while leaving the other

unchanged at saturation concentrations.

Fig. 1 Sequence alignment of A. vinelandii AvPTA-1 (ACO79604)

and AvPTA-2 (ACO80246) and related enzymes. The sequences

included are Bacillus subtilis pta (CAB15793), Methanosarcinathermophila pta (AAA72041), Salmonella typhimurium pta

(AAL21239), E. coli pta (AAC75357), Lactobacillus sanfrancisensispta (BAB19267), E. coli eutD (AAC75511), Salmonella typhimuriumeutD (AAL21360) and A. vinelandii AvPTA-3 (ACO80523). Multiple

sequence alignment was performed using ClustalW [42]. Black boxesindicate identical amino acids while grey boxes indicate similar.

Black dots indicate amino acids involved in catalysis of Methano-sarcina thermophila pta [26]

c

3656 Mol Biol Rep (2011) 38:3653–3663

123

Mol Biol Rep (2011) 38:3653–3663 3657

123

For the forward reaction (acetyl-CoA forming direction),

with acetyl-P as a variable substrate (Fig. 3a), AvPTA-1

showed substrate inhibition kinetics (Ki = 5.2 mM) with

Km = 1 ± 0.4 mM and Vmax = 0.1 ± 0.02 lmol s-1 lg-1.

With CoA as a variable substrate (Fig. 3b), AvPTA-1 showed

inhibition kinetics (Ki = 0.1 mM) with Km = 0.2 ± 0.1

mM and Vmax = 0.1 ± 0.06 lmol s-1 lg-1. These charac-

teristics are within the range of other bacterial and archaea

species (Brenda, http://www.brenda-enzymes.org/).

For the reverse reaction (acetyl-P forming direction)

with acetyl-CoA as a variable substrate (Fig. 3c), AvPTA-1

showed Michaelis–Menten kinetics with Km = 0.05 ±

0.01 mM and Vmax = 0.003 ± 0.001 lmol s-1 lg-1, while

with phosphate as a variable substrate (Fig. 3d), AvPTA-1

showed inhibition kinetics (Ki = 19 mM) with Km = 11 ±

9 mM and Vmax = 0.004 ± 0.002 lmol s-1 lg-1. These

characteristics, as well, are within the range of other bac-

terial and archaea species (Brenda, http://www.brenda-

enzymes.org/).

AvPTA-2, on the other hand, for the forward reaction

(acetyl-CoA forming direction), with acetyl-P as a variable

substrate (Fig. 4a), showed substrate inhibition kinetics

(Ki = 12 mM) with Km = 0.5 ± 0.1 mM and Vmax =

0.004 ± 0.0004 lmol s-1 lg-1. With CoA as a variable

substrate (Fig. 4b), AvPTA-2 showed inhibition kinetics

(Ki = 0.04 mM) with Km = 0.3 ± 0.1 mM and Vmax =

0.04 ± 0.02 lmol s-1 lg-1, as well within the range of

other species (Brenda, http://www.brenda-enzymes.org/).

For the reverse reaction (acetyl-P forming direction) we

could not detect any enzyme activity for AvPTA-2 under

our experimental conditions.

Oligomeric state analysis of AvPTA-1 and AvPTA-2

In order to gain an insight into the oligomeric state of

recombinant AvPTA-1 and AvPTA-2, we performed gel fil-

tration analysis. The recombinant AvPTA-1 was estimated as

a *163 kDa peak, probably representing a dimer, since the

theoretical molecular mass of AvPTA-1 is *77 kDa. Simi-

larly, recombinant AvPTA-2 was estimated as a *165 kDa

peak, probably representing a dimer as well, since the the-

oretical molecular mass of AvPTA-2, is *76 kDa.

A. vinelandii’s acetate metabolism under various

physiological and growth conditions

The effect of growth phase (exponentially or stationary

grown cells), carbon source (acetate or glucose) and source

of fixed nitrogen (ammonium supplied either by nitrogen

fixation or exogenously supplied to the medium) on the

accumulation of gene transcripts involved in acetate

assimilation or dissimilation was examined by real-time

RT-qPCR. Cells from the exponential and the stationary

phases were harvested and the relative gene expression

levels were calculated against rich LB medium grown cells

from the same growth phases.

Figure 5 presented a simplified model of the acetate

metabolism pathways examined in the present study. We

suppose that acetyl-CoA can be metabolized into acetate by

the phosphate acetyltransferase (pta, E.C 2.2.1.8,

Avin_34550, Avin_41130) and acetate kinase (Ack-A, E.C

2.7.2.1, Avin_34560, Avin_41120) pathway. An acylphos-

phatase (AcPh, E.C 3.6.1.7, Avin_36780) may possibly

convert acetyl-P into acetate while the acetyl-Co/acetate

interconversion could also be made by the acetyl-CoA syn-

thetase (Acs-A, E.C 6.2.1.1, Avin_34530, Avin_42750).

Moreover, a thiamine diphosphate-dependent pyruvate

dehydrogenase (poxB, E.C 1.2.2.2, Avin_08480) can pro-

duce acetyl-P from pyruvate.

When A. vinelandii was exponentially grown on glucose

minimal medium as the primary carbon source, Avacs-A1

transcript levels were elevated (P \ 0.05) (Fig. 6c) while

Avpta-1 and Avpta-2 were stably expressed (Fig. 6a, b).

Growth on acetate minimal medium, on the other hand,

Fig. 2 SDS-PAGE analysis of expression and purification of

recombinant AvPTA-1 and AvPTA-2. a SDS-PAGE analysis of the

soluble cytoplasmic fraction from E. coli BL21 (DE3) cells

overexpressing AvPTA-1 lane 1, and the elution fraction after the

Ni–NTA purification of the recombinant AvPTA-1 lane 2. b SDS-

PAGE analysis of the soluble cytoplasmic fraction from E. coli BL21

(DE3) cells overexpressing AvPTA-2 lane 1, and the elution fraction

after the Ni–NTA purification of the recombinant AvPTA-2 lane 2

3658 Mol Biol Rep (2011) 38:3653–3663

123

decreased Avacs-A1 expression (P \ 0.05) (Fig. 6c) while

Avpta-1 and Avpta-2 remained stably expressed although

they showed increased expression compared to the glucose

growth (P \ 0.05, Anova) (Fig. 6a, b). The presence of

ammonium in the glucose minimal medium down-regu-

lated Avacs-A1 expression (P \ 0.05) (Fig. 6c) while its

presence in the acetate minimal medium up-regulated

Avpta-2 expression (P \ 0.05) (Fig. 6b). Avacs-A2

retained its basal levels of expression during exponential

growth in all conditions tested (Fig. 6d).

During A. vinelandii stationary phase growth on either

glucose or acetate minimal media most of the studied genes

showed a tendency for down-regulation (Fig. 6). However,

during the stationary phase, Avacs-A1 retained its basal level

Fig. 3 Phosphate acetyltransferase activity of the forward (acetyl-

CoA forming direction) and reverse reaction (acetyl-P forming

direction) catalysed by AvPTA-1. a Acetyl-CoA forming direction.

The CoA was kept constant at 0.125 mM, while the acetyl-P

concentration was varied. b Acetyl-CoA forming direction. The

acetyl-P was kept constant at 2.5 mM, while the CoA concentration

was varied. c Acetyl-P forming direction. The phosphate was kept

constant at 12.5 mM, while the acetyl-CoA concentration was varied.

d Acetyl-P forming direction. The acetyl-CoA was kept constant at

0.1 mM, while the phosphate concentration was varied. Mean values

were obtained from three independent replicates and error barsrepresent standard errors. Non-linear regression analysis was used in

order to fit the data to curves using GraphPad Prism v5.0 (GraphPad

Software, San Diego, CA)

Fig. 4 Phosphate acetyltransferase activity of the forward (acetyl-

CoA forming direction) catalysed by AvPTA-2. a Acetyl-CoA

forming direction. The CoA was kept constant at 0.025 mM, while

the acetyl-P concentration was varied. b Acetyl-CoA forming

direction. The acetyl-P was kept constant at 2 mM, while the CoA

concentration was varied. Mean values were obtained from three

independent replicates and error bars represent standard errors. Non-

linear regression analysis was used in order to fit the data to curves

using GraphPad Prism v5.0 (GraphPad Software, San Diego, CA)

Mol Biol Rep (2011) 38:3653–3663 3659

123

of expression when ammonium was present in the acetate

minimal medium (Fig. 6c) while Avacs-A2 retained its basal

level of expression in the acetate minimal medium (Fig. 6d).

Finally, we could not detect either AvAcPh or AvpoxB

expression under these growth and physiological

conditions.

Discussion

Acetate is abundant in soil and other environments as a

product of many fermentative and degradative processes

[1–3] and since it constitutes the growth substrate for

methane producing archaea as well, it represents a major

intermediate in the global carbon cycle [5]. Phosphate

acetyltransferase rapidly converts acetyl-CoA via acetyl-P

to acetate as an overflow pathway or converts exogenous

acetate back to acetyl-CoA [14]. Two main classes of

phosphate acetyltransferase enzymes have been identified

with class I enzymes to be *350 amino acids in length,

whereas class II enzymes *700 amino acids [21].

AvPTA-1 and AvPTA-2 from the soil, nitrogen-fixing

A. vinelandii, belong to the Class II enzymes, with their

N-terminal regions containing Bio-D and DRTGG domains

Fig. 5 Simplified model for the acetate metabolism pathways

examined in the present study. Pta phosphate acetyltransferase,

Ack-A acetate kinase, AcPh acylphosphatase, Acs-A acetyl-CoA

synthetase, poxB thiamine diphosphate-dependent pyruvate

dehydrogenase

Fig. 6 Relative expression levels of Avpta-1 (a) Avpta-2 (b) Avacs-

A1 (c) and Avacs-A2 (d) in A. vinelandii cultures grown in various

Burk’s minimal media (BG Burk’s with glucose, BGN Burk’s with

glucose and ammonium, BA Burk’s with acetate, BAN Burk’s with

acetate and ammonium) versus A. vinelandii cultures grown in LB

full medium as determined by RT-qPCR. Fold change was calculated

according to the equation described in the Materials and methods with

normalization against recA. The data are the mean of three biological

replications while the bars represent standard errors. Up-pointingarrow indicate up-regulation (P \ 0.05), down-pointing arrow indi-

cate down-regulation (P \ 0.05) and rectangular indicate not statis-

tically significant regulation (P [ 0.05), calculated with the REST

software [35]

3660 Mol Biol Rep (2011) 38:3653–3663

123

while their C-terminal domains correspond to the pta

domain.

Our data demonstrate that both AvPTA-1 and AvPTA-2

show phosphate acetyltransferase activity in the acetyl-

CoA forming direction while for the acetyl-P forming

direction we were able to measure enzyme activity only for

the AvPTA-1 isoform. AvPTA-1 activity is greater in the

acetyl-CoA forming direction (Vmax = 0.1 ± 0.02 l-mol s-1 lg-1) than the acetyl-P forming direction

(Vmax = 0.003 ± 0.001 lmol s-1 lg-1) while AvPTA-2

activity in the acetyl-CoA forming direction was lower

than the AvPTA-1 activity at the same direction

(Vmax = 0.004 ± 0.0004 vs. 0.1 ± 0.02 lmol s-1 lg-1).

These data taken together suggest that AvPTA-1 has lower

affinity to acetyl-P, as shown by a Km for acetyl-P that is

approximately one order of magnitude higher than that of

AvPTA-2 (1 ± 0.4 vs. 0.5 ± 0.1 mM). In contrast,

AvPTA-1 has higher affinity than the AvPTA-2 to CoA

(0.2 ± 0.1 vs. 0.3 ± 0.1 mM). The different enzyme

kinetics of the isoforms suggest that the pta gene products

may serve different biological roles under certain

metabolic conditions which are imposed by the availabil-

ity of carbon sources and/or maintenance of acetyl-P

concentration.

In most bacteria, pta and ack-A genes are organized into

a single operon and they are involved in the maintenance of

the intracellular acetyl-CoA and acetyl-P pools [38].

However, the conversion of acetyl-CoA to acetate and ATP

often does not go to completion so cells maintain a sig-

nificant pool of acetyl-P, which serves as a storage mole-

cule of carbon, phosphate and energy as well as a global

signal [39]. Our biochemical and in silico analyses revealed

that A. vinelandi genome contains two functional phos-

phate acetyltransferase isoforms as well as two putative

acetate kinases and acetyl-CoA synthetases. Orthologs

neighbourhood analyses revealed that in A. vinelandi the

genes coding for these enzymes are organized into two pta-

ack-A operons, Avpta-1-Avack-A1 and Avpta-2-Avack-A2.

Furthermore, Avacs-A1 is also adjacent to the former

operon while Avacs-A2 is distant from the latter operon.

Our results show that the two phosphate acetyltransfer-

ase genes are constitutively expressed during the expo-

nential growth and the encoded enzymes show different

kinetic constants for the acetate catabolism as well as the

acetate production routes. So the acetate assimilation/dis-

similation system could function under a wide range of

acetate concentrations allowing A. vinelandii to adapt and

survive under the prevailing and/or changing environ-

mental conditions. Further, the allosteric regulation of the

Class II PTA enzymes by pyruvate, phosphoenolpyruvate,

NADH and ATP [21, 29, 40] indicates the central role

of that enzyme in the carbon and energy status of the

cell. Moreover, deletion of the E. coli phosphate

acetyltransferase resulted in increased sensitivity to envi-

ronmental changes and de-regulation of the central

metabolism [41]. Nevertheless, the existence of paralogs

for phosphate acetyltransferase, acetate kinase and acetyl-

CoA synthetase in A. vinelandii adds an extra level of

complexity in acetate metabolism of this soil bacterium.

A. vinelandii is among the few bacterial species able to

fix atmospheric nitrogen under aerobic conditions. Thus,

investigation of the expression pattern of genes involved in

acetate metabolism under conditions where ammonium is

either exogenously supplied or produced via nitrogen fix-

ation would provide valuable information as to the survival

and growth of these bacterial species in the soil environ-

ment. Our data demonstrated that when A. vinelandii was

exponentially grown under nitrogen-fixing conditions, on

glucose minimal medium as the primary carbon source,

Avacs-A1 transcript levels were elevated while Avpta-1 and

Avpta-2 were stably expressed. Growth on acetate minimal

medium, on the other hand, decreased Avacs-A1 expression

while Avpta-1 and Avpta-2 remained stably expressed

although they showed increased expression compared to

the glucose growth. These data taken together suggest that

rapidly growing and nitrogen-fixing A. vinelandii uses the

pta pathway when acetate is available, and the acs-A

pathway when glucose is available as a carbon source. The

presence of fixed nitrogen (ammonium) in the glucose

minimal medium down-regulated Avacs-A1 expression

while its presence in the acetate minimal medium up-reg-

ulated Avpta-2 expression indicating that the availability of

fixed nitrogen in combination with available carbon source

determines whether the acs-A or the pta pathway is used

for acetate assimilation.

During A. vinelandii stationary phase growth under

nitrogen-fixing conditions, on either glucose or acetate

minimal media, most of the studied genes showed a ten-

dency for down-regulation. However, during the stationary

phase, Avacs-A1 retained its basal level of expression when

ammonium was present in the acetate minimal medium

while Avacs-A2 retained its basal level of expression in the

acetate minimal medium, indicating the critical role of the

acs-A acetate assimilation system during growth on

acetate.

In conclusion, we provide evidence that A. vinelandii

genome contains two functional phosphate acetyltransfer-

ase isoforms as well as two differentially transcribed

acetyl-CoA synthetase isoforms. The co-expression of both

Avpta genes in combination with the homo-dimeric form of

both enzymes raises questions whether or not hetero-

dimeric PTA enzymes are formed in vivo and whether they

display any enzyme activity. Further studies on the func-

tional characterization of these enzymes by inactivation of

either of the genes or both genes would better clarify their

specific roles during acetate metabolism.

Mol Biol Rep (2011) 38:3653–3663 3661

123

References

1. Buckel W (1999) Anaerobic energy metabolism. In: Lengler JW,

Drews G, Chlegel HG (eds) Biology of the procaryotes. Thieme,

Stuttgart, pp 278–326

2. Glissmann K, Weber S, Conrad R (2001) Localization of pro-

cesses involved in methanogenic degradation of rice straw in

anoxic paddy soil. Environ Microbiol 3:311–502

3. Conrad R, Klose M, Claus P (2002) Pathway of CH4 formation in

anoxic rice field soil and rice roots determined by 13C-stable

isotope fractionation. Chemosphere 47:797–806

4. Jensen HL (1954) The Azotobacteriaceae. Bacteriol Rev

18:195–214

5. Ferry JG (1993) Fermentation of acetate. In: Ferry JG (ed)

Methanogenesis: ecology, physiology, biochemistry and genetics.

Chapman and Hall, New York, pp 304–334

6. Clark DP, Cronan JE (1996) Two-carbon compounds and fatty

acids as carbon sources. In: Neidhardt FC, Curtiss R III, Ingra-

ham JL, Lin ECC, Low KB, Magasanik B, Reznikoff WS, Riley

M, Schaechter M, Umbarger HE (eds) Escherichia coli and

Salmonella: cellular and molecular biology, 2nd edn. ASM Press,

Washington, pp 343–357

7. JJr Cronan, LaPorte D (2006) Tricarboxylic acid cycle and glyoxylate

bypass. In: Curtiss R III et al (eds) EcoSal-Escherichia coli and

Salmonella: cellular and molecular biology. ASM Press, Washington

8. Erb TJ, Berg IA, Brecht V, Muller M, Fuchs G, Alber BE (2007)

Synthesis of C5-dicarboxylic acids from C2-units involving

crotonyl-CoA carboxylase/reductase: the ethylmalonyl-CoA

pathway. Proc Natl Acad Sci USA 104:10631–10636

9. Kornberg HL (1966) The role and control of the glyoxylate cycle

in Escherichia coli. Biochem J 99:1–11

10. Zarzycki J, Schlichting A, Strychalsky N, Muller M, Alber BE,

Fuchs G (2008) Mesaconyl-coenzyme A hydratase, a new

enzyme of two central carbon metabolic pathways in bacteria.

J Bacteriol 190:1366–1374

11. Starai VJ, Escalante-Semerena JC (2004) Acetyl-coenzyme A

synthetase (AMP forming). Cell Mol Life Sci 61:2020–2030

12. Brown TD, Jones-Mortimer MC, Kornberg HL (1977) The

enzymic interconversion of acetate and acetyl-coenzyme A in

Escherichia coli. J Gen Microbiol 102:327–336

13. El-Mansi M (2005) Free CoA-mediated regulation of interme-

diary and central metabolism: a hypothesis which accounts for

the excretion of alpha-ketoglutarate during aerobic growth of

Escherichia coli on acetate. Res Microbiol 156:874–879

14. El-Mansi M, Cozzone AJ, Shiloach J, Eikmanns BJ (2006)

Control of carbon flux through enzymes of central and interme-

diary metabolism during growth of Escherichia coli on acetate.

Curr Opin Microbiol 9:173–179

15. Wolfe AJ (2005) The acetate switch. Microbiol Mol Biol Rev

69:12–50

16. El-Mansi EM, Holms WH (1989) Control of carbon flux to

acetate excretion during growth of Escherichia coli in batch and

continuous cultures. J Gen Microbiol 135:2875–2883

17. Kleman GL, Strohl WR (1994) Acetate metabolism by Esche-richia coli in high-cell-density fermentation. Appl Environ

Microbiol 60:3952–3958

18. Blake R, O’Brien TA, Gennis RB, Hager LP (1982) Role of the

divalent metal cation in the pyruvate oxidase reaction. J Biol

Chem 257:9605–9611

19. Abdel-Hamid AM, Attwood MM, Guest JR (2001) Pyruvate

oxidase contributes to the aerobic growth efficiency of Esche-richia coli. Microbiology 147:1483–1498

20. Pruss BM, Wolfe AJ (1994) Regulation of acetyl phosphate

synthesis and degradation, and the control of flagellar expression

in Escherichia coli. Mol Microbiol 12:973–984

21. Brinsmade SR, Escalante-Semerena JC (2007) In vivo and in

vitro analyses of single-amino acid variants of the Salmonellaenterica phosphotransacetylase enzyme provide insights into the

function of its N-terminal domain. J Biol Chem 282:12629–

12640

22. Liu Y, Leal NA, Sampson EM, Johnson CLV, Havemann GD,

Bobik TA (2007) PduL is an evolutionarily distinct phospho-

transacylase involved in B12-dependent 1, 2-propanediol degra-

dation by Salmonella enterica Serovar typhimurium LT2.

J Bacteriol 189:1589–1596

23. Kyrtopoulos SA, Satchell DP (1972) Kinetic studies with phos-

photransacetylase. IV. Inhibition by products. Biochim Biophys

Acta 276:383–391

24. Pelroy RA, Whiteley HR (1972) Kinetic properties of phospho-

transacetylase from Veillonella alcalescens. J Bacteriol

111:47–55

25. Bock AK, Glasemacher J, Schmidt R, Schonheit P (1999) Puri-

fication and characterization of two extremely thermostable

enzymes, phosphate acetyltransferase and acetate kinase, from

the hyperthermophilic eubacterium Thermotoga maritime. J Bac-

teriol 181:1861–1867

26. Lawrence SH, Luther KB, Schindelin H, Ferry JG (2006)

Structural and functional studies suggest a catalytic mechanism

for the phosphotransacetylase from Methanosarcina thermophila.

J Bacteriol 188:1143–1154

27. Xu QS, Jancarik J, Lou Y, Kuznetsova K, Yakunin AF, Yokota

H, Adams P, Kim R, Kim SH (2005) Crystal structures of a

phosphotransacetylase from Bacillus subtilis and its complex

with acetyl phosphate. J Struct Funct Genomics 6:269–279

28. Shimizu M, Suzuki T, Kameda KY, Abiko Y (1969) Phospho-

transacetylase of Escherichia coli B, purification and properties.

Biochim Biophys Acta 191:550–558

29. Campos-Bermudez VA, Bologna FP, Andreo CS, Drincovich MF

(2010) Functional dissection of Escherichia coli phosphotransa-

cetylase structural domains and analysis of key compounds

involved in activity regulation. FEBS J 277:1957–1966

30. Kennedy C, Gamal R, Hummprey R, Ramos J, Brigle K, Dean D

(1986) The nifH, nifM, and nifN genes of Azotobacter vinelandii:characterization by Tn5 mutagenesis and isolation from pLARF1

gene banks. Mol Gen Genet 205:318–325

31. Lawrence SH, Ferry JG (2006) Steady-state kinetic analysis of

phosphotransacetylase from Methanosarcina thermophila. J Bac-

teriol 188:1155–1158

32. Jahn CE, Charkowski AO, Willis DK (2008) Evaluation of iso-

lation methods and RNA integrity for bacterial RNA quantitation.

J Microbiol Methods 75:318–324

33. Ruijter JM, Ramakers C, Hoogaars WM, Karlen Y, Bakker O,

van den Hoff MJ, Moorman AF (2009) Amplification efficiency:

linking baseline and bias in the analysis of quantitative PCR data.

Nucleic Acids Res 37:e45

34. Pfaffl MW (2001) A new mathematical model for relative

quantification in real-time RT-PCR. Nucleic Acids Res 29:e45

35. Pfaffl MW, Horgan GW, Dempfle L (2002) Relative expression

software tool (REST) for group-wise comparison and statistical

analysis of relative expression results in real-time PCR. Nucleic

Acids Res 30:e36

36. Altschul SF, Madden TL, Schaffer AA, Zhang J, Zhang Z, Miller

W, Lipman DJ (1997) Gapped BLAST and PSI-BLAST: a new

generation of protein database search programs. Nucleic Acids

Res 25:3389–3402

37. Marchler-Bauer A, Anderson JB, Chitsaz F, Derbyshire MK,

DeWeese-Scott C, Fong JH, Geer LY, Geer RC, Gonzales NR,

Gwadz M, He S, Hurwitz DI, Jackson JD, Ke Z, Lanczycki CJ,

Liebert CA, Liu C, Lu F, Lu S, Marchler GH, Mullokandov M,

Song JS, Tasneem A, Thanki N, Yamashita RA, Zhang D, Zhang

3662 Mol Biol Rep (2011) 38:3653–3663

123

N, Bryant SH (2009) CDD: specific functional annotation with

the Conserved Domain Database. Nucleic Acids Res 37D:205–

210

38. Singh-Wissmann K, Ferry JG (1995) Transcriptional regulation

of the phosphotransacetylase-encoding and acetate kinase-

encoding genes (pta and ack) from Methanosarcina thermophila.

J Bacteriol 177:1699–1702

39. Wolfe AJ (2010) Physiologically relevant small phosphodonors

link metabolism to signal transduction. Curr Opin Microbiol

13:204–209

40. Suzuki T (1969) Phosphotransacetylase of Escherichia coli B,

activation by pyruvate and inhibition by NADH and certain

nucleotides. Biochim Biophys Acta 191:559–569

41. Castano-Cerezo S, Pastor JM, Renilla S, Bernal V, Iborra JL,

Canovas M (2009) An insight into the role of phosphotransa-

cetylase (pta) and the acetate/acetyl-CoA node in Escherichiacoli. Microb Cell Fact 8:54

42. Chenna R, Sugawara H, Koike T, Lopez R, Gibson TJ, Higgins

DG, Thompson JD (2003) Multiple sequence alignment with the

Clustal series of programs. Nucleic Acids Res 31:3497–3500

Mol Biol Rep (2011) 38:3653–3663 3663

123