Molecular basis of cellulose biosynthesis disappearance in submerged
culture of Acetobacter xylinum
Alina Krystynowicz1, Maria Koziołkiewicz1, Agnieszka Wiktorowska-Jezierska1,
Stanisław Bielecki1, Emilia Klemenska2, Aleksander Masny2 and
Andrzej Płucienniczak2
1Institute of Technical Biochemistry,Technical University of Lodz, Łódź, Poland; 2Institute of Biotechnology and
Antibiotics, Warszawa, Poland; e-mail: stanb@p.lodz.pl
Received: 13 July, 2005; revised: 25 July, 2005; accepted: 28 July, 2005
available on-line: 1 September, 2005
Acetobacter xylinum strains are known as very efficient producers of bacterial cellulose which,
due to its unique properties, has great application potential. One of the most important problems
faced during cellulose synthesis by these bacteria is generation of cellulose non-producing
cells, which can appear under submerged culture conditions. The reasons of this remain unknow.
These studies have been undertaken to compare at the molecular level wild-type, cellulose producing
(Cel+) A. xylinum strains with Cel– forms of cellulose-negative phenotype. Comparison of
protein profiles of both forms of A. xylinum by 2D electrophoresis allowed for the isolation of
proteins which were produced exclusively by either Cel+ or Cel– cells. Sequences of peptides derived
from these proteins were aligned with those of proteins deposited in databases. This analysis
revealed that Cel– cells lacked two enzymes: phosphoglucomutase and glucose-1-phosphate
uridylyltransferase, which generates UDP-glucose being the substrate for cellulose synthase.
DNA was analyzed by ligation-mediated PCR carried out at low denaturation temperature (PCRMP).
Two DNA fragments of different thermal stability (218 and 217 bp) were obtained from the
DNA of Cel+ and Cel– forms, respectively. The only difference between these Cel– and Cel+ DNA
fragments is deletion of one T residue. Alignment of those two sequences with those deposited
in the GenBank database revealed that similar fragments are present in the genomes of some
bacterial cellulose producers and are located downstream from open reading frames (ORF) encoding
phosphoglucomutase. The meaning of this observation is discussed.
Keywords: bacterial cellulose, Acetobacter xylinum, 2-D electrophoresis, PCR-MP, phosphoglucomutase,
UDP-glucose pyrophosphorylase
Presented at the International Review Conference on Biotechnology, Vienna, Austria, November 2004.
Abbreviations: Cel+ cells, wild-type cellulose-producing cells of Acetobacter xylinum; Cel– cells, non-reverting and cellulose-nonproducing
forms of A. xylinum; c-di-GMP, cyclic di-guanosine monophosphate; CS, cellulose synthase; CTAB,
cetyltrimethylammonium bromide; DTT, dithiothreitol; PCR-MP, ligation-mediated PCR performed at low denaturation
temperatures; TAE, Tris/acetate/EDTA buffer.
Bacterial cellulose has found multiple applications
in various fields owing to its unique physicochemical
and mechanical properties (Ring et al.,
1986; Ross et al., 1991). However, the scale of its
production, processing and use is relatively small
because of problems with selection of sufficiently efficient
producers and costs of culture media.
The most efficient producers of bacterial cellulose
are the Gram-negative Acetobacter xylinum
rods (reclassified as Gluconacetobacter xylinus) (Yamada,
2000). They occur singly, in pairs or in chains,
reproduce by binary fission, are motile by flagella,
and do not form endospores. Under limiting conditions,
A. xylinum strains form involution forms, i.e.
swollen or elongated filaments. Under conditions
appropriate for bacterial growth these forms atrophy
or fragment to shorter pieces which enables
recovery of normal cells. The optimum temperature
for A. xylinum growth is 25–30°C, and optimum pH
ranges from 5.4 to 6.2. A. xylinum produces celluVol.
52 No. 3/2005, 691–698
on-line at: www.actabp.pl
692 2005A. Krystynowicz and others
lose on the surface of liquid and solid culture media.
Gelatinuous, leather-like mats formed on the
surface of liquid culture media under stationary
culture conditions contain bacterial cells entrapped
in a network of cellulose fibers. Under agitated culture
conditions, deposition of pellicle is disrupted
and cellulose forms irregular granules stellate and
fibrous strands (Bielecki et al., 2001). On agar media
A. xylinum forms colonies with equal or undulate
edges, transparent or white, smooth or rough, flat or
convex.
A. xylinum strains are prone to spontaneous
mutations yielding cellulose non-producing cells,
which is one of the major problems facing commercial
exploitation of bacterial cellulose biosynthesis.
The appearance of Cel– forms in agitated cultures
was first described by Schramm and Hestrin (1954)
who isolated three different types of A. xylinum cells
distinguished by morphology of colonies and efficiency
of cellulose biosynthesis :
— Type I: wild-type, cellulose-producing (Cel+) cells;
— Type II: cellulose-nonproducing forms (Cel–) capable
of reverting through passages;
— Type III: non-reverting cellulose-nonproducing
forms (Cel–).
The morphology of Cel+ and Cel– colonies is different.
Colonies of Cel– cells are rough, flat, slimy with
undulate edges, in contrast to those of the Cel+ type,
which are spherical with smooth edges, gelatinous
and convex.
The frequency of Cel+ to Cel– conversion depends
on culture conditions and their changes. Cellulose-producing
cells dominate in stationary cultures
and produce on the surface of culture medium
a thick cellulose mat, called a pellicle, in which the
embedded bacterial cells have contact with the oxygen-rich
liquid/air interface. Sufficient and uniform
aeration of liquid culture media under agitated culture
conditions favors spontaneous appearance of
Cel– cells, which become dominating in the population.
Thus aeration of culture media is believed to
be a factor against discriminating Cel+ A. xylinum
cells (Leisinger et al., 1966).
Studies on the effect of culture medium composition
on the efficiency of cellulose synthesis under
agitated culture conditions revealed that the highest
yields of this biopolymer were achieved in media
supplemented with ethanol (2%, v/v). According to
Son et al. (2001) this phenomenon resulted not from
a change in metabolism but from the lack of conversion
of Cel+ cells to Cel– ones. Hence, the spontaneous
change of A. xylinum Cel+ cells to the cellulosenegative
phenotype in agitated cultures can be decreased
by addition of ethanol to culture media (Son
et al., 2001; Krystynowicz et al., 2002).
Cellulose synthesis by A. xylinum cells is catalyzed
by four key enzymes:
● glucokinase (EC 2.7.1.2), responsible for phosphorylation
on C-6 of glucose,
● phosphoglucomutase (EC 5.4.2.2), which catalyzes
isomerization of glucose-6-phosphate to glu-
cose-1-phosphate,
● glucose-1-phosphate uridylyltransferase (EC
2.7.7.9), which synthesizes UDP-glucose (UDPG),
● cellulose synthase (EC 2.4.1.12), which produces
cellulose from UDP-glucose.
Glucose-1-phosphate uridylyltransferase (also
known as UDPG pyrophosphorylase) is thought to
play an important role in cellulose synthesis because
the Cel– forms of A. xylinum lack this enzyme (Valla
et al., 1989). However, the genetics and regulation of
transcription of this enzyme in bacteria producing
cellulose or related polysaccharides have not been
explained.
According to Valla et al. (1987), also plasmids
can be involved in cellulose biosynthesis. Based on
electrophoretic separations of plasmid DNA isolated
from A. xylinum it was found that nine of thirteen
isolated mutants displayed changes in plasmid DNA
profile as compared to the wild-type strain.
It was also reported that insertion elements
could be involved in polysaccharide synthesis in
Pseudomonas atlantica (Bartle� & Silverman, 1989)
and Xanthomonas campestris (Ho�e et al., 1990). An
unstable polysaccharide production by Zoogloea ramigera
resulted from rearrangement of its DNA (Easson
et al., 1987). The presence of insertion element
IS1031 (950 bp) was detected in A. xylinum ATCC
23769. Cel– mutants of this strain possessed two or
more IS1031 elements, and furthermore, their DNA
was rearranged within the IS elements, as compared
to cellulose-producing wild-type strain (Coucheron,
1991). Therefore, the loss of capability of some A.
xylinum strains to produce cellulose is supposed to
result from dislocation of insertion elements and inactivation
of gene(s) responsible for cellulose synthe-
sis.
The presented studies have been undertaken
to compare wild A. xylinum E25 strain and Cel– forms
possessing cellulose-negative phenotype at the molecular
level. Genome DNA was isolated from Cel+
and Cel– A. xylinum E25 cells, and compared by PCRMP
(PCR-melting profiles) analysis. Independently,
protein profiles of both types of cells were compared
by means of two-dimensional electrophoresis. Spots
corresponding to the proteins inherent for only one
type of the cells were excised from gels, digested
and sequenced. The peptide sequences determined
in this way were subsequently aligned with protein
sequences deposited in databases. Two enzymes involved
in the synthesis of various glucose-containing
polysaccharides, namely phosphoglucomutase
and glucose-1-phosphate uridylyltransferase, were
identified as lacking in the Cel– phenotype. Possible
reasons of this phenomenon are discussed.
Vol. 52 693Molecular aspects of bacterial cellulose synthesis
MATERIALS AND METHODS
Microorganism. A. xylinum E25 strain from
the pure culture collection of the Institute of Technical
Biochemistry of the Technical University of Lodz
was used for the studies. It was maintained on agar
slants at 4oC.
Culture medium. The Schramm and Hestrin
(1954) medium (SH medium), containing (g/l):
glucose 20.0, yeast extract 5.0, bactopeptone 5.0,
Na2HPO4 2.7, citric acid 1.15, and MgSO4 × 7H2O 5.7,
with initial pH adjusted to 5.7 was used throughout
the cultures.
Culture conditions. Inoculum was prepared
by transferring a single A. xylinum Cel+ or Cel– colony
from SH agar medium into a 50-ml Erlenmeyer
flask containing 10 ml of liquid SH medium. This
cell suspension (10 ml) was added into a 500 ml Erlenmeyer
flask containing 100 ml of a fresh SH medium.
Agitated cultures were incubated at 30oC on a
rotary shaker at 90 r.p.m.
Selection of non-reverting mutant. A non-reverting
form of A. xylinum E25 was obtained by reiterated
passages under agitated culture conditions.
The cultures were carried out for 48 h at 30oC on a
rotary shaker at 90 r.p.m. This process was repeated
three times (3 passages). Cell suspension from each
passage was spread on SH agar medium and incubated
for 6 days at 30oC. Colonies with morphology
characteristic for Cel– forms were suspended in 5 ml
of liquid SH medium and incubated for 3 days at
30oC to test if the revertants were able to form the
cellulose pellicle. Non-reverting forms were isolated
from cellulose-free culture broths and used in further
studies.
Sample preparation for 2D electrophoresis.
A single colony of Cel+ or Cel– A. xylinum E25 was
suspended in 5 ml of SH medium and incubated for
48 h. From this culture, a 0.5 ml inoculum was added
do 10 ml of SH medium (in a 50 ml Erlenmayer
flask) and incubated for 48 h at 30oC. Bacterial
cells were harvested by centrifugation (9000 r.p.m.,
20 min, 4oC), washed with 40 mM Tris/HCl and
disrupted by sonication (3 × 2 min). Proteins were
separated using 2-D Clean-Up Kit (Amersham Biosciences)
and suspended in a solution containing: 7
M urea, 2 M thiourea, 4% Chaps, 1% DTT and 0.8%
ampholites pH 3–10, according to Westemeier et al.
(2002).
Two-dimensional gel electrophoresis. Twodimensional
electrophoresis was carried out in an Ettan
apparatus (Amersham Biosciences). In the first
dimension the separation of proteins was based on
their isoelectric points (Immobiline DryStrip gel pH
3–10), and in the second dimension the proteins
were separated by SDS/PAGE gel (12.5%, 200 × 260
mm) according to the protocol of an Amersham
Pharmacia Biotech Technical Manual (1999). The
analyzed samples contained 75 µg of proteins which
were visualized by Silver Staining Kit (Sigma).
DNA isolation. DNA was isolated from Cel+
and Cel– cells of A. xylinum E25 according to Maniatis
et al. (1989). Bacterial cells were pelleted by
centrifugation (1300 × g, 15 min), washed with 10
ml TGE (25 mM Tris/HCl, 50 mM glucose, 10 mM
EDTA, pH 8) and again centrifuged (1300 × g, 15
min). The pellet was suspended in 10 ml TGE supplemented
with lysozyme (10 mg/ml) and incubated
at room temperature for 10 min. Next, proteinase K
(100 µg/ml) and 1% SDS were added and the mixture
was kept at 37oC for 30 min. Then, 2 ml of 5
M aqueous NaCl solution and 1.5 ml of 10% CTAB
in 0.7 M NaCl were added to the mixture, which
was incubated at 65oC for 20 min. A�er mixing with
equal volume of phenol/chloroform, the mixture was
centrifuged (3600 × g, 10 min), extracted with a double
volume of chloroform/isoamyl alcohol (24:1, v/v),
carefully mixed with 2.5 volumes of cold (–20oC)
ethanol (96%), and le� for 15 min. DNA was collected
by centrifugation, the pellet washed 2 times
with 70% ethanol, centrifuged at 3600 × g for 10 min
and dried.
PCR-MP (melting profiles). The PCR-MP
procedure was carried out as described by Masny
and Płucienniczak (2003). Bacterial DNA prepared
as described above was digested with restriction
endonuclease HindIII, extracted with phenol/chloroform,
precipitated, the precipitate was dissolved
in ligation mixture containing two oligonucleotide
adapters: 5’-CTCACTCTCACCAACGTCGAA-3’
(POWIH) and 5’-AGCTTTCGACGTTGG-3’ (HIL)
(20 pmol each), in a total volume of 20 µl ligation
buffer (66 mM Tris/HCl, pH 8.5, 6.6 mM MgCl2,
10 mM DTT, 66 mM ATP; Amersham Pharmacia
Biotech). The mixture was heated in a water bath
for 2 min at 56oC and cooled for 10 min at room
temperature. Subsequently, 1 µl of T4 DNA ligase
(1U/µl) was added and the samples were incubated
overnight at 16oC.
Then, PCR reaction was carried out in an MJ
Research PTC200 thermocycler at denaturation temperatures
(Td) 80, 81 and 82oC. The reaction mixture
contained 50 pmol of POWBAGCT primer 5’-CTCACTCTCACCAACGTCGAAAGCTT-3’,
100 µmol
each dNTPs and 1 µl of ligation mixture in 50 µl of
PCR buffer. Samples (8 µl out of 50 µl) were loaded
on 6% polyacrylamide gel (bisacrylamide/acrylamide
= 1:60) with TAE buffer. The gel was stained with
ethidine bromide and visualized in UV at 302 nm, according
to the protocol of Masny and Płucienniczak
(2003). DNA fragments of different thermal stability
were isolated from the gel (Dybczyński & Plucienniczak,
1988), digested with HindIII restriction endonuclease,
ligated with the pBluScriptSK– plasmid
(Stratagene) and cloned in Escherichia coli, strain
NM522. The nucleotide sequences of the inserts were
694 2005A. Krystynowicz and others
determined in the Institute of Biochemistry and Biophysics
(PAS, Warsaw, Poland).
Protein identification. Proteins were identified
by peptide mass fingerprinting using MALDITOF
mass spectrometry and by partial peptide sequencing
with ion trap electrospray mass spectrometry.
Trypsin autodigestion peaks were exluded from
the database searching. Molecular mass data were
obtained from EXPASY (h�p://www.matrixscience.
com/). Monoisotopic peptide masses were used to
search protein databases to match and subsequently
identify individual protein spots.
RESULTS
PCR-MP
Figure 1 presents a profile of DNA fragments
liberated by HindIII restrictase from the genomes
under investigation and amplified at low Td’s according
to the procedure described by Masny and
Płucienniczak (2003). It appeared that in the case of
the cellulose producer strain (Cel+ form) one of the
DNA fragments migrating just above 242 bp was
less stable than the corresponding fragment from
Cel–. The DNA fragment from Cel+ cells was amplified
at the denaturation temperature of 81oC, while
its counterpart from Cel– became visible when the
Td was 1oC higher. The less stable fragment (from
Cel+ cells) has a stretch of four T-residues (positions
76–79, Fig. 2), while its counterpart from Cel– contains
at the same positions three T residues. One can
suggest that the deletion of a single T residue in the
Cel– fragment is responsible for its increased thermal
stability as compared to that of the corresponding
Cel+ DNA fragment.
Two-dimensional electrophoresis
Figures 3A and 3B present intracellular proteins
(75 µg per gel) isolated from the wild A. xylinum
E25 (Cel+) strain and its Cel– counterpart. Electrophoretic
separation was carried out in the pH
range from 4 to 7 in 12% polyacrylamide gel. Proteins
produced only by Cel+ cells and not synthesized
by the Cel– mutant, as well as proteins found
only in Cel– mutant and lacking in wild-type strain
are marked on the gels.
Protein identification
Proteins unique to Cel+ or Cel– cells were excised
from the gel and sequenced. The results of the
analysis of peptides derived from intracellular A. xylinum
E25 (Cel+) proteins supposedly involved in cellulose
biosynthesis and missing in cells of the cellulose-nonproducing
mutant are collected in Table 1.
These peptides are similar to amino-acid sequences
of the following enzymes:
— phosphoglucomutase from: Gluconacetobacter
xylinus, Mycobacterium tuberculosis (CDC1551), Bifidobacterium
longum (NCC2705 and DJO10A), Streptomyces
avermitilis (MA-4680), Pseudomonas fluorescens
(PfO-1), Desulfovibrio vulgaris subsp. vulgaris str., and
Leifsonia xyli subsp. xyli str. (CTCB07),
— glucose-1-phosphate uridylyltransferase from:
Gluconacetobacter xylinus and Acetobacter pasteurianus.
DISCUSSION
Cellulose non-producing forms of A. xylinum
were for the first time described in 1954 (Schramm
& Hestrin, 1954). An influence of culture conditions
on the frequency of Cel– phenotypes (Leisinger et
al., 1966; Son et al., 2001; Krystynowicz et al., 2002)
and activities of enzymes involved in the synthesis
of this polysaccharide (Valla et al., 1989) have been
reported by several groups. An influence of plasmid
DNA and insertion elements in the A. xylinum
genome on the efficiency of cellulose synthesis was
also described (Valla et al., 1987; Coucheron, 1991).
However, the reasons of the appearance of
cellulose non-producing A. xylinum forms in agitated
cultures remain obscure. One can suppose that this
effect results from different expression of genes encoding
enzymes responsible for glucose metabolism
and/or cellulose synthesis. Wong et al. (1990) identified
in A. xylinum the bcs operon encoding four proteins
essential for bacterial cellulose synthesis. The
first gene in this operon, bcsA, encodes the regulatory
subunit of cellulose synthase (CS), which binds
c-di-GMP acting as bacterial second messanger and
activator in the cellulose synthesis process (Wong et
al., 1990). The second gene, bcsB, encodes the catalytic
subunit of CS, which binds the substrate, UDPFigure
1. Electrophoretic pa�ern a�er PCR-MP of DNA
preparations from Cel– and Cel+ bacteria (lanes – and +,
respectively).
Experiments for each temperature were repeated two
times. Td’s of the PCR are marked above electrophoretic
lines. DNA bands taken for cloning and nucleotide sequencing
are marked with arrows. Lanes marked M, DNA
markers: 110+111, 147, 190, 242, 331, 404, 476 and 489 base
pairs.
Vol. 52 695Molecular aspects of bacterial cellulose synthesis
glucose, and presumably catalyzes the polymerization
of 1,4-β-�-glucan. Two other genes belonging
to this operon, dgc and pdeA, encode diguanylate
cyclase and phosphodiesterase A, which are responsible
for the synthesis and degradation of c-di-GMP,
respectively (Ross et al., 1991). Coucheron (1991)
reported insertion of the IS1031 element upstream
from the transcription start of this operon, which resulted
in the deficiency of cellulose synthesis.
Our studies carried out with the use of the
PCR-MP approach (Masny & Płucienniczak, 2003)
indicate that genomic libraries from Cel+ and Cel–
strains contain DNA fragments of different thermal
stability. In the case of the cellulose producer (Cel+
form) a DNA fragment migrating just above 242 bp
is less stable than the corresponding fragment from
Cel– cells. A comparison of their sequences has revealed
that the thermally less stable fragment (Cel+)
has array of four T-residues (positions 76–79, Fig. 2),
while its counterpart from Cel– cells contains at the
same positions only three T residues.
Alignment of the 218 nt sequence identified in
our studies with those deposited in the GenBank database
(National Center for Biotechnology Information)
reveals that several genes of Acetobacter xylinum
and Gluconacetobacter oxydans contain in their 3’-terminal
parts (downstream from their ORFs) nucleotide
sequences similar to the fragment sequenced in
our studies. The 2,702,173 nt-long Gluconacetobacter
oxydans genome contains five regions showing significant
homology to this fragment (Table 4). However,
it should be noticed that all these sequences are
shorter than 218 nt (see Fig. 2). Because the 218 nt
fragment contains inverted repeats (see Fig. 2), one
can speculate that the corresponding RNA arising
during transcription can form stem-loop or hairpin
secondary structures (see Fig. 4). It is known that
these types of RNA structures can regulate gene expression
by their involvement in the transcriptional
terminator/antiterminator mechanism (Turner et al.,
1994; Yanofsky, 2000). It cannot be excluded that the
loss of the single T residue results in a change of the
RNA secondary structure which may influence transcription
of the genes involved in cellulose synthesis.
Figure 2. The nucleotide sequence of the 218 nt element from Cel+ A. xylinum E25 strain isolated a�er PCR-MP
(shown in capital le�ers) and its comparison with the corresponding Gluconacetobacter xylinus and Acetobacter europaeus
nucleotide sequences.
The array of T residues where deletion of one T occurs in Cel– is shown in bold. Inverted repeat sequences are shown
by gray background. The GenBank accession numbers of the sequences under comparison are shown in square brackets.
Figure 3. 2-D separation of a lysate of Acetobacter xylinum
E25 cells.
Proteins from (Cel–) (A) and wild-type (Cel+) (B) were
separated by 2-D electrophoresis and visualized by silver
staining. The pH gradient (horizontal direction) runs from
pH 3–10 and the SDS/PAGE separation (perpendicular direction)
was performed in 12.5% gel. The arrows indicate
the proteins identified as phosphoglucomutase (spots 3
and 4) and UDP-glucose pyrophosphorylase or glucose-1phosphate
uridylyltransferase (spot 9).
696 2005A. Krystynowicz and others
Table 1. Results of analysis of peptide fragments derived from gel slices numbered 3, 4 and 9 (according to Fig. 3).
Gel slice number 3 Gel slice number 4 Gel slice number 9
phosphoglucomutase (PGM) phosphoglucomutase (PGM) glucose-1-phosphate uridylyltransferase
(UDPGP)
amino acids: 555;
Mr: 59654
amino acids: 555;
Mr: 59654
amino acids: 284
Mr: 31792
protein ID: P38569
(swissprot); gi|585669
(according to NCBI)
protein ID: P38569
(swissprot); gi|585669
(according to NCBI)
protein ID: P27897
(swissprot); gi|7381245
(according to NCBI)
score: 1394
queries matched: 36
score: 1735
queries matched: 44
score: 207
queries matched: 5
sequence coverage: 68% sequence coverage: 76% sequence coverage: 19%
Table 2. Localization of DNA fragments homologous to the 218 nt element
Bacterial strain
GenBank
Accession Number
(literature reference)
Region of
homology
within the 218 nt
DNA
Element-coordi-
nates
Region of
homology
in bacterial DNA,
data from
GenBank-coordi-
nates
Genes in the closest proximity to the
fragment homologous to the 218 nt
element; (gene coordinates; name of
the protein; protein id)
Gluconacetobacter
xylinus b
AB071166
(Tajima et al., 2001)
1–86 716–799 1137..3356; β-glucosidase
Gluconacetobacter
xylinus b
AB091059
(unpublished)
1–86 378–461 808..3036; β-galactosidase
Acetobacter xy-
linum
L24077.1
(Brautaset et al., 1994)
1–75 1942–2012 135..1812; phosphoglucomutase
Gluconobacter
oxydans 621H
CP000009.1
(Prust et al., 2005)
56–1 1137409–1137463 1136433..1137404; putative oxidoreductase;
protein id: AAW60805.1 and
1137522..1137968; hypothetical protein;
protein id: AAW60806.1
Gluconobacter
oxydans 621H
CP000009.1
(Prust et al., 2005)
154–208 1137409–1137467 1136433..1137404; putative oxidoreductase;
protein id: AAW60805.1 and
1137522..1137968; hypothetical protein;
protein id: AAW60806.1
Gluconobacter
oxydans 621H
CP000009.1
(Prust et al., 2005)
1–56 797775–797829 795730..797730; oligopeptidase; protein
id: AAW60504.1 and 797839..798789;
putative multidrug efflux pump;
protein id: AAW60505.1
Gluconobacter
oxydans 621H
CP000009.1
(Prust et al., 2005)
1–55 916535–916588 915033..916508; NADPH-dependent
�-sorbose reductase; protein id:
AAW60623.1 and 916634..917890 (c);
UDP-N-acetylglucosamine 1-carboxy-
vinyltransferase;
protein id: AAW60624.1
Gluconobacter
oxydans 621H
CP000009.1
(Prust et al., 2005)
218–149 916521–916594 915033..916508; NADPH-dependent
�-sorbose reductase; protein id:
AAW60623.1 and 916634..917890 (c);
UDP-N-acetylglucosamine 1-carboxy-
vinyltransferase;
protein id: AAW60624.1
Acetobacter xy-
linus
AB010645
(Nakai et al., 1998)
84–13 16568–16638 12448..14655; β-glucosidase;
protein id: BAA31467.1
Acetobacter xy-
linus
AB010645
(Nakai et al., 1998)
1–60 15344–15402 12448..14655; β-glucosidase;
protein id: BAA31467.1
Acetobacter euro-
paens
Y08696.1
(Thurner et al., 1997)
60–1 5991–6049 3604..5925; aldehyde dehydrogenase;
protein id: BAA00408.1
Acetobacter euro-
paens
Y08696.1
(Thurner et al., 1997)
148–218 5989–6062 3604..5925; aldehyde dehydrogenase;
protein id: BAA00408.1
Acetobacter po-
lyoxogenes
D00521.1
(Tamaki et al., 1989)
62–1 2623–2683 236..2557; aldehyde dehydrogenase
precursor; protein id: BAA00408.1
Acetobacter xy-
linum
Y18467.1
(Edwards et al., 1999)
1–86 1041–1123 32..997; glucosyl transferase;
protein id: CAB44443.1
Vol. 52 697Molecular aspects of bacterial cellulose synthesis
A comparison of the protein profiles of wildtype
(Cel+) A. xylinum strains and their Cel– mutants
revealed some differences. Cel– cells do not
synthesize two key enzymes involved in cellulose
biosynthesis, namely phosphoglucomutase and glucose-1-phosphate
uridylyltransferase. The lack of
glucose-1-phosphate uridylyltransferase in cellulose
non-producing cells was reported earlier (Valla et
al., 1989). Our studies proved that A. xylinum cells
of Cel– phenotype are also deprived of phosphoglucomutase.
Although it is known that phosphoglucomutase
and glucose-1-phosphate uridylyltransferase
(or UDP-glucose pyrophosphorylase) play a critical
role in cellulose synthesis, their genetics and regulation
have not been studied in detail.
Taking into account the results of the PCRMP
analysis and 2D electrophoresis as well as the
alignment of the nucleotide and peptide sequences,
one can suggest that the cellulose deficiency in A.
xylinum Cel– cells can result from the lack of expression
of the genes encoding phosphoglucomutase and
glucose-1-phosphate uridylyltransferase. Our finding
that in the Cel– form the ORF encoding phosphoglucomutase
may be flanked by a DNA fragment of a
sequence different than that in Cel+ cells can indicate
a possible transcriptional repression of the gene
expression.
Acknowledgements
This work was financially supported by grant
No. 2 P04B 017 26 from the State Commi�ee for Scientific
Research (KBN, Poland).
REFERENCES
Bielecki S, Krystynowicz A, Turkiewicz M, Kalinowska
H (2005) Bacterial cellulose. In Biotechnology of Biopolymers,
Steinbu�el A, Doi Y, eds, vol. 1, pp 381–434.
Willey-VCH, Weinheim.
Bartle� DH, Silverman M (1989) Nucleotide sequences of
IS492, a novel insertion sequences causing variation in
extracellular polysaccharide production in the marine
bacterium Pseudomonas atlantica. J Bacteriol 171: 1763–
1766.
Brautaset T, Standal R, Fjaervik E, Valla S (1994) Nucleotide
sequence and expression analysis of the Acetobacter
xylinum phosphoglucomutase gene. Microbiology
140: 1183–1188.
Coucheron DH (1991) An Acetobacter xylinum insertion sequence
element associated with inactivation of cellulose
production. J Bacteriol 173: 5723–5731.
Dybczyński I, Płucienniczak A (1988) A protocol for DNA
fragment extraction from polyacrylamide gels. Biotechniques
6: 924–926.
Easson DD Jr, Sinskey AJ, Peoples OP (1987) Isolation of
Zoogloea ramigera I-16 M exopolysaccharide biosynthetic
genes and evidence for instability within this region.
J Bacteriol 169: 4518–4524.
Edwards KJ, Jay AJ, Colquhoun IJ, Morris VJ, Gasson MJ,
Griffin AM (1999) Generation of a novel polysaccharide
by inactivation of the aceP gene from the acetan
biosynthetic pathway in Acetobacter xylinum. Microbiology
145: 1499–1506.
Ho�e B, Roth-Arnold I, Puhler A, Simon R (1990) Cloning
and analysis of a 35.3 kb DNA region involved in exopolysaccharide
production by Xanthomonas campestris.
J Bacteriol 172: 2804–2807.
Krystynowicz A, Czaja W, Wiktorowska-Jezierska A, Goncalves-Miśkiewicz
M, Turkiewicz M, Bielecki S (2002)
Factors affecting the yield and properties of bacterial
cellulose. J Ind Microbiol Biotechnol 29: 189–195.
Leisinger T, Wiemken A, E�linger L (1966) Űber cellulosefreie
Mutanten von Acetobacter xylinus. Arch Microbiol
54: 21–36.
Maniatis T, Sambrook J, Fritsch EF (1989) Molecular Cloning.
A Laboratory Manual. 2 edn.
Masny A, Płucienniczak A (2003) Ligation mediated PCR
performed at low denaturation temperatures – PCR
melting profiles. Nucleic Acids Res 31: e114.
Nakai T, Moriya A, Tonouchi N, Tsuchida T, Yoshinaga
F, Horinouchi S, Sone Y, Mori H, Sakai F, Hayashi T
(1998) Control of expression by the cellulose synthase
(bcsA) promoter region from Acetobacter xylinum BPR
2001. Gene 213: 93–100.
Protein Electrophoresis, Technical manual (1999) Amersham
Pharmacia Biotech.
Prust C, Hoffmeister M, Liesegang H, Wiezer A, Fricke
WF, Ehrenreich A, Go�schalk G, Deppenmeier U (2005)
Complete genome sequence of the acetic acid bacterium
Gluconobacter oxydans. Nat Biotechnol 23: 195–200.
Ring DF, Nashed W, Dow T (1986) Liquid loaded pad for
medical applications. US Patent 45884000.
Ross P, Mayer R, Benziman M (1991) Cellulose biosynthesis
and function in bacteria. Microbiol Rev 55: 35–58.
Schramm M, Hestrin S (1954) Factors affecting production
of cellulose at the air/liquid interface of a culture of
Acetobacter xylinum. J Gen Microbiol 11: 123–129.
Son HJ, Heo MS, Kim YG, Lee SJ (2001) Optimization of
fermentation conditions for the production of bacterial
cellulose by a newly isolated Acetobacter sp. A9 in shaking
cultures. Biotechnol Appl Biochem 33: 1–5.
Tajima K, Nakajima K, Yamashita H, Shiba T, Munekata
M, Takai M (2001) Cloning and sequencing of the betaglucosidase
gene from Acetobacter xylinum ATCC 23769.
DNA Res 8: 263–269.
Tamaki T, Horinouchi S, Fukaya M, Okumura H, Kawamura
Y, Beppu T (1989) Nucleotide sequence of the
membrane-bound aldehyde dehydrogenase gene from
Acetobacter polyoxogenes. J Biochem (Tokyo) 106: 541–
544.
Thurner C, Vela C, Thony-Meyer L, Meile L, Teuber M
(1997) Biochemical and genetic characterization of the
acetaldehyde dehydrogenase complex from Acetobacter
europaeus. Arch Microbiol 168: 81–91.
Turner RJ, Lu Y, Switzer RL (1994) Regulation of the Bacillus
subtilis pyrimidine biosynthetic gene cluster by an
autogenous transcriptional a�enuation mechanism. J
Bacteriol 176: 3708–3722.
Westemeier R, Naven T (2002) Proteomics in Practise. A Laboratory
Manual of Proteome Analysis. Wiley-VCH.
Wong HC, Fear AL, Calhoon RD, Eichinger GH, Mayer
R, Amikam D, Benziman M, Gelfand DH, Meade JH,
Emerick AW, Bruner R, Ben-Bassat A, Tal R (1990) Genetic
organization of the cellulose synthase operon in
Acetobacter xylinum. Proc Natl Acad Sci USA 87: 8130–
8134.
Valla S, Coucheron DH, Kjosbakken J (1987) The plasmids
of Acetobacter xylinum and their interaction with the
host chromosome. Mol Gen Genet 208: 76–83.
Valla S, Coucheron DH, Fjaervik E, Kjosbakken J, Weinhouse
H, Ross P, Amikam D, Benziman M (1989) Clon-
698 2005A. Krystynowicz and others
ing of a gene involved in cellulose biosynthesis in Acetobacter
xylinum: complementation of cellulose-negative
mutant by the UDPG pyrophosphorylase structural
gene. Mol Gen Genet 217: 26–30.
Yamada Y (2000) Transfer of Acetobacter oboediens Sokollek
et al. 1998 and Acetobacter intermedius Boesch et al.,
1998 to the genus Gluconacetobacter as Gluconacetobacter
oboediens comb. nov. and Gluconacetobacter intermedius
comb. nov. Int J Syst Evol Microbiol 50: 2225–2227.
Yanofsky C (2000) Transcription a�enuation: once viewed
as a novel regulatory strategy. J Bacteriol 182: 1–8.