Polish Journal of Microbiology

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N. gonorrhoeae 220 RifRNalR, 13; N. gonorrhoeae WR302 RifRNalR, 14; ..... tional analysis, performed with a series of mutated plasmid mini-derivatives, showed that the replicator ...... K r u g e r R., S.A. R a k o w s k i and M. F i l u t o w i c z.
POLSKIE TOWARZYSTWO MIKROBIOLOGÓW POLISH SOCIETY OF MICROBIOLOGISTS

Polish Journal of Microbiology formerly

Acta Microbiologica Polonica

2006 POLSKIE TOWARZYSTWO MIKROBIOLOGÓW

EDITORS Miros³awa W³odarczyk (Editor in Chief) Jaros³aw Dziadek, Anna Kraczkiewicz-Dowjat, Anna Skorupska, Hanna Dahm El¿bieta Katarzyna Jagusztyn-Krynicka (Scientific Secretary) EDITORIAL BOARD President: Andrzej Piekarowicz (Warsaw, Poland) Katarzyna Brzostek (Warsaw, Poland), Ryszard Chróst (Warsaw, Poland), Waleria Hryniewicz (Warsaw, Poland), Miros³aw Kañtoch (Warsaw, Poland), Donovan Kelly (Warwick, UK), Tadeusz Lachowicz (Wroc³aw, Poland), Wanda Ma³ek (Lublin, Poland), Zdzis³aw Markiewicz (Warsaw, Poland), Gerhard Pulverer (Cologne, Germany), Geoffrey Schild (Potters, Bar, UK), Wac³aw Szybalski (Madison, USA), Torkel Wadström (Lund, Sweden), Jadwiga Wild (Madison, USA) EDITORIAL OFFICE Miecznikowa 1, 02-096 Warsaw, Poland tel. 48 (22) 55 41 318, fax 48 (22) 55 41 402 e-mail pjm@ biol.uw.edu.pl Archives of Acta Microbiologica Polonica, from 2004 Polish Journal of Microbiology online www.microbiology.pl\pjm\ at PTM Journals online www.microbiology.pl Visit the home page to browse contents, gallery, links page and instructions to authors in HTML and PDF formats Editorial correspondence should be addressed to Editors of Polish Journal of Microbiology 02-096 Warsaw, Miecznikowa 1, Poland Correspondence regarding subscription and spedition of Polish Journal of Microbiology should be addressed to National Institute of Public Health, Division of Clinical Microbiology and Prevention of Infections 00-725 Warsaw, Che³mska 30/34 tel. 48 (22) 841 33 67, fax 48 (22) 841 29 49, e-mail: [email protected]

QUARTERLY OF POLISH SOCIETY OF MICROBIOLOGISTS, PUBLISHED WITH THE FINANCIAL SUPORT OF THE MINISTRY OF EDUCATION AND SCIENCE

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Front cover: Colonies of bacteria with nitrifying activity isolated from landfill leachate (courtesy of Renata Matlakowska Ph.D.)

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Polish Journal of Microbiology formerly Acta Microbiologica Polonica

2006, Vol. 55, No 4

CONTENTS ORIGINAL PAPERS

Analysis of the filamentous bacteriophage genomes integrated into Neisseria gonorrhoeae FA1090 chromosome

PIEKAROWICZ A., MAJCHRZAK M., K£Y¯ A., ADAMCZYK-POP£AWSKA M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 251

Replication system of plasmid pMTH4 of Paracoccus methylutens DM12 contains an enhancer

SZYMANIK M., WELC-FALÊCIAK R., BARTOSIK D., W£ODARCZYK M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261

Acquisition of iron by enterococci; some properties and role of assimilating ferric iron reductases

LISIECKI P., MIKUCKI J. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271

Susceptibility of Listeria monocytogenes strains isolated from dairy products and frozen vegetables to antibiotics inhibiting murein synthesis and to disinfectants

POPOWSKA M., OLSZAK M., MARKIEWICZ M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279

Partial characterization and optimization of production of extracellular "-amylase from Bacillus subtilis isolated from culturable cow dung microflora

SWAIN M.R., KAR S., PADMAJA G., RAY R.C. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289

Production of tannase through submerged fermentation of tannin-containing plant extracts by Bacillus licheniformis KBR6

DAS MOHAPATRA P.K., MONDAL K.C., PATI B.R. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 297

In vitro activity of synthetic antimicrobial peptides against Candida KAMYSZ W., NADOLSKI P., KÊDZIA A., CIRIONI O., BARCHIESI F., GIACOMETTI A., SCALISE G., £UKASIAK J., OKRÓJ M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303

Effects of culture conditions on production of extracellular laccase by Rhizoctonia practicola

JANUSZ G., ROGALSKI J., BARWIÑSKA M., SZCZODRAK J. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 309

Growth of Penicillum verrucosum and production of ochratoxin A on nonsterilized wheat grain incubated at different temperatures and water content

CZABAN J., WRÓBLEWSKA B., STOCHMAL A., JANDA B. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321

Detection of cytomegalovirus and Helicobacter pylori DNA in arterial walls with grade III atherosclerosis by PCR

KILIC A., ONGURU O., TUGCU H., KILIC S., GUNEY C., BILGE Y. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333

Detection of methicillin resistance in hospital environmental strains of coagulase-negative staphylococci

NOWAK T., BALCERCZAK E., MIROWSKI M., SZEWCZYK E.M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339

VOLUME 55 CONTENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 345 AUTHOR INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 348 ACKNOWLEDGMENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349 INSTRUCTIONS TO AUTHORS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351

Polish Journal of Microbiology 2006, Vol. 55, No 4, 251– 260

Analysis of the Filamentous Bacteriophage Genomes Integrated Into Neisseria gonorrhoeae FA1090 Chromosome ANDRZEJ PIEKAROWICZ*, MICHA£ MAJCHRZAK, ANETA K£Y¯ and MONIKA ADAMCZYK-POP£AWSKA

Department of Virology, Institute of Microbiology, Warsaw University, Warsaw, Poland Received 21 September 2006, accepted 10 October 2006 Abstract Bioinformatic analysis of the genome sequence of Neisseria gonorrhoeae revealed presence of four specific prophage islands. Based on the similarity with other DNA phage sequences they seem to belong to the filamentous ssDNA phages group. Phages belonging to this group are also present in the genome of Neisseria meningitidis. The nucleotide and amino acids sequence of NgoM6 and NgoM7 show similar genetic organization and high homology on DNA and amino acid level. The NgoM9 contains only part of the genomes of the NgoM6-8 prophages. Several functionally same genes of different origin are duplicated, with no homology to their counterparts in phages NgoM6, NgoM7 and NgoM9. The prophage sequences of nucleotides of NgoM6 and NgoM7 contain specific blocks of genes responsible for phage DNA replication and structural proteins. Comparative analysis at nucleotide and amino acid level suggests that these sequences can encode functionally active phages. The genetic organization of the NgoM6 suggests that it can serve as a prototype of filamentous phage of N. gonorrhoeae. Presence of the genomic ssDNA of these phages in the cultures of N. gonorrhoeae confirms this conclusion. K e y w o r d s: prophage sequences, ssDNA bacteriophages

Introduction The sequencing of the bacterial genomes revealed that presence of integrated viral genomes (prophages) is a common phenomenon. As much as 51 of the 82 genomic sequences published since year 2003 (Casjens, 2003) carry prophages, and these contain 230 recognizable putative prophages (for review see, Casjens, 2003; Canchaya et al., 2004; Brüssow et al., 2004). Prophages can constitute up to 10– 20% of bacterium’s genome and are major contributions between individual species (Casjens, 2003). The acquisition of prophages would be an irrelevant process for the evolution of bacteria if phages did not transfer genes useful to the lysogens (Brüssow et al., 2004). A lot of these genes play an important role in the pathogenicity or in general fitness of the bacterial host (Brüssow et al., 2004; Canchaya et al., 2004; Casjens, 2003). This influence of the presence of prophages on phenotypic properties is termed lysogenic conversion. The important role is played by so called morons, defined as extra genes in the prophage sequences; they do not have a phage function but may act as fitness factors for the lysogen. The list of such putative and known morons of phages and phage-like elements found in pathogenic bacteria is very long and representing a broad group of genes (Brüssow et al., 2004). Among them there are type III effector proteins or CT, the principal virulence factor of Vibrio cholerae (Brüssow et al., 2004). The analysis of the prophages suggested that after being integrated into bacterial genomes, they undergo a complex decay process which involve inactivating point mutations, genome rearrangements, modular exchanges, invasion by further mobile DNA elements, and massive DNA deletions (Canchaya et al., 2004; Casjens, 2003). However, the final answer whether the particular gene is in fact inactivated has to be proven experimentally, since it was shown that the demaged by deletion genes encoding the RM system in Neisseria gonorrhoeae can be biologically active (Piekarowicz et al., 2001). * Corresponding author: A. Piekarowicz: Institute of Microbiology, Warsaw University, Miecznikowa 1, 02-096 Warsaw, Poland; phone + 48 22 55 41417; e-mail: [email protected]

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4

The presence of the Mu-like prophage sequences was detected in the genomes of serogroup A strains of the epidemic subgroups I, III, IV-1 and VI of Neisseria meningitidis (Masignani et al., 2001; Morgan et al., 2002; Klee et al., 2000). A ~35-kb region designated Pnm1, is inserted within a putative gene encoding an ABC-type transporter. This region contains 46 open reading frames, 29 of which are collinear and homologous to the genes of Escherichia coli Mu phage. The two additional Mu-like sequences were found in N. menigitidis serotype A (Morgan et al., 2002) and the sequence homologous to Pnm1 was found in the genome of serogroup B N. meningitidis, Haemophilus influenzae which is being named FluMu and Deinococcus radiodurans, but not in genome of N. gonorrhoeae FA1090 (Masignani et al., 2001). Recently, the presence of filamentous prophages genomes was detected in N. menigitidis and it was shown that these prophages could be excised resulting in the production of the biologically active phages (Bille et al., 2005). In this report, we present the data indicating the presence of filamentous bacteriophages genomes integrated into N. gonorrhoeae strain FA1090 chromosome. These are able to produce active phages whose genomes can be detected in the cultures of N. gonorrhoeae strains. Experimental Materials and Methods Bacterial strains, plasmids and growth conditions. Escherichia coli K-12 strain XL-1 Blue MRF’ [)(mcrA)183)(mcrCBhsdSMR-mrr)173 endA1 supE44 thi-1 recA1 gyrA96 relA1 lac [F’ proAB lacIqZ )M15Tn10(Tetr)]] was used during manipulation with pH17 plasmid. This strain was grown at 37°C in Luria-Bertani (LB) medium (Sambrook et al., 1989). Antibiotics included in media were used at the following final concentrations (µg/ml): ampicillin 100, tetracycline 10. Neisseria strains were grown in standard gonococcal medium (designated GCP if broth and GCK if agar) (Difco laboratories) plus growth supplements (White and Kellogg, 1965) and 0.042% sodium bicarbonate if in broth or in a 37°C CO2 incubator. N. gonorrhoeae FA1090 strain was obtained from Dr D.C. Stein’s collection. Chromosomal DNAs of N. gonorrhoeae, Neisseria lactamica and N. meningitidis strains were obtained from Dr D.C. Stein (University of Maryland, USA). Plasmid pUC19 was purchased from Fermentas. Detection of the prophage and phage nucleotides sequences. The plasmid pH17 carrying the 5430 bp HindIII fragment of N. gonorrhoeae FA1090 was obtained by chance during cloning of the genes encoding MTases (Piekarowicz et al., 1991). Sequencing of the insert fragment showed that it contains the internal fragment of the NgoM6 spanning from the 1 082 100 to 1 087 500 bp. The dot blot hybridization was carried out using this fragment of DNA as a probe. The total DNA (0.1 µg) extracted from the bacterial cells was used in the hybridization experiments for the detection of the filamentous phage DNAs. The DNA was separated in 0.8% agarose gel and then transferred onto previously charged nylon membranes as described in Sambrook et al. (1989). The hybridization was carried out using DIG labeled HindIII fragment obtained from the pH17 DNA as a probe and under standard Southern blotting conditions, final washing was carried out for 30 min in 2 × SSC buffer (all manipulations were performed at room temperature). The DIG labeling, hybridization and detection was carried out according to the manufacturer’s instructions (La Roche). Presence of the prophage sequences was determined by PCR reactions, that were carried out using Pfu polymerase (MBI Fermentas) and according to the manufacturer’s instructions. The reaction mixture (25 µl) contained template chromosomal DNA or “crude phage preparations” DNA; reaction buffer; dATP, dGTP, dCTP and dTTP (0.2 mM each); forward and reverse primers (0.2 µM each) and 0.5 units of polymerase. Sequences of the primers used to amplify the fragment of prophage DNA were: 5'-AACTTTCGACGTGTCCGGGAACTTATGGAGACGGA-3' and 5'-AAGGCGCGGTGCTGATTGTTGGCGAAGCGCACTA-3'. PCR conditions were as follows: 5 min at 95, followed by 2 cycles of 30 sec at 94°C, 45 sec at 48°C, and 2 min at 72°C, followed by 29 cycles of 30 sec at 95°C, 45 sec at 62°C, and 3 min at 72°C (with final elongation – 10 min at 72°C). The PCR product (1050 bp) corresponds to the chromosomal sites localized within the NGO1138 sequence which encodes the homologue of CTXM Zot protein. The specificity of this product was confirmed by sequencing. The primers used for PCR amplification were obtained from IBB (Warsaw, Poland). All routine cloning procedures was carried out in accordance to protocols described by Sambrook et al. (1989). Enzymes and chemicals. Restriction enzymes were purchased from MBI Fermentas and New England Biolabs. T4 DNA ligase, Pfu DNA polymerase and DNA size markers were purchased from MBI Fermentas. Kits for the DNA clean-up and plasmid DNA isolation were purchased from A&A Biotechnology, Gdansk, Poland. All the chemicals used were reagent grade or better and they were obtained from Sigma and ICN, unless otherwise noted. Computer analysis. DNA and protein sequences were compared with the GenBank and SWISS-PROT databases on the BLAST server, hosted by the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/blast). The N. gonorrhoeae strain FA1090 genomic sequence was obtained from the University of Oklahoma’s Advanced Center for Genome Technology (http:// www.genome.ou.edu/gono.html). Both the N. meningitidis strain Z2491 (serogroup A) (http://www.sanger.ac.uk/Projects/ N_meningitidis) and N. meningitidis strain FAM 18 (http://www.sanger.ac.uk/Projects/N_meningitidis) sequences were obtained from Sanger Institute. Other comparisons were performed using the BLAST tools at the NCBI web site (http://www.ncbi.nlm.nih.gov/ BLAST) or http://www.cbs.dtu.dk/services/TMHMM. Phage techniques. The secreted form of phage and its DNA was prepared by standard phage preparation techniques (Sambrook et al., 1989). Bacteria were sedimented by centrifugation from 200 ml of culture exponentially growing in GC medium, and a total DNA extracted from the cells was dissolved in 200 µl of TE (10 mM Tris-HCl, 1 mM EDTA-Na pH, 8.0) buffer. After filtration through 0.45 µm filters, the supernatant was treated with DNase I and RNaseA (25 µg/ml each) for 3 h at 20°C. Particles were precipitated by addition of NaCl to final concentration 1 M and PEG 8000 to a final concentration of 10%. The mixture was incubated at 4°C overnight and the phage particles were precipitated by centrifugation at 12 000 × g for 30 min. DNA from the presumptive phage particles was extracted with phenol, and the precipitated material was redisolved in 200 µl of TE buffer.

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Filamentous bacteriophage genomes in Neisseria gonorrhoeae

Results Identification of the prophage sequences in the genome of N. gonorrhoeae FA1090. Annotations of the complete genome sequence of N. gonorrhoeae FA1090 (Accession number: AE004969) revealed four probable prophage islands, designated as NgoM6 to NgoM9, containing several ORFs with striking amino acid similarities to known functional phage proteins (Table I). The nucleotide sequences as well as amino acid sequences of NgoM6–9 show high similarity to the filamentous ssDNA phages, especially to those able to integrate into the chromosome of the host (Davis and Waldor, 2003; Bille et al., 2005). Table I Localization of the filamentous bacteriophages integrated into Neisseria gonorrhoeae FA1090 genome Phage

Nucleotide sequence coordinatesa

Length of the DNA sequence (bp)

NgoM6

1080185 – 1088060

NgoM7

1215967 – 1222383

NgoM8

1103017 – 1109444

NgoM9

1599378 – 1606537

a Coordinates

ORF annotations (Acc. No AE004969)

Number of ORFs

8 235

NGO1137 – NGO1146

13

7 416

NGO1262 – NGO1270

12

6 427

NGO1164 – NGO1170

8

6 754

NGO1641 – NGO1648

9

are relative to the DNA sequence contained at GenBank Acc. No AE004969

Nucleotide sequence and genomic organization. The DNA and amino acid sequences of both NgoM6 and NgoM7 show similar genetic organization and high homology (50–90% of identity) to each other. NgoM8 lacks an equivalent of ORF1 (NGO 1146), ORF2 and ORF3 (present in NgoM6 and NgoM7) show lower level of homology (Fig. 1A). However, the ORF2 and ORF3 of NgoM6 lack the homology (both on DNA and amino acid levels) to their equivalents present in NgoM7 and NgoM8 (Fig. 1B). The NgoM9 contains only part of the genomes of the NgoM6–8 prophages. Several functionally same genes of different origin are duplicated, with no homology to their counterparts in phages NgoM6, NgoM7 and NgoM9. (Fig. 1B). The genetic organization of the NgoM6 suggests that it can serve as a prototype of filamentous phage of N. gonorrhoeae. The nucleotide sequence of NgoM6 consists of 8240 nucleotides and has mean G+C content of 46.0%, which is lower than G + C content of the host N. gonorrhoeae (54%). In the chromosome of N. gonorrhoeae the 5’ end of the NgoM6 prophage is flanked by the gene encoding glucosyl transferase (ORF1136) while the 3’ is flanked by the ORF1149 encoding O-serin-succinylohomosulfohydrolase. The NgoM7, NgoM8 and NgoM9 prophage sequences are flanked on 5’ end by the 136 bp long sequence that is also present in NgoM6 prophage DNA (Fig. 1A) but in different localization. The ORF11 (NGO1137) of NgoM6 encoding integrase, has the opposite orientation than the corresponding genes in other prophages and the 136 bp sequence is present on the other flank of this gene. On the 3’ end of the NgoM9 sequence the ORF11 (NGO1648) encoding transposase is followed by the ORF1649 encoding the putative dsDNA phage protein. The 3’ ends of NgoM7 and NgoM8 sequences are flanked by NGO1273 encoding tetraadenosinophosphate and NGO1173 encoding VSR protein, respectively. We were unable to determine any homology between the 3’ ends of all four prophage sequences and between 3’ and 5’ ends of each sequence. The CTXM and VGJM phages, prototypes of the lysogenic filamentous phages, contain three modules essential for phage production (Davis et al., 2000; Campos et al., 2003). The five classical core genes of CTXM encode proteins needed for virion structure and assembly, the replication module encodes RstA, RstR and RstB proteins; RstA and RstR mediate replication of phage DNA and regulation of rstA expression respectively (Kimsey and Waldor, 2004; Mekalanos, 1983). The role (if there is any) of RstB in CTXM virion production has not yet been determined; it was only shown that RstB contributes to phage integration (Waldor and Mekalanos, 1996). The third module, the assembly one, encodes several genes needed for phage maturation. Their corresponding genes in VGJM are represented by ORF81, ORF44, ORF29, ORF493, ORF80, ORF384 (virion structure and assembly); ORF359, ORF112, ORF58, ORF67, ORF136, ORF 154 (replication and expression). The CTXM genome lacks an integrase; instead, its integration depends on the chromosome encoded tyrosine recombinases XerC and XerD (McLeod and Waldor, 2004). The NgoM6 – NgoM9 prophage nucleotide sequences posses the ORFs that encode putative transposase. The gene shares homology with Pfam 02371 transposase 9 and Pfam 01548 transposase 20 families (Table II). N. gonorrhoeae contains several copies of this gene that may encode a site-specific recombinase and play

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Fig. 1. Comparison of the organization of the NgoM6-NgoM9 prophages with that of Vibrio cholerae bacteriophages CTXM and VGJM N. meningiditis MDAZ2491. (A) Linear maps of CTXM, VGJM, MDAZ2491 and NgoM6-9 phages, aligned by using the first base of the replication initiator gene of CTXM as an orientation point. Arrows oriented in the direction of transcription represent ORFs or genes. Open arrows represent the replication module, solid arrows represent the structural module, arrows with left cross-hatching represent an assembly module, an arrow with black dots represents the integrase and the arrows with vertical lines represent the ORFs encoding the repressors. The arrow with right cross-hatching represents the ORFs with unknown function. The ORFs of VGJM, CTXM genes and MDA Z2491 are designated according to the published papers (Campos et al., 2003; Bille et al., 2005). The lengths of the products encoded (in amino acids) are indicated below each arrow. The solid boxes represent 136 bp sequence present on one flank of ORFs encoding integrase gene common for all four NgoM phages. (B) Similarity between the DNA nucleotide sequences of filamentous phages of N. gonorrhoeae. The open boxes represent the region of high homology (99% of identity), the solid boxes the regions of lack of homology (below 5% of identity).

a role in the integration and the excision of the ssDNA phage genome within the chromosome of N. gonorrhoeae. Its homologues, present in the genome of filamentous prophage in N. meningitidis, are responsible for their integration and excision from the chromosome (Bille et al., 2005). A computer-aided homology search was performed for the ORFs presumed to encode functional viral genes (designation presented in Table II). In NgoM6 it showed the same type of genetic organization as in other filamentous phages, like Vibrio phages CTXM, VGJM or N. meningitidis MDA (ssDNA prophage) sequence (Campos et al., 2003; Bille et al., 2005). The common property of these phages is the ability to integrate into the host chromosome. The function of these genes was also deduced from the homology to genes of other filamentous phages and by comparison of the presence of intermembrane motives (TM) (Table III). Within the putative replication module, (Fig. 1A; note that the maps of the NgoM6-9 on this figure are presented in the opposite orientation than in the chromosome of N. gonorrhoeae in order to aid in comparison with other phages) we identified ORF1 (NGO1146). The peptide encoded by ORF1 is homologous to the RstA protein of CTXM (26% identity and 44% positives over 234 residues), which is necessary for phage replication (Waldor et al., 1997). ORF1 is also similar in terms of size and position to genes of other filamentous phages, which were mapped at the same relative position as the gII gene in Ff phages (data not shown). ORF1 could be homologous to PII protein of filamentous phages necessary for rolling-circle replication of phage genomes (Model and Russel, 1988). There is no homologue of ORF NGO1 in NgoM8 or NgoM9. We were unable to identify the peptide that would be homologous to rstB or gene encoding ssDNA

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Filamentous bacteriophage genomes in Neisseria gonorrhoeae Table II Properties of the NgoM6 ORF’s amino acid sequences Protein Length (aa)

Gene

ORF1, NGO 1146

423

pMU1_p3 (Eikenella corrodens) plasmid replication ini- tiation factor (YP_245390), 1 × 10–103; RstA1 protein (pfam02486), 8 × 10–38; phage [Vibrio cholerae Prophage] replication protein RstA (NP_231106.1), 1 × 10–11 (COG2946), 4×10–143

Na N

ORF2, NGO 1145

103

NSb

NS

ORF3, NGO 1144

67

NS

NS

N

ORF4, NGO 1143

78

NS

NS

Y (2)

ORF5, NGO 1142

98

NS

NS

ORF6, NGO 1141

110

NS

NS

ORF7, NGO 1140

506

NS

neisserial TspB virulencefactor

Y (1)

ORF8, NGO 1139

107

NS

NS

Y (2)

ORF9, NGO 1138

361

pUM1_p7 (YP_245394.1), 2×10 –32

zonular occludens toxin (Zot) [Eikenella corrodens] (pfam05707), 7 × 10 –39

Y (1)

ORF10

107

NS

NS

320

transposase [Escherichia coli IS621], (BAC768887.1), 3 × 10 –37

transposase (COG3547), 1 × 10–17

ORF11, NGO 1137

a

Presence and number of transmembrane domains (TMHMM)

Conserved domains (BlastP)

Protein homologies (BlastP)

ORF12

23

NS

NS

ORF13

86

NS

NS

N, lack of TM domains;

b Y,

presence of TM domains;

c NS,

c

N

Y (1)

no significant homologies were found

Table III Comparison of the predicted transmembrane helix motives in some of the proteins encoded by filamentous phages Phage/protein (aa) CTXM, Zot (399)

Number of predicted TMHs 0

Outside

TMhelix 1

Inside

TMhelix2

Outside

1 – 399

–

–

–

–

VgJM, ORF384 (Zot) (384)

1

1– 213

214 – 233

234 – 384

–

–

NgoM9, ORF9* (395)

1

225–395

205–224

1–204

–

–

NgoM6, ORF 9361

0

1 – 361

–

–

–

–

CTXM, gIII

1

1 – 374

375 – 394

395 – 395

1

30 – 493

7 – 29

1–6

–

–

–

– –

CTX

(395)

VgJM, ORF493 (493) NgoM6, ORF7 (507)

1

1 – 483

484 – 506

507 – 507

CTXM, gIIICTX (395)

1

1 – 374

375 – 394

395 – 395

VgJM, ORF493 (493)

1

30 – 493

7 – 29

1– 6

–

NgoM6, ORF7 (507)

1

1 – 483

484 – 506

507 – 507

–

–

CTXM, Ace VgJM, ORF8

2

1 – 17

15 – 37

38 – 64

65 – 87

88 – 96

NgoM6, ORF8 (80)

1

1 – 43

44 – 60

67 – 80

–

–

NgoM9, ORF8 (98)

2

1 – 17

18 – 40

41 – 60

61 – 83

84 – 98

* NgoM9 encodes two copies of ORF9, which differ slightly in number of amino acid residues.

binding protein within the putative replication module. The ORF2 and ORF3 are located in the position of the rstB and psh genes of CTXM phage that participate in DNA replication (Waldor et al., 1997). These two genes do not show the presence of transmembrane domains (data not shown), this property is common for the structural and assembly proteins of the filamentous phages. Thus, it is possible that they are in fact engaged in the replication process. In addition, the ORF6 that does not have TM domain, is flanked by the ORF5 and ORF12 with such domains. Instead, this ORF shows the presence of Zinc-finger motive what could suggest its participation in replication or transcription control processes (Klug and Schwabe, 1995).

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Within the putative structural module of NgoM6, we identified the ORFs 4, 5, 12, 7 and 8; sizes and positions of these sequences are similar to those of genes encoding capsid structural proteins of Ff phages, CTXM, VGJM and other previously described filamentous phages (Campos et al., 2003). The ORF4 and 8 could play the same role as the cep and ace genes of CTXM phage (Canchaya et al., 2004). These ORFs are predicted to encode proteins with transmembrane domains. The results support the idea that they form the module of structural genes (Fig. 1A) which encode the capsid proteins of NgoM6. ORF7 (NGO1140) is homologous to N. meningitidis and N. gonorrhoeae TspB protein belonging to pfm05616 family. This family consists of several TspB virulence factor proteins. This ORF is located at the same relative position as gIII of the Ff phages and gIIICTX of CTXM phage and its homologue (ORF384) in VGJM phage. Its size is comparable to the sizes of these genes that encode pIII, a minor capsid protein that recognizes and interacts with the receptors and co receptors of these phages (Armstrong et al., 1981; Heilpern and Waldor, 2003; Lubkowski et al., 1999). Therefore, ORF7 (NGO1140) could be a homologue of gIII in CTXM. The ORF7 (NGO1140) shows very high homology (92% identity) to homologous genes in NgoM7– 9 phages. ORF8 (NGO 1139) shows some degree of identity (about 25% identity, 50% positives over 70 residues) to other ssDNA phages of Acetinobacter, Nitrosomonas europea, Xylella fastidiosa and Pseudomonas phage Pf3). Within the third putative module of NgoM6, the assembly module, we found ORF9 (NGO1138) belonging to pfam05707 of proteins, whose product is homologous to the pI protein of Pseudomonas phage Pf3 (27% identity over 344 residues) and to the Zot protein of CTXM (19% identity and 39% positives over 209 residues) as well to ORF6 of MDA Z2491. Based on its size and position, ORF9 also corresponds to the gI gene, which encodes pI in Ff phages, needed for assembly and secretion of viral particles (Marvin, 1998; Russel, 1995; Russel et al., 1997; Campos et al., 2003; Bille et al., 2005).

Fig. 2. Detection of the presence of filamentous DNA sequences in different strains of Neisseria. (A) Dot blot hybridization of the NgoM6 probe with a total DNA isolated from different Neisseria strains. 1; N. gonorrhoeae 220, 2; N. gonorrhoeae FA1090, 3; N. gonorrhoeae WR302-1, 4; N. gonorrhoeae WR302, 5; N. gonorrhoeae MS11, 6; N. gonorrhoae MS11 (different isolate), 7; N. gonorrhoeae Pgh3-2, 8.; N. lactamica 5841, 9; N. meningitidis 13113, 10; N. meningitidis 53414, 11; N. gonorrhoeae FA5100, 12; N. gonorrhoeae 220 RifRNalR, 13; N. gonorrhoeae WR302 RifRNalR, 14; N. gonorrhoeae 220 RifRNalR (different isolate), 15; N. lactamica 5841, 16; N. gonorrhoeae 1291. (B) Detection of the presence of phage sequences by PCR methods. 1; N. gonorrhoeae 1291, 2; N. gonorrhoeae 220, 3; N. gonorrhoeae FA1090, 4; N. gonorrhoeae WR302-1, 5; N. gonorrhoeae WR302, 6; N. lactamica 5841, 7; N. meningitidis 13113, 8; positive control, pH17, 9; N. gonorrhoeae MS11.

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Fig. 3. Detection of the extrachromosomal replicative and extracellular forms of phage DNA. N. gonorrhoeae total DNA separated in 0.8% agarose gel (A) and hybridized (B) with the NgoM6 probe. 1; N. gonorrhoeae 220, 2; N. gonorrhoeae FA1090, 3; N. gonorrhoeae WR302-1, 4; N. gonorrhoeae WR302. M; DNA marker, (C). The prophage DNA is secreted in the nuclease-resistant form. Treatment with DNaseI had no significant effect on the material amplifiable by the NgoM6 primers. PCR amplification was performed on DNA from phage preparation. 1; phage DNA (N. gonorrhoeae FA1090) treated with S1 nuclease, 2; phage DNA (N. gonorrhoeae FA1090) treated with DNaseI, 3; phage DNA (N. gonorrhoeae FA1090) treated with AciI, 4; phage DNA (N. gonorrhoeae FA1090).

The Zot protein encoded by ORF9 shows only partial homology to the Zot proteins of NgoM7 and 8 (37% identity, 55% positive over 350 residues) and also with both Zot proteins of NgoM9 (395 residues). What is more important, all of these Zot proteins show the homology to phage CTXM encoded Zot protein only at N-terminal end and not at the C-terminal ends. However, only NgoM9-encoded Zot protein do possess TM motive (Table III). Most of the structural and assembly proteins of NgoM6 show the overall structures of TM domains similar to the homologous protein of other filamentous phage (data not presented). However, in some cases they show important differences. The proteins ORF7 (NgoM6), gIII (f1), gIIICTX and ORF493 (VGJM) all have one TM domain but in the ORF493 it is located in N-terminal end of protein while in all others at the opposite end. The ORF 8 (NgoM6), ace (CTXM) and ORF80 (VGJM) proteins have two TM domains while their homolog in f1 phage (gVI) three domains (Table III). The proposed assembly and secretion module of NgoM6 do not have a homologue of the gIV of the Ff phages, similarly to VGJM (Campos et al., 2003) and MDAZ2491 (Morgan et al., 2002 ). Function of this gene in the development of CTXM , which is critical for phage morphogenesis (Opalka et al., 2003) is substituted by cell function of the gene EpsD (Campos et al., 2003). We were unable to identify the fourth, regulatory module. The ORF13 which is located in the position corresponding for the repressor gene encoded by CTXM phage but does no show homology to any of the known protein. Like the rstR of CTXM the ORF13 is transcribed in opposite direction of transcription of the rest of the phage genes. RstR protein is a repressor that regulates transcription of the initiator replication protein, RstA, and in turn regulates the expression of all phage genes (Davis et al., 2002; Kimssey and Waldor, 2004). We do not know the function of the ORF10, which is present in the genome of all Ngo ssDNA phages and in VGJM, but was not identified in MDAZ2491 (Fig. 1A). Presence of the prophage sequences in different N. gonorrhoeae strains. In N. meningitidis the phages homologous to NgoM6 are present predominantly in the hypervirulent isolates (Bille et. al., 2005). On the other hand, the Mu-like prophage sequences are present in all N. meningitidis strains (Masignani et al., 2001). The N. gonorrhoeae is the causative agent of the sexually transmitted disease called gonorrhoea. In the male, this is typically associated with a purulent discharge from the urethra. However, in women, infection of the cervix is often asymptomatic while the spread of the gonococci up the urinary tract or invasion across the epithelial layers can cause additional complications such as pelvic inflammatory diseases (PID) and disseminated gonococcal infection (DGI) (Jordan et al., 2005). There are experimental strains of N. gonorrhoeae commonly used in the laboratory, which were isolated either from the patient with DGI (FA1090) or with UG (F62 and MS11) (Jordan et al., 2005).

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The dot blot hybridization experiment (Fig. 2), where the 5.5 kb fragment of NgoM6 sequence was used as a probe, indicated that probably all strains of N. gonorrhoeae, except some mutants, contain the phage sequence, either the whole or its fragment. This sequence was not present in non-pathogenic strain of Neisseria lactamica. The presence of the NgoM6 was also confirmed by the formation of the PCR products specific for the zot gene of NgoM6 (Fig. 2B). Presence of the filamentous phages in N. gonorrhoeae cultures. The replication of the CTXM family of phages proceeds without the excision of the integrated genome in the form of the circular intracellular form of DNA (Waldor et al., 1997). The presence of such phage DNA form inside the lysogenic cells may serve as an indicator of the presence of the biologically active phage. The hybridization studies carried out with the total DNA isolated from the N. gonorrhoeae FA1090 cells failed however to show the presence of the prophage sequences in the form of free DNA molecules inside the cells (Fig. 3B). The failure to detect the presence of free phage DNA inside the cells could be due to limited sensitivity of our hybridization method or due to very low level of the phage DNA replication. In N. meningitidis, the genomic DNA of the filamentous phages was also detected as a single positive strand present in “crude phage preparations” (Bille et al., 2005), indicating the secretion of these phages from the cells and the functionality of the phage genome. In similar experiments, we have also shown the presence of free ssDNA specific for NgoM6 (Fig. 3C). This result indicates that also in N. gonorrhoeae FA1090, the prophage sequences of the filamentous phages can replicate and produce the progeny. Our attempts to obtain propagation of these phages on different strains of Neisseria have failed so far. Discussion Recent discovery of the biologically active filamentous phages in N. menigiditis and the results described in this paper indicate the presence of a new group of such phages. Comparison of the deduced sequence of NgoM6 with the published genetic maps of variuos filamentous phages integrated into the chromosome of N. meningitidis reveals their great similarity to the CTXM and other phages of Vibrio (Davis et al., 2000; Campos et al., 2003). Both groups of phages have a modular genome organization in which three modules can be distinguished: the replication module, the structural module and the assembly module. We were not able to identify the presence of the gene encoding phage repressor. Such repressor gene is present in all other filamentous phages able to integrate into the chromosome of the host like CTXM, VGJM, Vf33 or Cf1c (Brüssow et al., 2004) but absent in non-lysogenic phages. Its presence is required for maintenance of the lysogenic state and regulation of the gene expression (Kimsey and Waldor, 2004). The NgoM6 has a distinctive region of about 800 nucleotides that is not significantly homologous to any entry in databases and is absent in N. meningitidis filamentous phage genomes (Bille et al., 2005). Such region is present in VGJM phage genome; it encodes the ORF of unknown function. This region can be homologous to intergenic (IG) region present in non lysogenic phages containing signals for replication, packaging of DNA and the strong transcription terminator (Webster, 1994). Integration of CTXM DNA and probably several other lysogenic phages (VGJM, Cf1c, Cf16-v1) depends on the activity of the host encoded tyrosine recombinanses XerCD (Huber and Waldor, 2002; Campos et al., 2003) and the presence of the attP and attB sites within the phage genome and within the chromosome of bacteria, respectively. While one XerCD binding site in attP spans the short core region, the other site is approximately 80 bp away. Although integration occurs at the core XerCD binding site in attP, the second site is required for CTXM integration (McLeod and Waldor, 2004). In the chromosome of N. meningitidis the sequences corresponding to ssDNA prophages are inserted into 20 bp inverted repeats (dRS3) and are flanked by the sequence ATTCCCNC and GNGGGGAAT. These sequences can represent analogous integration core region as characterized for CTXM phage. No such sequences were found on the flanks of N. gonorrhoeae ssDNA prophages. However, the same 136 bp DNA sequence is present at the end of the ORFs encoding the phage integrase in all filamentous N. gonorrhoeae phages. This sequence can serve as a region responsible for binding of the integrase and the site of recombination. Like in N. meningitidis there are several ssDNA phage islands within the genome of N. gonorrhoeae strain FA1090. The same explanation given by Bille et al. (2005) can be valid for this phenomenon. These are as follows: (i) genomic rearrangements that result in duplication, (ii) horizontal transfer by transformation and insertion through recombination into the same type of repeat sequence (iii) better induction of immunity to superinfection. The presence of several copies of different types of prophage sequences acquired during evolution and coding for several restriction-modification systems gave them a total protection against

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phage infection. This can be an evolutional method for shifting the balance between phages and bacteria for favors of the host. Is there any biological role of the filamentous prophage sequences in N. gonorrhoae? The simpliest explanation would be, as in case of V. cholerae strains, that phage proteins increases the pathogenic potential of the bacteria. In N. meningitidis, the homologous filamentous prophages are mainly integrated into chromosomes of strains that show increased invasiveness (Bille et al., 2005). In N. gonorrhoae, the strains that seem not to possess such phages are not recognized as less pathogenic than those that have these sequences. There is one similarity in sequence between CTXM and the genomes of the N. meningitidis and N. gonorrhoeae filamentous phages; it lies within the genes encoding the Zot and Zot homologue proteins. The details analysis show that only the N-terminal end (from residue 9 to 196) of ORF9 (NGO1138) exhibits the homology with N-terminal end of Zot of CTXM (residues 9 to 205). The enterotoxin function of Zot has been assigned to its C-terminal region (Di Perro et al., 2001) that has no homology to the C-terminal end of ORF9 (NGO1138). This fact would argue with enterotoxic properties of the ORF9. The Zot protein encoded by ORF9 (NGO1138) (361 residues) is smaller then Zot of CTXM (399 residues) what can be responsible for the lack of homology between these two proteins. However, the C-terminal region of the ORF9 (NGO1647) in NgoM9 phage that has the larger number of amino acids (399) does not share homology with the C-terminal end of CTXM Zot protein. It does not preclude the possibility that the Zot encoded by ORF9 (NGO1138) has other toxic properties different than Zot of classical CTXM. This suggestion is supported by the observation that the Zot protein (361 aa) encoded by the filamentous NgoM phages, similarly to the Zot of CTXM, and contrary to the Zot of VgJM does not show the presence of TM motive (Table III), what could suggest that their structures are similar. The low homology between the Zot proteins encoded by NgoM6–9 phages suggests that their acquisition by N. gonorrhoeae was in fact several independent procesess that happened several times in the evolution of the bacteria. The two essential phage genes of CTXM phage, cep and ace that have their homologs in NgoM6 (ORFs 4 and 8), encode a core pilin. The products of these two genes do not function as toxins during V. cholerae infection (Canchaya et al., 2004) what may also be true for N. gonorrhoeae and its filamentous phages. To our knowledge it is the first description of the filamentous bacteriophage genomes integrated into N. gonorrhoeae but their role for N. gonorrhoeae pathogenicity is still not known. Acknowledgements. The Faculty of Biology Grant No 501/64-1005/4 supported this work.

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Polish Journal of Microbiology 2006, Vol. 55, No 4, 261– 270

Replication System of Plasmid pMTH4 of Paracoccus methylutens DM12 Contains an Enhancer MICHA£ SZYMANIK1, RENATA WELC-FALÊCIAK2, DARIUSZ BARTOSIK and MIROS£AWA W£ODARCZYK*

Department of Bacterial Genetics, Institute of Microbiology, Warsaw University, Warsaw, Poland Received 19 September 2006, revised 17 October 2006, accepted 23 October 2006 Abstract The replication system of plasmid pMTH4 (22 kb) of dichloromethane-degrading Paracoccus methylutens DM12 (Alphaproteobacteria) has been cloned within a mini-replicon pMTH100 (4.7 kb) and preliminarily characterized. Functional analysis, performed with a series of mutated plasmid mini-derivatives, showed that the replicator region consists of three elements: (i) gene repA coding for a replication initiation protein RepA, (ii) origin of replication (oriV), placed in the promoter region of repA and containing a set of imperfect directly repeated sequences (iterons) together with putative DnaA and IHF-binding DNA sequences as well as (iii) an enhancer (0.65 kb) upstream of oriV. We showed that the enhancer was not crucial for plasmid replication, however, it was necessary to assure the proper plasmid copy number. Additionally its presence has increased the strength of a determinant of incompatibility (located within the oriV region) as well as the level of transcription carried from the repA promoter. The enhancer region was shown not to encode any proteins or promoter sequences. We speculate that this region might constitute a site of binding of plasmid or host-encoded proteins that are able to interact with the origin, which positively regulates the initiation of replication. K e y w o r d s : Paracoccus methylutens, enhancer, iteron, plasmid replication

Introduction The majority of plasmids of Gram-negative bacteria replicate according to the so called theta model. Among them five classes have been distinguished (Bruand et al., 1993; Meijer et al., 1995), including the iteron-containing plasmids (class A). Replication initiation of class A plasmids depends on a plasmid-encoded initiation replication protein (Rep). The DNA region comprising the origin of replication of these plasmids contains several direct nucleotide repeats (iterons) recognized by Rep protein as well as host-encoded DnaA and IHF (and occasionally Fis, Ici, SeqA) binding sites (reviewed in Kruger et al., 2004). The well known representatives of the class A plasmids are: F, P1, R6K, pSC101, RK2 or pPS10 (Chattoraj, 2000). Saturation of the iteron sequences with monomeric forms of Rep proteins as well as binding of the host proteins results in a local destabilization of DNA sequence at the AT rich region of replication origin, which results in open complex formation (Messer et al., 2001). The frequency of formation of the complex is an important factor that regulates plasmid copy number. In some cases (R6K, pSC101 and pCD3.4) it has been shown that the class A replicator regions encode additional regulatory elements involved in regulation of initiation of replication (van Belkum and Stiles, 2006; Kelley et al., 1992; Miller and Cohen, 1993; Sugiura et al., 1993). In all cases the enhancers do not encode for additional proteins engaged directly in the regulatory process but encode sequences recognized by plasmid- or host-encoded proteins that interact with the oriV. Paracoccus methylutens DM12, the natural host of plasmid pMTH4, is a Gram-negative, oxidative, facultatively methylotrophic bacterium, able to utilize many compounds commonly considered as a highly * Corresponding author: M. W³odarczyk, Department of Bacterial Genetics, Institute of Microbiology, Warsaw University, Miecznikowa 1, 02-096 Warsaw, Poland; e-mail: [email protected] 1 Current address: Adamed Sp z o.o., Pieñków 149, 05-152 Czosnów, Poland 2 Current address: Department of Parasitology, Institute of Zoology, Warsaw University, Miecznikowa 1, 02-096 Warsaw, Poland

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toxic (eg. dichloromethane, dichloroamine, methanol, methylamine, and formate (Doronina et al., 1998). We have found that the strain harbors 4 native plasmids, and one of them – pMTH4 (not detected in our preliminary search; Baj et al., 2000) has become the object of our interest. Plasmid pMTH4 is ~22 kb in size. A mini-replicon of this plasmid has been constructed (pMTH100 – carries a 4.7 kb HindIII restriction fragment of pMTH4 and a kanamycin resistance cassette) and shown to be stably maintained in different representatives of the Alphaproteobacteria. In silico analysis of the nucleotide sequence of pMTH100 allowed us to distinguish two potential structural modules: (1) REP – coding for replication system, (2) STA – carrying stabilizing module as well as a truncated insertion sequence (IS) (Szymanik et al., 2004). The REP module carries (i) a repA gene whose putative product shows similarity to the replication initiation proteins of several iteron plasmids and (ii) the putative origin of replication (oriV) located upstream of repA gene, containing a set of directly repeated sequences (iterons) as well as presumptive DnaA and IHF binding sites (Fig. 1). The iterons R1-R3 (18 bp) are identical, while the other two R4 (17 bp in length) and R5 (19 bp) contain some differences at the nucleotide sequence. The distances between R1 and R2 as well as between R2 and R3 are identical (4 bp), while the distance between R4 and R5 is as long as 31 bp (Szymanik et al., 2004). The STA module of pMTH100 (~1000 bp) is located upstream of REP and carries two putative genes staA and staB (Fig. 1A) which represent a proteic toxin-antitoxin system, with staA gene encoding antitoxin and staB toxin products, respectively (Szymanik, 2006). The present paper deals with the functional characteristics of pMTH4 replication system, which led to the identification of an enhancer region that seems to positively influence the process of initiation of plasmid replication. Experimental Materials and Methods Bacterial strains, plasmids, media and growth conditions. Bacterial strains and plasmids used (but not constructed) in this work are listed in Table I. Plasmids constructed in this work are described under Results and in the appropriate figures. Bacteria were grown in Luria-Bertani (LB) medium (Sambrook and Russell, 2001) at 30°C (P. pantotrophus KL100) or 37°C (E. coli strains). When necessary, the medium was supplemented with antibiotics: kanamycin (Km) – 50 µg/ml, rifampicin (Rif) – 50 µg/ml, tetracycline (Tc) – 0.1 µg/ml for P. pantotrophus KL100 with derivatives of pABW2 (minimal selective concentration), 0.5 µg/ml for P. pantotrophus KL100 with derivatives of pRK415, and 20 µg/ml for E. coli.

Table I Bacterial strains and plasmids used (but not constructed) in this study Relevant characteristics

Source or reference

Bacterial strains E. coli DH5"

F– M80d lacZ )M15 (lacZYA-orgF)U169 deoR recA1 endA1 hsdR17 phoA Hanahan, 1983 supE44 8– thi-1 gyrA96 relA1 (host strain for helper plasmid pRK2013)

E. coli TG1

supE44 hsd)5 thi )(lac-proAB) F’ [traD36 proAB+ lacIq lacZ)M15] (host strain for recombinant plasmids)

Sambrook and Russell, 2001

P. pantotrophus KL100

Rifr derivative of P. pantotrophus DSM 11073 deprived of its natural plasmid pKLW1; contains pKLW2 (>400 kb)

Bartosik et al., 2002

pABW2

Tcr; mobilizable cloning vector based on pBGS18, oriT of RK2

Bartosik et al., 1997

pBGS18

Km ; cloning vector, ColE1 origin of replication

Spratt et al., 1986

pCM132

Km r; mobilizable promoter probe vector (promoter-less lacZ gene) carrying ColE1 and RK2 replicator regions, oriT of RK2

Marx and Lindstrom, 2001

pKRP11

Source of Kmr cassette

Reece and Phillips, 1995

pMTH100

Kmr; mini-replicon of pMTH4, composed of 4692-bp HindIII fragment of pMTH4 (coordinates 1 – 4692*) and Km r cassette

Szymanik et al., 2004

pRK415

Tcr; mobilizable broad-host-range cloning vector, oriT and origin of replication of RK2

Keen et al., 1988

pRK2013

Km r; helper plasmid carrying RK2 tra genes

Ditta et al., 1980

Plasmids

* – Acc. no. AY337272

r

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DNA manipulation. DNA techniques including plasmid purification, digestion with restriction enzymes, ligation and agarose gel electrophoresis were conducted according to standard procedures (Sambrook and Russel, 2001). All enzymes were purchased from Fermentas and used according to the manufacturer’s instructions. PCR amplification. For amplification of the origin fragment of pMTH100, containing all iterons, DNA box, and incomplete IHF binding site (ORILP fragment), the following pair of forward and reverse oligonucleotide primers was used: ORIL (5’GCGATATCACATTCTGTTCCAGAAGCGG-3’) and ORIP (5’-ATGGATCCCATCCGTTTGCCTGGCTGTT-3’). The introduced EcoRV (ORIL) and BamHI (ORIP) restriction sites are underlined. Amplification was performed in a Mastercycler Personal (Eppendorf) using the above primers, OptiTaq polymerase from EURX (with supplied buffer) and template DNA (pMTH100). The reaction mixture (25 µl) contained 10 ng template DNA, 2 mM MgCl2, 200 µM dNTPs, 50 pmol of each primer and 0.5 u OptiTaq polymerase. The amplification cycle was: 94°C for 5 min, followed by 35 cycles of 94°C for 1 min, 60°C for 0.5 min and 72°C for 1 min; the last cycle was followed by a final extension step of 10 min. Electroporation and transformation. Electroporation was carried out at 2500 V, 25 F and 400 S for P. pantotrophus KL100 in a gene pulser apparatus (Bio-Rad), based on modified Bio-Rad procedure (Wlodarczyk et al., 1994). Electrotransformants were selected on solidified LB medium supplemented with the appropriate antibiotic. Competent cells for transformation of E. coli TG1 were prepared and transformed as described by Kushner (1978). Triparental mating. For triparental mating three overnight cultures (cells harvested by centrifugation and washed to remove antibiotics) of the donor strain E. coli TG1 (carrying a mobilizable plasmid), the plasmid-less P. pantotrophus KL100 strain or P. pantotrophus KL100 strain carrying Kmr minireplicon of pMTH4 as the recipient, and E. coli DH5" strain carrying the helper plasmid pRK2013, were mixed at a ratio of 1:2:1. An aliquot of 100 µl of this mixture was spread on a plate of LB medium. After overnight incubation at 30°C, the bacteria were washed off the plate and suitable dilutions were plated on selective LB medium containing rifampicin (selective marker of the recipient strain) and kanamycin or rifampicin, kanamycin and tetracycline to select for transconjugants. Transcription and translation in vitro assay. The in vitro transcription and translation reaction was done with the E. coli Extraction System for Circular DNA kit (Promega) according to the manufacturer’s procedure. The obtained proteins were separated as described by Schagger and von Jagow (1987) and visualized according to the kit manufacturer’s procedure. $-galactosidase assay. $-galactosidase activity in P. pantotrophus KL100 cell extracts was measured by the conversion of o-nitrophenyl-$-D-galactopyranoside (ONPG) to nitrophenol as described by Miller (1972), with slightly modified bacterial cells lysis procedure – the addition to 100 µl of bacterial culture of 50 µl TE containing lysozyme in concentration 6 mg/ml and 2.5 µl of 0.5 M EDTA (pH 8.0). The samples were then incubated 5 min at room temperature. $-galactosidase activity assays were carried out in triplicate. Incompatibility testing. The incompatibility characteristics of two plasmids were examined by conjugational transfer (triparental mating) of tested recombinant Tcr plasmids based on vector pRK415 (containing PCR-amplified or restriction fragments of pMTH100) into the recipient strain (P. pantotrophus KL100) carrying pMTH105 (Kmr) mini-replicon. Transconjugants were selected for the incoming and resident plasmids. The plasmid patterns of the transconjugants were verified by screening 10 colonies using a rapid alkaline extraction procedure and agarose gel electrophoresis. The incompatibility behaviour of pMTH105 (which coexisted in trans with pRK415-based plasmids) was tested during growth for approximately 30 bacterial generations (72 h). Every 24 h cultures were diluted in fresh medium with tetracycline and without kanamycin. At this time intervals, samples were diluted and plated onto solid medium with tetracycline and without kanamycin. Two hundred colonies were tested for the presence of the Kmr marker by replica plating. The retention of pMTH105 was defined by determining the percentage of kanamycin-resistant colonies among 200 Tc r clones (containing a pRK415-based plasmid). Plasmid stability. The stability of plasmids (Kmr) during growth in non-selective conditions was tested as described previously (Bartosik et al., 2002). Briefly, every 24 h stationary phase cultures were diluted in fresh medium without antibiotic selection and cultivated for approximately 30 generations (72 h). At time intervals, samples were diluted and plated onto solid medium in the absence of selective antibiotic. Two hundred colonies were tested for the presence of the Km r marker by replica plating. The retention of plasmids, after approximately 30 generations, was determined from the percentage of kanamycin-resistant colonies among 200 clones.

Results Identification of the minimal replicon of pMTH4. In silico analysis of the nucleotide sequence of the mini-replicon pMTH100 allowed to distinguish a putative replication system (REP) composed of repA gene and origin of replication, placed upstream of repA, containing iteron-like repeated sequences. To prove the importance of the repA product in pMTH100 replication, mutational analysis was performed. To this end an E. coli-Paracoccus spp. shuttle vector pMTH102 was constructed, composed of (i) an E. coli-specific pBGS18 vector (Kmr; 3.7 kb; unable to replicate in paracocci) and (ii) the replicator region of pMTH4 (non-functional in E. coli) encoded by the 4.7 kb HindIII fragment of the mini-replicon pMTH100 (Fig. 1B). This plasmid (constructed in E. coli) could be easily introduced by electroporation into P. pantotrophus KL100 (a strain routinely used in our laboratory as a host for paracoccal plasmids). It appeared that a short not-in-frame insertion introduced within the XhoI restriction site of repA (4 bp) completely abolished the ability of the this mutated plasmid (pMTH102; Fig. 1B) to replicate in P. pantotrophus, as concluded from the lack of Kmr electroporants of the KL100 strain. This points to the crucial role of RepA protein in plasmid replication.

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Fig. 1. Genetic organization of mini-replicon pMTH100. The distinguished replication module (REP), stabilization module (STA), and truncated IS are appropriately marked. A. Nucleotide sequence of the iteron-containing region. The putative ribosome binding sequence (rbs), integration host factor (IHF) and DnaA (DnaA) binding boxes, –35 and –10 hexamers of the repA promoter and iterons (R1-R5) are marked. The deduced amino acid sequence of the N-terminal part of RepA is given below the sequence in a single-letter amino acid code in the first position of each codon. The numbers on the right refer to the deposited nucleotide sequence (Acc. no. AY337272). B. Localization of the minimal replicon of plasmid pMTH100. The restriction fragments shown in this figure were cloned into pBGS18 vector or ligated with the Kmr cassette derived from pKRP11. Not-in frame mutation introduced into repA is marked by an asterisk. * – plasmid replicating in P. pantotrophus KL100 (+) or unable to replicate (–).

In order to define the minimal replicon of pMTH100 we constructed diminished forms of the plasmid. To do this, two selected restriction fragments of pMTH100 (i) a 3 kb SacI-HindIII fragment and (ii) SmaI fragment (2 kb) (Fig. 1B) were ligated with a kanamycin resistance cassette derived from plasmid pKRP11, and the ligation mixtures were introduced by electroporation into P. pantotrophus strain KL100. In both cases electroporants were obtained, containing mini-derivatives pMTH104 and pMTH103, respectively. As shown in Fig 1B, the smallest mini-derivative (pMTH103) carried the repA gene, and the AT rich, iteroncontaining region (289 bp) as well as terminal part of a putative insertion sequence located downstream of repA (most probably non-essential for replication). Plasmid pMTH103 might be thus considered a minimal replicon of pMTH4. This confirmed the initial in silico predictions. Localization of the replication origin of pMTH4 and identification of an enhancer. To confirm experimentally that origin of replication of pMTH100 (and entire pMTH4) is located upstream of the repA several selected restriction fragments of pMTH100 (Fig. 2) containing the presumptive oriV were cloned into pABW2 vector (mobilizable Tcr derivative of pBGS18) and introduced by triparental mating into P. pantotrophus KL100 containing the pMTH100, with the hope that mini-replicon present in recipient would provide in trans RepA protein enabling the replication of the pABW2-derivatives. In control experiments we showed that plasmid pMTH232 (pABW2 carrying mutated version the 4.7 kb HindIII fragment of pMTH100, recovered from plasmid pMTH102; repA mutant) was capable of autonomous replication; the analyzed oriV can be thus trans activated by pMTH100. In contrast, empty vector pABW2 (carrying E. coli-specific replicator region) could not be introduced into the paracoccal hosts, which proved its inability to replicate in them. The results of the performed analysis are summarized in Fig. 2. Shortly, (i) no Tcr transconjugants of KL100 (pMTH100) were selected with pMTH239 (pABW2 containing amplified by PCR 202 bp region of pMTH100 carrying all the iterons, DnaA box, and incomplete IHF box) (Fig. 2), (ii) Tcr transconjugants were obtained upon conjugation of pMTH233, pMTH237 and pMTH235 (the plasmids contain the complete predicted origin, together with the adjacent sequences of pMTH100 of different length; Fig. 2). However, an autonomous form of the plasmid was visible after agar-

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Fig. 2. Defining of the origin of replication of pMTH100 by construction of two plasmid system [pABW2 (Tcr) + pMTH100 (Kmr)]. The restriction fragments of pMTH100, shown in this figure, were cloned into pABW2 vector and the resulting plasmids were introduced into P. pantotrophus KL100 (pMTH100). Insertion mutations introduced into repA and ORF5 are indicated with asterisks. Grey box indicates location of the iterons-containing region. * (+) – obtained Rifr Kmr Tcr transconjugants of P. pantotrophus KL100 (pMTH100) carrying the introduced plasmid; (–) – lack of transconjugants; ** (+) – the introduced plasmid visible after agarose gel DNA electrophoresis; (–) – plasmid DNA not visible after ethidium bromide staining.

ose gel DNA electrophoresis exclusively in case of pMTH235. We speculated that this might result from deviations in copy number of the three tested plasmids. Out of the tested plasmids pMTH235 contains the largest DNA fragment (0.65 kb) adjacent to the 5’end of the predicted iteron-containing origin (Fig. 2). The obtained results suggested that this region (called the enhancer) might be responsible for determination of the accurate pMTH100 copy number. Analysis of ORF5 carried by the enhancer. The enhancer region carries complete ORF5, which is one of the longest ORFs (642 bp) distinguished in pMTH100 sequence by in silico analysis (Szymanik et al., 2004). To find out whether it encodes a protein product that is expressed, the coupled in vitro E. coli-derived transcription-translation system was used. For this purpose a restriction fragment SacI-XhoI of pMTH100 (Fig. 2) containing the iteron region together with the adjacent enhancer was cloned, in opposite orientations, into pABW2 vector, to yield pMTH235 and pMTH236. Both plasmids together with pABW2 (as a control) were used for the analyses. Visualization of the obtained proteins showed that the enhancer region does not encode any protein products that are expressed in E. coli (data not shown). Since the transcription-translation system used by us is based on E. coli cellular components it was still probable that ORF5 might be preferentially expressed in the native paracoccal host. This might be, for instance, due to the presence of specific promoters that are functional in paracocci but not in E. coli. To test the importance of the putative ORF5 product in the activity of the enhancer, plasmid pMTH240 was constructed. This plasmid was nearly identical to the described above pMTH235 – it carried however a 2 bp insertion generated within the terminal part of ORF5 (Fig. 2). The mutated plasmid was introduced into P. pantotrophus KL100 containing pMTH100. In this case we observed trans activation of the cloned origin of pMTH240, as observed previously for pMTH235. Since no differences in copy number were detected (as judged from the level of DNA recovery of pMTH240 and pMTH235 from obtained KL100 transconjugants) it seems probable that ORF5 does not encode a protein that is crucial for enhancer activity. Influence of the enhancer on expression carried from repA promoter. Analysis of the nucleotide sequence of the REP region of pMTH100 allowed to distinguish a probable promoter of the repA gene (PrepA), located 70 bp upstream of the repA coding sequence (Fig. 1A). The predicted promoter was amplified by PCR within a 105 bp DNA fragment and inserted upstream of the promoter-less reporter lacZ gene in a broad host range promoter probe vector pCM132, to generate a transcriptional fusion. The resulting plasmid pMTH246 (Fig. 3A) was introduced into P. pantotrophus KL100 and $-galactosidase activity assay was used to examine promoter strength. The obtained results confirmed the presence of a weak promoter within the cloned fragment, as judged from the increased level of $-galactosidase activity, compared with the enzyme activity observed for the negative control – the empty vector pCM132 (Fig. 3B). To test whether the enhancer influence expression driven from the repA promoter, several selected DNA fragments of pMTH100 (all containing P repA with upstream sequences of different length – 0.1 kb, 0.4 kb,

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Fig. 3. Search for the promoters encoded within the REP module of pMTH100. A. Several selected restriction fragments were cloned into promoter probe vector pCM132. The orientation of the cloned fragments refer to the lacZ gene of pCM132 is shown by arrows. B. $-galactosidase activity in P. pantotrophus KL100 cells containing pCM132-derived plasmids listed in panel A.

0.85 kb and 1.6 kb) were cloned in a proper orientation into pCM132 vector, to yield pMTH245, pMTH247, pMTH248 and pMTH250, respectively (Fig. 3A). Analysis of the $-galactosidase activities determined in strain KL100 (Fig. 3B) allowed to conclude that the longer the enhancer region in the tested plasmids, the higher the enzyme activity (see pMTH245, pMTH247, pMTH248 in Fig. 3B). It also appeared that the presence of DNA sequences adjacent to the 5’ end of the enhancer region did not additionally increase $-galactosidase expression driven by PrepA in comparison to that specified by the enhancer itself (compare pMTH250 and pMTH248 in Fig. 3B). By construction of an additional plasmid pMTH251 (pCM132 carrying 1.15 kb SmaI fragment of pMTH100, lacking PrepA but containing the overall enhancer) we showed that enhancer itself does not encode internal promoters, as judged from the lack of the $-galactosidase activity (Fig. 3B). Additional analysis showed that the analyzed replicator region (including enhancer) does not carry promoter sequences placed in the opposite orientation towards P repA. The presence of such promoters might suggest formation of antisense RNA or small regulatory RNAs involved in regulation of the initiation of pMTH100 replication (see plasmids pMTH244 and pMTH249 in Fig. 3A). Within the analyzed region we did not identify any putative terminators of transcription, whose presence might block $-galactosidase expression carried from the searched promoters. Enhancer modulates the strength of a determinant of incompatibility (inc). In general two incompatible plasmids (i.e. carrying related replication or partitioning systems) cannot be stably maintained in a bacterial cell in the absence of selective pressure. To identify the incompatibility determinant within pMTH100, several selected DNA fragments of the mini-replicon (covering the whole mini-replicon; see Fig. 4) were cloned into the broad-host-range vector pRK415 (Tcr; compatible with pMTH100) in E. coli TG1 and the resulting constructs were transferred by triparental mating into P. pantotrophus KL100 containing pMTH4 mini-derivative. It was expected that the presence of the inc region on the incoming plasmid would cause the exclusion of the residing parental replicon. In the above experiments mini-replicon pMTH100 could not be used as the residing plasmid, since it encodes a stabilizing system that act on the toxin-antitoxin (TA) principium (Szymanik et al., 2004; Szymanik, 2006), which is responsible for post-segregational elimination of plasmid-less cells from bacterial population. The presence of such a system would exert a killing effect against the cells from which the plasmid was removed by reason of incompatibility. To avoid this, the mini-replicon pMTH105 was used as the recipient plasmid, which is a pMTH100 deletion derivative deprived of the functional TA system. Despite the inactivation of the TA locus, pMTH105 was very stably maintained in the bacterial population (after 72 h of growth in non-selective conditions 97% of the cells still carried the mini-replicon). The performed analysis revealed that pMTH100 carries at least one determinant of incompatibility (see pMTH201 in Fig. 4). The minimal region exerting the Inc phenotype was shown to be present within a 0.4 kb

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Fig. 4. Identification and characterization of incompatibility determinants of pMTH100. Selected DNA fragments of the mini-replicon were cloned in pRK415 vector and introduced into P. pantotrophus KL100 (pMTH105). The incompatibility of the introduced pRK415-derivatives toward pMTH105 are shown on the right: (–) compatible plasmids; (+) incompatible plasmids. The number of “+” indicates the strength of the observed incompatibility [e.g. (+) – the weakest incompatibility; (++++) lack of the residing plasmid (pMTH105) tested after 72 h of growth in non-selective conditions]. Insertion mutations introduced into repA or ORF5 are marked by asterisks. The putative IHF binding site, DnaA box and iterons have been appropriately indicated.

SmaI-XhoI restriction fragment (pMTH210; Fig. 4). Interestingly, a part of this restriction fragment (202 bp), containing all the iterons, DnaA box and incomplete IHF box (pMTH220), did not express incompatibility towards pMTH105. The presence of enhancer at the 5’ end of the minimal inc region strengthens the Inc phenotype (see plasmids pMTH218 and pMTH215 in Fig. 4). However, the enhancer itself does not express incompatibility (see pMTH209 in Fig. 4). It was also shown that mutation within ORF5 of enhancer did not influence the strength of the observed incompatibility (see pMTH215 and pMTH221 in Fig. 4). Interestingly, the obtained results demonstrated that the presence within the analyzed DNA fragments of the functional repA increases incompatibility, which points to the regulatory role of RepA protein in the regulation of plasmid replication (compare pMTH211 and pMTH212 as well as pMTH208 and pMTH214 in Fig. 4). Discussion The replication system of pMTH4 has a typical structure for the plasmid iteron-containing replicator regions (e.g. Chattoraj, 2000; Kruger et al., 2004). It contains a gene coding for replication initiation protein RepA and origin of replication placed upstream of the repA gene. The origin of pMTH4 encodes five putative iterons R1-R5, as well as two sequences similar to the IHF binding sequence and the DnaA box of E. coli. These sequences are conserved within a group of related replication systems carried by plasmids: (i) pALC1 of Paracoccus alcaliphilus (Bartosik et al., 2001), (ii) pRS241a of Rhodobacter sphaeroides (Acc. no. EAP62516) and (iii) pSD20 from Ruegeria sp. (Zhong et al., 2003), which suggests that they are functional and may play an important role in plasmid replication. In each of the listed above plasmids the iterons are placed upstream of the rep gene, however their number and arrangement within the origins varies (data not shown). Among the iterons of pMTH4 two groups could be distinguished: (i) identical repeats R1-R3 and (ii) R4 and R5 repeats placed within the repA promoter region. The R1, R2 and R3 repeats are separated by 22 bp

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– a distance which seems to enable the location of these sequences on the same side of the DNA helix, which consequently might enable cooperative binding of the Rep protein molecules (del Solar et al., 1998). It is thus probable that R1-R3 repeats are functional iterons involved in initiation of plasmid replication. The R4 and R5 repeats are not identical. The R4 carries a putative –10 hexamer of the repA promoter, while R5 is placed close to rbs, which suggests their putative role in regulation of expression of the repA gene. In this case the binding of RepA proteins to R4 and R5 might block (or decrease) transcription driven from PrepA promoter. Analogous negative regulation caused by replication proteins and iterons is observed in other iteron-containing replication systems, e.g. of prophage P1 (Sozhamannan and Chattoraj, 1993). The true role of the particular R1-R5 repeats in pMTH4 replication and replication regulation needs, however, to be experimentally verified. In this study we confirmed experimentally that the origin of replication of pMTH4 is located within the iteron-containing region. We also showed the presence of an additional element – termed an enhancer (present at the 5’ end of the origin within a 0.65 bp DNA region), which seems to regulate the process of initiation of plasmid replication. This region does not encode any proteins, therefore we speculate that it might contain a specific sequence or sequences which are recognized by a not yet identified plasmid- or host-encoded factors. As shown, the enhancer increases (i) the plasmid copy number, (ii) the strength of determinant of incompatibility (placed within the origin) and (iii) activity of the repA promoter. Enhancers are not commonly found in iteron-containing plasmids. So far such elements have been shown to be present merely in three plasmids: (i) pSC101 Salmonella panama (Miller and Cohen, 1993; Sugiura et al., 1993), (ii) R6K E. coli (Kelley et al., 1992) and (iii) pCD3.4 Carnobacterium divergens (van Belkum and Stiles, 2006). In all cases, the frequency of replication initiation is increased in the presence of the enhancer, this being manifested by higher plasmid copy number. The molecular basis of this phenomenon is not common for the studied plasmids. In the case of R6K, the enhancer (106 bp) is placed upstream of the origin of replication and contains a DnaA box. It is assumed that after the binding of protein IHF within the origin, subsequent bending of DNA enables direct interactions between the DnaA molecules bound to the enhancer and the proteins (DnaA and Rep) bound to the origin of replication (Lu et al., 1998). The enhancer of pCD3.4 carries a single iteron, which is not involved in initiation of replication but its presence increases plasmid copy number. In this case it is speculated that a replication protein binds to the enhancer region and then may probably interact with protein Rep molecules bound to the origin region (van Belkum and Stiles, 2006). Interestingly, the replicator region of plasmid pSC101 encodes two enhancers: one called par, placed upstream of the origin of replication (approx. 200 bp), and the second one placed between the iterons and a rep gene and composed of inverted repeats IR1 and IR2, which are the binding site for the Rep protein. Gyrase can bind to the par region – the protein can then change the negative superhelicity of DNA, resulting in destabilization of the DNA helix within the origin, which may stimulate the initiation of replication (Ohkubo and Yamaguchi, 1995). The common feature of the enhancers is thus the presence of sequences recognized by proteins that either interact with analogous proteins bound to the origin, or influence the structure of the origin. Enhancers are not homologous elements, for this reason they cannot be distinguished as a result of nucleotide sequence analysis. Furthermore, their mode of action has been not unequivocally determined. It is probable that DnaA or Rep mediated interactions between enhancer and origin might potentially hinder the access of Rep protein dimers to the origin (dimers negatively regulate the process of initiation of replication), which may lead to the prolonged time of the life of the open complex, and in effect bring about the initiation of a larger number of replication rounds. In this case negative regulation would be blocked by handcuffing, in which a key role would seem to be played by Rep protein dimers. These speculations might be confirmed by the results obtained for R6K. It was shown that R6K enhancer counteracts inhibition of initiation of replication caused by excess of replication protein B (such conditions favour the occurrence of dimers that inhibit the replication process) (Wu et al., 1994). In this study we have shown that the enhancer of pMTH4 increases the activity of the repA promoter, however it itself does not encode any promoter sequences. It is probable that increased activity of P repA (and consequently increased level of Rep protein) might potentially enhance the level of initiation of replication, however, such a directly proportional dependence might be observed exclusively in the absence of handcuffing. It would be interesting to know whether the activity of PrepA would be increased in the presence in trans of a RepA protein, which might potentially negatively regulate expression of repA gene at the level of transcription. It is also probable that the increased activity of PrepA, may be a secondary effect, caused by the potential binding of cellular proteins to the enhancer with subsequent destabilization of the helix with the origin region, thus facilitating the access of RNA polymerase.

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Fig. 5. Hypothetical model for functioning of the replication system of pMTH4. The RepA molecules are marked with the letter “R”. The RepA protein binds to the iterons (boxes with arrows), which activates initiation of replication (+) or negatively regulates expression of the repA gene (–). The DnaA and IHF molecules are appropriately indicated. A putative, not yet identified protein binding to the enhancer and able to interact with the origin, is marked with an “X”. The –35 and –10 hexamers of the repA promoter (PrepA) are indicated. See text for details.

Based on the obtained results and data concerning other enhancer-containing replicons we constructed a hypothetical model of functioning of the replication system of pMTH4 (Fig. 5). It assumes that (i) RepA proteins binds the R1-R3 iterons, which activates the origin, (ii) RepA also binds the R4 and R5 repeats, which negatively regulate expression of the repA gene, (iii) DnaA protein binds within the origin and directs chromosomally encoded replication proteins to the oriV, (iv) IHF binds within the origin and bends DNA, which consequently enable interactions between not yet identified putative proteins bound in enhancer with proteins bound within the origin. This might result in destabilization of the DNA helix within the origin, which facilitates open complex formation (or prolongs the time of its life). This directly translates to increased frequency of initiation of replication, and thus an increase in plasmid copy number. The iteron-containing replicons are usually low copy number plasmids. Their stable maintenance is determined by the presence of the partitioning systems (par), which are responsible for proper distribution of plasmid copies to daughter cells at cell division. All the plasmids that are closely phylogenetically related with pMTH4 (pALC1, pRS241a, pSD20) carry the partitioning modules placed in the close proximity to their replication systems. Genomic analysis of plasmid pMTH4 revealed that this replicon does not encode the par region (data not shown), therefore the only stabilizing system of this plasmid (placed within the sta region) is a toxin-antitoxin cassette, which is responsible for the post-segregational elimination of plasmid-less cells. It is thus probable that the pMTH4 copies are randomly segregated at cell division. Thus if pMTH4 was a low copy number plasmid, then each round of replication would be followed by the formation of a large number of plasmid-less cells being eliminated from the population by the action of the toxin-antitoxin system. This would ultimately lead to elimination of the whole bacterial population. The presence of enhancer that increases plasmid copy number seems to compensate for the lack of an active partitioning system in the plasmid. Literature B a j J., E. P i e c h u c k a, D. B a r t o s i k and M. W l o d a r c z y k. 2000. Plasmid occurrence and diversity in the genus Paracoccus. Acta Microbiol. Pol. 49: 265–270. B a r t o s i k D., A. B i a l k o w s k a, J. B a j and M. W l o d a r c z y k. 1997. Construction of mobilizable cloning vectors derived from pBGS18 and their application for analysis of replicator region of a pTAV202 mini-derivative of Paracoccus versutus pTAV1 plasmid. Acta Microbiol. Pol. 46: 387–392. B a r t o s i k D., M. W i t k o w s k a, J. B a j and M. W l o d a r c z y k. 2001. Characterization and sequence analysis of the replicator region of the novel plasmid pALC1 from Paracoccus alcaliphilus. Plasmid 45: 222–226. B a r t o s i k D., J. B a j, A.A. B a r t o s i k and M. W l o d a r c z y k. 2002. Characterisation of the replicator region of megaplasmid pTAV3 of Paracoccus versutus and search for plasmid-encoded traits. Microbiology 148: 871–881. B r u a n d C., E. L e C h a t e l i e r, S.D. E h r l i c h and L. J a n n i e r e. 1993. A fourth class of theta-replicating plasmids: the pAM1 family from Gram-positive bacteria. Proc. Natl. Acad. Sci. USA 90: 11668–11672. C h a t t o r a j D.K. 2000. Control of plasmid DNA replication by iterons: no longer paradoxical. Mol. Microbiol. 37: 467–476. d e l S o l a r G., R. G i r a l d o, M.J. R u i z - E c h e v a r r i a, M. E s p i n o s a and R. D i a z - O r e j a s. 1998. Replication and control of circular bacterial plasmids. Microbiol. Mol. Biol. Rev. 62: 434–464. D i t t a G., S. S t a n f i e l d, D. C o r b i n and D.R. H e l i n s k i. 1980. Broad host-range DNA cloning system for Gram-negative bacteria: construction of a bank of Rhizobium meliloti. Proc. Natl. Acad. Sci. USA 77: 7347–7351. D o r o n i n a N.V., Y.A. T r o t s e n k o, V.I. K r a u s o v a and N.E. S u z i n a. 1998. Paracoccus methylutens sp. nov. – a new aerobic facultatively methylotrophic bacterium utilizing dichloromethane. Syst. Appl. Microbiol. 21: 230–236.

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H a n a h a n D. 1983. Studies on transformation of Escherichia coli with plasmids. J. Mol. Biol. 166: 557–580. K e e n N.T., S. T a m a k i, D. K o b a y a s h i and D. T r o l l i n g e r. 1988. Improved broad-host-range plasmids for DNA cloning in gram-negative bacteria. Gene 70:191–197. K e l l e y W.L., I. P a t e l and D. B a s t i a. 1992. Structural and functional analysis of a replication enhancer: separation of the enhancer activity from origin function by mutational dissection of the replication origin ( of plasmid R6K. Proc. Natl. Acad. Sci. USA 89: 5078–5082. K r u g e r R., S.A. R a k o w s k i and M. F i l u t o w i c z. 2004. Participating elements in the replication of iteron-containing plasmids, pp. 25–45. In: B.E. Funnell and G.J. Phillips (eds), Plasmid Biology, ASM Press, Washington, D.C. K u s h n e r S.R. 1978. An improved method for transformation of E. coli with ColE1-derived plasmids. pp. 17–23. In: H.B. Boyer and S. Nicosia (eds), Genetic Engineering, Elsevier/North-Holland, Amsterdam. L u Y.B., H.J. D a t t a and D. B a s t i a. 1998. Mechanistic studies of initiator-initiator interaction and replication initiation. EMBO J. 17: 5192–5200. M a r x C.J. and M.E. L i d s t r o m. 2001. Development of improved versatile broad-host-range vectors for use in methylotrophs and other Gram-negative bacteria. Microbiology 147: 2065–2075. M e i j e r W.J.J., A.J. d e B o e r, S. v a n T o n g e r e n, G. Ve n e m a and S. B r o n. 1995. Characterization of the replication region of the Bacillus subtilis plasmid pLS20: a novel type of replicon. Nucleic Acids Res. 23: 3214–3223. M e s s e r W., F. B l a e s i n g, D. J a k i m o w i c z, M. K r a u s e, J. M a j k a, J. N a r d m a n n, S. S c h a p e r, H. S e i t z, C. S p e c k, C. W e i g e l, G. W e g r z y n, M. W e l z e c k and J. Z a k r z e w s k a - C z e r w i n s k a. 2001. Bacterial replication initiator DnaA. Rules for DnaA binding and roles of DnaA in origin unwinding and helicase loading. Biochimie 83: 5–12. M i l l e r J.H. 1972. Experiments in Molecular Genetics. Cold Spring Harbor, New York. M i l l e r C.A. and S.N. C o h e n. 1993. The partition (par) locus of pSC101 is an enhancer of plasmid incompatibility. Mol. Microbiol. 9: 695–702. O h k u b o S. and K. Y a m a g u c h i. 1995. Two enhancer elements for DNA replication of pSC101, par and a palindromic binding sequence of the Rep protein. J. Bacteriol. 177: 558–565. R e e c e K.S. and G.J. P h i l l i p s. 1995. New plasmids carrying antibiotic-resistance cassettes. Gene 165: 141–142. S a m b r o o k J. and D.W. R u s s e l l. 2001. Molecular Cloning. A Laboratory Manual. Cold Spring Harbor, New York. S c h a g g e r H. and G. v o n J a g o w. 1987. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166: 368–379. S o z h a m a n n a n S. and D.K. C h a t t o r a j. 1993. Heat shock proteins DnaJ, DnaK, and GrpE stimulate P1 plasmid replication by promoting initiator binding to the origin. J. Bacteriol. 175: 3546–3555. S p r a t t B.G., P.J. H e d g e, S. t e H e e s e n, A. E d e l m a n and J.K. B r o o m e - S m i t h. 1986. Kanamycin-resistant vectors that are analogues of plasmids pUC8, pUC9, pEMBL8 and pEMBL9. Gene 41:337–342. S u g i u r a S., S. O h k u b o and K. Y a m a g u c h i. 1993. Minimal essential origin of plasmid pSC101 replication: requirement of a region downstream of iterons. J. Bacteriol. 175: 5993–6001. S z y m a n i k M., D. B a r t o s i k and M. W l o d a r c z y k. 2004. Genetic organization of the basic replicon of plasmid pMTH4 of a facultatively methylotrophic bacterium Paracoccus methylutens DM12. Curr. Microbiol. 48: 291–294. S z y m a n i k M. 2006. Ph.D. Thesis. Warsaw University, Warsaw. v a n B e l k u m M.J. and M.E. S t i l e s. 2006. Characterization of the theta-type plasmid pCD3.4 from Carnobacterium divergens, and modulation of its host range by RepA mutation. Microbiology 152: 171–178. W l o d a r c z y k M., E.K. J a g u s z t y n - K r y n i c k a, D. B a r t o s i k and I. K a l i n o w s k a. 1994. Electroporation of Thiobacillus versutus with plasmid DNA. Acta Microbiol. Pol. 43: 223–227. W u F., I. L e v c h e n k o and M. F i l u t o w i c z. 1994. Binding of DnaA protein to a replication enhancer counteracts the inhibition of plasmid R6K ( origin replication mediated by elevated levels of R6K B protein. J. Bacteriol. 176: 6795–6801. Z h o n g Z., R. C a s p i, D. H e l i n s k i, V. K n a u f, S. S y k e s, C. O’ B y r n e, T.P. S h e a, J.E. W i l k i n s o n, C. D e L o u g h e r y and A. T o u k d a r i a n. 2003. Nucleotide sequence based characterizations of two cryptic plasmids from the marine bacterium Ruegeria isolate PR1b. Plasmid 49: 233–252.

Polish Journal of Microbiology 2006, Vol. 55, No 4, 271– 277

Acquisition of Iron by Enterococci: Some Properties and Role of Assimilating Ferric Iron Reductases PAWE£ LISIECKI* and JERZY MIKUCKI

Chair of Biology and Biotechnology, Department of Pharmaceutical Microbiology, Medical University, £ódŸ, Poland Received 29 May 2006, revised 30 August 2006, accepted 12 September 2006 Abstract Ferric iron reductases activities have been occurred in 91% of investigated enterococci strains. Maximum activity occurred with coenzyme NADH as the reductant and the presence of cofactor FMN was necessary. Mg(II) ions has not stimulated reductases activity. Treatment of cells with proteolytic enzymes had not effect on iron reduction. The whole cells and cell fraction – cytoplasmic membrane and cytoplasm showed Fe(III) – reducing activity. The highest specific activity was associated with cytoplasm. The activity in cytoplasmic membrane was not related to iron concentration in the growth medium. In cytoplasm the activity was stimulated after growth in low-iron medium. Ferric iron reductases of enterococci characterized the broad substrate specificity. The iron in form of ferric ammonium citrate, lactoferrin and ferrioxamine B were the best iron sources for enterococcal ferric iron reductases. K e y w o r d s: Enterococcus spp., ferric iron reductases, iron acquisition

Introduction Reduction of complexed Fe (III) to Fe (II) decreases its affinity to the carrier: stability constant K = 1030 of Fe(III)-siderophore after reduction amounts to K = 108 (Silver and Walderhaug, 1992; Clarke et al., 2001). The complex dissociates and iron is released in the Fe(II) form easily assimilated by bacteria. In the cell in the presence of ferrochelatase it is used for the synthesis of haeme and non haeme iron-containing proteins. It is also a corepressor of Fur protein – a negative repressor controlling expression of iron uptake and transport systems (Clarke et al., 2001; Schröder et al., 2003). The ferric iron reductases are used for assimilating Fe(III) for the purpose of intracellular incorporation into protein. They have occurred in all bacteria excluding a small group of lactic acid bacteria (Schröder et al., 2003). Assimilating ferric iron reductases can be localized in bacterial cytoplasm, cytoplasmic membrane and periplasmic space (Schröder et al., 2003). Some pathogenic bacteria release ferric iron reductases to the environment or expose them on the cell surface (Barchini and Cowart, 1996; Deneer et al., 1995; Homuth et al., 1998). These reductases permit bacteria to reduce in the environment iron Fe(III) free and bound to various carriers. The cytoplasmic reductases release Fe(II) by reduction from Fe(III)-siderophore complex introduced to the cell (Ratledge and Dover, 2000). Assimilating iron ferric reductases were detected in a number of pathogenic and facultative pathogenic bacteria (Barchini and Cowart, 1996; Coves and Fontecave, 1993; Johnson et al., 1991; Le Faou and Morse, 1991). However, no data are available on ferric iron reductases in enterococci. The part of own research we have published earlier (Lisiecki and Mikucki, 2005). The results of these studies reveal more information on the occurrence, some properties and the role of these enzymes. * Corresponding autor: P. Lisiecki, Chair of Biology and Biotechnology, Dept. Pharmaceutical Microbiology, Medical University, Pomorska 137, 90-235 £ódŸ, Poland; phone./fax. + 48 42 677 93 00; e-mail: [email protected]

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Experimental Materials and Methods Bacterial strains. One hundred twenty strains of Enterococcus genus were used. Forty-nine of them originated from our own collection (Department of Pharmaceutical Microbiology, Medical University, £ódŸ) , and rest from the National Institute of Public Health in Warsaw, Microbiology Department Collegium Medical University of Toruñ in Bydgoszcz, Czech National Collection of Type Cultures (CNCTC), Deutsche Sammlung von Mikroorganismen und Zellkulturen (DSMZ) and American Type Culture Collection (ATTC). Enterococci from our collection, were identified with API-STREP system (bioMerieux). Suspensions density and viable count. The optical density of suspension and cultures was measured in UV/VIS Cary 1 (Varian) spectrophotometer at 580 nm. It was expressed in form of ODU (optical density unit) corresponding to absorbance value A580 = 1.0. Mc Farland scale (Difco) and calibrated loops were used also. Viable count was estimated by using serial dilutions in buffered 0.85% NaCl, pH 7.2 and standard plate methods on 4% Trypticase Soy Agar (Difco). Growth media. Strains were grown on medium containing (per litre): 3 g Casamino Acids, Vitamin Free (Difco); 3 g Yeast Extraxt (Difco); 3 g KH2PO4; 5 g NaCl; 1 g NH4Cl; 0.09 g MgCl2; 0.01 g CaCl2; 12.1 g Tris (BDH) and 20 g glucose. pH of medium was adjusted to 7.2. Concentration of iron was reduced using polyaminocarboxylate resin Chelex 100 (200 – 400 mesh, BioRad). In some experiments the medium was supplemented with 5 µM haeme (Sigma) and 0.1 µM haemoglobin (Sigma). Solid medium contained 1.5% Noble Agar N° 1 (Oxoid). Mueller-Hinton 2 agar medium contained 50 µM o-phenantroline (Fluka) was used also. Growth conditions. Strains were initially iron-starved for 18 hours at 37°C on Mueller-Hinton 2 agar medium (Difco) containing 50 µM o-phenantroline (Fluka). The optical density of prepared suspension corresponded to standard N° 2 of Mc Farland scale and contained 6×10 8 cells per ml. This starved suspensions were used to inoculate (5% v/v) media of different iron content: Fe + with 10–4 M of iron in the form of FeSO4 × 7H2O and Fe– (Chelex) subject to Chelex 100 (BioRad) resin and containing approximately 1.2 × 10–6 M – 3.5 × 10–6 of iron. Cultures were incubated at 37°C for 24 hours under constant shaking and centrifuged (9500 × g, 15 min, 4°C). The cells were washed with buffered 0.155 M NaCl, pH 7.2 and supernatant was filtered through the membrane filter (0.22 µm, Millipore). Protoplast formation and fractionation. Protoplast of enterococci was prepared with lysozyme (Serva) according to the method of Zorzi et al. (1996) and Lindberg (1981). The protoplast were lysed in 0.01 M TRIS-HCl (pH 7.2) buffer, DNase (1 µg/ml; Sigma) and RNase (2.5 µg/ml; Sigma) were added and the suspension was incubated in an ice water bath for 30 min. The protoplast lysate was pre-centrifuged (3000 × g, 15 min, 4°C) the pellet discarded and the supernatant was dialysed against deionized water and centrifuged at 100 000 × g for 60 min at 9°C (Beckman L8-70M) to obtain membrane (pellet) and cytoplasmic (supernatant) fraction. The protein content was estimated by the method of Lowry et al. (1951) and both fraction were lyophilized. Ferric iron reductase assay. The reduction of Fe(III) was assayed with ferrozine [3-(2-pirydyl)-5,6-bis(4-phenylsulfonic acid)1,2,3-triazine] (Sigma) as Fe(II) trapping reagent (Deneer et al., 1995). Ferrozine is water soluble, does not react with Fe(III) iron. The ferrous complex of ferrozine has an absorbance maximum at 562 nm (Stookey, 1970; Cowart et al., 1993). Ferric iron reductases activity detection. Standardized suspension of starved enterococci was spotted with calibrated loop (0.01 ml) on the surface of solid medium. After 24 h of incubation in 37°C the plate was flooded with 15 ml of 0.8% agarose solution (Sigma) containing: 2 mM ferrozine (Sigma), 50 µM NADH (Sigma), 50 µM FMN (Sigma) and 100 µg/ml of substrate – ferric ammonium citrate (Sigma). The plates were incubated in 37°C for 3 h in darkness. The violet zone around the bacteria proved the presence of Fe(II)-ferrozine complex and, indirectly, iron reductase activity (Mazoy and Lemos, 1999). Determination of ferric iron reductases activity of whole cells. The cells were suspended in 8 ml of medium without MgCl2, CaCl2 and glucose and 1 ml of suspension was withdrawn to measure optical density of at 580 nm. The reaction mixture in final volume of 7 ml contained: bacteria, 50 µM NADH (Sigma), 3 µM FMN (Sigma) and 2 mM ferrozine (Sigma). In some experiments the reaction mixture contained 10 mM MgCl2 (Sigma). After adding 100 µl substrate of reduction (100 µg/ml final concentration) polystyrene tubes were shaked and incubated in 37°C. After 10, 20 and 30 minutes 1 ml of reaction mixture was withdrawn, centrifuged (9500 × g, 15 min, 4°C) and absorbance at 562 nm was measured in spectrophotometer UV/VIS Cary 1 (Varian). The control of assay contained all of the reagents except source of enzyme. The amount of Fe(II)-ferrozine complex was calculated as difference between the absorbance value of whole reaction mixture and control. Specific activity of ferric iron reductases was expressed as amount of µmol Fe(II)-ferrozine complex formed per ODU (optical density units) per minute (µmol Fe(II)-ferrozine/ min/ODU) (Deneer et al., 1995). The results represent the means of three separate experiments. Determination of ferric iron reductases activity in cell fractions. The sources of enzyme was cell fractions – membrane or cytoplasm. The reaction mixture contained in final of 2 ml volume: 0.5 ml of source of enzyme, 250 µM NADH (Sigma), 50 µM FMN (Sigma) and 250 µM ferrozine (Sigma) and of 0.01 M TRIS-HCl (pH 7.4) buffer. After adding 10 µl substrate of reduction (100 µg/ml final concentration) to polystyrene cuvettes the change in absorbance at 562 nm after 10, 20 and 30 minutes was measured in spectrophotometer UV/VIS Cary 1 (Varian). The assay was performed at 37°C. The control of assay contained all of the reagents except source of enzyme. The amount of Fe(II)-ferrozine complex was calculated as difference between the absorbance value of whole reaction mixture and control. Specific activity of ferric iron reductases was expressed as amount of µmol Fe(II)-ferrozine complex formed per g of protein per minute [µmol Fe(II)-ferrozine/min/g of protein] (Mazoy and Lemos, 1999). The results represent the means of three separate experiments. Determination of iron concentration. Iron concentration in media was determined as described by Gadia and Mehra (1977) with ferrozine (Sigma) using UV/VIS Cary 1 (Varian) spectrophotometer at the 526 nm. Substrate of reduction reactions. The following substrates were used: ferric ammonium sulphate (Sigma), ferric nitrate (POCH), ferric ammonium citrate (Sigma), ferric versenate (POCH), ferric pyruvate (Sigma), transferrin (Sigma), lactoferrin (Sigma), haemoglobin (Sigma), ferritin (Sigma) and ferrioxamin B (Sigma). Treatment of cells with proteolytic enzymes. This experiment was carried out according to Deneer et al. (1995). The cells were suspended in 5 ml of 0.1 M Tris/HCl (pH 7.4) buffer supplemented with 1 mM MgCl 2 (Sigma) and proteinase K (0.5 mg/ml)

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(Sigma) or trypsine (0.5 mg/ml) (Sigma) and the suspension was incubated at 37°C for 45 min. The cells were washed twice with buffered 0.155 M NaCl (pH 7.2) and resuspended in medium without salt and glucose and assayed for iron reductase activity. Statistical analysis. Statistical analysis was performed with the Statistica PL computer programme (StatSoft) and nonparametric test of Kruskal-Wallis and U Mann-Whitney (Dixon and Massey, 1951). Statistical significance was defined as p < 0.05.

Results Screening of strains of enterococci on the solid medium with ferrozine as Fe(II) iron trapping agent showed that 111 strains (91%) possessed reductase activity towards ferric ammonium citrate as a substrate. These data were confirmed by ferrozine test with resting cells. The studied strains differed in the activity in a wide range between 0.06 µmol of Fe(II)-ferrozine/min/ODU and 1.03 µmol of Fe(II)-ferrozine/min/ODU. There was no correlation between origin of strain, its species affiliation and reductase activity (p> 0.05). For further investigation 6 strains with high reductase activity were selected: 4 out of them were derived from clinical material – E. faecalis BY 56 and BD 123 and E. faecium 97-0 and BY 9, and two from the environment – E. saccharolyticus DSM 20726 and E. sulfureus DSM 6905. Only E. saccharolyticus DSM 20726 reduced Fe(III) in absences of exogenous coenzymes such as NADH or NADPH and FMN cofactor. In the remaining strains NADH or NADPH alone did not activate ferric reductases. The presence of FMN as cofactor was necessary and activated ferric iron reductase itself but the most effectively when NADH was a donor of electrons (Table I). Magnesium ions did not affect a degree of reduction (p> 0.05). Fe (III) deficiency at the concentration of 3.5× 10 –7 M in the medium did not influence the activity of ferric reductase in whole cells (p > 0.05) (Table II). The presence of ferroporphyrin-haemin in the concentration of 5 µM or haemoglobin in the concentration of 0.1 µM did not change the activity also (p> 0.05). Reduction of Fe (III) connected with a carrier required its contact with the cell. Separation of lactoferrin from the growing cells by a dialyzing membrane hindered its reduction. Modification of proteins on the Table I Effect of NADH, NADPH and FMN as reductans on ferric reductase activity in whole cells of enterococci Ferric iron reductese activity* –

NADH (50 µM)

NADPH (50 µM)

FMN (3 µM)

NADH + FMN

NADPH + FMN

E. faecalis BD 123

0

0

0

0.046

0.18

0.07

E. feacalis BY 56

0

0

0

0.043

0.09

0.075

E. faecium BY 9

0

0

0

0.14

0.21

0.09

E. faecium 97–0

0

0

0

0.08

0.11

0.12

1.55

1.50

1.10

2.50

2.90

3.20

0

0.28

0.032

0.32

0.36

0.14

Strains

E. sulfureus DSM 6905 E. saccharolyticus DSM 20726

* expressed as µM Fe(II)-ferrozine/min/ODU

Table II Effect of iron concentration in medium on ferric iron reductase activity in whole cells of enterococci Strains

Ferric iron reductese activity* Fe + medium

Fe- (Chelex) medium

E. faecalis BD 123

0.35

0.37

E. feacalis BY 56

0.14

0.12

E. faecium BY 9

0.25

0.11

E. faecium 97–0

0.15

0.12

E. sulfureus DSM 6905

1.78

1.70

E. saccharolyticus DSM 20726

0.28

0.18

* expressed as µM Fe(II)-ferrozine/min/ODU

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Lisiecki P. and J. Mikucki Table III Effect of trypsin or proteinase K treatment of cells on ferric iron reductase activity in enterococci Ferric iron reductese activity* Strains

Modificated cell

Intact cells

Trypsin

Proteinase K 0.15

E. feacalis BD 123

0.18

0.22

E. faecalis BY 56

0.03

0.01

0.04

E. faecium BY 9

0.01

0.02

0.01

E. faecium 97–0

0.02

0.02

0.04

E. sulfureus DSM 6905

0.06

0.12

0.05

E. saccharolyticus DSM 20726

0.14

0.25

0.03

* expressed as µM Fe(II)-ferrozine/min/ODU

surface of the cells decomposed by trypsin or proteinase K did not influence the activity of ferric reductase activity (p> 0.05) (Table III). The concentrated and non-concentrated culture supernatants did not show ferric iron reductase activity. It was present in whole cells and two subcellular fractions: cytoplasm and cytoplasmic membrane obtained after cells protoplastization and lysis of protoplasts. Ferric reductase activity in the cytoplasm was higher either in cells grown in the medium with iron deficiency [Fe- (Chelex)] and its abundance (Fe+) (p< 0.05). It amounted in mean value to 63 and 41 µmol of Fe(II)-ferrozine/min/g of protein) when corresponding values of activity in cytoplasmic membranes amounted to12.0 and 7.2 µmol of Fe(II)-ferrozine min/g of protein. Iron deficiency during the growth caused a significant increase in the activity of ferric iron reductases only in the cytoplasmic fraction (p < 0.1) (Table IV). Ferric iron reductases were characterized by wide substrate specificity. They reduced inorganic compounds of iron – ferric ammonium sulfate and ferric nitrate, organic compounds – ferric ammonium citrate, ferric versenate and ferric pyruvate, body sources of iron – transferrin, lactoferrin, hemoglobin and ferritin, and bacterial siderophore-ferrioxamine B (data not shown). No differences were found in reductase activity of strains towards all substrates (p > 0.05). Significant differences were detected only when groups of substrates were assessed (p < 0.05). Substrates reduced with the highest activity such as ferrioxamine B, lactoferrin and ferric ammonium citrate belonged to the first group (0.26, 0.18, 0.17 µmol of Fe(II)-ferrozine/ min/ODU, respectively). The second group involved ferric ammonium sulphate, ferric versenate and ferric nitrate (0.12, 0.09 and 0.042 µmol of Fe(II)-ferrozine min/ODU). The third group involved substrates reduced with the lowest activity: transferrin, ferric pyruvate, haemoglobin and ferritin (0.02, 0.019, 0.016, 0.016 µmol of Fe(II)-ferrozine/min/ODU).

Table IV Effect of iron concentration in medium on specific activity of ferric iron reductase in cell fraction of enterococci Ferric iron reductese activity* Cytoplasm

Strains

Cytoplasmic membrane

Fe +

Fe- (Chelex)

Fe +

Fe- (Chelex)

E. faecalis BD 123

49.0

140.0

10.0

8.0

E. feacalis BY 56

17.0

27.0

13.0

13.0

E. faecium BY 9

35.0

25.0

6.0

10.0

E. faecium 97–0

26.0

38.0

5.0

6.0

E. sulfureus DSM 6905

66.0

56.0

1.0

15.0

E. saccharolyticus DSM 20726

53.0

91.0

8.0

20.0

* expressed as µM Fe(II)-ferrozine/min/g of protein

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Discussion Assimilating ferric iron reductases are widely spread in Gram-positive and Gram-negative bacteria (Johnson et al., 1991; Le Faou and Morse, 1991; Deneer et al., 1995; Coves and Fontecave, 1993; Barchini and Cowart, 1996; Homuth et al., 1998). In more than 90% of enterococci of various species and origin reductase activity of ferric ammonium citrate was found. Bacterial ferric reductases were characterized by wide substrate specificity. They reduced Fe(III) in inorganic and organic compounds, complexes with natural and synthetic chelators, bound to the host body carriers and even free Fe(III) ions (Schröder et al., 2003; Ratledge and Dover, 2000). Enterococci reduced Fe(III) of inorganic and organic compounds, siderophores and body iron carriers also. The ability to reduce Fe(III) of lactoferrin was crucial for colonization of the macroorganism but a minor participation of the rest of body carriers-transferrin, hemoglobin and ferritin in the reduction was enigmatic. Assimilating ferric iron reductases use NADH and NADP coenzymes and sometimes glutathione as electron donors. The presence of free cofactors in the form of flavins – FMN, FAD or riboflavin is necessary for the reduction (Schröder et al., 2003). NADH as a coenzyme and FMN as a cofactor occur more often in bacteria (Schröder et al., 2003; Ratledge and Dover, 2000). Resting cells of enterococci did not reduce ferric ammonium citrate even in the presence of NADH or NADPH. Only free flavin (FMN) stimulated ferric reduction, and endogenous coenzymes associated with cell surface were electron donors. Exogenous coenzymes stimulated ferric reduction only in the presence of flavin (FMN), and NADH did it stronger as compared with NADPH. Same as in E. coli (Fontecave et al., 1987) and P. aeruginosa (Halle and Meyer, 1992) free flavin FMN could be reduced by enterococci and in this form served as direct, chemical reductant for numerous substrates. This reductase would be in fact a flavin reductase, not a ferric reductase. Assimilating ferric iron reductases are constitutive enzymes. In any studies a regulating role of iron was not noted in their production (Arceneaux and Byers,1980; Johnson et al., 1991; Deneer et al., 1995; Cowart, 2002). Activity of ferric iron reductases of whole enterococcal cells was not regulated by iron availability. Some bacterial ferric reductases are released to the environment (Homuth et al., 1998). Most often ferric reduction occurs when iron ions or their carriers contact with reductases exposed on the cell surface. In test with ferrozine only iron Fe(II) bound to the surface of resting cells and released to the environment after reduction was detected because this compound did not penetrate through cytoplasmic membrane (Stookey, 1970; Cowart et al., 1993; Deneer et al., 1995). Enterococci like Listeria monocytogenes (Deneer et al., 1995) did not reduce Fe (III) in lactoferrin when were separated from the substrate by a dialysis membrane so, the contact between substrate and cell was necessary. Modification of the cell surface of enterococci by means of the breakdown of proteins neither eliminated nor inhibited cooperation between cells and substrates same as in Legionella pnemophila (Johnson et al., 1991) and Listeria monocytogenes (Deneer et al., 1995). Enterococcal reductases can be localized deeper in the cell cover and less available for proteolytic enzymes. Bacterial assimilating ferric iron reductases can be detected in the cytoplasmic membrane, periplasmic space and cytoplasm (Schröder et al., 2003). They can be found in all subcellular fractions (Schröder et al. 2003; Cowart, 2002) and differ in the use of electron donors (Schröder et al., 2003). Location of reductases in the cytoplasmic membrane and periplasmic space has a strategic role: permits a contact with the substrate. Reductases occurring in the growth environment can be enzymes actively secreted or because of loose binding to the membrane and periplasmic spaces released in to the growth medium (Barchini and Cowart, 1996; Deneer et al., 1995). Detection of more and more exogenous ferric iron reductases allowed us to draw a conclusion that iron acquisition by means of their parcipitation could be a common bacterial strategy. However enterococci neither released from the cell surface nor secreted ferric reductases to the environment. It diminished their strategic role in the efficacy of iron acquisition. Ferric iron reductases occur in enterococci in two subcellular fractions: cytoplasm and cytoplasmic membrane. It is a localization of ferric reductases in the majority of bacteria. In enterococci as in other bacteria these reductases create two groups of enzymes: constitutive, connected with cytoplasmic membrane and inductive occurred in cytoplasm. Specific activity of iron ferric reductase in cytoplasm was higher as compared to the cytoplasmic membrane enzymes. Recently published data on cooperation between siderophores and ferric iron reductases can change our attitude towards their significance for iron acquisition by bacteria (Vartivarian and Cowart, 1999; Cowart, 2002). Siderophores are induced chelators while assimilating reductases are constitutive enzymes which synthesis in contradistinction to siderophores takes place independently of iron availability. Siderophores have been classified as secondary metabolites recently, because their production occurs in late phases of growth such as stationary and death phase. Ferric iron reductases are produced at all phases of growth.

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These features cause that they can play more spectacular role as compared with siderophores in mobilization of Fe(III) sources in the environment of bacteria. Therefore, the presence of at least two separate pathways of iron acquisition by bacteria, which produce siderophores and ferric iron reductases, have been postulated. Enterococci also belong to this group. In conditions of iron availability in the environment ferric iron reductases transform Fe(III) into Fe(II) which can be transported to the cell through Nramp transporter (natural resistance-associated macrophages proteins) or/and a pathway of simple diffusion (Halle and Meyer, 1992). With limited iron Fe(III) availability and therefore decreasing Fe(II) cell concentration derepression of siderophore system occurs and with participation of these chelators a second way of iron acquisition is opened. Siderophores bind free and complexed Fe(III) and also free Fe(II) after reduction. Binding of the latter to siderophores leads to their immediate oxidation to Fe (III) and Fe (III)-siderophores complex can be transferred to cytoplasm. The action of these two ways requires the presence of constitutive ferric iron reductases (Cowart, 2002). In enterococci a tight binding of reductases to cell can be a limitation in their ascribed action. Therefore they cannot mobilize Fe(III) sources through reduction outside the cell. They reduce substrates only adjoining to the cell surface and Fe(II) transportation into cytoplasm could take place with simple diffusion. Grampositive bacteria such as L. monocytogenes (Barchini and Cowart, 1996) and Gram-negative as E. coli and P. aeruginosa (Vartivarian and Cowart, 1999) make use of such individual way of Fe(II) ions transport, separated from the way of transport of Fe (III)-siderophore complexes. Cytoplasmic ferric iron reductases in enterococci are probably an induced enzymes. They can be a final part of induced siderophore system. After introduction with a receptor of Fe(III)-siderophore complex to cytoplasm they release iron through its reduction to Fe(II) which in the presence of ferrochelatase can be built in haeme and non-haeme proteins. Cytoplasmic ferric reductases dependent on NADH or NADPH for Fe(III)-schizokinen, ferrioxamine B and Fe(III)-aerobactin were detected in Mycobacterium, Bacillus and Acinetobacter genera (Clarke et al., 2001; Ratledge and Dover, 2000) Acknowlegment: This research was supported by grant from Medical University in £ódŸ No 502-13-224

Literature A r c e n e a u x J.E. and B.R. B y e r s. 1980. Ferrisiderophore reductase activity in Bacillus megaterium. J. Bacteriol. 141: 715–721. B a r c h i n i E. and R.E. C o w a r t. 1996. Extracellular iron reductase activity produced by Listeria monocytogenes. Arch. Microbiol. 166: 51–57. C l a r k e T.E, L.W. T a r i and H.J. Vo g e l. 2001. Structural biology of bacterial iron uptake systems. Curr. Top. Med. Chem. 1: 7–30. C o w a r t R.E., F.L. S i n g l e t o n and J.S. H i n d. 1993. A comparison of bathophenanthrolinedisulfonic acid and ferrozine as chelators of iron(II) in reduction reactions. Anal. Biochem. 211: 151–155. C o w a r t R.E. 2002. Reduction of iron by extracellular iron reductases: implications for microbial iron acquisition. Arch. Biochem. Biophys. 400: 273–281. C o v e s J. and M. F o n t e c a v e. 1993. Reduction and mobilization of iron by a NAD(P)H: flavin oxidoreductase from Escherichia coli. Eur. J. Biochem. 211: 635–641. D i x o n W.J. and F.J. M a s s e y. 1951. Introduction to statistical analysis. McGraw-Hill Book Co., New York. D e n e e r H.G., V. H e a l e y and I. B o y c h u k. 1995. Reduction of exogenous ferric iron by a surface-associated ferric reductase of Listeria spp. Microbiology 141: 1985–1992. F o n t e c a v e M., R. E l i a s s o n and P. R e i c h a r d. 1987. NAD(P)H: flavin oxidoreductase of Escherichia coli. A ferric iron reductase participating in the generation of the free radical of ribonucleotide reductase. J. Biol. Chem. 262: 12325–12331. G a d i a M.K. and M.C. M e h r a. 1977. Rapid spectrophotometric analysis of total and ionic iron in the µg range. Microchemica Acta (Wien) 11: 413–418. H a l l e F. and J.M. M e y e r. 1992. Ferrisiderophore reductases of Pseudomonas. Purification, properties and cellular location of the Pseudomonas aeruginosa ferripyoverdine reductase. Eur. J. Biochem. 209: 613–620. H o m u t h M., P. Va l e n t i n - W e i g a n d, M. R o h d e and G.F. G e r l a c h. 1998. Identification and characterization of a novel extracellular ferric reductase from Mycobacterium paratuberculosis. Infect. Immun. 66: 710–716. J o h n s o n W., L. Va r n e r and M. P o c h. 1991. Acquisition of iron by Legionella pneumophila: role of iron reductase. Infect. Immun. 59: 2376–2381. L e F a o u A.E. and S.A. M o r s e. 1991. Characterization of a soluble ferric reductase from Neisseria gonorrhoeae. Biol. Met. 4: 126–131. L i n d b e r g M. 1981. Genetic studies in Staphylococcus aureus using protoplast: cell fusion and transformation, p. 535–541. In: J. Jeljaszewicz (ed.), Staphylococcus and staphylococcal infection, Zentralbl. Bakteriol. Suppl. 10, G. Fischer Verlag, Stuttgard, New York.

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L i s i e c k i P. and J. M i k u c k i. 2005. Assimilatory ferric reductases in enterococci (in Polish). Med. Doœw. Mikrobiol. 57: 359–368. L o w r y H., N.J. R o s e b r o u g h, A. L e w i s F a r r and R.J. R a n d a l l. 1951. Protein measurment with the Folin phenol reagent. J. Biol. Chem. 193: 265–275. M a z o y R. and M. L e m o s. 1999. Ferric-reductase activities in Vibrio vulnificus biotype 1 and 2. FEMS Microbiol. Lett. 172: 205–211. R a t l e d g e C. and L.G. D o v e r. 2000. Iron metabolism in pathogenic bacteria. Annu. Rev. Microbiol. 54: 881–941. S c h r ö d e r I., E. J o h n s o n and S. d e V r i e s. 2003. Microbial ferric iron reductases. FEMS Microbiol. Rev. 27: 427–447. S i l v e r S. and M. W a l d e r h a u g. 1992. Gene regulation of plasmid – and chromosome-determined inorganic ion transport in bacteria. Microbiol. Rev. 56: 195–228. S t o o k e y L.L. 1970. Ferrozine – a new spectrophotometric reagent for iron. Anal. Biochem. 42: 779–781. V a r t i v a r i a n S.E. and R. C o w a r t. 1999. Extracellular iron reductases: identification of a new class of enzymes by siderophoreproducing microorganisms. Arch. Biochem. Biophys. 364: 75–82. Z o r z i W., X.Y. Z h o u, O. D a r d e u n e, J. L a m o t t e, D. R a z e, J. P i e r r e, L. G u t m a n n and J. C o y e t t e. 1996. Structure of the low-affinity Penicillin-Binding Protein 5 PBP fm in wild-type and highly penicillin-resistant strains of Enterococcus faecium. J. Bacteriol. 178: 4948–4957.

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Polish Journal of Microbiology 2006, Vol. 55, No 4, 279– 288

Susceptibility of Listeria monocytogenes Strains Isolated from Dairy Products and Frozen Vegetables to Antibiotics Inhibiting Murein Synthesis and to Disinfectants MAGDALENA POPOWSKA* MARTA OLSZAK and ZDZIS£AW MARKIEWICZ

Department of General Microbiology, Institute of Microbiology, Warsaw University, Warsaw, Poland Received 17 September 2006, accepted 12 October 2006 Abstract The susceptibility of 96 strains of Listeria monocytogenes isolated from food to antibiotics and disinfectants currently used in human therapy, veterinary, medicine and food industry was determined by a standard operating procedure – broth dilution method. Antimicrobial agents included the $-lactams ampicillin and penicillin, the lantibiotic nisin, and the disinfectants benzalkonium chloride and chlorhexidine gluconate. Among the studied strains we found 13 strains with 8-fold, 7 strains with 16-fold and 2 strains with 32-fold decreased susceptibility to ampicillin, as determined by MIC, compared to wild type reference strain. Interestingly, the mentioned strains were isolated from frozen vegetables and soups, none of the isolates from dairy products showed any elevated resistance to the studied antimicrobial agents. The occurrence in food products of strains with increased resistance to ampicillin is disquieting, especially since $-lactams are the most frequent antibiotic of choice in the therapy of infections caused by the pathogen. K e y w o r d s: Listeria monocytogenes, food-borne pathogens, antibiotic and disinfectant resistance, autolysis, autolysins

Introduction Listeria monocytogenes (Hamon et al., 2006) is an opportunistic Gram-positive bacterium ubiquitous in nature: it occurs in waters, soil, rotting vegetation, animal faeces and wastewaters and is also isolated from faecal samples from healthy individuals – the bacterium has been found to be present in 5% of studied adults (Farber and Peterkin, 1991). L. monocytogenes is also found in such food products as fresh vegetables, milk, fish, poultry and meat. The spread of listeriae is related to the use of wastewaters for irrigation, which results in the presence of the bacteria on plants and products of animal origin. L. monocytogenes is an intracellular pathogen of humans and animals and is the causative agent of listerioses in the form of sporadic and epidemic infections (Portnoy et al., 2002; Hamon et al., 2006). Thirteen serotypes of L. monocytogenes have been identified and on average 90% of clinical infections are caused by serotypes Ia, Ib and IVb, the latter dominating in Europe (Schlech, 2000). However, a recent study embracing L. monocytogenes strains isolated from various sources in Poland has shown that the most prevalent is serotype IIa (Paciorek et al., 2006). The group at greatest risk for L. monocytogenes infection includes pregnant women, newborns and immunocompromised individuals, resulting in death in 25– 30% of cases (Hamon et al., 2006). The therapy of choice in the case of listeriosis involves a $-lactam, such as penicillin or ampicillin alone or in combination with an aminoglycoside, usually gentamicin (Charpentier and Courvalin, 1999). Alternative treatments, especially in the case of sensitivity to penicillin, involve trimethoprim in combination with a sulfonamide, such as sulfamethoxazole (Hof, 2004). L. monocytogenes is naturally susceptible to penicillins, amino penicillins, carboxypenicillins, ureidopenicillins, carbapenems (e.g. imipenem) (Espaze and Reynaud, 1988; Troxler et al., 2000) but is relatively resistant to monobactams (aztreonam) and certain third generation cephalosporins, e.g. cefotaxim and ceftizoxim (Espaze and Reynaud, 1988). Poulsen et al. (1988) reported that 156 strains of L. monocytogenes * Corresponding author: M. Popowska, Department of General Microbiology, Institute of Microbiology, Warsaw University, 02-096 Warsaw, Miecznikowa 1, Poland; phone + 48 22 55 41 320; e-mail: [email protected]

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from humans during 27 years (1958–1985) were susceptible to 12 antibiotics. The first case of a clinical strain of L. monocytogenes resistant to ampicillin was described in 1984 (Rapp et al., 1984; Pollock et al., 1986). More recent papers describe L. monocytogenes strains showing multiple resistance, mainly to aminoglycosides, chloramphenicol (Facinell et al., 1991; Tsakris et al., 1997), tetracycline and penicillin (Walsh et al., 2001; Poyart-Salmeron et al., 1992), erythromycin (Roberts et al., 1996), as well as trimethoprim (Charpentier et al., 1995), sulfamethoxazole and rifampin (Abrahim et al., 1998). The occurrence of resistant strains is usually related to the acquisition of transposons or plasmids carrying resistance determinants and transfer occurs mainly in the digestive tracts of animals and humans, frequently Enterococcus spp. and Streptococcus spp. (Charpentier and Courvalin, 1999; Charpentier et al., 1995). The transfer of vancomycin resistance from enterococci to L. monocytogenes by conjugation under laboratory conditions has also been shown (Biavasco et al., 1996). Recently 2 cases of listeriosis caused by penicillin-resistant L. monocytogenes and 6 by strains resistant to ampicillin were described but these strains, like the ampicillinresistant one mentioned above, were never characterized (Safdar and Armstrong, 2003). The resistance of some L. monocytogenes isolates to popularly used disinfectants, especially chlorine compounds such as chlorhexidine and benzalkonium chloride, has been described. The mechanism of adaptation or resistance to benzalkonium chloride is associated with efflux pumps (Soumet et al., 2005). Two efflux pumps have been described in L. monocytogenes: MdrL can extrude antibiotics (macrolides and cefotaxime), heavy metals and ethidium bromide (Mata et al., 2000) and Lde is associated with fluoroquinolone resistance and partly responsible for resistance to acridine orange and ethidium bromide (Godreuil et al., 2003). A recent study has shown that MdrL is at least partly responsible for adaptation to benzalkonium chloride (Romanova et al., 2006). The aim of the present study was to determine the susceptibility to ampicillin, penicillin, nisin, benzalkonium chloride and chlorhexidine of 96 food isolates of L. monocytogenes. All these agents are or have been used in medicine, veterinary medicine and the food industry. We have also attempted to elucidate the reason for the reduced susceptibility of several of the studied strains to ampicillin. Experimental Material and Methods Bacterial strains. We studied 96 strains of L. monocytogenes isolated from frozen food (vegetables and soups) collected by Sanepid in Bydgoszcz (WSSE, Bydgoszcz). All strains were classified as L. monocytogenes according to the Polish norm PN-EN ISO 11290-1:1999 and further characterized in our Department by streaks on Palcam and OCLA agar plates (Oxoid). We also did a haemolysis test on TSYEB agar plates (Tryptone Soy Agar slants with 3% yeast extract; BioMérieux; BTL) containing 5% (v/v) sheep blood. The reference strain used in these studies was L. monocytogenes EGD. Bacteria were grown in TSYEB at 37°C with mild shaking (120 rpm) to mid-exponential phase of growth. Strains were stored at 4°C on TSYEB agar plates. Antimicrobial agents. The antibiotics tested were: ampicillin, penicillin, nisin (Sigma) and the disinfectants: benzalkonium chloride (alkyl dimethyl benzyl ammonium chloride) and chlorhexidine gluconate (Sigma). Antimicrobial stock solutions were prepared in distilled water and stored at –20°C. Broth dilution method for minimal inhibitory concentration (MIC) determinations. MICs of antimicrobial agents for food isolates were determined by broth microdilution method of the National Committee for Clinical Laboratory Standards (NCCLS, 2003), currently the Clinical and Laboratory Standards Institute (CSLI). The final concentrations of the antibiotics were 0.0468 – 12 µg/ml and for nisin 1 – 40 µ/ml. Each MIC determination was repeated two times. Isolation of cell wall. L. monocytogenes cells were harvested by centrifugation (6 000 × g, 15 min at 4°C) and resuspended in 1/40 of the original culture volume of ice-cold saline. Glass beads (diameter 150 – 215 mm; Sigma) were added (1 g per 1 ml cell suspension) and ten 1 minute bursts of ultrasound waves were employed in VCX-600 ultrasonicator (Sonics and Materials, USA) at amplitude 20%. The crude cell wall preparation was sedimented by centrifugation in a Beckman centrifuge (25 min, 100 000 × g at 4°C). The cells were washed in appropriate buffer and then resuspended in it. Autolysis of crude cell wall preparations from L. monocytogenes. After sonication as above, the cell walls were sedimented by centrifugation, washed in 10 or 50 mM Tris-HCl buffer and resuspended in the same buffer pre-warmed to 37oC. The suspension was incubated with shaking at 37 oC and changes in absorbance were followed at 600 nm. Autolysis of cells. Cultures of L. monocytogenes strains were grown to early exponential phase (OD 600 0.20) in TSB broth. To determine the effect of the antibiotic on growth, 10 times the MIC of penicillin for each of the individual strains, or 0.5 times the MIC of nisin was added, and the response of the culture was followed spectrophotometrically (Novaspec II spectrophotometer LKB-13 Pharmacia) while continuing incubation at 37°C with shaking. To determine the effect of Triton X-100 or SDS, the cells were grown to OD600 ~ 0.6, harvested and resuspended in 50 mM Tris-HCl (pH 7.5), 0.1% (v/v) Triton X-100 or 1% (v/v) SDS. Lysis of the cell suspension at 37°C was followed spectrophotometrically at 600 nm. Induced cell lysis. Cultures of L. monocytogenes strains were grown to mid-exponential phase (OD 600 0.80) in TSB broth. Cells were harvested by centrifugation as above and resuspended in 50 mM Tris-HCl (pH 7.5) buffer pre-warmed to 37°C. To determine the effect of muramidase-induced lysis lysosyme or cellosyl was added to final concentration 2 mg/ml, and the lysis of the culture was followed spectrophotometrically while continuing incubation at 37°C with shaking.

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Extraction of surface-associated autolytic enzymes. Cells of L. monocytogenes were harvested by centrifugation, washed in saline and resuspended in an ice-cold solution containing 4 M LiCl and 0.5 mM phenylmethylsulfonylfluoride (PMSF) in 50 mM Tris-HCl buffer, pH 7.0. The suspension was stirred for 30 min with the use of a magnetic bar in an ice bath, after which the cells were harvested (20 000 × g, 10 min at 4°C). The supernatant was transferred to dialysis tubing with cutoff value 3 500 (Spectrum) and dialyzed for 12 hours at 4°C against 50 mM Tris-HCl, pH 7.0 containing 100 mM LiCl, with several changes of the buffer. Alternatively, cells of L. monocytogenes were extracted with 1% (v/v) SDS at room temperature for 5 min, after which the cells were harvested (20 000 × g, 10 min at 4°C). The supernatant was analyzed on 12% SDS-polyacrylamide gel and renaturing gel. Haemolytic activities. Bacteria were cultured on sheep blood agar at 37°C for 36 h. L. innocua and L. ivanovii were used as a negative and positive control, respectively. After incubation, the narrow ring of $-hemolysis produced by the strains was compared with that of L. monocytogenes EGD. Plasmid DNA isolation. Plasmid DNA from E. coli was isolated and purified with the Plasmid Miniprep Plus kit (A&A Biotechnology). The procedures for the isolation of plasmid and chromosomal DNA from L. monocytogenes were performed as previously described (McLaughlan and Foster, 1998), starting with digestion of the bacterial cell wall in 5–10 mg/ml lysozymecontaining GTE buffer for 1 h at 37°C. Penicillin binding protein (PBP) assay. The PBP pattern of L. monocytogenes was visualized by fluorography following electrophoresis of cell membrane proteins incubated with [H 3]-benzylpenicillin, as described in detail elsewhere (Korsak et al., 2002).

Results Antimicrobial resistance. We analysed 96 strains of L. monocytogenes isolated from frozen meal and dairy products study for susceptibility to antimicrobial agents. Eighteen of the strains were just as sensitive to ampicillin as the reference strain (EGD) used (MIC 0.156 µg/ml), 14 were more sensitive to the antibiotic (0.039–0.078 µg/ml) and 65 of the strains showed reduced susceptibility to ampicillin (0.3125–5.0 µg/ml). Two of the latter strains were resistant to 5 µg ampicillin/ml, which is 32 times the MIC for EGD. None of the isolated strains showed any significantly altered susceptibility to nisin, the MIC values for the antibiotic against the isolates being similar as for EGD. Twelve of the studied strains were as susceptible to chlorhexidine as EGD (MIC 1.25 µg/ml) whereas all the remaining strains had two to 4-fold reduced susceptibility to the disinfectant (MIC 2.5– 5 µg/ml). In the case of benzalkonium chloride, the sensitivity of 37 of the strains to the compound was the same as that of EGD (MIC 1.25 µg/ml). Only 7 of the strains were two to 4-fold less sensitive (MIC 2.5– 5 µg/ml) (Table I). For further, more detailed analysis, six of the tested strains (Table II) were selected. Worth note is that strain was 11 times less sensitive to penicillin than EGD, 16 times less susceptible to ampicillin and 2 times less susceptible to both studied disinfectants. Similar values were obtained for strain 21 which, however, it was not more resistant to penicillin. Literature data indicate that one of the main reasons for the resistance of L. monocytogenes to antibiotics is the presence of mobile genetic elements carrying resistance genes. Multiple resistance of L. monocytogenes has been shown to be related to the presence of specific genes carried by plasmids pIP811, pUBX1 and pWDB100, whereas resistance to trimethoprim is determined by dihydrofolate reductase gene (dfrD), present in the 3.7 kb broad host range plasmid pIP823 (Charpentier and Courvalin, 1999). In this study we attempted to identify putative genetic elements that could carry genes responsible for the observed elevated resistance to ampicillin. L. monocytogenes has recently been shown to carry a chromosomal Table I Susceptibility of L. monocytogenes isolates to ampicillin and disinfectants MIC (µg/ml)

Number of strains

MIC (µg/ml)

Ampicillin

Number of strains

Chlorhexidine

0.039

11

1.25

12

0.078

3

2.5

43

0.156

18

5.0

7

0.3125

18

0.625

25

1.25

37

1.25

13

2.5

6

2.5

7

5.0

1

5.0

2

Benzalkonium chloride

282

4

Popowska M. et al. Table II Antimicrobial susceptibility of selected strains of L. monocytogenes MIC (µg/ml)

Strain designation

Origin

ampicillin

penicillin G

chlorhexidine

benzalkonium chloride

nisin

1

cauliflower

2.5

0.7

5.0

2.5

9

6

vegetables with rice

5.0

0.12

2.5

2.5

10

21

vegetables soup

2.5

0.06

5.0

5.0

11

58

green peas

5.0

0.12

2.5

2.5

10

105

dairy product

0.156

0.06

2.5

1.25

10

121

dairy product

0.156

0.03

2.5

1.25

11

EGD

collection of Institute of Microbiology

0.156

0.06

2.5

1.25

11

gene Lmo0540 that may code a putative class C $-lactamase (Guinane et al., 2006) but we have never been able to demonstrate any-lactamase activity in this bacterium using a number of different approaches (PoroœG³uchowska, 2004). Similarly, to our knowledge no chromosomally coded L. monocytogenes $-lactamase activity has been described by any other laboratory. Subsequently, we used a specific procedure for the isolation of plasmid DNA from L. monocytogenes strains, involving a phenol step, followed by electrophoresis in 1% agarose gel. In spite of repeated attempts in no case any plasmid preparations were obtained from the selected L. monocytogenes strains. In the next stage we analysed the PBP pattern of the studied strains in order to see if any quantitative or qualitative differences explaining the increased resistance to ampicillin could be observed. No differences were observed (data not presented). In turn, we decided to carry out a physiological analysis aimed to demonstrate changes in the protein composition of the cell wall and/or muropeptide composition of the cell A

90 80

OD600 (%)

OD600 (%)

70 60 50 40 30 20 10 0 0

30

60

90

EGD

1

120 21

150 105

180 121

210 Time (min)

B 180 160

OD600 (%)

OD600 (%)

140 120 100 80 60 40 20 0 0

30

60

90 EGD

120

150

1

21

180 105

210

240 121

270

300

Time (min)

Fig. 1. Effect of antibiotics at 10 times the MIC on the growth of L. monocytogenes EGD and selected isolates A) penicillin; B) nisin

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A 120

% of OD600

% of OD600

100 80 60 40 20 0 0

30

60

90

120 150 180 210 240 270 300 330 360 390 420 450 EGD

B

1

21

90

120

105

121

Time (min)

120

% of OD600

% of OD600

100 80 60 40 20 0 0

30 EGD

60 1

21

105

150

180

121 Time (min)

Fig. 2. Autolysis of cells of L. monocytogenes EGD and selected strains induced by detergents A) Triton X-100; B) SDS

wall murein, as possible determinants for the observed resistance to $-lactams, i.e. impeding the penetration of the antibiotic to the cytoplasmic membrane. Autolysis of whole cells and isolated cell walls. A significant effect of penicillin G on the growth of the studied strains was observed – all the isolates were less susceptible compared to EGD at ten times the MIC for the individual strains, and this was very evident already after 60 min of the experiment. Strain 1 grew faster by about 37% and the remaining strains by about 23% (Fig. 1A). Similarly, all the studied isolates demonstrated strongly reduced sensitivity to nisin at 0.5 times the MIC compared to strain EGD. The growth rate of strain 21, for which the MIC of nisin was equal to that of the reference strain, was faster by about 74% than EGD (Fig. 1B). All the strains were also slightly less susceptible to the action of Triton X-100, compared to strain EGD, but the rate of their autolysis was similar. The least susceptible to the action of the detergent were strains 21 and 121, which autolysed slower by 10% and 17% after 450 min, respectively, compared to EGD (Fig. 2A). When SDS was used all the studied strains were less susceptible to the action of the detergent than the wildtype strain. The autolysis of strain 121 was about 30% slower, and that of the remaining strains by about 25% slower than observed for EGD (Fig. 2B). Murein isolated from the strains 1, 21 and 121 showed faster autolysis kinetics than for that from strain EGD – in the case of strain 1 by as much as 24%. A similar tendency was observed for murein from strains 21 and 121, which lysed 8% faster than murein from strain EGD. Only murein from strain 105 showed a slightly slower rate of autolysis (by about 4%) (Fig. 3). Cell lysis. The cells of all the studied strains lysed much faster in the presence of hen egg white lysozyme than EGD. The rate was faster by 12% for strain 121 and by 7% for strain 1 (Fig. 4A). Lysis of whole cells by cellosyl was somewhat faster (by 4 to 16%) for three of the strains, but strain 21 was lysed slower by about 6 % compared to the wild type EGD (Fig. 4B). Analysis of surface protein composition. Analysis of the protein fraction isolated with the use of SDS suggests similar protein composition of strain 1 and the reference strain EGD, as well as between the pairs of strains 58 and 105, and 6 and 21 (Fig. 5). The profiles of the reference strain and strain 1 lack a protein

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with mass about 60 kDa (designation c, Fig. 5) and there is also much less of a protein (or two proteins with similar mass) located between 20.1 kDa and 14.4 kDa (designation e, Fig. 5). The surface protein fraction of strains 58 and 105 contains a clearly visible protein with mass approx. 94 kDa (designation a, Fig. 5), this protein being produced by the remaining strains in much smaller copy number. Strains 6 and 21 are characterized by a decidedly different profile, which lacks a protein between 94 kDa, a 67 kDa (designation b, Fig. 5) present in the remaining strains as well as two proteins between 67 kDa and 43 kDa, that are visible in strains 1, 58 and 105 (designation d, Fig. 5). Analysis of the surface fraction isolated using LiCl shows multi-point similarity in the protein composition for the reference strain and strain 1, as opposed to strains 6, 21, 58 and 105 (Fig. 6). Three proteins that were A

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present only in the reference strain and strain 1, located between 94 kDa, and 67 kDa were observed (designation a, b, c, Fig. 6) as well as two additional ones in the protein profile of the above strains – approx. 65 kDa (designation d, Fig. 6) and approx. 23 kDa (designation g, Fig. 6). Significant quantitative and/or qualitative differences were also observed among proteins in the ca. 60 kDa to ca. 40 kDa range (designation e, Fig. 6). An additional protein with mass ca. 38 kDa, or its increased amount compared to the reference strain, that was not present in strain 1, was observed in the case of strains 6, 21, 58 and 105 (designation f, Fig. 6). Discussion An undesirable consequence of the use, frequently unwarranted, of antimicrobials is the occurrence and spread of strains of pathogenic bacteria resistant to antibiotics and/or disinfectants in the environment. This problem is particularly challenging in the case of food-borne pathogens such as L. monocytogenes, which accounts for 28% of food-related deaths in the USA (Mead et al., 1999). The incidence of listeriosis has increased over the past two decades throughout the world. Several large food-borne outbreaks of listeriosis

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have been reported in numerous countries, including England, Germany, Sweden, New Zealand, Switzerland, Australia, France, and the US. L. monocytogenes strains resistant to antibiotics were reported in the eighties (e.g. Rapp et al., 1984; Poulsen et al., 1988). Since then, Listeria spp. isolated from food, the environment or in sporadic cases of human listeriosis have been shown to be resistant to one or several antibiotics and the number of reports of isolates of L. monocytogenes resistant to $-lactam antibiotics, the antibiotics of choice for the treatment of listeriosis, is slowly but systematically growing (Rapp et al., 1984; Walsh et al., 2001; Chen et al., 2001; Prazak et al., 2002; Safdar and Armstrong, 2003). The data published by Prazak et al. (2002) indicate that 87% of multiply resistant strains were resistant to penicillin and some of these in addition, also to gentamicin, the second antibiotic used together with a $-lactam in combined chemotherapy of listeriosis. An equally disturbing phenomenon is the adaptation or resistance of L. monocytogenes to benzalkonium chloride and other disinfectants (Romanova et al., 2006). The data for Poland are scarce, this being due, among others, to infrequent screening for the occurrence of L. monocytogenes. A fairly recent study involving 73 strains of L. monocytogenes isolated from a variety of clinical and environmental sources screened for resistance to ampicillin, penicillin, gentamicin, erythromycin, clarithromycin, sulfisoxazole and trimethoprim found that the strains were susceptible to the antibiotics studied, except for the resistance of 30.1% of the strains to sulfisoxazole (Paciorek, 2004). In our earlier studies, we showed that L. monocytogenes can readily develop resistance to $-lactams under antibiotic pressure as well as under transposon mutagenesis conditions (Poroœ-G³uchowska and Markiewicz, 2003a; 2003b). The pathogen is capable of acquiring antibiotic resistance genes from enterococci and streptococci through movable genetic elements such as transposons and plasmids and resistance genes also come from other Gram-positive and Gram-negative bacteria through conjugative mobilization (de Niederhausern et al., 2004; Bertrand et al., 2005). In this study, a collection of 96 L. monocytogenes strains isolated from dairy products and frozen foods was screened for susceptibility to commonly used antibiotics and biocides. Analysis of the growth rate of the strains isolated from food products at 37°C in TSB did not indicate any significant differences compared to EGD, in spite of their being subjected to thermal processing (pasteurization, freezing). This seems to indicate that temperature variations were not a factor contributing towards observed resistance (Asselt and Zwietering 2006). Of the strains studied, 68% showed reduced susceptibility to ampicillin, and for 2 of these the MIC of ampicillin was approx. 32-fold higher than for strain EGD. Interestingly, the resistance of these strains to penicillin was in a much lower range, from 2 to 12-fold the MIC for EGD. In our in vitro attempts to generate ampicillin resistant derivatives of L. monocytogenes EGD mentioned above (Poroœ-G³uchowska and Markiewicz, 2003a; 2003b), the highest value was similar to that observed for two of the studied isolates, that is 32-fold. Attempts to identify the cause of ampicillin resistance of the food-borne strains were negative and, as mentioned in the introduction, no resistance mechanism was identified for any of the other $-lactam resistant isolates mentioned in the literature. An interesting observation was the behaviour of the isolates showing lower susceptibility to penicillin to the presence of the antibiotic at 10 times the individual MICs. Penicillin is well known to kill L. monocytogenes after a lag phase of several hours, without accompanying lysis (Chen et al., 1996; Popowska et al., 1999). This is reflected as a plateau in the growth curve after the addition of penicillin at concentrations above the MIC. The isolates, however, continued to grow, as is clearly visible in Fig. 1A. Nisin does not affect the growth of L. monocytogenes the way penicillin does, but nevertheless in this case the isolates grew much faster at 0.5 times the MIC for the individual strains than the reference strain during the time of the experiment. These observations demand further investigation. In the case of benzalkonium chloride 16% of the strains were characterized by from 2 to 4-fold reduced susceptibility, whereas as many as 84% of the isolates were equally susceptible to the compound as EGD. The ratio was practically quite the opposite for chlorhexidine in which case 82% of the studied strains showed 2 to 4-fold reduced susceptibility to the disinfectant whereas 18% were just as susceptible to the biocide as EGD. It is assumed that reduced susceptibility to disinfectants can be a result of altered permeability of the bacterial cell wall, e.g. structural modifications, thickness, or a consequence of the action of active pumps (e.g. MdrL), which detoxify the cell (Mata et al., 2000; Romanova et al., 2006). Since we ruled out $-lactamase activity and changes in the PBPs of L. monocytogenes as the reasons for the resistance of some of the isolated strains to ampicillin and penicillin, and it is known that changes in cell wall permeability may affect the resistance of some bacteria to antibiotics (e.g. Sieradzki and Markiewicz, 2004), we decided to examine the cell wall murein of the mutants. A comparison of the muropeptide profiles of the mutants obtained after HPLC chromatography of a muramidase digest of murein did not reveal any significant changes in the murein of the strains showing

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elevated resistance to $-lactams. There were some minor changes in the murein of the strains with reduced susceptibility to the disinfectants, but in view of only two-fold reduction in the susceptibility of the strains to the compounds, these observations were not followed up. Differences were, however, observed when crude murein preparations from the cells were subjected to conditions promoting autolysis. Murein from strain 105 reproducibly autolysed slower than that from EGD, whereas murein from strains 1, 21 and 121 autolysed much faster than murein from the reference strain. These results may point to differences in the modification of murein, such as extent of N-acetylation of aminosugars or the presence or absence of O-acetyl groups which may affect the activity of the autolytic enzymes in the cell wall (Popowska, 2004). Alternatively, the studied strains may lack certain autolytic enzymes or the enzymes in question may be less or more stringently regulated. These conclusions are supported by the differences in the kinetics of lysis of intact L. monocytogenes cells induced by Triton X-100 and SDS. Similar differences were also observed when L. monocytogenes cells were treated with egg white lysozyme or the muramidase cellosyl. Both enzymes not only hydrolyze bonds in murein but also induce the activity of autolysins, especially lysozyme, which is a protein with cationic nature (Leitch and Willcox, 1999). Altogether, in the light of these observations further experiments are being planned. The greatest differences between the strains were found when the surface protein profiles of the cells were compared, regardless of extraction method used. Compared to EGD, the studied isolates either had new bands not observed in the reference strains or lacked certain proteins. These differences may reflect, for instance, the absence of certain autolytic activities or the presence of proteins whose synthesis is induced under temperature stress conditions and which may determine altered permeability of the cell wall. Much remains to be elucidated, but the main conclusion of these studies is that strains of L. monocytogenes resistant to $-lactam antibiotics are becoming increasingly more ubiquitous in the environment, especially in food products. The increasing incidence of antibiotic resistance in L. monocytogenes, in combination with the widespread use of antibiotics in medicine and various branches of industry may have significant future clinical implications for the treatment of listeriosis. Literature A b r a h i m A., A. P a p a, N. S o u l t o s, I. A m b r o s i a d i s and A. A n t o n i a d i s. 1998. Antibiotic resistance of Salmonella spp. and Listeria spp. isolates from traditionally made fresh sausages in Greece. J. Food Prot. 61: 1378–380 A s s e l t E.D. and M.H. Z w i e t e r i n g. 2006. A systematic approach to determine global thermal inactivation parameters for various food pathogens. Int. J. Food. Microbiol. 107: 73–82. B e r t r a n d S., G. H u y s, M. Y d e, K. D’ H a e n e, F. T a r d y, M. V r i n t s, J.K. S w i n g s and J.M. C o l l a r d. 2005. Detection and characterization of tet(M) in tetracycline-resistant Listeria strains from human and food-processing origins in Belgium and France. J. Med. Microbiol. 54: 1151–1156. B i a v a s c o F., E. G i o v a n e t t i, A. M i e l e, C. V i g n a r o l i, B. F a c i n e l l i and P.E. Va r a l d o. 1996. In vitro conjugative transfer of VanA vancomycin resistance between Enterococci and Listeriae of different species. Eur. J. Clin. Microbiol. Infect. Dis. 15: 50–59. C h a r p e n t i e r E. and P. C o u r v a l i n. 1999. Antibiotic resistance in Listeria spp. Antimicrob. Agents Chemother. 43: 2103–2108. C h a r p e n t i e r E., G. G e r b a u d, C. J a c q u e t, J. R o c o u r t and P. C o u r v a l i n. 1995. Incidence of antibiotic resistance in Listeria species. J. Infect. Dis. 172: 277–281 C h e n L.J., J. W a n g and R.E. L e v i n. 1996. Effect of benzylpenicillin on the viability and osmotic sensitivity of Listeria monocytogenes. Lett. Appl. Microbiol. 22: 10–12. E s p a z e E.P. and A.E. R e y n a u d. 1988. Antibiotic susceptibilities of Listeria: in vitro studies. Infection 16 Suppl 2: S160–164. F a c i n e l l i B., E. G i o v a n e t t i, P.E. V a r a l d o, P. C a s o l a r i and U. F a b i o. 1991. Antibiotic resistance in foodborne Listeria. Lancet 338: 1272. F a r b e r J.M. and P.I. P e t e r k i n. 1991. Listeria monocytogenes, a food-borne pathogen. Microbiol. Rev. 55: 476–511. G o d r e u i l S., M. G a l i m a n d, G. G e r b a u d, C. J a c q u e t and P. C o u r v a l i n. 2003. Efflux pump Lde is associated with fluoroquinolone resistance in Listeria monocytogenes. Antimicrob. Agents Chemother. 47: 704–708. H a m o n M., H. B i e r n e and P. C o s s a r t. 2006. Listeria monocytogenes: a multifaceted model. Nat. Rev. Microbiol. 4: 423–434. G u i n a n e C.M., P.D. C o t t e r, R.P. R o s s and C. H i l l. 2006. Contribution of penicillin-binding protein homologs to antibiotic resistance, cell morphology, and virulence of Listeria monocytogenes EGDe. Antimicrob. Agents Chemother. 50: 2824–2828 H o f H. 2004. An update on the medical management of listeriosis. Expert Opin. Pharmacother. 5: 1727–1735. K o r s a k D., J.J. Z a w a d z k a, M.E. Œ i w i ñ s k a and Z. M a r k i e w i c z. 2002. Penicillin-binding proteins of Listeria monocytogenes – a re-evaluation. Acta Microbiol. Pol. 51: 5–12. L e i t c h E.C. and M.D. W i l l c o x. 1999. Elucidation of the antistaphylococcal action of lactoferrin and lysozyme. J. Med. Microbiol. 48: 867–871. M a t a M.T., F. B a q u e r o and J.C. P e r e z - D i a z. 2000. A multidrug efflux transporter in Listeria monocytogenes. FEMS Microbiol. Lett. 187: 185–188.

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M c L a u g h l a n A.M. and S.J. F o s t e r. 1998. Molecular characterization of an autolytic amidase of Listeria monocytogenes EGD. Microbiology 144: 1359–1367. M e a d P.S., L. S l u t s k e r, V. D i e t z, L.F. M c C a i g, J.S. B r e s e e, C. S h a p i r o, P.M. G r i f f i n and R.V. T a u x e. 1999. Food-related illness and death in the United States. Emerg. Infect. Dis. 5: 607–625. N C C L S. 2003. Methods for dilution antimicrobial susceptibility testing for bacteria that grow aerobically; approved standard – sixth edition. NCCLS document M7-A6. NCCLS, Wayne, Pennsylvania. d e N i e d e r h a u s e r n S., C. S a b i a, P. M e s s i, E. G u e r r i e r i, G. M a n i c a r d i and M. B o n d i. 2004. Glycopeptideresistance transferability from vancomycin-resistant enterococci of human and animal source to Listeria spp. Lett. Appl. Microbiol. 39: 483–489. P a c i o r e k J. 2004. Antimicrobial susceptibilities of Listeria monocytogenes strains isolated from 2000 to 2002 in Poland. Pol. J. Microbiol. 53:279–281. P a c i o r e k J., C. J a c q u e t, C. S a l c e d o, M. D o u m i t h, J.A. V a z q u e z and P. M a r t i n. 2006. Genotypes of Listeria monocytogenes strains isolated from 2000 to 2002 in Poland. Pol. J. Microbiol. 55: 31–35 P o l l o c k S.S., T.M. P o l l o c k and M.J. H a r r i s o n. 1986. Ampicillin-resistant Listeria monocytogenes meningitis. Arch. Neurol. 43: 106–107 P o p o w s k a M., M. K l o s z e w s k a, S. G ó r e c k a and Z. M a r k i e w i c z. 1999. Autolysis of Listeria monocytogenes. Acta. Microbiol. Pol. 48: 141–52. Popowska M. 2004. Analysis of the peptidoglycan hydrolases of Listeria monocytogenes: multiple enzymes with multiple functions. Pol. J. Microbiol. 53: Suppl: 29–34. P o r o œ - G ³ u c h o w s k a J. and Z. M a r k i e w i c z. 2003a. Antimicrobial resistance of Listeria monocytogenes. Acta Microbiol. Pol. 52: 113–29. P o r o œ - G ³ u c h o w s k a J. and Z. M a r k i e w i c z. 2003b. Ampicillin resistance in Listeria monocytogenes acquired as a result of transposon mutagenesis. Acta Microbiol. Pol. 52: 131–42. P o r o œ - G ³ u c h o w s k a J. 2004. PhD Thesis. Warsaw University P o r t n o y D.A., V. A u e r b u c h and I.J. G l o m s k i. 2002. The cell biology of Listeria monocytogenes infection: the intersection of bacterial pathogenesis and cell-mediated immunity. J. Cell Biol. 3:409–414. P o u l s e n P.N., A. C a r v a j a l, A. L e s t e r and J. A n d r e a s e n. 1988. In vitro susceptibility of Listeria monocytogenes isolated from human blood and cerebrospinal fluid. A material from the years 1958–1985, APMIS 96: 223–228. P o y a r t - S a l m e r o n C., P. T r i e u - C u o t, C. C a r l i e r, A. M a c G o w a n, J. M c L a u c h l i n and P. C o u r v a l i n. 1992. Genetic basis of tetracycline resistance in clinical isolates of Listeria monocytogenes. Antimicrob. Agents Chemother. 36: 463–466. P r a z a k M.A., E.A. M u r a n o, I. M e r c a d o and G.R. A c u f f. 2002. Antimicrobial resistance of Listeria monocytogenes isolated from various cabbage farms and packing sheds in Texas. J. Food Prot. 65: 1796–1799 R a p p M.F., H.A. P e r s h a d s i n g h, J.W. L o n g and J.M. P i c k e n s. 1984. Ampicillin-resistant Listeria monocytogenes meningitis in a previously healthy 14-year-old athlete. Arch. Neurol. 41: 1304. R o b e r t s M.C., B. F a c i n e l l i, E. G i o v a n e t t i, P.E. V a r a l d o. 1996. Transferable erythromycin resistance in Listeria spp. isolated from food. Appl. Environ. Microbiol. 62: 269–270. R o m a n o v a N.A., P.F. W o l f f s, L.Y. B r o v k o and M.W. G r i f f i t h s. 2006. Role of efflux pumps in adaptation and resistance of Listeria monocytogenes to benzalkonium chloride. Appl. Environ. Microbiol. 72: 3498–503. S a f d a r A. and D. A r m s t r o n g. 2003. Antimicrobial activities against 84 Listeria monocytogenes isolates from patients with systemic listeriosis at a comprehensive cancer center (1955–1997). J. Clin. Microbiol. 41: 483–485. S c h l e c h W.F. 2000. Foodborne listeriosis. Clin. Infect. Dis. 31: 770–775. S i e r a d z k i K. and Z. M a r k i e w i c z. 2004. Mechanism of vancomycin resistance in methicillin resistant Staphylococcus aureus. Pol. J. Microbiol. 53: 207–214. S o u m e t C., C. R a g i m b e a u and P. M a r i s. 2005. Screening of benzalkonium chloride resistance in Listeria monocytogenes strains isolated during cold smoked fish production. Lett. Appl. Microbiol. 41: 291–296. T r o x l e r R., A. v o n G r a e v e n i t z, G. F u n k e, B. W i e d e m a n n and I. S t o c k. 2000. Natural antibiotic susceptibility of Listeria species: L. grayi, L. innocua, L. ivanovii, L. monocytogenes, L. seeligeri and L. welshimeri strains. Clin. Microbiol. Infect. 6: 525–535. T s a k r i s A., J. D o u b o y a s and L.S. 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Partial Characterization and Optimization of Production of Extracellular "-amylase from Bacillus subtilis Isolated from Culturable Cow Dung Microflora MANAS R. SWAIN1, SHAKTIMAY KAR1, GOURIKUTTI PADMAJA 2 and RAMESH C. RAY1* 1 Regional

Center of Central Tuber Crops Research Institute, Bhubaneswar, India Tuber Crops Research Institute, Sreekarium, Trivandrum, India

2 Central

Received 27 June 2006, received 4 September 2006, accepted 15 September 2006 Abstract Studies of "-amylase production by Bacillus subtilis (CM3) isolated earlier from cow dung microflora, were carried out. The optimum temperature, pH and incubation period for amylase production were 50–70°C, 5.0–9.0 and 36 h, respectively. Enzyme secretion was very similar in the presence of any of the carbon sources tested (soluble starch, potato starch, cassava starch, wheat flour, glucose, fructose, etc.). Yeast extract and ammonium acetate (1%) as nitrogen sources gave higher yield compared to other nitrogen sources (peptone, malt extract, casein, asparagine, glycine, beef extract), whereas ammonium chloride, ammonium sulphate and urea inhibited the enzyme activity. Addition of Ca+2 (10 – 40 mM) to the culture medium did not result in further improvement of enzyme production, whereas the addition of surfactants (Tween 20, Tween 40, Tween 80, and sodium lauryl sulphate) at 0.02% resulted in 2 – 15% increase in enzyme production. There were no significant variations in enzyme yield between shaked-flask and laboratory fermentor cultures. The purified enzyme is in two forms with molecular mass of 18.0 ± 1 and 43.0 ± 1 kDa in native SDS-PAGE. K e y w o r d s: Bacillus subtilis; cow dung microflora; extracellular "-amylase

Introduction Various microorganisms are associated with rumen microflora of cattle, sheep, buffalos and goats, which are largely responsible for digestion in these animals (Ware et al., 1988). The general culturable microflora of the cattle gut involves Bacillus, Bifidobacterium, Lactobacillus, and yeasts (Wallace and Newbold, 1993). According to EEC directive 70/524, several microorganisms have been authorized as additives for feedstuffs (Abe et al., 1995; Auclair, E., 2006 http://resources.ciheam.org/om/pdf/c54/01600010.pdf). These microorganisms belong to different species: Bacillus subtilis, Bacillus cereus, Bacillus licheniformis, Enterococcus faecium and Saccharomyces cerevisiae. All these strains have a positive effect on the health of different animal species such as beef cattle, dairy cows, pigs and rabbits (Breul, 1998; Guo et al., 2006). In our previous study it was found that B. subtilis strains were one of the predominant groups of bacteria isolated from culturable cow dung microflora (Swain and Ray, 2006). These strains exhibited several beneficial attributes, which include biocontrol of pathogenic fungi, i.e. Fusarium oxysporum and Botryodiplodia theobromae, plant growth promotion, sulphur oxidation, solubilization of rock phosphorus and production of industrially important enzymes (amylase and cellulase) in vitro. Among the starch hydrolyzing enzymes that are produced on an industrial scale, thermostable "-amylases are of significant commercial interest. "-Amylases (E.C. 3.2.1.1) randomly hydrolyze "-1,4, glycosidic linkages in starch and its partial hydrolysis products. Bacteria belonging mainly to the genus Bacillus have been widely used for the commercial production of thermostable "-amylase (Tonkova, 2006). The most important characteristic of thermophilic amylase producers is their ability to produce the enzyme with higher operational stability and longer shelf-life. The "-amylases presently used in starch saccharification * Corresponding author: C. Ray, Cen. Tuber Crops Res. Inst., Bhubaneswar 751019, Orissa, India; e-mail: [email protected]

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require Ca2+ for their activity and/or stability. There is a continuous search for microorganisms producing "-amylase which do not require Ca2+ (Tonkova, 2006). The present study was carried out to explore the possibility of production of thermostable Ca 2+ independent "-amylase by B. subtilis strains isolated from cow dung microflora, to characterize the enzyme, and to investigate conditions for its production. Experimental Materials and Methods B. subtilis strain. The B. subtilis strain CM3 isolated from culturable cow dung microflora (Swain and Ray, in press) was used in this study. The culture was maintained on Nutrient Agar slants at 4°C. Production of "-amylase. The production of "-amylase from B. subtilis strain CM3 was carried out in a basal medium with following composition: 1% soluble starch, 0.2% yeast extract 0.5% peptone, 0.05% MgSO4, 0.05% NaCl, 0.015% CaCl2 and 2% agar (pH adjusted to 7.0 before autoclaving). The medium (100 ml taken in 250 ml Erlenmeyer flasks) was inoculated with 2% (1 × 106 CFU/ml) of 24 h seed culture and incubated with shaking (150 rpm) and 50°C for 36 h in an orbital shaker-incubator (Remi Pvt. Ltd, Bombay, India). Different carbon and nitrogen sources (1%) were used for optimization of nutritional factors. The various carbon sources tested were: soluble starch, potato starch, sweet potato starch, cassava starch, wheat flour, glucose, fructose, maltose, lactose and sucrose. Different levels of soluble starch (0.5 – 3.0%) were also used. The nitrogen sources tested were: yeast extract, peptone, malt extract, casein, asparagine, glycine, beef extract, ammonium chloride, ammonium sulphate, urea and ammonium acetate. Different concentrations of yeast extract (0.25 – 3.0%) and ammonium acetate (0.25 – 3%) were also used. The physical parameters, including pH of the medium (4.0 – 11.0), temperature of incubation (40 – 90°C), and incubation period (12 – 60 h) were also tested. The effect of different surfactants (0.02%): Tween 20, Tween 40, Tween 80 and sodium lauryl sulphate on amylase activity was studied. Similarly, the effect of Ca2+ ions (10 – 40 mM) was also tested. All the experiments were carried out in triplicate and mean data with standard deviations (±) were calculated. At the end of the incubation period (36 h for all experiments except the experiment in which incubation period was studied), the cell-free enzyme supernatant was obtained by centrifugation at 8000 × g for 15 min at 4°C. Enzyme stability at various temperatures and pH was also studied by incubating cell-free supernatants at different temperatures (40 – 90°C) and assay buffer pH (4.0 – 10.0). The pH of 4.0 – 5.0 was maintained with acetate buffer (0.2 M) while pH of 6.0 – 8.0 and 9.0 – 10.0 was achieved with phosphate (0.1 M) buffer and borax-NaOH (0.05 M) buffers, respectively. B. subtilis CM3 was also cultivated in a 2-liter fermentor (Model Biostat B, B. Braun, Germany) with a working volume of 1l (Glass Jar Vessel) containing "-amylase production medium (pH 7.0) at 60°C and 150 rpm. The aeration level was maintained at 1vvm (volume of air/unit volume of the medium/min). The enzyme production and growth of the organism were compared with shaked-flask cultures incubated at 60°C and 150 rpm in an orbital shaker incubator. B. subtilis growth. The growth of B. subtilis strain CM3 was determined by measuring the optical density of the growth medium at 600 nm in a UV-Vis spectrophotometer (Cecil Instruments, UK). Thin-layer chromatographic analysis. The products liberated by the action of amylase on starch were identified by spotting the starch digest and standard sugars (glucose and maltose) on a silica gel plate activated at 80°C for 30 min. The plates were developed in butanol:ethanol:water (50:30:20) and dried overnight at 32 ± 2°C. The individual sugar(s) were visualized with acetone-silver nitrate solution (0.1 ml saturated solution of AgNO3 in 20 ml of acetone). Amylase assay. The amylase assay was based on the reduction in blue colour intensity resulting from enzyme hydrolysis of starch (Palanivelu, 2001). The reaction mixture consisted of 0.2 ml enzyme (cell free supernatant), 0.25 ml of 0.1% soluble starch solution and 0.5 ml of phosphate buffer (0.1 M, pH 6.8) incubated at 50°C for 10 min. The reaction was stopped by adding 0.25 ml of 0.1 N HCl and colour was developed by adding 0.25 ml of I/KI solution (2% KI in 0.2% I). The optical density (OD) of the blue colour solution was determined using a UV-Vis spectrophotometer (Cecil Instruments, UK) at 690 nm. One unit of enzyme activity is defined as the quantity of enzyme that caused 0.01% reduction of blue colour intensity of starch iodine solution at 50°C in one min per ml (Palanivelu, 2001). The optimal activity and stability of the partially purified enzyme (described in the following section) at various pH values (5.0 – 10.0) and temperature (40 – 80°C) were studied. Partial purification of the enzyme. "-Amylase was partially purified by ammonium sulphate fractionation followed by dialysis and gel filtration chromatography. A total of 100 ml of bacterial culture filtrate was centrifuged at 8000 × g for 20 min at 4°C to remove the cells. The supernatant was brought to 50% ammonium sulphate saturation at 4°C in an ice bath. The precipitated protein was collected by centrifugation at 8000 × g at 4°C and dissolved in a minimum volume of phosphate buffer (0.1 M; pH, 6.0). The enzyme solution was dialyzed at 4°C against the same buffer for 24 h at 4°C with continuous stirring and three changes of the same buffer. The DEAE cellulose ion exchange column was pre-equilibrated with the same buffer. The dialysate was concentrated with a rotary evaporator at 50°C and applied to the DEAE cellulose column at a flow rate of 0.6 ml/min with 200 ml linear NaCl gradient (0 to 1.0 M). Fractions of 10 ml were collected and each fraction was analyzed for protein concentration and "-amylase activity. The active fractions were pooled and concentrated with a rotary evaporator at 50°C. The final enzyme solution was taken for Sodium Dodecyl Sulphate-Polyacrylamide Gel Electrophoresis (SDS-PAGE). Electrophoresis and molecular mass determination. SDS-PAGE was performed with 12% polyacrylamide gel using a Mini GEL electrophoresis system (Model No 0502, Bangalore Genei Pvt. Ltd., Bangalore, India) as described by Laemmli (1970). The bacterial proteins were stained with 0.2% Coomassie Brilliant Blue. The molecular mass of the partially purified amylase was estimated using standard ‘protein markers’ (PMW-M) of known molecular mass (14.3 – 97.4 kDa) (Bangalore Genei Pvt. Ltd., Bangalore, India).

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Results

5000

0.45

4500

0.4

4000

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1500 1000

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Enzyme activity (Units)

Amylase production by B. subtilis strain CM3 started in the log phase of growth and maximum enzyme production was achieved during the stationary phase (36 h) of the growth of the organism (Fig. 1). Further, this strain produced amylase optimally at growth temperature of 50– 70°C (Fig. 2 A). To examine the thermostability of the enzyme, the enzyme solution buffered at pH 6.8 was incubated at various temperature (40– 90°C) for 30 min. The maximum activity was 4900– 4960 units at temperature of 60– 70°C (Fig. 2 A).

0 12

24

36

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Growth

Fig. 1. Effect of incubation period on growth and "-amylase production by B. subtilis strain CM3

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production

stability

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Stability

Fig. 2. Effect of temperature (A) and pH (B) on "-amylase production and stability by B. subtilis strain CM3

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Swain M.R. et al. Table I Effect of different carbon sources (starch and sugar) and nitrogen sources (inorganic and organic) on "-amylase production by B. subtilis strain CM3 Carbon sources

Enzyme production (Units)

Nitrogen sources

Enzyme production (Units)

Soluble starch

4870.50 ±103.4

Peptone

4865.23 ± 106.2

Potato starch

4970.50 ± 098.0

Casein

4462.00 ± 093.23

Sweet potato starch

4923.50 ± 121.1

Malt extract

1088.23 ± 065.2

Cassava starch

4923.50 ± 110.4

Yeast extract

4918.56 ± 121.0

Wheat flour

4970.50 ± 103.4

Beef extract

4491.13 ± 095.2

Glucose

4970.50 ± 087.2

Asparagine

4751.56 ± 103.6

Fructose

4978.54 ± 056.6

Glycine

4311.00 ± 056.3

Maltose

4976.20 ± 00.0

Ammonium chloride

857.00 ± 103.2

Lactose

4967.81 ± 121.3

Ammonium sulphate

1065.15 ± 069.6

Sucrose

4957.08 ± 103.2

Urea

107.50 ± 106.5

Ammonium acetate

4938.32 ± 103.2

± Standard deviations

When the crude enzyme was heated at 90°C for 30 min, 80% of the original enzyme activity was lost. Using thin-layer chromatographic analysis, the end products of starch hydrolysis detected were glucose and maltose which suggested an endo-mode of action for the amylase, i.e. "-amylase (data not shown). Following optimization of parameters, the optimum pH for enzyme production was in the range of 5.0– 9.0 (4950– 5180 units/ml culture medium) and stability was in the range of 5.0– 8.0 (3455– 3947 units)

A

Enzyme activity (Units)

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1

1.5

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Soluble starch concentration (%)

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Yeast extract

0.5

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centratA Ammonium acetate (concentration – %)

Fig 3. Effect of different concentrations (%) of soluble starch (A), yeast extract and ammonium acetate (B) on "-amylase production by B. subtilis strain CM3

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43 000

AII

20 100 AI 14 300

A

B

Fig. 4. Determination of molecular weight by SDS-PAGE. (A) molecular mass markers: (97.0–14.3 kDa) (B) "-amylase(s) from B. subtilis strain CM3. Isozymes AI and AII

(Fig. 2B). The organism could grow and produce almost equal amount of amylase in medium containing different carbon sources (1% w/v), i.e. soluble starch, glucose, fructose, wheat flour, etc. (Table I). Moreover, among the various starch concentrations used, 1% soluble starch gave maximum enzyme yield (Fig. 3A). Similarly, enzyme production was more efficient in medium containing organic nitrogen sources, i.e. peptone, yeast extract, beef extract, etc., as compared with inorganic nitrogen sources (ammonium acetate was the exception) (Table I). The enzyme production increased concomitantly with increase in concentrations of yeast extract and ammonium acetate up to 1% level, beyond that there was a gradual decline (Fig. 3B). Amylase production increased in culture medium due to addition of surfactants such as Tween 20, Tween 40, Tween 80 and sodium lauryl sulphate. While the medium without surfactant (control) yielded 4756 units of amylase, 5290, 5416, 5104 and 5020 units of enzyme were produced in medium containing 0.02% of Tween 20, Tween 40, Tween 80 and sodium lauryl sulphate, respectively. Addition of Ca2+ ion (10– 40 mM) had no significant difference (increase or decrease) on "-amylase activity. The enzyme production varied in the range of 4965– 5011 units in medium with or without Ca2+ ion. "-Amylase was partially purified using ammonium sulphate fractionation. The crude extract contained 327.23 mg/ml protein and showed a specific activity of 14.53 units/mg protein. After partial purification, the specific activity increased to 39.61 units/mg protein with a yield of 19% and three fold purification. 0.3

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Incubation period (h) Fermentor enzyme production

Shake flask enzyme production

Fermentor B.subtilis growth

Shake flask B.subtilis growth

Fig. 5. Enzyme production and growth of B. subtilis strain CM3 in laboratory fermentor and shake flask cultures

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Swain M.R. et al.

4

Electrophoretic studies showed that there were two forms of the "-amylase (AI and AII) and the molecular mass of partially purified enzymes were approximately 18 000 ± 1000 (AI) and 43 000 ± 1000 (AII) Da, respectively (Fig. 4). The partially purified enzyme showed similar pH (7.0) and temperature (60°C) optima and stability as the crude enzyme preparation (data not shown) When B. subtilis strain CM3 was cultivated in a laboratory fermentor, peak activity (4685 units) was obtained at 36 h which was similar to the shake-flask cultures (4487 units) (Fig. 5) while the cell density attained in the fermentor (0.253 at 600 nm) was similar to that in the shake flasks (0.249 at 600 nm). Discussion Members of the genus Bacillus produce a large variety of extracellular enzymes, of which amylases are of particularly significant industrial importance. Similar to other Bacillus species, i.e. B. coagulans, B. thermooleovorans, B. licheniformis, etc., B. subtilis strain CM3 was found to produce "-amylase maximally at the optimal growth temperature of 50– 60°C (Babu and Satyanarayana, 1993; Malhotra et al., 2000; Das et al., 2004; Najafi et al., 2005). The pH range (5.0– 9.0) was found to be optimal for amylase production by B. subtilis CM3 as also reported for B. brevis (Tsvetkov and Emanuilova, 1989), B. coagulans, B. licheniformis (Krishnan and Chandra, 1983), B. thermooleovorans (Malhotra et al., 2000) and B. subtilis (Das et al., 2004). Cultural conditions have a profound influence on amylase production. Enzyme production was maximal when the cell population entered into stationary phase of growth. Similar findings have been recorded for several other Bacillus species i.e. Bacillus amyloliquefaciens (Roychoudhary et al., 1989), B. thermooleovorans (Malhotra et al., 2000), and B. subtilis (Baig et al., 1984; Najafi et al., 2005). Amylase production by this strain was constitutive since biosynthesis of the enzyme took place not only in the presence of starch but also with other carbon sources. Moreover, the amylase yield was similar in all types of carbon sources such as soluble starch, potato starch, glucose, maltose, sucrose, etc. and therefore, this was not considered to reflect inducibility (Tonkova, 2006). Starch at a concentration of 1% (w/v) supported optimal enzyme production, followed by a decline at higher concentrations (Fig. 2A). This can be attributed to the high viscosity of culture broth at such concentrations, which interferes with O 2 transfer leading to limitation of dissolved O2 for the growth of bacteria (Rukhaiyar and Srivastava, 1995). Among nitrogen sources, organic nitrogen supported higher amylase secretion in comparison with inorganic nitrogen sources, with ammonium acetate as the only exception. Moreover, urea, ammonium chloride and ammonium sulphate at 1% level inhibited (78.1– 97.5%) enzyme activity. However, there was no difference in the growth of B. subtilis grown with any of these nitrogen sources. Similar results were obtained for other Bacillus spp. i.e., B. licheniformis (Aiyer, 2004), B. subtilis (Haq et al., 2002), B. thermooleovorans (Narang and Satyanrayan, 2001) and B. coagulans (Babu and Satyanrayan, 2001). In contrast, Das et al. (2004) reported maximum "-amylase production by B. subtilis DM-03 obtained by using ammonium chloride as the nitrogen source. Further, there was no significant variations in the enzyme yield among organic nitrogen sources (beef extract, peptone, yeast extract, etc.) incorporated at 1% level in the basal medium. Similar results were reported for B. thermooleovorans (Malhotra et al., 2000), B. stearothermophilus (Davies et al., 1980) and B. amyloliquefaciens (Babu and Satyanarayana, 1993). The concentration of yeast extract or ammonium acetate was also critical for obtaining maximum enzyme yield. The enzyme levels were high at 0.5– 1.0% levels and declined sharply thereafter as in the case of B. coagulans (Malohotra et al., 2000). The decline in amylase production at increased nitrogen concentration could be due to the lowering of pH of the production medium or the induction of protease, which suppresses the amylolytic activity (Tonkova, 2006). It is generally known that surfactants often increase enzyme secretion and production (Ray et al., 1990) but the explanation how they act to increase enzyme yield is largely conjectural (Reese and Maguire, 1969). In this study, the various surfactants (0.02%) applied enhanced amylase activity (2– 15%) over control (no surfactant). The increase enzyme accumulation might be due to increase in cell membrane permeability (Rao and Satyanarayana, 2003) and/or modification (swelling) of starch (Moorthy, 2002). Similar results were found in the case of other microorganisms, i.e. Thermomyces lanuginosus (Arnesen et al., 1998), Bacillus circulans (Palit and Banerjee, 2001), etc. For production and stability of amylase of many Bacillus spp., addition of Ca2+ ion is often necessary (Tonkova, 1991; 2000). In this study, addition of Ca2+ had no effect at all on enzyme activity. Ca2+ independent "-amylase from Bacillus spp. have been reported by several authors (Malhotra et al., 2000; Kumar et al., 1990; Mamo et al., 1999). Ca2+ independent amylase merits consideration for starch liquefaction, especially in the manufacture of fructose syrup, where Ca2+ is a known inhibitor of glucose isomerase (Tonkova, 2006).

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Control environmental conditions are critical in achieving higher concentration or yield of any microbial product. Since B. subtilis strain CM3 could grow and produced "-amylase over a wide range of pH (5.0– 9.0) and temperature (40– 60°C), this might be the reason for not obtaining a higher yield in the fermentor in comparison with shaked flask culture. Several bacteria, i.e. Lactobacillus plantarum, L. casei, L. acidophilus, B. subtilis, Enterococcus diacetylactis, etc. were isolated from the lower part of the gut of cow (Ware et al., 1988). Other than these, the cow rumen contains various species of Bacillus and Bifidobacterium as well as yeasts (S. cerevisiae) for better rumen fermentation (Kung, L. Jr., www.das.psu.edu/dairynutrition/documents/kung.pdf.), which form the initial culturable microflora of cow dung (Swain and Ray, in press). Among the culturable cow dung microflora, B. subtilis strains were found to be the most common bacteria (Swain and Ray, 2006). The starch-digesting characteristics of these organisms, i.e. pH, 5.0– 9.0; temperature, 40– 60°C and capable of growing in various carbon and organic nitrogen sources may be useful parameters for the bacteria to digest starch as a component of feed under a wide-range of gut environmental conditions. Preliminary studies have also shown that this organism possesses endo- and exo- cellulase activities (Swain and Ray, 2006). The multi-complex enzyme (amylase and cellulase) activities of B. subtilis will be useful to the cattle in digesting a wide range of feed rich in starch and cellulose. There were wide variations in molecular mass of amylases of different Bacillus spp., i.e. B. coagulans and B. subtilis (42 800– 67 000 Da.) (Babu and Satyanarayan, 1993; Ozcan et al., 2001; Das et al., 2004), Bacillus spp.(97000 Da) (Kim et al., 1995). In this study, the two forms of amylase (AI and AII) of B. subtilis strain CM3 had molecular masses of 18 ± 1 and 43 ± 1 kDa, respectively. In a recent study, Najafi et al. (2005) reported molecular mass of "-amylase by a strain of B. subtilis AX 20 was 14.9 kDa. In the present study, the "-amylase of B. subtilis strain CM3 isolated from cow dung is Ca2+ independent and thermostable and can therefore be of importance for the starch-processing industries. Further work is in progress in our laboratory on the use of this enzyme in cassava (Manihot esculanta L.) and sweetpotato (Ipomoea batata L.) starch saccharification for production of ethanol, organic acid and other bioproducts. Acknowledgments. Financial assistance from the Indian Council of Agricultural Research, New Delhi, India (Project No 8(39)/ 2003-Hort II dated 7 June, 2004) is sincerely acknowledged. The authors thank Dr. T.K. Danger, Senior Scientist (Plant Pathology, Central Rice Research Institute, Cuttack, Orissa, India) for suggestions.

Literature A b e F., N. I s h i b a s h i and S. S h i m a m u r a. 1995. Effect of administration of Bifidobacteria and lactic acid bacteria to newborn calves and piglets. J. Dairy Sci. 78: 2838–2846. A r n e s e n S., S.H. E r i k s e n, J. O l s e n and B. J e n s e n. 1998. Increased production of "-amylase from Thermomyces lanuginosus by the addition of Tween 80. Enzyme Microb. Technol. 23: 249–252. Auclair E., http://resources.ciheam.org/om/pdf/c54/01600010. pdf. Accessed on 20 th January, 2006. A i y e r P.V.D. 2004. Effect of C:N ratios on "-amylase production by Bacillus licheniformis SPT 27. African J. Biotech. 3: 519–522. B a b u K.R. and T. S a t y a n a r a y a n a. 1993. Parametric optimization of extracellular "-amylase production by thermophilic Bacillus coagulans. Folia Microbiol. 38: 77–80. B a i g M.A., J. P a z l a r o v a and J. Vo t r u b a. 1984. Kinetics of "-amylase production in a batch and fedbatch culture of Bacillus subtilis. Folia Microbiol. 29: 359–364. B r e u l S. 1998. Les Probiotiques en alimentation animale. Med. Chir. Dig. 27: 89–91. D a v i e s P.E., D.L. C o h e n and A. W h i t a k e r. 1980. The production of "-amylase in batch and chemostat culture of Bacillus stearothermophilus. Antonie Van Leeuwenhock 46: 391–398. D a s K., R. D o l e y and A.K. M u k h e r j e e. 2004. Purification and biochemical characterization of thermostable, alkaliphilic, extracellular "-amylase from Bacillus subtilis DM-03, a strain isolated from the traditional food of India. Biotechnol. Appl. Biochem. 40: 291–298. G u o X, D. L i, W. L u, X. P i a o and X. C h e n. 2006 Screening of Bacillis as potential and subsequent confirmation of the in vitro effectiveness of Bacillus subtilis. Antonie. Van Leeuwenhoek 90: 139–146. H a q I.U., S. R a n i, H. A s h r a f and M.A. Q a d e e r. 2002. Biosynthasis of alpha amylases by chemically treated mutant of Bacillus subtilis. J. Biol. Sci. 2:73–75. K i m T.U., B.G. G u, J.Y. J e o n g, S.M. B y u n and Y.C. S h i n. 1995. Purification and characterization of a maltotetraoseforming alkaline "-amylase from an alkalophilic Bacillus strain GM8901. Appl. Environ. Microbiol. 61: 3105–3112. K r i s h n a n T. and A.K. C h a n d r a. 1983. Purification and characterization of "-amylase from Bacillus licheniformis CUMC305. Appl. Environ. Microbiol. 46: 430–437. K u n g L.Jr. A direct fed microbes and enzyme for dairy cows. www.das.psu.edu/dairynutrition/documents/kung.pdf. Assessed on 10th February 2005

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Kumar S.U., F. Rehana and K. Nand. 1990. Production of an extracellular thermostable calcium-inhibited "-amylase by Bacillus licheniformis MY10. Enzyme Microb. Technol. 12: 714–716. L a e m m l i U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680–685. M a l h o t r a R., S.M. N o o r w e z and T. S a t y a n a r a y a n a. 2000. Production and partial characterization of thermostable and calcium independent "-amylase of extreme thermophile Bacillus thermooleovorans NP54. Lett. Appl. Microbiol. 31: 378–384. M a m o G., B.A. G a s h e and A. G e s s e s s e. 1999. A highly thermostable amylase from a newly isolated thermophilic Bacillus sp. WN11. J. Appl. Microbiol. 86: 57–560. M o o r t h y S.N. 2002. Physicochemical and functional properties of tropical tuber starches: a review. Starch/Starke 54:559–592. N a j a f i M.F., D. D e o b a g k a r and D. D e o b a g k a r. 2005. Purification and characterization of an extracellular "-amylase from Bacillus subtilis AX20. Protein Expr. Purif. 41: 349–354. N a r a n g S. and T. S a t y a n a r a y a n. 2001. Thermostable "-amylase production by an extreme thermophile Bacillus thermooleovorans. Lett. Appl. Microbiol. 32: 31–35. O z c a n N., A. A l t I n a l a n and M.S. E k a i n c l. 2001. Molecular cloning of an "-amylase gene form Bacillus subtilis RSKK264 and its expression in Escherichia coli and in Bacillus subtilis. Turk J. Anim. Sci. 25: 197–201. P a l a n i v e l u P. 2001. Analytical Biochemistry and Separation Techniques. Kalamani Printers, Madurai, India. P a l i t S. and R. B a n e r j e e. 2001. Optimization of extracellular parameters for recovery of "-amylase from the fermented bran of Bacillus circulans GRS313. Braz. Arch. Biol. Technol. 44: 147–151. R a y R.C., G. P a d m a j a and C. B a l a g o p a l a n. 1990. Extracellular rhodanese production by Rhizopus oryzae. Zentralbl. Microbiol. 145: 259–268. R e e s e E.T. and A. M a g u i r e. 1969. Surfactants as stimulants of enzyme production by microorganisms. Appl. Microbiol. 17: 81–114. R o y c h o u d h a r y R.S., S.U.J. P a r u l e k a r and W.A. W e i g a n d. 1989. Cell growth and "-amylase production characterization of Bacillus amyloliquefaciens. Biotechnol. Bioeng. 33: 197–206. R a o J.L.U. and T. S a t y a n a r a y a n a. 2003. Enhanced secretion and low temperature stabilization of a hyperthermostable and Ca2+-independent "-amylase of Geobacillus thermolevorans by surfactants. Lett. App. Microbiol. 36: 191–198. R u k h a i y a r R. and S.K. S r i v a s t a v a. 1995. Effect of various carbon substrates on "-amylase production from Bacillus species. World J. Microbiol. Biotechnol. 10: 76–82. T o n k o v a A. 1991. Effect of glucose and citrate on "-amylase production in Bacillus licheniformis. J. Basic. Microbiol. 31: 217–222. T o n k o v a A. 2006. Microbial starch converting enzymes of the "-amylase family. In: Ray R.C., and Wards O.P. (eds), pp. 421–472, Microbial Biotechnology in Horticulture, Science Publishers, Enfield, New Hampshire, USA. T s v e t k o v V.T. and E.I. E m a n u i l o v a. 1989. Purification properties of heat stable "-amylase from Bacillus brevis. Appl. Microbiol. Biotechnol. 31: 246–248. W a l l a c e R.T. and C.J. N e w b o l d. 1993. Rumen fermentation and its manipulation: the development of yeast culture as feed additives. In: Lyons, T.P. (ed.). Biotechnology in Feed Industry, pp. 173–192, Alltech Technical Publications, Kentucky, USA. W a r e D.R., P.L. R e a d and E.T. 1988. Lactation performance of two large diary herds fed Lactobacillus acidophilus strain BT 1386. J. Dairy Sci. 71: 219–222.

Polish Journal of Microbiology 2006, Vol. 55, No 4, 297– 301

Production of Tannase through Submerged Fermentation of Tannin-containing Plant Extracts by Bacillus licheniformis KBR6 PRADEEP K. DAS MOHAPATRA, KESHAB C. MONDAL and BIKAS R. PATI*

Department of Microbiology, Vidyasagar University, Midnapore, West Bengal, India Received 28 July 2006, revised 17 August 2006, accepted 13 October 2006 Abstract Tannins are water-soluble polyphenolic compounds found in plants as secondary metabolites. The presence of these substances in the barks of eight different plants was initially examined and their crude extracts were used separately as a substrate for production of tannase through submerged fermentation by Bacillus licheniformis KBR6. Tannase production as well as biodegradation of the substrate reached a maximum within 15 to 18 h against crude tannin extract obtained from Anacardium occidentale. Among different concentrations of the crude tannin tested, 0.5% (w/v) induced maximum synthesis of enzyme. Tannase production was higher by almost two-fold in the presence of crude tannin compared to pure tannic acid used as a substrate. It seems that industrial production of tannase, using bark extract of A. occidentale can be a very simple and suitable alternative to presently used procedures. K e y w o r d s: Bacillus licheniformis, plant tannins, submerged fermentation

Introduction Tannins are the fourth most abundant plant constituent after cellulose, hemicellulose and lignin (Swain, 1965). Generally tannins are accumulated as secondary metabolites in the bark and heartwood of plants. Although these substances have negligible value for growth, they play a great role in the immunity of plants. Tannin protects the vulnerable parts of the plants from microbial attack by inactivating viruses and invasive microbial extracellular enzymes (Field and Lettinga, 1992). In spite of its antimicrobial effect, some organisms use this compound as a nutrient for growth, utilizing tannase-hydrolyzing enzyme (Lewis and Starkey, 1969). Tannase (tannin-acyl-hydrolase, E.C: 3.1.1.20) has been known to hydrolyze the ester and depside linkages of hydrolysable tannins into glucose and gallic acid. Nowadays, the enzyme has wide applications in food, beverage, brewing, cosmetics and chemical industries (Lekha and Lonsane, 1997). It is used mainly for the preparation of gallic acid, instant tea, acron wine, coffee flavoured soft drinks, high-grade leather tannin, clarification of beer and fruit juice, detannification of food and to clean-up highly polluting tannin from the effluent of leather industry (Lekha and Lonsane, 1997). Gallic acid, a hydrolytic product of tannin, has different uses like preparation of trimethoprim, pyrogallol, propyl gallate, dyes, etc. (Hadi et al., 1994; Mukherjee and Banerjee, 2003). Most of the reported tannase-producing organisms are fungi (Aoki et al., 1976; Bhat et al., 1998; Mondal et al., 2001c; Ramirez-Coronel et al., 2003) and only a few are bacteria (Deschamps et al., 1983; Mondal and Pati, 2000; Mondal et al., 2001a). Many authors studied tannase production by these organisms in the medium containing pure tannic acid acting as both inducer as well as available carbon source. Pure tannic acid is a very costly substrate and is not suitable for large-scale production of the enzyme. In this respect crude tannin could be cost effective and suitable for the commercial production of the enzyme. Agro-residues and forest products are generally considered the best source of tannin-rich substrate (Pandey et al., 1999). * Corresponding author: B.R. Pati, Department of Microbiology, Vidyasagar University, Midnapore 721 102, West Bengal, India; fax: (091)03222-275329; e-mail: [email protected]

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Production of tannase by Rhizopus oryzae and Aspergillus foetidus from the powdered fruits of Terminalia chebula and Caesalpinia digyna has been reported (Mukherjee and Banerjee, 2004). In this regard there are no reports on bacterial tannase production using tannin containing any agro-based substrates. In the present publication we are reporting for the first time the production of tannase by Bacillus licheniformis KBR6 through submerged fermentation of crude tannin extracted from the barks of various forest plants. Experimental Materials and Methods Microorganism. The non-pathogenic tannase-producing soil bacterium, Bacillus licheniformis KBR 6 (IMI: 379224) described earlier (Mondal and Pati, 2000) was used in the present study. Preparation of inoculum. Inoculum was prepared by growing a loopful amount of stock culture of the bacterium in 50 ml sterile tannic acid medium (pH 5.0) at 35°C for 20 h. Composition of the medium was (g%, w/v): tannic acid, 1; NH 4Cl, 0.3; KH2PO4, 0.05; K2HPO4, 0.05 and MgSO4, 0.05. Extraction of crude tannins. Collected barks of different forest plants from south West Bengal, India, were cut into small pieces and dried in hot air-oven at 60°C for 24 h. The barks (50 g) were then mixed with distilled water (200 ml) and kept at room temperature overnight. After soaking, the mixture was boiled for 10 min. The filtered solutions were used as source of crude tannin (Schanderi, 1970). Detection of tannin by paper chromatography. Presence of tannin in the plant extract was confirmed through paper chromatographic analysis. A descending mode of solvent system containing n-butanol, acetic acid and water (4:1:5) was used for the study. Detection of the spot was made by FeCl3 (0.1 g % in 30% methanol) as colouring spray reagent (Mondal and Pati, 2000) and confirmed after comparing it with standard tannic acid. Measurement of tannin biodegradation. The tannin content of the crude plant extracts was measured before and after fermentation by Folin-Denis method (Schanderi, 1970). The crude extract (0.2 ml) was initially diluted with 8.3 ml of distilled water and then mixed with 0.5 ml of Folin-Denis reagent. After proper mixing, 1 ml of 15% (w/v) Na 2CO3 was added to it and kept in the dark for 30 min at room temperature. The absorbency of tannin was measured at 700 nm (SL 171 MINI SPEC, ELICO, INDIA) and its concentration was calculated using pure tannic acid as standard. Mode of fermentation. Tannase production by B. licheniformis KBR6 was achieved through submerged fermentation of crude tannin at 35°C in a rotary shaker (200 rpm). Different concentrations of tannin were prepared by diluting the measured crude tannin with distilled water. The pH of the medium was adjusted to 5.0 after sterilization. Fermentations were carried out separately in individual 250 ml Erlenmeyer flasks containing 50 ml medium with 1% (v/v) fresh inoculum. The cell-free fermented broth was used as the source of the enzyme. The growth of the organism in culture media was monitored by measuring dry weight of the biomass (mg/ml). All experiments were done in triplicate and data presented as mean ± SE. Assay of tannase. Tannase activity in the fermented medium was determined by the colorimetric method of Mondal et al. (2001b). For assay, 0.1 ml of enzyme was incubated with 0.3 ml of substrate tannic acid (1.0% w/v in 0.2 M citrate buffer, pH 5.0) at 50°C for 30 min. The reaction was terminated by the addition of 3 ml BSA solution (1 mg/ml), which also precipitates the residual tannic acid. A control reaction was done side by side using heat-denatured enzyme. The tubes were then centrifuged (5000 × g, 10 min) and precipitate was dissolved in 2 ml of SDS-triethanolamine (1% w/v, SDS in 5% v/v, triethanolamine) solution. Absorbency was measured at 530 nm after addition of 1 ml of FeCl 3 (0.13 M). The specific extinction co-efficient of tannic acid at 530 nm was 0.577 (Mondal et al., 2001b). Using this co-efficient, one unit of tannase activity is defined as the amount of enzyme required to hydrolyze 1 mM substrate (tannic acid) in 1min at 50°C and pH 5.0.

Results and Discussion The selection of a substrate for large-scale enzyme production by fermentation depends upon its availability and cost. In this regard several low cost agro-forest residues were used as substrates for obtaining the desired fermented product (Pandey et al., 1999). Different substrates (tannin containing plant extracts) were tested in this experiment for production of tannase through submerged fermentation by B. licheniformis KBR6. Tannin contents of the bark of some commonly available plants were initially examined by paper chromatography and quantified by colorimetric method (Table I). Among the eight plant species tested, the maximum amount of tannin was found in the extract of the bark of Acacia auriculiformis. On the basis of tannin content, the studied plants can be arranged in the following order: Acacia auriculiformis > Casuarina equisetifolia > Psidium guazava > Anacardium occidentale > Delonix regia > Eucalyptus tereticornis > Cassia fistula > Ficus benghalensis (Table I). Tannase production by B. licheniformis KBR6 was studied using different crude tannins as submerged fermentation media. It has been found that extract of A. occidentale was the best for induction of tannase (0.62 ± 0.04 U/ml) and as much as 73% of tannin in the culture media was degraded by it (Table I). Enzyme production by the organism was found to reach a maximum within 15– 18 h of growth in all the tannin

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Production of tannase by B.licheniformis Table I Measurement of tannin content in crude plant extract, tannin biodegradation and tannase production by B. licheniformis KBR6 Tannin content in the crude extract (g% w/v)

Biodegradation (%) of tannins through fermentation

Tannase production (U/ml)

Acacia auriculiformis

1.33 ± 0.18

27 ± 3.60

0.32 ± 0.09

Anacardium occidentale

0.65 ± 0.10

73 ± 4.65

0.62 ± 0.04

Casuarina equisetifolia

0.85 ± 0.09

34.5 ± 5.33

0.06 ± 0.10

Cassia fistula

0.43 ± 0.16

64.4 ± 1.88

0.52 ± 0.01

Delonix regia

0.54 ± 0.09

21 ± 3.38

0.12 ± 0.07

Eucalyptus tereticornis

0.45 ± 0.12

58 ± 6.10

0.17 ± 0.02

Ficus benghalensis

0.39 ± 0.20

53 ± 1.58

0.40 ± 0.11

Psidium guazava

0.73 ± 0.03

0.08 ± 0.01

0.32 ± 0.06

Plant source

The organism was grown for 18 h at specified conditions, which are described in Materials and Methods. The amount of tannase production is significantly correlated with the biodegradation of crude tannins (correlation coefficient, r = + 0.54).

extracts except for Eucalyptus, (Fig. 1). A similar type of time related enzyme production by the same organism was also reported with pure tannic acid as substrate (Mondal and Pati, 2000). Tannase production by the organism was found to be maximal in the extract of A. occidentale compared to other plant extracts (Fig. 1). It is not clear to us why the production of enzyme in the extract of A. occidentale is high, but we assume that there may be some inducing factors that accelerate enzyme synthesis. Enzyme production was also studied at different concentrations (0.5– 2.0 %, w/v) of crude tannins. It was observed that a specific concentration of crude tannin from a plant influenced enzyme production the strongest (Table II). In our experiments a maximal amount of enzyme was produced in medium containing 0.5% (w/v) of crude tannin of A. occidentale, but the highest enzyme production (Mondal et al., 2000) was observed when 1.5% (w/v) of pure tannic acid was used as substrate in the medium The concentration of tannin is thus a very important determining factor for tannase biosynthesis for most fungi and bacteria (Lekha and Lonsane, 1997; Mondal et al., 2000; Banerjee et al., 2001). The actual mode of tannase induction in a particular concentration of tannin has not been properly explained until now. Lewis and Starkey (1969) mentioned that higher concentrations of tannin lead to non-reversible bonds with surface proteins and impair the metabolism as well as growth of the organism. One of the most striking observations in this experiment is that enzyme production was increased about four-fold in the medium containing basal salt with crude tannin (0.5%) compared to medium containing crude tannin extract (0.5%) of A. occidentale alone (Table III). This result revealed that some specific microelements (salts and ions) are probably essential

1

Enzyme activity (U/ml)

Acacia auriculiformis 0.8

Anacardium occidentale Casuarina equisitifolia

0.6

Cassia fistula Delonix regia

0.4

Eucalyptus sp. 0.2

Ficus benghalensis Psidium guazava.

0 6

9

12

15

18

21

24

Time (h)

Fig. 1. Time course of tannase production by Bacillus licheniformis KBR6 using different plant extracts (crude tannin) as substrate

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Das Mohapatra P.K. et al. Table II Effect of different concentrations of crude tannin from different plants on tannase production (mean ± SD) Concentration of tannin (%)

Tannase (U/ml)

Acacia auriculiformis

0.5 1 2

0.33 ± 0.09 0.42 ± 0.11 0.35 ± 0.19

Anacardium occidentale

0.5 1 2

1.23 ± 0.29 0.86 ± 0.08 0.83 ± 0.23

Casuarina equisetifolia

0.5 1 2

1.01 ± 0.11 1.12 ± 0.15 1.02 ± 0.14

Cassia fistula

0.5 1 2

0.55 ± 0.25 0.83 ± 0.14 0.84 ± 0.30

Delonix regia

0.5 1 2

0.36 ± 0.02 0.89 ± 0.42 0.10 ± 0.32

Eucalyptus tereticornis

0.5 1 2

0.20 ± 0.15 0.10 ± 0.01 0.210 ± 0.22

Ficus benghalensis

0.5 1 2

0.50 ± 0.31 0.83 ± 0.20 0.85 ± 0.08

Psidium guazava

0.5 1 2

0.30 ± 0.25 0.86 ± 0.05 0.78 ± 0.16

Plant source

Table III Comparative study of tannase production in pure tannic acid and crude tannin extract as substrates by Bacillus licheniformis KBR6 Substrates in fermentation medium

Growth (mg/ml)

Tannase (U/ml)

Crude tannin (0.5 g/100 ml)

0.82 ± 0.18

0.17 ± 0.06

Crude tannin (0.5 g/100 ml) + Basal salts*

1.12 ± 0.22

0.66 ± 0.12

Tannic acid (1.5 g/100 ml) containing enriched medium**

0.58 ± 0.12

0.31 ± 0.13

* Composition (g%, w/v): NH 4Cl, 0.3; KH2PO4, 0.05; K 2HPO4, 0.05 and MgSO4, 0.05. ** Composition (g%, w/v): NH 4Cl, 0.3; KH 2PO4, 0.05; K 2HPO4, 0.05; MgSO 4, 0.05; glucose, 0.01%; alanine, 0.01%; pyridoxine, 0.003%. Fermentation was carried out at specified condition for 18 h.

for growth as well as enzyme synthesis by B. licheniformis KBR6. Both growth of the organism and enzyme production increased two-fold when it was grown in salt containing crude tannin extract rather than enriched pure tannic acid medium. All these beneficial effects of the plant extract of A. occidentale make it promising as one of the best as well as cheaper substrates for the large scale production of microbial tannase. In conclusion, tannase has now been extensively used in different biochemical industries. The selected bacterium used in this study is able to synthesize high amounts of tannase through fermentation of crude tannin of A. occidentale. Exploitation of these plant extracts could be a source of cheaper substrate for industrial production of microbial tannase. Acknowledgement. Financial support by the All India Council for Technical Education, New Delhi, India is acknowledged.

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Literature A o k i K., R. S h i n k e and H. N i s h i r a. 1976. Purification and some properties of yeast tannase. Agric. Biol. Chem. 40: 79–85. B a n e r j e e D., K.C. M o n d a l and B.R. P a t i. 2001. Production and characterization of extracellular and intracellular tannase from newly isolated Aspergillus aculeatus DBF9. J. Basic Microbiol. 41: 313–316. B h a t T.K., B. S i n g h and O.P. S h a r m a. 1998. Microbial degradation of tannins – a current perspective. Biodegradation 25: 43–357. D e s c h a m p s A.M., G. O t u k and J.M. L e b e a u l t. 1983. Production of tannase and degradation of chestnut tannin by bacteria. J. Ferment. Technol. 61: 55–59. F i e l d J.A. and G. L e t t i n g a. 1992. Toxicity of tannic compounds to microorganisms, p. 673–692. In: Hemingway RW and Laks E (ed.) Plant Polyphenols: Synthesis, Properties, Significance. Plenum Press, New York. H a d i T.A., R. B a n e r j e e and B.C. B h a t t a c h a r y a. 1994. Optimization of tannase biosynthesis by a newly isolated Rhizopus oryzae. Bioprocess Eng. 11: 239–243. L e k h a P.K. and B.K. L o n s a n e. 1997. Production and application of tannin acyl hydrolase: state of the art. Adv. Appl. Microbiol. 44: 215–260. L e w i s J.A. and R.L. S t a r k e y. 1969. Decomposition of plant tannins by some soil microorganisms. Soil Science 107: 235–241. M o n d a l K.C. and B.R. P a t i. 2000. Studies on the extracellular tannase from newly isolated Bacillus licheniformis KBR6. J. Basic Microbiol. 40: 223–232. M o n d a l K.C., R. B a n e r j e e and B.R. P a t i. 2000. Tannase production by Bacillus licheniformis. Biotechnol. Lett. 20: 767–769. M o n d a l K.C., D. B a n e r j e e, R. B a n e r j e e and B.R. P a t i. 2001a. Production and characterization of tannase from Bacillus cereus KBR6. J. Gen. Appl. Microbiol. 47: 263–267. M o n d a l K.C., D. B a n e r j e e, M. J a n a and B.R. P a t i. 2001b. Colorimetric assay method for determination of the tannin acyl hydrolase (EC 3.1.1.20) activity. Anal. Biochem. 295: 168–171. M o n d a l K.C., S. S a m a n t a, S. G i r i and B.R. P a t i. 2001c. Distribution of tannic acid degrading microorganisms in the soil and comparative study of tannase from two fungal strains. Acta Microbiol. Pol. 50: 75–82. M u k h e r j e e G. and R. B a n e r j e e. 2003. Production of gallic acid, Biotechnological routes (Part 1). Chimica Oggi/Chemistry Today 21: 59–62. M u k h e r j e e G. and R. B a n e r j e e. 2004. Biosynthesis of tannase and gallic acid from tannin rich substrates by Rhizopus oryzae and Aspergillus foetidus. J. Basic Microbiol. 44: 42–48. P a n d e y A., P. S e l v a k u m a r, C.R. S o c c o l and P. N i g a m. 1999. Solid-state fermentation for the production of industrial enzymes. Curr. Sci. 77:149–162. R a m i r e z - C o r o n e l A., A. D a r v i l l, G. V i n i e g r a - G o n z a l e z and C. A u g u r. 2003. A novel tannase from Aspergillus niger with $-glucosidase activity. Microbiology 149: 2941–2946 S c h a n d e r i S.H. 1970. In: Methods in Food Analysis. Academic Press. New York. p709. S w a i n T. 1965. Plant Biochemistry, J. Bonner and J.E. Varner, eds. Academic Press. New York.

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Polish Journal of Microbiology 2006, Vol. 55, No 4, 303– 307

In Vitro Activity of Synthetic Antimicrobial Peptides Against Candida WOJCIECH KAMYSZ 1*, PIOTR NADOLSKI1, ANNA KÊDZIA2, OSCAR CIRIONI3, FRANCESCO BARCHIESI3, ANDREA GIACOMETTI3, GIORGIO SCALISE3, JERZY £UKASIAK4 and MARCIN OKRÓJ5 1 Department

of Inorganic Chemistry, Medical University of Gdañsk, Gdañsk, Poland; of Oral Microbiology, Medical University of Gdañsk, Gdañsk, Poland; 3 Institute of Infectious Diseases and Public Health, Università Politecnica delle Marche, Ancona, Italy; 4 Department of Physical Chemistry, Medical University of Gdañsk, Gdañsk, Poland; 5 Department of Cell Biology, Medical University of Gdañsk, Gdañsk, Poland 2 Department

Received 6 June 2006, revised 27 September 2006, accepted 9 October 2006 Abstract Yeast-like fungi are the most common cause of fungal infections in humans. Actually, in the age of opportunistic infections and increasing resistance, development of modern antifungal agents becomes a very important challenge. This paper describes synthesis and antimicrobial assay of four naturally occurring peptide antibiotics (aurein 1.2, citropin 1.1, temporin A, uperin 3.6) and three chemically engineered analogues actually passing clinical trials (iseganan, pexiganan, omiganan) against Candida strains isolated from patients with infections of the oral cavity or respiratory tract. The peptides were synthesized using solid-phase method and purified by high-performance liquid chromatography. Biological tests were performed using the broth microdilution method. The antifungal activity of the peptide antibiotics was compared to that of nystatin and amphotericin B. We found synthetic peptides to be generally less potent than amphotericin B or nystatin. However, some of the naturally occurring peptides still retained reasonable antifungal activities which were higher than these of iseganan, pexiganan or omiganan. We think that the naturally occurring peptide antibiotics included in our study can be a good matrix for development of novel antifungal compounds. K e y w o r d s: antimicrobial activity, antimicrobial peptides; fungi

Introduction Nowadays superficial fungal infections emerge as an important problem. Many fungal species, among others Candida, reside either in oral flora or on the skin and mucous membranes. However, these commensal fungi may become pathogenic under certain circumstances, such as immunity disorders in the presence of a particular host factor (Zuber et al., 2000). Neutropenia, cancer, chemotherapy and HIV are additional risk factors for opportunistic infections (Karabinis et al., 1988; Punzon et al., 2002). It is estimated that the yeast-like fungi Candida account for approximately 15% of all nosocomial infections (Vazquez, 2003). The unfavourable efficacy/toxicity ratio of some antifungals (e.g., amphotericin B), limitations in usage (e.g., azoles in the case of neutropenia) and appearance of resistant strains represent a major challenge for developing new methods of treatment (Marchetti et al., 2003). Moreover a comparison of the susceptibility of fungi in these days to that recorded earlier shows a dramatic increase of resistant strains (Nierebinska et al., 1992). New mechanisms of resistance are developed and are propagated rapidly among pathogenic strains. Accumulating resistance factors lead to a multidrug resistance phenotype (MDR), affecting an increasing number of strains (Vanden Bossche et al., 1998; Prasad et al., 2002). Since fungi belong to Eukarya domain, their mechanism of gene expression, replication and some structural features are similar to those occurring in human cells. This is the reason why developing of new, highly specific antifungals may be * Corresponding author: W. Kamysz, Department of Inorganic Chemistry, Faculty of Pharmacy, Medical University of Gdañsk, Al. Gen. Hallera 107, 80-416 Gdañsk, Poland; phone: + 48 58 3493221; fax: + 48 58 3493224; e-mail: [email protected]

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Kamysz W. et al.

difficult. Therefore, researchers look for new agents with different chemical structures and different mechanisms of action to treat fungal infections. Recently much attention has been devoted to natural antimicriobial peptides which are produced by living organisms and act as a very ancient line of innate immunity. Herein we present the results of our studies on anticandidal activity of some naturally occurring antimicrobial peptides: aurein 1.2, citropin 1.1, temporin A and uperin 3.6 (Chia et al., 1999; Rozek et al., 2000; Wegener et al., 1999; Conlon et al., 2004), and peptides actually passing clinical trials: iseganan, omiganan and pexiganan (Kamysz, 2005). Their fungicidal activities are compared to those of two commonly used antifungal drugs, amphotericin B and nystatin. Experimental Materials and Methods Fungal strains The yeasts were isolated from patients with infections of the oral cavity and respiratory tract (Medical University of Gdañsk). The material was inoculated onto the Sabouraud agar (Becton-Dickinson) and incubated under aerobic conditions at room temperature for 72 hours. All Candida strains included in the study were identified according to the classification proposed by de Paiva Martins et al. (2002). We identified 18 strains of Candida albicans, 3 strains of Candida tropicalis, 2 strains of Candida kefyr, 2 strains of Candida krusei, 5 strians of Candida parapsilosis and 4 strains of Candida glabrata. Antibacterial compounds. All the peptides included in the study were synthesized manually by the solid-phase method on Polystyrene AM-RAM resin (0.66 mmol/g; Rapp Polymere, Germany) using the 9-fluorenylmethoxycarbonyl (Fmoc) chemistry (Fields and Noble, 1990). The peptides were synthesized by the following procedure: (i) 5 and 15 min deprotection steps using 20% piperidine in dimethylformamide (DMF) in the presence of 1% Triton; (ii) the coupling reactions carried out with the protected amino acid (Fmoc-AA) diluted in a DMF/N-methyl-2-pyrrolidone (NMP) (1:1, v/v) mixture in the presence of 1% Triton using diisopropylcarbodiimide (DIC) as the coupling reagent in the presence of 1-hydroxybenzotriazole (HOBt) (Fmoc-AA/DIC/HOBt, 1:1:1) for 2 h. The completeness of each coupling reaction was monitored by the chloranil test (Christensen, 1979). The peptides were cleaved from the solid support by trifluoroacetic acid (TFA) in the presence of water (2.5%), and triisopropylsilane (2.5%) as scavengers. The cleaved peptides were precipitated with diethyl ether. Iseganan was cyclized by air oxidation as described in (Chen et al., 2000). The peptides were purified by high-performance liquid chromatography (HPLC) on a Knauer two-pump system with a Kromasil C8 column 10 × 250 mm (5 µm particle diameter, 100 Å pore size) with a flow rate of 5 ml/min, absorbance at 226 nm. The resulting fractions of purity greater than 95 – 98% were tested by HPLC. The peptides were analyzed by matrix-assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF). Sequences, net charges and molecular masses of particular peptides are shown in Table I.

Table I Amino acid sequences of antimicrobial peptides Peptide

Sequence

Molecular mass

Net charage of peptide

Naturally occurring antimicrobial peptides Aurein 1.2

GLFDIIKKIAESF-NH 2

1 479.8 Da

+1

Citropin 1.1

GLFDVIKKVASVIGGL-NH2

1 614.9 Da

+2

Temporin A

FLPLIGRVLSGIL-NH2

1 396.7 Da

+2

Uperin 3.6

GVIDAAKKVVNVLKNLF-NH 2

1 827.2 Da

+3

Chemically engineered peptides Omiganan

ILRWPWWPWRRK-NH2

1 779.2 Da

+5

Iseganan

RGGLCYCRGRFCVCVGR-NH 2

1 904.3 Da

+5

Pexiganan

GIGKFLKKAKKFGKAFVKILKK-NH 2

2 477.2 Da

+10

Nystatin and amphotericin were obtained from Sigma-Aldrich. Antifungal susceptibility testing. The yeasts were tested by the broth microdilution method, performed according to the CLSI (M27-A2) standard guidelines (CLSI guideline, 2002). The antimicrobial peptides, nystatin and amphotericin B, were used to obtain final drug concentrations ranging from 0.25 to 512 µg/ml. A standard inoculum of Candida was diluted to a final concentration of 0.5–1.5 × 103 CFU/ml in microtiter plates. The broth microdilution testing was performed in sterile flat-bottomed 96-well microplates which contained 100 µl of each tested drug (concentration 1 mg/ml). At the beginning of experiment 100 µl of a suspension containing a final concentration of fungi was dispensed into wells of each row containing diluted antifungal agents. Drug-free purity controls and growth controls were included for each experiment. The plates were incubated at 35°C for 48 h. The MIC assumed as the lowest drug concentration which inhibited the growth of tested fungi. Experiments were performed in triplicates.

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Synthetic peptides activity against Candida

Results The antibiotics used commonly in antifungal therapies; nystatin and amphotericin B, tested against our clinical strains were several times more active than the peptide antibiotics included in the study, and their MIC values varied between 0.125 and 8 µg/ml. In some cases, the activity of the peptide antibiotics was found to be promising. In particular, uperin 3.6 and aurein 1.2 exhibited relatively high antifungal activity against C. albicans (Table II). Similarly, the antifungal activity of uperin 3.6, much higher than that of other peptide antibiotics, was noticed for non C. albicans strains (Table III). For 2 out of 3 strains of C. tropicalis, Table II The activity of antimicrobial peptides and conventional antifungals against C. albicans strains Minimal inhibitory concentration (MIC) (µg/ml) Candida albicans (18)

Agent

Range

50%

90%

Aurein 1.2

16–64

16

32

Citropin 1.1

8–256

64

128

Temporin A

16–256

64

128

Uperin 3.6

4–128

16

32

Iseganan

2–128

64

128

Omiganan

32–128

64

128

Pexiganan

16–512

128

256

Amphotericin

0.125–4

0.5

1

Nystatin

0.25–8

2

4

Table III The activity of antimicrobial peptides and conventional antifungals against fungi strains other than C. albicans Minimal inhibitory concentration (MIC) (µg/ml) Candida tropicalis (3) Range

Candida kefyr (2) Range

Candida krusei (2) Range

Candida parapsilosis (5) Range

Candida glabrata (4) Range

16–64

32–256

32–128

16–128

16–64

Citropin 1.1

16–64

64–256

8–64

16–128

2–64

Temporin A

32–256

64–256

64–256

32–512

32–128

Uperin 3.6

2–32

64–256

16–64

8–64

4–32

Agent

Aurein 1.2

Iseganan

32–128

64–256

32–128

2–64

8–128

Omiganan

32–128

128–512

32–128

64–128

64–128

Pexiganan

32–128

128–512

32–128

8–128

16–64

Amphotericin

0.25–0.5

0.5

0.5–1

0.25–2

0.25–1

2–8

1–2

1–2

1–4

2–8

Nystatin

the MIC value of uperin 3.6, it was below 16 mg/ml, whereas that for 90% or 75% strains of C. albicans and C. glabrata respectively was below 16 µg/ml. The peptide antibiotics currently under clinical trials exhibited generally lower activities than the naturally occurring sequences. The MIC range of tested substances for particular strains are presented in Tables II and III. Discussion Summing up our results, it can be stated that generally the peptide antibiotics are several times less active against fungi than the conventional drugs tested, nystatin and amphotericin B. Among the peptides tested, uperin 3.6 was the most effective one towards all Candida strains except for C. kefyr, the finding that

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cannot be considered as a coincidence. Thus, this 17-residue peptide originally isolated from Australian toadlet, Uperoleia mjobergii, appears to be a promising antifungal compound. Interestingly, the remaining naturally occurring peptides exhibit higher anticandida activities than those currently under clinical trials (omiganan, pexiganan and iseganan) as drugs for the treatment of oral mucositis, lung infections, infected diabetic ulcers and catheter infections (Kamysz, 2005). This shows that the peptide antibiotics possess a broad spectrum of activity and even those peptides not previously considered as being worth attention can turn out to be useful in the case of particular pathogens. Fungal infections caused by Candida spp. are especially difficult in treatment. Candida species reside on mucosal surfaces of about 60% of human population. However, in certain conditions, they can cause superficial mucosal infections such as vaginitis,thrush, and esophagitis. At present, an increasing number of Candida and Candida-like infections can be observed. On the other hand, also the number of fungal pathogens resistant to common therapeutic antifungals does increase (Branchini et al., 1998; Barchiesi et al., 1998; Wong-Beringera et al., 2001). Such a situation becomes dangerous, especially in the age of opportunistic infections. For instance, almost a half of AIDS patients develop oral candidiasis and most of responsible strains are drug-resistant (Law et al., 1994; Georgopapadakou, 1998). A similar problem concerns patients undergoing cancer chemotherapy and immunosupression (de Paiva Martins et al., 2002). These immunocompromised patients are also susceptible to severe systemic infections. Taken together, candidiasis is an emerging problem and new effective and safe antifungals are absolutely needed. The naturally occurring peptides studied herein seem to be a good matrix for development of novel antifungal compounds. Modification of their structure by substitution of particular amino acid residues, introduction of D-amino acids or non-peptide entities, coupling of fatty acid chain or shortening the sequence without affecting cytocidal properties could lead to activity enhancement, improvement of pharmacological parameters and finally cost-effectiveness. Literature B a r c h i e s i F., L.F. D i F r a n c e s c o, P. C o m p a g n u c c i, D. A r z e n i, A. G i a c o m e t t i and G. S c a l i s e. 1998. In-vitro interaction of terbinafine with amphotericin B, fluconazole and itraconazole against clinical isolates of Candida albicans. J. Antimicrob. Chemother. 41: 59–65. B r a n c h i n i M.L., F.H. A o k i, A.L. C o l o m b o, H. T a g u c h i, K. Y a m a m o t o and M. M i y a z i. 1998. Effect of antifungal agents on Candida spp. and Pichia anomala isolated from oropharyngeal candidiasis of AIDS patients in a University Hospital in Brazil. Braz. J. Infect. Dis. 2: 187–196. C h e n J., T.J. F a l l a, H. L i u, M.A. H u r s t, C.A. F u j i i, D.A. M o s c a, J.R. E m b r e e, D.J. L o u r y, P.A. R a d e l, C. C h e n g C h a n g, L. G u and J.C. F i d d e s. 2000. Development of protegrins for the treatment and prevention of oral mucositis: structure-activity relationships of synthetic protegrin analogues. Biopolymers 55: 88–98. C h i a B.C., J.A. C a r v e r, T.D. M u l h e r n and J.H. B o w i e. 1999. The solution structure of uperin 3.6, an antibiotic peptide from the granular dorsal glands of the Australian toadlet, Uperoleia mjobergii. J. Pept. Res. 54: 137–145. C h r i s t e n s e n T. 1979. Qualitative test for monitoring coupling completeness in solid phase peptide synthesis using chloranil. Acta Chem. Scand. Series B. (Org. Chem. Biochem.) 33: 763–776. C L S I. 2002. Antifungal susceptibility testing of yeasts. Approved standard-second edition. Document M-27A2. Clinical and Laboratory Standards Institute, Wayne, Pa. C o n l o n J.M., J. K o l o d z i e j e k and N. N o w o t n y. 2004. Antimicrobial peptides from ranid frogs: taxonomic and phylogenetic markers and a potential source of new therapeutic agents. Biochim. Biophys. Acta 1696: 1–14. d e P a i v a M a r t i n s C.A., C.Y. K o g a - I t o and A.O.C. J o r g e. 2002. Presence of Staphylococcus spp. and Candida spp. in the human oral cavity. Braz. J. Microbiol. 33: 236–240. F i e l d s G.B. and R.L. N o b l e. 1990. Solid phase peptide synthesis utilizing 9-fluorenylmethoxycarbonyl amino acids. Int. J. Pept. Protein Res. 35: 161–214. G e o r g o p a p a d a k o u N.H. 1998. Antifungals: mechanism of action and resistance, established and novel drugs. Curr. Opin. Microbiol. 1: 547–557. K a m y s z W. 2005. Are antimicrobial peptides an alternative for conventional antibiotics? Nucl. Med. Rev. Cent. East Eur. 8: 78–86. K a r a b i n i s A., C. H i l l, B. L e c l e r c q, C. T a n c r e d e, D. B a u m e and A. A n d r e m o n t. 1988. Risk factors for candidemia in cancer patients: a case-control study. J. Clin. Microbiol. 26: 429–432. L a w D., C.B. M o o r e, H.M. W a r d l e, L.A. G a n g u l i, M.G. K e a n e y and D.W. D e n n i n g. 1994. High prevalence of antifungal resistance in Candida spp. from patients with AIDS. J. Antimicrob. Chemother. 34: 659–68. M a r c h e t t i O., P. M o r e i l l o n, J.M. E n t e n z a Vo u i l l a m o z, M.P. G l a u s e r, J. B i l l e and D. S a n g l a r d. 2003. Fungicidal synergism of fluconazole and cyclosporine in Candida albicans is not dependent on multidrug efflux transporters encoded by the CDR1, CDR2, CaMDR1, and FLU1 genes. Antimicrob. Agents Chemother. 47: 1565–1570. N i e r e b i ñ s k a E., Z. G w i e Ÿ d z i ñ s k i, S. U r b a n o w s k i and M. P r a c z. 1992. Phenomenon of increased resistance of yeast-like fungi to nystatin. Wiad. Lek. (in Polish) 45: 427–429. P r a s a d R., S.L. P a n w a r and M. S m r i t i. 2002. Drug resistance in yeasts an emerging scenario. Adv. Microb. Physiol. 46: 155–201.

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P u n z o n C., S. R e s i n o, J.M. B e l l o n, M.A. M u n o z - F e r n a n d e z and M. F r e s n o. 2002. Analysis of the systemic immune response in HIV-1-infected patients suffering from opportunistic Candida infection. Eur. Cytokine Netw. 13: 215–223. R o z e k T., K.L. W e g e n e r, J.H. B o w i e, I.N. O l v e r, J.A. C a r v e r, J.C. W a l l a c e and M.J. T y l e r. 2000. The antibiotic and anticancer active aurein peptides from the Australian Bell Frogs Litoria aurea and Litoria raniformis the solution structure of aurein 1.2. Eur. J. Biochem. 267: 5330–5341. V a n d e n B o s s c h e H., F. D r o m e r, I. I m p r o v i s i, M. L o z a n o - C h i u, J.H. R e x and D. S a n g l a r d. 1998. Antifungal drug resistance in pathogenic fungi. Med. Mycol. 36: 119–128. V a z q u e z J.A. 2003. Treatment of candidiasis in hospitalized patients. Curr. Treatm. Options Infect. Dis. 5: 495–506. W e g e n e r K.L., P.A. W a b n i t z, J.A. C a r v e r, J.H. B o w i e, B.C. C h i a, J.C. W a l l a c e and M.J. T y l e r. 1999. Host defence peptides from the skin glands of the Australian blue mountains tree-frog Litoria citropa. Solution structure of the antibacterial peptide citropin 1.1. Eur. J. Biochem. 265: 627–637. W o n g - B e r i n g e r a A., J. H i n d l e r, L. B r a n k o v i c, L. M u e h l b a u e r and L. S t e e l e - M o o r e. 2001. Clinical applicability of antifungal susceptibility testing on non-Candida albicans species in hospitalized patients. Diagn. Microbiol. Infect. Dis. 9: 025–31. Z u b e r T.J. and K. B a d d a m. 2001. Superficial fungal infection of the skin. Where and how it appears help determine therapy. Postgrad. Med. 109: 117–120, 123–126, 131–132.

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Polish Journal of Microbiology 2006, Vol. 55, No 4, 309– 319

Effects of Culture Conditions on Production of Extracellular Laccase by Rhizoctonia praticola GRZEGORZ JANUSZ1, JERZY ROGALSKI1, MAGDALENA BARWIÑSKA1 and JANUSZ SZCZODRAK 2* 1 Department

2Department

of Biochemistry, Maria Curie-Sk³odowska University, Lublin, Poland of Industrial Microbiology, Maria Curie-Sk³odowska University, Lublin, Poland

Received 10 July 2006, revised 22 September 2006, accepted 9 October 2006 Abstract It was found that the soil-dwelling fungus Rhizoctonia praticola 93A was capable to produce laccase in submerged cultures. Effects of culture conditions on the enzyme biosynthesis in shaken flask and aerated bioreactor cultures were evaluated to improve the yields of the process. Production of extracellular laccase was considerably intensified by the addition of Cu2+ to a carbon-limited and nitrogen-sufficient culture medium (C/N = 0.98). When an optimized medium containing glucose (2 g/l) and L-asparagine (1.5 g/l) was used and enzyme synthesis was stimulated by addition of 5 µM Cu2+ before inoculation, maximal laccase activities obtained in a batch cultivation were, approximately, 1000 nkat/l. Under these conditions, addition to the medium of the aromatic inducer 2,5-xylidine (1 mM) led to a 10-fold increase in laccase activity. Laccase productivity in shaken flask cultures was also enhanced (to more than 4000 nkat/l on day 3) by using a medium with the initial pH of 7.5. Such a high value of the optimal medium pH for laccase production by R. praticola is exceptional among the ligninolytic fungi. In fermenter fungal cultures supplemented with cupric ions, the highest laccase activity (about 4000 nkat/l after 3 days’ cultivation) was reached after 24-h incubation using a bioreactor with the aeration rate of 2 l/min, the agitation speed of 200 rpm, and a constant medium pH of 8.0. K e y w o r d s: Rhizoctonia praticola, laccase, inducers, shaken and fermenter cultures

Introduction Laccases (EC 1.10.3.2, p-diphenol:dioxygen oxidoreductases) are copper-containing enzymes which use molecular oxygen to oxidise various aromatic and non-aromatic compounds by the radical-catalysed reaction mechanism (Leonowicz et al., 2001). Because of their broad substrate specificity, soluble or immobilised laccases can potentially be used in textile dye bleaching, pulp delignification, effluent detoxification, production of washing powder, removal of phenolics from effluents, treatment of must, wine and fruit juices, transformation of steroids and antibiotics and in biosensors (Mayer and Staples, 2002). Laccases are commonly found in plants, fungi, insects, and bacteria. The best known laccases producers and a major source of these enzymes are ligninolytic organisms, such as white-rot fungi (Claus, 2004). Unfortunately, production of laccases by fungi is associated with secondary metabolism, the main drawback of which is limited yield of the enzyme under the growth-limiting conditions (Moreira et al., 2000). At present, research and application are somewhat stymied by rather low yields of the enzyme (Gianfreda et al., 1999), as well as by difficulties in efficient heterologous overexpression of laccases in an active form (Jönsson et al., 1997). The problem of increasing the yield of ligninolytic enzymes in fungal cultures is, therefore, of constant interest to researches (Tien and Kirk, 1984). Fungal laccase production is influenced by many typical culturing parameters, such as medium composition, carbon and nitrogen ratio, pH, temperature, and aeration ratio (Arora and Gill, 2001). In white-rot fungi, extracellular laccases are constitutively produced in small amounts (Bollag and Leonowicz, 1984), but their production can be considerably enhanced by a wide variety of inducing substances, mainly aromatic or phenolic compounds related to lignin or lignin derivatives (Farnet et al., 1999). In addition, laccase * Corresponding author: J. Szczodrak, Department of Industrial Microbiology, UMCS, Akademicka 19, 20-032 Lublin, Poland; phone: + 48 81 5375909; fax: +48 81 5375959; e-mail: [email protected]

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production can be modulated by nitrogen and carbon concentrations in the culture medium, as well as by heavy metal ions, especially Cu2+ (Galhaup and Haltrich, 2001; Galhaup et al., 2002). Rhizoctonia praticola is a soil-dwelling plant-pathogenic fungus which causes root rot. It produces an inducible laccase belonging to the class of blue oxidases (Xu et al., 1998; Janusz, 2005). The laccase from this source has a unique property of detoxifying chlorinated phenolic pollutants at a broad range of pH (pH 3.0– 10) and temperatures (5– 55°C) (Shuttleworth et al., 1986; Dec and Bollag, 1990; Cho et al., 1999). The extracellular enzyme from R. praticola had been well characterised (Bollag et al., 1979), but operational conditions favorable to the production of increased amounts of laccase in submerged fungal cultures had not been investigated in detail. Keeping in mind the potential applications of laccase in various bioprocesses, the present study was undertaken to find the best culture conditions for the overproduction of laccase by R. praticola in shaken flasks and aerated fermenter cultures, and to search for the most effective inducers of the enzyme synthesis. The results of this study may contribute to commercialisation of the production of this valuable enzyme. Experimental Materials and Methods Chemicals. Syringaldazine (4-hydroxy-3,5-dimethoxybenzaldehyde azine), ferulic acid (4-hydroxy-3-methoxycinnamic acid), veratric acid (3,4-dimethoxybenzoic acid), o-anisidine (2-methoxyaniline), p-anisidine (4-methoxyaniline), and antifoam B emulsion were supplied by Sigma-Aldrich (St. Louis, MO, USA). 2,5-Xylidine (2,5-dimethylaniline) was purchased from Fluka (Buchs, Switzerland), while L-asparagine came from Merck (Darmstadt, Germany). All other products were of a reagent or an analytical grade and were purchased locally. Fungal strain, media, and culture conditions. Rhizoctonia praticola was obtained from the culture collection of the Laboratory of Soil Microbiology (Pennsylvania State University, USA), and is deposited in the fungal strains collection of the Department of Biochemistry (Maria Curie-Sk³odowska University, Poland) under the strain number 93A. Stock cultures of fungus were stored at 4°C on GPY agar slants (per litre: 1.0 g of glucose, 0.5 g of peptone, 0.1 g of yeast extract, 20 g of agar). For inoculations, pieces of mycelium taken from agar slants were grown for 7 days at 28°C in stationary conical flasks with the Lindeberg-Holm (pH 5.5) (Lindeberg and Holm, 1952) or the Czapek-Dox medium (pH 6.8) (Leonowicz et al., 1984). Mycelial mats were subsequently collected and broken in a Waring blender (three times for 15 s at 10 000 rpm), and the homogenates were used as inocula for shaken flask and aerated bioreactor cultures. After inoculation with 2.5% (v/v) mycelial suspension, the shaken flask cultures were run for up to 14 days at 28°C in 100-ml wide mouth Erlenmeyer flasks (each with 40 ml of the Lindeberg-Holm or the Czapek-Dox medium) placed on an orbital rotary shaker at 180 rpm. Out of the media tested for laccase production, the medium developed by Lindeberg and Holm (1952) was chosen for the experiments and optimised with respect to the initial pH, and carbon (0.5, 1.0, 2.0, 5.0, 10, 15 g/l glucose), nitrogen (0.25, 0.5, 1.0, 1.5, 2.0, 5.0, 10 g/l L-asparagine) and copper concentrations (CuSO 4 × 5 H2O over the range of 0 – 300 µM). Copper concentration in the unsupplemented medium was 0.2 µM. Putative aromatic laccase inducers (ferulic and veratric acids, o- and p-anisidines, and 2,5-xylidine) were dissolved in ethanol as stock solutions and sterilised by filtration (Sterivex-GS filter unit, 0.22 µm; Millipore Corp.). These were added to the growing fungal cultures on the third day of incubation, so that their final concentration in the optimised Lindeberg-Holm medium was 0.02, 1.0, or 5.0 mM. The concentration of ethanol in the growth medium was always less than 0.5% and an equivalent amount of ethanol was added to control flasks without aromatic inducers. Bioreactor-scale cultivations were performed at 28°C in a 2.5 l glass fermenter (BioFlo III, New Brunswick Scientific, Edison, NY, USA) containing 2 l of the optimised Lindeberg-Holm medium. The fermenter was sterilised (121°C, 30 min) and seeded with mycelial suspension (10% of the total volume). The culture was run for 5 days at an aeration rate of 1 or 2 l air/min and stirrer speeds of 100, 200, or 300 rpm. Antifoam B emulsion was added to break the foam. The pH was either not regulated, or was automatically maintained at 7.5, 8.0, and 8.5 value. Samples of the culture media were harvested from shaken flasks or the fermenter at specified time intervals and analysed for laccase activity and pH. Submerged cultures were performed in three independent experiments, and analyses were carried out at least in duplicate. The values reported here are mean values with standard deviations being less than 10% in all cases. Assays. Laccase activity in the culture supernatant was measured spectrophotometrically at 525 nm in a Shimadzu UV-Vis 160A spectrophotometer (Tokyo, Japan) using syringaldazine as a substrate (Leonowicz and Grzywnowicz, 1981). Enzyme and substrate blanks were included. One unit (nano katal, nkat) of laccase activity was defined as the amount of enzyme catalysing the production of one nanomole of coloured product [quinone, gM = 65 000/(M × cm)] per second at 25°C and pH 7.4. The activity was expressed as nano katals per litre of culture medium (nkat/l).

Results and Discussion It had been reported by other authors that secretion of laccase by different white-rot fungi was strictly dependent upon growth conditions (Niku-Paavola et al., 1990; Arora and Gill, 2001; Hess et al., 2002). Our experiments were undertaken for determining the effect of different media, initial pH, nitrogen, carbon and

4

311

Production of laccase by Rhizoctonia praticola

inducer concentrations, as well as agitation and aeration rates of the culture on the increase of laccase production by R. praticola in shaken flask and aerated bioreactor cultures. Enzyme production in shaken cultures. The first step of the studies was determination of the optimum medium composition for extracellular laccase synthesis by R. praticola. Fungal shaken flask cultures were run for 14 days using two culture media (Czapek-Dox or Lindeberg-Holm) commonly used for the production of ligninolytic enzymes by fungi. Typical culture profiles are depicted in Fig. 1. The results clearly showed that in the Lindeberg-Holm medium laccase productivity reached the maximum level (on 8th day) faster than in the Czapek-Dox medium (on 11th day). Enzyme activity in the Lindeberg-Holm medium declined very slowly towards the end of the incubation, in contrast to the Czapek-Dox medium, where a sharp decrease was observed. Taking these results into consideration, the Lindeberg-Holm medium was chosen for further experiments. To determine the optimum C/N ratio in the Lindeberg-Holm medium, different variants of this medium were prepared. The concentration of carbon (added as glucose) varied from 0.5 to 15 g/l (2.78– 83.34 mM) while that of nitrogen (added as L-asparagine) from 0.25 to 10 g/l (1.89– 75.69 mM). The C/N ratio ranged from 0.037 to 44.10. Titres of laccase were measured for 14 days in shaken flask cultures. Fig. 2 illustrates a topographic dependence of laccase activity upon glucose and L-asparagine concentrations. The results indicated that enzyme activity reached its maximum (420 nkat/l on day 6) in cultures with a C/N ratio of 0.98, i.e., containing 2 g/l glucose (11.12 mM) and 1.5 g/l L-asparagine (11.34 mM), respectively. These optimum C and N concentrations were used in further studies. Under these nutritional conditions (a strict carbon limitation and a sufficient amount of nitrogen), the enzyme productivity was six times higher than that obtained in the original Lindeberg-Holm medium used as a control (glucose 10 g/l, L-asparagine 1 g/l; C/N ratio of 7.35). It was found that an increase in glucose concentration from 5 to 15 g/l significantly suppressed laccase activity. In contrast, a rise in nitrogen concentration (from 0.25 to 2.0 g/l) enhanced enzyme synthesis yields. On the basis of the above data, we postulate that synthesis of laccase by R. praticola is the result of a carbon-based regulatory mechanism rather than an outcome of nitrogen limitation. Also, studies of Kwon and Anderson (2001) revealed that expression of laccases in a wheat-pathogenic Fusarium proliferatum isolate was regulated by the nutritional status of the fungus and was greater on a carbon-limited than on a nitrogen-limited medium. Opposite results were obtained in batch and fed-batch cultivations of Trametes

70

Laccase activity (nkat/l)

60

50

40

30

20

10

0 1

2

3

4

5

6

7

8

9

10

11

12

13

14

Cultivation time (days) Fig. 1. Time course of laccase production in Lindeberg-Holm (l) and Czapek-Dox ( o) media during shaken flask cultures of Rhizoctonia praticola.

312

4

cn aregi Agspi n

p ara

( g) i onn(g/l t rraattio cnecennt

eo cno

/l) ion (g entratt i o n ( g e conecn t r a

-1 l )

420 400 380 360 340 320 300 280 260 240 220 200 180 160 140 120 100 80 60 40 20 0

Laccase activity (nkat/l)a

Janusz G. et al.

s c Cluecoc o n

-1 l )

G lu c o s

Fig. 2. Effect of nitrogen (L-asparagine) and carbon (glucose) concentrations on laccase activity of R. praticola grown in shaken flask cultures in Lindeberg-Holm medium. a The

values given are maximum activities reached during cultivation.

pubescens, in case of which high glucose and peptone levels in the medium (40 g/l and 10 g/l, respectively) gave rise to maximum activity of extracellular laccase (Galhaup et al., 2002). Higher nitrogen levels are often required in order to enhance laccase production (Gianfreda et al., 1999; Pointing et al., 2000; Galhaup and Haltrich, 2001; Chen et al., 2003), but with certain fungi nitrogen-limited culture conditions stimulate the formation of this enzyme (Niku-Paavola et al., 1990; Pointing et al., 2000; Baldrian and Gabriel, 2002). The influence of Cu2+ concentration on the production of extracellular laccase in 10-day shaken flask cultures of R. praticola is shown in Fig. 3. Total Cu2+ concentrations varied within the range of 0 to 300 µM, and these ions were added to the culture medium before inoculation with fungal suspension. As the data obtained demonstrate, all Cu2+-containing cultures of R. praticola showed higher laccase activity and produced the enzyme within a shorter incubation period (3 days) when compared with Cu2+-free cultures where maximum laccase production was delayed until day 6. Maximum laccase activity (about 1 000 nkat/l) for this fungus was observed in the presence of 5 µM Cu2+. Increased concentrations of copper (from 10 to 300 µM Cu2+) repressed laccase production (to 54% at 300 µM Cu2+). Cupric ions had been found to be strong stimulants of laccase activity also in experiments carried out by Giardina et al. (1999) as well as Galhaup et al. (2002), in which up to 50 times higher levels of the enzyme were obtained in induced, compared to non-induced, cultures. In the present study, both the time of Cu 2+ ions supplementation and Cu2+ ions concentration were important for obtaining an increased laccase activity. Their supplementation before the inoculation of culture resulted in markedly increased laccase titres. The optimal copper concentration for the enzyme production by R. praticola 93A in shaken flask cultures was 5 µM. Thus, approximately 2.5 times higher yields of the enzyme were obtained in cultures containing Cu2+ as compared to Cu2+-free cultures. The optimal Cu2+ concentration was significantly lower than that (2.0 mM, added after 4 days of incubation) reported by Galhaup and Haltrich (2001) for submerged cultures of T. pubescens, but was still within the range of 2 to 600 µM used in typical cultivation media for the production of laccase both in wild-type and recombinant strains of different basidiomycete fungi (Farnet et al., 1999; Palmieri et al., 2000; Chen et al., 2003). In another study, Baldrian and Gabriel (2002) showed that laccase production by Pleurotus ostreatus increased 8 times during stationary cultivation in nitrogen-limited medium supplemented with 1.0 mM CuSO4 after 12 day incubation. It had also been reported (Palmieri et al., 2000) that the induction of laccase in P. ostreatus occurred when the fungus was cultivated in a nutrient-rich medium supplemented with 150 µM CuSO4 at the time of inoculation. A Cu2+ dose of 1.0 mM was also required for enhancement of laccase synthesis by T. multicolor in bioreactor cultures (Hess et al., 2002).

4

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Production of laccase by Rhizoctonia praticola

1000

Laccase activity (nkat/l)

800

600

400

200

0 0

5

10

15

25

50

100

150

200

300

Cu2+ ions concentration (µM) Fig. 3. Effect of Cu2+ concentration on the production of laccase by R. praticola in shaken flask cultures. Cu2+ ions were added to the medium before inoculation with the fungus (on day 0). Values represent maximum enzyme activity measured on day 6 (control without Cu2+ ) and 3 (samples with Cu2+ ).

Production of laccase is very often enhanced by phenolic and aromatic compounds related to lignin or lignin derivatives such as guaiacol or ferulic acid (Gianfreda et al., 1999). The question arises whether the same substances affect the formation of extracellular laccase by R. praticola in a similar way. To check this, several putative inducers that have a proven effect on laccase synthesis in other fungi were added at varying concentrations (0.02, 1.0 and 5.0 mM) to an actively growing culture of R. praticola on the third day of incubation. Results of these shaken-flask experiments are summarised in Table I. Three out of five different compounds tested, namely, xylidine, and o- and p-anisidines, showed an inductive effect on laccase production at almost all concentrations used. They increased enzyme activity to the maximum level (approximately 10, 2, and 4 times, respectively) at the concentration of 1.0 mM compared to the control culture medium containing no aromatic compound. However, the particular maxima of laccase activity were shifted to day 7 (for xylidine and o-anisidine) or even day 9 (in the case of p-anisidine) compared to the control, which reached its maximum after 3 days of cultivation. Addition of the other substances, i.e., ferulic and veratric acids, to the culture medium reduced laccase levels significantly, and veratric acid, used at the dose of 5.0 mM, caused a slight increase in enzyme titres. Xylidine, the most effective inducer, stimulated laccase production in R. praticola by a factor of 10 at the concentration of 1.0 mM. This contrasts with the results of earlier investigations on this fungus (Bollag and Leonowicz, 1984), in which the authors showed that R. praticola laccase was not affected by xylidine when grown at 24 oC in sugar-rich shaken and stationary cultures. The difference in the findings might be attributed to the higher concentration of xylidine used in the cited experiments. On the other hand, our results were consistent with those of other studies carried out for several fungal strains by different research groups (Galhaup and Haltrich, 2001; Jang et al., 2002; Nyanhongo et al., 2002; Chen et al., 2003; Rancaño et al., 2003), which reported that xylidine, one of the most common and most often used stimulants, elevated laccase production in various Trametes species and in Volvariella volvacea. Moreover, our results are also in accordance with investigations performed by Rogalski et al. (1991), who revealed that out of several tested laccase elicitors, xylidine led to the highest laccase activities in Phlebia radiata. Anisidines are known to induce laccase in Rhizoctonia species (Shuttleworth et al., 1986; Crowe and Olsson, 2001), and such specific induction may operate via receptor-mediated transcriptional activation (Fernandez-Larrea and Stahl, 1996). As expected, we also found p-anisidine to be a powerful inducer of laccase in R. praticola. On the other hand, ferulic and veratric acids repressed laccase production significantly in this fungus although these substances are effective inducers for other fungi, such as V. volvacea,

314

Table I Laccase activity in shaken flask cultures of R. praticola exposed to various aromatic compoundsa

Inducer

Enzyme activity (nkat/l) Cultivation time (day) 2

3

4

5

6

7

8

9

0.00

841.5 ± 61.1

960.1 ± 63.3

764.0 ± 62.6

658.0 ± 45.9

653.0 ± 35.1

572.0 ± 39.0

532.0 ± 32.3

494.6 ± 31.3

Ferulic acid

0.02

–

445.6 ± 34.0

384.2 ± 26.5

365.7 ± 27.2

329.0 ± 22.8

280.6 ± 19.4

245.6 ± 20.5

234.3 ± 20.3

1.00

–

373.1 ± 27.4

328.9 ± 29.1

351.7 ± 32.8

272.7 ± 22.4

226.7 ± 15.8

220.6 ± 20.9

116.4 ± 11.2

5.00

–

146.7 ± 9.2

114.2 ± 9.6

85.0 ± 6.1

50.6 ± 2.5

39.0 ± 3.4

41.8 ± 2.8

0.0 ± 0.0

0.02

–

605.9 ± 43.1

528.1 ± 35.1

495.0 ± 41.2

442.7 ± 31.4

472.7 ± 32.2

242.4 ± 23.9

377.9 ± 25.8

1.00

–

648.2 ± 44.7

582.6 ± 47.4

528.8 ± 40.7

511.1 ± 31.8

475.8 ± 40.7

409.5 ± 35.0

383.9 ± 27.5

Xylidine

o-Anisidine

p-Anisidine

5.00

–

727.4 ± 50.1

857.7 ± 59.9

722.8 ± 59.4

659.5 ± 43.7

578.3 ± 50.0

550.4 ± 35.1

504.4 ± 37.5

0.02

–

1342.8 ± 71.9

1592.6 ± 106.1

1441.8 ± 95.2

1240.7 ± 82.6

1225.3 ± 86.5

904.9 ± 56.9

828.5 ± 49.6

1.00

–

360.5 ± 16.5

468.0 ± 29.9

1015.8 ± 71.2

2942.1 ± 193.4

9254.1 ± 498.7

7283.5 ± 449.9

4029.4 ± 302.7

5.00

–

900.7 ± 51.3

857.8 ± 54.7

811.9 ± 58.9

740.0 ± 47.9

609.1 ± 44.1

603.1 ± 31.5

599.8 ± 30.1

0.02

–

857.2 ± 52.9

1205.3 ± 60.1

938.3 ± 58.4

901.7 ± 56.1

857.3 ± 39.9

685.8 ± 40.4

583.0 ± 34.6

1.00

–

49.2 ± 4.6

481.7 ± 39.9

535.7 ± 37.7

536.5 ± 22.9

1875.4 ± 114.9

1028.3 ± 89.8

624.2 ± 48.6

5.00

–

37.1 ± 1.9

42.1 ± 2.5

42.8 ± 2.2

55.7 ± 3.5

78.8 ± 5.5

29.6 ± 1.6

22.5 ± 1.42

0.02

–

1242.7 ± 61.8

1611.5 ± 91.6

1563.1 ± 99.4

1502.2 ± 92.2

1269.9 ± 88.6

1191.0 ± 71.2

864.5 ± 65.0

1.00

–

489.1 ± 35.1

782.9 ± 50.1

791.0 ± 44.9

1392.5 ± 65.3

2027.9 ± 122.3

3246.4 ± 246.4

3478.9 ± 216.7

5.00

–

873.7 ± 54.8

1106.3 ± 64.8

923.0 ± 64.9

729.1 ± 45.5

717.4 ± 59.2

509.2 ± 32.3

62.0 ± 5.4

Janusz G. et al.

Control

Veratric acid

a

Concentration (mM)

Lindeberg-Holm medium with optimised carbon and nitrogen concentrations and 5 µM Cu2+ (added before inoculation, i.e., on day 0) was used. Each inducer was introduced into the medium on the third day of cultivation.

4

4

315

Production of laccase by Rhizoctonia praticola 4500

A

Laccase activity (nkat/l)

4000 3500 3000 2500 2000 1500 1000 500 0

0

1

2

3

4

5

6

7

Time (day) 9.5

B

9

Initial pH

8.5 8 7.5 7 6.5 6 5.5 0

1

2

3

4

5

6

7

Time (day) Fig. 4. The effect of initial medium pH (A) and changes in the pH of culture medium as a function of time (B) on laccase biosynthesis during shaken flask cultures of R. praticola in optimised Lindeberg-Holm medium containing Cu2+ (5 µM, added before inoculation). Initial pH of the original medium was 5.5. Symbols: pH 5.5 ( o), pH 6.0 (●), pH 6.5 ( ), pH 7.0 (▲), pH 7.5 (¨), pH 8.0 (n), pH 8.5 ( ).

¨



Marasmius quercophilus, Pleurotus eryngii, and P. radiata (Rogalski and Leonowicz, 1992; Muñoz et al., 1997; Farnet et al.,1999; Chen et al., 2003). In preliminary experiments, the pH of the original Lindeberg-Holm medium was 5.5. To find the most suitable conditions for laccase production in shaken flask cultures, we determined enzyme activity at pHs between 5.5 to 8.5. It follows clearly from the data in Fig. 4 that the initial medium pH of 7.5 was the most conducive to enzyme production, yielding, in shaken flask cultures, over 4000 nano katals of laccase per litre of culture broth after 3 days of cultivation. This activity was approximately three times higher than that obtained at the initial pH of 5.5 (about 1400 nkat/l).

316

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Janusz G. et al. 1200

9

800

8.5 pH

Laccase activity (nkat/l)

1000

600

400

8

200

0

7.5 0

1

2

3 Time (day)

4

5

Fig. 5. Laccase production (▲, ●) and changes in pH (∆, o) during fermenter cultures of R. praticola in optimised Lindeberg-Holm medium containing Cu2+ (▲, ∆) and in a Cu2+-free medium (●, o). Cu 2+ ions (5 :M) were added to the medium before inoculation. Culture conditions: pH-value not controlled (the initial medium pH was 7.5); stirrer speed 300 rpm; aeration rate 1 l/min.

The results also demonstrated the high correlation between laccase production and changes in the medium pH during cultivation. In all cases, the enzyme was detected in the medium when the fungus alkalised the medium to a pH above 7.5. Such a high value of the initial medium pH (7.5), which is indispensable for the production of laccase by R. praticola, is exceptional among ligninolytic fungi. Most fungal laccases reach their maximum activity when the initial pH of the nutrient medium ranges from 4 to 6 (Galhaup et al., 2002; Jang et al., 2002; Chen et al., 2003). A newly isolated strain of T. modesta resembles R. praticola in requiring a relatively high initial pH (i.e. 6.95) to produce the highest titres of laccase (Nyanhongo et al., 2002); however, the laccase from R. praticola is of particular interest because it also has an unusually high pH optimum for its catalytic activity. Most fungal laccases have optima in the acidic region (below pH 5), whereas laccase isolated from R. praticola as well as enzymes obtained lately from Melanocarpus albomyces and Pleurotus ostreatus have their optimum activity at a neutral pH (Bollag and Leonowicz, 1984; Shuttleworth et al., 1986; Kiiskinen et al., 2002; Pozdnyakova et al., 2004 ). Enzyme production in fermenter cultures. Laccase production by R. praticola was also carried out in 2-litre batches in a 2.5 l fermenter. Biosynthesis conditions were fixed, taking into account the previous results obtained in agitated flask cultures (the optimised Lindeberg-Holm production medium with the determined Cu2+ dose and C/N ratio, initial pH of the medium, and incubation temperature). During the first stage, we compared laccase activities achieved by R. praticola in bioreactor cultures performed on the optimised Lindeberg-Holm medium with and without cupric ions. As Fig. 5 shows, the fungus reached the highest titres of laccase activity (over 1000 nkat/l) when it was cultivated in the presence of the fixed dose of Cu2+. This activity was two times higher than in the Cu 2+-free culture, but still much lower (even by 75%) than that obtained in optimised shake flask cultures. In the fermenter culture, however, the enzyme productivity reached maximum levels within a shorter incubation period (2 days). In order to increase laccase yields in aerated fermenter cultures, some operating parameters affecting enzyme production by R. praticola were optimised. The effects of increasing the stirrer speed (from 100 to 300 rpm) and the aeration rate (from 1 to 2 l per min) on laccase production are shown in Fig. 6. The results indicate that laccase activity reached maximum levels (2400 nkat/l on day 2) in cultures with the aeration rate of 2 l/min and the agitation speed of 200 rpm. The effects of stirrer speed and aeration rate agree with data obtained for fungal laccases in other studies (Moreira et al., 2000; Galhaup et al., 2002; Rancaño et al., 2003). It is significant that in all fermenter cultures carried out without pH regulation, the fungus alkalised the medium during cultivation, and pH rose from 7.5 to above 8.8. Therefore, the influence of stabilisation of

4

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Production of laccase by Rhizoctonia praticola 2200

9.2

A

1800

8.8

1600 1400

8.4

1200

pH

Laccase activity (nkat/l)

2000

1000

8

800 600

7.6

400 200 0 0

1

2

3

4

5

7.2

Time (day)

B

2400

9

2200 1800 8.5

1600 1400

pH

Laccase activity (nkat/l)

2000

1200 1000 8

800 600 400 200

7.5

0 0

1

2

3

4

5

Time (day) Fig. 6. Time course of laccase production (▲, ■, ●) and changes in pH (∆, ¨, o) during fermenter cultures of R. praticola in optimized Lindeberg-Holm medium. The fungal cultures were run at the aeration rates of 1 (A) and 2 l (B) air per min and with the stirrer speeds of 100 (■, ¨), 200 (▲, ∆), and 300 rpm (●, o). Cu2+ ions (5 :M) were added to the medium before inoculation. Culture conditions: pH value not controlled (the initial medium pH was 7.5).

medium pH after 24 h incubation on laccase activity had to be estimated (Fig. 7). The obtained data show that the use of an automatic pH control set at pH 8 significantly induced laccase productivity. Under these conditions, the highest enzyme activity of 4000 nkat/l was reached after 3-day incubation. It was almost two times higher than that obtained in a fermenter culture with a non-stabilised pH value. Consequently, the application of optimised medium and culture conditions as well as the use of a bioreactor maintaining the pH value on the same level enabled us to obtain an increased laccase yield (in average 4000 nkat/l) within a short time (2– 3 days), both in shake flask and aerated fermenter cultures of R. praticola. In conclusion, the present study reveals that the soil-dwelling plant-pathogenic fungus R. praticola, strain 93A, represents a new source of extracellular laccase which requires high values of the initial medium pH (7.5 to 8.0) to produce the highest titres of the enzyme. The study also shows that an appropriate combination of culture conditions, i.e., the use of a C-limited medium supplemented with an adequate dose of Cu2+ ions before inoculation, stabilisation of medium pH, and maintenance of stirring speed and aeration rate at

318

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Laccase activity (nkat/l)

3500 3000 2500 2000 1500 1000 500 0 0

1

2

3 Time (day)

4

5

6

Fig. 7. Effect of stabilisation of medium pH on laccase production by R. praticola in fermenter cultures in optimised Lindeberg-Holm medium. The pH was automatically maintained at a value of 7.5 (o), 8.0 (●) and 8.5 (∆) after 24-h incubation. Cu2+ ions (5 :M) were added to the medium before inoculation. Culture conditions: stirrer speed 200 rpm; aeration rate 2 l/min.

the optimised level increases laccase production by R. praticola 93A within a short period of incubation. These results applied in a large scale fermenter culture could prove to be of an economic advantage. Future experiments with the use of immobilised mycelium could lead to determining the optimal conditions for repeated batch or continuous fungal cultures, resulting in another potential cost reduction in laccase production. Acknowledgments. This work was financially supported by the Polish State Committee for Scientific Research (Grant PBZKBN-098/T09/2003).

Literature A r o r a D.S. and P.K. G i l l. 2001. Effects of various media and supplements on laccase production by some white rot fungi. Biores. Technol. 77: 89–91. B a l d r i a n P. and J. G a b r i e l. 2002. Copper and cadmium increase laccase activity in Pleurotus ostreatus. FEMS Microbiol. Lett. 206: 69–74. B o l l a g J.-M. and A. L e o n o w i c z. 1984. Comparative studies of extracellular fungal laccases. Appl. Environ. Microbiol. 48: 849–854. B o l l a g J.-M., R.D. S j o b l a d and S.-Y. L i u. 1979. Characterization of an enzyme from Rhizoctonia praticola which polymerizes phenolic compounds. Can. J. Microbiol. 25: 229–233. C h e n S., D. M a, W. G e and J.A. B u s w e l l. 2003. Induction of laccase activity in the edible straw mushroom, Volvariella volvacea. FEMS Microbiol. Lett. 218: 143–148. C h o N.-S., J. R o g a l s k i, M. J a s z e k, J. L u t e r e k, M. W o j t a s - W a s i l e w s k a, E. M a l a r c z y k, M. F i n k - B o o t s and A. L e o n o w i c z. 1999. Effect of coniferyl alcohol addition on removal of chlorophenols from water effluent by fungal laccase. J. Wood Sci. 45: 174–178. C l a u s H. 2004. Laccases: structure, reactions, distribution. Micron 35: 93–96. C r o w e J.D. and S. O l s s o n. 2001. Induction of laccase activity in Rhizoctonia solani by antagonistic Pseudomonas fluorescens strains and a range of chemical treatments. Appl. Environ. Microbiol. 67: 2088–2094. D e c J. and J.-M. B o l l a g. 1990. Detoxification of substituted phenols by oxidoreductive enzymes through polymerization reactions. Arch. Environ. Contamin. Toxicol. 19: 543–550. F a r n e t A.-M., S. T a g g e r and J. L e P e t i t. 1999. Effects of copper and aromatic inducers on the laccases of the white-rot fungus Marasmius quercophilus. C.R. Acad. Sci. Paris, Sciences de la vie/Life Sciences 322: 499–503. F e r n a n d e z - L a r r e a J. and U. S t a h l. 1996. Isolation and characterization of a laccase gene from Podospora anserina. Mol. Gen. Genet. 252: 539–551.

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G a l h a u p C. and D. H a l t r i c h. 2001. Enhanced formation of laccase activity by the white-rot fungus Trametes pubescens in the presence of copper. Appl. Microbiol. Biotechnol. 56: 225–232. G a l h a u p C., H. W a g n e r, B. H i n t e r s t o i s s e r and D. H a l t r i c h. 2002. Increased production of laccase by the wooddegrading basidiomycete Trametes pubescens. Enzyme Microb. Technol. 30: 529–536. G i a n f r e d a L., F. X u and J.-M. B o l l a g. 1999. Laccases: a useful group of oxidoreductive enzymes. Biorem. J. 3: 1–25. G i a r d i n a P., G. P a l m i e r i, A. S c a l o n i, B. F o n t a n e l l a, V. F a r a c o, G. C e n n a m o and G. S a n n i a. 1999. Protein and gene structure of a blue laccase from Pleurotus ostreatus. Biochem. J. 34: 655–663. H e s s J., C. L e i t n e r, C. G a l h a u p, K.D. K u l b e, B. H i n t e r s t o i s s e r, M. S t e i n w e n d e r and D. H a l t r i c h. 2002. Enhanced formation of extracellular laccase activity by the white-rot fungus Trametes multicolor. Appl. Biochem. Biotechnol. 98–100: 229–241. J a n g M.Y., W.R. R y u and M.H. C h o. 2002. Laccase production from repeated batch cultures using free mycelia of Trametes sp. Enzyme Microb. Technol. 30: 741–746. J a n u s z G. 2005. Comparative studies of fungal laccases. Ph.D. Thesis, Maria Curie-Sklodowska University, Lublin, Poland. J ö n s s o n L.J., M. S a l o h e i m o and M. P e n t t i l ä. 1997. Laccase from the white-rot fungus Trametes versicolor: cDNA cloning of lcc1 and expression in Pichia pastoris. Curr. Genet. 32: 425–430. K i i s k i n e n L.L., L. V i i k a r i and K. K r u u s. 2002. Purification and characterization of a novel laccase from the ascomycete Melanocarpus albomyces. Appl. Microbiol. Biotechnol. 59: 198–204. K w o n S.-I. and A.J. A n d e r s o n. 2001. Laccase isozymes: production by an opportunistic pathogen, a Fusarium proliferatum isolate from wheat. Physiol. Mol. Plant Pathol. 59: 235–242. L e o n o w i c z A., N.-S. C h o, J. L u t e r e k, A. W i l k o l a z k a, M. W o j t a s - W a s i l e w s k a, A. M a t u s z e w s k a, M. H o f r i c h t e r, D. W e s e n b e r g and J. R o g a l s k i. 2001. Fungal laccase: properties and activity on lignin. J. Basic Microbiol. 41: 185–227. L e o n o w i c z A., R.U. E d g e h i l l and J.-M. B o l l a g. 1984. The effect of pH on the transformation of syringic and vanillic acids by the laccases of Rhizoctonia praticola and Trametes versicolor. Arch. Microbiol. 137: 89–96. L e o n o w i c z A. and K. G r z y w n o w i c z. 1981. Quantitative estimation of laccase forms in some white-rot fungi using syringaldazine as a substrate. Enzyme Microb. Technol. 3: 55–58. L i n d e b e r g G. and G. H o l m. 1952. Occurrence of tyrosinase and laccase in fruit bodies and mycelia of some Hymenomycetes. Physiol. Plant. 5: 100–114. M a y e r A.M. and R.C. S t a p l e s. 2002. Laccase: new functions for an old enzyme – a review. Phytochemistry 60: 551–565. M o r e i r a M.T., A. T o r r a d o, G. F e i j o o and J.M. L e m a. 2000. Manganese peroxidase production by Bjerkandera sp. BOS55. 2. Operation in stirrer tank reactors. Bioprocess Eng. 23: 657–661. M u ñ o z C., F. G u i l l é n, A.T. M a r t i n e z and M.J. M a r t i n e z. 1997. Induction and characterization of laccase in the ligninolytic fungus Pleurotus eryngii. Curr. Microbiol. 34: 1–5. N i k u - P a a v o l a M.-L., E. K a r h u n e n, A. K a n t e l i n e n, L. V i i k a r i, T. L u n d e l l and A. H a t a k k a. 1990. The effect of culture conditions on the production of lignin modifying enzymes by the white-rot fungus Phlebia radiata. J. Biotechnol. 13: 211–221. N y a n h o n g o G.S., J. G o m e s, G. G ü b i t z, R. Z v a u y a, J.S. R e a d and W. S t e i n e r. 2002. Production of laccase by a newly isolated strain of Trametes modesta. Biores. Technol. 84: 259–263. P a l m i e r i G., P. G i a r d i n a, C. B i a n c o, B. F o n t a n e l l a and G. S a n n i a. 2000. Copper induction of laccase isoenzymes in the ligninolytic fungus Pleurotus ostreatus. Appl. Environ. Microbiol. 66: 920–924. P o i n t i n g S.B., E.B.G. J o n e s and L.L.P. V r i j m o e d. 2000. Optimization of laccase production by Pycnoporus sanguineus in submerged liquid culture. Mycologia 92: 139–144. P o z d n y a k o v a N.N., J. R o d a k i e w i c z - N o w a k and O.V. T u r k o v s k a y a. 2004. Catalytic properties of yellow laccase from Pleurotus ostreatus D1. J. Mol. Cat. B- Enzymatic 30: 19–24. R a n c a ñ o G., M. L o r e n z o, N. M o l a r e s, S. R o d r i g u e z C o u t o and Á. S a n r o m á n. 2003. Production of laccase by Trametes versicolor in an airlift fermentor. Proc. Biochem. 39: 467–473. R o g a l s k i J. and A. L e o n o w i c z. 1992. Phlebia radiata laccase forms induced by veratric acid and xylidine in relation to lignin peroxidase and manganese-dependent peroxidase. Acta Biotechnol. 12: 213–221. R o g a l s k i J., T. L u n d e l l, A. L e o n o w i c z and A. H a t a k k a. 1991. Influence of aromatic compounds and lignin on production of ligninolytic enzymes by Phlebia radiata. Phytochemistry 30: 2869–2872. S h u t t l e w o r t h K.L., L. P o s t i e and J.-M. B o l l a g. 1986. Production of induced laccase by the fungus Rhizoctonia praticola. Can. J. Microbiol. 32: 867–870. T i e n M. and T.K. K i r k. 1984. Lignin-degrading enzyme from Phanerochaete chrysosporium: Purification, characterization, and catalytic properties of a unique H 2O2-requiring oxygenase. Proc. Natl. Acad. Sci. USA 81: 2280–2284. X u F., R.M. B e r k a, J.A. W a h l e i t h n e r, B.A. N e l s o n, J.R. S h u s t e r, S.H. B r o w n, A.E. P a l m e r and E.I. S o l o m o n. 1998. Site-directed mutations in fungal laccase: effect on redox potential, activity and pH profile. Biochem. J. 334: 63–70.

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Polish Journal of Microbiology 2006, Vol. 55, No 4, 321–331

Growth of Penicillium verrucosum and Production of Ochratoxin A on Nonsterilized Wheat Grain Incubated at Different Temperatures and Water Content JANUSZ CZABAN1* , BARBARA WRÓBLEWSKA1, ANNA STOCHMAL2 and BOGDAN JANDA2 1 Department

of Agricultural Microbiology, 2 Department of Biochemistry and Crop Quality, Institute of Soil Science and Plant Cultivation – State Research Institute, Pu³awy, Poland Received 9 June 2006, revised 1 September 2006, accepted 12 September 2006 Abstract

The results of two experiments with wheat grain inoculated with Penicillium verrucosum are reported. In Experiment I, wheat grain, containing 10, 20 and 30% water, was incubated for 2 weeks at 10, 15, 21 and 28°C. In Experiment II, wheat grain, containing 14, 16, 18, 20 and 22% water, was incubated for 2 weeks at 10, 15, and 20°C. At initial moisture content (IMC) of the wheat grain up to 16% neither P. verrucosum growth nor ochratoxin A (OTA) formation were observed. In the range of IMC from 18% to 22% both the fungal growth and OTA synthesis were distinct, and the parameters were higher at higher temperature in the range 10 – 21°C. A temperature of 28°C was probably too high for proper metabolism of the fungus, including OTA formation. OTA formation was distinctly related to P. verrucosum abundance in the temperature range 10 – 21°C, expressed both as the counts of fungal colony forming units (CFU) on agar DYSG medium and diameters of the fungal colonies growing around the wheat kernels placed on the surface of DYSG medium. OTA formation and abundance of P. verrucosum were negatively correlated with the percentage of wheat kernels, placed on DYSG medium, with growing colonies of fungi different from P. verrucosum. CFU counts of P. verrucosum on the wheat grain were significantly related to the diameter of the fungal colonies growing around the wheat kernels placed on DYSG medium. The relationship is described by an exponential regression equation. K e y w o r d s: Penicillium verrucosum abundance, ochratoxin A, wheat grain

Introduction Penicillium verrucosum Dierckx is the only known species of Penicillium able to produce nefrotoxic, carcinogenic, teratogenic and immunotoxic ochratoxin A (OTA) on grain and cereal products (Elmholt et al., 1999; Lund and Frisvad, 2003). Growth of P. verrucosum appears to be the major cause of grain contamination by OTA in European countries (Arroyo et al., 2005; Lund and Frisvad, 2003; Miller, 1995). Aspergillus ochraceous and several related species that are considered rare on grain also produce OTA (Miller, 1995; Pardo et al., 2004; Ramos et al., 1998). There is the general opinion that species of Aspergillus genus are important in OTA production in warmer regions, while in colder regions, especially of temperate climates, only the activity of P. verrucosum is significant (Elmholt and Hestbjerg, 1999; Lindblad et al., 2004; Park et al., 2005). Moisture content and temperature are the most important variables in determining growth and rate of mycotoxin production by fungi in stored grain ecosystems (Cairns-Fuller et al., 2005; Pardo et al., 2004; Ramos et al., 1998). P. verrucosum is a typical xerophilic storage species that is able to thrive at relatively low water activities (Arroyo et al., 2005; Axberg et al., 1997; Moss, 1996; Ramakrishna et al., 1996). Various other fungal species present on grain differ in their growth responses to temperature and water activity of the substrate (Haasum and Nielsen, 1998; Moss, 1996; Pardo et al., 2004; Ramakrishna et al., * Corresponding author: J. Czaban, Dept. of Agricultural Microbiology, Institute of Soil Science and Plant Cultivation – State Research Institute, 8 Czartoryskich, 24-100 Pu³awy, Poland; e-mail: [email protected]

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4

1996). Ramakrishna et al. (1996) reported, on the basis of their own results and results of the cited authors, that some of these fungi might influence the colonization of cereal grains by P. verrucosum and subsequent OTA production, but all those studies were conducted with sterilized grain. Therefore, there is a lack of information about growth of P. verrucosum and OTA production by the fungus competing with natural microbial contaminants on nonsterilized grain under different environmental conditions. Development of the selective and diagnostic medium DYSG for the detection of P. verrucosum in foods and feeds (Frisvad et al., 1992) offers opportunities for estimating the abundance of the fungus in mixed populations of fungi, using the classical dilution plating technique or direct plating (Elmholt et al., 1999; Czaban and Wróblewska, 2006). The objective of this study was to determine the effects of different temperatures and the moisture contents on growth of a strain of P. verrucosum, isolated from Polish rye, and on its ability to ochratoxin A formation on nonsterile winter wheat grain, inoculated with the fungus. Experimental Materials and Methods Fungal strain of P. verrucosum was isolated from grain of rye (LPH63 DE), harvested in 2002. The fungal growth media and conditions. P. verrucosum was isolated and grown on DYSG medium (Lund and Frisvad, 2003; Czaban and Wróblewska, 2006). Grain of winter wheat cv. Mewa and cv. Finezja, harvested in 2004 year, were used in Experiment I and Experiment II, respectively. Basing on of the method described by Lund and Frisvad (2003), the grain was not contaminated with P. verrucosum and other ochratoxinogenic fungi. The grain contains 6.1% (after drying at 40°C) and 12.1% water for Mewa and Finezja, respectively. The initial moisture content (IMC) of the grain was adjusted to 10, 20 and 30% in Experiment I, and to 14, 16, 18, 20 and 22% in Experiment II by adding sterile water (and suspension of P. verrucosum spores – see below) to beakers, containing 100 g samples of the grain. The samples were mixed, placed into closed plastic bags and kept for 24 hours in a refrigerator at 4°C. The beakers with moistened grain were shaken several times during the cold storage and the grain was mixed again to distribute moisture and the inoculum of P. verrucosum before incubation. The grain in each beaker was inoculated with 1 ml of spore suspensions containing 0.80 × 107 or 0.64 × 107 spores, to obtain 2 × 10 5 or 1.6 × 105 spores per 1 g of wheat grain in Experiment I and Experiment II, respectively. The beakers with inoculated grain were incubated in ventilated plastic bags in darkness at 10, 15, 21 and 28°C in Experiment I, and at 10, 15 and 20°C in Experiment II, for two weeks. Determination of fungal abundance on the incubated grain. After the incubation, fungi were isolated from the grain by shaking with a solution containing 0.85% NaCl, 0.1% peptone and 0.1% Tween 80 for 30 min (Frisvad, 1986b). The number of colony forming units (CFU) of both P. verrucosum and all fungi or only P. verrucosum in the resulting suspensions were determined by dilution plating on Martin’s (Martin, 1950) and DYSG (Lund and Frisvad, 2003; Czaban and Wróblewska, 2006) media, respectively. The Petri dishes with Martin’s medium were incubated at 27°C for 4 days. The plates with DYSG medium were incubated in the dark at 20°C for 5 days. After 7 days P. verrucosum colonies had developed their characteristic terracotta-colored pigmentation on the DYSG reverse, caused by synthesis of an anthraquinone (Elmholt et al., 1999; Frisvad et al., 2005). The determinations were done with four replicates. Abundance of P. verrucosum on the wheat grain was also determined by the measurement of diameters of P. verrucosum colonies growing around the wheat kernels placed on agar DYSG medium, according to Czaban and Wróblewska (2006). Twenty seven incubated wheat kernels were placed on DYSG medium (9 kernels per 1 Petri dish). The diameters of the fungal colonies were measured across the width of the kernels after 5 days of incubation in the dark at 20°C. Although Czaban and Wróblewska (2006) used 22.5°C for incubation of the Petri dishes with wheat kernels contaminated with P. verrucosum, temperature of 20°C was chosen in present studies, because it is recommended in standardized method No. 152 of The Nordic Committee on Food Analysis for growth of the fungus (NMKL, 2005). Competitive relation of P. verrucosum and other fungi was assessed on the basis of the percentage of twenty-seven incubated wheat kernels, placed on agar DYSG medium, with growing colonies (beside the kernels) of fungi different from P. verrucosum. Radial mycelial growth of P. verrucosum was measured on 9 cm diameter Petri plates with agar DYSG medium, after 3, 5, 7, 9, 11 and 14 days of incubation at 10, 15, 21 and 28°C (Experiment A) and after 14 days at 15, 22.5, 25, 27.5 and 30°C (Experiment B). Each plate was point-inoculated in the center with a suspension of fungal spores. The measurements were done in five replicates. Ochratoxin A (OTA) extraction. After the incubation, the wheat grain, collected from the beakers, was ground and carefully mixed. Samples (10 g) of the mixture were homogenized for 1 min with 80 ml of 80% acetonitrile and then extracted for 16 h at room temperature with occasional shaking. The extract was filtered through filter paper. The residual material was incubated for 1 h with 50 ml of 80% acetonitrile, and then it was sonicated for 3 min and filtered through filter paper. The both filtrates were combined and evaporated at a temperature not exceeded 40°C to remove acetonitrile. The condensed extract was diluted with water and cleaned-up by a SEP-PAK C18 microcolumn, which was rinsed with water and then with 100% methanol. The extracts were evaporated to dryness and solubilized in 1 ml of 100% methanol. The samples were kept in dark vials at 5°C. OTA content determination. Measurements with a HPLC Waters system equipped with a 474 fluorescence detector (excitation 230 nm, emission 470 nm) were done. The samples were separated using a C18 Eurospher-100 column (5 µm, 250 × 4 mm) at 35°C. Sample volume was 50 µl. The analysis was performed under isocratic conditions at a flow rate of 1 ml/min of the mobile phase (CH 3CN:H2O:H3PO4; 54:45:1) for 15 min. Ochratoxin A (Sigma) was used as a standard. The retention time of OTA was approximately 7.8 min. The measurement was done in triplicates. Statistical evaluations. The data were subjected to one-way analysis of variance and the means were separated with Student’s t-test (at P = 0.0001 for OTA concentration and fungal colony determinations, and at P = 0.05 for fungal CFU number determina-

4

Penicillium verrucosum growth and OTA production

323

tions). Although concentrations of OTA and CFU numbers are shown on a logarithmical scale in Fig. 1 and 2, only basic values were used in these statistical analyses. Statistical evaluation of significant differences between the percentages of wheat grains surrounded by colonies of fungi different from P. verrucosum on DYSG medium were calculated according to Czaban and Wróblewska (2006). For estimation of all examined relationships, linear correlation analysis as well as linear and exponential regression analyses were applied. Together with correlation coefficients (r), the probabilities (P) and the number of replicates (n) are presented. The determination coefficient (R2) is presented with linear and exponential regression equations.

Results and Discussion In the presented studies, the wheat grain was inoculated with a large number of P. verrucosum spores, 10 times greater than the CFU number of other fungi on the grain, to obtain a distinct dominance of P. verrucosum at the beginning of the incubation period to be sure that P. verrucosum would grow and produce OTA on nonsterile grain. In Experiment I, three levels of IMC – initial moisture content (10, 20 and 30%) of the incubated wheat grain (after inoculation with P. verrucosum) were chosen. The first level (10%) of IMC in the grain was chosen as too low for growth of all fungi. As expected, P. verrucosum and other fungi did not reveal any symptoms of growth or metabolic activities on the grain with IMC 10%. In this case, the numbers of the fungal CFU were at similar level as the inoculation dose (Fig. 1A, 1B), and P. verrucosum colony diameters around the kernels placed on the DYSG medium were small (Fig. 1C). Also, no OTA was detected in the grain (Fig. 1E). The next level (20%) of IMC in the grain was chosen as close to “the boundary value”, which enables xerophilic storage moulds (including P. verrucosum) to grow, but it prevents the growth of field fungi (French et al., 1989). Indeed, except for the grain incubated at 10°C, the intensive growth of P. verrucosum was noted in all series with this moisture content, expressed as high CFU numbers of the fungus (Fig 1A and 1B) and its big colony diameters (Fig. 1C). However, distinct growth of fungi different from P. verrucosum on the grain was not observed (Fig. 1B). Only single colonies could be detected on plates with Martin’s medium in the case of lower dilutions, so their CFU numbers could not be determined, because it was masked by high numbers of P. verrucosum. The low percentage (in comparison to the series with 10% and 30% IMC) of the wheat kernels (placed on DYSG medium) with colonies of fungi different from P. verrucosum (Fig. 1D) also confirms the distinct domination of P. verrucosum in the series with 20% IMC at 15, 21 and 28°C. In these series, very high numbers of CFU of P. verrucosum (Fig. 1A and 1B) and large diameter of the fungal colonies (Fig. 1C) were related to very high production of OTA by the fungus at 15 and 21°C, but not at 28°C (Fig. 1E). This temperature was probably too high for efficient metabolism of the fungus. This assumption was confirmed by radial mycelial growth in Experiment A (Fig. 1F) and in Experiment B (Fig. 1G) on agar DYSG medium. Although in Experiment A the diameters of P. verrucosum colonies were the largest at 28°C, the characteristic for the fungus terracotta dye production on the reversal side of its colonies was inhibited to a certain degree. At this temperature, the obverse of the fungal colony was also changed. In radial mycelial growth Experiment B the terracotta dye production was also decreased at 27.5°C and completely inhibited at 30°C. Data of Experiment I are consistent with results of Myslivec and Tuite (1970) and Myslivec et al. (1975). They reported that P. cyclopium could not grow at temperatures higher than 30°C. Similarly, Northolt et al. (1979) found that strains of P. viridicatum and P. cyclopium could not produce OTA at 31°C. The correct present taxonomic classification of these strains most likely should be P. verrucosum (Domsch et al., 1986; Frisvad, 1986a; Frisvad and Filtenborg, 1989). Results of some studies (Ramakrishna et al., 1996) with a strain of P. verrucosum obtained from the Rothamsted Experimental Station culture collection confirm that a temperature of 30°C is not favourable to OTA formation by this fungus and its growth. After 2-weeks of incubation of wheat grain with the highest level of IMC (30%) distinct increases of CFU number of molds differing from P. verrucosum was noted (Fig. 1B). Good growth of these fungi reduced growth of P. verrucosum. CFU numbers of P. verrucosum were lower in comparison to series with 20% IMC (Fig. 1A and 1B). It was particularly noticeable in the case of the wheat grain incubated at 28°C. Probably, the competition effect of P. verrucosum on the grain was weak at this temperature. Although the inoculum dose of P. verrucosum was 10 times greater than the CFU number of other fungi, no one colony of this fungus was detected on Martin’s medium (Fig. 1B) and diameters of P. verrucosum colonies surrounding wheat kernels placed on DYSG medium were even smaller than those of the series with 10% IMC (Fig. 1C). Also, just as in the case of 20% IMC, OTA concentration in the grain incubated at 28°C was much lower than in the grain incubated at 15°C and 21°C (Fig. 1E). As mentioned in the previous paragraph,

324

log of CFU numbers on 1 g of grain

8

34.6

32.8 gh

123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123

A

7,5

fg

13.9

7 0.26

6,5

0.25

5,5

0.79 0.25

0.29

6

a

a

a

123 123 123 123 123 123 123 123 123 123 123 123 123 123 123

5 4,5 4 3,5

c

0.24 a

11.9

h 5.09 de 1.44 d

ef 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12

0.50 b

Inoc.

10ºC 15ºC 21ºC 28ºC

10ºC 15ºC 21ºC 28ºC

10ºC 15ºC 21ºC 28ºC

dose

10% 10% 10% 10%

20% 20% 20% 20%

30% 30% 30% 30%

8 log of fungal CFU numbers on 1 g of grain

4

Czaban J. et al.

gh

B

7,5

h

g

g Other fungi

7

f

P. verrucosum e

6,5

d

6

c

c

5,5

b

b

5 ?

4,5

?

?

?

4

a

3,5 Start

10ºC 15ºC 21ºC 28ºC

10ºC 15ºC 21ºC 28ºC

10ºC 15ºC 21ºC 28ºC

10% 10% 10% 10%

20% 20% 20% 20%

30% 30% 30% 30%

d

18

C

Fungal colony diameter in mm

16 14 12 10 8 6 4 2 0

ab

ab

b

b

123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123

ab

10°C 10ºC 15°C 15ºC 21°C 21ºC 28°C 28ºC 10% 10% 10% 10%

d

123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123 123

d

10°C 10ºC 15°C 15ºC 21°C 21ºC 28°C 28ºC 20% 20% 20% 20%

d

c b

12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12

a

10°C 10ºC 15°C 15ºC 21°C 21ºC 28°C 28ºC 30% 30% 30% 30%

4

325

Penicillium verrucosum growth and OTA production 100 90

D

Percentage of kernels

80 70 60 50 40 30 20 10 0 10ºC

15ºC 21ºC

28ºC

10ºC

15ºC

21ºC

28ºC

10ºC

15ºC 21ºC

28ºC

10%

10%

10%

20%

20%

20%

20%

30%

30%

30%

10%

e 4483

3,5

E

3 2,5 2 1,5

10.7

b

1 0,5

a

a

a

e

e

12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12

a

0

4393 1070 c

225

11.5

b

4372

275 c

15ºC 21ºC

28ºC

10ºC

15ºC

21ºC

28ºC

10ºC

15ºC 21ºC

28ºC

10%

10%

10%

20%

20%

20%

20%

30%

30%

30%

10%

50

50

45

F

10ºC 10°C

40

45

35

21ºC 21°C

30

28°C 28ºC

25 20 15

35 30 25 20 15

10

10

5

5

0

0 1

2

3

4

d

G

40

15ºC 15°C

0

d

12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12 12

10ºC

Colony diameter in mm

(A) CFU numbers of P. verrucosum on DYSG medium (the numbers in squares are the CFU × 106 per 1 g of grain); (B) CFU numbers of P. verrucosum and other fungi on Martin’s medium. (In experimental series with 20% grain water content the number of other fungi could not be determined, because it was masked by high number of P. verrucosum); (C) Diameters of P. verrucosum colonies growing around the wheat kernels placed on DYSG medium; (D) Percentage of wheat kernels placed on DYSG medium with colonies of fungi different from P. verrucosum growing by the kernels (The vertical lines represent 95% confidence intervals of the proportions); (E) OTA production by P. verrucosum (the numbers in squares are OTA concentration in ng per 1g of grain). The values of any columns with different letters are significantly different at P < 0.05 (Fig. 1A and 1B), P 0.05). Detection rate of CMV DNA was not significantly different between atherosclerotic and non-atherosclerotic vascular wall specimens (p > 0.05). Table II The presence of H. pylori and CMV DNA in the arteries of non- and atherosclerotic groups

Parameter

H. pylori (%) CMV (%)

Coronary artery Grade III Nonatheroscle- atherosclerotic group rotic group (n = 6) (n=24)

Carotid artery Grade III Nonatheroscle- atherosclerotic group rotic group (n = 12) (n = 18)

p

p

Abdominal aorta artery Grade III Nonatheroscle- atherosclep rotic group rotic group (n = 11) (n = 19)

4 (66.7)

3 (12.5)

0.016

2 (16.7)

3 (16.7)

1.000

8 (72.7)

6 (31.6)

0.029

3 (50)

4 (16.7)

0.120

5 (41.7)

7 (38.9)

1.000

3 (27.3)

9 (47.4)

0.442

Discussion It has been proposed that atherosclerosis is caused by inflammatory process and microorganisms could be the prime initiators of this process. Most authors investigating the endovascular presence of infectious agents used directional coronary atherectomy. Therefore, they were limited to small samples, possibly underestimating the true incidence of pathogens, so they could not identify co-infections within the coronary arteries. In terms of potential pathogenic effects, these agents may act involving both direct effects on arterial walls and indirect effects mediated via the circulation (Libby et al., 1997). Results obtained from animal studies (Fabricant et al., 1978), sero-epidemiologic observations (Patel et al., 1995; Rechciñski et al., 2005), pathologic-based investigations (Maass et al., 1998), and data from small-randomised clinical trials (Gupta et al., 1997) have provided evidence of direct pathogenic involvement of infectious agents in the process of atherogenesis. Several microorganisms including CMV and H. pylori have recently been related to the pathogenesis of atherosclerosis (Carlsson et al., 2000). The CMV, a herpesvirus restricted exclusively to the human host, may predispose to the development of atherosclerosis (Reinhardt et al., 2003; Horne et al., 2003). Moreover, it can be inferred from serological studies that there is an epidemiological link between CMV infection and atherosclerosis (Nieto et al., 1996; Reinhardt et al., 2002), but the data remain controversial (Siscovick et al., 2000). In atherosclerotic vascular walls, the presence of CMV DNA has been demonstrated by the detection of viral antigen and nucleic acids in the cultured smooth muscle cells from coronary

336

Kilic A. et al.

4

artery plaque material (Melnick et al., 1994). CMV nucleic acid was extracted from atherosclerotic femoral arteries and abdominal aortas in one study (Hendrix et al., 1991). It has been shown that 90% of advanced atherosclerotic lesions (Grade III) obtained at surgery were positive for CMV nucleic acids, whereas 50% of Grade I lesions obtained at autopsy from age- and sex-matched patients were positive (Hendrix et al., 1990). In our study, CMV DNA was found in both non-atherosclerotic and atherosclerotic artery tissue, but no significant association between the atherosclerotic groups and non-atherosclerotic groups were observed (P > 0.05). It can be inferred from this result that CMV could be in vessels, but it may not be a significant factor for atherosclerotic process. The Gram-negative curved bacillus, H. pylori, occurs endemically in worldwide, and it is estimated that up to 50% of the world’s population is infected (Sinha et al., 2002). H. pylori colonizes the gastrointestinal tract and is associated with chronic gastritis, peptic ulcer disease, and gastric cancer (Takashima et al., 2002). H. pylori has also been sero-epidemiologically linked to coronary heart disease and atherosclerosis by causing chronic infections (Heuschmann et al., 2001). H. pylori genome has been shown in diseased arterial segments (Danesh et al., 1999). However, two studies failed to demonstrate any evidence of H. pylori in the atherosclerotic plaques of abdominal aortic aneurysms (Blasi et al., 1996) and carotid arteries (Chiu et al., 1997) of patients being sero-positive for H. pylori. It has been shown that H. pylori has an important role in the pathogenesis of atherosclerosis, especially in Turkey, where infection is prevalent and conventional risk factors fail to explain the high prevalence of atherosclerotic vascular disease (Farsak et al., 2000). H. pylori DNA has also been found in 53% of carotid atherosclerotic plaques by demonstrating the microorganism in lesions by specific immunohistochemistry (Ameriso et al., 2001). In our study, the H. pylori DNA has been found in both non-atherosclerotic and atherosclerotic artery tissues. However, there was a significant association between H. pylori infection and atherosclerosis for coronary and abdominal aorta arteries (P < 0.05), whereas no significant association was found for carotid arteries (P > 0.05). H. pylori infection rates change markedly among distinct populations. Consistent with previous findings of high seroprevalences in less developed countries, Turkish people have been reported to be a high-risk population (Porsch-Ozcurumez et al., 2003). In highly risky countries like Turkey, H. pylori may play a role in the development of atherosclerosis. In conclusion, in this study, a relation between H. pylori and atherosclerosis but not CMV was shown. 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Polish Journal of Microbiology 2006, Vol. 55, No 4, 339– 343

Detection of Methicillin Resistance in Hospital Environmental Strains of Coagulase-negative Staphylococci TOMASZ NOWAK1, EWA BALCERCZAK2, MAREK MIROWSKI2 and ELIGIA M. SZEWCZYK 1* 1 Department 2 Molecular

of Pharmaceutical Microbiology, Medical University, £ódŸ, Poland Biology Laboratory, Department of Pharmaceutical Biochemistry, Medical University, £ódŸ, Poland

Received 3 July 2006, revised 24 September 2006, accepted 29 September 2006 Abstract The aim of this study was to evaluate methicillin resistance detection methods currently used when studying coagulasenegative staphylococci (CoNS). The resistance to oxacillin of 142 strains from seven species of CoNS isolated from the Intensive Care Unit environments was tested. The methods used were: disc diffusion test with cefoxitin (FOX 30) and oxacillin (OX1), oxacillin agar screen test with 6 mg/l of oxacillin (MHOXA), latex test for PBP2a (LA) and detection of mecA via PCR. One hundred and one isolates were methicillin-resistant in at least one of methods used, but only 74 were mecA-positive. Of the 68 mecA-negative strains: two were positive by OX1, the LA and MHOXA methods; three by the LA and MHOXA; and 22 only by OX1 test. Most of these strains were from the novobiocin-resistant CoNS group. The results obtained for all tested strains using FOX 30 showed complete concordance with the presence of the mecA gene. K e y w o r d s: cefoxitin disc test, coagulase-negative staphylococci, methicillin resistance

Introduction The number of nosocomial infections, particularly bloodstream infections caused by coagulase-negative staphylococci (CoNS) has increased in recent years (Marshall et al., 1998). The majority of clinical isolates are resistant to $-lactam antibiotics mainly thought to be due to methicillin (oxacillin) resistance mechanism (Pfaller et al., 1998). More than 70% of CoNS isolates worldwide are resistant to oxacillin (Diekema et al., 2001). Staphylococcus epidermidis is often isolated from clinical specimens but other species including novobiocin-resistant ones can also cause serious infection (Ishihara et al., 2001; Mastroianni et al., 1995; Okudera et al., 1991; Tselenis-Kotsowilis et al., 1982). It is vital that the appropriate antimicrobial therapy is applied for patients with these infections as inaccurate detection of oxacillin resistance can lead to important adverse clinical consequences. Furthermore, due to the emergence of vancomycin-resistant staphylococci, it is important for clinical laboratories to distinguish between oxacillin-susceptible (MSCNS) and oxacillinresistant coagulase-negative staphylococci (MRCNS) strains in order to limit unnecessary use of vancomycin. Thus, accurate and rapid detection of methicillin resistance in CoNS is essential for the success of this strategy. Laboratories use various tests to determine methicillin resistance, including: disc diffusion, oxacillin agar screening (MHOXA), broth microdilution, agar dilution and the latex agglutination (LA) test for PBP2a. Detection of mecA by PCR is still considered the “gold standard”, but this methodology is not feasible in every laboratory. Detection of the gene product – protein PBP2a as a marker for methicillin resistance is recommended as an alternative for mecA PCR by the National Committee for Clinical Laboratory Standards (NCCLS, 2002). In 2003, Ferreira et al. (2003) showed the accuracy of the MHOXA method with 4 mg/l of oxacillin and detection of PBP2a by the LA test for detection of oxacillin susceptibilities of CoNS strains, suggesting both methods as good alternatives for mecA PCR. Since 1999 use of the agar screen test has not been * Corresponding author: E.M. Szewczyk, Department of Pharmaceutical Microbiology, Medical University, Pomorska 137, 90-235 £ódŸ, Poland; phone: + 48 42 677 93 01; e-mail: [email protected]

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recommended by the NCCLS (1999), while Kohner et al. (1999) support the use of the MHOXA method for CoNS strains as well as for Staphylococcus aureus strains. In 2004, the NCCLS recommended a new disc diffusion test with 30 µg of cefoxitin for detection of oxacillin resistance as a further attempt to detect the correlation between oxacillin resistance detection and the presence of the mecA gene (NCCLS, 2004). Our study evaluated the oxacillin susceptibilities of 142 well-characterized strains of a wide group of CoNS novobiocin-susceptible and novobiocin-resistant species isolated from hospital environments. Four different phenotypic methods were compared with mecA gene assay to examine the resistance and the usefulness of these methods for the detection of methicillin resistance in CoNS with special attention to novobiocinresistant strains which haven’t previously been discussed as they are seldom isolated and studied. Experimental Materials and Methods Bacterial strains and identification. Coagulase-negative staphylococci: 142 well-characterized strains of seven species belonging to novobiocin-resistant group (Staphylococcus saprophyticus, Staphylococcus cohnii and Staphylococcus xylosus) and novobiocin-susceptible group (Staphylococcus epidermidis, Staphylococcus hominis, Staphylococcus haemolyticus, Staphylococcus warneri) were tested. Strains were collected in 1997 and in 2003 from the hospital environments of the Intensive Care Unit at the Teaching Paediatric Hospital in £ódŸ, Poland. Identification was performed using standard criteria with particular reference to Kloos and Bannerman (1999) and Freney et al. (1999). The API StaphSystem (bioMerieux, Marcy l’Etoile, France) was used as a parallel method of identification. Disc diffusion test. All strains were tested with 30 µg cefoxitin disc (FOX 30) and with 1 µg oxacillin (OX 1) disc on MuellerHinton Agar 2 (bioMerieux). Plates were incubated for 24 h in ambient air at 33 – 35°C (for cefoxitin, results may be reported after 18 h if resistant) according to the Clinical and Laboratory Standards Institute (CLSI, formerly NCCLS) (CLSI, 2005). Oxacillin agar screen test (MHOXA). Strains were plated on Mueller-Hinton agar (Difco Laboratories, Detroit, USA) supplemented with 4% NaCl containing 6 mg/l of oxacillin using a cotton swab dipped into a 0.5 McFarland standard suspension of each test strain according to the procedures outlined in the CLSI guidelines for S. aureus (CLSI, 2005). Oxacillin resistance was demonstrated by bacterial growth after 48 h of incubation at 35°C. Detection of PBP2a (DR900A – Oxoid, Basingstoke Hampshire, England). Strains were grown on Columbia agar with 5% sheep blood plates with a 1 µg/ml oxacillin disc placed on the main inoculum. After overnight incubation, cells that grew around the disc were used to perform the test. The test was carried out according to the manufacturer’s instructions using 1.5 × 109 CFU/ml inoculum of bacteria. Oxacillin-susceptible S. aureus ATCC 29213 and oxacillin-resistant S. aureus ATCC 43300 were used as control organisms. Detection of mecA gene. Staphylococcal DNA was extracted by lysostaphin and heating at 99°C as described by van Griethuysen et al. (1999). The mecA gene was detected by PCR according to the manufacturer’s instructions (DNA-GDANSK II, Gdansk, Poland). A 533-bp fragment of the mecA gene was amplified in a thermal controller (Tpersonal; Biometra, Göttingen, Germany) and then revealed by electrophoresis on 1.5% agarose gel at 120 V for 40 min.

Results One hundred and one (71%) of 142 tested strains belonging to seven species of CoNS were methicillinresistant according to at least one of methods used in this research (Table I), but only 68 presented the same results in all methods used in this research (Table II). Among the 142 CoNS strains 74 were mecA-positive and 68 were mecA-negative. Table I The number of strains suspected of showing methicillin resistance among tested species on the basis of at least one of methods used Species

Novobiocin-susceptible

Number of strains

Methicillin-susceptible

Methicillin-resistant

S. epidermidis

38

21

17

S. hominis

18

8

10

7

1

6

2

1

1

73

9

64

S. haemolyticus S. warneri S. cohnii

Novobiocin-resistant

S. xylosus

2

1

1

S. saprophyticus

2

0

2

142

41

101

TOTAL

4

341

Methicillin resistance in coagulase-negative staphylococci Table II Detection of methicillin resistance by the recommended methods compared to mecA PCR Number of strains

a b c d

Methicillin resistance demonstration PCR mecA

LA test PBP2aa

FOX30b

MHOXAc

OX1d

68

+

+

+

+

+

22

–

–

–

–

+

3

–

+

–

+

–

6

+

–

+

+

+

2

–

+

–

+

+

LA test PBP2a, the latex agglutination test for PBP2a FOX30, disc diffusion test with 30 µg of cefoxitin MHOXA, oxacillin agar screen test with 6 mg/l of oxacillin OX 1, disc diffusion test with 1 µg of oxacillin

Table III Credibility of currently used tests for determining the methicillin resistance in particular species of staphylococci Methicillin resistance demonstration Species S. epidermidis S. hominis

S. haemolyticus S. warneri

Number of strains

PCR mecA

LA test PBP2a

FOX30

MHOXA

OX1

16

+

+

+

+

+

1

–

+

–

+

+

7

+

+

+

+

+

2

–

+

–

+

–

1

–

+

–

+

+

6

+

+

+

+

+

1

+

+

+

+

+

38

+

+

+

+

+

19

–

–

–

–

+

6

+

–

+

+

+

1

–

+

–

+

–

S. xylosus

1

–

–

–

–

+

S. saprophyticus

2

–

–

–

–

+

S. cohnii

The results obtained for all tested strains using cefoxitin disc diffusion showed complete concordance with the presence of the mecA gene. Of the 68 mecA-negative strains: two were positive by OX1, the LA and MHOXA methods; three by the LA and MHOXA; and 22 only by OX 1 test (Table II). Our research did not show the correlation between methicillin resistance estimated by expression of protein PBP2a and presence of mecA gene in every strain tested (Table II and III). Almost 18% of strains presented methicillin resistance although mecA gene was not found. Such results occurred in the oxacillin disc test (19 S. cohnii strains, two S. saprophyticus and one S. xylosus), but also in MHOXA method and latex agglutination (two strains S. hominis and one strain S. cohnii). One S. epidermidis and one S.hominis strains were positive in both the oxacillin tests and in the LA test. Although mecA gene was present, the LA test revealed methicillin susceptibility in six strains of S. cohnii (Table III). Discussion Numerous studies have been conducted to determine the optimal methods for phenotypic detection of methicillin resistance in CoNS. The disc diffusion method currently recommended by the CLSI for all staphylococci is the 30 µg cefoxitin disc method (CLSI, 2005). In our study, all of the MRCNS detected as

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methicillin-resistant by this method possessed the mecA gene. Our results were in full concordance with data from Stierna-Johsen et al. (2005) who tested 110 CoNS isolates from monomicrobial bacteraemia. In our experiments many strains possessed the mecA gene and this was in agreement in all cases with cefoxitin resistance. The high percentage positive results, when using other phenotypic methicillin-resistance tests, suggest that resistance to methicillin may be connected with other mechanisms (genes) in these cases. Among the species most often isolated from clinical sources, the group of novobiocin-susceptible staphylococci, a discordance where results were positive and mecA was absent, applied to only 12% of tested strains. Among novobiocin-resistant staphylococci this discordance applied to more than 34% of strains. In the group of novobiocin-resistant strains S.cohnii dominated in hospital environments (Szewczyk et al., 2000). Six strains of these species in current study also did not demonstrate methicillin resistance in LA test although they were mecA and other phenotypic tests positive. Our results clearly show heterogeneity in the coagulase-negative staphylococci group. It seems however, that phenotypic testing with cefoxitin, which easily detects the methicillin resistance of all CoNS species, may be used to determine this feature. Nevertheless, negative opinions appearing in publications cannot be overlooked. Frigatto et al. (2005) observed discrepant results between oxacillin and cefoxitin when testing CoNS by disc diffusion and suggested that the detection of low-level methicillin resistance in CoNS by the cefoxitin disc could be problematic. Methicillin resistance in novobiocin-resistant staphylococci detected by phenotypic methods has already been mentioned in some articles concerning the optimisation of such methods for all CoNS (Ramotar et al., 2001; Tenover et al., 1999). However, these results usually came from a limited range of novobiocin-resistant strains and remained outside the main interest of the papers’ authors and were left unclear. Analysing studies of Suzuki et al. (1992), York et al. (1996) and Hussain et al. (2000) we noticed that such results have not lead to discussion regarding a few species of staphylococci, mainly: S. saprophyticus, S. cohnii, S. xylosus and S. lugdunensis. These data, however, confirmed our observations. Undertaking studies of CoNS strains specifically isolated from hospital environments allowed us to perform a unique review of a large novobiocin-resistant group relatively seldom isolated from clinical materials, a case in point being S. cohnii (Basaglia et al., 2003; Gauduchon et al., 2003). Kloos (1997) suggested that the ability of S. cohnii and S. hominis to share plasmids and antibiotic resistance genes may indicate that these species can act as a bridge for genetic exchange by horizontal gene transfer between novobiocin-susceptible Staphylococcus epidermidis group and novobiocin-resistant Staphylococcus saprophyticus group. This suggests a clear potential for interspecies spread of antibiotic resistance in CoNS and forces us to consider results relating to novobiocin-resistant species in the hospital. While the availability of PCR techniques is limited, the cefoxitin disk diffusion test is preferable to the other phenotypic methods, including LA test for detection of MRCNS and seems to be suitable for routine use. The MHOXA test with 6 mg/l of oxacillin, routinely used for S. aureus testing, gives more accurate results than the LA test which, in turn, is better than the oxacillin disc method in classifying mecA-negative CoNS as oxacillin susceptible. It may be possible that methicillin resistance of the six strains of S. cohnii, which were negative in LA test, was due to the presence of proteins other than PBP2a or different unknown mechanisms. The recent suggestion of Chandran and Rennie (2005) to discontinue routine antibiotic susceptibility testing (AST) of CoNS isolates from blood cultures because of common methicillin resistance, in light of the data put forward by us, could be seen as very controversial. Another conclusion drawn was that the CoNS group is not homogenous and the mechanisms of resistance to $-lactam antibiotics remains, to a large extent, unexplained. Therefore, we believe that the detection of susceptibility or resistance to methicillin even when using the cefoxitin disc diffusion test should not be considered the only possible option when leading to choices in antimicrobial therapy. Acknowledgments. This research was presented as a poster during the IX Symposium Advances in Medicine of Infection, December 2005, Warsaw, Poland. This study was supported by grant from the Medical University of £ódŸ No. 502-13-074.

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Y o k o t a. 1992. Survey of methicillin-resistant clinical strains of coagulase-negative staphylococci for mecA gene distribution. Antimicrob. Agents Chemother. 36: 429–434. S z e w c z y k E.M., A. P i o t r o w s k i and M. R ó ¿ a l s k a. 2000. Predominant staphylococci in the intensive care unit of a paediatric hospital. J. Hosp. Infect. 45: 145–154. T e n o v e r F.C., R.N. J o n e s, J.M. S w e n s o n, B. Z i m m e r, S. M c A l l i s t e r and J.H. J o r g e n s e n. 1999. Methods for improved detection of oxacillin resistance in coagulase-negative staphylococci: results of a multicenter study. J. Clin. Microbiol. 37: 4051–4058. T s e l e n i s - K o t s o w i l i s A.D., M.P. K o l i o m i c h a l i s and J.T. P a p a v a s s i l i o u. 1982. Acute pyelonephritis caused by Staphylococcus xylosus. J. Clin. Microbiol. 16: 593–594. v a n G r i e t h u y s e n A., M. P o u w, N. v a n L e e u w e n, M. H e c k, P. W i l l e m s e, A. B u i t i n g and J. K l u y t m a n s. 1999. Rapid slide latex agglutination test for detection of methicillin resistance in Staphylococcus aureus. J. Clin. Microbiol. 37: 2789–2792. Y o r k M.K., L. G i b b s, F. C h e h a b and G.F. B r o o k s. 1996. Comparison of PCR detection of mecA with standard susceptibility testing methods to determine methicillin resistance in coagulase-negative staphylococci. J. Clin. Microbiol. 34: 249–253.

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345 Polish Journal of Microbiology formerly Acta Microbiologica Polonica

2006, Vol. 55, No 1–4

CONTENTS Vol. 55 (1– 4), 2006 No 1 IN MEMORIAM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

5

ORIGINAL PAPERS

Tuberculosis bacilli still posing a threat. Polymorphism of genes regulating anti-mycobacterial properties of macrophages DRUSZCZYÑSKA M., STRAPAGIEL D., KWIATKOWSKA S., KOWALEWICZ-KULBAT M., RÓ¯ALSKA B., CHMIELA M., RUDNICKA W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

7

Mycobacterium bovis BCG mycobacteria – new application KOWALEWICZ-KULBAT M., PESTEL J., BIET F., LOCHT C., TONNEL A.B., DRUSZCZYÑSKA M., RUDNICKA W. . . .

Application of DNA markers to estimate genetic diversity of Mycobacterium tuberculosis strains KORZEKWA K., POLOK K., ZIELIÑSKI R. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

13

19

Potential role of LPS in the outcome of Helicobacter pylori related diseases GRÊBOWSKA A., RECHCIÑSKI T., B¥K- ROMANISZYN L., CZKWIANIANC E., MORAN A., DRUSZCZYÑSKA M., KOWALEWICZ-KULBAT M., OWCZAREK A., DZIUBA M., KRZEMIÑSKA-PAKU£A M., P£ANETA-MA£ECKA I., RUDNICKA W., CHMIELA M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Genotypes of Listeria monocytogenes strains isolated from 2000 to 2002 in Poland PACIOREK J., JACQUET C., SALCEDO C., DOUMITH M., VAZQUEZ J.A., MARTIN P. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Purification and characterization of cytolitic toxins produced by Aeromonas hydrophila and Aeromonas veronii biotype sobria strains KRZYMIÑSKA S., KAZNOWSKI A., SPYCHA£A H. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Effect of neuraminidase on adherence of Pseudomonas aeruginosa to human buccal epithelial cells. Inhibition of adhesion by monosaccharides WOLSKA K., ZABIELSKA K., JAKUBCZAK A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Interferon alpha in the establishment of latency by herpes simplex virus type I strain tr. WIECZOREK M., LITWIÑSKA B. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Nutrient modulated alkaline phosphatase and associated processes in diazotrophic cyanobacteria PANDEY M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

The isolation of microorganisms capable of phenol degradation PRZYBULEWSKA K., WIECZOREK A., NOWAK A., POCHRZ¥SZCZ M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Effectiveness of biodegradation of petroleum products by mixed bacterial populations in liquid medium at different pH values

25 31

37

43 49 53 63

BOSZCZYK-MALESZAK H., ZABOST A., WOLICKA D., KACIESZCZENKO J. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

69

INSTRUCTIONS TO AUTHORS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

75

No 2 IN MEMORIAM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

83

ORIGINAL PAPERS

Genetic characterisation of the cjaAB operon of Campylobacter coli WYSZYÑSKA A., PAW£OWSKI M., BUJNICKI J., PAWELEC D., van PUTTEN J.P.M., BRZUSZKIEWICZ E., JAGUSZTYN-KRYNICKA E.K. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

85

Natural mannose-binding lectin (MBL) down regulates phagocytosis of Helicobacter pylori STRAPAGIEL D., GRÊBOWSKA A., RÓ¯ALSKA B., B¥K-ROMANISZYN L., CZKWIANIANC E., P£ANETA-MA£ECKA I., RECHCIÑSKI T., RUDNICKA W., CHMIELA M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

95

Phenotypic and genotypic characteristic of Pseudomonas aeruginosa strains isolated from hospitals in the north-west region of Poland

CZEKAJ£O-KO£ODZIEJ U., GIEDRYS-KALEMBA S., MÊDRALA D. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103

346 Detection of enterotoxic Bacillus cereus producing hemolytic and non hemotylic enterotoxins by PCR test

O£TUSZAK-WALCZAK E., WALCZAK P., MODRAK R. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113

Optimization and purification of alkaline proteases produced by marine Bacillus sp. MIG newly isolated from Eastern Harbour of Alexandria

M.K. GOUDA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119

Novel yeast cell dehydrogenase activity assay in situ

BER£OWSKA J., KRÊGIEL D., KLIMEK L., ORZESZYNA B., AMBROZIAK W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127

In vitro activity of caspofungin against planktonic and sessile Candida sp. cells

SEREFKO A., CHUDZIK B., MALM A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133

Enhancement of oil degradation by co-culture of hydrocarbon degrading and biosurfactant producing bacteria

KUMAR M., LEON V., De SISTO MATERANO A., ILZINS O.A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139

Biotranformation of phosphogypsum on distillery decoctions (preliminary results)

WOLICKA D., KOWALSKI W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 147

Survival of Proteus mirabilis O3 (S1959), O9 and O18 strains in normal human serum (NHS) correlates with the diversity of their outer membrane proteins (OMPs)

FUTOMA-KO£OCH B., BUGLA-P£OSKOÑSKA G., DOROSZKIEWICZ W., KACA W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153

Lack of an association between Helicobacter infection and autoimmune hepatitis in children DZIER¯ANOWSKA-FANGRAT K., NILSSON I., WONIAK M., JӏWIAK P., RO¯YNEK., WOYNAROWSKI M., SOCHA J., LJUNGH A., WADSTROM T. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157 INSTRUCTIONS TO AUTHORS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161

No 3 IN MEMORIAM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 ORIGINAL PAPERS

Bactericidal activity of normal bovine serum (NBS) directed against some Enterobacteriaceae with sialic acid-containing lipopolysaccharides (LPS) as a component of cell wall

BUGLA-P£OSKOÑSKA G., DOROSZKIEWICZ W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169

Characterization of coagulase-negative staphylococci isolated from cases of ostitis and osteomyelitis

WILK I., EKIEL A., K£UCIÑSKI P., KRZYSZTOÑ-RUSSJAN J., MARTIROSIAN G. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175

Bioactive compounds from Streptomyces nasri and its mutants with special reference to proteopolysaccharides

GOHAR Y., BESHAY U., DABA A., HAFEZ E. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179

Hydrophobicity and biofilm formation of lipophilic skin corynebacteria

KWASZEWSKA A.K., BREWCZYÑSKA A., SZEWCZYK E.M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189

Iron supply of enterococci by 2-oxoacids and hydroxyacids

LISIECKI P., MIKUCKI J. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195

Growth, ferrous iron oxidation and ultrastructure of Acidithiobacillus ferrooxidans in the presence of dibutyl phthalate

MATLAKOWSKA R., SKUDLARSKA E., SK£ODOWSKA A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 203

Urea and ureolytic activity in lakes of different trophic status

SIUDA W., CHRÓST R.J. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211

Characterization of selected groups of microorganisms occurring in soil rhizosphere and phyllosphere of oats

REKOSZ-BURLAGA H., GARBOLIÑSKA M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 227

A simple, direct plating method, alternative to dilution plating, for estimation of the abundance of Penicillium verrucosum on incubated cereal grain

CZABAN J., WRÓBLEWSKA B. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237

L-forms of Staphylococcus epidermidis induced by penicillin

WRÓBLEWSKA J., JANICKA G., GOSPODAREK E., SZYMANKIEWICZ M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 243

INSTRUCTIONS TO AUTHORS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245

No 4 ORIGINAL PAPERS

Analysis of the filamentous bacteriophage genomes integrated into Neisseria gonorrhoeae FA1090 chromosome

PIEKAROWICZ A., MAJCHRZAK M., K£Y¯ A., ADAMCZYK-POP£AWSKA M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 251

Replication system of plasmid pMTH4 of Paracoccus methylutens DM12 contains an enhancer

SZYMANIK M., WELC-FALÊCIAK R., BARTOSIK D., W£ODARCZYK M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261

347 Acquisition of iron by enterococci; some properties and role of assimilating ferric iron reductases

LISIECKI P., MIKUCKI J. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271

Susceptibility of Listeria monocytogenes strains isolated from dairy products and frozen vegetables to antibiotics inhibiting murein synthesis and to disinfectants

POPOWSKA M., OLSZAK M., MARKIEWICZ M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279

Partial characterization and optimization of production of extracellular "-amylase from Bacillus subtilis isolated from culturable cow dung microflora

SWAIN M.R., KAR S., PADMAJA G., RAY R.C. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289

Production of tannase through submerged fermentation of tannin-containing plant extracts by Bacillus licheniformis KBR6

DAS MOHAPATRA P.K., MONDAL K.C., PATI B.R. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 297

In vitro activity of synthetic antimicrobial peptides against Candida KAMYSZ W., NADOLSKI P., KÊDZIA A., CIRIONI O., BARCHIESI F., GIACOMETTI A., SCALISE G., £UKASIAK J., OKRÓJ M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303

Effects of culture conditions on production of extracellular laccase by Rhizoctonia practicola

JANUSZ G., ROGALSKI J., BARWIÑSKA M., SZCZODRAK J. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 309

Growth of Penicillum verrucosum and production of ochratoxin A on nonsterilized wheat grain incubated at different temperatures and water content

CZABAN J., WRÓBLEWSKA B., STOCHMAL A., JANDA B. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321

Detection of cytomegalovirus and Helicobacter pylori DNA in arterial walls with grade III atherosclerosis by PCR

KILIC A., ONGURU O., TUGCU H., KILIC S., GUNEY C., BILGE Y. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333

Detection of methicillin resistance in hospital environmental strains of coagulase-negative staphylococci

NOWAK T., BALCERCZAK E., MIROWSKI M., SZEWCZYK E.M. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339

VOLUME 55 CONTENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . AUTHOR INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ACKNOWLEDGMENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . INSTRUCTIONS TO AUTHORS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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348 Polish Journal of Microbiology formerly Acta Microbiologica Polonica

2006, Vol. 55, No 1–4

Polish Journal of Microbiology 2006, Vol. 55, 1–4. Author Index Adamczyk-Pop³awska M. 251 Ambroziak W. 127 B¹k-Romaniszyn L. 25, 95 Balcerczak E. 329 Barchiesi F. 303 Bartosik D. 261 Barwiñska M. 309 Ber³owska J. 127 Beshay U. 179 Biet F. 13 Bilge Y. 333 Boszczyk-Maleszak H. 69 Brewczyñska A. 189 Brzuszkiewicz E. 85 Bugla-P³oskoñska G. 153, 169 Bujnicki J. 85 Chmiela M. 7, 25, 95 Chróst R.J. 211 Chudzik B. 133 Cirioni O. 303 Czaban J. 237, 321 Czekaj³o-Ko³odziej U. 103 Czkwianianc E. 25, 95 Daba A. 179 Das Mohapatra P.K. 297 de Sisto Materano A.139 Doroszkiewicz W. 153, 169 Doumith M. 31 Druszczyñska M. 7, 13, 25 Dzier¿anowska-Fangrat K. 157 Dziuba M. 25 Ekiel A. 175 Futoma-Ko³och B. 153 Garboliñska M. 227 Giacometti A. 303 Gierdys-Kalemba S. 103 Gohar Y. 179 Gospodarek E. 243 Gouda M.K. 119

Grêbowska A. 25, 95 Guney C. 333 Hafez E. 179 Ilzins O.A. 139 Jacquet C. 31 Jagusztyn-Krynicka E.K. 85 Jakubczak A. 43 Janda B. 321 Janicka G. 243 Janusz G. 309 JóŸwiak P. 157 Kaca W. 153 Kacieszczenko J. 69 Kamysz W. 303 Kar S. 289 Kaznowski A. 37 Kêdzia A. 303 Kilic A. 333 Kilic S. 333 Klimek L. 127 K³uciñski P. 175 K³y¿ A. 251 Korzekwa K. 19 Kowalewicz-Kulbat M. 7, 13, 25 Kowalski W. 147 Krêgiel D. 127 Krzemiñska-Paku³a M. 25 Krzymiñska S. 37 Krzysztoñ-Russjan J. 175 Kumar M. 139 Kwaszewska A.K. 189 Kwiatkowska S. 7 Leon V. 139 Lisiecki P. 195, 271 Litwiñska B. 49 Ljungh A. 157 Locht C. 13 £ukasiak J. 303

Majchrzak M. 251 Malm A. 133 Markiewicz Z. 279 Martin P. 31 Martirosian G. 175 Matlakowska R. 203 Mêdrala D. 103 Mikucki J. 195, 271 Mirowski M. 339 Modrak R. 113 Mondal K.C. 297 Moran A. 25 Nadolski P. 303 Nilsson I. 157 Nowak A. 63 Nowak T. 339 Okrój M. 303 Olszak M. 279 O³tuszak-Walczak E. 113 Onguru O. 333 Orzeszyna B. 127 Owczarek A. 25 Paciorek J. 31 Padmaja G. 289 Pandey M. 53 Pati B.R. 297 Pawelec D. 85 Paw³owski M. 85 Pestel J. 13 Piekarowicz A. 251 P³aneta-Ma³ecka I. 25, 95 Pochrz¹szcz M. 63 Polok K. 19 Popowska M. 279 Przybulewska K. 63 Ray R.C. 289 Rechciñski T. 25, 95 Rekosz-Burlaga H. 227 Rogalski J. 309

Ró¿alska B. 7, 95 Ro¿ynek E. 157 Rudnicka W. 7, 13, 25, 95 Salcedo C. 31 Scalise G. 303 Serefko A. 133 Siuda W. 211 Sk³odowska A. 203 Skudlarska E. 203 Socha J. 157 Spycha³a H. 37 Stochmal A. 321 Strapagiel D. 7, 95 Swain M.R. 289 Szczodrak J. 309 Szewczyk E.M. 189, 339 Szymanik M. 261 Szymankiewicz M. 243 Tonnel A.B. 13 Tugcu H. 333 van Putten J.P.M. 85 Vazquez J.A. 31 Wadstrom T. 157 Walczak P. 113 Welc-Falêciak R. 261 Wieczorek A. 63 Wieczorek M. 49 W³odarczyk M. 261 Wolicka D. 69, 147 Wolska K. 43 Woynarowski M. 157 WoŸniak M. 157 Wróblewska B. 237, 321 Wróblewska J. 243 Wyszyñska A. 85 Zabielska K. 43 Zabost A. 69 Zieliñski R. 19

349 Polish Journal of Microbiology formerly Acta Microbiologica Polonica

2006, Vol. 55, No 1–4

ACKNOWLEDGEMENTS The Editors of Polish Journal of Microbiology wish to express their gratitude to the following colleagues from the various fields of microbiology, who have reviewed the manuscripts submitted to our Journal in 2006. Tadeusz Antczak, Jadwiga Baj, Jacek Bardowski, Dariusz Bartosik, Jacek Bielecki, Hanna BoszczykMaleszak, Katarzyna Brzostek, Adam Choma, Aleksander Chmiel, Ryszard Chróst, Stanis³aw Ciesielski, Bronis³aw Cymborowski, Jerzy D³ugoñski, Maria Doligalska, W³odzimierz Doroszkiewicz, Nadzieja Drela, Adam Dubin, Jaros³aw Dziadek, Jan Fiedurek, Andrzej Gamian, Marek Gniadkowski, Renata Godlewska, Marcin Go³êbiewski, Stefania Giedrys-Kalemba, Anna Grabiñska-£oniewska, Krzysztof Grzywnowicz, Monika Hryniewicz, Waleria Hryniewicz, El¿bieta K. Jagusztyn-Krynicka, Wirginia Janiszowska, Halina Kalinowska, Henryk Ko³oczek, Gra¿yna Korczak-Kowalska, Teresa Korni³owicz-Kowalska, Jolanta Krzysztoñ-Russjan, Józef Kur, Jan Kwiatkowski, Felicja Meisel-Miko³ajczyk, Zdzis³aw Markiewicz, Gajane Martirosian, Jacek Miêdzobrodzki, Magdalena Miko³ajczyk-Chmiela, Gra¿yna M³ynarczyk, Krystyna Olañczuk-Neyman, Jaros³aw Paciorek, Andrzej Piekarowicz, Janusz Popowski, Anna Przondo-Mordarska, Gabrielle Reichmann, Barbara Ró¿alska, Ewa Sadowy, Edward Siñski, Aleksandra Sk³odowska, Tomasz S³omczyñski, Aleksander Œwi¹tecki, Jolanta Szych, Ewa Trafny, Stanis³awa Tylewska-Wierzbanowska, Grzegorz Wêgrzyn, Miros³awa W³odarczyk, Krystyna I. Wolska, Marta Wrzosek, Jan Zimowski, Zofia Zwolska

350

Polish Journal of Microbiology formerly Acta Microbiologica Polonica

2006, Vol. 55, No 4

Instructions to authors I. General information Polish Journal of Microbiology publishes descriptions of all aspects of basic and applied research that focuses on topics of basic research of practical value in microbiology. Topics that are considered include microbiology of a genetic and molecular nature, foods, agriculture, industry, biotechnology, microbial ecology, public health and basic biological properties of bacteria, viruses, and simple eukaryotic microorganisms. Submit manuscripts directly to the Editorial Office, Polish Journal of Microbiology. The manuscript should be accompanied by a covering letter stating the address, fax number, e-mail of the corresponding author and “running head” of the manuscript (no longer than 47 characters). Submit two complete copies of each manuscript, including figures and tables. The manuscript should be either the original typescript from jet or laser printer (not dot matrix). Accepted papers are copy-edited as word-processor files, so authors are asked to provide their paper in this form on a disk when they submit the revised version. The text should be edited in Word 7 or higher or ASCII. Submission of figures in TIF or CorelDraw Format is appreciated. All manuscripts are subjected to peer review by the editors, by members of the editorial board and by qualified outside reviewers. When a manuscript is returned to the authors for modification, it should be returned to the editor within 2 months; otherwise it may be considered withdrawn. 15 reprints are sent free to the first author. II. Preparation of the manuscript The original paper should be divided into the following sections written in sequence: Abstract, Introduction, Experimental: Materials and Methods and Results, Discussion, Acknowledgments, Literature. Type every portion of the manuscript double spaced with left hand margins, including figure legends, tables, table footnotes, and literature cited (type Literature sections on separate pages), and number all pages in sequence, including the abstract, tables and figure legends. The literature section must include all cited work. Arrange the citations in alphabetical order by first authors. Key words (no more than five) and the suggestion of runing head should be included. A paper in the form of a short communication must have an abstract of no more than 100 words. Do not use section headings in the body of the “Communication”; report introduction, methods, results, and discussion in a single section. The text should be concise, and the number of figures and tables should be kept to a minimum. Material and methods should be described in the text, not in figure legends or table footnotes. Present acknowledgments as in full-length papers. Minireviews are published in areas of particular interest and importance. They are usually invited, but authors wishing to submit a minireview should contact the scientific editor for further information. Before writing a manuscript authors are advised to consult a current issue of Polish Journal of Microbiology and carefully read the detailed “Instruction to authors” printed in number 1 of every volume in order to be familiar with the literature citations, preparation of figures and tables and the rules concerning chemical, biochemical, genetic etc. nomenclature recommended.

Polish Journal of Microbiology, formerly Acta Microbiologica Polonica, is a broad-based microbiological journal covering: General Microbiology, Bacterial Physiology and Genetics, Virology, Clinical Bacteriology, Environmental Microbiology and Biotechnology. Polish Journal of Microbiology publishes mainly original research articles including short communications but also minireviews, book reviews and press releases. All the papers should be prepared following “Instructions to Authors” which is always included in the first issue of every year. The papers appear usually within 3 months after final acceptance. Polish Journal of Microbiology is currently indexed by: BIOSIS/Biological Abstracts, EMBASE/Excerpta Medica, Index Medicus/MEDLINE, Cambridge Scientific Abstracts (series: Biological Sciences, Microbiology A, B, C), SCOPUS, Index Copernicus. Submission of manuscripts in various areas of Microbiology is very welcome.

Advertising offer The editorial office of the Polish Journal of Microbiology encourages our Readers to place advertisements in our quarterly. Polish Journal of Microbiology reaches scientists working in various research institutions and Universities and involved in different areas of microbiology, such as General Microbiology, Molecular Microbiology, Clinical Microbiology, Environmental Microbiology, Biotechnology. We ensure rapid appearance of the ad. Price of black and white ad on inside cover: half page = 150 PLN full page = 300 PLN Price of ad inside an issue half page = 75 PLN full page = 150 PLN Please send file with graphical layout to the editorial office of the Polish Journal of Microbiology, 02-096 Warsaw, Miecznikowa 1, phone 48 (22) 55 41 302

Do cz³onków Polskiego Towarzystwa Mikrobiologów Zarz¹d P.T.M. uprzejmie prosi wszystkich cz³onków o wp³atê zaleg³ych i bie¿¹cych sk³adek cz³onkowskich. Zachêcamy tak¿e do zaprenumerowania “Polish Journal of Microbiology”.