Polish Journal of Microbiology

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POLSKIE TOWARZYSTWO MIKROBIOLOGÓW POLISH SOCIETY OF MICROBIOLOGISTS

Polish Journal of Microbiology

I am pleased to inform you that Polish Journal of Microbiology has been selected for coverage in Thomson Scientific products and customers information services. Beginning with No 1, Vol. 57, 2008 information on the contents of the PJM is included in: Science Citation Index Expanded (ISI) and Journal Citation Reports (JCR)/Science Edition. Stanis³awa Tylewska-Wierzbanowska Editor in Chief

2009

Polish Journal of Microbiology formerly Acta Microbiologica Polonica

2009, Vol. 58, No 3

CONTENTS MINIREVIEW Communication between microorganisms as a basis for production of virulence factors

GOSPODAREK E., BOGIEL T., ZALAS-WIÊCEK P. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

ORIGINAL PAPERS Identification and molecular modeling of a novel lipase from an Antarctic soil metagenomic library

CIEŒLIÑSKI H., BIA£KOWSKA A., TKACZUK K., D£UGO£ÊCKA A., KUR J., TURKIEWICZ M. . . . . . . . . . . . . . . . . . . . . 199

Use of zwitterionic type of detergent in isolation of Escherichia coli O56 outer membrane proteins improves their two-dimensional electrophoresis (2-DE)

BUGLA-P£OSKOÑSKA G., FUTOMA-KO£OCH B., SKWARA A., DOROSZKIEWICZ W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205

Purification and biochemical characteristic of a cold-active recombinant esterase from Pseudoalteromonas sp. 643A under denaturing conditions

D£UGO£ÊCKA A., CIEŒLIÑSKI H., BRUDZIAK P., GOTTFRIED K., TURKIEWICZ M., KUR J. . . . . . . . . . . . . . . . . . . . . . 211

Usefulness of PCR method for detection of Leishmania in Poland MYJAK P., SZULTA J., DE ALMEIDA M.E., DA SILVA A.J., STEURER F., LASS A., PIETKIEWICZ H., NAHORSKI W.L., GOLJAN J., KNAP J., SZOSTAKOWSKA B. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219

Biofilm formation as a virulence determinant of uropathogenic Escherichia coli Dr+ strains

ZALEWSKA-PI¥TEK B.M., WILKANOWICZ S.I., PI¥TEK R.J., KUR J.W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 223

The first detection of Babesia canis canis in Ixodes ricinus ticks (Acari, Ixodidae) collected in urban and rural areas in northern Poland

CIENIUCH S., STAÑCZAK J., RUCZAJ A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231

Genetic variability of Czech and German RHD virus strains

HUKOWSKA-SZEMATOWICZ B., PAWLIKOWSKA M., DEPTU£A W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237

Usefulness of PCR melting profile method for genotyping analysis of Klebsiella oxytoca isolates from patients of a single hospital unit

STOJOWSKA K., KA£U¯EWSKI S., KRAWCZYK B. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 247

Genetic features of clinical Pseudomonas aeruginosa strains

WOLSKA K., SZWEDA P. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255

Effect of ciprofloxacin and N-acetylcysteine on bacterial adherence and biofilm formation on ureteral stent surfaces

EL-FEKY M.A., EL-REHEWY M.S., HASSAN M.A., ABOLELLA H.A., ABD EL-BAKY R.M., GAD G.F. . . . . . . . . . . . . . . . 261

Isolation and identification of a new fungal strain for amylase biosynthesis

BAKRI Y., MAGALI M., THONART P. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269

SHORT COMMUNICATION The cross-reactivity of Shewanella fidelis lipopolysaccharide with anti-Proteus antibodies

KWINKOWSKI M., GRABOWSKI S., KONIECZNA I., NAZARENKO E.L., KACA W. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275

INSTRUCTION TO AUTHORS AVAILABLE AT www.microbiology.pl/pjm

Polish Journal of Microbiology 2009, Vol. 58, No 3, 191–198 MINIREVIEW

Communication Between Microorganisms as a Basis for Production of Virulence Factors EUGENIA GOSPODAREK*, TOMASZ BOGIEL and PATRYCJA ZALAS-WIÊCEK

Department of Microbiology, Nicolaus Copernicus University in Toruñ, Collegium Medicum in Bydgoszcz, Poland Received 25 June 2009, revised 24 July 2009, accepted 30 July 2009 Abstract Quorum sensing (QS), or cell-to-cell communication in bacteria, is achieved through the production and subsequent response to the accumulation of extracellular signal molecules called autoinductors. The main role of QS is regulation of production of virulence factors in bacteria. Bacterial pathogenicity is often manifested by the expression of various cell-associated and secreted virulence factors, such as exoenzymes, toxins and biofilm. In bacteria, the expression of virulence factors is controlled coordinately by the global regulatory QS systems, which includes the AI-1/LuxIR-, AI-2/LuxS-, AI-3/QsC-, AIP/Agr-based systems. The regulation of production of virulence factors is extremely complex and many components influence it. K e y w o r d s: autoinductor, pathogenesis, quorum sensing, virulence factors

Introduction One of the fundamental adaptation abilities of microorganisms is sensitivity to changes in the environment and to their behaviour by managing for survival purposes. While the environment of microorganisms changes, the reaction is often programmed by changes in the expression of genes, resulting from the improvement of survival. Pathogenesis may be considered as bacterial adaptation towards survival and development in a new niche in a host organism. During pathogenesis, numerous virulence factors are expressed and used in different ways to support the survival of bacteria in the host organism. Many different factors may influence the evolution and course of pathogenesis, that is temperature, pH, chemical compounds from the host. Moreover, numerous natural microflora should be added to host factors, which may also detect signals deriving from other bacterial species. During this process, known as quorum sensing (QS), bacteria produce signal called autoinductor (AI), which is secreted to environment. Every AI, reaching critical concentration causes growth of the bacterial population and induces changes in the expression of genes resulting from switching life cycle and bacterial metabolism. Some bacteria use

QS to detect one another when the community they belong to reaches population numerous enough to activate a “profitable” virulent way of life, i.e. biofilm formation. AI-based systems, which do not react strictly with QS model, are also used by bacteria, i.e. pathogenic bacteria utilize signal molecules to detect the presence of a host’s microflora, indicating this way that the pathogen is localized in the specific niche as colonization. In these cases the detected signal is not transferred by bacteria – “invader”, but it rather informs of the high population density. Hence, bacteria that colonize a host do not have to produce or secrete a signal, only to detect it. There are known four common systems of bacterial cell-to-cell communication. Two of them, where autoinductor-1 (AI-1) and autoinductor-3 (AI-3) are used, exist in Gram-negative bacteria while Gram-positive have their own system based on an autoinducing polypeptide (AIP). The fourth system, which utilizes autoinductor-2 (AI-2), is present in both Gram-positive and Gram-negative bacteria and it is a very common signaling system. All the systems begin producing and secreting the autoinductor to the environment by either pathogen or natural microflora. Detection of these chemically different autoinductors and changes in genes’ expression is specific to all of the systems.

* Corresponding author: E. Gospodarek, Department of Microbiology, Nicolaus Copernicus University in Toruñ, Collegium Medicum in Bydgoszcz, 9 M. Sklodowska-Curie St, 85-094 Bydgoszcz, Poland; phone: (+48) 52 5854480; e-mail: [email protected]

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QS signal molecules, although largely considered as effectors of QS-dependent gene expression, are also emerging as multifunctional molecules that influence life, development and death in single and mixed microbial populations and impact significantly the outcome of host-pathogen interactions (Williams and Cámara, 2009). QS is a key regulatory system, which itself is affected by many other regulators (Girand and Bloemberg, 2008). AI-1/LuxIR-based system The first finding of this system is associated with explanation of the generation of luminescence by Vibrio fischeri. That is why LuxI/LuxR-based system is still a model QS-system, on which the others base. It includes LuxI, which produces N-acyl homoserine lactone (AHL) called AI-1 and LuxR, transcription factor responsible for gene expression control in the presence of AI. LuxI and its homologues produce AI by fatty acid chain transfer from acylated acyl carrier protein (ACP) to S-adenosylonationine (SAM), releasing AHL and methylthioadenosine (Schaefer et al., 1996). AHL produced by LuxI homologues contain different fatty acids remains, what favours recognition of specific ACP by synthases and results from species- and genusspecific signal. After LuxI synthesis, AI-1 diffuses through cell barriers and is released to environment. That is why, every single bacterium simultaneously produces AI-1 in growing population, and its concentration in the environment grows along with population growth. At high cell density, AI-1 concentration is high enough to diffuse inside the cell, where it bonds LuxR. After AI-1 binding, LuxR activates transcription of luxCDABEGH-operon by binding Lux-boxes localized in promoter (Devine et al., 1989). The product of this operon, luciferase, catalyzes chemical reaction, which gives luminescence. Presence of LuxI/LuxR-system homologues has been proved in many Gram-negative bacteria, able to produce specific AHL. Among some bacterial species like Pseudomonas aeruginosa, Serratia marcescens, opportunistic pathogens, mechanisms of communication control pathways responsible for expression of different virulence factors. Both species, able to form biofilm on biomaterials, cause chronic infections in hospitalized patients. The nature of biofilms makes these infections more resistant to antibiotic treatment than planktonic bacteria cells. P. aeruginosa rods contain two AHL-based QS systems: LuxI/LuxR for biofilm formation control and production of extracellular enzymes, as well as transcription by different QS system, RhII/RhIR enriching by additional control

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level through AHL-signaling system (De Kievit and Iglewski, 2000). Both systems play a role in virulence of lasI and rhII strains, as mutants with one or two mutations. These strains colonize lungs of infant mouse more weakly than wild type (Pearson et al., 2000). LasI synthesize oxo-C12-homoserine lactone (HSL), which activates LasR, while RhII produces C4-HSL, activating RhIR (Schuster and Greenberg, 2006). Ueda and Wood (2009) discern a mechanism by which QS controls biofilm formation by screening 5850 transposon mutants P. aeruginosa PA14 for altered biofilm formation. This screen identified the PA3885 mutant, which had 147-fold more biofilm than the wild-type strain. Loss of PA3885 decreased swimming, abolished swarming, and increased attachment, although this did not affect production of rhamnolipids. The PA3885 mutant also had a wrinkly colony phenotype, formed pronounced pellicles, had substantially more aggregation, and had 28-fold more exopolysaccharide production. Expression of PA3885 in trans reduced biofilm formation and abolished aggregation. Whole transcriptome analysis showed that loss of PA3885 activated expression of the pel locus, an operon that encodes for the synthesis of extracellular matrix polysaccharide. Genetic screening identified that loss of PelABDEG and PA1120 protein (which contains a GGDEF-motif) suppressed the phenotypes of the PA3885 mutant, suggesting that the function of the PA3885 protein is to regulate 3.5-cyclic diguanylic acid (c-di-GMP) concentrations as a phosphatase since c-di-GMP enhances biofilm formation by activating PelD, and c-di-GMP inhibits swarming. Loss of PA3885 protein increased cellular c-di-GMP concentrations; hence, PA3885 protein is a negative regulator of c-di-GMP production. Purified PA3885 protein has phosphatase activity against phosphotyrosine peptides and is translocated to the periplasm. Las-mediated QS positively regulates expression of the PA3885 gene. The PA3885 protein responds to AHL signals and likely dephosphorylates PA1120, which leads to reduced c-di-GMP production. This inhibits matrix exopolysaccharide formation, which leads to reduced biofilm formation (Ueda and Wood, 2009). P. aeruginosa forms a biofilm in lung tissue in cystis fibrosis (CF) patients and oxo-C12-HSL particles can be detected in sputum. In vitro studies indicate that oxo-C12-HSL is conducive to IL-8 production by bronchial epithelial cells, through induction of the transcription factors NF-kB and AP-2. In vivo studies using the adult mouse acute lung refection model indicate that P. aeruginosa ) lasI/rhlI mutant, defective for the production of oxo-C12-HSL and C4-HSL, is attenuated for disease in mice. Oxo-C12-HSL, but not C4-HSL, induces a potent inflammatory response in vivo. Expression analysis indicated increased expression of the chemokines MIP-2 (macrophage inflamma-

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tory protein), MCP-1 (monocyte chemotactic protein), MIP1-beta, IP-10, CTCA 3) T cell activation gene 3, IL-1 alpha and IL-6 (Pacheco and Sperandio, 2009). Activated T cells exposed to oxo-C12-HSL showed an increased production of IFN comparable to that induced by the control IL-12, but not IL-4, suggestion that oxo-C12-HSL induces a Th1 inflammatory response. These data suggest that oxo-C12-HSL stimulates inflammation in most tissues. Oxo-C12-HSL induces apoptosis in bone marrow derived macrophages and neutrophiles (Tateda et al., 2003). Although there is an increasing evidence that oxo-C12-HSL manipulates host cell signaling, the mechanism of this regulation remains unknown. Culture positivity for lasR mutant P. aeruginosa may serve as marker of early CF adaptive change of prognostic significance. Furthermore, as LasR inactivation alters susceptibility to antibiotics, infection with las mutant P. aeruginosa may impact response to therapy (Hoffman et al., 2009). In P. aeruginosa, the expression of virulence genes is coordinately controlled by the global regulatory QS systems, which includes the las and rhl systems, as well as the Pseudomonas quinolone signal (PQS) system (Liang et al., 2008). Phenazine compounds are among the virulence factors under the control of both the rhl and PQS systems. Gene PA0964, designated pmpR (pqsR-mediated PQS regulator), has been identified as novel regulator of the PQS system. It belongs to a large group of widespread conserved hypothetical proteins with unknown function, the YebC protein family (Pfam family DUF28). It negatively regulates the QS response regulator pgsR of the PQS system by binding at its promoter region. Alongside phzA1 expression and phenazine and pyocyanin production, a set of virulence factors genes controlled by both rhl and the PQS were shown to modulated by PmpR. Swarming motility and biofilm formation are also significantly affected. The results added another layer of regulation in the rather complex QS systems in P. aeruginosa and demonstrated a clear functional clue for the YebC family proteins. Nakamura et al. (2008) investigated extracellular lipopolysaccharide (LPS) and DNA in the supernatants of culture solutions from PAO1, the wild-type P. aeruginosa, and those of QS mutants. As compared to that of las QS mutants, the amount of LPS and DNA released was significantly higher in PAO1 and in las QS mutants complemented with N-(3-oxododecanoyl) homoserine lactone. QS is among the regulators involved in the release of extracellular LPS and DNA. S. marcescens also has a homologous system LuxI/ LuxR. This system in the presence of AHL, produced by Serratia spp. turns on the control of many pathways connected with S. marcescens strains pathogenesis, in comparison with wild type strain. Synthase SmaI

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AHL-deficient strain proves decrease in swarming motion, weaker adhesion (probably due to biofilm formation defect) and lower production level of extracellular casein, chitinase and hemolysin (Coulthurst et al., 2006). These defects are repaired by AHL addition, what indicates that these properties are caused directly or indirectly by cell-to-cell communication. Studies suggest that QS systems are used to control the density of own population, but it does not always happen. Escherichia coli and Salmonella enterica serovar Typhimurium do not have LuxI-equivalents, hence they do not produce AI-1, but encode LuxR-homologues, called SdiA. When overproduced, they exhibit negative interaction on genes that influence cell adhesion of enterohemorrhagic E. coli strains (EHEC) (Kanamaru el al., 2000). Positive interaction is observed in case of different genes localized on S. Typhimurium virulence plasmid, including rck gene, producing protein involved in host immune response evasion (Ahmer et al., 1998). Although exact role of SdiA in pathogenesis is unclear, protein allows EHEC and S. Typhimurium strains to change genes’ expression in response to the presence of AI-1, produced by other bacteria (Michael et al., 2001). It may have a particular significance for these pathogens, because of colonization gastrointestinal tract, where AI-1 may be present as a product of host’s natural microflora. AI-2/LuxS-based system Not only AI-1-based system is available for cellto-cell communication. Numerous Gram-negative and Gram-positive bacteria species have a system that detects extracellular signal known as autoinductor-2 (AI-2). This signal differs from the above described AHL-system and is synthesized by SAM metabolism. LuxS changes ribose – homocysteine into homocysteine and 4.5-dihydroxy-2.3-pentanedione (DPD) component, which in the presence of water, forms different furanone-derivatives. AI-2, like AI-1, is released by bacteria and accumulated in the cells’ environment. Two mechanisms of AI-2 detection, both different from recognition of AI-1, have been described. First of them, described in V. harveyi, is specific to borate diester forming AI-2. This mechanism detects AI-2 in periplasm by binding signal molecules to proteins binding specific autoinductor, LuxP. AI-2/LuxP-complex reacts subsequently with LuxQ-kinase sensor, initiating phosphate-transfer cascade by deactivation of negative response of LuxO regulator in luciferase production and luminescence. Among pathogenic V. cholerae strains LuxO inhibits the production of transcription regulator HapR at low cell density, which further inhibits expression of some virulence factors (Zhu et al., 2002). These

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bacteria have reversed “intuition” in virulence factors expression at low population density. To identify AIregulated target genes in V. cholerae El Tor, strain responsible for the current cholera pandemic, luciferase expression was assayed in an AI– strain carrying a random lux transcriptional reporter library in the presence and absence of exogenously added AI. Twenty three genes were identified and shown to require the QS transcription factor, HapR, for their regulation. Several of the QS-dependent target genes, annotated as encoding hypothetical proteins, in fact encode HD-GYP proteins, phosphodiesterases that degrade the intracellular second messenger cyclic dimeric GMP (c-di-GMP), which is important for controlling biofilm formation. Indeed, overexpression of a representative QS-activated HD-GYP protein in V. cholerae El Tor reduced the intracellular concentration of c-di-GMP, which in turn decreased exopolysaccharide production and biofilm formation. The V. cholerae classical biotype which caused previous cholera pandemics and is HapR–, controls c-di-GMP levels and biofilm formation by the VieA signaling pathway. The VieA pathway is dispensable for biofilm formation in V. cholerae El Tor but that restoring HapR in V. cholerae classical biotype reestablishes QS-dependent repression of exopolysaccharides production (Hammer and Bassler, 2009). AI-2 is used by E. coli and S. Typhimurium rods in different ways. These bacteria import AI-2 by the mechanism similar to capturing ribose by rbs-system. Distinctive LuxP/LuxQ-system, Lsr system regulated by LuxS, induces cellular response by transport of the AI-2 to cell cytoplasm. This process starts with recognition of the signal by periplasmic protein, LsrB, which binds to R-THMF form of AI-2. One bond of the transporter, Lsr ABC, contains LsrA and LsrC and allows AI-2 insertion to the cell, where it is phosphorylated by LsrK. Phosphorylated AI-2 form reacts with transcription repressor LsrR what extinguishes operon Isr (Taga et al., 2001). Although not demonstrated directly, this mechanism or a similar one, possibly results from decrease of the regulation of additional operons in the presence of phosphorylated form of AI-2. LuxS/AI-2 systems have been detected among numbers of Gram-positive and Gram-negative bacteria species, what indicates that AI-2 system allows common communication among species (Xavier and Bassler, 2003). Considering a complicated nature of most natural bacteria communities, for example, in the mammalian gastrointestinal tract, this signal may be used for regulation of genes required to survive in the presence of other bacteria. Pathogenic bacteria may use the same signals for activating mammalian cells, which let them gain an advantage in the same environment. The role of AI-2 in pathogenesis of bacteria other than Vibrio spp. is unclear. Studies using luxS-mutants have demonstrated

the growth of the expression level of pili and type III secretion mechanism in EHEC strains only when autoinductor is present (Sperandio et al., 1999, 2001). It has been noticed that direct influence of the phosphorylated AI-2 or bonded AI-2 on these genes never affects the response of the regulator. Many regulatory systems control virulence-associated traits in Staphylococcus epidermidis. LuxS mutant S. epidermidis shows increased biofilm formation in vitro and enhanced virulence in rat model of biofilm-associated infection. On the contrary, inactivation of luxS in various S. aureus strains has been reported not to affect virulence-associated traits. Externally added AI-2 almost completely restored gene expression patterns of the wild-type strain in the luxS mutant strain S. epidermidis. S. epidermidis regulates virulence-associated factors in addition to metabolism in an AI-2-dependent manner. There was dramatic AI-2-dependent alternation of pro-inflammatory phenol-soluble modulin (PSM) expression (Li et al., 2008). PSMs have been recently recognized as key proinflammatory and immune evasion factors in S. epidermidis and S. aureus. AI-3/QsC-system AI-3, different from AI-2-system, was first described as a component of used growth medium, which activates expression of genes responsible for EHEC adhesion to eukaryotic cells, the result of which is rearrangement of actin proteins (Sperandio et al., 2003). Structure and synthesis of that signal is unclear. Like AI-1, it represents family of particles. Results indicate that LuxS influences some pathways, because AI-3 production is less efficient in luxS mutants. Other studies suggest that lack of AI-3 production in the same mutants is a result of change in a cell metabolism. When oxaloacetate was used instead of SAM as methionine precursor and L-aspartate was added to growth medium to reduce requirements of oxaloacetate, AI-3 production was reversed, but no result on AI-2 production was observed (Walters et al., 2006). These studies also indicate that numerous commensal bacteria, i.e. nonpathogenic E. coli and Enterobacter cloacae and pathogenic species like Shigella spp., Salmonella spp. and Klebsiella spp. produce AI-3. It suggests that AI-3 may represent other interspecies signal, which has not been detected in Grampositive bacteria. The role of AI-3 in commensal communities is still vague. AI-3 detection is possible using 2-complements system composed of QseC-kinase sensor and QseBregulator response. In the presence of periplasmic AI-3, QseC is autophosphorylated first and transfers this phosphate on QseB afterwards, which activates

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genes responsible for pili biosynthesis and motility by decrease in regulation of genes regulator responsible for host’s cilium flhDC (Clarke et al., 2006). AI-3 presence is also connected with the process of adhesion and effacement (attaching and effacing, AE) of lesion by EHEC – phenomenon connected with increased regulation of 5 different loci of enterocyte effacement (LEE) operons localized on the EHEC chromosome (Sperandio et al., 2003). Cascade responsible for these genes’ regulation is still unclear, probably requires QseA, LysR – family regulator that influences cell-to-cell communication and directly increases LEE genes regulation (Sperandio et al., 2002). On the basis of these observations, it is suggested that intestinal pathogens may utilize AI-3 produced by host’s microflora and pili overexpression as well as motility regulation is necessary to penetrate intestinal mucous membrane rich in epithelial cells. Proteins encoded by LEE genes are crucial to contact with epithelial cells and make pathogens able to adhere and finally colonize eukaryotic cells. QseBC cascade is also responsible for adrenaline and noradrenaline signals, existing in gastrointestinal tract (Clarke et al., 2006). It suggests that QseC is responsible simultaneously for bacteria and host signals. Adrenergic receptors on eukaryotic cells may react to AI-3 in like they do to adrenaline and noradrenaline. Intestinal bacteria may have an advantage in this kind of communication for intestinal epithelial cells preparation for colonization. AIP/Agr-system This system of cell-to-cell communication occurs in Gram-positive bacteria. It is based on the prototype Agr system, first described in S. aureus strains. Gram-positive bacterial systems utilize polypeptide signals. They are bi-functional: they influence an organism which produces them as autoinductors and other organisms as inhibitors. That signal, called AIP, is encoded by agrD genes. After translation, propeptide AgrD is directed to membrane by N-terminal sequence of the signal. Near membrane, AgrB, membrane-related endopeptidase cuts C-terminal of the propeptide. N-terminal part of the propeptide, including signal sequence, is excluded by peptidase SpsB signal. Eventually, C-terminal part of the transformed polypeptide is covalently bonded with centrally placed cysteine to a ring form thiolactone with free N-end. Both structures are required for appropriate function of AIP. After reaching environment, AIP is recognized by AgrC-signal receptor. This protein contains transmembrane rest ended with N-sequence responsible for recognizing specific AIP and histidine kinase N-rest, which in the presence of specific AIP, phos-

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phorylates the response of AgrA regulator. Phosphorylated AgrA activates transcription of selected genes by direct bond of repeated regions localized in promoter. Uniqueness of AIP/Agr system is the fact that AIP produced by one Staphylococcus spp. strain may interfere into Agr system of another one. This double role as activator and inhibitor is connected with interaction between AIP and AgrC. Cyclic structure of AIP is required for interactions with AgrC, but the N-terminal part is responsible for AgrC activation. Removal of this ending (tarl) results from a universal inhibitor, which bonds AgrC, but is not capable to activate Agr system (Lyon et al., 2000). Agr system is connected with pathogenesis of numerous Gram-positive bacteria. Agr system is basic QS element among staphylococci. During the switch from late logarithmic to stationary phase, decrease in the level of expression of a few surface membrane proteins and increase of numerous virulence factors are observed. About 150 genes are under its control. It is found in about 14 species/subspecies of staphylococci. Its role, as a QS mediator, depends on the type of infection, time and environment which host provides. Agr expression is dynamic. Agr is exposed only in dynamically growing cell populations. Its activity is observed in two phases: phase I – first three hours after infection, important for abscess growth, phase II – 48-72 hours afterwards. Agr inactivation is connected with increase in resistance to antibiotics and small colony variants (SCV). SCV growth in the presence of AIP of isogenic strain restores or partially restores Agr activity. S. aureus strains, deprived of Agr system, have a decreased ability to cause osteomyelitis on both mice and rabbit models (Blevins et al., 2003; Gillaspy et al., 1995). AIP genes regulation by AgrA results from production and secretion of numerous S. aureus toxins, for instance, hemolysin alpha-, beta- and delta-, serine protease and toxic shock syndrome toxin (TSST-1). The S. aureus genome encodes a number of transcriptional factors, the Sar (staphylococcal accessory regulator) family of proteins, including SarA, SarR, SarS, SarT, SarV, and Rot. SarA is transcriptional activator for the Agr system, as well as a transcriptional regulator that activates or represses a number of staphylococcal genes. SarA, for example, is a repressor of spa (staphylococcal protein A) and an upregulator for the fibronectin-binding protein A. SarA is also required for biofilm formation. SarR binds to the sarA promoter region to downregulate transcription from its P1promoter and thus reduces SarA protein expression. SarS is a positive regulator of spa transcription. SarT has been shown to positively regulate sarS transcription and negatively regulate expression of hla (which encodes alpha-hemolysin) and sarU. SarU is proposed to be a positive regulator of agr expression.

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SarV is thought to be an important regulator in the autolytic pathway of S. aureus. SarX is a negative regulator of Agr. AgrA has been shown to be an activator of microcapsule synthesis, nuclease expression, and norA transcription but represses the expression of alpha-toxin, coagulase, protease, protein A, and certain genes involved in autolysis (Hsieh et al., 2008). Hemolysin secretion and eukaryotic cells damage by protease facilitate staphylococci to adhere, while TSST-1 is a superantigen that bonds nonspecific receptors of the T-cells and eventually yields strong immune response caused by systemic cytokine production. Furthermore, high toxins regulation and high AIP concentration are responsible for decreased level of proteins exposed to the cell surface, like fibronectin binding proteins and protein A, two proteins connected with staphylococci adhesion. That drop of cell adhesion ability may impact negatively on the architecture of populations related to biofilm. Bacteria cells are released from their populations and as planktonic populations migrate to other structures of the host body. If they find an available place, they adhere again and multiply, causing secondary infections. QS regulation in S. epidermidis has two major characteristics that reflect the adaptation to stationary growth phase: 1) up-regulation of virulence, resistance, stress and other factors that are needed for survival under suboptimal conditions and 2) down-regulation processes, such as translation and cell division that are typical of exponentially growing cells (Yao et al., 2006). Biofilm formation is a key virulence determinant during chronic staphylococcal infections, particularly in infection by S. epidermidis. The low activity of agr is a characteristic feature of the main part of S. epidermidis biofilms, whereas exposed regions in a biofilm express agr at a higher level. Agr is differentially expressed during the course of infection. Enterococcus faecalis is another Gram-positive bacterium that uses the AIP signal. These streptococci utilize a 2-component system corresponding to the Agr system in Staphylococcus spp. When AIP is detected, the cells produce and secrete two extracellular proteases: gelatinase (GeIE) and serine protease (SprE) (Qin et al., 2000). The role of these two proteins in pathogenesis is still unclear, but gelatinase facilitates cells migration into the host organism (Zeng et al., 2005). E. faecalis translocation through epithelial cells of the colon may allow the infection of vascular and lymphatic system, migration to other places, causing secondary infections. AIP-based system is not the only one in E. faecalis QS. Cytolysin production by the enterococci is repressed at the level of transcription by two proteins: membrane connected sensor, CylR1 and DNA connected proteins, CylR2 (Haas et al., 2002). When cytolysin concentration outside the cell is low, CylR2

represses genes encoding subunits of cytolysins, cylLL and cylLS, by cylL promoter bond. CylR1 monitors cytolysin concentration outside the cell and while the threshold concentration is reached, CylR1 changes the ability to bond DNA by CylR2. It results from CylR2 secretion by cylL promoter and increase in the cytolysin production. Cytolysin is regulated in a manner dependent on population, by means of its detection rather than AIP detection. Streptococcus mutans uses the QS signaling system, which is dependent on competence stimulating peptide (CSP), to regulate diverse physiological activities including bacteriocin production, genetic transformation, and biofilm formation. The expression of QS-associated genes was increased 3.4-5.3-fold by CSP in biofilms. Cell viability of S. mutant grown in biofilms is affected by the CSP-dependent QS system (Zhang et al., 2009). Summary It is becoming more understandable that bacterial cell-to-cell communication plays an important role in interactions between bacteria and host. This is true for both natural symbiotic microflora and pathogenic “invading” bacteria. A great number of bacteria employ QS for regulation of various phenotypes as a part of their pathogenic or symbiotic lifestyles. In the case of a host’s microflora, signaling particles are utilized to control density of the population for a better response coordination of numerous population, frequently occurring in the created, complex biofilm. Some pathogens use also signaling particles to coordinate a reaction dependent on population in case of a huge number of bacteria needed for colonization. They also use signaling particles produced by other populations, like natural microflora as indicator of their presence in host organism. Using that kind of strategy, the “passing” bacteria activate pathogenic genes only when they are in the environment where pathogenesis is successful. It secures bacteria from pointless using of additional part of energy required for pathogenesis. A higher relatedness between the bacteria infecting a host (lower strain diversity) will lead to more prudent exploitation of the host, and hence lower virulence. A higher relatedness will favor higher levels of cooperation that in turn allows the host to be exploited more efficiently, and hence a higher virulence (Rumbaugh et al., 2009). QS molecule signals may modulate the physiology of other microbes in as-yetundiscovered interspecies interactions (Shank and Kolter, 2009). QS signal molecules, although largely considered as effectors of QS-dependent gene expression are also emerging as multifunctional molecules that influence life, development and death in single

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Communication between microorganisms and virulence factors production

and mixed microbial populations and impact significantly the outcome of host-pathogen interactions (Williams and Cámara, 2009).

Literature Ahmer B.M., J. van Reeuwijk, C.D. Timmers, P.J. Valentine and F. Heffron. 1998. Salmonella Typhimurium encodes an SdiA homolog, a putative quorum sensor of the LuxR family, that regulates genes on the virulence plasmid. J. Bacteriol. 180: 1185–1193. Blevins J.S., M.O. Elasri, S.D. Allmendinger, K.E. Beenken, R.A. Skinner, J.R. Thomas and M.S. Smeltzer. 2003. Role of sarA in the pathogenesis of Staphylococcus aureus musculoskeletal infection. Infect. Immun. 71: 516–523. Clarke M.B., D.T. Hughes, C. Zhu, E.C. Boedeker and V. Sperandio. 2006. The QseC sensor kinase: a bacterial adrenergic receptor. Proc. Natl. Acad. Sci. USA 103: 10420–10425. Coulthurst S.J., N.R. Williamson, A.K. Harris, D.R. Spring and G.P. Salmond. 2006. Metabolic and regulatory engineering of Serratia marcescens: mimicking phage mediated horizontal acquisition of antibiotic biosynthesis and quorum-sensing capacities. Microbiology 152: 1899–1911. De Kievit T.R. and B.H. Iglewski. 2000. Bacterial quorum sensing in pathogenic relationships. Infect. Immun. 68: 4839–4849. Devine J.H., G.S. Shadel and T.O. Baldwin. 1989. Identification of the operator of the lux regulon from the Vibrio fischeri strain ATCC7744. Proc. Natl. Acad. Sci. USA 86: 5688–5692. Gillaspy A.F., S.G. Hickmon, R.A. Skinner, J.R. Thomas, C.L. Nelson and M.S. Smeltzer. 1995. Role of the accessory gene regulator (agr) in pathogenesis of staphylococcal osteomyelitis. Infect. Immun. 63: 3373–3380. Girand G. and G.V. Bloemberg. 2008. Central role of quorum sensing in regulating the production of pathogenicity factors in Pseudomonas aeruginosa. Future Microbiol. 3: 97–106. Haas W., B.D. Shepard and M.S. Gilmore. 2002. Two component regulator of Enterococcus faecalis cytolysin responds to quorum-sensing autoinduction. Nature 415: 84–87. Hammer B.K. and B.L. Bassler. 2009. Distinct sensory pathway in Vibrio cholerae El Tor and classical biotypes modulate cyclic dimeric GMP levels to control biofilm formation. J. Bacteriol. 191: 169–177. Hoffman L.R., H.D. Kulasekara, J. Merson, L.S. Houston, J.L. Burns, B.W. Ramsey and S.I. Miller. 2009. Pseudomonas aeruginosa lasR mutant are associated with cystic fibrosis lung disease progression. J. Cyst. Fibros. 8: 66–70. Hsieh H.Y., Ch.W. Tseng and G.C. Stewart. 2008. Regulation of Rot expression in Staphylococcus aureus. J. Bacteriol. 190: 546–554. Kanamaru K., K. Kanamaru, I. Tatsuno, T. Tobe and C. Sasakawa. 2000. SdiA, an Escherichia coli homologue of quorum-sensing regulators, controls the expression of virulence factors in enterohaemorrhagic Escherichia coli O157:H7. Mol. Microbiol. 38: 805–816. Li M., A.E. Villaruz, V. Vadyvaloo, D.E. Sturdevant and M. Otto. 2008. AI-2-dependent gene regulation in Staphylococcus epidermidis. BMC Microbiol. 8: 1–9. Liang H., L. Li, Z. Dong, M.G. Surette and K. Duan. 2008. The YebC family protein PA0964 negatively regulates the Pseudomonas aeruginosa quinolone signal system and pyocyanin production. J. Bacteriol. 190: 6217–6227. Lyon G.J., P. Mayville, T.W. Muir and R.P. Novick. 2000. Rational design of a global inhibitor of the virulence response in Staphylococcus aureus, based in part on localization of the site

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Polish Journal of Microbiology 2009, Vol. 58, No 3, 199–204 ORIGINAL PAPER

Identification and Molecular Modeling of a Novel Lipase from an Antarctic Soil Metagenomic Library HUBERT CIEŒLIÑSKI*1A, ANETA BIA£KOWSKA2A, KAROLINA TKACZUK2, ANNA D£UGO£ÊCKA1, JÓZEF KUR1 AND MARIANNA TURKIEWICZ2 1 Department

of Microbiology, Chemical Faculty, Gdañsk University of Technology, Poland of Technical Biochemistry, Technical University of £ódŸ, Poland

2 Institute

Received 2 July 2009, revised 17 July 2009, accepted 25 July 2009 Abstract In this work, we present the construction of a metagenomic library in Escherichia coli using pUC19 vector and environmental DNA directly isolated from Antarctic topsoil and screened for lipolytic enzymes. Screening on agar supplemented with olive oil and rhodamine B revealed one clone with lipolytic activity (Lip1) out of 11000 E. coli clones. This clone harbored a plasmid, pLip1, which has an insert of 4722 bp that was completely sequenced from both directions. Further analysis of the insert showed three open reading frames (ORFs). ORF2 encoded a protein (Lip1) of 469 amino acids with 93% identity to the uncultured Pseudomonas sp. lipase LipJ03. Amino acid sequence comparison and phylogenetic analysis indicated that Lip1 lipase was closely related to family I subfamily 3. Furthermore, we present a three-dimensional model of lipase Lip1 which was generated based on the two known structures of mesophilic lipases from Pseudomonas sp. MIS 38 (PML lipase, PDB; 2Z8X) and Serratia marcescens (SML lipase, PDB: 2QUB). Finally, we report the results of comparisons between lipase Lip1 and mesophilic lipases and point out similarities and differences in the catalytic site and in other parts of the analyzed structures. K e y w o r d s: cold-adapted lipase, metagenomic library, molecular modeling

Introduction Metagenomics, the study of genomic content DNA isolated from environmental samples, has proved particularly useful for the analysis of uncultured bacteria because greater than 99% of microbes found in natural environments cannot be cultured with currently available technologies (Schloss and Handelsman, 2003; Streit et al., 2004; Daniel, 2004). Over the last few years, the metagenome-based strategy has been commonly used to explore some novel enzymes and molecules for biotechnological and pharmaceutical applications, such as lipases (Wei et al., 2008; Henne et al., 2000; Lee et al., 2004; Elend et al., 2007; Jeon et al., 2009), esterases (Jeon et al., 2008), cellulases (Kim et al., 2008), xylanases (Hu et al., 2008) and novel antibiotics turbomycin A and B (Gillespie et al., 2002) from different environments. One approach in the search for novel biocatalysts is to generate the metagenomic library from soils that are known to harbor a high level of microbial diversity

and thus, potentially a wide diversity of biocatalysts (Curtis and Sloan, 2005). A further development of this approach is to create the metagenomic library from an environment that has been subjected to extreme conditions in the likelihood that enzymes from such an environment will be able to function under those extreme conditions (Torsvik and Øvreås, 2002). Among the extreme environments, antarctic soil constitutes an attractive biocatalytic resource because of its abundance and broad diversity of cold-adapted microorganisms. Such microorganisms have developed adaptive mechanisms to perform their metabolic functions at low temperatures by incorporating unique features mainly in their proteins. Compared to proteins from mesophiles, cold-adapted proteins show decreased ionic interactions and hydrogen bonds, possess fewer hydrophobic groups and more charged groups ion on their surface as well as longer surface loops. Due to these modifications, at low temperatures psychrophilic proteins lose their rigidity and gain increased structural flexibility for enhanced catalytic

* Corresponding author: H. Cieœliñski, Department of Microbiology, Gdañsk University of Technology, Narutowicza 11/12, 80-952 Gdañsk, Poland; phone: (+48) 58 347 16 05; fax: (+48) 58 347 18 22; e-mail: [email protected] Hubert Cieœliñski and Aneta Bia³kowska contributed equally to this work.

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function. The adaptive properties of psychrophilic enzymes are their high specific activity at low temperatures, a relatively low apparent temperature optimum for activity and a rather relatively high thermolability (Hoyoux et al., 2004; Georlette et al., 2004; D’Amico et al., 2006; Siddiqui and Cavicchioli, 2006). This unique ability of extremophilic biocatalysts to catalyze reactions at low or moderate temperatures offers great industrial and biotechnological potential (Gerday et al., 2000; Cavicchioli et al., 2002). One of the methods for prospecting these molecules with high potential for downstream application is metagenome-based approaches (Elend et al., 2007). Recently, increasing attention has focused on a search for cold-adapted lipolytic enzymes. These enzymes are expected to be applied as additives to detergents used at low temperatures and biocatalysts for biotransformation of labile compounds at low temperature. Their molecular flexibility is higher relative to enzymes from mesophilic organisms and makes them particularly useful in stereoselective reactions, carried out in organic solvents and at low temperatures and yielding almost exclusively the isomer produced through conversion having the lower activation energy (Cavicchioli et al., 2002). In the present study, we constructed a library of environmental DNA from soil sample collected from Adelie penguin rookery, which is located in the neighborhood of Henryk Arctowski Polish Antarctic Station at King George Island. The screening for novel lipolytic enzymes from uncultured soil microorganisms revealed one clone, Lip1, with a strong lipolytic activity. Further analysis revealed that the Lip1 clone harbored a novel lip1 gene encoded lipase belonging to family I subfamily 3. The three-dimensional structure of this lipase has not yet been determined but we have modeled the 3D structure of metagenomic enzyme (Lip1) on the basis of the sequence of its gene. We analyzed the amino acid composition of lipase in comparison to mesophilic counterparts and characterized the architecture of the catalytic site The features potentially responsible for cold-adaptation and stabilization at low temperatures were also identified. Experimental Materials and Methods

Bacterial strains, plasmids and growth media. E. coli TOP10F’ (Invitrogen) was used as the host strains for metagenomic DNA library construction and screening for lipolytic active clones. Plasmid pUC19 (Invitrogen) was used as DNA vector for metagenomic DNA library construction. The E. coli strain was grown on LB medium (Sambrook and Russel, 2001), supplemented with 100 µg/ml of ampi-

3

cillin, 0.1 mM isopropyl-$-D-thiogalactopyranoside (IPTG), 2% (vol/vol) olive oil and 0.001% (wt/vol) rhodamine B (the fluorescent dye). Colonies were incubated for 1 day at 37°C, followed by incubation for 2 days at 25°C. Orange fluorescent halos around lipase-positive E. coli strains could be seen when these plates were exposed to UV light of 312 nm (Kouker and Jaeger, 1987). Ampicillin, IPTG and rhodamine B were purchased from Sigma-Aldrich (USA). Sampling. The analyzed soil sample was collected from Adelie penguin rookery (S 62°09’46’’, W 58°27’42’’) which is located in the neighborhood of Henryk Arctowski Polish Antarctic Station at King George Island. Sampling depth was 5 cm. Next, the soil sample was sieved to remove penguin dung debris and particulate matter larger than 2 mm and was kept without disturbance at 4°C until analysis. General DNA manipulations. Restriction enzymes, T4 DNA ligase were purchased from Fermentas (Lithuania). Restriction enzymes and other DNAmodifying enzymes were used according to the manufacturer’s recommendations. The reagents for PCR and various oligodeoxynucleotides were purchased from DNA-Gdansk II (Poland). Metagenomic DNA isolation from Antarctic soil sample. Soil samples of 5 g were mixed with 13.5 ml of DNA extraction buffer (100 mM Tris-HCl [pH 8.0], 100 mM sodium EDTA [pH 8.0], 100 mM sodium phosphate [pH 8.0], 1.5 M NaCl, 1% CTAB) and 25 µl of lysozyme (10 mg/ml, A&A Biotechnology, Poland) in 15 ml Falcon tubes by horizontal shaking at 225 rpm for 30 min at 37°C. Next, 50 µl of proteinase K (20 mg/ml, A&A Biotechnology, Poland) was added to the sample and incubated again at the same conditions as described above. After the shaking treatment, 1.5 ml of 20% SDS was added, and the samples were incubated in water bath at 65°C for 2 h with gentle end-over-end inversions every 15 to 20 min. The supernatants were collected after centrifugation at 6 000× g for 10 min at room temperature and transferred into 50-ml centrifuge tubes. The soil pellets were extracted two more times by adding 4.5 ml of extraction buffer and 0.5 ml of 20% SDS, vortexing for 10 s, incubating at 65°C for 10 min, and centrifuging as before. Supernatants from the three cycles of extractions were combined and mixed with an equal volume of chloroform: isoamyl alcohol (24:1, vol/vol). The aqueous phase was recovered by centrifugation and precipitated with 0.6 volume of isopropanol at room temperature for about 24 h. The pellet of crude nucleic acids was obtained by centrifugation at 16 000× g for 20 min at room temperature, washed with cold 70% ethanol, and resuspended in sterile deionized water, to give a final volume of 500 µl. The extracted DNA was further purified by Genomic Mini kit (A&A Biotechnology, Poland), followed by final purification with Genomic AX Bacteria kit (A&A

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Molecular modeling of cold-adapted lipase

Biotechnology, Poland). The pure DNA samples were stored frozen at –20°C until it was required. Metagenomic library construction and screening for lipolytic activity. The pUC19 library of the purified DNA was constructed by the following procedures. DNA samples from Antarctic soil were used to construct metagenomic DNA library following partial digestion with HindIII and BamHI, optimized to maximize fragments in the 1–5 kbp size range. The fragments of 1–5 kbp resolved in an agarose gel were excised and concentrated. The BamHI- and HindIIIdigested pUC19 DNA was overhanged using T4 DNA ligase (Epicentre, USA) to genomic DNA library digested with the same restriction endonucleases, and E. coli TOP10F’ (Invitrogen, USA) were transformed with the ligation products. The lipase activities of the transformants were tested on Luria-Bertani agar containing: 100 µg/ml of ampicillin, 0.1 mM isopropyl$-D-thiogalactopyranoside (IPTG), 2% (vol/vol) olive oil and 0.001% (wt/vol) rhodamine B (the fluorescent dye) designed as LB-lipRB medium. Analysis of the lipolityc active E. coli clone. The E. coli clone showing orange halos was inoculated into a 250 ml flask containing 50 ml of LB medium (100 µg/ml ampicillin). After overnight incubation at 37°C, the cells were harvested by centrifugation at 5 000 × g for 15 min and washed twice with sterile distilled water. Next, the plasmid DNA was isolated using Plasmid Mini isolation kit (A&A Biotechnology, Poland), retransformed into E. coli TOP10F’ and the new clones were examined on the LB-lipRB medium for lipase activity. The plasmid DNA from the one of the positive clones (pink fluorescence at UV light 312 nm) was isolated as described above and designed as pLip1. Next, a few samples with plasmid pLip1 DNA were digested with selected restriction enzymes to create restriction maps of examined plasmid. The nucleotide sequence of the DNA insert from pPINK-UV plasmid was determined using an ABI 3730 xl/ABI 3700 sequencing technology (Genomed, Poland). The open reading frames were detected using the ORF search tool provided by the National Center for Biotechnology Information (NCBI). The predicted function of ORFs was annotated using BlastX search toward the NCBI nonredundant protein database (Altschul et al., 1997). Sequence data analysis and molecular modeling. GenBank was scanned for related sequences by the BLAST algorithm. Nucleotide and amino-acid sequences were aligned by using the computer program ESPript (Gouet et al., 1999). Protein structure prediction was carried out via the GenSilico MetaServer (http://gensilico.pl/meta/, Kurowski and Bujnicki, 2003), which is a convenient gateway to a number of publicly available online services for secondary structure prediction.

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The alignment between the sequences of metagenomic lipase Lip1 and its mesophilic counterparts from the database was the first step of homology modelling. The set of alternative models was generated using both MODELLER (Sali et al., 1995) and SWISS-MODEL (Guex and Peitsch, 1997). The models were scored using MetaMQAPII metaserver (Paw³owski et al., 2008). The final model was obtained using the ‘Frankenstein’s Monster’ approach (Kosinski et al., 2003), which comprises cyclic realignment in poorly scored regions and merging the best scoring fragments. The sequence-structure fit in the final model was also evaluated using MetaMQAPII and ProQ server (Paw³owski et al., 2008), while the stereochemical parameters were assessed with PROCHECK (Laskowski et al., 1993). Modeling of the lipase Lip1 was performed using templates selected based on the results of the Genesilico MetaServer. The templates used for the modeling were the crystal structures of two lipases from Pseudomonas sp. MIS 38 (Angkawidjaja et al., 2007) and Serratia marcescens (Meier et al., 2007). Nucleotide sequence accession number. The sequences of the lip1 gene was deposited in the GenBank database with accession number GQ352455. Results and Discussion Construction and screening of a metagenomic library. A metagenomic library consisting of ~11,000 clones was constructed using DNA isolated from soil sample collected from Adelie penguin rookery which is located in the neighborhood of Henryk Arctowski Polish Antarctic Station at King George Island. To screen for lipase-producing clones, the pUC19derivaite clones were plated onto LB-lipRB medium. One clone with lipolytic activity was detected and designated as LIP1. Molecular analysis of pLIP1. The insert of the recombinant plasmid recovered from Lip1 clone was sequenced. The nucleotide sequence analysis revealed the presence of 3 ORFs longer than 100 amino acid residues that exhibited similarities to genes annotated with predicted functions (Table I). The ORF’s sequence analysis with using BlastX search toward the NCBI nonredundant protein database revealed that ORF2 encoded protein with high sequence similarity (93% identity) with lipase LipJ03 from uncultured Pseudomonas sp. strain. The Lip1 protein consisted of 469 amino acids with a deduced molecular mass (Mw) of 49 372 Da. Further analysis of lipase Lip1 sequence revealed that the enzyme belonged to family I subfamily 3 of lipases. The program BLAST which was used to screen for sequences of lipolytic enzymes deposited in NCBI

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Cieœliñski H. et al. Table I Sequence analysis of the ORFs encoded in metagenomic insert of pLip1

ORF #

Length (amino acids)

1 2 3

235 469 442

Putative function (most similar homologue)

Putative source organism

Accession number

hypothetical protein PFLU3139 Pseudomonas fluorescens SBW25 YP_002872714 lipase uncultured Pseudomonas sp. AAU12351 outer membrane protein Pseudomonas fluorescens BAA88494

showed that the metagenomic antarctic lipase displayed the highest sequence identity to the lipase from the non-culturable bacterium Pseudomonas (93% sequence identity) (Jiang et al., 2006). Molecular modeling. Bioinformatic analysis of metagenomic lipase Lip 1 molecule revealed the similarity in its sequence to other lipases with known structures, isolated from culturable bacteria. Among

% Identity/ similarity

E value

99/99 93/97 57/78

3e–126 0.0 8e–88

the proteins of known crystallographic structure, Lip1 showed the maximum sequence identity to PML Pseudomonas sp. MIS 38 lipase (72%) and SML lipase from Serratia marcescens (63%) (Fig. 1) that served as templates for modeling of the metagenomic lipase structure using the “Frankenstein monster” approach. The antarctic lipase Lip1 consists of 469 amino acid residues. In contrast to PML and SML enzymes, Lip 1

Fig. 1. Sequence alignment of the metagenomic Antarctic lipase and lipases from Pseudomonas sp. MIS38 (PML lipase, PDB; 2Z8X) and Serratia marcescens (SML lipase, PDB: 2QUB) that were templates used for structure modeling (conserved sequences are marked).

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Molecular modeling of cold-adapted lipase

Fig. 2. Model of Lip1 lipase (the successive fragments of the molecule with defined secondary structure are marked in different colors: dark blue from the N-terminus and red at the C-terminus; the domains formed by these fragments are also shown in the model).

is characterized by a lack of 138 amino acids in the polypeptide chain. The evidence that the lack of a fragment of the polypeptide chain is possible was provided by an earlier experiment of Jiang et al. (2006) who isolated from the non-culturable bacterium Pseudomonas the gene of the homologous enzyme (by using the genome-walking approach), expressed it in the yeast Pichia pastoris and obtained the biologically active protein. This means that the lacking structural element in lipase Lip1 and lipase LipJ03 from non-culturable Pseudomonas strain is not characteristic of all lipases belonging to family I.3 and it is not essential for retaining their activity. The Lip1 lipase consists of two domains (Fig. 2). The N-terminal domain shows a modified "/$ hydrolase fold and it is rich in "-helices, while C-domain contains two $-roll motifs, laterally stacked together forming the so-called sandwich, similar to that of SML and PML. In the C-terminal part of C-domain Lip1 there are several repeats of the RTX motif – GGXG XDX(U)X (U, hydrophobic amino acids), which are responsible for activity and folding of this group of lipolytic enzymes (Fig. 3). This repeated motif of PML and SML has been proposed to function as an intramolecular chaperone, because deletion or mutation of this motif generates inactive proteins, which are incompletely folded (Meier et al., 2007; Angkawidjaja et al., 2007). The active site of lipases is defined by a canonical catalytic triad, which in Lip1 consists of Ser207, Asp255,

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Fig. 3. The model structure of Lip1 lipase (clusters of Gly residues in C-terminal domain are marked in dark blue).

and His 313. These residues superimpose quite well with the corresponding amino acids from other lipases, e.g. lipases from Pseudomonas sp. MIS 38 or Serratia marcescens (Fig. 4).

Fig. 4. Active center of the metagenomic Lip1 lipase (catalytic amino acid residues are marked with colors).

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Conclusions. In this paper we present the identification, isolation of a new lipase lip1 gene from Antarctic soil metagenomic library and analysis of the model in silica of lipase encoded by lip1 gene. The results indicate that lipase encoded by gene lip1 showed a lack of 138 amino acids in the polypeptide chain in comparison to the structures of well characterized mesophilic analogs: PML lipase from Pseudomonas sp. MIS 38 and SML lipase from Serratia marcescens. On the other hand, the analyzed lipase showed high sequence similarity (93%) to cold-adapted LipJ03 lipase from non-culturable Pseudomonas strain with optimum temperature activity 35°C and pH 8.0, respectively (Jiang et al., 2006). The results presented in this paper encourage us to further study the lip1 gene product. To this end we intend to construct a recombinant E. coli strain for Lip1 lipase production and purify the recombinant enzyme for further characterization. Literature Altschul S.F., T.L. Madden, A.A. Schaffer, J. Zhang, Z. Zhang, W. Miller and D.J. Lipman. 1997. Gapped BLAST and PSIBLAST: a new generation of protein database search programs. Nucleic Acids Res. 25: 3389–3402. Angkawidjaja C.,  D.J. You,  H. Matsumura,  K. Kuwahara, Y. Koga,  K. Takano and S. Kanaya. 2007. Crystal structure of a family I.3 lipase from Pseudomonas sp. MIS38 in a closed conformation. FEBS Lett. 581: 5060–5064. Cavicchioli R., K.S. Siddqui, D. Andrews and K.R. Sowers. 2002. Low-temperature extremophiles and their applications. Curr. Opin. Biotechnol. 13: 253–261. Curtis T.P. and W.T. Sloan. 2005. Exploring microbial diversity – a vast below. Science 309: 1331–1333. D’Amico S., T. Collins, J.C. Marx, G. Feller and Ch. Gerday. 2006. Psychrophilic microorganisms:challenges for life. EMBO 7: 385–389. Daniel R. 2004. The soil metagenome – a rich resource for the discovery of novel natural products. Curr. Opin. Biotechnol. 15: 199–204. Elend C., C. Schmeisser, H. Hoebenreich, H.L. Steele and W.R. Streit. 2007. Isolation and characterization of a metagenomederived and coldactive lipase with high stereospeciWcity for (R)-ibuprofen esters. J. Biotechnol. 130: 370–377. Georlette D., V. Blaise, T. Collins, S. D’Amico, E. Gratia, A. Hoyoux, J.C. Marx, G. Sonan, G. Feller and C. Gerday. 2004. Some like it cold: biocatalysis at low temperatures FEMS Microbiol. Rev. 28: 25–42. Gerday Ch., M. Aittaleb, M. Bentahir, J.P. Chessa, P. Claverie, T. Collins, S. D’Amico, J. Dumont, G. Garsoux, D. Georlette, A. Hoyoux, T. Lonhienne, M.A. Meuwis, and G. Feller. 2000. Cold-adapted enzymes: from fundamentals to biotechnology. Trends Biotechnol. 18: 103–107. Gillespie D.E., S.F. Brady, A.D. Bettermann, N.P. Cianciotto, M.R. Liles, M.R. Rondon, J. Clardy, R.M. Goodman and J. Handelsman. 2002. Isolation of antibiotics turbomycin A and B from a metagenomic library of soil microbial DNA. Appl. Environ. Microbiol. 68: 4301–6. Gouet P., E. Courcelle, D.I. Stuart and F. Metoz. 1999. ESPript: multiple sequence alignments in PostScript. Bioinformatics. 15: 305–8. Guex N. and M.C. Peitsch. 1997. SWISS-MODEL and the SwissPdbViewer: an environment for comparative protein modeling. Electrophoresis 18: 2714–2723.

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Henne A., R.A. Schmitz, M. Bömeke, G. Gottschalk and R. Daniel. 2000. Screening of environmental DNA libraries for the presence of genes conferring lipolytic activity on Escherichia coli. Appl. Environ. Microbiol. 66: 3113–6. Hoyoux A., V. Blaise, T. Collins, S. D’Amico, E. Gratia, A.L. Huston, J.C. Marx, G. Sonan, Y. Zeng, G. Feller and Ch. Gerday. 2004. Extreme catalysts from low-temperature environments. J. Biosci. Bioeng. 98: 317–330. Hu Y., G. Zhang, A. Li, J. Chen and L. Ma. 2008. Cloning and enzymatic characterization of a xylanase gene from a soil-derived metagenomic library with an efficient approach. Appl. Microbiol. Biotechnol. 80: 823–30. Jeon J.H., J.T. Kim, S.G. Kang, J.H. Lee and S.J. Kim. 2008. Characterization and its potential application of two esterases derived from the Arctic sediment metagenome. Mar. Biotechnol. (NY). Sep 24. [Epub ahead of print] Jeon J.H., J.T. Kim, Y.J. Kim, H.K. Kim, H.S. Lee, S.G. Kang, S.J. Kim and J.H. Lee. 2009. Cloning and characterization of a new cold-active lipase from a deep-sea sediment metagenome. Appl. Microbiol. Biotechnol. 81: 865–74. Jiang Z., H. Wang, Y. Ma and D. Wei. 2006. Characterization of two novel lipase genes isolated directly from environmental sample. Appl. Microbiol. Biotechnol. 70: 327–32. Kim S.J., C.M. Lee, B.R. Han, M.Y. Kim, Y.S. Yeo, S.H. Yoon, B.S. Koo and H.K. Jun. 2008. Characterization of a gene encoding cellulase from uncultured soil bacteria. FEMS Microbiol. Lett. 282: 44–51. Kosinski J., I.A. Cymerman, M. Feder, M.A. Kurowski, J.M. Sasin and J.M. Bujnicki. 2003. A „Frankenstein’s monster” approach to comparative modeling: merging the finest fragments of Fold-Recognition models and iterative model refinement aided by 3D structure evaluation. Proteins, 53:369–379. Kouker G. and K.E. Jaeger. 1987. Specific and sensitive plate assay for bacterial lipases. Appl. Environ. Microbiol. 53: 211–213. Kurowski M.A. and J.M. Bujnicki. 2003. GeneSilico protein structure prediction meta-server. Nucleic Acids Res. 31: 3305–7. Laskowski R.A., M.W. MacArthur, D.S. Moss and J.M. Thornton. 1993. PROCHECK – a program to check the stereochemical quality of protein structures. J. Appl. Cryst. 26: 283–291. Lee S.W., K. Won, H.K. Lim, J.C. Kim, G.J. Choi and K.Y. Cho. 2004. Screening for novel lipolytic enzymes from uncultured soil microorganisms. Appl. Microbiol. Biotechnol. 65: 720–6. Meier R.,  T. Drepper,  V. Svensson,  K.E. Jaeger and U. Baumann. 2007. A calcium-gated lid and a large beta-roll sandwich are revealed by the crystal structure of extracellular lipase from Serratia marcescens. J. Biol. Chem. 282: 31477–31483. Pawlowski M., M.J. Gajda, R. Matlak and J.M. Bujnicki. 2008. MetaMQAP: a meta-server for the quality assessment of protein models. BMC Bioinformatics 9: 403. Sali A., L. Potterton, F. Yuan, H. van Vlijmen and M. Karplus. 1995. Evaluation of comparative protein modeling by MODELLER. Proteins 23:318–326. Sambrook J. and D.W. Russell. 2001. Molecular Cloning; a Laboratory Manual, 3rd . Cold Spring Harbour, NY: Cold Spring Harbour Laboratory. Siddiqui K.S. and R. Cavicchioli. 2006. Cold-adapted enzymes. Annu. Rev. Biochem. 75:403–433. Schloss P.D. and J. Handelsman. 2003. Biotechnological prospects from metagenomics. Curr. Opin. Biotechnol. 14: 303–310. Streit W.R., R. Daniel and K.E. Jaeger. 2004. Prospecting for biocatalysts and drugs in the genomes of non-cultured microorganisms. Curr. Opin. Biotechnol. 15: 285–290. Torsvik V. and L. Øvreås. 2002. Microbial diversity and function in soil: from genes to ecosystems. Curr. Opin. Microbiol. 5: 240–245. Wei P., L. Bai, W. Song and G. Hao. 2008. Characterization of two soil metagenome-derived lipases with high specificity for p-nitrophenyl palmitate. Arch. Microbiol. 191: 233–240.

Polish Journal of Microbiology 2009, Vol. 58, No 3, 205–209 ORIGINAL PAPER

Use of Zwitterionic Type of Detergent in Isolation of Escherichia coli O56 Outer Membrane Proteins Improves their Two-Dimensional Electrophoresis (2-DE) GABRIELA BUGLA-P£OSKOÑSKA*, BO¯ENA FUTOMA-KO£OCH, ALEKSANDRA SKWARA, W£ODZIMIERZ DOROSZKIEWICZ

Department of Microbiology, Institute of Genetics and Microbiology, University of Wroc³aw, Poland Received 3 July 2009, revised 24 July 2009, accepted 30 July 2009 Abstract Escherichia coli O56 were originally isolated from infected humans. Here it is reported that using the zwitterionic detergent (Zwittergent Z 3-14) to isolate outer membrane proteins (OMPs) from Escherichia coli O56 is suitable for their separation by two-dimensional electrophoresis (2-DE) using pH 3-10 immobilized pH gradient IPG strips (BIO-RAD). K e y w o r d s: Escherichia coli O56, outer membrane proteins, two-dimensional electrophoresis

Introduction Proteomics is a powerful tool that is used in many parts of biology. One of them is electrophoretic study of outer membrane proteins (OMPs) of Gram-negative bacteria. These structures of the bacterial cells are the keys that interact with the environment. It has been discovered that changes in OMPs expression can be a reason for the resistance of Gram-negative bacteria to the bactericidal action of serum (Bugla et al., 2004; Bugla-P³oskoñska and Doroszkiewicz, 2006; Futoma et al., 2005; Mielnik et al., 2001). The resistance of bacteria to serum’s lytic activity as a result of the expression of many virulence factors may be essential in the development of sepsis and septic shock. Taylor and Parton (Taylor and Parton, 1976) proved that a 46 kDa OMP plays a decisive role in Escherichia coli resistance to serum. Studies of Kroll and co-authors (Kroll et al., 1983) and Taylor and Parton (Taylor and Parton, 1976) have shown that treating E. coli cells with sera generates changes in their composition of OMPs. Proteomics proves that OMPs are virulence factors of many diseases, for example proteins: OmpA, IbeA, IbeB, IbeC, AslA, TraJ of E. coli are involved in meningitidis infections (Badger and Kim, 1998).

E. coli O56 were originally isolated from infected humans from mesenterial lymph node (Orskow et al., 1977). It was determined (Mielnik et al., 2001; BuglaP³oskoñska and Doroszkiewicz, 2006), that bacteria of this serotype are sensitive to the bactericidal activity of normal cord serum (NCS) and normal bovine serum (NSB). Gamian and co-authors (Gamian et al., 1994) have shown that sialic acid is a component of the O-specific part of the polysaccharide chains of E. coli O56. Sialic acids may contribute to the pathogenicity of the microorganisms by mimicking host tissue components (Vimr and Lichtensteiger, 2002). On the other hand, sialic acid as a component of bacterial capsules activates the complement system in serum. This was shown for encapsulated Streptococcus agalactiae, which is the most common cause of neonatal sepsis and meningitis (Aoyagi et al., 2008). Recent investigations are also based on finding and testing the protective potential of OMPs against life-threating invasive bacterial infections caused i.e. Neisseria meningitidis serogroup B (Jessouroun et al., 2004) and Pseudomonas aeruginosa (Sorichter et al., 2009). Proteomics gives possibilities to find molecular candidates for vaccines. Witkowska and co-workers (Witkowska et al., 2006) proved that a 38 kDa OMP

* Corresponding author: G. Bugla-P³oskoñska, Department of Microbiology, Institute of Genetics and Microbiology, University of Wroc³aw, Przybyszewskiego 63/77, 51-148 Wroc³aw, Poland; phone: (+48) 71-3756306; fax: (+48) 71-3252151, e-mail: [email protected]

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present in most Enterobacteriaceae species is a protein that generates immunological response in human organisms and is a good candidate for creating a vaccine against such species as Escherichia coli, Shigella flexneri, Klebsiella pneumoniae or Proteus vulgaris. Hamid and Jain (Hamid and Jain, 2008) showed that immunization of mice with a 49 kDa OMP gives them 100% survival after being treating with a lethal dose of Salmonella Typhimurium. For any bacterial proteomic study sample preparation is a crucial step and is a critical influential factor in isoelectric focusing (IEF). The aim of this paper is to present a 2-DE procedure in conjunction with the isolation of bacterial OMPs using the zwitterionic detergent – Zwittergent Z 3-14®. Experimantal Materials and Methods

Bacterial cell culture. Escherichia coli O56 PCM 2372 from the collection of the Institute of Immunology and Experimental Therapy, Wroc³aw, Poland, PCM (NCTC 9056-National Collection of Type Cultures, Central Public Health Laboratory, London) were inoculated into 50 ml of Brain Heart Infusion broth (Difco) in 200 ml shake flasks and left to grow at 37°C for 18 h. Isolation of outer membrane proteins (OMPs). The procedure of isolation of OMPs was done according to Murphy and Bartos (Murphy and Bartos, 1989) with minor modifications. Bacterial cells from an overnight culture were harvested (4000 rpm at 4°C for 15 min) and the pellet was suspended in 1.25 ml of buffer b [1 M sodium acetate (POCh), 0.001 M $-mercaptoethanol (Merck)]. Then 11.25 ml of a water solution containing 5% (w/v) Zwittergent Z 3-14 (Calbiochem) and 0.5 M CaCl2 (POCh) was added. This mixture was stirred at room temperature (RT) for 1 h. To precipitate nucleic acids, 3.13 ml of 96% (v/v) cold ethanol (POCh) was added very slowly. The mixture was then centrifuged at 12 300 rpm at 4°C for 10 min. The proteins in the supernatant were precipitated by the addition of 46.75 ml of 96% (v/v) cold ethanol and centrifuged at 12 300 rpm at 4°C for 20 min. The pellet was left to dry at ambient temperature and then suspended in 2.5 ml of buffer Z [0.05% (w/v) Zwittergent Z 3-14, 0.05 M Trizma-Base (Sigma) and 0.01 M EDTA (Sigma), pH 8.0] and stirred at RT for 1 h. The solution was kept at 4°C overnight and centrifuged at 8 700 rpm at 4°C for 10 min. OMPs were present in the soluble fraction of buffer Z after the centrifugation. The preparations of OMPs in the soluble fraction of buffer Z were checked for the presence of succinic dehydrogenase activity, a marker for cytoplasmic

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membranes, using the method described by Rockwood et al. (1987). Protein quantification. Protein quantification was performed with the BCA Protein Assay Kit (PIERCE®) according to Smith et al. (Smith et al., 1985) with bovine serum albumine (BSA) (Sigma) as the standard. The Pierce BCA Protein Assay is a detergentcompatible formulation based on bicinchoninic acid (BCA) for the colorimetric detection and quantification of total protein. The purple-colored reaction product (reduction Cu2+ to Cu+1 by protein in an alkaline medium) exhibits a strong absorbance at 562 nm with increasing protein concentration over a broad range: 20–20,000 µg/ml) (PIERCE Instruction). Removal of salts, buffers and small ionic contaminants from buffer Z containing OMPs. In the next stages of preparation a 2-DE Sample Preparation for Soluble or Insoluble Proteins (PIERCE) was used according to the manufacturer’s instructions. After the desalting procedure (Protein Desalting Spin Columns, PIERCE) the samples of OMPs were suspended in a buffer (PIERCE) that was compatible for 2-DE. Two-dimensional electrophoresis (2-DE). The OMPs from E. coli O56 were separated on a series of 7 cm pH 3–10 immobilized pH gradient (IPG) strip. The electrophoresis separation of proteins was performed essentially as described by O’Farrell (O’Farrell, 1975). 2-DE was carried out with the PROTEAN® IEF Cell (BioRad). The main reagents for 2-DE were purchased from Bio-Rad and basically used according to the manufacturer’s instructions (BioRad Instruction Manual). Rehydratation. IPG strips (7 cm) were rehydrated prior to isoelectric focusing. The rehydratation step was performed outside the PROTEAN® IEF Cell (passive rehydratation). A total amount of 169 µg of OMPs was suspended in 125 µl of rehydratation buffer (8M urea, 0.5% CHAPS, 10mM DTT). The rehydrated strips were positioned in the focusing tray and covered with mineral oil (BioRad). The time of the rehydratation process amounted 16 hours. Isoelectric focusing was conducted for: Step 1: 250 V, 20 min (linear); Step 2: 4 000 V, 120 min (linear); Step 3: 4 000 V, 160 min (rapid). Total volt-hours parameter reached 14 000 V-hr. After IEF the IPG strips were removed from the focusing tray and were transferred into rehydratation/ equilibration buffer for 20 min (BioRad). SDS-PAGE. For the second dimension the IPG strips were applied onto a 9–12.5% gradient SDSpolyacrylamide gel (PAGE) using 1% (w/v) agarose in the running buffer. Gels were electrophoresed according to Laemmli (Laemmli, 1970). Tricine was used instead of glycine in the electrophoresis buffer [0.05 M Trizma Base, 0.05 M tricine, 0.1% (w/v) sodium dodecyl sulfate (SDS), pH = 8.2]. The gels were run at 5°C and at 35 V for 125 min. After that they were

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stained for 24 h with coloidal Coomassie Brilliant Blue R-250 (Merck). The gel images were scanned using PDQuest 2-D Analysis Software v. 8.0.1 (BioRad). The tests were repeated three times. Result and Discussion Detection and identification of OMPs is of particular interest as they play important functions in signal transduction pathways, bacterial-host interactions, and other processes. However, detection of hydrophobic OMPs in 2-DE gels is associated with certain limitations (Fountoulakis, 2005). The poor solubility of hydrophobic OMPs accounts for their absence from the 2D gel map, but the addition of zwitterionic detergents can improve protein solubilization (Shaw and Riederer, 2006). The anionic nature of sodium dodecyl sulfate (SDS) detergents generally limits their effectiveness for proteomic analyses. Zwitterionic detergents have found widespread use in 2-DE (Luche et al., 2003; Henningsen et al., 2002). Zwitterionic detergents lack conductivity and electrophoretic mobility and are also suited for breaking protein-protein interactions (Srirama, 2001). Sample preparation is a very crucial step in 2-DE. Some modifications (use of other detergents, different way of sample preparation) were introduced into the 2-DE protocol suggested by O’Farrell (O’Farrell, 1975) and BioRad (Instruction Manual, 2008) which significantly impaired the resolution of proteins. In this case, the detergent Zwittergent 3-14® was used for the isolation of OMPs and not Nonidet P-40. Additionally, in the methods proposed by O’Farrell isoelectric focusing gels were made in glass tubing. The samples of bacterial OMPs were tested for the presence of biological membranes. The preparations were assayed for the enzymatic activity of succinic dehydrogenase, a marker for the cytoplasmic membrane. The Zwittergent-extracted OMPs from E. coli O56 contained no detectable activity of succinic dehydrogenase which confirms that they were free of membrane contaminations. Each sample of OMPs, appropriately prepared for 2-DE separation (i.e. desalting), was run in triplicate: three gels were obtained from E. coli O56 samples. Fig. 1 shows representative examples of the OMPs separated on a 2-DE gel, to which 169 µg of total OMPs protein (per gel) was applied. Approximately 59 (a), 62 (b), and 63 (c) protein spots were detected on Coomassie Brilliant Blue R-250 gels (Fig. 1). Fig. 2 shows the master gel which was generated on the basis of the (a), (b), and (c) gels visualisations (Fig. 1). The master gel shows 63 separated spots of OMPs of E. coli O56. The obtained results show that Zwittergent Z 3-14® is suitable for the isolation of OMPs from E. coli O56. This paper confirms that detergents of this type may

Fig. 1. 2-DE profile of OMPs of E. coli O56 isolated with Zwittergent Z 3-14®. OMPs were separated using pH 3–10 IPG strips (BioRad) and 9–12.5% SDS-PAGE. Gels were stained with the Coomassie Brilliant Blue R-250 (Merck) and preliminarily analyzed with PDQuest 2-D Analysis Software v. 8.0.1 (BioRad).

be used in the isolation of OMPs and subsequently in 2-DE with IPG strips. During 2-DE analysis we marked the main spots and were able to visualize even 63 individual protein spots. This method is a promising tool for the characterisation of bacterial virulence factors as OMPs.

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Fig. 2. Master of three 2-DE gel profiles (from Fig. 1 a, b, c) of OMPs of E. coli O56 isolated with Zwittergent Z 3-14® and analyzed with PDQuest 2-D Analysis Software v. 8.0.1 (BioRad).

2-DE methodology has a potential for the rapid development of specific, safe, and highly efficacious vaccines against infection caused by E. coli in humans and livestock. 2-DE electrophoretic methods have been successfully used for the separation of, among others, Escherichia coli (Molloy et al., 2000), Salmonella Typhimurium (Hamid and Jain, 2008), Brucella abortus (Connolly et al., 2006), Edwardsiella tarda (Kawai et al., 2004), Shigella flexneri (Peng et al., 2004), and Leptospira interrogans (Cullen et al., 2002) OMPs, but none of the authors used zwitterionic detergents in the stages of OMPs preparation. They used buffer with urea (10 mM Tris-HCl, pH 7.5, 10 mM EDTA, and 6M urea) (Hamid and Jain, 2008), detergent TRITON X-114 (Cullen et al., 2002), sodium lauryl sarcosinate (Peng et al., 2004). We conclude that Zwittergent Z 3-14® detergent is as an effective detergent for the isolation of the OMPs of E. coli O56 and can be adapted to 2-DE using IPG strips. Acknowledgments We would like to thank Prof. Andrzej Gamian for providing the Escherichia coli O56 strains. We thank mgr Ilona Dudka for her contribution to this study. This work was supported by a grant from the University of Wroc³aw – 2931/W/IGMI/08 – to Dr Gabriela Bugla-P³oskoñska.

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Polish Journal of Microbiology 2009, Vol. 58, No 3, 211–218 ORIGINAL PAPER

Purification and Biochemical Characteristic of a Cold-Active Recombinant Esterase from Pseudoalteromonas sp. 643A under Denaturing Conditions ANNA D£UGO£ÊCKAa, HUBERT CIEŒLIÑSKIa, PIOTR BRUDZIAK1, KAROLINA GOTTFRIED, MARIANNA TURKIEWICZ2 and JÓZEF KUR*

Department of Microbiology, Gdañsk University of Technology, Gdañsk, Poland of Physical Chemistry, Gdañsk University of Technology, Gdañsk, Poland 2 Institute of Technical Biochemistry, £ódŸ, Poland

1 Department

Received 3 July 2009, revised 14 July 2009, accepted 15 July 2009 Abstract In this paper production of a cold-active esterase EstA from the Antarctic bacterium Pseudoalteromonas sp. 643A in E. coli expression system was described. The purification and biochemical characteristic of EstA were performed in the presence of urea and then compared with results obtained for the esterase with no addition of urea and isolated from the native source. In both cases the cold-active enzyme displayed similar properties. However, the differences concerning thermal activity were observed. The optimal temperature for recombinant esterase in the presence of urea (1 M) was about 15°C lower in comparison with enzyme isolated from the native source. Furthermore, the EstA was found to be more thermolabile in denaturant conditions. The differences were presumably caused by slightly changed protein structure in the presence of urea. The preservation of activity of EstA dissolved in buffer containing 8M urea suggests that the protein structure is retained and it does not undergo dramatic changes due to high urea concentration. This thesis was confirmed with FT-IR data. K e y w o r d s: Pseudoalteromonas sp., cold-active enzyme; esterase, denaturing conditions, urea

Introduction Lipases and esterases, collectively known as lipolytic enzymes, hydrolyze hydrophobic long- and shortchain carboxylic acid esters, respectively (Arpigny and Jaeger, 1999; Fojan et al., 2000; Singh et al., 2006). They are also able to catalyze esterification, transesterification and enantioselective hydrolysis reactions (Pandey et al., 1999). Due to the lack requirement for cofactors, stability in organic solvents, chemo-, regioand stereoselectivity lipolytic enzymes are important biocatalysts in many branches of industry. They are widely used in food industry, manufacturing of cosmetics and detergents, fats and oils processing, syntheses of fine chemicals and pharmaceuticals, in paper making, sewage treatment plants, and in polymer decomposition (Jaeger and Reetz, 1998; Jaeger et al., 1999; Pandey et al., 1999; Sharma et al., 2001; Jaeger and Eggert, 2002). Nowadays an increasing attention has been drawn to potential applications of cold-active lipolytic enzymes isolated from psychro-

philic and psychrotrophic microorganisms (Rashid et al., 2001; Kulakova et al., 2004; Ryu et al., 2006) due to the high catalytic activity at low temperature and low thermostability. Our group has isolated a psychrotrophic bacterium Pseudoalteromonas sp. strain 643A, producing a novel cold-active esterase EstA belonging to the GDSL family of bacterial lipolytic enzymes (Cieœliñski et al., 2007). The subclass of GDSL esterases and lipases is characterized by a broad substrate specificity and regiospecificity, so these enzymes could be used in synthesis and hydrolysis reactions in many branches of industry (Lo et al., 2003; Akoh et al., 2004). We have isolated and sequenced the gene encoding esterase EstA of Pseudoalteromonas sp. 643A and characterized the purified enzyme from the native source. However, its heterologous overexpression in E. coli was not satisfying either due to formation of inclusion bodies and its probable integration in E. coli cell membrane (Cieœliñski et al., 2007), or ineffective extracellular secretion (D³ugo³êcka et al., 2008).

* Corresponding author: J. Kur, Gdañsk University of Technology, Department of Microbiology, Narutowicza 11/12, 80-952 Gdañsk, Poland; phone: (+48) 58 347 23 02; fax: (+48) 58 347 18 22; e-mail: [email protected] a Anna D³ugo³êcka and Hubert Cieœliñski contributed equally to this work.

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In this study we describe a new E. coli expression system in which the cold-active esterase was produced without its signal sequence to be accumulated in E. coli cells and to avoid its integration into the cell membrane. The recombinant esterase was purified under denaturing conditions (8 M urea) and its biochemical characteristics were assayed without removing the urea from the reaction mixture. Moreover, we estimated the optimal concentration of urea with respect to purified enzyme activity and the changes in the enzyme structure. To this end, the EstA protein structure was determined using the FT-IR technique in the presence and absence of urea. Experimental Material and Methods

Bacterial strains, plasmids, growth conditions. E. coli TOP10F’ (Invitrogen) and BL21(DE3) (Novagen) were used as the host strains for DNA manipulation and gene expression, respectively. Plasmid pUC19 (Invitrogen) was used for subcloning, and pET22b(+) (Novagen) was used as expression vector. Plasmid pLipo1 (Cieœliñski et al., 2007) was used for PCR amplifications of estA gene. The E. coli strain was grown on LB medium (Sambrook and Russel, 2001), supplemented with ampicillin (100 µg/ml), and IPTG (0,5 mM) (Sigma). IPTG (isopropyl-$-D-thiogalactopyranoside) was used as expression inducer. General DNA manipulations. DNA manipulations were carried out according to the standard procedures (Sambrook and Russel, 2001) or manufacturer’s recommendations. Restriction enzymes were purchased from Fermentas and DNA ligase was purchased from Epicentre. DNA polymerase Pwo and other PCR reagents were purchased from DNA Gdañsk II s.c. Kits for plasmid isolation and DNA purification were purchased from A&A Biotechnology. Electrophoresis. Protein fractions were analysed by sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) on 15% gel slabs and stained with Commassie blue (Walker, 1996). The amount of recombinant protein was evaluated by the optical densitometry of SDS-PAGE gels using the Quantity One program (Bio-Rad) with the bovine serum albumin (Sigma) as a standard. Expression of the recombinant esterase gene in E. coli and protein purification. Primers used for amplification of the estA gene of Pseudoalteromonas sp. 643A were: EstHis-F 5’-ATACATATGCACCATCAT CATCATCAT G A C A A C A C G AT T T TA ATA C ACGGAG-3’ (containing NdeI recognition site and sequence encoding poliHis domain) and Est-R 5’CCCAAGCTTTTAGACGTTATTTAACCAC-3’

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(containing HindIII recognition site) were used. The boldface parts of primer sequences are complementary to the nucleotide sequences of the Pseudoalteromonas sp. 643A EstA esterase gene, whereas added recognition sites for restriction endonucleases are underlined. The obtained PCR product, encoded EstA esterase devoid of its signal sequence and with Histag domain at the N-termini, was cloned into SmaI site of pUC19 vector, resulting in recombinant plasmids of pUC19-EstAHis. The correctness of the constructed plasmid was confirmed by DNA sequencing using ABI 3730 xl/ABI 3700 sequencing technology (AGOWA GmbH). Expression plasmid for production of His-tagged esterase EstA was constructed by excising the estA gene from pUC19-EstAHis by NdeI and HindIII and ligated into the pET22b(+) plasmid digested with the same endonucleases to give the expression plasmid pET22b(+)-EstAHis. The E. coli strain Bl21(DE3) cells transformed with pET22b(+)EstAHis were grown for 16 h at 37°C in LB medium containing ampicillin (0.1 mg/ml). The preculture was inoculated (1%) into fresh LB medium containing ampicillin and cultivation was continued at 37°C to OD600 of 0.5. The culture was then supplemented with 0.5 mM IPTG and grown for 3 h at 30°C to achieve the overexpression of GDSL-esterase gene. Cells were harvested by centrifugation at 16 000× g for 10 min, resuspended 50 mM Tris-HCl buffer pH 8.0 containing 300 mM NaCl, 5 mM imidazole, and 8 M urea, disrupted by sonification, and cell debris was removed by centrifugation at 16 000× g for 10 min. The enzyme was purified on His-Bind Resin (Novagen) using denaturing conditions, with a gravity flow of 2 ml min–1. For washing 50 mM Tris-HCl buffer pH 8.0 containing 300 mM NaCl, 5 mM imidazole, and 8 M urea was used and esterase EstA-His was eluted with a 50 mM Tris-HCl buffer pH 8.0 containing 300 mM NaCl, 250 mM imidazole, and 8 M urea. Protein was then dialyzed against 50 mM Tris-HCl buffer pH 8.0 containing 8 M urea (for activity assays) or against 20 mM phosphate buffer pH 8.0 (for FT-IR). Enzyme assays. Esterase activity was determined spectrophotometrically by measurements of concentration of p-nitrophenol (at 405 nm) released from p-nitrophenyl butyrate. Reaction mixtures contained 5 µl of 50 mM substrate solution and 5 µl of enzyme solution buffered in 100 µl of 50 mM Tris-HCl pH 7.5 were incubated at 20°C for 5 min. To determine urea influence on EstA activity substrate solution and enzyme solution were buffered in 50 mM Tris-HCl pH 7.5 containing urea at different concentrations. However the characterization of EstA esterase under denaturing conditions was performed for the final concentration 0.4 M of urea in the reaction mixture. One unit of esterase activity is equivalent to 1 mmol of p-nitrophenol released from the p-nitrophenyl bu-

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Cold-active esterase from Antarctic bacterium

tyrate in 1 min at 20°C. For routine enzyme assay, p-nitrophenyl butyrate (pNPB) (Sigma) was used as esterase substrate. To determine enzyme specificity various p-nirophenyl derivatives were used. Shortchain fatty acid esters of p-nitrophenol (p-nitrophenyl acetate, p-nitrophenyl butyrate, p-nitrophenyl caprylate, and p-nitrophenyl decanoate) were dissolved in acetonitrile whereas long-chain fatty acid esters (p-nitrophenyl palmitate and p-nitrophenyl stearate) were dissolved in n-hexane. FT-IR measurements and analysis. Protein pellet suspension (1ml) dialyzed against 20 mM phosphate buffer pH 8.0 was resuspended (not solubilized) in 100 µl of 20 mM phosphate buffer pH 8.0 or completely solubilized in 20 mM phosphate buffer pH 8.0 containing 8 M urea. To prepare the calibration curve ten solutions of urea were prepared in the range 0 to 9 M urea (pH 8.0). The refractive index of each sample was measured with thermostated refractometer (PZO). High urea concentration excluded the usage of classical transmission cell. Instead, attenuated total reflection accessory (ATR) had been used, which allowed to obtain satisfactory spectra of protein and proteinurea solutions. The spectra were recorded on Nicolet 8700 spectrometer (Thermo Electron Co.), using the Golden Gate single-reflection ATR accessory (Specac). The temperature during measurements was kept at 25 ± 0.1°C using electronic temperature controller (Specac). For each spectrum 128 scans were collected with a resolution of 4 cm–1. The spectrometer source was on Turbo mode during measurement. The spectrometer and the ATR accessory were purged with dry nitrogen to diminish water vapour contamination of spectra. All ATR spectra were water vapour subtracted and corrected using advanced ATR correction algorithm (part of the OMNIC software). The absorbance value at 1160 cm–1 was used to determine the urea concentration in protein sample. The corresponding urea spectrum was interpolated using Yanusz.ab program run under Grams/32 (Galactic Industry Corporation) and subtracted from protein spectrum. The reference water spectrum was subtracted from the protein spectra to obtain straight horizontal line in the range of 1900–1720 cm–1. The spectra were smoothed with a 13-point Savitzky-Golay algorithm. The difference spectra were analysed using the OMNIC (Thermo Electron Corporation) and Grams/32 programs (Galactic Industry Corporation). Results and Discussion Gene cloning, expression, and purification of EstA. EstA esterase was produced as a His-tagged protein at its N-terminus. Additionally, to avoid accumulating in cell membrane or in periplasm of E. coli,

Table I EstA activity at different urea concentrations. Urea concentation [M]

Specific activity [U × mgEstA–1]

0.08 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 2 4 6 8

387.0 ± 2.34 431.0 ± 2.53 464.0 ± 2.19 464.0 ± 2.50 420.0 ± 2.78 420.0 ± 2.11 409.0 ± 4.19 464.0 ± 2.43 442.0 ± 2.32 464.0 ± 2.17 663.0 ± 2.45 420.0 ± 2.54 132.0 ± 2.03 126.0 ± 2.80 99.0 ± 3.23

Relative activity [%] 58 65 70 70 63 63 62 70 67 70 100 63 20 19 15

Average ± SD, (n = 3).

the enzyme was devoid of its signal sequence. However, the enzyme accumulated in the cytoplasm as inclusion bodies, which were easily solubilized in buffers containing 8 M urea. Surprisingly, esterase activity was at high level and was found to precipitate during dialysis against buffers without urea. The EstA esterase shown the highest activity in the presence of 1 M urea whereas it partially lost the activity at lower or higher denaturant concentrations (Table I). The applied expression system was quite efficient, giving about 140 mg of esterase from 1 L induced culture. As a result of purification in ‘denaturant’ conditions about 80 mg of electrophoretically homogenous protein was obtained, with a specific activity of 420 U mgEstA–1 (Fig. 1),which was the highest activity in comparison with all previous attempts (Table II). The enhancement of activity in the presence of urea at low concentration was also observed for some other enzymes (Kaufman, 1968; Narayanasami et al., 1997; Zhang et al., 1997; Kumar et al., 2003; Shahnawaz et al., 2007). The presence of urea (at low concentration) in the environment of the protein causes some slight conformal changes in the neighborhood of the active site. These changes in the secondary and tertiary protein structure might promote the more open enzyme conformation (Deshpande et al., 2001; Kumar et al., 2003) or make the active site more flexible (Zhang et al., 1997). The esterase EstA Pseudoalteromonas sp. 643A which possesses its esterolytic activity in the presence of urea might be useful due to its simple purification procedure for some biotechnological applications: there is no need to remove the denaturant after solubilisation of inclusion bodies. However, it is important

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Fig. 1. Purification of EstA esterase. Lane 1 – Unstained Protein MW Marker (Fermentas); lane 2 – E. coli BL(DE3), cell fraction; lane 3 – E. coli BL(DE3) + pET22b(+)-EstAHis, soluble fraction of total lysate; lane 4 – fraction of unbounded proteins; lane 5 – fraction of protein collected during washing; lane 6 – protein fraction after purification and dialysis.

to evaluate the optimal concentration of urea with respect to enzyme concentration in the reaction condition e.g. 1 M urea for recombinant EstA enzyme. On the other hand, for some applications the purification of EstA might be unnecessary – the E. coli proteins coproduced with EstA should be inactive in the presence of denaturing agent. Characterization of EstA esterase at denaturing conditions. The optimum temperature for activity of

recombinant enzyme was 20°C. Esterase retained above 40% of maximum activity at 0°C and lost it rapidly above 20°C (Fig. 2a). It was found to be stable at the temperature near to 0°C and it was completely unstable after 1 h incubation at 30°C (Fig. 2b). The optimum pH range for the enzyme activity and stability is from 7.0 to 8.5 (Fig. 2c, d). Studies of the substrate specificity of purified recombinant EstA esterase performed by comparing enzymatic activity

Table II Comparison of various methods for EstA isolation and purification. Data relate to protein purified from 1 L culture. Total protein Total activity Specific activity [mg] [U] [U/mgEstA]

Isolation and purification EstA from a native source – Pseudoalteromonas sp. 643A Recombinant EstA produced in E. coli cells, purified from inclusion bodies Recombinant EstA co-produced in E. coli with ABC transporters of Pseudoalteromonas sp., purified from culture medium Recombinant EstA produced in E. coli cells, purified in urea-containing buffer

Table III EstA activity for p-nitrophenyl esters. Substrate p-nitrophenyl acetate p-nitrophenyl butyrate p-nitrophenyl caprylate p-nitrophenyl decanoate p-nitrophenyl palmitate p-nitrophenyl stearate

No. of C atoms in alkyl chain 2 4 8 10 16 18

Relative activity [%] 62 100 21 3 1 0

Reference

3.78

10.8

51

Cieœliñski et al., 2007 Cieœliñski et al., 2007; unpublished data

23

0.21

0.01

28

362

10.3

D³ugo³êcka et al., 2008; unpublished data

80

3.3 104

420

This study

towards various chromogens (p-nitrophenyl derivatives) revealed that the preferred substrate for EstA esterase is p-nitrophenyl butyrate, but it hydrolyses also esters of shorter- (p-nitrophenyl acetate) and longer-chain (p-nitrophenyl caprylate) fatty acids (Table III). Enzyme was strongly or completely inhibited by 5 mM Cd2+, Co2+, Cu2+, Ni2+, and Zn2+ ions, whereas Ca2+ and Mg2+ ions were found to activate it (Table IV). Furthermore, 5 mM thiol compounds strongly decreased esterase activity (Table V). Recombinant esterase was also activated by EDTA,

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Cold-active esterase from Antarctic bacterium

Fig. 2. Effects of temperature and pH on EstA activity and stability. (a) – temperature dependence of EstA activity. (b) – thermal stability after 1 h incubation of EstA at various temperatures; (c) – pH dependence of EstA activity; (d) – pH-stability after 1 h incubation of EstA at various pH; comparatively, results for native EstA esterase (Cieœliñski et al., 2007) were added and indicated with dashed lines, results for recombinant EstA esterase with continuous line.

but it can be easily explained by chelating Ni2+ residues used in purification step. The performed biochemical characteristic of recombinant EstA revealed that enzyme produced in E. coli cells and dissolved in buffer with urea possesses simi-

lar properties (Tables III–V) to esterase isolated from the native source without addition of urea (Cieœliñski et al., 2007). The main difference is the temperature dependence activity (Fig. 2a) and thermal stability (Fig. 2b), respectively. Recombinant EstA in the

Table IV Effects of selected chemicals on EstA activity. The enzyme was incubated for 1 h at 20°C with the reagent (5 mM) and the residual activity was assayed under standard conditions. Reagent

Ca2+

Cd2+

Co2+

Cu2+

Mg2+

Mn2+

Ni2+

Zn2+

None

Residual activity [%]

103

18

30

12

189

69

4

8

100

Table V Effects of selected chemicals on EstA activity. The enzyme was incubated for 1 h at 20°C with the reagent (5 mM) and the residual activity was assayed under standard conditions. Reagent

Residual activity [%]

2-mercaptoethanol

13

DTT

10

EDTA

281

Glutathione-SH

20

PMSF

23

None

100

presence of urea was found to show the highest activity at the temperature of 20°C and it completely lost its stability after 1 h incubation at 30°C. In contrast, enzyme isolated from a native source without addition of urea displayed its maximal activity at the temperature of 35°C and was much more stable at higher temperatures- the complete inactivation occurred after 1 h incubation at 65°C. Another difference is that the recombinant enzyme in the presence of urea is active and stable in more narrow range (Fig. 2c, 2d). These differences were probably connected with the esterase EstA structure, which was slightly changed in the

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Fig. 3. Decomposition of amide I band into Gaussian-Lorentzian product components. (a) the amid I band (solid line) of EstA in 20 mM phosphate buffer pH 8.0; (b) EstA dissolved in 8M urea in 20 mM phosphate buffer, pH 8.0; the components (dashed line) are described with corresponding secondary structures (Arrondo and Goni, 1999; Barth and Zscherp, 2002; Natalello et al., 2005).

presence of urea (Desphande et al., 2001; Grinberg et al., 2008). The decreasing of the optimum temperature for the EstA activity in the presence of urea might be another advantage of using the enzyme for some specific applications. However, to explain the role of this denaturant in decreasing of the optimal temperature for enzyme activity, the esterase EstA structure without addition of urea has to be solved. Analysis of the EstA structure in the presence of urea. In the amide I region of protein spectra (1600–1700 cm–1) it is possible to distinguish several overlapped bands characteristic to various structures (e.g. $-sheets, "-helix) (Arrondo and Goni, 1999; Barth

and Zscherp, 2002). By means of different mathematical methods (second derivative, Fourier self-deconvolution) the amide I band can be resolved into individual components, allowing describing the secondary structure of a given protein. The peak-fitting technique allows describing the amide I band shape with a few synthetic bands of specified shape. Such resolved amide I band and its components allow to ascribe the percentage participation of particular secondary structure in the overall structure of a given protein (Barth and Zscherp, 2002). Figure 3a reveals that the main structural components of the EstA protein in 20 mM phosphate buffer are both "-helices (band near 1655 cm–1)

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Cold-active esterase from Antarctic bacterium

and $-sheets (band near 1630 cm–1). The data would seem to suggest that the protein consists of twice as many "-helices as $-sheets (34% and 17%, respectively). This is not quite consistent with available structure data of analogous E. coli thioesterase I/protease I/lysophospholipase L1 (PDB no.: 1IVN_A) which consists of 47–56% "-helices and 11–14% $-sheets, depending on secondary structure assignment algorithm. The arising differences probably result from the form of measured sample which was a precipitated protein suspended in phosphate buffer, not dissolved. It is possible that intermolecular interactions contribute in the $-sheet characteristic bandwidth (van de Weert et al., 2001). However, the spectrum of this protein still exhibits a shape characteristic for "/$ proteins. The preservation EstA activity dissolved in phosphate buffer containing 8 M urea suggests that the structure of this protein is retained and it does not undergo dramatic changes due to high urea concentration. This thesis can be confirmed with the FT-IR data. No band characteristic for protein denaturation can be seen in the spectra (Haris and Severcan, 1999; van de Weert et al., 2001; Natalello et al., 2005). Figure 3b suggests that secondary structure of the protein still can be described as consisting of " and $ structures. The proportions of both are similar to the protein suspended only in buffer, though exact analysis of bands area indicates, that participation of "-helix and $-sheet bands in the whole amide I band area is higher (42% and 20%, respectively). These are closer to the available structure of E. coli thioesterase I, particularly the "-helix content. Although the products of amide I band decomposition must be taken with great care, the results can suggest that structure of EstA protein might be slightly different than the E. coli thioesterase, especially the amount of $-sheet structure seems to be a little higher. Though, these results must be confirmed with other techniques. Conclusions. Research of recombinant lipolytic enzymes is significantly limited by their tendency to accumulate in the cytoplasm as inactive inclusion bodies (Chung et al., 1991; Zhuo et al., 2005). An active lipase or esterase can be then obtained by dissolving the inclusion bodies and then refolding the enzyme in buffer with denaturing agent, e.g. urea. In standard enzyme purification procedures the denaturing agent is removed at the next purification step to promote protein renaturation. However, in many cases the efficiency of the refolding procedure is low. Therefore, in our study we decided to modify the standard procedure of refolding the EstA esterase. In this aim we analyzed the impact of urea concentration on the structure (FT-IR measurements and analysis) and activity of the analyzed esterase. We found that the active EstA esterase could be effectively retrieved from

inclusion bodies and purified by immobilized metal affinity chromatography under denaturing conditions without removing the denaturing agent completely, which might enable the use of the EstA enzyme for some biotechnological applications. Literature Akoh C.C., G.C. Lee, Y.C., Liaw, T.H. Huang and J.F. Shaw. 2004. GDSL family of serine esterases/lipases. Prog. Lipid. Res. 43: 534–552. Arpigny J.L. and K.E. Jaeger. 1999. Bacterial lipolytic enzymes: classification and properties. Biochem. J. 343: 177–183. Arrondo J.L. and F.M. Goni. 1999. Structure and dynamics of membrane proteins as studied by infrared spectroscopy. Prog. Biophys. Mol. Biol. 72: 367–405. Barth A. and C. Zscherp. 2002. What vibrations tell us about proteins. Quart. Rev. Biophys. 35: 369–430. Chung G.H., Y.P. Lee, O.J. Yoo and J.S. Rhee. 1991. Cloning and nucleotide sequence of thermostable lipase gene from Pseudomonas fluorescens SIK W1. Appl. Microbiol. Biotechnol. 35: 237–241. Cieœliñski H., A.M. Bia³kowska, A. D³ugo³êcka, M. Daroch, K.L. Tkaczuk, H. Kalinowska, J. Kur and M. Turkiewicz. 2007. A cold-adapted esterase from psychrotrophic Pseudoalteromonas sp. strain 643A. Arch. Microbiol. 188: 27–36. Deshpande R.A., A.R. Kumar, I..Khan and V. Shankar. 2001. Ribonuclease Rs from Rhizopus stolonifer: lowering of optimum temperature in the presence of urea. Biochim. Biophys. Acta 1545: 13–19. D³ugo³êcka A., H. Cieœliñski, M. Turkiewicz, A.M. Bia³kowska and J. Kur. 2008. Extracellular secretion of Pseudoalteromonas sp. cold-adapted esterase in Escherichia coli in the presence of Pseudoalteromonas sp. components of ABC transport system. Protein Expr. Purif. 62: 179–84. Fojan P,. P.H. Jonson, M.T.N. Petersen and S.B., Petersen. 2000. What distinguishes an esterase from a lipase: A novel structural approach. Biochimie. 82: 1033–1041. Grinberg V.Y., T.V. Burova, N.V. Grinberg, T.A. Shcherbakova, D.T. Guranda, G.G. Chilov and V.K. Svedas. 2008. Thermodynamic and kinetic stability of penicillin acylase from Escherichia coli. Biochim. Biophys. Acta 1784: 736–746. Haris P.I. and F. Severcan. 1999. FTIR spectroscopic characterization of protein structure in aqueous and non-aqueous media. J. Mol. Catal. B: Enz. 7: 207–221. Jaeger K.E., B.W. Dijkstra and M.T. Reetz. 1999. Bacterial biocatalysts: molecular biology, three-dimensional structures, and biotechnological applications of lipases. Annu. Rev. Microbiol. 53: 315–351. Jaeger K.E. and T. Eggert. 2002. Lipases for biotechnology. Curr. Opin. Biotechnol. 13: 390–397. Jaeger K.E. and M.T. Reetz. 1998. Microbial lipases form versatile tools for biotechnology. Trends Biotechnol. 16: 396–403. Kaufman B.T. 1968. Studies on dihydrofolic reductase. III. Activation of the chicken liver enzyme by urea and thiourea. J. Biol. Chem. 243: 6001–6008. Kulakova L., A. Galkin, T. Nakayama, T. Nishino and N. Esaki. 2004. Cold-active esterase from Psychrobacter sp. Ant300; gene cloning, characterization, and the effects of Gly→Pro substitution near the active site on its catalytic activity and stability. Biochim. Biophys. Acta 1696: 59–65. Kumar A.R., S.S. Hegde, K.N. Ganesh and M.I. Khan. 2003. Structural changes enhance the activity of Chainia xylanase in low urea concentrations. Biochim. Biophys. Acta 1645: 164–171.

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Lo Y.C., S.C. Lin, J.F. Shaw and Y.C. Liaw. 2003. Crystal structure of Escherichia coli thioesterase I/protease I/lysophospholipases L1: consensus sequence blocks constitute the catalytic center of SGNH-hydrolases through a conserved hydrogen bond network. J. Mol. Biol. 330: 539–551. Narayanasami R., J.S. Nishimura, K. McMillan, L.J. Roman, T.M. Shea, A.M. Robida, P.M. Horowitz and B.S.S. Masters. 1997. The influence of chaotropic reagents on neuronal nitric oxide synthase and its flavoprotein module, urea and guanidine hydrochloride stimulate NADPH-cytochrome c reductase activity of both proteins. Nitric Oxide 1: 39–49. Natalello A., D. Ami, S. Brocca, M. Lotti and S.M. Doglia. 2005. Secondary structure, conformational stability and glycosylation of a recombinant Candida rugosa lipase studied by Fourier-transform infrared spectroscopy. Biochem. J. 385: 511–517. Pandey A., S. Benjamin, C.R. Soccol, P. Nigam, N. Krieger and V.T. Soccol. 1999. The realm of microbial lipases in biotechnology. Biotechnol. Appl. Biochem. 29, 119–131. Rashid N., Y. Shimada, S. Ezaki, H. Atomi and T. Imanaka. 2001. Low-temperature lipase from psychrotrophic Pseudomonas sp. strain KB700A. Appl. Environ. Microbiol. 67: 4064–4069. Ryu H.S., H.K. Kim, W.C. Choi, M.H. Kim, S.Y. Park, N.S. Han, T.K. Oh and J.K. Lee. 2006. New cold-adapted lipase from Photobacterium lipolyticum sp. nov. that is closely related to filamentous fungal lipases. Appl. Microbiol. Biotechnol. 70: 321–326.

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Polish Journal of Microbiology 2009, Vol. 58, No 3, 219–222 ORIGINAL PAPER

Usefulness of PCR Method for Detection of Leishmania in Poland PRZEMYS£AW MYJAK1*, JOANNA SZULTA1, MARCOS E. de ALMEIDA2, ALEXANDRE J. da SILVA2, FRANCIS STEURER2, ANNA LASS1, HALINA PIETKIEWICZ1, WAC£AW L. NAHORSKI1, JOLANTA GOLJAN1, JÓZEF KNAP3, BEATA SZOSTAKOWSKA1 1 Chair

of Tropical Medicine and Parasitology, Interfaculty Institute of Maritime and Tropical Medicine in Gdynia, Medical University of Gdañsk,, Poland, 2 Parasitc Diseases Branch, Division of Parasitic Diseases, Centers for Disease Control and Prevention, Atlanta, GA, USA, 3 Department of Environmental Hygiene and Parasitology, Institute of Agricultural Medicine, Lublin, Poland Received 2 July 2009, revised 15 July 2009, accepted 23 July 2009 Abstract Leishmania parasites are the etiological agents of leishmaniosis, with severe course and often fatal prognosis, and the global number of cases has increased in recent decades. The gold standards for the diagnosis of leishmaniosis are microscopic examinations and culture in vitro of the different clinical specimens. The sensitivity of these methods is insufficient. Recent development in specific and sensitive molecular methods (PCR) allows for detection as well as identification of the parasite species (subspecies). The aim of the study was to estimate the usefulness of molecular methods (PCR) for detection of Leishmania species and consequently for the implementation of such methods in routine diagnostics of leishmaniosis in Polish patients returning from endemic areas of the disease. In our investigations we used 54 known Leishmania positive DNA templates (from culture and clinical specimens) received from the CDC (Atlanta, GA, USA). Moreover, 25 samples of bone marrow, blood or other tissues obtained from 18 Polish individuals suspected of leishmaniosis were also examined. In PCR we used two pairs of primers specific to the conserved region of Leishmania kinetoplast DNA (kDNA) minicircle (13A/13B and F/R). Using these primers we obtained amplicons in all DNA templates from the CDC and in three Polish patients suspected for Leishmania infection. In one sample from among these cases we also obtained positive results with DNA isolated from a blood specimen which was previously negative in microscopic examinations. K e y w o r d s: leishmaniosis in Poland, molecular diagnosis (PCR)

Introduction Leishmaniosis is a parasitic infection that occurs in 88 tropical and subtropical countries except for Australia and Oceania. The disease is caused by several protozoan species of the genus Leishmania and is characterized by a wide variety of clinical forms from cutaneous (CL) and mucocutaneous (MCL) to visceral (VL) form with severe course and often fatal prognosis (Berman, 1997; Eddleston et al., 2006; Salata, 1993; Schwartz et al., 2006). Geographical distribution and the risk of getting infection are different for each form of the disease. The highest number of cutaneous leishmaniosis cases is observed in Afghanistan, Algeria, Iran, Iraq, and Saudi Arabia as well as in countries of Middle America

and South America where mucocutaneous form occurs with similar frequency. The visceral form of the disease is often reported on the African continent, in Southeast Asia and Middle America and South America. In Europe cutaneous and visceral leishmaniosis occur in the countries of Mediterranean Sea Basin. It is estimated that about 12 million people are infected worldwide, and there are recorded up to 2 million of new cases every year (Stefaniak et al., 2003). In Poland only a few cases are reported annually. The disease is diagnosed in individuals who had visited endemic areas. The reservoirs of Leishmania spp. are humans as well as domestic and wild animals. In the countries of the Mediterranean Sea Basin dogs are the main source of endemic persistence of the disease. Visceral

* Corresponding author: P. Myjak, Department of Tropical Parasitology, Interfaculty Institute of Maritime and Tropical Medicine, Medical University of Gdañsk, Powstania Styczniowego 9b, 81-519 Gdynia, Poland; phone: (+48) 058 3493740; fax: (+48) 058 6223354; e-mail: [email protected]

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leishmaniosis is caused by L. donovani, L. infantum or L. chagasi species, depending on the region. Cutaneous leishmaniosis is caused mainly by L. tropica or L. major, while the mucocutaneous form by L. braziliensis complex (Guerin et al., 2002; Salata, 1993; Schwartz et al., 2006). Parasites are transmitted by mosquitoes of Phlebotomus and Lutzomyia species, known as sandflies. Especially susceptible to infection are children, elderly people and immunocompromised persons (HIV infected, after organ transplantation, pharmacologically – immunosuppressed patients). Lately, the number of people infected with this parasite has increased among patients with HIV/AIDS. Protozoans of Leishmania spp. have a predilection to immune system cells, particularly to the spleen, liver and bone marrow, causing severe damage of these organs. The process often progresses slowly and in an apparent way resulting in deep lesions in affected sites (Berman, 1997; Guerin et al., 2002). Patients with leishmaniosis suffer from fever, which tends to be sustained or intermittent, progressive weakness, hepatosplenomegaly with signs of splenomegaly and lymphadenopathy. Accompanying pancytopenia often leads to misdiagnosis of hematologic malignancy and delays the institution of the appropriate treatment. The bases of the diagnosis of leishmaniosis are microscopic examinations of the biopsy material form spleen, bone marrow, lymph nodes or skin ulcers. In VL the most valuable is examination of the material from biopsy of the spleen, however it poses for patients a big risk of complications. The sensitivity of this method is insufficient. It is possible to culture the parasites in vitro using special media, most often NNN medium, which is more sensitive than the microscopic investigation (Boelaert et al., 2007; Reithinger and Dujardin, 2007). Out of immunological methods, the skin test (Montenegro test) is used or serological tests that turn out to be of little value in the case of cutaneous or mucocutaneous leishmaniosis. In order to identify the species isoenzymatic profile or monoclonal antibodies may be used. However, these methods need prior multiplication of the parasites using in vitro culture (Reithinger and Dujardin, 2007). The treatment of leishmaniosis is difficult, longlasting and expensive. Pentavalent antimonials, amphotericin B, pentamidyne, ketoconazole are still in use together with recently introduced miltefosine. Of prime importance is the treatment of concomitant illnesses such as tuberculosis, HIV/AIDS and others. Untreated visceral leishmaniosis leads to death in 90% of cases. Therefore, early and modern diagnosis is needed in order to implement effective treatment (Guerin et al., 2002; Olliario et al., 2005; Eddleston et al., 2006). Reithinger and Dujardin (2007) claim that for parasite detection in laboratories in countries of nonen-

demic areas there is a trend to prefer molecular diagnostics. Recent development in specific and sensitive molecular methods (PCR) allows for detection as well as identification of the parasite species (subspecies) (Belli et al., 1998; Marques et al., 2001; Reithinger and Dujardin, 2007). Therefore, the aim of this study was to estimate the usefulness of molecular methods (PCR) for detection of Leishmania spp. and consequently for their implementation in routine diagnostics of leishmaniosis in Polish patients arriving from endemic areas of the disease. Experimental Material and Methods

Leishmania infantum constantly cultured in vitro in Philips medium was used as a positive control in PCR assay. Specimens. The following known DNA samples received from the CDC (Atlanta, GA, USA) were investigated: 26 templates isolated from in vitro culture obtained from infected persons, 28 templates isolated directly from clinical specimens from patients with leishmaniosis and one sample isolated from the blood of an infected dog. These samples were characterized using Isoenzyme Analysis based on the CAE method (Cellulose Acetate Electrophoresis) developed by Kreutzer (Kreutzer et al., 1983; 1987; Kreutzer, 1996), using 6PGDH and GPI Isoenzymes. The obtained templates belonged to the most important species of Leishmania such as: L. donovani, L. infantum, L. chagasi, L. major, L. tropica and L. braziliensis. Moreover, 25 samples of bone marrow, blood or other tissues obtained from 18 Polish citizens suspected of leishmaniosis were also investigated. The samples were sent to the Department of Tropical Parasitology (Interfaculty Institute of Maritime and Tropical Medicine of Medical University of Gdañsk) in order to perform routine diagnostic examinations in the laboratory, after which they were frozen at –20°C. Written informed consents were obtained from all patients. In three cases amastigota forms of Leishmania spp. were detected in microscopic examination of the smears from bone marrow aspirates. DNA extraction. DNA was extracted from 100 µl of Leishmania in vitro culture (100 µg of tissue) or 100 µl of whole EDTA-stabilized bone marrow (blood) collected from Polish patients (kept frozen at –20°C) with the use of the Genomic Mini Kit or Blood Mini Kit (A&A Biotechnology, Gdynia, Poland), respectively. At the CDC (Atlanta, GA, USA) DNA was isolated using QIAamp DNA Micro Kit (Qiagen, Chatsworth, CA, USA). All DNA tem-

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Results and Discussion

Table I PCR master mixes (in µl) Primers

Reagent

13A/13B

PCR buffer for RUN polymerase dNTP 2,5 mM Primer 1 10 mM Primer 2 10 mM Tag Polymerase RUN 1U/µl Distilled water DNA template Total

F/R

2

2

2 1 1 0.625 16.375 13.375 2 5 (1:10) (1:5) 25

2 1 1 1 16 2 (1:5 or 1:10) 25

In parentheses (used dilutions of templates received from the CDC)

Table II PCR conditions Primers Step

13A/13B

F/R

Temp. °C Time m Temp. °C Time m Initial denaturation Denaturation Annealing Extension Final extension Number of cycles

94 94 52 72 72

3 1 1 1 10 40

94 94 60 72 72

5 1 1 0.5 5 30

plates were extracted according to the manufacturer’s instructions. PCR amplification. Two pairs of primers were used in this study: 13A (5’ GTG GGG GAG GGG CGT TCT-3’) and 13B (ATT TTA CAC CAA CCC CCA GTT-3’) (Bell et al., 1998) as well as F (5’ GGG (G/T)AG GGG CGT TCT (G/C)CG AA-3’ and R (5(G/C)(G/C)(G/C)(A/T)CT AT(A/T) TTA CAC CAA CCC C-3’) (Marques et al., 2001) that are specific to conserved region of all Leishmania species kinetoplast DNA (kDNA) minicircle. The length of amplified PCR products is about 120 bp. The composition of amplification reaction mixture as well as PCR conditions are presented in Table I and II, respectively. DNA amplifications were performed in the 7600 Gold thermocycler (Applied Biosystems, USA).

DNA isolated from promastigota forms obtained from in vitro culture of Leishmania allowed for optimization of the PCR reaction (data not shown). The DNA templates received from the CDC (Atlanta, USA) were used to perform PCR with different primers in the Polish laboratory. In PCR tests performed with these primer pairs using reference material from the CDC, the presence of appropriate Leishmania DNA fragments was detected in all 54 examined samples (Table III). However, a significant imperfection of these primers is their inability to identify Leishmania species. The methodology devised on the base of templates achieved from the CDC (Atlanta, USA) allowed for introducing PCR method into diagnostics of Polish patients. Examination of clinical samples taken from Polish patients gave positive results of PCR performed with the use of two pairs of primers in 3 cases (Table IV). Those were the patients (returning from Georgia, Turkey and Portugal) seropositive in Indirect Fluorescence Antibody test (IFA) and in which amastigota forms of the parasite were found in smears from bone marrow aspirates. In one sample from these cases we also obtained positive results with DNA isolated from a blood sample which was previously negative in microscopic examinations (Table IV). These patients were previously suspected of hematological neoplasms for several months and treatment was introduced in one case. In case of the other 15 persons suspected of leishmaniosis, where no parasites were found as well as the results of IFA tests were negative, none of the pair of primers gave a positive result in PCR reaction (Table IV). In diagnostics of leishmaniosis the most important is detection of the parasite or its DNA. Of prime importance is also identification of a particular species (subspecies), because the therapeutic response is species and, perhaps, even strain specific (Reithinger and Dujardin, 2007). For the routine diagnostics of leishmaniosis in Poland we suggest the adoption of primers encoding kDNA minicircle. It occurs in a high copy number in the cell and in consequence makes

Table III PCR results of templates obtained from the CDC Leishmania species Specimen Culture Clinical Total

L. infantum/ L. donovani L. infantum chagasi 2 4 5 3 4 5 5

L. major

L. tropica

5 5 10

7 6 13

L. braziliensis 8 4 12

Species not determined 4 1 5

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Myjak P. et al. Table IV PCR results of samples obtained from Polish patients Culture or Patient

Day from first Microscopic examination results

DNA isolation from

promastigota – amastigota – – amastigota – amastigota – – – – – –

in vitro culture blood bone marrow blood bone marrow bone marrow bone marrow bone marrow bone marrow bone marrow bone marrow bone marrow blood other tissues

positive control DJ

HR

CA N (7) N (5) N (3)

0 3 3 38 0 626 0 33 78 125 0 0 0

PCR results with primers 13A/13B

F/R

+ + + – – + – + – – – – – –

+ + + – – + – + – – – – – –

IFA results n.d. 1:160 n.d. n.d. n.d. 1 : 80 1 : 40 1 : 80 n.d. 1 : 80 1 : 20–1 : 40 – – –

+ = positive; – = negative; n.d. = not done; DJ = patients’ initials; N = number of patients

the reaction more sensitive. The best are the primers F/R, because primers 13A/13B may give additional unspecific products with templates isolated from clinical specimens. In conclusion, the results of this study indicate that the tested pairs of primers can be used for routine diagnostics of leishmaniosis in Poland. Acknowledgement The authors thank Alicja Rost and Ewa Zieliniewicz for excellent technical assistance.

Literature Belli A., B. Rodriguez, H. Aviles and E. Harris. 1998. Simplified polymerase chain reaction detection of new world Leishmania in clinical specimens of cutaneous leishmaniasis. Am. J. Trop. Med. Hyg. 58: 102–109. Berman J.D. 1997. Human leishmaniasis: clinical, diagnostic and chemotherapeutic developments in the last 10 years. Clin. Infect. Dis. 24: 684–703. Boelaert M., S. Bhattacharya, F. Chappuis, S.H. El Safi, A. Hailu, D. Mondal, S. Rijal, S. Sundar, M. Wasunna and R.W. Peeling. 2007. Evaluation of rapid diagnostic tests: visceral leishmaniasis. Nat. Rev. Microbiol. November: S30–S39. Eddleston M., R. Davidson, R. Wilkinson and S. Pierini (eds.) 2006. Oxford Handbook of Tropical Medicine, 2nd ed. Visceral leishmaniasis (kala-azar): 226–227. Oxford University Press.

Guerin P.J., P.L. Olliaro, S. Sundar, M. Boelaert, S.L. Croft, P. Desjeux, M.K. Wasunna and A.D. Bryceson. 2002. Visceral leishmaniasis: current status of control, diagnosis and treatment and a proposed research and development agenda. Lancet Infect. Dis. 2: 494–501. Kreutzer R.D. 1996. Genetic similarity among Central and South American populations of Leishmania (Viannia) braziliensis. Am. J. Trop. Med. Hyg. 55: 106–110. Kreutzer R.D., M.E. Semko, L.D. Hendricks and N.Wright. 1983. Identification of Leishmania spp. by multiple isozyme analysis. Am. J. Trop. Med. Hyg. 32: 703–715. Kreutzer R.D., N. Souraty and M.E. Semko. 1987. Biochemical identities and differences among Leishmania species and subspecies. Am. J. Trop. Med. Hyg. 36: 22–32. Marques M.J., A.C. Volpini, O. Genaro, W. Mayrink and A.J. Romanha. 2001. Simple form of clinical sample preservation and Leishmania DNA extraction from human lesions for diagnosis of American cutaneous leishmaniasis via polymerase chain reaction. Am. J. Trop. Med. Hyg. 65: 902–906. Olliaro P.L., P.J. Guerin, S. Gerstl, A.A. Haaskjold, J.A. Rottingen and S. Sundar. 2005. Treatment options for visceral leishmaniasis: a systemic review of clinical studies done in India, 1980–2004. Lancet Infect. Dis. 5: 763–767. Reithinger R. and J.C. Dujardin. 2007. Molecular diagnosis of leishmaniasis: current status and future applications. J. Clin. Microbiol. 45: 21–25. Salata R.A. 1993. Leishmaniasis. pp. 28–35. In: Mahmoud A.A.F. (ed.). Tropical and Geographical Medicine, Companion handbook. McGraw-Hill, Inc. Schwartz E., C. Hatz and J. Blum. 2006. New World cutaneous leishmaniasis in travellers. Lancet Infect. Dis. 6: 342–349. Stefaniak J., Paul M., Kacprzak E. and Koryna-Karcz B. 2003. Visceral leishmaniasis (in Polish). Przegl. Epidemiol. 57: 341–348.

Polish Journal of Microbiology 2009, Vol. 58, No 3, 223–229 ORIGINAL PAPER

Biofilm Formation as a Virulence Determinant of Uropathogenic Escherichia coli Dr+ Strains BEATA M. ZALEWSKA-PI¥TEK*, SABINA I. WILKANOWICZ, RAFA£ J. PI¥TEK and JÓZEF W. KUR

Department of Microbiology, Gdañsk University of Technology, Poland Received 2 July 2009, revised 9 July 2009, accepted 11 July 2009 Abstract Urinary tract infections are the most common health problem affecting millions of people each year. Uropathogenic Escherichia coli (UPEC) strains are the major factor causing lower and upper urinary tract infections. UPEC produce several virulence factors among which are surface exposed adhesive organelles (pili/fimbriae) responsible for colonization, invasion and amplification within uroepithelial cells. The virulence of the uropathogenic E. coli Dr+ IH11128 is associated with Dr fimbriae belonging to the Dr family of adhesins (associated with diarrhea and urinary tract infections) and a DraD protein capping the linear fiber at the bacterial cell surface. In this study we revealed that biofilm development can be another urovirulence determinant allowing pathogenic E. coli Dr+ to survive within the urinary tract. E. coli strains were grown in rich or minimal media, allowed to adhere to abiotic surfaces and analyzed microscopically by staining of cells with cristal violet. We found that both Dr fimbriae and DraD, exposed at the cell surface in two forms, fimbria-associated or fimbria non-associated, (DraE+/DraD+, DraE+/DraD– or DraE–/DraD+ E. coli strains) are required for biofilm formation. Additionally, we demonstrated the biofilm formation capacity of E. coli strains deficient in the surface secretion or production of the DraE adhesin. K e y w o r d s: E. coli Dr+, aggregation, biofilm, dra gene cluster, uropathogenic

Introduction Many bacterial strains live in sessile communities called biofilms. Biofilms are compact microbial communities consisting of organisms adherent to each other and a target surface (Geesey et al., 1977; Costerton et al., 1994; 1995). Biofilms are formed by a number of punctuated microcolonies separated from each other by liquid channels responsible for supplying nutrients (influx) and removal of the metabolic products (efflux). Biofilm development occurs in a few steps. First, the planktonic bacteria sessile on a biotic/abiotic surface adhere and create a cluster. Then the cells form antimicrobial resistant colonies. Some cells probably leave the biofilm colonies to settle on a surface and create new agglomerations (Costerton et al., 1994; 1995). In fact, bacterial biofilms can establish on any solid surface of inorganic or organic nature spanning a wide spectrum of environments (Schembri et al., 2001; Schembri and Klemm, 2001). Autoaggregation of bacterial cells can create more beneficial conditions for

colonization (Soto et al., 2007; Naves et al., 2008). Diseases connected with the ability of bacteria to form biofilms are generally chronic and difficult to treat because of biofilm resistance to antimicrobial agents compared to their planktonic (free-living) counterparts. The greater resistance to antibiotic therapy than the planktonic colonies is probably the effect of lack of penetration of biofilms by antibiotics (Costerton et al., 1999). Escherichia coli strains causing prostatitis produce biofilm communities in vitro more frequently than uropathogenic strains involved in cystitis and pyelonephritis (Soto et al., 2007). In those infections type 1 pili are considered to be an important factor in the first step of biofilm formation. Type 1 pili mediate binding to the host receptor and invasion of bacteria into bladder epithelial cells which activate a gene cascade essential for the formation of intracellular bacterial communities (Anderson et al., 2003; Justice et al., 2004). Urinary tract infections (UTI) are among the most common bacterial infections in humans and affect millions of people each year. The most frequent etiologic

* Corresponding author: B. Zalewska-Pi¹tek, Department of Microbiology, Gdañsk University of Technology; G. Narutowicza 11/12, 80-952 Gdañsk, Poland; phone/fax (+48) 58 3471822; e-mail: [email protected]

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agent of urogenital infections is uropathogenic E. coli, UPEC accounting for 65 to 90% of cases (Millar and Cox, 1997). Urovirulence factors associated with UPEC include surface (type 1 and P pili, S and F1C fimbriae, Dr family of adhesins) and exported virulence determinants (toxins, siderophores). Tissue invasion and biofilm formation can be other urovirulence factors which allow the bacterial cells to survive and persist a long time within the upper or lower urinary tract (Oelschlaeger et al., 2002; Emödy et al., 2003; Arisoy et al., 2006; Yamamoto, 2007). E. coli strains bearing the Dr family of adhesins account for 40% of pylenophritis cases in the third trimester of pregnancy, 50% of chronic diarrhea cases in children and 20% of recurrent urinary tract infections in young women (Foxman et al., 1995; Goluszko et al., 1997). The Dr family includes fimbrial (DraE adhesin) and afimbrial adhesins (AFA – I, –II, –III and –IV) (Servin, 2005; van Loy et al., 2002). The dra gene clusters share a highly conserved region, including the draF, draA, draB, draC, and draD genes (Nowicki et al., 1989). The draE gene, highly heterogeneous within the Dr family of adhesins, is the adhesin-encoding gene (Zalewska-Pi¹tek et al., 2008). Dr haemaglutinin is the only member of the Dr family of adhesins that has the unique ability to bind to DAF and type IV collagen receptors. The DraD protein binds to $1 integrins, a common receptor for AfaD invasins (Garcia et al., 1996; Plancon et al., 2003; Zalewska-Pi¹tek et al., 2008). The interactions with the host receptors are essential for the subsequent internalization of bacterial cells and the maintenance of chronic infections (Plancon et al., 2003; Cota et al., 2006). The adherence capacity of E. coli Dr+ can also stimulate bacterial aggregation, which can be the first step of biofilm development and formation of cell microcolonies (Zalewska-Pi¹tek et al., 2008). In this paper, we present the role of Dr fimbriae and DraD protein as virulence factors exposed at the surface of uropathogenic E. coli Dr+ strains, in biofilm formation under different nutrient conditions. The investigations involved the use of E. coli strains (of laboratory and clinical origin) expressing Dr fimbriae with or without DraD as a capping fimbrial subunit (DraE+ and DraD+/DraE+ and DraD–, respectively). Because the DraD protein can be expressed in two forms, fimbria-associated and fimbria non-associated, as a protein adhesive sheath surrounding the bacterial cells, it was very important to reveal the predisposition of UPEC to form structured bacterial communities in vitro without the fimbrial urovirulence determinant (DraE– and DraD+). The ability of the selected E. coli strains to create aggregates of bacterial cells was confirmed by analysis of biofilm growth in rich or minimal media and light microscopy of cells stained with cristal violet (CV).

Experimental Materials and Methods

Bacterial strains, plasmids and reagents. The expression of genes encoded by a dra gene cluster was carried out in laboratory Escherichia coli BL21(DE3) (Novagen, Nottingham, UK), or clinical E. coli IH11128 (Goluszko et al., 1997), DR14 and DR14/gspD strains (Zalewska-Pi¹tek et al., 2008). E. coli BL21(DE3) is a 8DE3 strain, which carries the gene for T7 polymerase under lacUV5 promoter control. E. coli IH11128 is a strain of clinical origin (isolated from human with pylenophritis) bearing Dr fimbriae (Goluszko et al., 1997). E. coli DR14 is an insertional draC mutant strain, described previously (Goluszko et al., 1997). The E. coli DR14/gspD strain is a gspD mutant (with a gspD gene knockout) of E. coli DR14, described previously (Zalewska-Pi¹tek et al., 2008). The laboratory E. coli strains harboring the dra gene cluster and its mutants with an inactivation in the draE, draD or draC genes grew in either LB or M63 minimal medium supplemented with 0.2% glucose and 1% LB. The clinical E. coli strains were cultivated only in M63 minimal glucose medium. Plasmids pCC90 carrying the dra gene cluster with its promoter region and regulatory genes (draF-draA) upstream of a draB gene deleted, and pCC90D54stop (the DraE-negative mutant), with a mutated draE gene were provided by S. Moseley, University of Washington, Seattle, WA, USA (Carnoy and Moseley, 1997). Plasmids pCC90DraDmut (the DraD-negative mutant), with a mutated draD gene, and pCC90DraCmut (the DraC-negative mutant), with a mutated draC gene were described previously (Zalewska et al., 2005). PVC (polyvinyl chloride) dishes (96-well; Falcon 3911 microtest III flexible assay plates) and glass coverslips were obtained from Becton Dickinson (Franklin Lakes, NJ). 1% crystal violet (CV), a dye which stains attached cells but not PVC was purchased from Merck (Darmstadt, Germany). Growth of biofilm in rich and minimal media. Ten ml cultures of the E. coli strains of interest were grown in LB at 37C° without shaking for 24 h. Bacterial strains were then subcultured (1:100) into 200 µl of fresh LB and minimal medium M63 supplemented with 0.2% glucose and 1% LB (M63 minimal glucose) in 96-well plates. The cells were grown for an additional 24 h at 37°C without shaking. After that time planktonic cells were removed by three-time rinsing with PBS (phosphate-buffered saline) and then the biofilm was stained with 200 µl of 1% (w/v) CV. After washing, the A 531 was determined. Strains presenting a blank corrected mean absorbance value of > 0.1 were considered as positive.

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Biofilm formation of uropathogenic E.coli Dr+ strains

Light microscopy of biofilm. Strains were grown in LB for 24 h at 37°C without shaking. Then the bacterial cultures were subcultured into LB or M63 minimal glucose media. The biofilm assays were performed by using either polystyrene 6-well culture dishes with glass coverslips. Two ml of LB or M63 minimal glucose media were add to each well simultaneously with 40 µl of the bacterial overnight culture. The culture dishes were incubated overnight at 37C° without shaking. After 24 h the wells were washed 3 times with PBS. Biofilm formation was visualized by first fixing the bacteria with 10% (v/v) formalin for 10 min and then staining with 300 µl of 1% CV for 5 min, washing with PBS and air-drying. The cover slides were removed from the wells, mounted on microscopic slides and observed by a light microscopy using a 40 × objective lens and a Olympus BX-60 microscope. Results Biofilm formation under various growth conditions (rich and minimal environments). The ability of uropathogenic E. coli Dr+ and Dr– strains to form biofilms was determined through the growth of bacteria in either LB or M63 minimal glucose media on PVC plates. Bacterial cells attached to the abiotic surfaces were visualized after staining with CV (observation of stained purple cells) (Fig. 1, part A and B). In our investigations we used 8 different E. coli strains of laboratory and clinical origin. The laboratory E. coli BL21(DE3) strain (Fig. 1a) and the same strain transformed with pCC90 encoding the dra cluster without a regulatory region (the DraE and DraD positive strain) (Fig. 1b) were used as a negative and positive control of biofilm formation, respectively. The E. coli BL21(DE3)-pCC90DraDmut containing the dra gene

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cluster with an inactivated draD gene (the DraD negative strain) (Fig. 1c) was employed to reveal the role of fimbrial polymer composed of only the DraE adhesin, without the DraD capping subunit (DraE + and DraD–), in formation of bacterial communities. The E. coli BL21(DE3)-pCC90DraCmut and E. coli BL21(DE3)-pCC90D54stop harboring the dra operon with an inactivated draC or draE gene, respectively (the DraC and DraE negative strains) (Fig. 1d-e) were used to determine whether the DraD protein nonassociated with Dr fimbriae might also be important for the adherence to abiotic surfaces under static growth conditions. The results obtained were compared with the clinical E. coli IH1128 bearing Dr fimbriae (Fig. 1g), its insertional draC mutant DR14 (DraC serves as a polymerization platform of Dr fimbriae) (Fig. 1h) and D14/gspD with the gspD gene disruption (GspD is responsible for the DraD secretion across the outer membrane on the bacterial cell surface) (Fig. 1f). In case of the above laboratory E. coli strains we could observe biofilm formation in both LB (Fig. 1be, part A) and glucose-minimal medium (Fig. 1b-e, part B). On the contrary the clinical strains revealed the effect of bacterial aggregation only in a glucoseminimal medium (Fig. 1g-h) because of the methylation-dependent phase-variation mechanism which controls the expression of the genes encoding urovirulence determinants and allowing the alternation between Dr fimbriae plus (phase ON), and Dr fimbriae minus (phase OFF) states. The biofilm phenotypes were exhibited by E. coli strains harboring the whole dra gene cluster and also its mutants with an inactivation in the draE or draC genes (expressing only DraD protein at the bacterial cell surface) or the draD gene (with surface exposition of Dr fimbriae). Additionally, the moderately visible differences in biofilm formation was shown in case of bacterial strains expressing Dr fimbriae with (Fig. 1b) or without the

Fig. 1. Analysis of growth of biofilm in rich LB (A) and glucose-minimal medium (B). E. coli biofilms were stained with CV and the A531 of each CV sample was determined. Each bar represents the mean ± SEM from three independent experiments: a) E. coli BL21(DE3); b) E. coli BL21(DE3)-pCC90; c) E. coli BL21(DE3)-pCC90DraDmut; d) E. coli BL21(DE3)-pCC90DraCmut; e) E. coli BL21(DE3)-pCC90D54stop; f) E. coli DR14/gspD; g) E. coli IH11128; h) E. coli DR14.

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DraD as a capping fimbrial domain (Fig. 1c) in comparison with the strains exposing only DraD (DraE+/ DraD+, DraE+/DraD– or DraE–/DraD+ E. coli strains, respectively) (Fig. 1d-e, h) on the cell surfaces. Microscopic analysis of biofilm formation. Both laboratory and clinical E. coli strains were analyzed by the light microscopy to confirm the role of Dr fimbriae and DraD protein during biofilm formation. The strains were grown for 24 h in LB or glucose-minimal media (laboratory and clinical strains, respectively) on glass coverslips, stained with CV and observed under the light microscopy. The bacteria forming biofilm were visible as cell clusters (Fig. 2). The effect of bacterial aggregation was not exhibited by the E. coli BL21(DE3) (Fig. 2A) and E. coli DR14/gspD with the draC and gspD gene knockout (Fig. 2F). The E. coli BL21(DE3)-pCC90 and the wild-type E. coli IH11128 strains (Fig. 2B and G) were observed as the dark clusters of cells. The same effect was displayed by the E. coli BL21(DE3)-pCC90DraCmut and E. coli DR14 (the DraC negative strains) (Fig. 2D and H), E. coli BL21(DE3)-pCC90D54stop (the DraE negative strain) (Fig. 2E) or E. coli BL21(DE3)-pCC90DraDmut (the DraD negative strain) (Fig. 2C). Additionally, in case of DraE– and DraD+ (Fig. 2D-E and H) or DraE+ and DraD– E. coli strains (Fig. 2C), the clusters of cells could be visualized as interspersed layers of various density. The microscopic studies revealed that Dr fimbriae are critical for the interaction of E. coli with abiotic surfaces. We also showed the instrumental role of the DraD protein, located alone at the bacterial cell surface or associated with the fimbrial organelles, in the development of biofilm. Discussion UTI are the most common urologic diseases in humans. Uropathogenic E. coli strains are the most frequent etiologic agent of UTI. The strains exhibit various virulence determinants required for the process of colonization of lower or upper urinary tract and persistence of the bacterial infections. Among these urovirulence factors are adhesive subunits, the structural elements, of homo- or heteropolymeric structures exposed at the surface of bacterial cells (Foxman et al., 1995; Arisoy et al., 2006; Yamamoto et al., 2007). The adhesive structures can be also associated with tissue invasion and biofilm development during the course of UTI. Biofilm formation is classified as a pathogenic determinant of uropathogenic E. coli strains which allows bacteria to adhere to any abiotic or biological surfaces (Costerton et al., 1995; Soto et al., 2007). One of the E. coli urovirulence factors is the Dr family of adhesins responsible for lower and upper UTI, especially in young women and children.

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This adhesive family includes DraE, DaaE, AfaE-I, AfaE-III, AfaE-V, NfaE and others (Servin, 2005). DraE adhesin, a subject of our studies, is encoded by the dra gene cluster which also contains draF, draA, draB, draC, draD and draP genes (Nowicki et al., 1989). In clinical E. coli Dr+ strains DraE forms linear polymers capped at the tip by the DraD protein, enabling the bacteria to enter and replicate within epithelial cells (Anderson et al., 2004; Pi¹tek et al., 2005; Zalewska et al., 2005). The internalization of E. coli by eukariotic cells via a zipper-like mechanism is dependent on dynamic microtubules, lipid rafts, and "5$1 integrin. The interactions with the host receptors (the initial step in colonization) and invasion are required for recurrent and chronic UTI (Goluszko et al., 1997; Guignot et al., 2001; Kansau et al., 2004). In this study, we demonstrated for the first time the biofilm development by uropathogenic E. coli strains harboring the dra gene cluster. The formation of bacterial communities by laboratory and clinical E. coli strains was analyzed under rich and minimal growth conditions. It was connected with the expression of virulence determinants often regulated by the various environmental stimuli. Some of the stimuli can suppress the transcription of the virulence genes such as fimbrial genes (Mekalanos, 1992; White-Zigler et al., 2000). In case of clinical E. coli IH11128 and DR14 isolates, containing the whole dra gene cluster with a regulatory region including the draF-draA genes, biofilm formation was not observed in LB medium. The results obtained were in accordance with earlier published investigations which revealed that rich medium, LB, repressed transcription of pap, daa and fan gene clusters encoding type P pili, F1845 fimbriae and K99 fimbriae, respectively (White-Zigler et al., 2000). The analyzed operons were also under the control of phase-variation mechanism, which allows the bacteria a transition between two expression states: phase ON characterized by expression of pili/fimbriae and phase OFF blocking the expression of pili/fimbriae (Bilge et al., 1993; van der Woude et al., 1992). The mentioned mechanism is controlled at the transcription level and is associated with the formation of specific DNA methylation patterns of GATC boxes. The GATC I site is non-methylated in phase ON whereas the GATC II box is non-methylated in phase OFF (van der Woude et al., 1992; van der Woude and Low, 1992). The exactly role of repression effect of LB medium is not determined. However, it is assumed that the bacterial growth in this medium increases the rate at which the cells expressing fimbriae alternate to phase OFF (White-Ziegler et al., 2000). The dra gene cluster of E. coli Dr+ also contains two GATC boxes (spaced by 102 bp) between draF and draA genes which are conserved among pili/fimbrial operons of

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Biofilm formation of uropathogenic E.coli Dr+ strains

Fig. 2. Biofilm development on glass surfaces examined by a light microscopy. E. coli biofilms were formed through growth in LB or glucose-minimal medium (laboratory and clinical E. coli strains harboring a dra gene cluster and its mutants, respectively) and stained with CV. (A) E. coli BL21(DE3); (B) E. coli BL21(DE3)-pCC90; (C) E. coli BL21(DE3)-pCC90DraDmut; (D) E. coli BL21(DE3)-pCC90DraCmut; (E) E. coli BL21(DE3)-pCC90D54stop; (F) E. coli DR14/gspD; (G) E. coli IH11128; (H) E. coli DR14.

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uropathogenic E. coli strains (van der Woude et al., 1992). A high degree of homology between the regulatory proteins and the presence of the highly homologues regions with the GATC sites of the studied gene clusters can suggest the same mechanism of transcription regulation of the dra operon. Because the development of biofilm is strictly associated with attachment of bacterial cells to the abiotic or biological surfaces (using surface exposed adhesive organelles) we could not exhibit the formation of bacterial communities by clinical E. coli IH11128 strain not expressing Dr fimbriae in rich LB medium (transition from phase ON to phase OFF and abrogation of the dra transcription). On the contrary the laboratory E. coli BL21(DE3) strains transformed with the plasmids encoding the dra gene cluster, without the draFdraA regulatory region, and its mutants with an inactivated draE or draC (the DraE negative and DraD positive strains) and draD (the DraD negative and DraE positive strain) showed biofilm formation in either LB and glucose-minimal medium after staining the cells with CV. Furthermore, both DraE and DraD were shown to be required for adherence to abiotic surfaces under static growth conditions. The laboratory E. coli strain deficient in the production of the DraD protein was able to form a biofilm but the strains deficient in the surface secretion or production of the DraE showed a greater capacity for this process. The observed differences in development of biofilm were not so clearly visible for clinical E. coli IH1128 and DR14 which can suggest a role of another factor besides the DraE and DraD stimulating the bacterial aggregations within urinary tract. The results obtained were confirmed by the light microscopy of laboratory and clinical E. coli strains harboring the dra gene cluster. Cells attached to the abiotic surfaces were observed as clusters containing cells which were in physical contact, sometimes forming interspersed layers. On a basis of the investigations performed we can hypothesize that Dr fimbriae capped with the DraD can be critical for the biofilm development as a resistant barrier against hydrodynamic environment of urinary tract. However, the role of DraD in this process is also very significant. Earlier examinations showed that the bacteria exposing at the surface Dr fimbriae with or without the DraD as a tip subunit were able to stimulate a strict adhesion to Hela cells surrounding their whole surfaces (Zalewska-Pi¹tek et al., 2008). On the contrary the surface expression of DraD without Dr fimbriae induced aggregation of bacterial cells often adjacent in certain parts of HeLa cells which can suggest the effect of compensation of lack of Dr fimbriae at the cell surface. The same results were obtained by analysis of polystyrene beads coated with DraD or Dr fimbriae (Zalewska-Pi¹tek et al., 2008). Therefore, it is possible to assume that

the adhesion of E. coli Dr+ strains to host receptors located at the surface of epithelial cells of upper urinary tract mediates the invasion resulting in the formation of bacterial communities by intracellular interactions with the participation of DraD protein. Further studies are needed to determine the exactly role of Dr fimbriae and DraD of uropathogenic E. coli strains in biofilm development. Acknowledgement This work was supported by the Ministry of Science and Higher Education, grant N401 156 32/3040 to B.Z.-P.

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Polish Journal of Microbiology 2009, Vol. 58, No 3, 231–236 ORIGINAL PAPER

The First Detection of Babesia EU1 and Babesia canis canis in Ixodes ricinus Ticks (Acari, Ixodidae) Collected in Urban and Rural Areas in Northern Poland STELLA CIENIUCH*, JOANNA STAÑCZAK and ANNA RUCZAJ

Department of Tropical Parasitology, Interfaculty Institute of Maritime and Tropical Medicine, Medical University of Gdañsk, Gdynia, Poland Received 5 July 2009, revised 13 July 2009, accepted 14 July 2009 Abstract Ixodes ricinus, the most commonly observed tick species in Poland, is a known vector of such pathogenic microorganisms as TBE viruses, Borrelia burgdorferi sensu lato, Anaplasma phagocytophilum, Rickettsia helvetica, Babesia divergens and B. microti in our country. Our study aimed to find out whether this tick can also transmit other babesiae of medical and veterinary importance. DNA extracts of 1392 ticks (314 nymphs, 552 male and 526 female ticks) collected in urban and rural areas in the Pomerania province (northern Poland), were examined by nested PCR for the detection of Babesia spp., using outer primers: 5-22F and 1661R, and inner primers: 455-479F and 793-772R, targeting specific fragment of 18S rRNA gene. Overall, at least 1.6% ticks were found to be infected with babesial parasites. In the case of nymphs, the minimal prevalence was 0.6%, and it was approx. 3-times lower than in adults (1.9%). Percentages of infected males and females were comparable (2.0% vs. 1.7%). Sequences of 15/22 PCR-derived fragments of 18S rRNA gene demonstrated 100% similarities with the sequence of Babesia EU1 (proposed name B. venatorum) (acc. no. AY046575) (n = 13) and with B. canis canis (acc. no. AY321119) (n = 2), deposited in the GenBank database. The partial 18S rDNA sequences of Babesia EU1 and B. c. canis obtained by us from I. ricinus have been deposited in GenBank, accession nos. GQ325619 and GQ325620, respectively. The results obtained suggest the possible role of I. ricinus as a source of microorganisms, which have been identified as agents of human and canine babesiosis, respectively, in Europe. To our knowledge this is the first report on the occurrence of Babesia EU1 and B. c. canis in I. ricinus in Poland. K e y w o r d s: Babesia canis canis, Babesia EU1, Ixodes ricinus, canine babesiosis, human babesiosis, Poland

Introduction Ixodes ricinus is a widely distributed tick species in Europe, including Poland, where serves as a vector and reservoir of various pathogens causing diseases in animals and humans. It may transmit such emerging infectious diseases like: Lyme borreliosis, human granulocytic anaplasmosis, and babesiosis. The latter is caused by piroplasms of the genus Babesia, ticktransmitted obligatory parasites of mammalian red blood cells. Of them, mainly two species are responsible for human babesiosis. In the North America, the disease is predominantly caused by B. microti, a rodent parasite transmitted by I. scapularis ticks, while in Europe most cases have been attributed to B. divergens, a cattle parasite, transmitted by I. ricinus. The first demonstrated, fatal case of human babesiosis in the world was reported in Europe, in 1957,

in an asplenic man in the former Yugoslavia (Škrabalo and Deanoviè, 1957). Since then, approximately 400 confirmed cases of human babesiosis have been reported in the United States (Kjemtrup and Conrad, 2000). In Europe the disease is considerably less prevalent and for over 50 years just about 40 cases have been noted (Homer et al., 2000), including two human infections with B. microti recently reported in patients from Switzerland and Germany (MeerScherrer et al., 2004, Hildebrandt et al., 2007). However, during the last decade of the XXth century, new species of Babesia and Babesia-like microorganisms have been identified as potential infectious agents. In North America, these pathogens include: B. duncani (Conrad et al., 2006), previously referred to as strains WA1-and CA1 (Persing et al., 1995, Quick et al., 1993, Herwaldt et al., 2004), and the MO1 type parasites (Herwaldt et al., 1996). In Europe, the new

* Corresponding author: S. Cieniuch, Department of Tropical Parasitology, Interfaculty Institute of Maritime and Tropical Medicine, Medical University of Gdañsk, 9B Powstania Styczniowego, 81-519 Gdynia; phone (+48) 58 349 37 41; fax: (+48) 58 622 33 54; e-mail: [email protected]

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Babesia EU1 (proposed name B. venatorum) was isolated for the first time from two asplenic patients in Austria and Italy (Herwaldt et al., 2003), and later reported also in patient from Germany (Häselbarth et al., 2007). Its competent vectors are I. ricinus ticks (Becker et al., 2009) while cervids, primarily roe deer, are considered as potential reservoir hosts (Bonet et al., 2007, Duh et al., 2005). While no cases of human babesiosis have been so far reported in Poland, canine babesiosis is a growing veterinary problem both in our and other European countries. The disease is primarily caused by B. gibsoni and B. canis, which is divided into three subspecies namely B. canis vogeli, B. canis rossi and B. canis canis (Uilenberg et al., 1989). Among them, the latter is responsible for most infections in dogs throughout Europe (Adaszek and Winiarczyk, 2008, Duh et al., 2004, Földvári and Farkas, 2005), where is transmitted mainly by the hard tick Dermacentor reticulatus (Földvári et al., 2007, Rar et al., 2005b, Zahler et al., 2000). In Poland, canine babesiosis is frequently noted in dogs between the Vistula River and the San River basin (north-eastern, eastern and south-eastern areas) (Adaszek and Winiarczyk, 2008, Sobczyk et al., 2005), which is the distribution range of D. reticulatus in our country. The majority of cases noted by us in the Tri-City agglomeration (Gdañsk, Sopot and Gdynia, northern Poland) (data not published) also concern dogs travelling with their owners to the areas where D. reticulatus is common. However, in a few cases, dogs suffering from babesiosis did not leave the Pomerania province, where the meadow ticks do not occur. Thus, they might acquire infection only by I. ricinus bite. As to date there has been no reports that I. ricinus ticks can take part in circulation of B. canis canis in nature, a survey was carried out to detect this and other babesial parasites in ticks from urban and rural recreational areas in northern Poland. Experimental

DNA extraction. An ammonium hydroxide (NH4OH) method was used to extract DNA from crushed nymphs and adult I. ricinus (Rijpkema et al., 1996). All adult ticks were processed separately, while nymphs individually or in pools of 2–3 specimens. The obtained lysates were stored at –20°C until examined. The quality of isolated samples was confirmed by PCR with primers specific to 28S rRNA gene of genus Ixodes. Amplification of DNA of Babesia spp. For Babesia spp., a nested PCR was performed with outer primers: 5-22F and 1661R, and inner primers: 455-479F and 793-772R, targeting specific fragment of 18S rRNA gene. An outer primer pair amplified nearly full-length gene, while an inner primer pair was originally designed to amplify an approximately ~ 340-bp fragment from B. gibsoni (Asian serotype), B. canis vogeli, B. canis rossi and B. canis canis (Birkenheuer et al., 2003). Primary reaction used 2 µl of a tick template in a total volume of 20 µl reaction mixture, while nested amplification used 1 µl of the primary PCR product in a total volume of 20 µl. Dog blood samples positive for B. c. canis, confirmed by the analysis of sequences of the PCR products, and double distilled water were used as positive and negative controls, respectively. The conditions of PCR and nested PCR were as described earlier (Birkenheuer et al., 2003). All PCR reactions were carried out in a GeneAmp® PCR System 9700 (Applied Biosystems 850, Foster City, CA, USA). Obtained PCR products were analyzed after electrophoresis in 2% agarose gel stained with ethidium bromide. Sequencing of PCR products. The PCR products of chosen positive samples were purified using CleanUp purification kit (A&A Biotechnology, Gdynia, Poland) and sequencing reaction were carried out using ABI Prism® Big Dye™ Terminator v.3.1 Cycle Sequencing Kit. Then, the obtained products were sequenced with ABI Prism 310 Genetic Analyser (Applied Biosystem 850, Forster City, CA, USA) according to the manufacturer’s protocol. Sequences were compared with gene sequences deposited in GenBank database using NCBI BLAST network service.

Materials and Methods

Tick sampling. In April – September 2008, ticks were collected by flagging lower vegetation in three different collection sites localised in urban forests of the Tri-City agglomeration (Gdynia – Kolibki, Gdynia – Chwarzno, Gdañsk – Jaœkowa Dolina) and in one site in a wooded, recreational area in the vicinity of the village Sulêczyno (Kaszuby region) (northern Poland). In the laboratory, ticks were identified by species (based on morphological characteristics) and stage of development, killed by rapid immersion in a hot water and preserved in 70% ethanol for further investigations.

Results To detect babesial parasites, a nested PCR was performed with outer primers: 5-22F and 1661R, and inner primers: 455-479F and 793-772R. Although inner primers were originally designed to amplify an approximately ~ 370-bp fragment 18S rRNA gene from B. gibsoni (Asian serotype) and ~ 340 bp of B. canis (Birkenheuer et al., 2003), we proved in our assays that they may also amplify DNA of Babesia EU1. The results of these studies are given in Table I.

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Transmission of babesiosis by Ixodes vicinus in northen Poland Table I Prevalence of infection with Babesia spp. in nymphs and adult Ixodes ricinus ticks collected in urban and rural forested areas in the Pomerania province (northern Poland) in 2008 Area

Site

Gdañsk Jaœkowa Dolina

Urban

Gdynia Kolibki

Gdynia Chwarzno

Urban area – total

Rural

Sulêczyno

Urban and rural area – total

Tick stage

No examined

Adults: females males Nymphs Subtotal Adults: females males Nymphs Subtotal Adults: females males Nymphs Subtotal Adults: females males Nymphs Total Adults: females males Nymphs Subtotal Adults: females males Nymphs

102 41 61 78 180 422 209 213 139 561 356 176 180 56 412 880 426 454 273 1153 198 100 98 41 239 1078 526 552 314 1392

In total, 1392 I. ricinus (314 nymphs, 552 male and 526 female ticks) were collected in urban and rural areas in the Pomerania province, northern Poland. Altogether, 1262 tick lysates, including individual adults (n = 1078) and nymphs (n = 103), as well as 32 pools containing two nymphs (n = 64) and 49 pools consisting three nymphs (n = 147), were examined for the presence of Babesia spp. Of these, specific DNA (Fig. 1) was identified in 22 samples (1.7%), i.e. in 20 adult I. ricinus and in two pools of nymphs. There were no significant differences between the percent of infected males and females (2.0% vs. 1.7%). In case of nymphs, assuming that only one of them in each positive pool was infected, the minimal prevalence was 0.6%, and it was approx. 3-times lower than in adults (1.9%). Overall, at least 1.6% ticks were found to be infected with babesial parasites. In urban forests, Babesia spp. was detected in two sites in the city of Gdynia: Kolibki (2.9%) and Chwarz-

Number / % positive Babesia EU1 B. canis canis Babesia spp. 0/0 0/0 0/0 0/0 0/0 9 / 2.1 5 / 2.4 4 / 1.9 0 / 0.0 9 / 1.6 2 / 0.6 1 / 0.6 1 / 0.6 0 / 0.0 2 / 05 11 / 1.3 6 / 1.4 5 / 1.1 0 / 0.0 11 / 1.0 2 / 1.0 1 / 1.0 1 / 1.0 0 / 0.0 2 / 0.8 13 / 1.2 7 / 1.3 6 / 1.1 0 / 0.0 13 / 0.9

0/0 0/0 0/0 0/0 0/0 2 / 0.5 1 / 0.5 1 / 0.5 0 / 0.0 2 / 0.4 0/0 0/0 0/0 0/0 0/0 2 / 0.2 1 / 0.2 1 / 0.2 0 / 0.0 2 / 0.2 0/0 0/0 0/0 0/0 0/0 2 / 0.2 1 / 0.2 1 / 0.2 0 / 0.0 2 / 0.1

0/0 0/0 0/0 0/0 0/0 4 / 0.9 0 / 0.0 4 / 1.9 1 / 0.7 5 / 0.9 0 / 0.0 0 / 0.0 0 / 0.0 1 / 1.8 1 / 0.2 4 / 0.5 0 / 0.0 4 / 0.9 2 / 0.7 6 / 0.5 1 / 0.5 1 / 1.0 0 / 0.0 0 / 0.0 1 / 0.4 5 / 0.5 1 / 0.2 4 / 0.7 2 / 0.6 7 / 0.5

Total 0/0 0/0 0/0 0/0 0/0 15 / 3.6 6 / 2.9 9 / 4.2 1 / 0.7 16 / 2.9 2 / 0.6 1 / 0.6 1 / 0.6 1 / 1.8 3 / 0.7 17 / 1.9 7 / 1.6 10 / 2.2 2 / 0.7 19 / 1.6 3 / 1.5 2 / 2.0 1 / 1.0 0 / 0.0 3 / 1.3 20 / 1.9 9 / 1.7 11 / 2.0 2 / 0.6 22 / 1.6

no (0.7%), while in the city of Gdañsk, in Jaœkowa Dolina, no infected specimens were reported. In total, 1.6% infection level in urban area was slightly higher than infection rate noted in rural environment, near Sulêczyno village – 1.3%. The highest percentage of infected ticks was observed in April – 2.9%, and then gradually decreased to 1.6% in May and June, 1% in July and 0% in August and September. Fifteen of 22 positive samples were sequenced. Analysis of nucleotide sequences of 340 bp obtained from the 18S rRNA gene revealed that 13 of them showed 100% similarity to Babesia EU1 (acc. no. AY046575), while two others were 100% homologous to B. canis canis (acc. no AY072926, AY321119) deposited in the GenBank database. The GenBank accession numbers for the partial sequences we generated of the 18S rRNA gene for the babesial organisms are as follows: Babesia EU1 – GQ325619, B. canis canis – GQ325620.

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Fig. 1. Agarose gel electrophoresis analysis of nested PCR products obtained with inner primer set targeting 18S rRNA gene of Babesia spp. (455-479F and 793-772R). Lane 1 – pUC19/MspI marker, lane 2 – negative control, lane 3 and 14 – positive controls (Babesia canis canis isolated from dog), lane 4–13 and 15–26-tick samples. Lane 7 and 21 – positive samples, confirming the presence of Babesia sp DNA.

Discussion It has been already documented that in Europe I. ricinus harboured B. divergens and B. microti infections (Blaschitz et al., 2008, Casati et al., 2006, Duh et al., 2001, Foppa et al. 2002, Wielinga et al., 2009). These two pathogens were also detected in ticks in our country (Siñski et al. 2006, Skotarczak and Cichocka, 2001, Stañczak et al., 2004). This is the first study reporting the occurrence of a new species – Babesia EU1 (proposed name B. venatorum) in I. ricinus ticks in Poland. Moreover, we reported also for the first time the detection of B. canis canis, the parasite associated primarily with Dermacentor reticulatus, in the same tick species. Overall, Babesia spp. DNA was detected in 1.6% of the examined I. ricinus. The 13/15 partial sequences of the 18S rRNA gene generated from ticks showed to be identical to the corresponding gene of EU1 (AY046575) isolated from two patients in Austria and Italy (Herwaldt et al., 2003). The two gene sequences showed 100% homology to B. c. canis, isolates from infected dogs from Warsaw area (AY321119) (Sobczyk et al., 2005) and Croatia (AY072926) (Cacciò et al., 2002), and 99.4% identity to B. c. canis sequence (AY 649326) found in D. reticulatus from western Siberia, Russia ( Rar et al., 2005a), due to a GA → AG inversion status at position 150 and 151. According to the presence of inversion and the restriction pattern, Adaszek and Winiarczyk (2008) proposed to classify European B. c. canis isolates in two group: group A – GA variant, cut by HincII restriction enzyme and

3

group B – AG variant, uncut when digested. Babesia c. canis detected by us in I. ricinus belongs to the group A. So far D. reticulatus, which occurs in the east and central region of our country, has been considered as a main vector of canine babesiosis both in Poland and other European countries, where is frequently found on dogs. For instance, 64.6% ticks collected in Warsaw veterinary clinics from dogs presented for veterinary care, were identified as D. reticulatus (Zygner and Wêdrychowicz, 2006). Of them, B. c. canis was detected in 9.5% male (13/137) and 11.9% female (29/244) ticks (Zygner et al., 2008). Moreover, in D. reticulatus (n = 144) originating from dogs from Budapest and other locations in Hungary at least 29.9% samples were positive (Földvári et al., 2007). On the other hand, babesial DNA was detected in 0.3% unfed adults in Germany (Naucke, 2007) and in 3.6% ± 2.0% questing adult D. reticulatus collected in Novosibirsk and Omsk regions in western Siberia, Russia (Rar et al., 2005a). In our study, questing I. ricinus ticks were infected with B. c. canis only in 0.1%. However, even such a low infection level indicates that they can play a role in the circulation of the etiological agent of canine babesiosis in areas where meadow ticks do not occur, including Pomerania province (northern Poland). Moreover, sheep ticks are competent vectors of Babesia EU1 (Bonnet et al., 2007). It has been recovered from I. ricinus from Slovenia (2.2%) (Duh et al., 2005), The Netherlands (0.9%) (Wielinga et al., 2009) and Switzerland (Casati et al., 2006), but there have been no evidence that this species occurs in our country. Thus, we demonstrated, for the first time in Poland the natural infection of I. ricinus ticks with EU1. The calculated overall minimal infection level was 0.9%. Babesia EU1 was detected in ticks collected in rural and urban sites. In both areas a wide range of vertebrate tick host occurred. The presence of roe deer (Capreolus capreolus) is especially important as this species is considered the main vertebrate reservoir of EU1. A survey conducted in Slovenia showed that 21.6% roe deer tested were infected with EU1 (Duh et al., 2005), and this rate was comparable to 23% noted in C. capreolus in France (Bonnet et al., 2007). Roe deer are important hosts both for nymphs and female I. ricinus ticks (Adamska, 2008, Tälleklind and Jaenson, 1997), which may acquire infection during feeding on infected animal. Babesia EU1 is transmitted transtadially and transovarially by ticks, thus they may serve also as reservoirs of pathogen (Bonnet et al., 2007). The prevalence of Babesia EU1 infection in questing ticks collected by us increased approx. 3-fold from nymphal to adult stage, that suggests that adults acquired parasites when feeding as nymphs on infected host, the most probably being roe deer. Babesia spp. were detected in I. ricinus collected both in the rural area and in two localities in the for-

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Transmission of babesiosis by Ixodes vicinus in northen Poland

ests of the city of Gdynia, where the anthropopression was relatively low. However, no infected ticks were found in the third urban site – Jaœkowa Dolina in the city of Gdañsk, characterised by intensive degradation of this area by the human activity. The overall infection rate in ticks from urban and rural areas were comparable (1.6% vs. 1.3%). It seems that diversity of habitats and a wide range of vertebrate tick host in the suburban and urban forests of the TriCity agglomeration create as suitable conditions for development and survival of I. ricinus as forests in rural area. So far it has been demonstrated that ticks occurring there were infected with Borrelia burgdorferi s.l., Anaplasma phagocytophilum and B. microti (2.3%) (Stañczak et al., 2004). Our study showed that these ticks may also harbour Babesia EU1 and B. c. canis. The results of this study confirm the existence of natural foci of human and canine babesiosis in rural (Kaszuby region) and urban forests of Tri-City (the Pomerania province), and indicate a potential risk for people and their dogs living or visiting these areas to contract Babesia EU1 and B. c. canis, respectively. Although, taking into consideration the low percent of infected ticks (~0.9% vs. ~0.1%) the risk of transmission of both pathogens is not significant, they should be added to the list of potentially dangerous microorganisms transmitted by ticks in Poland. Further systematic sampling is needed because knowledge of their prevalence in I. ricinus ticks and distribution is insufficient. Literature Adamska M. 2008. Infestation of game animals from north-western Poland by common tick (Ixodes ricinus) (in Polish)]. Wiad. Parazytol. 54: 31–36. Adaszek £. and S. Winiarczyk. 2008. Molecular characterisation of Babesia canis canis isolates from naturally infected dog in Poland. Vet. Parasitol. 152: 235–241. Becker C.A.M., A. Bouju-Albert, M. Jouglin, A. Chauvin and L. Malandrin. 2009. Natural transmission of zoonotic Babesia spp. by Ixodes ricinus ticks. Emerg. Infect. Dis. 15: 320–322. Birkenheuer A.J., M.G. Levy and E.B. Breitschwerdt. 2003. Development and evaluation of a seminested PCR for detection and differentiation of Babesia gibsoni (Asian Genotype) and B. canis DNA in canine blood samples. J. Clin. Microbiol. 41: 4172–4177. Blaschitz M., M. Narodoslavsky-Gföller, M. Kanzler, G. Stanek and J. Walochnik. 2008. Babesia species occurring in Austrian Ixodes ricinus ticks. Appl. Environ. Microbiol. 74: 4841–4846. Bonnet S., M. Jouglin, M. L’hostis and A. Chauvin. 2007. Babesia sp. EU1 from roe deer and transmission within Ixodes ricinus. Emerg. Infect. Dis. 13: 1208–1210. Cacciò S.M., B. Antunovic, A. Moretti, V. Mangili, A. Marinculic, R.R. Baric, S.B. Slemenda and N.J. Pieniazek. 2002. Molecular characterisation of Babesia canis canis and Babesia canis vogeli from naturally infected European dogs. Vet. Parasitol. 106: 285–292. Casati S., H. Sager, L. Gern and J.C. Piffaretti. 2006. Presence of potentially pathogenic Babesia sp. for human in Ixodes ricinus in Switzerland. Ann. Agric. Environ. Med. 13: 65–70.

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Conrad P.A., A.M. Kjemtrup, R.A. Carreno, J. Thomford, K. Wainwright, M. Eberhard, R. Quick, S.R. Telford 3 rd and B.L. Herwaldt. 2006. Description of Babesia duncani n.sp. (Apixomplexa: Babesiidae) from humans and its differentiation from other piroplasms. Int. J. Parasitol. 36: 779–789. Duh D., M. Petrovec and T. Avšiè-Županc. 2001. Diversity of Babesia infecting European sheep ticks (Ixodes ricinus). J. Clin. Microbiol. 39: 3395–3397. Duh D., M. Petrovec and T. Avšiè-Županc. 2005. Molecular characterization of human pathogen Babesia EU1 in Ixodes ricinus ticks from Slovenia. J. Parasitol. 91: 463–465. Duh D., M. Petrovec M., A. Bidovec and T. Avšiè-Županc. 2005. Cervids as babesiae host in Slovenia. Emerg. Infect. Dis. 11: 1121–1123 Duh D., N. Tozon, M. Petrovec, K. Strašek and T. Avšiè-Županc. 2004. Canine babesiosis in Slovenia: Molecular evidence of Babesia canis canis and Babesia cais vogeli. Vet. Res. 35: 363–368. Foppa, I.M., P.J. Krause, A. Spielman, H. Goethert, L. Gern, B. Brand and S.R. Telford III. 2002. Entomologic and serologic evidence of zoonotic transmission of Babesia microti, eastern Switzerland. Emerg. Infect. Dis. 8: 722–726. Földvári G. and R. Farkas. 2005. Babesia canis canis in dogs from Hungary: Detection by PCR and sequencing. Vet. Parasitol. 127: 221–226. Földvári G., M. Márialigeti, N. Solymosi, Z. Lucács, G. Majoros, J.P. Kósa and R. Farkas. 2007. Hard ticks infesting dogs in Hungary and their infection with Babesia and Borrelia species. Parasitol. Res. 101: 25–34. Häselbarth K., A.M. Tenter, V. Brade, G. Krieger and K.P. Hunfeld. 2007. First case of human babesiosis in Germany – Clinical presentation and molecular characterisation of the pathogen. Int. J. Med. Microbiol. 289, S1: 111–120 Herwaldt B.L., S. Cacciò, F. Gherlinzoni, H. Aspöck, S.B. Slemenda, P. Piccaluga, G. Martinelli, R. Edelhofer, U. Hollenstein, G. Poletti and others. 2003. Molecular characterization of a nonBabesia divergens organism causing zoonotic babesiosis in Europe. Emerg. Infect. Dis. 9: 942–948. Herwaldt B.L., G. de Bruyn, N.J. Pieniazek, M. Homer, K.H. Lofy, S.B. Slemenda, T.R. Fritsche, D.H. Persing and A.P. Limaye. 2004. Babesia divergens-like infection, Washington State. Emerg. Infect. Dis. 10: 622–629. Herwaldt B.L., D.H. Persing, E.A. Précigout, W.L. Goff, D.A. Mathiesen, P.W. Taylor, M.L. Eberhard and A.F. Gorenflot. 1996. A fatal case of babesiosis in Missouri: identification of another piroplasm that infects humans. Ann. Intern. Med. 124: 643–650. Hildebrandt A., K.P. Hunfeld, M. Baier, A. Krumbholz, S. Sachse, T. Lorenzen, M. Kiehntopf, H.J. Fricke and E. Straube. 2007. First confirmed autochthonous case of human Babesia microti infection in Europe. Eur. J. Clin. Microbiol. Infect. Dis. 26: 595–601. Homer M.J., I. Aguilar-Delfin, S.R. Telford III, P.J. Krause and D.H. Persinig. 2000, Babesiosis. Clin. Microbiol. Rev. 13: 451–69. Kjemtrup A.M. and P.A. Conrad. 2000. Human babesiosis: an emerging tick-borne disease. Int. J. Parasitol. 30: 1323–1337. Meer-Scherrer L., M. Adelson, E. Mordechai, B. Lottaz and R. Tilton. 2004. Babesia microti infection in Europe. Curr. Microbiol. 48: 435–437. Naucke T. 2007. Dermacentor reticulatus in Germany and the spread of canine babesiosis. Proceedings of 2nd Canine VectorBorne Diseases (CVBD) Symposium. Mazara del Vallo, Sicily, Italy, 25– 28 April 2007. p. 26–28. Persing D.H., B.L. Herwaldt, C. Glaser, R.S. Lane, J.W. Thomford, D. Mathiesen, P.J. Krause, D.F. Phillip and P.A. Conrad. 1995. Infection with a babesia-like organism in northern California. N. Engl. J. Med. 332: 298–303.

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Quick R.E., B.L. Herwaldt, J.W. Thomford, M.E. Garnett, M.L. Eberhard, M. Wilson, D.H. Spach, J.W. Dickerson, S.R. Telford 3rd, K.R. Steingart and others. 1993. Babesiosis in Washington State: a new species of Babesia? Ann. Intern. Med. 119: 284–290. Rar V.A., N.F. Fomenko, A.K. Dobrotvorsky, N.N. Livanova, S.A. Rudakova, E.G. Fedorov, V.B. Astanin and O.V. Morozova. 2005a. Tickborne pathogen detection, western Siberia, Russia. Emerg. Infect. Dis. 11: 1708–1715. Rar V.A., T.G. Maksimova, L.P. Zakharenko, S.A. Bolykhina, A.K. Dobrotvorsky and O.V. Morozova. 2005b. Babesia DNA detection in canine blood and Dermacentor reticulatus ticks in south-western Siberia, Russia. Vector-Borne Zoonotic Dis. 5: 285–287. Rijpkema S., D. Golubic, M. Molkenboer, N. Verbreek-de Kruif and J. Schellekens. 1996. Identification of four genomic groups of Borrelia burgdorferi sensu lato in Ixodes ricinus ticks in a Lyme borreliosis endemic region of northern Croatia. Exp. Appl. Acarol. 20: 23–30. Siñski E., A. Bajer, R. Welc, A. Pawe³czyk, M. Ogrzewalska and J.M. Behnke. 2006. Babesia microti: Prevalence in wild rodents and Ixodes ricinus ticks from the Mazury Lakes District of north-eastern Poland. Int. J. Med. Microbiol. 296, S1: 137–143. Skotarczak B. and A. Cichocka. 2001. Isolation and amplification by polymerase chain reaction DNA of Babesia microti and Babesia divergens in ticks in Poland. Ann. Agric. Environ. Med. 8: 187–189. Škrabalo Z. and Z. Deanoviè. 1957. Piroplasmosis in man. Report on a case. Docum. Med. Geogr. Trop. (Amsterdam) 9: 11–16.

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Sobczyk A.S., G. Kotomski, P. Górski and H. Wedrychowicz. 2005. Usefulness of touch-down PCR assay for the diagnosis of atypical cases Babesia canis canis infections in dogs. Bull. Vet. Inst. Pulawy. 49: 407–410. Stanczak J., R.M. Gabre, W. Kruminis-Lozowska, M. Racewicz and B. Kubica-Biernat. 2004. Ixodes ricinus as a vector of Borrelia burgdorferi sensu lato, Anaplasma phagocytophilum and Babesia microti in urban and suburban forests. Ann. Agric. Environ. Med. 11: 109–114. Tälleklind L. and T.G.T. Jaenson. 1997. Infestation of mammals by Ixodes ricinus ticks (Acari; Ixoidae) in south-central Sweden. Exp. Appl. Acarol. 21: 755–771. Uilenberg G., F.F. Franssen, N.M. Perie and A.A. Spanjer. 1989. Three groups of Babesia canis distinguished and a proposal for nomenclature. Vet. Q. 11: 33–40. Wielinga P.R., M. Fonville, H. Sprong, C. Gaasenbeck, F. Borgsteede and J.W.B. van der Giessen. 2009. Persistent detection of Babesia EU1 and Babesia microti in Ixodes ricinus in The Netherlands during a 5-year surveillance: 2003–2007. Vector-Borne Zoonotic Dis. 9: 119–122. Zahler M., T. Steffenz, S. Lutz, W.C. Hahnel, H. Rinder and R. Gothe. 2000. Babesia canis und Dermacentor reticulatus in Munchen, ein neuer Naturherd in Deutschland. Tierarztl. Prax. 28: 116–120. Zygner W., S. Jaros and H. Wêdrychowicz. 2008. Prevalence of Babesia canis, Borrelia afzelii, and Anaplasma phagocytophilum infection in hard ticks removed from dogs in Warsaw (central Poland). Vet. Parasitol. 153: 139–142. Zygner W. and H. Wêdrychowicz. 2006. Occurrence of hard ticks in dogs from Warsaw area. Ann. Agric. Environ. Med. 13: 355–359.

Polish Journal of Microbiology 2009, Vol. 58, No 3, 237–245 ORIGINAL PAPER

Genetic Variability of Czech and German RHD Virus Strains BEATA HUKOWSKA-SZEMATOWICZ* , MA£GORZATA PAWLIKOWSKA and WIES£AW DEPTU£A

Department of Microbiology and Immunology, Faculty of Natural Science, University of Szczecin, Poland Received 22 June 2009, revised 15 July 2009, accepted 17 July 2009 Abstract RHD (rabbit haemorrhagic disease) virus (RHDV) is the aetiological factor of the haemorrhagic disease of rabbits and is currently present on all continents. RHDV is a small, envelope-free virus containing genetic material in the form of a 7437-nucleotide long RNA strand. Studies indicate that genetic variability of RDHV strains originating from various parts of the world is approximately 14%, regardless the time and place of isolation. The aim of this study was to evaluate the genetic variability of 6 RHD virus strains from the Czech Republic (CAMPV-561, CAMPV-562, CAMPV-558) and Germany (Frankfurt, Wika, Rossi) based on analysis of fragment of a gene coding a nonstructural p30 protein. The largest variability of nucleotide sequences within the studied fragment was found for the Rossi strain and CAMPV-562 (13.5%) and CAMPV-558 (13.5%), Wika and Frankfurt (12.1%), and CAMPV-561 and Wika (11.2%). Among the Czech strains the largest genetic distance was noted for strains CAMPV-558 and Iowa (0.130/0.140), and in the case of the German strains, for Frankfurt and Iowa (0.123/0.132). A homology tree constructed based on a fragment of a p30 protein-coding gene divided the 14 analysed strains into IV groups of 88% homology. Phylogenetic relationships also divided the tested strains into 4 genetic groups (G1-G4). The larger genetic distance exists between the Czech and German strains and the American ones, and the smallest between them and the European strains. K e y w o r d s: RHD virus, genetic variability, phylogenetic analysis, polyprotein p30

Introduction RHD (rabbit haemorrhagic disease) virus (RHDV) belongs to the Caliciviridae family and constitutes an aetiological factor of rabbit plague (rabbit haemorrhagic disease). The first outbreak of the disease caused by RHDV was reported in 1984 in China (Liu et al., 1984). The first European foci of infection were noted in Italy in 1986, in former Czechoslovakia in 1987, in Germany in 1988, in France and Spain in 1989, and in Poland at the turn of 1987 and 1988 (HukowskaSzematowicz, 2006). At present the disease has swept through all the continents. RHDV is a small, envelope-free virus, 28–40 nm in size, with cubic symmetry. It contains a linear, singlestranded RNA made of 7437 nucleotides (Meyers et al., 1991, Meyers et al., 2000; Wirblich et al., 1996). There are two reading frames in the RHD virus genome: a longer – ORF1 (7034 nucleotides), coding non-structural proteins, structural capsid protein VP60; and a shorter – ORF2 (353 nucleotides), coding a VP12

protein of yet unknown function (Meyers et al., 1991, Meyers et al., 2000, Wirblich et al., 1996). Detailed genetic map of the RHD virus indicates within ORF1 sequences coding six non-structural proteins (p16, p23, p37 [helicase], p30, TCP [protease], polymerase), VPg protein and structural capsid protein VP60 in the following order: NH2-p16-p23-p37(helicase)-p30VPg-TCP(protease)-polymerase-Vp60-COOH (Wirblich et al., 1996). Currently, in the GenBank database (GenBank, 2009) there are 32 RHDV strains with completely sequenced genome (German, Spanish, French, Czech, British, New Zealand, American, Korean, Chinese, Japanese and from Bahrain and Saudi Arabia) and 37 RHDV strains with complete sequence of the structural capsid protein – VP60 (Chinese, Irish, French, Mexican, New Zealand, German, British and Korean). GenBank contains also over 200 registered RHD virus strains in which various fragments of their genome are sequenced, including a fragment coding the non-structural p30 protein. Long-term observations of course of the disease, and – most of all – results of

* Corresponding author: B. Hukowska-Szematowicz, Department of Microbiology and Immunology, Faculty of Natural Science, University of Szczecin, ul. Felczaka 3c, 71-412 Szczecin, Poland; phone (+48) 91 4441592; e-mail: [email protected]

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RHD virus strains’ tests performed using molecular biology methods (Asgari et al., 1999; Bildt et al., 2006; Capucci et al., 1998; Chrobociñska, 2007; Chrobociñska and Mizak, 2007; Farnos et al., 2007; Fitzner et al., 2001, Fitzner, 2005; Fitzner and Kêsy, 2003; Forrester at al., 2006a; Forrester at al., 2006b; Gould et al., 1997; Hukowska-Szematowicz, 2006; Gall Le et al., 1998; Gall-Recule Le et al., 2003; Matiz et al., 2006; Milton et al., 1992; Moss et al., 2002; McIntosh et al., 2007, Nowotny et al., 1997; NiedŸwiedzka-Rystwej et al., 2009; Pawlikowska et al., 2009; Rasschaert et al., 1994; Schirrmeier et al., 1999; Tian et al., 2007) show low level of genetic variability of RHDV of approximately 14%. According to Fitzner and Kêsy (2003), significant evolutionary conservation of the RHD virus genome is associated, among others, with rapid course of the disease and very frequent exchange of generations of susceptible rabbits. Both factors do not facilitate fixation of phenomena leading to the virus variability. The aim of this study was to evaluate the genetic variability of 6 strains of RHD virus from the Czech Republic (CAMPV-561, CAMPV-562, CAMPV-558) and Germany (Frankfurt, Wika, Rossi) based on analysis of fragment of a gene coding the non-structural p30 protein. Experimental Materials and Methods

RHD virus strains. Three Czech strains were used (CAMPV-561, CAMPV-562, CAMPV-558) identified in 1988–1996, prepared in freeze-dried form according to a procedure described previously by Fitzner

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(Fitzner et al., 1996). Moreover, 3 German strains (Rossi, Wika, Frankfurt) from 1996–2002 were used, obtained from the liver of experimentally infected rabbits (Table I). Isolation of viral RNA. Complete RNA of RHD virus was isolated from lyophilisates (in the case of Czech strains) and from 30% liver homogenates in buffered normal saline (in the case of German strains) using the RNA set Total RNA (A&A Biotechnology, Poland) according to the provided protocol. Reverse transcription (RT) reaction – cDNA synthesis. Complementary cDNA strand were obtained on a matrix of viral RNA, using reverse transcriptase enzyme (M-MLV Reverse Transcriptase, Invitrogen, USA). 25 µl of the reaction mixture contained: 1.0 µl of specific antisense starter (C2) at 100 mM concentration (Metabion GmbH, Germany), 1.0 µl of dNTPs nucleotide blend at 25 mM concentration (Promega, USA), 0.5 µl of reverse transcriptase enzyme M-MLV RT (Invitrogen, USA), 2.0 µl of 5-fold concentrated RT-PCR buffer (Invitrogen, USA), 0.5 µl DTT 0,1 M (Invitrogen, USA), 1.0 µl RNase inhibitor RNase OUT (Invitrogen, USA), 14 µl of water for molecular biology (Eppendorf, Germany) and 5.0 µl of RNA of an appropriate RHD virus strain. Before reaction mixture was prepared, RNA of six tested RHDV strains was heated for 5 minutes at 65°C, and then stored on ice until the mixture was prepared. RT-PCR was conducted in a T-gradient Thermocycler (Biometra, Germany) using the following temperature-time profile: 25°C for 10 minutes, 37°C for 60 minutes, 95°C for 5 minutes and 4°C for 1 minute. Resulting cDNA was stored at 2–8°C for further analyses. Starters. Starters suggested by Guittre (Guittre et al., 1996) based on complete sequence of the RHDV-FRG virus genome, developed by Meyers

Table I List of studied RHD virus strains and RHD reference strains obtained from GenBank Strain

Year of identification

Country

GenBank accession number

CAMPV-561 CAMPV-562 CAMPV-558 WIKA FRANKFURT ROSSI IOWA NY-01 BAHRAIN pJG-DD06 BS SD FGR V-351

1996 1992 1988 1996 1996 2002 2000 2001 2000 2006 1989 1989 1988 1987

Czech Republic Czech Republic Czech Republic Germany Germany Germany USA USA Bahrain Germany Italy France Germany Czech Republic

FJ232000 FJ232001 FJ232002 in preparation for submission to GenBank in preparation for submission to GenBank in preparation for submission to GenBank AF 258618 EU 003581 DQ189077 EF363035 X87607 Z29514 M67473 U54983

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Genetic variability of RHD virus strains

(Meyers et al. 1991), and allowing amplification of fragment of the p30 protein-coding gene was used. Using starters: C1 (sense) 5’gttcacatatcgagggcgag 3’ and C2 (antisense) 5’gacagggtcccttgagtacc3’ a 490-bp fragment was amplified. Starter synthesis was performed by Metabion GmbH (Germany). PCR. 50 µl of reaction mixture contained: 2.0 µl of starters (1.0 µl of each C1 and C2) at 10 mM concentration each (Metabion GmbH, Germany), 1.0 µl of dNTPs blend at 10 mM concentration (Promega, USA), 5.0 µl of 10-fold concentrated PCR buffer, 1.0 µl of Taq DNA polymerase, 1.0 µl of 10-fold concentrated buffer for Taq DNA polymerase (Promega, USA), 38.0 µl of water for molecular biology (Eppendorf, Germany) and 2.0 µl of cDNA of an appropriate RHD virus strain (added to the reaction mixture in the end). PCR was conducted in a T-gradient Thermocycler (Biometra, Germany). The following temperature-time profile was used: preliminary denaturation 94°C – 2 minutes, 35 cycles involving denaturation (94°C – 30 seconds), starter affixing (50°C, 53°C or 55°C – depending on a strain – 1 minute), chain elongation (72°C – 2 minutes), final elongation (72°C for 5 minutes) and cooling the reaction mixture down to 4°C. Reaction products were stored at 4°C for further analyses. Electrophoresis of PCR products in agarose gel. Electrophoresis in 1.5% agarose gel (Prona, USA) dyed with ethidium bromide (Fermentas, Lithuania) was performed in order to visualise PCR products. Molecular mass marker GeneRuler 100 and 50 (Fermentas, Lithuania) was used for evaluation of size of products. Electrophoretic separation was conducted in 1.0-fold concentrated TBE buffer, at room temperature, with current voltage of 100V/cm of gel for 45 minutes, using a set for electrophoresis from BioRad (Germany). Storage and interpretation of results was completed using a UV visualisation set (Vilber Lourmat, France). Preparative amplification, purification and preparation of analysed fragments of RHD virus genome for sequencing. Following PCR results visualisation, mass PCR was performed along with electrophoretic separation, using conditions identical to those described above. Preliminary DNA isolation from gel was performed using a Gel OUT set (A&A Biotechnology, Poland) according to the manufacturer’s recommended procedure. Obtained samples were sent for automatic sequencing to Metabion GmbH, Germany and to the DNA Sequencing and Oligonucleotide Synthesis Laboratory at IBB PAN in Warsaw. Molecular phylogenetic analysis of RHD virus sequence. Nucleotide sequences of a p30 protein-coding gene fragment from six tested RHDV strains obtained from sequencing were compared to each other

and to eight homologous sequences obtained from Gene Bank (Table I). Sequence comparative analysis was performed using DNAMAN software, version 5.2.10 (Lynnon BioSoft, Canada). Based on comparison of nucleotide sequences of RHD virus strains matrices of distance and homology were constructed in DNAMAN software, constituting a method of transformation of biological sequences into mathematical data. The analysis used two methods of data transformation: observed divergence and maximum likelihood. The values of the resulting matrices were then graphically transformed into homology trees showing genetic relationships between analysed RHDV strains and expressed in %. Phylogenetic trees presenting hypothetical relationships and evolutionary relations between the analysed strains were constructed using observed divergence and maximum likelihood methods. A bootstrap method was used for evaluation of a program-generated phylogenetic tree. The method reports frequency of a given node occurrence in thousand newly-constructed trees (Baxevanis and Ouellette, 2004). Results PCR yielded amplification of 490-bp-long genome fragment which was subsequently sequenced. Obtained nucleotide sequences of 6 strains (CAMPV-561, CAMPV-562, CAMPV-558 and Wika, Rossi, Frankfurt) were compared to each other (a fragment of 430 nucleotides was compared) and to 8 homologous sequences obtained from GenBank (Table I). Variability of nucleotide sequence in the tested p30 protein-coding gene fragment for the German and Czech strains was 5.7%. Genetic divergence of the tested Czech and German strains was manifested by the occurrence of 81 polymorphic loci, with repeated transitions and few transversions. The highest heterogeneity of nucleotide sequence within the tested fragment was observed for the Rossi strain and CAMPV-562 and CAMPV-558 (13.5%), Wika and Frankfurt (12.1%), CAMPV-561 and Wika (11.2%), and the lowest was for CAMPV-562 and CAMPV-558 (2.1%), Frankfurt and CAMPV-561 (2.8%) and Wika and Rossi (3%). In turn, comparative analysis of three tested Czech strains and three German strains to 8 strains originating from USA, Bahrain, Germany, Italy, France and the Czech Republic showed variability as the level of 5.5%. Nucleotide sequence comparison between 14 RHDV strains indicated 96 polymorphic loci with transitions dominating over transversions; no deletions or insertions were noted. Constructed distance matrix (Table II) (presenting genetic distance for all sequence pairs in the set of analysed strains) showed that among the Czech strains

240

Table II Distance matrix for 14 RHD virus strains performed using the observed divergence method (A) and the maximum likelihood method (B) Strain

BAHRAIN

V-351

BS

SD

NY-01

IOWA

JPGDD06

CAMPV558

CAMPV561

CAMPV562

FRANKFURT

ROSSI

WIKA

0 0.074 0.005 0.063 0.067 0.119 0.128 0.084 0.021 0.067 0.019 0.067 0.133 0.121

– 0 0.074 0.053 0.072 0.107 0.121 0.047 0.084 0.026 0.084 0.044 0.123 0.114

– – 0 0.058 0.063 0.119 0.128 0.079 0.016 0.067 0.014 0.063 0.133 0.121

– – – 0 0.056 0.109 0.123 0.049 0.067 0.037 0.067 0.023 0.128 0.116

– – – – 0 0.102 0.116 0.079 0.072 0.070 0.072 0.056 0.114 0.114

– – – – – 0 0.023 0.116 0.121 0.105 0.121 0.109 0.033 0.021

– – – – – – 0 0.130 0.130 0.119 0.130 0.123 0.033 0.016

– – – – – – – 0 0.088 0.035 0.088 0.040 0.121 0.119

– – – – – – – – 0 0.077 0.021 0.072 0.135 0.123

– – – – – – – – – 0 0.077 0.028 0.123 0.112

– – – – – – – – – – 0 0.072 0.135 0.123

– – – – – – – – – – – 0 0.128 0.121

– – – – – – – – – – – – 0 0.030

– – – – – – – – – – – – – 0

0 0.078 0.005 0.065 0.070 0.127 0.138 0.087 0.021 0.070 0.019 0.070 0.144 0.130

– 0 0.078 0.055 0.075 0.114 0.130 0.048 0.088 0.026 0.088 0.045 0.133 0.122

– – 0 0.060 0.065 0.127 0.138 0.082 0.016 0.070 0.014 0.065 0.144 0.130

– – – 0 0.058 0.116 0.133 0.050 0.070 0.038 0.070 0.023 0.138 0.125

– – – – 0 0.109 0.125 0.082 0.075 0.072 0.075 0.057 0.122 0.122

– – – – – 0 0.023 0.124 0.129 0.111 0.130 0.116 0.033 0.021

– – – – – – 0 0.140 0.140 0.127 0.140 0.132 0.033 0.016

– – – – – – – 0 0.093 0.035 0.093 0.040 0.130 0.127

– – – – – – – – 0 0.080 0.021 0.075 0.146 0.132

– – – – – – – – – 0 0.080 0.028 0.132 0.119

– – – – – – – – – – 0 0.075 0.146 0.132

– – – – – – – – – – – 0 0.138 0.130

– – – – – – – – – – – – 0 0.031

– – – – – – – – – – – – – 0

Hukowska-Szematowicz B. et al.

A. FRG BAHRAIN V-351 BS SD NY-01 IOWA JPG-DD06 CAMPV-558 CAMPV-561 CAMPV-562 FRANKFURT ROSSI WIKA B. FRG BAHRAIN V-351 BS SD NY-01 IOWA JPG-DD06 CAMPV-558 CAMPV-561 CAMPV-562 FRANKFURT ROSSI WIKA

FRG

3

3

Table III Homology matrix for 14 RHD virus strains performed using the observed divergence method Strain

FRG

V-351

BS

SD

NY-01

IOWA

JPGDD06

CAMPV558

CAMPV561

CAMPV562

FRANKFURT

ROSSI

WIKA

FRG

100%

–

–

–

–

–

–

–

–

–

–

–

–

–

BAHRAIN

92.6%

100%

–

–

–

–

–

–

–

–

–

–

–

–

V-351

99.5%

92.6%

100%

–

–

–

–

–

–

–

–

–

–

–

BS

93.7%

94.7%

94.2%

100%

–

–

–

–

–

–

–

–

–

–

SD

93.3%

92.8%

93.7%

94.4%

100%

–

–

–

–

–

–

–

–

–

NY-01

88.1%

89.3%

88.1%

89.1%

89.8%

100%

–

–

–

–

–

–

–

–

IOWA

87.2%

87.9%

87.2%

87.7%

88.4%

97.7%

100%

–

–

–

–

–

–

–

JPG-DD06

91.6%

95.3%

92.1%

95.1%

92.1%

88.4%

87.0%

100%

–

–

–

–

–

–

CAMPV-558

97.9%

91.6%

98.4%

93.3%

92.8%

87.9%

87.0%

91.2%

100%

–

–

–

–

–

CAMPV-561

93.3%

97.4%

93.3%

96.3%

93.0%

89.5%

88.1%

96.5%

92.3%

100%

–

–

–

–

CAMPV-562

98.1%

91.6%

98.6%

93.3%

92.8%

87.9%

87.0%

91.2%

97.9%

92.3%

100%

–

–

–

FRANKFURT

93.3%

95.6%

93.7%

97.7%

94.4%

89.1%

87.7%

96.0%

92.8%

97.2%

92.8%

100%

–

–

ROSSI

86.7%

87.7%

86.7%

87.2%

88.6%

96.7%

96.7%

87.9%

86.5%

87.7%

86.5%

87.2%

100%

–

WIKA

87.9%

88.6%

87.9%

88.4%

88.6%

97.9%

98.4%

88.1%

87.7%

88.8%

87.7%

87.9%

97.0%

100%

Genetic variability of RHD virus strains

BAHRAIN

241

242

Hukowska-Szematowicz B. et al.

3

the largest genetic distance exists between strain pairs CAMPV-558 and Iowa (0.130 according to the observed divergence method/0.140 according to maximum likelihood method), CAMPV-558 and NY-01 (0.121/0.129); between CAMPV-561 and Iowa (0.119/0.127), CAMPV-561 and NY-01 (0.105/0.111) and between CAMPV-562 and Iowa (0.130/0.140). In turn, among German strains the highest distance was noted between strains: Frankfurt and Iowa (0.123/0.132), Frankfurt and NY-01 (0.109/0,116) and clear distance separates Rossi and Wika strains from the majority of the tested strains (0.114/0.119– 0.135/0.146) (Table II), except for NY-01 (0.021– 0.033) and Iowa (0.016–0.033), for which the distance was much smaller. Constructed homology matrix for 14 RHDV strains (Table III) showed over 97% homology between strains, including CAMPV-562 and V-351 (98.6%), CAMPV-558 and V-351 (98.4%), Wika and Iowa (98.4%), CAMPV-562 and FRG (98.1%), CAMPV558 and FRG (97.9%), Wika and NY-01 (97.9%), Frankfurt and BS (97.7%), CAMPV-561 and Bahrain (97.4%), Frankfurt and CAMPV-561 (97.2), Wika and Rossi (97%). Homology tree (Figure 1) constructed based on fragment of a p30 protein-coding gene divided 14 ana-

Fig. 2. Phylogenetic tree for 14 RHD virus strains performed using the observed divergence method (A) and the maximum likelihood method (B).

Fig. 1. Homology tree for 14 RHD strains performed using the observed divergence method.

lysed strains into IV groups (GI-GIV) of 88% homology. First one (GI) (with 98% homology) included two tested Czech strains: CAMPV-562, CAMPV-558 and FRG and V-351. The second group (GII) was formed solely by the French strain SD. The tested Czech strain CAMPV-561 and the German Frankfurt formed the third group (GIII), along with strains: Bahrain, BS and JPG-DD06 (96% homology). The fourth group (GIV) was composed of the tested German strains Wika and Rossi, and NY-01, Iowa (97% homology).

3

Genetic variability of RHD virus strains

Phylogenetic relationships presented in the tree (Figure 2) also divided tested strains into 4 genetic groups (G1-G4), corresponding to the groups in homology tree (Figure 1). The first genetic group (G1) included strains: FRG, V-351, CAMPV-558 identified during the first years (1987–1988) of the plague development in Europe (Germany, Czech Republic), and the strain CAMPV-562 (1992), position of which on the tree may suggest that it comes from the strain CAMPV-558. Group G2 was formed by the French strain SD (1989), which appeared to belong to the G1 group along with the strains identified in 1980s. The third genetic group (G3) gathered strains from 1996–2006, except for the Italian BS strain (1989). Evolutionary relationships in that group indicate that the Bahrain strain (2000) could come from European strains. Group G4 included tested German strains Wika and Rossi (1996) and American: NY-01 (2001) and Iowa (2000), constituting antigenic variants of RHDV. Topology of phylogenetic trees constructed both using observed divergence and maximum likelihood were very similar, and the bootstrap value was higher in case of the highest likelihood method, equal to 50–100, confirming very high reliability of the generated phylogenetic tree. Discussion The variability of nucleotide sequences within the compared sequences of Czech and German strains (5.7%) and variability of tested strains and those obtained from the GenBank (5.5%) noted in this study is similar to the variability noted by other researchers (0–14.0%), analysing sequence variability within the whole RHDV genome, or its fragments (Asgari et al., 1999; Bildt et al., 2006; Capucci et al., 1998; Chrobociñska, 2007; Chrobociñska and Mizak, 2007; Farnos et al., 2007; Fitzner et al., 2001, Fitzner, 2005; Fitzner and Kêsy, 2003; Forrester at al., 2006a; Forrester at al., 2006b; Gould et al., 1997; Hukowska-Szematowicz, 2006; Gall Le et al., 1998; Gall-Recule Le et al.; 2003; Matiz et al., 2006; Milton et al., 1992; Moss et al., 2002; McIntosh et al., 2007, Nowotny et al., 1997; NiedŸwiedzka-Rystwej et al., 2009; Pawlikowska et al., 2009; Rasschaert et al., 1994; Schirrmeier et al., 1999; Tian et al., 2007). The variability of nucleotide sequences within fragment of the p30 protein coding gene for the Czech and German strain pairs observed in this study ranged from 2.1% (CAMPV-562 and CAMPV-558) to 13.5% (Rossi, CAMPV-562 and CAMPV-558) and was higher compared to Polish strains SGM, KGM, PD, LUB, BLA, GSK, ¯D (1–8%) (Fitzner and Kêsy, 2003; Fitzner, 2005), and French strains (8.7%) (Gall Le et al., 1998). Lower variability of nucleotide sequences (0.1–9.0%)

243

was noted within the VP60 capsid protein coding gene (or its fragments) in RHDV strains originating from various parts of the world (Hukowska-Szematowicz, 2006; Pawlikowska et al., 2009). Considering the facts mentioned above it is reasonable to suppose that p30 non-structural protein coding region within the RHD virus genome shows higher variability compared to the VP60 structural protein coding region. Distance matrices presented here (Table II) show a clear genetic distance between strains originating from Europe and the USA. That phenomenon may be explained by the fact that viruses come from various continents and are separated in time. Distance matrices presented here and constructed according to the observed divergence and maximum likelihood methods proved to be reliable method of biological data interpretation. It should be noted that slightly higher matrix values were obtained when the observed maximum likelihood was used, which is consistent with assumption of the method basing on evaluation of phylogenetic distance taking into account mutation level evaluated from sequence comparison and determined by differences in sequences. And the observed divergence method is based on unchanged values obtained directly from sequence comparison by calculation of divergence coefficient for each pair of sequences (Hukowska-Szematowicz, 2006). On the other hand, the homology matrix (Table III) showed that European strains from 1980s tend to be more homologous to each other than to strains originating from other continents. Molecular phylogenetic analysis allowed separation of the tested strains into 4 genetic groups. It is impossible to indicate a criterion based on which the strains formed those genogroups. The results of studies (Bildt et al., 2006; Chrobociñska, 2007; Chrobociñska and Mizak, 2007; Farnos et al., 2007; Fitzner et al., 2001, Fitzner, 2005; Fitzner and Kêsy, 2003; Forrester at al., 2006a; Forrester at al., 2006b; Forrester et al., 2003; Hukowska-Szematowicz, 2006; Gall Le et al., 1998; Gall-Recule Le et al., 2003; Matiz et al., 2006; Moss et al., 2002; McIntosh et al., 2007, Nowotny et al., 1997; NiedŸwiedzka-Rystwej et al., 2009; Pawlikowska et al., 2009) on the phylogenesis of RHDV strains indicate that strains form groups depending on the time of their identification or geographic region. Strain groups having a common identification time, regardless of their geographic origin, are referred to as “clusters”. In the case of G1 and G3 it is possible to indicate the existence of those clusters, and strain composition in the G4 genogroup may be interpreted by both their origin and identification time. Evaluation of the genetic variability of the Czech and German RHD virus strains performed on a fragment of the p30 protein coding gene showed that tested strains are more variable compared to the Polish and French ones. A larger distance exists between

244

Hukowska-Szematowicz B. et al.

Czech and German strains and American ones, and the smallest between them and the European strains. Strain division into groups was associated with their identification time rather than geographic region. In 2008, RHD virus variability via recombination was described (Abrantes et al., 2008; Forrester et al., 2008), which will facilitate analysis of variability mechanisms of other RNA viruses. Acknowledgements This work was supported by grant No N308 03832/3662.

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Polish Journal of Microbiology 2009, Vol. 58, No 3, 247–253 ORIGINAL PAPER

Usefulness of PCR Melting Profile Method for Genotyping Analysis of Klebsiella oxytoca Isolates from Patients of a Single Hospital Unit KAROLINA STOJOWSKA1, STANIS£AW KA£U¯EWSKI2, BEATA KRAWCZYK1* 2 Department

1 Department of Microbiology, Gdañsk University of Technology, Gdañsk, Poland of Bacteriology, National Institute of Public Health – National Institute of Hygiene, Warszawa, Poland

Received 24 June 2009, revised 20 July 2009, accepted 21 July 2009 Abstract The development of rapid and simple typing methods is required in order to identify possible sources of human exposure to opportunistic pathogens. Klebsiella spp. belongs to a group of bacteria that are opportunistic pathogens responsible for an increasing number of multiresistant infections in hospitals. Recently, we showed the high genetic diversity of K. oxytoca using a large collection of strains isolated from the patients of several hospitals in Poland over a 50-year period. Our results showed that the internal transcribed spacer polymerase chain reaction method (ITS-PCR) is useful for the phylogenetic delineation of genetic groups in K. oxytoca and the high discriminatory power of the PCR melting profiles (PCR MP) method can be useful for epidemiological studies of K. oxytoca. In the present study the usefulness of PCR MP was tested on two sets of strains isolated from a single unit over a short period of time. The results revealed that PCR MP has a high discriminatory power and can be useful for epidemiological studies of closely related strains of K. oxytoca isolated from a single unit over a short period of time to identify the source, reservoirs and the tract of infection spread. The advantage of PCR MP for the above application was shown by using the procedure at increasing denaturation temperature during PCR to confirm genotyping results. Considering this feature and the high discriminatory power of PCR MP, as shown in this report for determination of the genetic similarities of consecutive K. oxytoca strains, we propose that PCR MP is one of the best techniques for short-term epidemiology analysis. K e y w o r d s: Klebsiella oxytoca, ITS-PCR, PCR MP, genotyping

Introduction Klebsiella spp. embraces opportunistic pathogens that cause a wide spectrum of severe diseases such as septicaemia, pneumonia, urinary tract infection, and soft tissue infection. Nosocomial Klebsiella infections are caused mainly by Klebsiella pneumoniae. Since the early 1980s, isolates of Klebsiella oxytoca have been recognized as clinically significant and an indication for therapy (Livermore et al., 1995). From year to year the number of infections caused by K. oxytoca has increased. Most reports regarding K. oxytoca have included epidemic cases that occurred in an outbreak (Garcia de la Torre et al., 1985; Hansen et al., 1988; Yinnon et al., 1996; Watanakunakorn and Jura, 1991; Korvick et al., 1992; Morgan et al., 1984; Ransjo et al., 1992). It is known that K. pneumoniae and K. oxytoca exhibit a high degree of genetic heterogeneity, as was demonstrated by capsular typing (Ørskov et al., 1984),

O-antigen variation (Mizuta et al., 1983), biotyping (Rennie and Durcan, 1974), protein electrophoretic profiling (Ferragut et al., 1989), multilocus enzyme electrophoresis (Combe et al., 1994), ribotyping (Bingen et al., 1993), randomly amplified polymorphic DNA (RAPD) analysis (Brisse and Verhoef, 2001), pulsedfield gel electrophoresis (Toldos et al., 1997) and diversity of $-lactamase genes (Fournier et al., 1996). Based on the sequence diversity of the K. oxytoca chromosomal $-lactamase and house-keeping genes, finally, six phylogenetic groups of K. oxytoca were determined (Fevre et al., 2005). In our previous study the diversity of K. oxytoca strains throughout over a 50-year period using internal transcribed spacer polymerase chain reaction (ITS-PCR) and PCR melting profiles (PCR MP) genotyping methods on a large collection of strains isolated from patients of several hospitals in Poland was analyzed retrospectively (Stojowska et al., 2009). Based on ITS-PCR method

* Corresponding author: B. Krawczyk, Gdañsk University of Technology, Department of Microbiology, Narutowicza 11/12, 80-233 Gdañsk, Poland; phone/fax: (+48) 58 3472383; e-mail: [email protected]

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six phylogenetic groups of K. oxytoca was also determined. Typing by PCR MP method showed higher level of genetic diversity. However, all K. oxytoca strains were also divided into six distinct branches. We found that the ITS-PCR and PCR MP methods are useful for phylogenetic delineation of genetic groups in K. oxytoca. The PCR MP method allows specific gradual amplification of genomic DNA in terms of thermal stability starting from less stable DNA fragments amplified at lower Td values to more stable ones amplified at higher Td values. Low Td during LM PCR leads to limited and specific amplification of a small number of less stable DNA fragments. The electrophoretic patterns of DNA fragments obtained after such amplifications are characteristic for the bacterial strain taken for DNA isolation. Using PCR MP we also have the possibility to increase the number of amplified DNA restriction fragments by increasing denaturation temperature during PCR. A steady increase in the number of amplified DNA fragments is dependent on denaturation temperature increase. The objective of this study was to show the use of the PCR MP technique for routine epidemiological study of K. oxytoca strains isolated in a short period of time in individual hospital unit. Experimental Materials and Methods

Bacterial strains. The isolates included in the study were sent from the State Institute of Hygiene collection (Poland). The strain collection comprised 14 strains of K. oxytoca isolated during 15 years (1992–2007) from patients of Neonatal Intensive Care Unit of a hospital in Warsaw (set A of strains) and 14 strains isolated in 1999 from patients of Neonatal Intensive Care Unit of a hospital in Bydgoszcz (set B of strains). Identification tests. All bacterial isolates were identified as K. oxytoca in the Department of Bacteriology at the National Institute of Public Health (Poland) by 40 non commercial classical biochemical tests. The final biochemical identification was carried out as described in Bergey’s Manual of Determinative Bacteriology (Holt et al., 1994). The methodological details of identification based on dulcitol and melezitose fermentation and sodium-potassium tartrate degradation were described previously (Ka³u¿ewski, 1967). DNA isolation. DNA isolations (from a single colony on a Columbia sheep blood agar plate) were carried out with the DNA Genomic Mini kit (A&A Biotechnology, Poland) according to the manufacture’s procedure with minor modifications. The DNA con-

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centration ranged between 100 and 200 ng/µl (measured by NanoDrop 1000 spectrophotometer, Thermo Scientific). Genotyping methods. ITS-PCR was performed according to the previously method described (Jensen et al., 1993) with slight modifications. The primers (G1: 5’-GAAGTCGTAACAAGG-3’ and L1: 5’-CAA GGCATCCACCGT-3’) were designed based on the sequences complementary to the conserved regions of the 16S and 23S rRNA genes of various bacterial species. In a Biometra T-gradient thermal cycler, PCRs were performed as follows: 25 cycles of denaturation at 95°C for 1 min, annealing at 55°C for 1 min and elongation at 72°C for 1 min. Prior to cycling, 5 min denaturation step at 95°C was included. After the last cycle, samples were incubated for 5 min at 72°C. The amplification products were submitted to 4% polyacrylamide gels electrophoresis in TBE buffer. The gels were stained with ethidium bromide, visualized under an ultraviolet transilluminator and photographed using a Versa Doc Imaging System version 1000 (BioRad). PCR MP was carried out according to the method described for E. coli isolates (Krawczyk et al., 2006) with slight modifications. Digestion reactions were performed under uniform conditions using approximately 50 ng of DNA sample and 5 U of the HindIII endonuclease (Fermentas, Lithuania). Following a 30 min incubation at 37°C, ligation mix comprising 2 µl of two oligonucleotides forming an adaptor (POW, 5’CTCACTCTCACCAACGTCGAC-3’; HINDLIG, 5’AGCTGTCGACGTTGG-3’; 20 pmol each), 2.5 µl of ligation buffer (10 × concentrated; Epicentre, Madison, WI), 0.5 µl of 25 mM ATP (Epicentre), and 5U of T4 DNA ligase (Epicentre, Madison, WI) was added and the samples were incubated for 1 h at room temperature. Next, the mixture was heated in a thermoblock for 10 min at 70°C and cooled for 10 min at room temperature. The PCR was carried out in a 50 µl reaction mixture containing 1 µl ligation solution, 5 µl 10×PCR buffer (100 mM Tris-HCl, pH 8.5, 500 mM KCl, 1% Triton X-100), 2 µl 50 mM MgCl2, 5 µl of a deoxynucleoside triphosphate mixture (concentration of each deoxynucleoside triphosphate, 2.5 mM), 2 U of Pwo polymerase (DNA Gdañsk II, Poland), and 50 pM of PowaAGCTT primer (5’-CTCACTCTCA CCAACGTCGACAGCTT-3’). In a Biometra T-gradient thermal cycler, PCRs were performed as follows: 7 min at 72°C to release unligated oligonucleotides HINDLIG and to fill in the single-stranded ends and create amplicons, followed by initial denaturation at 86°C for 90 s and 22 cycles of denaturation at 86°C for 1 min, and annealing and elongation at 72°C for 2 min. After the last cycle, samples were incubated for 5 min at 72°C. The denaturation temperature was calculated during the optimization experiments for several K. oxytoca isolates using a gradient

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thermal cycler (Biometra Tgradient Engine) with a gradient range from 85 to 89°C for the denaturation step as described above. The amplification products were submitted to 6% polyacrylamide gels electrophoresis in TBE buffer. The gels were stained with ethidium bromide, visualized under an ultraviolet transilluminator and photographed using a Versa Doc Imaging System version 1000 (BioRad). Fingerprint analysis and dendrogram constructions. The patterns obtained from the electropherograms were converted and analyzed using the Quanty One software, version 4.3.1 (Bio-Rad, USA). For ITS-PCR and PCR MP, the band positions in each gel were normalized using the M 100–1000 DNA ladder (DNA Gdansk, Poland). Band matching and isolate similarity was accomplished using Dice band-based coefficient of similarity, which provides the most accurate similarity results when compared with visual inspection of the fingerprint patterns (Carrico et al., 2005). A dendrogram was constructed using the unweighted pair-group method with arithmetic averages (UPGMA), which employs a sequential clustering algorithm in which the relationships are identified in order of similarity and the dendrogram was built in a stepwise manner (Carrico et al., 2005). The cut-off values for genotype definition were established as 95% and 90% for ITS-PCR and PCR MP, respectively. Results Genotyping of K. oxytoca strains isolated from a single clinical unit over 15-year period. ITS-PCR method was used to study genetic relationship between K. oxytoca strains, regarding to genetic diversity of K. oxytoca isolated in Poland over a 50-year period (Stojowska et al., 2009). Five different amplification profiles at 40% of similarity level were identified by ITS-PCR for 14 clinical K. oxytoca strains (set A) isolated from a single clinical unit (Fig. 1; Table I). Clustering analysis of electrophoretic patterns groups the K. oxytoca strains into two major clusters, KoX1 and KoY1 (names of genotype groups and genotypes as in previous study, Stojowska et al., 2009), suggest that the examined strains represent at least two different lineages. Low genetic diversity of K. oxytoca strains within groups (over than 75% of similarity) shows that these strains were probably closely related. One genotype group, KoX1, was markedly predominant, as this was represented by 12 clinical isolates (86%). The strains from KoX1 were classified as five different genotypes. One of them, KoX1-1, was predominant and was represented by 8 strains (57%). Two strains, which were classified as a group KoY1, show the same ITS-PCR amplification pattern (genotype KoY1-5). Strains of

K. oxytoca which were classified to KoX1-1 genotype were identified in the whole period of time examined (from 1992 to 2007). The KoY1-1 genotype was identified twice only in 1995. Genotyping by PCR MP showed high genetic diversity of K. oxytoca strains tested. As shown in Fig. 1 and Table I 13 different genotypes were distinguished, which were divided into four genotype groups: KoA (8 strains), KoC (2 strains), KoD (2 strains), and KoF (2 strains) at the cut-off level only 15% (names of genotype groups and genotypes as in previous study, Stojowska et al., 2009). The absence of a predominant strain and high genetic diversity suggest that none of those strains is endemic in hospital. Table I Clinical data of colonized/infected patients in 15 years of study period. Names of ITS-PCR and PCR MP genotypes are related to our previous study (Stojowska et al., 2009). No

Date

Isolate source

1 2 3 4 5 6 7 8 9 10 11 12 13 14

1992 1993 1995 1995 1996 1997 1999 1999 2000 2002 2003 2005 2007 2007

stool stool stool stool stool stool stool stool throat nose swabs stool stool stool stool

PCR MP genotype group

ITS-PCR genotype

KoX1-1 KoX1-1 KoY1-5 KoY1-5 KoX1-1 KoX1-11 KoX1-5 KoX1-1 KoX1-1 KoX1-1 KoX1-5 KoX1-1 KoX1-8 KoX1-1

KoA-1 KoA-2 KoF-1 KoF-1 KoA-3 KoD-1 KoC-2 KoA-4 KoA-5 KoA-6 KoC-2 KoA-7 KoD-2 KoA-8

Table II Clinical data of colonized/infected patients in 1 year of study period. Names of ITS-PCR and PCR MP genotypes are related to our previous study (Stojowska et al., 2009). No

Patient

1 2 3 4 5 6 7 8 9 10 11 12 13 14

P1 P2 P3 P3 P4 P4 P4 P4 P4 P5 P6 P7 P8 Hospital environment

Isolate source stool stool throat stool throat urine pus throat stool urine urine urine urine swab

ITS-PCR genotype

PCR MP genotype

KoX1-1 KoX1-1 KoX1-1 KoX1-1 KoX1-1 KoX1-1 KoX1-1 KoX1-1 KoX1-1 KoX1-3 KoX1-3 KoX1-3 KoX1-3 KoX1-3

KoA-1 KoA-1 KoA-1 KoA-1 KoA-1 KoA-1 KoA-1 KoA-1 KoA-1 KoC-1 KoC-1 KoC-1 KoC-1 KoC-1

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Fig. 1. Panel A: ITS – PCR (a) and PCR MP (b) electrophoresis patterns of K. oxytoca strains isolated from single clinical unit in Poland over a 15-year period. M – molecular size marker (100–1000 bp); from 1 to 14 – numbers of K. oxytoca strains. Panel B: Dendrogram of K. oxytoca strains based on ITS – PCR (a) and PCR MP (b) methods generated with Dice Coefficient (DC) and the UPGMA clustering method.

We found complete agreement between the grouping as defined by ITS-PCR and PCR MP. All strains from group KoA, KoC and KoD (typed by PCR MP) belong to KoX-1 group (typed by ITS-PCR) and two strains of KoF group belong to KoY group. Genotyping of K. oxytoca strains isolated from a single clinical unit over 1-year period. The typing of K. oxytoca isolated from one single clinical unit revealed small genetic diversity among 14 strains (set B) isolated from single clinical unit over a 1-year period (Fig. 2). Typing by both ITS-PCR and PCR MP grouped all strains into two clusters (Table II).

Each cluster grouped strains at 100% of similarity level (all strains in one cluster showed the same amplification pattern – the same genotype). Genotypes KoX1-1 and KoX1-3, distinguished by ITS-PCR showed 80% of similarity level and belonged to the same sub-group (KoX1). Genotypes KoA1-1 and KoC1-1 identified by PCR MP were not closely related (15% of similarity) and belonged to two different genotype groups. Clustering analysis based on ITS-PCR and PCR MP results indicates the presence of clonally dependent strains, which can be responsible for nosocomial infections. The existence of two

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Fig. 2. ITS – PCR (a) and PCR MP (b) electrophoresis pattern of K. oxytoca strains isolated from single clinical unit in Poland. M – molecular size marker (100–1000 bp); from 1 to 14 – numbers of K. oxytoca strains.

Fig. 3. PCR MP fingerprint at increasing denaturation temperatures (84°C, 86°C and 88°C) of K. oxytoca strains isolated from different patients and from the hospital environment. M – molecular size marker (100–1000 bp); P5, P6, P7, P8 – numbers of patients.

different groups of genotypes probably indicates two different sources of infection in this clinical unit. Typing by both ITS-PCR and PCR MP methods showed that K. oxytoca strain isolated from the hospital environment (the baby changing table) was classified into the same genotype as four other strains isolated from four patients (P5, P6, P7, P8) (Fig. 2;

Table II). To ensure that those 5 isolates belonging to the same genotype PCR-MP at increasing denaturation temperatures were carried out. A steady increase in the number of amplified DNA fragments, which is dependent on Td increase, was observed and still produced identical profiles for isolates belonging to the same genotype (Fig. 3). Based on this experiment we

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Fig. 4. PCR MP fingerprint at increasing denaturation temperatures (84°C, 86°C and 88°C) of K. oxytoca strains isolated from different sources of one patient. M – molecular size marker (100–1000 bp); from 5 to 9 – numbers of K. oxytoca strains isolated from patient P4.

can conclude that those five isolates of K. oxytoca were probably clonally related and the hospital environment was the source of nosocomial infection. Five K. oxytoca strains isolated from different sources of one patient (P4) were classified into the same genotype (both ITS-PCR and PCR MP) (Fig. 2, lines 5–9; Table II). The presence of clonally related strains in throat, urine, pus, faeces may indicate a spread of bacteria within patient. The PCR MP experiment at increasing denaturation temperatures (Fig. 4) confirmed that isolates suspected of playing a role in the spread of bacteria within patient were clonally related. Discussion Currently, nucleic acid-mediated techniques are more frequently applied and better appreciated than the phenotypically oriented approaches in taxonomy, epidemiology, and evolutionary studies. There are many different methods for determination of genetic variation among microbial isolates. Each of these methods has its technical and nucleic acid target dependent limitations, which should be taken into account when performing molecular typing studies and subsequently calculating strain relatedness. Furthermore, space and time need to be considered when

selecting the optimal molecular markers. Small-scale epidemiological studies require different approaches in comparison to the analysis of a worldwide or nationwide spread of certain microbes. Actually, macrorestriction analysis of genomic DNA followed by pulsed field gel electrophoresis (REA-PFGE) is regarded as “the gold standard” for molecular typing of many microorganisms (Van Belkum et al., 1998). However, problems such as time consuming analyses, cost limitations, electrophoretical resolution or the need for special equipments still remain to be solved to expand the practice of bacterial typing at the strain level. In our study we used two different genotyping methods, ITS-PCR and PCR MP, for determination of genetic variety of K. oxytoca strains isolated from patients of two different clinical units over a 15-year and 1-year period, respectively. Typing by ITS-PCR the clinical set A of K. oxytoca strains was divided into two groups of genotypes: KoX1 and KoY1. This suggests that the examined strains represented at least two different lineages. KoX1 group and KoX1-1 genotype were markedly predominant. This results confirmed our earlier results (Stojowska et al., 2009), where KoX1-1 strains of K. oxytoca were identified with the highest frequency during over 50-year period in Poland (since 1965). Based on our experiments we can conclude that ITS-PCR in comparison to PCR MP has lower level

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of discriminatory power and may be chosen to study phylogenetic delineation of genetic groups in K. oxytoca. The PCR MP method has a high discriminatory power and can be useful for epidemiological studies of closely related strains of K. oxytoca isolated from a single unit over a short period of time. This method is especially dedicated to identify of source, reservoirs, tract of infection spread and to distinguish between epidemic and endemic strains. The advantage of PCR MP for the above application is the possibility of increasing the number of amplified DNA restriction fragments by increasing denaturation temperature during PCR to confirm genotyping results. Literature Bingen E.H., P. Desjardins, G. Arlet, F. Bourgeois, P. Mariani Kurkdjian, N.Y. Lambert-Zechovsky, E. Denamur, A. Philippon and J. Elion. 1993. Molecular epidemiology of plasmid spread among extended broad-spectrum beta-lactamase-producing Klebsiella pneumoniae isolates in a pediatric hospital. J. Clin. Microbiol. 31: 179–184. Brisse S. and J. Verhoef. 2001. Phylogenetic diversity of Klebsiella pneumoniae and Klebsiella oxytoca clinical isolates revealed by randomly amplified polymorphic DNA, gyrA and parC genes sequencing and automated ribotyping. Int. J. Syst. Evol. Microbiol. 51: 915–924. Carrico, J. A., F.R. Pinto, C. Simas, S. Nunes, N.G. Sousa, N. Frazao, H. de Lencastre and J.S. Almeida. 2005. Assessment of bandbased similarity coefficients for automatic type and subtype classification of microbial isolates analyzed by pulsedfield gel electrophoresis. J. Clin. Microbiol. 43: 5483–5490. Combe M.L., J.L. Pons, R. Sesboue and J.P. Martin. 1994. Electrophoretic transfer from polyacrylamide gel to nitrocellulose sheets, a new method to characterize multilocus enzyme genotypes of Klebsiella strains. Appl. Environ. Microbiol. 60: 26–30. Ferragut C., K. Kersters and J. De Ley. 1989. Protein electrophoretic and DNA homology analysis of Klebsiella strains. Syst. Appl. Microbiol. 11: 121–127. Fevre C., M. Jbel, V. Passet, F.X. Weill, P.A.D. Grimont and S. Brisse. 2005. Six groups of the OXY beta-lactamase evolved over millions of years in Klebsiella oxytoca. Antimicrob. Agents. Chemother. 49: 3453–3462. Fournier B., P.H. Roy, P.H. Lagrange and A. Philippon. 1996. Chromosomal beta-lactamase genes of Klebsiella oxytoca are divided into two main groups, blaOXY-1 and blaOXY-2. Antimicrob. Agents. Chemother. 40: 454–459. Garcia de la Torre M., J. Romero-Vivas, J. Martinez-Beltran, A .Guerrero, M. Meseguer and E. Bouza. 1985. Klebsiella bacteremia: an analysis of 100 episodes. Rev. Infect. Dis. 7: 143–150. Hansen D.S., A. Gottschau and H.J. Kolmos. 1998. Epidemiology of Klebsiella bacteraemia: a case control study using Escherichia coli bacteraemia as control. J. Hosp. Infect. 38: 119–132.

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Holt J.G., N.R. Krieg, P.H.A. Sneath, J.T. Staley and S.T. Williams. 1994. Bergey’s Manual of Determinative Bacteriology. The Williams & Wilkins Co., Baltimore, Md. Jensen M.A., J.A. Webster and N. Straus. 1993. Rapid identification of bacteria on the basis of polymerase chain reactionamplified ribosomal DNA spacer polymorphisms. Appl. Environ. Microbiol. 59: 945– 952. Ka³u¿ewski S. 1967. Taxonomic position of indole-positive strains of Klebsiella. Exper. Med. Microbiol. 19: 327–35 Korvick JA, C.S. Bryan, B. Farber, T.R. Beam Jr, L. Schenfeld, R.R. Muder, D. Weinbaum, R. Lumish, D.N. Gerding, M.M. Wagener, et al. 1992. Prospective observational study of Klebsiella bacteremia in 230 patients: outcome for antibiotic combinations versus monotherapy. Antimicrob. Agents. Chemother. 36: 2639–2644 Krawczyk B, A. Samet, J. Leibner, A. Sledzinska and J. Kur. 2006. Evaluation of a PCR Melting Profile Technique for Bacterial Strain Differentiation. J. Clin. Microbiol. 44: 2327–32. Livermore D.M. 1995. b-Lactamases in laboratory and clinical resistance. Clin. Microbiol. Rev 8: 557–584. Mizuta K, M. Ohta, M. Mori, T. Hasegawa, I. Nakashima and N. Kato. 1983. Virulence for mice of Klebsiella strains belonging to the O1 group: relationship to their capsular (K) types. Infect. Immun. 40: 56–61. Morgan M.E., C.A. Hart and R.W. Cooke. 1984. Klebsiella infection in a neonatal intensive care unit: role of bacteriological surveillance. J. Hosp. Infect. 5: 377–385. Ørskov I. and F. Ørskov. 1984. Serotyping of Klebsiella. In: T. Bergan (ed.) Methods in Microbiology, Vol. 14, Academic Press Inc. New York, NY, pp. 143–164. Ransjo U., Z. Good, K. Jalakas, I. Kuhn, I. Siggelkow, B. Aberg and E. Anjou. 1992. An outbreak of Klebsiella oxytoca septicemia associated with the use of invasive blood pressure monitoring equipment. Acta. Anaesthesiol. Scand. 36: 289– 291. Rennie R.P. and I.B.R. Duncan. 1974. Combined biochemical and serological typing of clinical isolates of Klebsiella. Appl. Microbiol. 28: 534–539. Stojowska K., B. Krawczyk, S. Ka³u¿ewski and J. Kur. 2009. Retrospective analysis of the genetic diversity of Klebsiella oxytoca isolated in Poland over a 50-year period. Eur. J. Clin. Microbiol. Infect. Dis. (in press) Toldos C.M., G. Ortiz, M. Camara and M Segovia. 1997. Application of pulsed-field gel electrophoresis in an outbreak of infection due to Klebsiella oxytoca. J. Med. Microbiol. 46: 889–890. Van Belkum A., W. van Leeuwen, M.E. Kaufmann, B. Cookson, F. Forey, J. Etienne, R. Goering, F. Tenover, C. Steward, F. O’Brein, et al. 1998. Assessment of resolution and intercenter reproducibility of results of genotyping Staphylococcus aureus by pulsed-field gel electrophoresis of SmaI macrorestriction fragments: a multicenter multicenter study. J. Clin. Microbiol. 36: 1653–1659. Watanakunakorn C. and J. Jura. 1991. Klebsiella bacteremia: a review of 196 episodes during a decade (1980–1989). Scand. J. Infect. Dis. 23: 399–405. Yinnon A.M., A. Butnaru, D. Raveh, Z. Jerassy and B. Rudensky. 1996. Klebsiella bacteraemia: community versus nosocomial infection. QJM. 89: 933–941.

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Polish Journal of Microbiology 2009, Vol. 58, No 3, 255–260 ORIGINAL PAPER

Genetic Features of Clinical Pseudomonas aeruginosa Strains KATARZYNA WOLSKA1* and PIOTR SZWEDA2 1 Department

of Microbiology, Natural Faculty, University of Podlasie, Siedlce, Poland of Food Chemistry, Technology and Biotechnology, Chemistry Faculty, Gdansk University of Technology, Gdañsk, Poland

2 Department

Received 18 February 2009, revised 20 July 2009, accepted 31 July 2009 Abstract The genetic features of each isolate were determined by enterobacterial repetitive intergenic consensus (ERIC) primer sequences used in PCR and by searching for six virulence genes (alg D, las B, tox A, plc H, plc N, exo S). 49 (79%) of the isolates were distributed in three ERIC PCR subgroups and showed 62% of similarity. The remaining 13 strains generated unique patterns. The first subgroup was primarily composed of isolates from faeces, these strains indicated over 70% relationship with the next subgroup, and primarily contained strains isolated from wounds and bronchial washings and the last subgroup contained strains isolated from wounds and urine. The unique strains were isolated mainly from urine. Statistical analysis indicated that variations in distribution of virulence genes in P. aeruginosa isolates with respect to strain origin and genomic subgroups were not significant. In the group of 49 strains, 100% gave a positive reaction to alg D, las B and plc H genes, 91.8% to tox A and plc N genes and 83.7% to exo S gene. Among the strains that generated unique (ERIC-PCR) patterns, 69.2% gave a positive reaction to alg D gene, 84.6% to las B gene, 76.9% to tox A, plc N and plc H genes, and 46.15% to exo S gene. K e y w o r d s: Pseudomonas aeruginosa, ERIC-PCR, virulence genes

Introduction Pseudomonas aeruginosa is an opportunistic pathogen that is responsible for a wide range of infections. It is a common hospital-acquired pathogen and is responsible for ventilator-associated pneumonia in patients with underlying immune defects (Carratala et al., 1998), as well as wound and catheter-associated infections. Also intestinal colonization by P. aeruginosa could be a reservoir for invasive infections caused by this bacterium and for its dissemination (Zaborina et al., 2006). In healthy individuals, it is a leading cause of contact lens keratitis (Cheng et al., 1999), otitis externa (swimmer’s ear), and hot-tub folliculitis (Speert, 2002). Additionally, P. aeruginosa is the most common pathogen in lung infections affecting cystic fibrosis (CF) patients and is the leading cause of morbidity and mortality in this patient group (Govan and Deretic, 1996). The ability of P. aeruginosa to cause infection is further exacerbated by a high level of resistance to antibiotics, which makes Pseudomonas infections difficult to treat (Hancock and Speert, 2000). P. aeruginosa produces several virulent factors to colonize the cells of its host. Many of these factors are

controlled by regulatory systems involving cell-to-cell signaling (Van Delden and Iglewski, 1998). Among these are: exotoxin A, exoenzyme S, las B elastase, phospholipases C and alginate. Exotoxin A, encoded by the tox A gene, inhibits protein biosynthesis by transferring an ADP-ribosyl moiety to elongation factor 2 of eukaryotic cells. Exoenzyme S, encoded by the exo S gene, is also an ADP-ribosyltransferase that is secreted by a type-III secretion system directly into the cytosol of epithelial cells (Rumbaugh et al., 1999a). Las B elastase, a zinc metalloprotease encoded by the las B gene, attacks eukaryotic proteins such as collagen and elastin, and destroys the structural proteins of the cell (Toder et al., 1994). While phospholipids are hydrolyzed by two phospholipases C encoded by plc H and plc N genes (PLC-H and PLC N, respectively). Alginate, encoded by alg D gene, protects the bacterium from the host’s immune response and from antibiotics (Ostroff et al., 1990; Storey et al., 1997). Genetic methods have been used to explore the genetic diversity of various populations of P. aeruginosa strains. Various methods, such as pulsed-field gel electrophoresis, arbitrarily primed polymerase chain reaction, and ribotyping are currently available for genotyping of P. aeruginosa (Bennekov et al., 1996).

* Corresponding author: K. Wolska, Department of Microbiology, Natural Faculty, University of Podlasie in Siedlce, Prusa 12, 08-110 Siedlce, Poland; e-mail: [email protected]

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On the other hand, the enterobacterial repetitive intergenic consensus (ERIC) sequences are known to be dispersed throughout the prokaryotic genome, and polymerase chain reaction (PCR) studies of eubacteria have revealed that inter-ERIC distances and patterns are highly specific to the individual group of the same bacterial species (Versalovic et al., 1991). In the present study we aimed to determine the genetic features of clinical P. aeruginosa isolates by ERIC – based PCR (ERIC-PCR) and by assessing variations in the prevalence of six virulence genes (alg D, las B, tox A, plc H, plc N, exo S). Experimental Materials and Methods

Source of isolates and identification. A total of 62 strains of P. aeruginosa, were originally isolated from a variety of clinical specimens: faeces (26), urine (11), blood (1), bronchial washings (9), sputum (1), wound swab (9), throat swab (2), ulceration swab (1), swab from skin round tracheotomy (1) and from ear (1). The bacteria were obtained from 62 patients from different wards of the municipal hospital, main hospital and outpatients’ department in Siedlce (Poland), between December 2005 and March 2006. The strains were identified as P. aeruginosa according to biochemical patterns in the Api 20NE system (bio Mérieux). The reference strain NCTC 6749 was also examined. Stock cultures were stored in TSB (tripticase soy broth, Difco) containing 20% glycerol at –80oC. DNA extraction. Isolates were grown in trypticasesoya broth at 37°C for 24 h and DNA was extracted by using the Genomic DNA Pre Plus (A&A Biotechnology, Gdañsk, Poland).

Detection of virulence genes by PCR. The prevalence of virulence genes encoding alginate (alg D), Las B elastase (las B), haemolytic phospholipase C (plc H), non-haemolytic phospholipase C (plc N), exoenzyme S (exo S) and exotoxin A (tox A) was determined by PCR. The genes were amplified with primers selected on the basis of the published PAO1 sequence (Stover et al., 2000). The PCR mixture contained PCR buffer (10 mM Tris/HCl, 50 mM KCl, 1.5 mM MgCl2, pH 8.3), 200 µM of each dNTP (Boehringer), 12.5 pmol of each primer, DMSO at a final concentration of 4%, 1 U Ampli Tag DNA polymerase (Perkin Elmer) and 25 ng DNA template. The DNA was amplified in PTC-100 Programmable Thermo Controller (MJ Research) using the following protocol: 94°C for 3 min, 30 cycles of 94°C for 30 s, 55°C for 1 min and 72°C for 1 min 30 s, and 72°C for 5 min. Each gene was amplified separately. PCR products were separated in a 1% agarose gel for 1 h at 100 V, stained with ethidium bromide and detected by UV transillumination. Amplified genes were identified on the basis of fragment size (Table I). Enterobacterial Repetitive Intergenic Consensus (ERIC-PCR). ERIC primer sequences were used in PCR to detect differences in the number and distribution of these bacterial repetitive sequences in the bacterial genome. ERIC PCR was carried out using the primer sequences ERIC-1R, 5'-CACTTAGGGGTCC TCGAATGTA-3' and ERIC-2, 5'-AAGTAAGTGACT GGGGTGAGCG-3' to amplify the regions in the bacterial genome placed between the ERIC sequences. Dice coefficient was calculated and compared to evaluate similarity among strains through the use of BIOGENE software. The method was described in the previous study (Wolska and Szweda, 2008). Statistical methods. The distribution of virulence genes with respect to genomic groups or strain origin was compared using chi-square (82) test.

Table I Primers used for PCR amplification of virulence factors Gene alg D las B tox A plc H

Primer sequence (5'-3') alg DF – CGTCTGCCGCGAGATCGGCT alg DR – GACCTCGACGGTCTTGCGGA las BF – GGAATGAACGAAGCGTTCTCCGAC las BR – TTGGCGTCGACGAACACCTCG tox AF – CTGCGCGGGTCTATGTGCC tox AR – GATGCTGGACGGGTCGAG plc HF – GCACGTGGTCATCCTGATGC plc HR – TCCGTAGGCGTCGACGTAC

Product (bp) 313 284 270 608

Number of pair G + C 14 13 13 13 13 12 14 12

plc N

plc NF – TCCGTTATCGCAACCAGCCCTACG plc NR – TCGCTGTCGAGCAGGTCGAAC

481

14 13

exo S

exo SF – CGTCGTGTTCAAGCAGATGGTGCTG exo SR – CCGAACCGCTTCACCAGGC

444

14 13

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Results We used PCR to assess the prevalence of six virulence genes. PCR detected alg D, las B, tox A, plc H, plc N and exo S in 58 (93.55%), 60 (96.8%), 55 (88.7%), 59 (95.2%), 55 (88.7%) and 47 (75.8%) isolates, respectively. 40 (64.5%) isolates gave positive PCR results for all studied genes. In this group, 17 (65.4%) isolates were obtained from faeces, 5 (45.45%) isolates were obtained from urine, 7 (77.8%) from wound, 8 (88.9%) from bronchial washings and individual strains were isolated from sputum, skin and blood. The remaining strains (22–35.5%) gave a negative result for one or more genes. All studied genes were not detected in two strains; the first strain (number 2) was isolated from urine and the second one (42) was obtained from throat swab. The strain isolated from faeces (59) gave a negative PCR reaction to alg D, tox A, plc N and exo S genes. The next strain isolated from the same source (53) gave a negative PCR reaction to alg D, plc N and exo S genes, and the last strain of this origin (8) gave a negative PCR reaction to plc N and exo S genes. The other strains gave a negative result to one of the three following genes: exo S, tox A and plc N. Exo S gene was not presented in strains isolated from urine (4 isolates), faeces (1), throat swab (1), wound (2), bronchial washings (1) and ear (1). Tox A gene and plc N were not detected

in 2 and 3 strains isolated from faeces. The variations in distribution of virulence genes in P. aeruginosa isolates with respect to origin were not significant by the use the chi-square analysis (82 = 4.01) (Table II). Nearly all strains, with the exception of 2, 42 and 59 which gave negative reactions to nearly all studied virulence genes, showed 50% of similarity according to Dice coefficient values (Fig. 1). 41 (66.1% of all) isolates revealed a high similarity, over 70%. Dendrogram analysis enabled the division of strains into groups (50–62% of similarity), then subgroups (64–80%), genotypes (80–86%) and lastly subtypes (86–100%). 13 (21%) strains generated unique ERICPCR patterns. 49 (79%) isolates and reference strain of P. aeruginosa creating group 1 showed 62% of similarity. The chi-square analysis indicated that variations in distribution of P. aeruginosa isolates in genomic subgroups with respect to ecological origin were significant (82 = 80.186) (Table III). The most numerous clonal subgroup of 1, which was characterized by nearly 80% of similarity, consisted of 2 genotypes. This subgroup contained 29 isolates, among which, 24 (92.3%) were isolated from faeces and the individual strains in this group were isolated from wound, skin, bronchial washings, urine and throat swab. The mentioned subgroup demonstrated over 70% of similarity with the next clonal subgroup, which included 12 isolates situated in 2 types. Among

Table II Distribution of virulence genes in Pseudomonas aeruginosa isolates with respect to origin Virulence genes

Origin (No) of strains Faeces (26) Urine (11) Wound (9) Bronchial washings (9) Throat (2) Ulceration (1) Ear (1) Blood (1) Skin (1) Sputum (1) Reference (1)

alg D

las B

tox A

plc H

plc N

exo S

24 (92.3%) 10 (90.9%) 9 (100%) 9 (100%) 1 1 1 1 1 1 1

26 (100%) 10 (90.9%) 9 (100%) 9 (100%) 1 1 1 1 1 1 1

23 (88.5%) 9 (81.8%) 9 (100%) 9 (100%) 1 – 1 1 1 1 1

25 (96.1%) 10 (90.9%) 9 (100%) 9 (100%) 1 1 1 1 1 1 1

21 (80.8%) 10 (90.9%) 9 (100%) 9 (100%) 1 1 1 1 1 1 1

22 (84.6%) 6 (54.5%) 7 (77.8%) 8 (88.9%) – 1 – 1 1 1 1

Table III Distribution of the 63 Pseudomonas aeruginosa isolates in genomic subgroups with respect to origin Genomic subgroups (No of strains) 1 (29) 2 (12) 3 (9) The unique patterns (13)

Faecies (n = 26)

Urine (n = 11)

Wound (n = 9)

Bronchial washings (n = 9)

Throat (n = 2)

24 (92.3%) 1 (9.1%) 1 (11.1%) 1 (11.1%) 1 (50%) – 1 (8.3%) 5 (55.5%) 4 (44.4%) – – 3 (27.3%) 3 (33.3%) 1 (11.1%) – 2 (7.7%)

6 (54.5%)

–

3 (33.3%) 1 (50%)

Ulceration Ear Blood Skin Sputum Reference (n = 1) (n = 1) (n = 1) (n = 1) (n = 1) (n = 1) – 1 –

– 1 –

– – –

1 – –

– – 1

– – 1

–

–

1

–

–

–

258 Wolska K. and Szweda P.

Fig. 1. Dendrogram showing genetic relationships between 62 P. aeruginosa isolates.

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The results were obtained by ERIC-PCR analysis using two primers.

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Genetic features of clinical P. aeruginosa strains Table IV Distribution of virulence genes in Pseudomonas aeruginosa isolates with respect to genomic subgroups Virulence genes

Genomic subgroups (No of strains) 1 (29) 2 (12) 3 (9) The unique patterns (13)

alg D

las B

tox A

plc H

plc N

exo S

29 (100%) 12 (100%) 9 (100%) 9 (69.2%)

29 (100%) 12 (100%) 9 (100%) 11 (84.6%)

27 (93.1%) 11 (91.7%) 8 (88.9%) 10 (76.9%)

29 (100%) 12 (100%) 9 (100%) 10 (76.9%)

25 (86.2%) 12 (100%) 9 (100%) 10 (76.9%)

24 (82.7%) 11 (91.7%) 7 (77.8%) 6 (46.15)

these 12 isolates, 5 (55.6%) strains were obtained from wound, 4 (44.4%) from bronchial washings, 1 (9.1%) from urine and 1 from ulceration. Among 41 strains forming subgroups 1 and 2, 29 (70.7%) gave a positive reaction to all six genes. 11 (26.8%) strains gave a positive reaction to five genes and only 1 (2.4%) strain gave a positive reaction to four genes. All isolates of these subgroups gave positive reactions to alg D, las B and plc H genes. The prevalence of tox A gene in isolates of subgroup 1 was similar to isolates of subgroup 2. And the prevalence of exo S gene was higher in the isolates of group 2 than in group 1. The last 9 strains forming group 1, showed 62% of similarity with subgroups 1 and 2. Among which, there were isolates from urine (3 (27.3%) strains), wound (3 (33.3%)), sputum (1), bronchial washings (1 (11.1%)) and the reference strain of P. aeruginosa NCTC 6749. These strains gave a positive reaction to nearly all virulence genes with the exception of two strains isolated from urine. One gave a negative result for exo S gene and the second one gave a negative result for tox A gene. The remaining strains which were collected (13–21%) generated unique (ERIC-PCR) patterns, showing the genetic diversity of these bacteria. They were isolated from urine (6 (54.5%)), bronchial washings (3 (33.3%)), blood (1), faeces (2 (7.7%) and throat swab (1). Among these strains, 6 (46.15%) gave positive reactions to all virulence genes, but 7 (53.85%) gave negative results to one or more (from 3 to 6) virulence gene. The statistical analysis showed that variations in distribution of virulence genes in P. aeruginosa isolates with respect to genomic subgroups were not significant (82 = 4.01) (Table IV). Discussion The relationship between isolates was calculated by numerical analysis of genetic features determined by ERIC PCR. 79% of isolates were distributed in three ERIC PCR subgroups and showed 62% of similarity. The remaining strains generated unique patterns. The first subgroup was primarily composed of isolates from faeces that were strictly related with each other. These strains indicated over 70% relationship with the

next subgroup, and primarily contained strains isolated from wound and bronchial washings. Above all, the last subgroup contained strains isolated from wound and urine. The unique strains were isolated mainly from urine. Virulence genes were detected more frequently in strains isolated from bronchial washings and wound rather than in strains isolated from urine and faeces. However, the statistical analysis indicated that variations in distribution of virulence genes in P. aeruginosa isolates with respect to strain origin and genomic subgroups were not significant. This result is in agreement with the studies of Hamood et al. (1996) and Rumaugh et al. (1999b). They suggest that elastase, phospholipase C, exotoxin A and exoenzyme S are produced by P. aeruginosa isolates from the different sites of infection. The studies also demonstrated that the production of higher levels of elastase and phospholipase C is important in all types of infections. In some bacterial infections, complex systems of coordinate regulation control the expression of virulence genes (Mecalanos, 1992). Storey et al. (1997) found a positive correlation between alg D transcript accumulation and both las B and las A transcript accumulation levels. This indicates a common regulatory element in a cascade of regulators or a common environmental cue that triggers transcription. Numerous authors (Joly et al., 2005; Rietsch et al., 2005; Yahr et al., 1995) have documented the presence of a type III secretion system in P. aeruginosa that appears to play a major role in the virulence of this organism. Endimiani et al. (2006) showed that 100% of P. aeruginosa isolates from bloodstream infections were positive for the following genes: exo T, exo U, las B, plc H, plc N, tox A and nan 2. Exo S, exo Y and nan 1 genes were detected in 78.9%, 73.7% and 57.9% of isolates. Our data demonstrated that the studied genes: alg D, las B, tox A, plc H, plc N and exo S were present in 93.55%, 96.8%, 88.7%, 95.2%, 88.7% and 75.8% P. aeruginosa isolates, respectively. Feltman et al. (2001) and Ferguson et al. (2001) indicated that over 90% of clinical P. aeruginosa strains contain exo T and exo Y genes, but clinical isolates from urine frequently have the exo Y gene present at a relatively lower level-about 70%. Similarly, in our study exo S gene was rarely detected in P. aeruginosa strains isolated from urine. According

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to Lin et al. (2006), exo U gene of P. aeruginosa is the major contributor to cytotoxicity against mammalian cells. Others (Rumbaugh et al., 1999a) revealed that P. aeruginosa isolates recovered from patients suffering from urinary tract infections produced significant levels of exotoxin A. Over 80% of studied isolates from urine contained the tox A gene. In all probability, this toxin can play a significant role as a virulence factor of P. aeruginosa within catheter-associated urinary tract infections (Goldsworthy, 2008). Lanotte et al. (2004) demonstrated that 80% of all pulmonary isolates harboured exo S, indicating that this gene plays a major role in pulmonary infections. In our study also, strains isolated from bronchial washings showed the prevalence of exo S. Exo S gene was frequently associated with multi-drug resistant (MDR) strains which suggests that P. aeruginosa isolates showing MDR trails may be more virulent than susceptible strains (Endimiani et al., 2006). Results of Zaborina et al. (2006) showed that the multi-drug resistant P. aeruginosa clinical strains that disrupted the barrier function of cultured intestinal epithelial cells are characterized by the exo U gene. The presence of virulent strains of P. aeruginosa within the intestinal tract could be the major source of systemic sepsis and death among immuno-compromised patients. The above presented data has been founded in Poland. So far nobody has applied the ERIC-PCR method in discriminating clinical P. aeruginosa strains and evaluated the occurrence of such numerous groups of virulence genes. Literature Bennekov T., H. Colding, B. Ojeniyi, M.W. Bentzon and N. Hoiby. 1996. Comparison of ribotyping of Pseudomonas aeruginosa isolates from cystic fibrosis patients. J. Clin. Microbiol. 34: 202–204. Carratala J., B. Roson, A. Fernandez-Sevilla, F. Alcaide and F. Gudiol. 1998. Bacteremic pneumonia in neutropenic patients with cancer. Arch. Intern. Med. 158: 868–872. Cheng K.H., S.L. Leung, H.W. Hoekman, W.H. Beekhuis, P.G. Mulder, A.J. Geerards and A. Kijlstra. 1999. Incidence of contact-lens-associated microbial keratitis and its related morbidity. Lancet 354: 181–185. Endimiani A., B. Pini, A. Baj, F. Luzzaro and A. Toniolo. 2006. Bloodstream infections due to Pseudomonas aeruginosa: clinical outcome associated with pathogenesis-related genes. Abstracts 16th European Congress Clinical Microbiology and Infectious Disease 2006. Nice, France p. 1164. Feltman H., G. Schlert, S. Khan, M. Jain, L. Peterson and A.R. Hauser. 2001. Prevalence of type III secretion genes in clinical and environmental isolates of Pseudomonas aeruginosa. Microbiology 147: 2659–2669. Ferguson M.W., J.A. Maxwell, T.S. Vincent, J. da Silva and J.C. Olson. 2001. Comparison of the exo S gene and protein expression in soil and clinical isolates of Pseudomonas aeruginosa. Infect. Immun. 69: 2198–2210. Goldsworthy M.J.H. 2008. Gene expression of Pseudomonas aeruginosa and MRSA within a catheter-associated urinary tract infection biofilm model. Bioscience Horizons 1: 28–37.

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Govan J.R. and V. Deretic. 1996. Microbial pathogenesis in cystic fibrosis: mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol. Rev. 60: 539–574. Hamood A.N., J.A. Griswold and C.M. Duhan. 1996. Production of extracellular virulence factors by Pseudomonas aeruginosa isolates obtained from tracheal, urinary tract and wound infections. J. Surg. Research 61: 425–432. Hancock R.E. and D.P. Speert. 2000. Antibiotic resistance in Pseudomonas aeruginosa: mechanisms and impact on treatment. Drug. Resist. Updat. 4: 247–255. Joly B., M. Pierre, S. Auvin, F. Colin, F.B. Guery and M.O. Husson. 2005. Relative expression of Pseudomonas aeruginosa virulence genes analyzed by a real time RT-PCR method during lung infection in rats. FEMS Microbiol. Lett. 243: 271–278. Lanotte P., S. Watt, L. Mereghetti, N. Dartiguelongue, A. Raster-Lari, A. Goudeau and R. Quentin. 2004. Genetic features of Pseudomonas aeruginosa isolates from cystic fibrosis patients compared with those of isolates from other origins. J. Med. Microbiol. 53: 73–81. Lin H., S. Huang, H. Teng, D. Ji, Y. Chen and Y. Chen. 2006. Presence of the exo U gene of Pseudomonas aeruginosa is correlated with cytotoxicity in MDCK cells but not with colonization in BALB/c mice. J. Clin. Microbiol. 44: 4596–4597. Mecalanos J.J. 1992. Environmental signals controlling expression of virulence determinants in bacteria. J. Bacteriol.174: 1–7. Ostroff R.M., A.I. Vasil and M.L. Vasil. 1990. Molecular comparison of nonhemolytic and a hemolytic phospholipase C from Pseudomonas aeruginosa. J. Bacteriol. 172: 5915–5923. Rietsch A., I. Vallet-Gely, S.L. Dove and J.J. Mekalanos. 2005. ExsE, a secreted regulator of type III secretion genes in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. 102: 9006–8011. Rumbaugh K.P., B.S. Abdul, N. Hamood and J.A. Griswold. 1999. Analysis of Pseudomonas aeruginosa clinical isolates for possible variations within the virulence genes exotoxin A and exoenzyme S. J. Surg. Research 82: 95–105. Rumbaugh K.P., J.A. Griswold and A.N. Hamood. 1999. Pseudomonas aeruginosa strains obtained from patients with tracheal, urinary tract and wound infection: variations in virulence factors and virulence genes. J. Hosp. Infect. 43: 211–218. Speert D.P. 2002. Molecular epidemiology of Pseudomonas aeruginosa. Front. Biosci. 7: e354–e361. Storey D.G., E.E. Ujack, J. Mitchell and H.R. Rabin. 1997. Positive correlation of algD transcription to las B and las A transcription by populations of Pseudomonas aeruginosa in the lungs of patients with cystic fibrosis. Infect. Immun. 65: 4061–4067. Stover C.K., X.Q. Pham, A.L. Erwin, S.D. Mizoguchi, P. Warrener, M.J. Hickey, F.S.L. Brinkman, W.O. Hufnagle and D.J. Kowalik. 2000. Complete genome sequence of Pseudomonas aeruginosa PAO1, an opportunistic pathogen. Nature 406: 959–964. Toder D.S., S.J. Ferrell, J.L. Nezezon, L. Rust and B.H. Iglewski. 1994. Las A and las B genes of Pseudomonas aeruginosa: Analysis transcription and gene product activity. Infect. Immun. 62: 1320–1327. Van Delden C. and B.H. Iglewski 1998. Cell-to-cell signaling and Pseudomonas aeruginosa infections. Emerg. Infect. Dis. 4: 551–60. Versalovic J., T. Koeuth and J.R. Lupski. 1991. Distribution of repetive DNA sequences in eubacteria and application to fingerprinting of bacterial genomes. Nucleid Acids Res. 19: 6823–6831. Wolska K. and P. Szweda. 2008. A comparative evaluation of PCR ribotyping and ERIC PCR for determining the diversity of clinical Pseudomonas aeruginosa isolates. Pol. J. Microbiol. 57: 157–163. Yahr T.L., A.K. Hovey, S.M. Kulich and D.W. Frank. 1995. Transcriptional analysis of the Pseudomonas aeruginosa exoenzyme S structural gene. J. Bacteriol. 177: 1169–1178. Zaborina O., J.E. Kohler, Y. Wang, C. Bethel, O. Shevchenko, L. Wu, J.R. Turner and J.C. Alverdy. 2006. Identification of multi-drug resistant Pseudomonas aeruginosa clinical isolates that highly disruptive to the intestinal epithelial barrier. An. Clin. Microbiol. Antimicrob. 5: 1–14.

Polish Journal of Microbiology 2009, Vol. 58, No 3, 261–267 ORIGINAL PAPER

Effect of Ciprofloxacin and N-acetylcysteine on Bacterial Adherence and Biofilm Formation on Ureteral Stent Surfaces MOHAMED A. El-FEKY1, MOSTAFA S. El-REHEWY 1, MONA A. HASSAN1, HASSAN A. ABOLELLA2 REHAB M. ABD El-BAKY3* and GAMAL F. GAD3 1 Microbiology

Department, Faculty of Medicine, Assuit University, Assuit, Egypt Department, Faculty of Medicine, Assuit University, Assuit, Egypt 3 Microbiology Department, Faculty of Pharmacy, El-Minia University, El-Minia, Egypt 2 Urology

Received 2 February 2009, revised 25 July 2009, accepted 27 July 2009 Abstract The aim of this study was to evaluate the effect of ciprofloxacin (CIP), N-acetylcysteine (NAC) alone and in combination on biofilm production and pre-formed mature biofilms on ureteral stent surfaces. Two strains each of Staphylococcus aureus, Staphylococcus epidermidis, Escherichia coli, Klebseilla pneumoniae, Pseudomonas aeruginosa and Proteus vulgaris, recently isolated from patients undergoing ureteral stent removal and shown to be capable of biofilm production, were used in this study. The inhibitory effects of ciprofloxacin, N-acetylcysteine and ciprofloxacin/N-acetylcysteine combination were determined by static adherence assay. Ciprofloxacin (MIC and 2 MIC) and N-acetylcysteine (2 and 4 mg/ml) inhibited biofilm production by ≥ 60% in all tested microorganisms. Disruption of pre-formed biofilms of all tested microorganisms was found to be ≥ 78% in the presence of ciprofloxacin (MIC and 2 MIC) and ≥ 62% in the presence of N-acetylcysteine (2 and 4 mg/ml), compared to controls. Ciprofloxacin/N-acetylcysteine showed the highest inhibitory effect on biofilm production (94–100%) and the highest disruptive effect on the pre-formed biofilms (86–100%) in comparison to controls. N-acetylcysteine was found to increase the therapeutic efficacy of ciprofloxacin by degrading the extracellular polysaccharide matrix of biofilms. These data are statistically significant. The inhibitory effects of ciprofloxacin and N-acetylcysteine on biofilm production were also verified by scanning electron microscope (SEM). In conclusion, Ciprofloxacin/N-acetylcysteine combinations have the highest inhibitory effect on biofilm production and the highest ability to eradicate pre-formed mature biofilms. K e y w o r d s: Biofilm, ciprofloxacin, mature biofilm, N-acetylcysteine

Introduction Ureteral stents are commonly used in urologic practice (Mohan-pillai et al., 1999). Although urinary catheterization is valuable, it may lead to stent obstruction, stent migration, stent encrustation, stone formation and biofilm formation (Lojanapiwat, 2005). Biofilm is a population of cells growing on a surface and enclosed in an exopolysaccharide matrix which may lead to complete blocking to the lumen of the catheter (Desgrandchamps et al., 1997). Biofilm development initiates when bacteria transfer from a planktonic (free) existance to a lifestyle in which microorganisms are firmly attached to abiotic or biotic surfaces. After attachment, exopolysaccharide glycocalyx polymers are produced, a matrix inside which microcolonies increase and coalease to form biofilms

(Costerton et al., 1987). The longer the stents remain in place, the greater the tendency of these microorganisms to develop biofilms (Stickler, 1996). Several approaches have been studied to prevent the formation of biofilms. Some of these depend on coating catheters with silver, antiseptics or by producing radio-opacity by silicone material and some depend on the use of antimicrobial agents or non antimicrobial agents (Flowers et al., 1989). Ciprofloxacin is active against a wide range of urinary tract pathogens and able to inhibit bacterial colonization of urinary catheters in vitro (Reid et al., 1994). N-acetylcysteine (NAC) is a non-antibiotic drug that has antibacterial properties. It is a mucolytic agent that disrupts disulphide bonds in mucus and reduces the viscosity of secretions. NAC is widely used in medical practice via inhalation, oral and intravenous routs and has an excellent safety profile

* Corresponding author: Rehab M. Abd El-Baky, Department of Microbiology, Faculty of Pharmacy, El-Minia University,| El-Minia, Egypt; 56, Adnan El-Malky str., Ard Sultan, El-Minia, Egypt; phone: (+20) 123350610; e-mail: [email protected]

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(Kao et al., 2003). NAC was found to decrease biofilm formation by a variety of bacteria and reduces the production of extracellular polysaccharide matrix while promoting the disruption of mature biofilms (PézerGiraldo et al., 1997). Since the antimicrobial susceptibility of biofilm-associated bacteria is enhanced in disrupted biofilms (El-Azizi et al., 2005), it is conceivable that an antibiofilm/antimicrobial agent combination would be synergistic (Olofsson et al., 2003). In this work, we study the effect of ciprofloxacin, N-acetylcysteine each alone and in combination on biofilm formation on ureteral stents surfaces to select a suitable treatment for biofilm-associated infections to decrease hospitalization time and cost. Experimental Materials and Methods

Strains. Two strains each of Staphylococcus aureus, Staphylococcus epidermidis, Escherichia coli, Klebseilla pneumoniae, Pseudomonas aeruginosa and Proteus vulgaris recently isolated from urine samples and stent segments collected from patients undergoing ureteral removal, identified according to standard procedures (Benson, 2002 and Sheretz et al., 1990) and shown to be capable of biofilm production (Christensen et al., 1985), were employed in this study. Drugs. Preparation of stock solutions of ciprofloxacin (Bayer, Milan, Italy) was performed in accordance with the manufacturer’s instructions. Two concentrations (2 and 4 mg/ml) of NAC (Sedico, Egypt) were evaluated. Determination of minimum inhibitory concentrations (MIC) of ciprofloxacin. Minimum inhibitory concentrations of ciprofloxacin were determined by agar dilution method, according to Clinical Laboratory Standards Institute (CLSI) (2007). Effect of ciprofloxacin, N-acetylcysteine (alone and in combination) on the pre-formed biofilms on stent surfaces and on biofilm production by the tested microorganisms. Seven pieces of JJ uretral stent segments (1 cm length) were incubated in bacterial suspensions that contained 5× 105 – 1× 106 cfu/ml of bacteria in 5 ml of Trypticase soy broth (TSB, BBL, USA) to allow biofilm formation. After incubation at 37°C for 24 h, segments were removed and rinsed three times with phosphate buffer saline (PBS, 7.2) to remove non-adherent bacteria. To test the effect of the tested agents on the preformed mature biofilms, catheter segments were suspended for 24 h at 37°C in one of the following treatment solutions: saline (control), ciprofloxacin (MIC and 2 MIC), N-acetylcysteine (2 and 4 mg/ml) and CIP/NAC (MIC/2 mg/ml and 2 MIC/4 mg/ml).

After incubation, catheter segments were rinsed, placed in 10 ml fresh sterile saline and sonicated for 30 seconds to dislodge the sessile adherent cells. Serial dilutions of the sonicated saline were cultured. The number of sessile bacteria that indicates degree of adherence was determined by the viable count technique (Reid et al., 1994). To test the effect of the tested agents on bacterial adherence to stent surfaces, bacterial cultures in 5 ml TSB were washed, diluted with fresh TSB, standardized to contain 5× 105 – 1× 106 cfu/ml and distributed into test tubes. One of the following solutions: Ciprofloxacin (MIC and 2 MIC), N-acetylcysteine (2 and 4 mg/ml) and CIP/NAC (MIC/2 mg/ml and 2 MIC/4 mg/ml) were added to each tube. Normal saline was added to control tubes. At the same time, one segment of ureteral stents was added to each tube. After 24 hours incubation, the number of viable adherent cells were determined as described before. Each assay was repeated at least three times. Data were expressed as mean cfu ± S.E.M. Scanning Electron Microscopy (SEM). Catheter segments were fixed in 2.5% (vol/vol) glutaraldehyde in Dulbecco PBS (PH 7.2) for 1.5 h, rinsed with PBS, and then dehydrated through an ethanol series. Samples were dried and gold-palladium coated. SEM examinations were made on a JSM-840 SEM (JEOL Ltd., Tokyo, Japan) (Soboh et al., 1995). Statistical analysis. One-Way ANOVA was employed to evaluate any significant difference between the values obtained without the drug (controls) and the values obtained in the presence of different drug concentrations. Differences were done using SPSS, 11 statistical software (SPSS Inc., Chicago, IL). Results A total of 12 biofilm-producing strains (2 strains of each microorganism) were isolated and identified from urine samples and stents segments collected from patients undergoing ureteral stent removal. Minimum inhibitory concentrations of ciprofloxacin. The MICs of ciprofloxacin were 1 and 2 µg/ml for S. aureus, S. epidermidis and P. vulgaris, 2 and 32 µg/ml for E. coli, 1 and 32 µg/ml for P. aeruginosa and 0.5 and 64 µg/ml for K. pneumoniae. Inhibition of biofilm production. The inhibitory effects of ciprofloxacin and N-acetylcysteine were found to be concentration dependent. Ciprofloxacin at MIC inhibited biofilm synthesis by ≥75.6% in S. aureus, S. epidermidis and P. vulgaris, ≥ 67% in P. aeruginosa and E. coli, 60–95.8% in K. pneumoniae (p< 0.05). At 2 MIC, reduction of biofilm synthesis was ≥89.3% in S. aureus, S. epidermidis, E. coli, P. aeruginosa and P. vulgaris and 80–95.8% in

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K. pneumoniae (P< 0.05). N-acetylcysteine showed a significant inhibitory effect (p