International Microbiology

Official journal of the Spanish Society for Microbiology Volume 17 · Number 3 · September 2014

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Ariza-Miguel J, Hernández M, FernándezNatal I, Rodríguez-Lázaro D Molecular epidemiology of methicillinresistant Staphylococcus aureus in a university hospital in northwestern Spain

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Spricigo DA, Cortés P, Moranta D, Barbé J, Bengoechea JA, Llagostera M Significance of tagI and mfd genes in the virulence of non-typeable Haemophilus influenzae

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Padilha IQM, Valenzuela SV, Grisi TCSL, Diaz P, de Araújo DAM, Pastor FIJ A glucuronoxylan-specific xylanase from a new Paenibacillus favisporus strain isolated from tropical soil of Brazil

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INTERNATIONAL MICROBIOLOGY www.im.microbios.org

2014 pp 131-184

Manrique-Ramírez P, Galofré-Milà N, Serrano M, Aragon V Identification of a class B acid phosphatase in Haemophilus parasuis

Number 3

López-Martínez G, Borrull A, Poblet M, Rozès N, Cordero-Otero R Metabolomic characterization of yeast cells after dehydration stress

Chiellini C, Maida I, Emiliani G, Mengoni A, Mocali S, Fabiani A, Biffi S, Maggini V, Gori L,Vannacci A, Gallo E, Firenzuoli F, Fani R Endophytic and rhizospheric bacterial communities isolated from the medicinal plants Echinacea purpurea and Echinacea angustifolia

Volume 17

RESEARCH ARTICLES

International Microbiology

INTERNATIONAL MICROBIOLOGY

Volume 17 · Number 3 · September 2014 · ISSN 1139-6709 · e-ISSN 1618-1905

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17(3) 2014 INDEXED IN

Agricultural and Environmental Biotechnology Abstracts; ASFA/Aquatic Sciences & Fisheries Abstracts; BIOSIS; CAB Abstracts; Chemical Abstracts; SCOPUS; Current Contents®/Agriculture, Biology & Environmental Sciences®; EBSCO; EMBASE/Elsevier Bibliographic Databases; Food Science and Technology Abstracts; ICYT/CINDOC; IBECS/BNCS; ISI Alerting Services®; MEDLINE®/Index Medicus®; Latindex; MedBioWorldTM; SciELO-Spain; Science Citation Index Expanded®/SciSearch® September 2014

Official journal of the Spanish Society for Microbiology

Publication Board

Editorial Board

Coeditors-in-Chief José Berenguer (Madrid), Autonomous University of Madrid Ricardo Guerrero (Barcelona), University of Barcelona

Juan Aguirre, Prince Edward Island University, Canada Ricardo Amils, Autonomous University of Madrid, Madrid, Spain Miguel A. Asensio, University of Extremadura, Caceres, Spain Shimshon Belkin, The Hebrew University of Jerusalem, Jerusalem, Israel Albert Bordons, University Rovira i Virgili, Tarragona, Spain Albert Bosch, University of Barcelona, Barcelona, Spain Javier del Campo, University of British Columbia, Vancouver, Canada Victoriano Campos, Pontificial Catholic University of Valparaíso, Chile Josep Casadesús, University of Sevilla, Sevilla, Spain Rita R. Colwell, Univ. of Maryland & Johns Hopkins Univ., Baltimore, MD, USA Katerina Demnerova, Inst. of Chem. Technology, Prague, Czech Republic Esteban Domingo, CBM, CSIC-UAM, Cantoblanco, Spain Mariano Esteban, Natl. Center for Biotechnol., CSIC, Cantoblanco, Spain Mariano Gacto, University of Murcia, Murcia, Spain Juncal Garmendia, Institute of Agrobiotechnology, Pamplona, Spain Olga Genilloud, Medina Foundation, Granada, Spain Steven D. Goodwin, University of Massachusetts, Amherst, MA, USA Juan C. Gutiérrez, Complutense University of Madrid, Madrid, Spain Johannes F. Imhoff, University of Kiel, Kiel, Germany Juan Imperial, Technical University of Madrid, Madrid, Spain John L. Ingraham, University of California, Davis, CA, USA Juan Iriberri, University of the Basque Country, Bilbao, Spain Roberto Kolter, Harvard Medical School, Boston, MA, USA Germán Larriba, University of Extremadura, Badajoz, Spain Rubén López, Center for Biological Research, CSIC, Madrid, Spain Bernard M. MacKey, University of Reading, Reading, UK Michael T. Madigan, Southern Illinois University, Carbondale, IL, USA Beatriz S. Méndez, University of Buenos Aires, Buenos Aires, Argentina Diego A. Moreno, Technical University of Madrid, Madrid, Spain Ignacio Moriyón, University of Navarra, Pamplona, Spain Juan A. Ordóñez, Complutense University of Madrid, Madrid, Spain José M. Peinado, Complutense University of Madrid, Madrid, Spain Antonio G. Pisabarro, Public University of Navarra, Pamplona, Spain Carmina Rodríguez, Complutense University of Madrid, Madrid, Spain Fernando Rojo, Natl. Center for Biotechnology, CSIC, Cantoblanco, Spain Manuel de la Rosa, Virgen de las Nieves Hospital, Granada, Spain Carmen Ruiz Roldán, University of Murcia, Murcia, Spain Claudio Scazzocchio, Imperial College, London, UK James A. Shapiro, University of Chicago, Chicago, IL, USA John Stolz, Duquesne University, Pittsburgh, PA, USA James Strick, Franklin & Marshall College, Lancaster, PA, USA Gary A. Toranzos, University of Puerto Rico, San Juan, Puerto Rico Antonio Torres, University of Sevilla, Sevilla, Spain José A. Vázquez-Boland, University of Edinburgh, Edinburgh, UK Antonio Ventosa, University of Sevilla, Sevilla, Spain Tomás G. Villa, Univ. of Santiago de Compostela, Santiago de C., Spain Miquel Viñas, University of Barcelona, Barcelona, Spain Dolors Xairó, Biomat, S.A., Grifols Group, Parets del Vallès, Spain

Associate Editors Mercedes Berlanga, University of Barcelona Mercè Piqueras, Catalan Association for Science Communication Nicole Skinner, Imperial College, London Wendy Ran, International Microbiology Secretary General Jordi Mas-Castellà, International Microbiology Managing Coordinator Carmen Chica, International Microbiology Specialized editors Josefa Antón, University of Alicante Susana Campoy, Autonomous University of Barcelona Ramón Díaz, CIB-CSIC, Madrid Josep Guarro, University Rovira i Virgili Enrique Herrero, University of Lleida Emili Montesinos, University of Girona José R. Penadés, Inst. of Mountain Livestock-CSIC, Castellon Jordi Vila, University of Barcelona Digital media Coordinator Núria Radó-Trilla, Institute for Catalan Studies Webmaster Jordi Urmeneta, University of Barcelona

Addresses Editorial Office International Microbiology C/ Poblet, 15 08028 Barcelona, Spain Tel. & Fax +34-933341079 E-mail: int.microbiol@microbios.org www.im.microbios.org Spanish Society for Microbiology C/ Rodríguez San Pedro, 2 #210 28015 Madrid, Spain Tel. +34-915613381. Fax +34-915613299 E-mail: sem@microbiologia.org www.semicrobiologia.org Institute for Catalan Studies C/ Carme, 47 08001 Barcelona, Spain Tel. +34-932701620. Fax +34-932701180 E-mail: int.microbiol@microbios.org © 2014 Spanish Society for Microbiology, Madrid, & Institute for Catalan Studies, Barcelona. Printed in Spain ISSN (print): 1139-6709 e-ISSN (electronic): 1618-1095 D.L.: B.23341-2004

The Spanish Society for Microbiology (SEM) is a scientific society founded in 1946 at the Jaime Ferrán Institute of the Spanish National Research Council (CSIC), in Madrid. Its main objectives are to foster basic and applied microbiology, promote international relations, bring together the many professionals working in this science, and contribute to the dissemination of science in general and microbiology in particular, among society. It is an interdisciplinary society, with about 1800 members working in different fields of microbiology.

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International Microbiology is a quarterly, open-access, peer-reviewed journal in the fields of basic and applied microbiology. It publishes two kinds of papers: research articles and complements (editorials, perspectives, books, reviews, etc.).

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CONTENTS International Microbiology (2014) 17:131-184 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

Volume 17, Number 3, Setember 2014 RESEARCH ARTICLES

López-Martínez G, Borrull A, Poblet M, Rozès N, Cordero-Otero R Metabolomic characterization of yeast cells after dehydration stress

131

Manrique-Ramírez P, Galofré-Milà N, Serrano M, Aragon V Identification of a class B acid phosphatase in Haemophilus parasuis

141

Ariza-Miguel J, Hernández M, Fernández-Natal I, Rodríguez-Lázaro D Molecular epidemiology of methicillin-resistant Staphylococcus aureus in a university hospital in northwestern Spain

149

Spricigo DA, Cortés P, Moranta D, Barbé J, Bengoechea JA, Llagostera M Significance of tagI and mfd genes in the virulence of non-typeable Haemophilus influenzae

159

Chiellini C, Maida I, Emiliani G, Mengoni A, Mocali S, Fabiani A, Biffi S, Maggini V, Gori L,Vannacci A, Gallo E, Firenzuoli F, Fani R Endophytic and rhizospheric bacterial communities isolated from the medicinal plants Echinacea purpurea and Echinacea angustifolia

165

Padilha IQM, Valenzuela SV, Grisi TCSL, Diaz P, de Araújo DAM, Pastor FIJ A glucuronoxylan-specific xylanase from a new Paenibacillus favisporus strain isolated from tropical soil of Brazil

175

Journal Citations Reports The 2012 Impact Factor of International Microbiology is 2,556.

The journal is covered in several leading abstracting and indexing databases, including the following ones: Agricultural & Environmental Bio­­technology Abstracts; ASFA/Aquatic Sciences & Fisheries Abstracts; BIOSIS; CAB Abstracts; Chemical Abstracts; SCOPUS; Current Contents/Agriculture, Biology & Environmental Sciences; EBSCO; EMBASE/Elsevier Bibliographic Databases; Food Science & Technology Abstracts; ICYT/CINDOC; IBECS/ BNCS; ISI Alerting Services; MEDLINE/Index Medicus; Latindex; MedBioWorld; PubMed; SciELO-Spain; Science Citation Index Expanded; SciSearch.

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Front cover legends Upper left. Analysis of a transfer mechanism exploited by the human immunodeficiency virus type 1 (HIV-1) to infect new target immune cells.

Electron microscopy micrograph showing the intimate contact between a mature dendritic cell exposed to HIV-1 (top) and a CD4+ T cell (bottom). HIV-1 particles captured by the mature dendritic cell are polarized to the cell-to-cell contact area, favoring CD4+ T cell infection throughout the formation of an infectious synapse. Micrograph by M. Teresa Fernández-Figueras and Núria Izquierdo-Useros, Pathology Dept. at HUGTIP and AIDS Research Institute IrsiCaixa, Barcelona, Spain. (Magnification, ca. 20,000×) Center. View of Cartagena de Indias, a Colombian city in the Caribbean Coast Region, declared a World Heritage Site by UNESCO in November 1984. Cartagena was founded on June 1, 1533, by Spanish commander Pedro de Heredia, in the former location of the indigenous Caribbean Calamarí village. Its historic center, known as the “Walled City”, conserves the architecture of a Spanish city from the Colonial times. With this cover, we wish commemorate the 22nd Latin American Congress of Microbiology and 4th Colombian Congress of Microbiology, to be held on November 5–8, 2014, in Cartagena, organized by Prof. Howard Junca (president of the ALAM) and Prof. Liliana Marcela Ochoa G. (president of the ACM). See p. A3, this issue.

Upper right. Transmission electron micrograph of Escherichia coli with amyloid inclusions of the prionoid REPA-WH1. Gold particles for immunodetection map the distribution of molecules of chaperone DnaK (Hsp70), involved in the conformational dynamics of the protein, generating, from globular amyloid aggregates, a variant amyloid that is less cytotoxic. Micrograph by Rafael Giraldo, Department of Cellular and Molecular Biology, CIB–CSIC, Madrid. (Magnification, ca.

32,000×)

ders of the phylum Axostylata, specifically Tri­cho­ monadida, Hypermastigida, and Oxymonadida.

Pho­tograph (dark-field microscopy) by Rubén Duro (Center for Microbiological Research, CIM, Barcelona). See covers of Int. Microbiol. vol. 14 (2011) and R. Guerrero, L. Margulis, M. Berlanga, Int. Microbiol. 15(2013):133-143. (Mag­nification, ca. 1500×) Lower right. Scanning electron micrograph of a 24-h mixed biofilm containing Candida albicans hyphae and blastoconidia of Candida glabrata.

Photo by Cristina Marcos Arias. Faculty of Medicine, Uni­versity of the Basque Country, UPV/EHU, Bizkaia Campus, Bilbao. (Magnification, ca. 5000×)

Lower left. Micrograph of Trychonympha sp., a protist from the intestine of the lower termite Reti­ culitermes grassei. Lower termites have a symbiotic protist–bacteria community in their hindgut, that allows them to digest cellulose. The protists belong to basal eukaryotic taxa, i.e., flagellate or-

Back cover: Pioneers in Microbiology Federico Lleras Acosta (1877–1938), Colombia Portrait of Federico Lleras Acosta (1877– 1932), considered the father of Colombian microbiology and pioneer of public health in his country, who contributed significantly to the development of modern medicine in Colombia at a time when clinical physicians still lacked confidence in the laboratory. He was born in Bogotá on April 27, 1877, from Federico Lleras Triana and Amalia Restrepo in a family that has given to Colombia scientists and other intellectuals as well as politicians (Lleras Acosta’s son, Carlos Alberto [1908–1994] was the president of Colombia in 1966–1977).

He studied at the School of Veterinary Medicine, which depended on the School of Medicine and Natural Sciences of the University of the United States of Colombia and had been founded in 1884 by Claude Véricel, a French veterinary doctor. Along with the first microscope, Véricel had taken to Colombia laboratory reagents and media for bacteriological cultures. Graduates from that school were indeed the Colombian first microbiologists and worked mainly in fields related to microbiology, including food control and hygiene, the production of sera and vaccines, infectious and parasitic disease diagnostics, and public health. Lleras Acosta, who defended his master thesis on “La inspección sanitaria de

las carnes” (“Sanitary inspection of meat”) in 1902, specialized in serology and bacteriology, invested his savings in buying a modern microscope and founded the first clinical laboratory in Bogotá. He worked in various fields related to medical microbiology and public health; studied carbuncle, which affected the cattle, and the presence of Koch’s rod in urine, analized the quality of water in Bogotá, contributed to the diagnostic of plague, had to face an epidemics of enterocolitis which affected children in Bogotá, and spent time and efforts in the search for a method to culture Mycobacterium leprae, the leprosy rod. In 1936, he reported the cultivation of an acid-fast bacillus from the blood of patients suffering from cutaneous leprosy, but there was a high controversy about the cultivation of the true etiological agent of leprosy, whose culture in the laboratory has not yet been achieved. He also worked to set up a serological reaction for leprosy diagnostic. In 1934, Colombia’s President Alfonso Pérez Pumarejo created the Laboratorio Central de Investigaciones de la Lepra (Central Laboratory for Research on Leprosy) and Lleras Acosta was appointed its director. His pioneer work in microbiology was recognized by the University of Antioquia—the oldest university in Colombia, founded in 1803—which awarded him an honorary doctorate. Lleras Acosta died in Marseille, France, on March 18, 1938, on his way to Cairo to participate in the IV International Conference on Leprosy. Several hospitals and research centers have been named after him, the first—Federico Lleras Acosta Institute for Medical Research—having been inaugurated the very year of his death.

Front cover and back cover design by MBerlanga & RGuerrero

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RESEARCH ARTICLE International Microbiology (2014) 17:131-139 doi:10.2436/20.1501.01.215. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Metabolomic charactetization of yeast cells after dehydration stress Gema López-Martínez¶, Anna Borrull¶, Montse Poblet, Nicolas Rozès, Ricardo Cordero-Otero* Department of Biochemistry and Biotechnology, University Rovira Virgili, Tarragona, Spain Received 28 January 2014 · Accepted 15 July 2014

Summary. In this study, we analyzed the metabolite features of the yeasts Saccharomyces cerevisiae, Naumovia castellii, and Saccharomyces mikatae. The three species are closely related genetically but differ in their tolerance of desiccation stress.

Specifically, we determined whether certain metabolites correlated with cell viability after stress imposition. The metabolomic profiles of these strains were compared before cell desiccation and after cell rehydration. In S. mikatae, the presence of lysine or glutamine during rehydration led to a 20% increase in survival whereas during dehydration the levels of both amino acids in this yeast were drastically reduced. [Int Microbiol 2014; 17(3):131-139] Keywords: Saccaromyces mikatae · Saccharomyces cerevisiae · Naumovia castellii · viability · dehydration stress · metabolite extraction · wild yeast

Introduction Among the broad group of biological studies known as “omics,” metabolomics focuses on the analysis of a large number of metabolites at an organism level [8]. In yeast populations, intracellular metabolome analysis has improved our understanding of cell metabolism and thus of cellular responses to external physiological conditions [27]. The metabolomic profile of a selected yeast strain with respect to its response to dehydration is of commercial interest. In the food industry, dried yeast (active dry yeast) formulations have gained widespread acceptance because of the greater genetic stability at room temperature, resulting in savings in transport and storage costs. However, the loss of cell viability during the indusCorresponding author: R. Cordero-Otero Departament de Bioquímica i Biotecnologia Universitat Rovira i Virgili Marcel·lí Domingo, 1 43007 Tarragona, Spain E-mail: ricardo.cordero@urv.cat *

¶

Those authors contributed equally.

trial drying processes and the resulting lower activity have to some extent hindered the development of a high-quality inoculum [23]. Thus, identifying the changes in metabolites that occur in response to desiccation stress would facilitate efforts aimed at optimizing the drying process. During desiccation, the yeast’s metabolic processes are in a suspended state [4,21] whereas during rehydration these organisms rapidly swell and return to active life.

The methodological steps required for metabolite detection and data analysis are well established [3,10,14]. VillasBôas et al. [29] described methods to improve yeast sample preparation, especially with respect to cold methanol quenching, metabolic extraction by chemical treatment, and sample concentration through solvent evaporation. The quenching step consists of a brief metabolic arrest, which is required to obtain a focused cell metabolome “snapshot” of the cellular response to environmental conditions. In microbial physiol­ogy studies, the most commonly applied cell quenching method involves immersion of the sample in a cold aqueous methanol solution [11]. An advantage of this method compared to environmental pH variations is that it allows the subsequent sepa-

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Int. Microbiol. Vol. 17, 2014

ration of extra- from intra-cellular fractions, which significantly reduces contamination between the two [9].

Other well-known and widely used extraction methods in yeast research include methanol-chloroform, pH-dependent, and boiling ethanol treatments; however, problems have been identified in each case, including the recovery of non-phosphorylated compounds, compound stability at extreme pH values, and compound thermolability, respectively [3]. In the case of metabolite extraction from Escherichia coli and Saccharomyces cerevisiae, these effects can be mitigated by extracting in 50% (v/v) methanol after a –80ºC cycle [13,29].

However, most of the methods for metabolite extraction have been optimized for laboratory strain batch cultures, which are much more easily manipulated than either industrial strains or cells from non-growing conditions, in which the cell wall becomes a problematic barrier that strongly impacts the repeatability of quantitative assays [25]. The majortiy of metabolite extraction protocols for microorganisms require a final sample concentration step, typically a lyophilization procedure, that prevents further modification of cell compounds by eliminating the water-miscible solvent mix from the sample [6].

In this study, we compared three species of yeast, Saccharomyces cerevisiae, Naumovia castellii, and Saccharomyces mikatae, that differ in their dehydration tolerance. The analysis described herein is a continuation of a previous study in which we explored the effect of changes in cell lipid composition on the viability of these yeast strains following rehydration [23]. Through metabolomic characterization, the aim of this study was to identify the putative cellular compounds involved in overcoming dehydration/rehydration stress. S. mikatae was from the sensu stricto group, which includes yeast species closely related to S. cerevisiae. N. castellii belongs to the sensu lato group, which includes heterogeneous yeast species that diverge more significantly from S. cerevisiae [16]. In tandem with these two strains, S. cerevisiae was analyzed.

These three species differ from each other in a very limited number of physiological characteristics, some of which may be controlled by single-gene mutations [22]. The metabolite features of these yeasts during the dehydration/rehydration process were analyzed using an optimized gas ���������������� chromatography-mass spectrometry (GC-MS) metabolomics method.

Materials and methods Strains, growth conditions, and desiccation-rehydration.

Overnight liquid cultures of the yeast species Saccharomyces cerevisiae (CECT-1477, from Burdeos sparkling wine), Naumovia castellii (CECT11356, from Finland soil), and Saccharomyces mikatae (CECT-11823, Japan

LÓPEZ-MARTÍNEZ ET AL.

monosporic culture) at an initial OD600 of 0.5 were used as the inoculants. All three strains were grown in shake flasks (170 rpm) in YPD [1% (w/v) yeast extract, 2% (w/v) bacto-peptone, and 2% (w/v) glucose (Cultimed, Barcelona, Spain)] at 28ºC for 24 h. The effects of lysine (1, 2, 5, and 10%), Na2HPO4 (2, 5 and 10%), and glutamine (2, 5 and 10%) on cell viability during rehydration were studied by adding each compound individually to the pure water basal condition. Desiccation-rehydration, and cell viability were determined by flow cytometry, performed as previously described [22].

Metabolite extraction. Each step of the extraction was optimized based on the methods of Roessner et al. [24]. Five × 108 live cells were immediately frozen in cold 70% ethanol and stored at –20ºC until further analysis. After cell quenching, three methods for intracellular metabolite extraction were evaluated by assessing the percentage of unbroken cells. The mean number of unbroken cells per ml was calculated according to the CFU (colony-forming units) of treated cells after taking into account the CFU per ml before cell breakage. Frozen cells were pelleted and resuspended in methanol-water (1:1, v/v) to a volume of 400 µl, as described by Villas-Bôas et al.

[29], with 10 µl of ribitol at 2 mg/ml (Sigma, Switzerland) added as an internal standard (IS). To optimize this step, samples were incubated at 90ºC either for 10 min, or for 5 min in the presence of 0.2-mm glass beads (BioSpec Products, USA) in a sonication bath (J.P. Selecta, Spain), or for 5 min in the presence of 0.5-mm acid-washed glass beads (~300 µl). The samples were then disrupted in a multitube bead-beater (BioSpec Products, USA) using five cycles of 1 min/beat, followed by 30-s rest for cooling. After centrifugation, the supernatant was dried in a SC110 speed vacuum system SC110 (Savant Instruments, USA) for 4 h. The dried residue was redissolved and derivatized for 1 h at 40ºC in 50 µl of 20 mg/ml methoxyamine hydrochloride in pyridine (Sigma, Japan; Fluka, India), followed by a 90-min treatment at 40ºC with 70 µl of N-methyl-N-(trimethylsilyl) trifluoroacetamide (Sigma, USA).

Gas chromatography-mass spectrometry analysis. Gas chromatography (GC) was performed using an Agilent Technologies Network GC system 6890N connected to an HP computer with the ChemStation software (Agilent Technologies). Compounds were detected using an inert mass selective detector (MSD, model 5975, Agilent Technologies). Two µl of the cell extract was injected at a split ratio of 20:1 into a DB-5HT column (30 m × 0.25 mm × 0.1 µm; Agilent Technologies) with an automatic injector (7683B, Agilent Technologies). Helium was used as the carrier gas at a constant flow of 1.0 ml/min. The injector temperature was 200ºC. The column oven temperature was initially held at 80ºC for 4 min and then increased first to 200ºC at a rate of 5ºC/min and then to 300ºC at a rate of 25ºC/min, where it was held for 7 min. The MSD transfer temperature was 300ºC. The MSD quadrapole and source temperatures were maintained at 180ºC and 280ºC, respectively. The MSD data were acquired in electronic ionization scan mode at 70 eV within the range of 35–650 amu after a solvent delay of 4 min. Postrun analysis was performed with the Agilent MSD Chemstation. The relative abundance of each identified compound was calculated according to the respective chromatographic peak areas corrected with respect to the IS peak.

These values were relativized to the 1 × 109 CFU, according to the cells plated and counted after 24 h of culture and desiccation/rehydration, and expressed as percentages.

Flow cytometry analysis. Flow cytometry was carried out using a CYFlow space instrument (PARTEC, Germany) fitted with a 22-mW ion laser for excitation (488 nm); a single emission channel (575-nm band-pass filter) was used for monitoring. Instrument control, data acquisition and data analysis were performed using FloMax software (Quantum Analysis, Germany).

The two-color fluorescent probe from the LIVE/DEAD yeast viability kit was used to label the cells. In this system, plasma-membrane integrity and fungal metabolic function are required to convert the intracellular yellow-green fluorescence of FUN1 into red-orange intravacuolar structures, Calcofluor White

DEHYDRATION STRESS IN YEAST

M2R labels cell-wall chitin with blue fluorescence regardless of metabolic state (Life Technologies). An overnight YPD culture of a Saccharomyces sp. strain was taken as a control of full viability (99% by FUN1 red-orange/Calcofluor white stain).

Microscopy. The cells were viewed with a Leica microscope (DM4000B, Germany) and a digital camera (Leica DFC300FX). The Leica IM50 software was used for image acquisition.

Statistical analysis. The data were analyzed by principal component analysis (PCA) using the SPSS 20.0 statistical software package. For further analysis, one-way ANOVA and the Tukey test were used with the same program. Statistical significance was set at P ≤ 0.05.

Results and Discussion Yeast cell viability after dehydration stress. Differences in the desiccation tolerance of the three closely related yeast species were assessed by flow cytometry. FUN1/ Calcofluor co-staining was performed to quantify cell metabolic activity and cell wall integrity (Fig. 1A). This staining protocol allowed metabolically active cells (live cells) to be distinguished from damaged or non-active cells (dead cells).

The mean number of viable cells after rehydration was calculated relative to cell viability before drying. After drying, the cells of each species were resuspended in either 10% trehalose or, as the reference condition, pure water to evaluate viability. The viability of cells in the deionized water condition was lower for S. mikatae and N. castellii than for S. cerevisiae (20%, 40% and 84%, respectively) whereas the viability of yeast cells dried in the presence of trehalose was increased by approximately 20% for all three species (Fig. 1A). Trehalose was previously shown to act as a membrane protector by reducing the membrane phase-transition temperature during the desiccation-rehydration process, without having a significant impact on cell metabolites [7,23].

Metabolite extraction. The metabolite fractions of the three strains were investigated to determine their dehydrationstress tolerance metabolite profiles of the yeasts. The extraction process was optimized for these strains based on the protocol of Roessner et al. [24]. After the cells were frozen to stop their metabolism, extracts were prepared using three different methods: thermal shock, sonication, and vortexing.

Figure 1B shows the cells treated according to these methods and then evaluated by microscopy and flow cytometry to quantify the unbroken cell fraction, both before and after treatment. Resuspending the cells in the presence of 0.2-mm

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glass beads after shaking with a mini-bead beater for 5 × 1 min with a 1-min rest between treatments resulted in 98% broken cells whereas thermal shock and sonication resulted in 30% and 20% un­broken cells, respectively.

Metabolomic profiling of yeast cells during the desiccation process. To understand the metabolic differences between S. mikatae, N. castellii, and S. cerevisiae cells before the imposition of stress, their metabolomic profiles before dehydration (BD) and after rehydration (AR) were compared. The cellular metabolites were analyzed by GC-MS, with 44 peaks identified and quantified for each strain and condition (BD and AR). These metabolites included amino acids, glycolytic compounds, sugars, fatty acids, and organic acids.

Multivariate data analysis was performed using PCA to examine the variations among metabolites between the BD and AR steps in S. cerevisiae, N. castellii, and S. mikatae (Fig. 2A, B, and C, respectively). Endogenous trehalose increased by 8.7fold, 5.5-fold, and 3-fold during stress imposition in S. mikatae, N. castellii and S. cerevisiae, respectively, but for reasons of scale the data are not included in Fig. 2. However, intracellular trehalose did not correlate with the desiccation tolerance of the three species, as previously reported by RodríguezPorrata et al. [23]. PCA was performed after data processing, including normalization according to the IS and CFU corrections (see Materials and methods).

The relative amounts of fatty acids, organic acids, and compounds resulting from glycolysis differed significantly in the three species, both BD and AR. By contrast, there were no significant differences in the BD amounts of amino acids and sugars. The largest differences occurred in N. castellii, whose metabolite profile during dehydration was the opposite of that of S. cerevisiae, which also had a 40% greater viability AR (Fig. 2A and Fig. 1A). Although phosphoric acid is toxic to yeast cells, the BD concentration in N. castellii was 4-fold higher than in the two other yeast strains and increased 1.5-fold during stress, whereas in S. mikatae and S. cerevisiae the concentration decreased by 3-fold (Fig. 2C). These results suggested that the viability of N. castellii would be negatively affected during stress due to inefficient detoxification and/or a poor capacity for metabolic esterification of phosphoric acid into nucleic acids, proteins, lipids, and sugars [18]. Yeast cells store phosphate in vacuoles as polyphosphate, which normally increases during the stationary growth phase [18]. No significant differences among the yeast species were recorded BD in terms of phosphate content (Fig. 3). However, between the BD and AR steps, the phosphate content of S. mikatae and S. cerevisiae

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B

Fig. 1. Yeast cell viability following air drying and rehydration. (A) Cells were dried to a moisture content of 5% at 28ºC. The scale of relative viability (%) indicates the percentage of experimental values for the different yeasts relative to the highest viability for Saccharomyces cerevisiae. The values shown represent the means of five independent samples ± SD, as evaluated by flow cytometry. The viability of cells dried in the presence (gray bars) and absence (white bars) of trehalose was evaluated. (B) Microscopy images show the broken S. cerevisiae cells after treatment, and the predominance of large cell fragments. White spots are the cell walls of empty cells, and dark spots are intact cells.

decreased by 70% and 30%, respectively, while it doubled in N. castellii (Fig. 3). Nonetheless, this greater accumulation of phosphate by N. castellii than by S. cerevisiae during AR did not enhance its dehydration tolerance, as reflected in the differ-

ences in viability: 60% vs. 95%, respectively (Fig. 1A). S. mikatae had a significantly lower content of phosphate after drying and rehydration, consistent with its low viability (36%). The main metabolic response of S. cerevisiae to osmotic pressure,

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Fig. 2. Metabolic variations between before drying (BD) and after rehydration (AR), as examined by principal component analysis, during stress imposition in (A) Saccharomyces cerevisiae, (B) Naumovia castellii, and (C) Saccharomyces mikatae. Principal component analysis revealed the largest variance between the BD and AR steps, as shown by the changes in the intracellular alanine concentration (compare A and C) during stress imposition.

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Fig. 3. Relative abundance of significant and stress-related compounds BD (white bars) and AR (grey bars) in Saccharomyces cerevisiae, N. castellii and S. mikatae. The values represent the means of three independent experiments. Results for each compound with statistically significant differences (P < 0.05) compared to the BD condition.

which fluctuates during dehydration and rehydration, is the cytoplasmic retention of glycerol [12,26]. In fact, in N. castellii and S. cerevisiae intracellular glycerol accumulation during stress imposition increased by 50%, whereas in S. mikatae it decreased by 50%, which further explains the low viability of these species (Fig. 3).

Our findings suggest a relationship between glycerol accumulation and desiccation tolerance, whereby a reduction in glycerol content during stress induction reduced cell viability.

Glycerol biosynthesis is an important side-reaction of the glycolytic pathway, as it provides the NAD+ necessary for cellular glycolysis activity [2]. Thus, according to our results, cells that synthesized glycerol during dehydration also had a backup supply of NAD+ that allowed rapid glycolytic activation when cell activity was resumed. The NAD+ generated is oxidized to maintain redox balance by the conversion of glyceraldehyde-3-phosphate to 1,3-bisphosphoglycerate by the enzyme glyceraldehyde-3-phosphate dehydrogenase (GAPDH). In the GAPDH glycolytic pathway, under stress conditions, glycolytic flux is strongly directed to glycerol accumu-

lation, depending on the kinetics of GAPDH [17]. Saccharomyces. cerevisiae is unusual in that three genes (TDH1–3) encode three different GAPDH isoenzymes, whereas other yeasts, such as C. albicans and K. marxianus, contain only one gene [1,15,28]. The TDH1 gene encodes a less efficient GAPDH isoenzyme but it is the only gene that is expressed under conditions of cell stress [5,17]. Additional experiments are required to determine whether TDH1 activation is responsible for glycerol accumulation during dehydration. Glycolysis is used by almost 800 yeast species in a sequence of enzymatic reactions that convert glucose to pyruvate, which, under aerobic conditions, is channelled into the mitochondria, where it enters the tricarboxylic acid cycle (TCA) and is eventually converted into acetyl-CoA. Alternatively, pyruvate acts as a precursor for a side-reaction in α-alanine synthesis [19]. Intracellular α-alanine increased by 3-fold and 2-fold in N. castelli and S. cerevisiae, respectively, while in S. mikatae there was a 2-fold decrease between the BD and AR steps (Fig. 3). Acetyl-Co-A branchpoint metabolites can be converted into citric acid or lipids and

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Fig. 4. Effect of rehydration on cell viability. Saccharomyces mikatae yeast cells were incubated at 37ÂşC for 30 min in pure water or in the presence of lysine (Lys), phosphate (Pho), or glutamine (Gln). The scale of relative viability (%) indicates the percentage of experimental values for the different yeasts relative to the non-complemented condition (H2O). The values are the means of three independent experiments. Results with statistically significant differences (P < 0.05) compared to the H2O condition are presented.

fatty acids, such as palmitoleic acid, which during dehydration increased in N. castellii and S. cerevisiae whereas in S. mikatae a 30% decrease was recorded. These results concerning palmitoleic acid and Îą-alanine indirectly suggest a greater reduction of pyruvate availability from the glycolytic pathway in S. mikatae than in the other two yeasts during stress induction. The only TCA cycle intermediate that showed significant variations during stress was succinic acid, whose levels increased by 9-fold and 2-fold in N. castellii and S. cerevisiae, respectively, while in S. mikatae the levels remained the same (Fig. 3). This accumulation of succinic acid in S. cerevisiae does not agree with previous results shown by Raab et al. [20]. The opposite is true in S. mikatae and could explain the insignificant differences between the BD and AR values while the high level of accumulation in N. castellii highlights its metabolic difference compared to the other two yeast species. In the TCA cycle, after three successive enzymatic steps, succinic acid is converted to oxaloacetate, another branch-point metabolite because it can be transaminated to form aspartate. The BD abundances of aspartic acid were similar in the three yeasts. However, in N. castellii and S. cerevisiae the AR levels of aspartic acid increased by 13-fold, but only by 3.5-fold in S. mikatae (Fig. 3). The changes in the abundances of aspartic acid and succinic acid

suggested that S. mikatae cells should lack TCA activity during dehydration and rehydration, in contrast to N. castellii and S. cerevisiae. The TCA intermediate 2-oxoglutarate is also a branch-point metabolite; it is converted through transamination to lysine and glutamate, which can be used to synthesize other amino acids, such as glutamine, proline, and arginine. In our study, the BD contents of lysine and glutamine were similar in the three yeast species, whereas during stress imposition the changes in the abundances of these amino acids did not correlate with putative changes in yeast TCA activities. Thus, S. mikatae contained 3-fold less lysine after stress induction, while in N. castellii and S. cerevisiae lysine increased by 3-fold and 2-fold, respectively. Glutamine decreased significantly in S. mikatae and S. cerevisiae, by 6-fold and 3-fold, respectively.

In fact, one of the two anaplerotic pathways responsible for replenishing the TCA cycle with intermediates, thereby maintaining its function, is the glyoxylate cycle, in which isocitrate dehydrogenase converts isocitrate into 2-oxoglutarate. The participation of both the TCA and glyoxylate cycles in providing 2-oxoglutarate to cells is not consistent which the changes in glutamine and lysine concentrations during stress induction.

These variations also did not correlate with the dehydration tolerance profiles of any of the yeast species.

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Effect of compound supplementation on yeast cell viability. We next sought to ascertain whether the low viability of S. mikatae after rehydration was due to low levels of lysine, Na2HPO4, or glutamine. Therefore, we tested several rehydration media in an attempt to overcome the drop in yeast viability during this process. In this experiment, all additives were resuspended in deionized water and pure water served as the reference condition for cell viability evaluation by flow cytometry. A statistically significant increase of 20% in cell survival driven by 1, 2, and 5% lysine or 10% glutamine supplementation was observed in S. mikatae (Fig. 4) but not in S. cerevisiae. Unexpectedly, phosphate supplementation was detrimental and increased cell death by 30% compared to the pure water reference condition (data not shown).

In conclusion, the changes that take place in cells that allowed them to overcome dehydration stress were investigated at the metabolomic level in S. cerevisiae, N. castellii, and S. mikatae. The extraction method for GC-MS analysis was optimized using a mini bead beater system, which enhanced cell breakage by ~20%. After data analysis and PCA, the observed variations in intracellular metabolites could be related to the form of central carbon metabolism, which offered a better understanding of the cellular metabolic differences between yeasts with respect to desiccation tolerance. Nevertheless, the poor survival of S. mikatae and N. castellii could not be ascribed to a single metabolite; rather, at least in N. castellii, the accumulation of both succinic acid and phosphoric acid by stress imposition might account for its low viability (60%). The metabolite profiles suggested the activity of the glycolytic pathway and TCA cycle in N. castellii and S. cerevisiae, but not in S. mikatae, which in the latter species could have led to a reduction in the levels of secondary metabolites required for viability following rehydration. Our results also demonstrated that the presence of lysine or glutamine during rehydration had a positive effect on the recovery of S. mikatae cell activity, although the benefits achieved by the supplementation of these compounds might be yeast dependent. Further investigations extending to other aspects such as global gene expression and proteomics will provide a better understanding of the cellular mechanisms involved in overcoming desiccation stress.

Acknowledgements. This work was financially supported by grants from the Spanish government (Projects AGL2010-22001-C02-02 and AGL2012-40018-C02-01, awarded to NR and RC, respectively). GL and AB also thank the Rovira i Virgili University for the doctoral fellowships FI-DGR 2012, AGAUR and 2009BRDI/12/12, respectively.

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Competing interest: None declared.

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OMICS 14:553-561 7. Eleutherio EC, Araujo PS, Panek AD (93) Role of the trehalose carrier in dehydration resistance of Saccharomyces cerevisiae. Biochim Biophys Acta 1156:263-266 8. Fiehn O, Kopka J, Dörmann P, Altmann T, Trethewey R, Willmitzer L (2000) Metabolite profiling for plant functional genomics. Nat Biotechnol 18:1157-1161 9. Gonzalez B, François J, Renaud M (1997) A rapid and reliable method for metabolite extraction in yeast using boiling buffered ethanol. Yeast 1:1347-1355 10. Kell BD (2004) Metabolomics and systems biology: making sense of the soup. Curr Opin Microbiol 7:296-307 11. Koning W de, van Dam K (1992) A method for the determination of changes of glycolytic metabolites in yeast on a subsecond time scale using extraction at neutral pH. Anal Biochem 204:118-123 12. Larsson K, Ansell R, Eriksson P, Adler L (1993) A gene encoding snglycerol 3-phosphate dehydrogenase (NAD+) complements an osmo­ sensitive mutant of Saccharomyces cerevisiae. Mol Microbiol 10:11011111 13. Maharjan RP, Ferenci T (2003) Global metabolite analysis: the influence of extraction methodology on metabolome profiles of Escherichia coli.

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RESEARCH ARTICLE International Microbiology (2014) 17:141-147 doi:10.2436/20.1501.01.216. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Identification of a class B acid phosphatase in Haemophilus parasuis Paula Manrique-Ramírez,1 Núria Galofré-Milà,1 Marta Serrano,1 Virginia Aragon1,2* Centre de Recerca en Sanitat Animal (CReSA), UAB-IRTA, Campus de la Universitat Autonoma de Barcelona, Bellaterra, Barcelona, Spain. 2Institut de Recerca i Tecnologia Agroalimentàries (IRTA), Barcelona, Spain

1

Received 8 April 2014 · Accepted 15 September 2014 Summary. An acid phosphatase activity was detected in the supernatant of Haemophilus parasuis, a Gram-negative pleomorphic bacillus and the causative agent of Glässer’s disease in pigs. To identify the gene responsible for the secreted activity, a genomic library of H. parasuis strain ER-6P was produced in Escherichia coli. Screening of the library allowed identification of two homologs to known phosphatases: PgpB and AphA. PgpB was predicted to be located in the bacterial membrane through six transmembrane domains while AphA was predicted to have a signal peptide. The aphA gene was cloned and expressed in E. coli. Characterization of H. parasuis AphA indicated that this protein belongs to the class B nonspecific acid phosphatases. AphA contained sequence signatures characteristic of this family of phosphatases and its activity was inhibited by EDTA. The optimal pH of recombinant AphA differed from that of the phosphatase activity found in H. parasuis supernatants. In addition, the phosphatase activity from H. parasuis supernatants was not inhibited by EDTA, indicating that H. parasuis AphA does not account for the phosphatase activity observed in the supernatants. Our results demonstrate the presence of a class B acid phosphatase (AphA) in H. parasuis and suggest that the bacterium would also secrete another, as yet unidentified phosphatase. [Int Microbiol 2014; 17(3):141-147] Keywords: Haemophilus parasuis · non-specific acid phosphatases · phosphatase activity · Glässer’s disease

Introduction Haemophilus parasuis is a Gram-negative pleomorphic bacillus and the etiological agent of Glässer’s disease in pigs, a severe systemic disease characterized by fibrinous polyserositis, polyarthritis, and meningitis that is responsible for important economic losses in the swine industry worldwide. H. parasuis has been shown to produce a variety of potential mem* Corresponding author: Virginia Aragon Centre de Recerca en Sanitat Animal (CReSA) Universitat Autònoma de Barcelona 08193 Bellaterra, Barcelona, Spain Tel. +34-935814494. Fax +34-935814490 E-mail: virginia.aragon@cresa.uab.es

brane-associated virulence factors [5], but little is known about the role of its acid phosphatases.

Acid phosphatases are ubiquitous in prokaryotes and eukary­ otes. Based on criteria such as specificity and optimum pH, phosphatases can be classified into several families. Thus, bacterial non-specific acid phosphatases (NSAPs) are categorized into three types (classes A, B and C) [3,23]. Reports indicate that in Gram-negative bacteria these enzymes play a critical role in numerous processes, including pathogenesis [2,6,8,9,16,19,20]. Acid phosphatases catalyze the transfer of a phosphoryl group from phosphomonoesters to water at an acidic pH. The class B acid phosphatases are a group of homo­ tetrameric secreted phosphohydrolases containing a 25-kDa polypeptide monomer. These enzymes are characterized by

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conserved sequence motifs including four conserved aspartate residues, which are characteristic of the molecular superfamily of phosphohydrolases referred to as “DDDD” [11,21, 22]. The class B signature sequence is divided into two domains, an N-terminal motif composed of FDIDDTVLFSSP and a C-terminal motif of YGD-(A/S)-DXD-[I/V] [18]. The aspartate nucleophile in this group is located in the N-terminal region of the signature sequence. These phosphatases are members of the haloacid dehalogenase (HAD) superfamily and include a large number of bacterial phosphatases. The first bacterial nonspecific class B acid phosphatase described was NapA from Morganella morganii [21], but others have been identified since then, such as the AphA phosphatase of Escherichia coli [14,22]. A napA homolog was also detected in the genome of Haemophilus influenzae [7]. Here we provide the first report of a class B acid phosphatase in H. parasuis.

Materials and methods Bacterial strains and growth conditions. Haemophilus parasuis strains (Table 1) were grown on chocolate agar plates (BioMerieux, Madrid, Spain) at 37ºC in an atmosphere of 5% CO2 or in brain heart infusion (BHI) broth when a liquid culture was needed (see below). All H. parasuis strains were stored frozen in BHI with 20% glycerol at –80ºC. The majority of the strains were stored after the minimum number of passages needed for isolation (usually two passages on plates). The passage numbers of strains IT29205, 9904108, and P015/96 and the reference strains Nagasaki and SW114 are not known. Escherichia coli BL21 was used as the host for recombinant plasmids and was grown in LB medium at 37ºC. LB medium was supplemented with ampicillin (100 μg/ml) or chloramphenicol (30 μg/ml) as required.

Preparation of supernatants. Culture supernatants were analyzed at different growth phases. Strains of H. parasuis were grown in BHI broth supplemented with isovitalex (BD, Madrid, Spain) at 37ºC in an orbital shaker (200 rpm). Bacterial growth was assessed by measuring the absorbance at 660 nm in a VIS7200 spectrophotometer (Dinko Instruments, Barcelona, Spain).

Supernatants were obtained by centrifugation followed by filtration (0.22 µm).

For some experiments; the supernatants were concentrated 100-fold through a Millipore YM10 ultrafiltration cell.

Phosphatase assays. Phosphatase activity was measured using the fluorescent substrate 4-methylumbelliferyl phosphate (MUP) [13,17]. Acid phosphatase activity was measured in 0.1 M sodium acetate buffer (pH 5.2) or in 0.1 M sodium citrate buffer (pH 5.2) and MUP at a final concentration of 0.6 mg/ml. The reaction was measured in a fluorometer (ASCENT Fluoroskan & FL; Thermo Labsystems). The fluorometer was programmed for incubation at 37ºC and for fluorescence measurements at different time points (usually, at 0, 10 min, 30 min, 1 h and 2 h). Fluorescence was measured at an excitation of 355 nm and an emission of 460 nm.

In standard phosphatase assays, 96-well microtiter plates were used with 50 μl of sample, 50 μl of 2× reaction buffer, and MUP. To measure phosphatase activity at different pHs, 0.1 M sodium citrate buffer was used for pH 4,

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5.2, and 6, and 0.05 M Tris buffer used for pH 7, 8, and 9. The MUP concentration was the same as above. An acid phosphatase from potato (Sigma-Aldrich) was used in the assays as a positive control.

The effect of ethylenediaminetetraacetic acid (EDTA) on phosphatase activity was assessed by the addition of 20 mM EDTA to the reaction. The effects of tartrate and molybdate on phosphatase activity were assayed by adding sodium tartrate or sodium molybdate to the reaction at 1, 10, 100, 1000 and 10,000 μM. Construction of a genomic library and screening. DNA from H. parasuis strain ER-6P (acid phosphatase positive) was extracted and partially digested with Sau3AI. The reaction was incubated at 37ºC for 2 min and then stopped with 40 mM EDTA. DNA fragments between 4 and 5 kb were selected and excised from an agarose gel and then purified using the Wizard SV gel and PCR clean-up system (Promega). Plasmid pACYC184 digested with BamHI and then dephosphorylated was used to clone the 4- to 5-kb genomic fragments from strain ER-6P, which were then introduced into E. coli BL21. The genomic library consisted of 2,880 clones, which was sufficient to ensure the representation of the H. parasuis genome. To screen for acid phosphatase activity, the individual clones from the genomic library were cultured in 96-well plates with 200 μl of medium. After incubation at 37ºC, the complete culture and the supernatant from each well were analyzed for phosphatase activity. Plasmid DNA from the positive clones was purified and the insert was sequenced with primers pACYCseq-F (5������������������� ′������������������ -ACTTGGAGCCACTATCGACTAC-3′) and pACYCseq-R (5′-CGGTGATGTCGGCGATA­TAGG-3′).

The complete sequence included in each clone was deduced by comparison with the H. parasuis genome sequence from strain SH0165 (�������������� GenBank������� accession number NC_011852.1).

Cloning of aphA homolog from Haemophilus parasuis.

Screening of the 4- to 5-kb genomic library and BLAST searches of the genome of H. parasuis SHO165 (accession number YP002476402.1) detected an aphA homolog in H. parasuis. The aphA gene, with its putative promoter region (predicted by BProm), was PCR-amplified from the strains described in Table 1, using primers APha-Fw-prom (5���������������������� ′��������������������� -ATGTTCCCTATAACCTATTGTG-3′) and Apha-Rev (5′-ATTAGTAGCTTGAATTTATAATAAC-3′). The resulting 900-bp PCR fragment was purified, cloned into pGemT-easy, and electroporated into E. coli BL21. Transformants were selected and the acid phosphatase activity of the clones was evaluated in complete cultures and in the supernatants. Inserts in the clones were sequenced and the orientation of the gene in each clone was checked. The presence of a putative signal peptide in the predicted amino acid sequence was analyzed using SignalP 4.1 and PSORT. Transmembrane domains and cellular location were analyzed using TMHMM Server v.2.0 and PSORT.

Results and Discussion Secreted acid phosphatase activity. An acid phosphatase activity was detected in the supernatant of H. parasuis, with a secretion peak in the early-stationary phase of bacterial growth and an optimal pH of 5–6 (Fig. 1). The activity of the supernatants could be preserved at –80ºC if 50% glycerol was added to the samples. Failure to add glycerol rendered the samples inactive after freezing at –80ºC. Although the activity may have been released into the supernatant after bacterial lysis, this was unlikely since the samples were obtained at the end of the

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Table 1. Strains of Haemophilus parasuis used in this study and their main characteristicsa Susceptibility tob Serovar

Serum

Nagasaki

5

R

Strain

Phagocytosis

Disease status

Reference or source

R

Glässer’s disease

Reference strain [10]

Strains from lesions ER-6P

15

R

R

Glässer’s disease

[4]

IT29205

4

R

R

Glässer’s disease

[1]

373/03A

7

I

R

Glässer’s disease

[1]

264/99

10

R

R

Glässer’s disease

[1]

2725

10

S

R

Glässer’s disease

[1]

228/04

5

R

R

Glässer’s disease

[12]

PV1-12

15

S

R

Glässer’s disease

[1]

CT175-L

15

S

ND

Glässer’s disease

[4]

PC4-6P

12

R

R

Glässer’s disease

[4]

9904108

4

S

R

Glässer’s disease

[1]

P015/96

5

R

R

Pneumonia

[12]

SW114

3

S

S

Healthy

Reference strain [10]

F9

6

S

S

Healthy

[12]

IQ1N-6

9

S

S

Healthy

[4]

ND14-1

7

S

S

Healthy

[4]

FL3-1

7

ND

ND

Healthy

[12]

FL1-3

10

S

I

Healthy

[12]

CA38-4

12

R

R

Glässer’s disease

[12]

MU21-2

7

S

S

Healthy

[12]

c

Nasal strains

VS6-2

15

I

S

Healthy

[12]

SC14-1

15

S

S

Healthy

[12]

SL3-2

10

S

S

Healthy

[4]

All strains are aphaA positive.

S, Sensitive; I, intermediate; R, resistant; ND, Not determined. c This strain did not reproduce disease when inoculated intranasally in colostrum-deprived piglets [1]. a b

exponential phase of growth, during which time no significant lysis occurred. Moreover, supernatants obtained after overnight incubation showed phosphatase activity at an alkaline pH (not shown), probably due to the release of an alkaline phosphatase after bacterial lysis.

A gradual loss of the acid phosphatase activity was observed after several in vitro passes of strain ER-6P on agar plates. This was the case after 6, 10, and 14 passes (ER-6Pp6, ER-6Pp10, and ER-6Pp14). After 14 passes, acid phosphatase activity in the supernatant of ER-6Pp14 was no longer detectable, even though

the growth rate of this strain in the laboratory was similar to that of the original ER-6P strain (not shown), recovered from the pericardium of a diseased pig and maintained frozen with no additional passages. We assume that, under standard laboratory conditions, H. parasuis does not require this acid phosphatase and therefore blocks its production. Similarly, a previous report had demonstrated the reduced expression of the H. parasuis capsule after in vitro passage of the bacterium [15]. The role of the acid phosphatase activity in infection, whether to obtain nutrients or as a virulence factor, remains to be determined.

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Fig. 1. Phosphatase activity in the supernatant prepared from Haemophilus parasuis strain ER-6P harvested at the beginning of the stationary phase of growth (gray bars) and in the supernatant of clone E. coli BL21 pGem-T Easy-aphA FL3-1#19 (black bars). Phosphatase activity was measured at different pH values, with MUP as the substrate. Phosphatase activity is represented as relative fluorescence units (RFU) per min. (Average of duplicate samples ± standard deviation.)

Screening of the genomic library. To identify the gene responsible for the phosphatase activity detected in H. pa­ra­suis supernatants, individual clones of the genomic library H. parasuis ER-6P were tested for the production of phosphatase activity in the presence of MUP. Sequencing of the phosphatasepositive clones yielded four clones with a class B acid phosphatase (aphA homolog gene) and one clone with a phosphatidylglycerophosphate phosphatase (pgpB homolog gene). The pgpB and aphA genes were PCR-amplified and cloned to subsequently determine their activity. Acid phosphatase activity was demonstrated in the pgpB clones (not shown). The predicted sequence of PgpB contained six transmembrane domains probably located in the cytoplasmic membrane. These data are in agreement with the location of other bacterial PgpB proteins in the cytoplasmic membrane. Thus, in the following, we focus our attention on the aphA homolog as a possible secreted phosphatase.

Analysis of the aphA gene from the available H. parasuis sequence demonstrated that the predicted protein had the features of a NSAP class B phosphatase, including the two signature motifs characteristic of these enzymes (Fig. 2). Also, a signal peptide was detected by SignalP in the predicted protein. The predicted molecular mass of 24.33 kDa for the monomer is characteristic of class B phosphatases [18].

The complete aphA gene, with the putative promotor region, from virulent and non-virulent strains of H. parasuis was PCR-amplified and sequenced (Table 1). No association between aphA and the putative virulence of the strains was observed, since the gene was detected by PCR in all of the analyzed strains (Table 1). Clones were produced with aphA from strains ER-6P, FL3-1, IQ1N-6, and ND14–1. All of the clones showed phosphatase activity, but only one aphA-containing clone, from strain FL3-1 (clone pGem-T Easy aphA FL3-1#19), secreted the activity into the culture supernatant, (Fig. 1). Sequencing revealed that the only difference between this clone and others from the same strain was the orientation of the insert in the plasmid, as the orientation of the insert in clone FL3-1#19 was opposite to that of the other inserts.

Thus, why the activity was secreted by this clone but not by the others is unclear.

Characterization of the AphA homolog from Haemophilus parasuis. The supernatant from BL21 pGem-T Easy aphA FL3-1#19 was analyzed at different pHs and the activities were compared to the activity in H. parasuis supernatants (Fig. 1). The activity produced by the H. parasuis AphA homolog had a different optimal pH than the activity of the H. parasuis supernatants, indicating that the two

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Fig. 2. Alignment of the predicted amino acid sequence of acid phosphatase AphA from different strains of Haemophilus parasuis (CT: CT175-L; ER: ER-6P; F9; FL: FL3-1; IQ: IQ1N-6; SW: SW114; ND: ND14-1; IT: IT29205; NG: Nagasaki; SH: SH0165). The catalytic domain characteristic of class B acid phosphatases is presented in two boxes; motif F-D-I-D-D-T-V-L-F-S-S-P, located in the N-terminal moiety, and motif Y-G-D-(A/S)-D-X-D-(IV), located near the C-terminus [23].

supernatant of strain ER-6P was only partially inhibited at 10 mM whereas there was no effect on the recombinant AphA phosphatase (Fig. 4).

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activities were produced by different enzymes. The optimal pH for H. parasuis Apha with MUP was around 7–8. This may seem high for an acid phosphatase, but the optimal pH of these enzymes varies with the substrate [18]; thus, the activity of AphA with its natural substrate may be maximal at an acidic pH. Since class B NSAPs require Mg2+, we evaluated the inhibitory effects of EDTA. The activity produced by clone pGem-T Easy aphA FL3-1#19 was indeed inhibited by EDTA (Fig. 3), in agreement with its class B phosphatase structure [18]. However, the activity in the supernatant of H. parasuis strain ER-6P was not affected by the addition of 20 mM EDTA to the reaction (not shown). Thus, besides having the characteristic domains of a type B acid phosphatase, the H. parasuis AphA homolog is inhibited by EDTA, in agreement with its classification within this group of phosphatases.

Therefore, the inhibition of Apha activity by EDTA is compatible with the requirement of divalent cations by class B phosphatases.

Tartrate was also tested as phosphatase inhibitor but it had no effect on the secreted activity in the supernatants of either H. parasuis ER-6P or the recombinant AphA, up to a concentration of 10 mM sodium tartrate. When molybdate concentrations up to 10 mM were similarly tested, the activity in the

Fig. 3. Inhibition of AphA by EDTA. Phosphatase activity in the supernatant of clone E. coli BL21 pGem-T Easy aphA FL3-1#19 with (black bars) and without (white bars) 20 mM EDTA, at different pH values. Phosphatase activity was measured with MUP as the substrate and is represented as relative fluorescence units (RFU) per min (average of duplicate samples ± standard deviation).

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cording to our results, phosphatase activity is not be associated with virulence, a role for this activity in bacterial survival during infection cannot be ruled out.

Acknowledgements. This study was funded by grant AGL2010-15232 from the Ministerio de Economía y Competitividad of the Spanish Government. We thank Ayub Darji, from CReSA, for critically reviewing this manuscript.

Competing interest: None declared.

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References

Fig. 4. Effect of molybdate on the phosphatase activity found in (A) the supernatant of Haemophilus parasuis ER-6P and (B) the supernatant from clone E. coli BL21 pGem-T Easy aphA FL3-1#19. Phosphatase activity is expressed as relative fluorescence units (RFU) per min (average of duplicate samples ± standard deviation). Sodium molybdate was used at different concentrations up to 10 mM.

The presence of the aphA gene in strains from different clinical backgrounds, including non-virulent nasal isolates, indicated that this enzyme was probably not a virulence factor. In addition, we did not detect any effect on the activation/inhibition of alveolar macrophages (cell-surface CD163, SLAI, SLAII, sialoadhesin and SWC3) after incubation of the cells with supernatant containing the AphA activity (not shown).

In summary, the different optimal pH and the different EDTA and molybdate susceptibilities between Apha and the activity found in H. parasuis supernatant suggested that Apha was not the activity found in the H. parasuis supernatant.

These results are consistent with the periplasmic location of AphA, as described in E. coli [14], or with the secretion of the Apha enzyme into the supernatant together with another, as yet unidentified phosphatase activity. Further experiments are required to identify the enzyme responsible for the phosphatase activity found in H. parasuis supernatants. Although ac-

1. Aragon V, Cerdà-Cuéllar M, Fraile L, Mombarg M, Nofrarias M, Olvera A, Sibila M, Solanes D, Segales J (2010) Correlation between clinico-pathological outcome and typing of Haemophilus parasuis field strains. Vet Microbiol 142:387-393 2. Aragon V, Kurtz S, Cianciotto NP (2001) Legionella pneumophila major acid phosphatase and its role in intracellular infection. Infect Immun 69:177-185 3. Bouchet B, Vanier G, Jacques M, Gottschalk M (2008) Interactions of Haemophilus parasuis and its LOS with porcine brain microvascular endothelial cells. Vet Res 39:42 4. Cerdà-Cuéllar M, Aragon V (2008) Serum-resistance in Haemophilus parasuis is associated with systemic disease in swine. Vet J 175:384-389 5. Costa-Hurtado M, Aragon V (2013) Advances in the quest for virulence factors of Haemophilus parasuis. Vet J 198:571-576 6. du Plessis EM, Theron J, Joubert L, Lotter T, Watson TG (2002) Characterization of a phosphatase secreted by Staphylococcus aureus strain 154, a new member of the bacterial class C family of nonspecific acid phosphatases. Syst Appl Microbiol 25:21-30 7. Fleischmann RD, Adams MD, White O, Clayton RA, Kirkness EF, Kerlavage AR, Bult CJ, Tomb JF, Dougherty BA, Merrick JM (1995) Wholegenome random sequencing and assembly of Haemophilus influenzae Rd. Science 269:496-512 8. Hoopman TC, Wang W, Brautigam CA, Sedillo JL, Reilly TJ, Hansen EJ (2008) Moraxella catarrhalis synthesizes an autotransporter that is an acid phosphatase. J Bacteriol 190:1459-1472 9. Jungnitz H, West NP, Walker MJ, Chhatwal GS, Guzmán CA (1998) A second two-component regulatory system of Bordetella bronchiseptica required for bacterial resistance to oxidative stress, production of acid phosphatase, and in vivo persistence. Infect Immun 66:4640-4650 10. Kielstein P, Rapp-Gabrielson VJ (1992) Designation of 15 serovars of Haemophilus parasuis on the basis of immunodiffusion using heat-stable antigen extracts. J Clin Microbiol 30:862-865 11. Leone R, Cappelletti E, Benvenuti M, Lentini G, Thaller MC, Mangani S (2008) Structural insights into the catalytic mechanism of the bacterial class B phosphatase AphA belonging to the DDDD superfamily of phosphohydrolases. J Mol Biol 384:478-488 12. Olvera A, Calsamiglia M, Aragon V (2006) Genotypic diversity of Haemophilus parasuis field strains. Appl Environ Microbiol 72:3984-3992 13. Omene JA, Glew RH, Baig HA, Robinson DB, Brock W, Chambers JP (1981) Determination of serum acid and alkaline phosphatase using 4-methylumbelliferyl phosphate. Afr J Med Med Sci 10:9-18 14. Passariello C, Forleo C, Micheli V, Schippa S, Leone R, Mangani S, Thaller MC, Rossolini GM (2006) Biochemical characterization of the class B acid phosphatase (AphA) of Escherichia coli MG1655. Biochim Biophys Acta 1764:13-19

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15. Rapp-Gabrielson VJ, Gabrielson DA, Schamber GJ (1992) Comparative virulence of Haemophilus parasuis serovars 1 to 7 in guinea pigs. Am J Vet Res 53:987-994 16. Reilly TJ, Baron GS, Nano FE, Kuhlenschmidt MS (1996). Characterization and sequencing of a respiratory burst-inhibiting acid phosphatase from Francisella tularensis. J Biol Chem 271:10973-10983 17. Robinson D, Willcox P. 1969. 4-Methylumbelliferyl phosphate as a substrate for lysosomal acid phosphatase. Biochim Biophys Acta 191:183-186 18. Rossolini GM, Schippa S, Riccio ML, Berlutti F, Macaskie LE, Thaller MC (1998) Bacterial nonspecific acid phosphohydrolases: physiology, evolution and use as tools in microbial biotechnology. Cell Mol Life Sci 54:833-850 19. Saha AK, Dowling JN, LaMarco KL, Das S, Remaley AT, Olomu N, Pope MT, Glew RH (1985) Properties of an acid phosphatase from Legio­nella micdadei which blocks superoxide anion production by human neutrophils. Arch Biochem Biophys 243:150-160

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20. Saleh MT, Belisle JT (2000) Secretion of an acid phosphatase (SapM) by Mycobacterium tuberculosis that is similar to eukaryotic acid phosphatases. J Bacteriol 182:6850-6853 21. Thaller MC, Lombardi G, Berlutti F, Schippa S, Rossolini GM (1995) Cloning and characterization of the NapA acid phosphatase/phosphotransferase of Morganella morganii: identification of a new family of bacterial acid-phosphatase-encoding genes. Microbiology 141:147-154 22. Thaller MC, Schippa S, Bonci A, Cresti S, Rossolini GM (1997) Identification of the gene (aphA) encoding the class B acid phosphatase/phosphotransferase of Escherichia coli MG1655 and characterization of its product. FEMS Microbiol Lett 146:191-198 23. Thaller MC, Schippa S, Rossolini GM (1998) Conserved sequence motifs among bacterial, eukaryotic, and archaeal phosphatases that define a new phosphohydrolase superfamily. Protein Sci 7:1647-1652

RESEARCH ARTICLE International Microbiology (2014) 17:149-157 doi:10.2436/20.1501.01.217. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Molecular epidemiology of methicillin-resistant Staphylococcus aureus in a university hospital in northwestern Spain Jaime Ariza-Miguel,1 Marta Hernández,1 Isabel Fernández-Natal,2,3 David Rodríguez-Lázaro1,4* Institute of Agricultural Technology of Castilla y León, Valladolid, Spain. 2University Hospital of Leon, Leon, Spain.

3 Institute of Biomedicine (IBIOMED), University of Leon, Leon, Spain.

4 Microbiology Section, Faculty of Sciences, University of Burgos, Burgos, Spain

1

Received 10 April 2014 · Accepted 30 September 2014 Summary. Continuous monitoring of methicillin-resistant Staphylococcus aureus (MRSA) is necessary to understand the clonal evolution of successful lineages. In this study, we identified the MRSA clones circulating in a Spanish hospital during a 2-year period, assessed their relationship with antimicrobial resistance profiles, and investigated the presence of the emerging community-associated and livestock-associated MRSA lineages (CA-MRSA, LA-MRSA). CC5-MRSA-IV isolates were the most frequently recovered, which supports the previously reported prevalence of this clone in Spanish hospitals. We observed ST125 isolates that harbored specific cassette chromosome recombinase (ccr) gene elements of the staphylococcal cassette chromosome mec (SCCmec) types IV and VI. That clone, which was first detected only recently, has increased resistance to erythromycin. Furthermore, 94% of the infections were caused by non-multiresistant isolates. Neither CA-MRSA nor LA-MRSA isolates were observed. These findings, along with related events over the last decade, suggest the establishment of a clonal endemic population in the Spanish clinical environment. [Int Microbiol 2014; 17(3):149-157] Keywords: methicillin-resistant Staphylococcus aureus (MRSA) · clonal population · molecular epidemiology · multilocus sequence typing

Introduction Staphylococcus aureus is an opportunistic pathogen that causes a wide range of diseases in humans, from mild skin infections to life-threatening diseases such as toxic shock syndrome, septicemia, endocarditis, and necrotizing pneumonia.

Methicillin-resistant Staphylococcus aureus (MRSA) was * Corresponding author: D. Rodríguez-Lázaro Microbiology Section. Faculty of Sciences University of Burgos Pl. Misael Bañuelos, s/n 09001 Burgos, Spain Tel. +34-637451100 E-mail: rodlazda@gmail.com

first described in 1960, a year after the introduction of methicillin into clinical practice to treat infections caused by penicillin-resistant strains of S. aureus [11]. Methicillin resistance is mediated by the presence of the mecA gene, which encodes an additional penicillin-binding protein (PBP2a or PBP2′) with low affinity for β-lactam antibiotics [15]. More recently, a novel methicillin-resistance determinant, mecALGA251 or mecC, has been described that has 70% homology at the DNA level with mecA [9]. Those genes are harbored in a highly diverse mobile genetic element, the staphylococcal cassette chromosome mec (SCCmec), whose transmission to ecologically successful methicillin-susceptible S. aureus strains (MSSA) led to the emergence of the ancestral MRSA lineages [12]. Up to 11 SCCmec types, defined by combining the ge-

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netic structures of the cassette chromosome recombinase (ccr) gene complex and the mec gene complex, have been described [www.sccmec.org/Pages/SCC_TypesEN.html]. Thus, MRSA isolates are represented not only by their genetic backgrounds, but also by the genetic structure of their SCCmec elements [8].

MRSA lineages have become a truly global health concern over the last few decades, as their worldwide spread has occurred very quickly. The prevalence of MRSA greatly varies among countries, provinces and hospitals but it has increased significantly since its appearance in 1960. In the USA, MRSA infections increased from 4% in the 1980s to 50% of all S. aureus infections in the late 1990s [24]. In Spain, MRSA infections increased from 1.5% in 1986 to 31.2% in 2002 [34]. In the worst cases, such as in Taiwan, in 2007 MRSA accounted for 80% of all S. aureus isolates causing nosocomial infections [36]. However, the scenario has become even more complex. Until the mid 1990s the presence of MRSA was restricted to the clinical environment (hospital-acquired MRSA, HA-MRSA), but since then MRSA strains have been recovered increasingly from the community (community-acquired MRSA, CA-MRSA) [33]. In contrast to HA-MRSA, CA-MRSA strains are frequently isolated from children and young people without previous health-care contact and they show specific genetic and phenotypic traits: they have a different genetic background, harbor a smaller SCCmec type IV or V, which provides them with a selective advantage in terms of faster replication times, present more virulence mediated by factors such as the Panton-Valentine leukocidin cytotoxin (PVL), and usually have a non-multiresistant antimicrobial profile [26].

The recurrent penetration of CA-MRSA into the clinical environment has blurred the boundary between HA-MRSA and CA-MRSA, making it more difficult to discriminate between them. In addition, a livestock-associated MRSA (LAMRSA) has been described. LA-MRSA strains may act as genetic reservoirs of resistance and could play a major role in the adaptive evolution of MRSA. In fact, the recently described methicillin resistance gene mecC was originally detected in LA-MRSA strains but is now increasingly recovered from clinical settings [9,27]. Continuous molecular epidemiologic surveillance of MRSA is imperative to trace the evolution and spread of successful MRSA clones, both for therapeutic reasons and for the implementation of initiatives aimed at controlling and preventing MRSA infections. Here we describe a molecular epidemiology study of MRSA isolates recovered from the University Hospital of Leon from 2007 to 2008. The genetic backgrounds of these isolates were charac-

ARIZA-MIGUEL ET AL.

terized by pulsed-field gel electrophoresis (PFGE) and multilocus sequence typing (MLST), and the structures of the respective SCCmec elements were determined. Furthermore, we investigated both the antimicrobial resistance profiles of CA-MRSA and the presence of their PVL genes, as markers.

By identifiying the MRSA clones circulating in the hospital, this study contributes to the knowledge of the MRSA lineages circulating in Spanish health-care setting.

Materials and methods Clinical information and bacterial isolates. Seventy Staphylo­ coccus aureus isolates recovered from patients at the University Hospital of Leon (Leon, northwestrn Spain), between 2007 and 2008 were determined to be MRSA based on antimicrobial susceptibility testing. The University Hospital of Leon is a 795-bed public teaching hospital. Forty-nine isolates were recovered from blood cultures (one of them was recovered also from bile); 14 from central venous catheter blood samples; 3 from peripheral vascular catheter blood samples; 1 from packed red blood cells; 1 from a drainage sample; 1 from synovial joint fluid; and 1 from ascites. The specimens were isolated and identified in conventional medium (mannitol salt agar, blood and chocolate agar plates) and commercial MicroScan microdilution panels (Siemens Diagnostics, Munich, Germany) and by real-time PCR [32] in a 7500 realtime PCR system platform (Applied Biosystems, Carlsbad, CA, USA). Methicillin resistance was confirmed in 68 of the 70 isolates by using internal amplification controls designed within the mecA gene [13,22]. Isolates SA13 and SA65, which were MRSA as revealed by antimicrobial susceptibility testing, tested negative for the presence of mecA. Repetition of the susceptibility tests confirmed both isolates as MSSA. Antimicrobial susceptibility testing. Antimicrobial susceptibility testing was performed by the microdilution method following the recommendations and minimal inhibitory concentration (MIC) breakpoints of the Clinical and Laboratory Standars Institute (CLSI) guidelines (2008). Susceptibility to the following 15 antimicrobial agents was tested: ampicillin, penicillin, oxacillin, amoxicillin/clavulanate, cefotaxime, erythromycin, clindamycin, teicoplanin, vancomycin, ciprofloxacin, imipenem, gentamicin, rifampicin, tetracycline, and cotrimoxazole. MRSA isolates were clustered in resistance profiles (RPs) according to their susceptibility to 10 antimicrobials: ampicillin, penicillin, erythromycin, teicoplanin, vancomycin, ciprofloxacin, imipenem, rifampicin, tetracycline, and cotrimoxazole. Isolates resistant to three or more antibiotics, in addition to β-lactams, were considered to be multiresistant.

Characterization of the genetic background. Genetic characterization of all S. aureus isolates was carried out by pulse-field gel electrophoresis (PFGE) and multilocus sequence typing (MLST). PFGE was performed following the method McDougal et al. [16] modified as follows: bacterial cells were suspended in PIV buffer (10 mM Tris-HCl, 1 M NaCl, pH 8) to an absorbance of 0.9–1.1 at 610 nm. Then, 200 µl of the adjusted culture was washed and resuspended in 300 µl of the same buffer. After the lysis step, the plugs were transferred into a new tube containing 1 ml ESP buffer (1% N-lauroyl sarcosine in 0.5 M EDTA pH 8, 100 µg proteinase K/ml) and incubated at 56ºC for 16–20 h. DNA fragment sizes were determined by comparing bands with a Lambda Ladder PFG Marker (New England Biolabs). PFGE profiles were analyzed using Bionumerics v.6.6 (Applied-Maths NV, SintMartens-Latem, Belgium) to describe genetic relationships among isolates.

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Dendograms were constructed using the Dice similarity coefficient and the unweighted pair group mathematical average (UPGMA) clustering algorithm. MLST was carried out as previously described [8]. DNA isolation was performed using the commercial kit QIAamp DNA mini kit (QIAGEN, Hilden, Germany) following the manufacturer’s instructions. To guarantee analysis of identical genetic material by PFGE and MLST, DNA from the same culture was used for both methods. Allelic profiles obtained by MLST were assigned by comparing the consensus sequences with the information already published in the S. aureus MLST database hosted at [http://saureus. mlst.net/]. Thereby, genotypes were constituted by seven numbers representing the unique allelic profiles at seven housekeeping genes. Data regarding the 70 isolates analyzed, as well as the novel allelic profiles were submitted to the MLST database. Allelic profiles were used to compare the S. aureus isolates recovered from our hospital with those previously analyzed from other geographic areas. Simpson’s index of diversity, which measures the probability that two unrelated strains sampled from the test population will be placed into different typing groups [10], was calculated to compare the discriminative power of PFGE typing versus MLST and for measuring genetic diversity among isolates using the comparing partitions website [http://darwin.phyloviz.net/ComparingPartitions/index.php?link=Home]. The adjusted Wallace coefficient for quantification of the agreement between PFGE typing and MLST was calculated on that website.

Typing and subtyping of the SCCmec element. The genetic structure of the SCCmec element was determined by multiplex-PCR carried out on a Veriti 96 Well thermal cycler and GeneAmp PCR system 9700 (Applied Biosystems) as described elsewhere [13,22]. These two molecular methods allowed discrimination of SCCmec types I, II, III, IV, V and VI, as well as the variants IA & IIIA. SCCmec IV was further subtyped into eight different subtypes, from IVa to IVh, by multiplex-PCR as previously described [18].

Detection of Panton-Valentine leukocidin virulence factors. The presence of the PVL genes (lukS-PV & lukF-PV) was investigated by conventional PCR as previously described [14]. Reference strain ATCC 49775 was used as the positive control.

Results Antimicrobial susceptibility testing. All 68 MRSA isolates were resistant to the β-lactam antibiotics tested (oxacillin, ampicillin, penicillin, cefotaxime and imipenem) as well

as to amoxicillin/clavulanate and ciprofloxacin. Furthermore, 42.6% of the isolates were resistant to erythromycin, 21% were resistant to clindamycin, 15.9% were resistant to gentamicin, 4.4% to rifampicin and 1.5% to both tetracycline and cotrimoxazole. Of the isolates resistant to erythromycin, 34.8% had the M phenotype. All isolates were susceptible to the glycopeptide antimicrobial agents vancomycin and teicoplanin. The isolates were clustered into five RPs, with all the isolates resistant to the β-lactam antibiotics tested and to at least one more antimicrobial agent (Table 1). Notably, the percentage of MRSA resistant to gentamicin increased from 5.7% in 2007 to 28.6% in 2008. Resistance to erythromycin, rifampicin, tetracycline, and cotrimoxazole also increased during the study period albeit to a lesser extent, while resistance to clindamycin decreased from 23.5% in 2007 to 17.9% in 2008 (Table 2).

Genetic background. Genetic characterization of all 70 S. aureus isolates by PFGE with the restriction enzyme SmaI revealed 27 different pulsotypes (25 pulsotypes among the 68 MRSA isolates), resulting in a Simpson’s index of diversity of 0.909, which indicated that, if two isolates were sampled randomly from the population, on 90.9% of the occasions they would fall into different types. Genotypes 19 and 26 accounted for 40% of the isolates (17.1% and 22.9%, respectively). A dendrogram constructed using the Dice similarity coefficient with the UPGMA clustering algorithm revealed four major clusters (arbitrarily designated from A to D) with a cutoff of 90% similarity. More than 81% of the isolates were grouped into these four genetic clusters. Genotypes that did not belong to these clusters were designed as “sporadic.” Cluster A comprised 18 ST5-MRSA-IV isolates, two novel sequence types (STs) derived from ST-5, and one ST125-MRSA-IV. Cluster B comprised ten ST125-MRSA-IV and one ST5-MRSA-IV.

Table 1. Resistance profiles of 68 methicillin-resistant Staphylococcus aureus isolates at the University Hospital of Leon, 2007 and 2008 Resistance profile

Antimicrobial agenta

% MRSA clones

RP0

Only to β-lactams

0

None

RP1

CIP

55.2

ST5-IV(10); ST8-IV(14); ST125-IV(12); ST125-IV/VI(2)

RP2

ERY, CIP

38.8

ST5-IV(10); ST8-IV(1); ST125-IV(4); ST125-IV/VI(8); ST228-I(1); ST2755-IV(1); ST2756-IV(1)

RP3

ERY, CIP, RIF

3

ST5-IV(2)

RP4

ERY, CIP, COT

1.5

ST8-IV(1)

RP5

CIP, RIF, TET

1.5

ST5-IV(1)

All isolates were resistant to the β-lactam antibiotics tested: oxacillin, ampicillin, penicillin, cefotaxime and imipenem). CIP, ciprofloxacin; ERY, erythromycin; RIF, rifampicin; COT, cotrimoxazole; TET, tetracycline.

a

151

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Table 2. Evolution of the resistance to antimicrobials by 68 methicillin-resistant Staphylococcus aureus isolates, University Hospital of Leon, 2007 and 2008 Antibioticsa

2007b

2008b

∆(2008–2007)c

Erythromycin

41.7

43.8

+2.1

Clindamicyn

23.5

17.9

–5.6

Gentamycin

5.7

28.6

+22.9

Rifampicin

2.8

6.3

+3.5

Tetracicline

0

3.1

+3.1

Cotrimoxazole

0

3.1

+3.1

All MRSA isolates were resistant to the β-lactam antibiotics tested: oxacillin, ampicillin, penicillin, cefotaxime, and imipenem.

They were also resistant to amoxicillin/clavulanate and ciprofloxacin, and were susceptible to vancomycin and teicoplanin. b Percentage of the mean values of antibiotic resistance. c Differences between the percentages of the mean values of antibiotic resistance in 2007 and 2008. a

Cluster C grouped nine ST125-MRSA-IV/VI isolates, and cluster D all sixteen ST8-MRSA-IV isolates (Fig. 1).

Overall, among the 68 MRSA isolates, six STs were determined by MLST analysis. Three of the six accounted for 95.6% of the isolates: ST-125 (38.2%), ST-5 (33.8%), and ST-8 (23.5 %). ST-228 and two novel STs were also detected, in one isolate each. Detected clones belonged to the clonal complexes CC5 (76.5 %) and CC8 (23.5 %). After submission to the S. aureus MLST database [http://saureus.mlst.net/], novel STs were designated as ST-2755 and ST-2756 (allelic profiles 314–4–1–4–12–1–10 and 1–4–1–4–12–1–320 at arcC, aroE, glpF, gmk, pta, tpi, and yqiL, respectively). The novel STs were very similar to ST-5: ST-2755 had a unique difference of a single nucleotide polymorphism (SNP) in nucleotide 39 of arcC, showing 99% similarity with arcC allele 1. However, this difference at the DNA level was not translated into the functional level since both codons encoded the amino acid threonine. In addition, ST-2756 had a SNP in nucleotide 256 of allele 10 of yqiL (99% similarity), where thymine is replaced by guanine. This change leads to a non synonymous substitution in which cysteine is substituted by glycine in the encoded protein. MSSA isolates SA 13 and SA 65 belonged to ST-15 and ST-5, respectively. In 2007, the most frequently recovered isolates belonged to ST-5 and ST-125 (38.9% each one), followed by ST-8 (19.4%). ST-2756 was recovered just one isolate. In 2008, isolates showing ST-125 were the most frequently isolated (37.5%), followed by those of ST-5 and ST-8 (28.1% each). ST-228 and ST-2755 were also sporadically observed in one isolate.

Genetic structure of the SCCmec element. The genetic structure of the SCCmec element was investigated in all S. aureus isolates. During the 2-year study period, 67 out of

68 MRSA isolates (98.5%) harbored SCCmec type IV; the remaining isolate (SA16) harbored SCCmec type I (1.5%).

Further subtyping of SCCmec IV revealed that all belonged to subtypes IVc/IVe. Note that, 38.5% of the ST125-MRSAIV isolates (14.9% of all isolates harboring SCCmec IV) also had specific ccr gene elements of SCCmec type VI (mecA gene complex class B and ccr gene complexes type 2 and 4; A2B2 and A4B4, respectively) and consequently were designated as ST125-MRSA-IV/VI. Furthermore, 37 out of the 67 (55.2%) SCCmec type IV elements contained an additional 381-bp DNA fragment according to the system previously described by Oliveira and Lencastre [22], corresponding to the presence of the kanamycin/neomycin/bleomycin resistance plasmid pUB110. The evolution of the MRSA clones circulating at our hospital over the 2-year study period are shown in Fig. 2.

Congruence between PFGE-typing and MLST. PFGE better discriminated among the 70 S. aureus isolates than MLST (Simpson’s index of diversity of 0.909 and. 0.701, respectively). The adjusted Wallace coefficient PFGE→MLVA was 0.883, with a 95% confidence interval (CI) of 0.765–1.000; the coefficient for MLVA→PFGE was 0.208 (CI 95%; 0.137– 0.279). These values implied that if two isolates were in the same pulsotype, they had an 88.3% chance of having the same MLST genotype; conversely, sharing the same MLST type was associated with only a 20.8% chance of having the same pulsotype. Two discordant results (group violations) were obtained

by PFGE typing. Isolate SA 45, belonging to ST-125, clustered closely with isolates showing ST-5. Conversely, isolate SA 29, which belonged to ST-5, was clustered in a branch closely related to other isolates from ST-125 (Fig. 1). The molecular typing results are summarized in Table 3.

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Fig. 1. Genetic relationships among 70 Staphylococcus aureus isolates based upon comparisons of the pulsed-field gel electrophoresis profiles obtained with the restriction enzyme SmaI. The dendrogram was produced using a Dice similarity coefficient matrix with the unweighted pair group method with arithmetic mean (UPGMA). Major clusters with a cutoff of 90 % similarity were arbitrarily designated from A to D. Scale bar indicates similarity values.

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Fig. 2. Clonal evolution of 68 methicillin-resistant Staphylococcus aureus isolates recovered from 2007 to 2008 at the University Hospital of Leon.

Panton-Valentine leukocidin genes. All 70 S. au­ reus isolates tested negative for the presence of the PVL genes (lukS-PV and lukF-PV). Type strain ATCC 49775, included as a positive control, gave the expected 433-bp DNA fragment in all PCRs.

Discussion In 2006, MRSA in Spain accounted for 30% of all S. aureus clinical isolates [5]. Until 1995, ST247-MRSA-I (classically known as the ‘Iberian clone’) had been the most frequently observed type in the clinical environment. However, this clone has been gradually replaced by others, and nowadays, ST5-MRSA-IV and ST125-MRSA-IV are the predominant clones in Spanish hospitals [28,34,35]. The results of our study are in agreement with that trend: in 2007�������������� –������������� 2008, the recovery rate of MRSA at the University Hospital of Leon reached 38.7%. Overall, ST5-MRSA-IV was the most frequently detected clone (33.8%), followed by ST125-MRSAIV (23.5%), ST8-MRSA-IV (23.5%), and ST125-MRSA-IV/ VI (14.7%). Moreover, ST228-MRSA-I and two novel STs were sporadically recovered, accounting for one isolate each (1.5%). ST125 isolates have rarely been recovered worldwide, according to the S. aureus MLST database [http://saureus.mlst.net]. In fact, only 7 out of the 5024 isolates available

from that database belong to ST125 (0.14%, on 08/27/2013).

Four of the isolates occurred in Spain, and the other three in Norway, Finland, and France, respectively. The reason for the apparent evolutionary success of clone ST125-MRSA-IV in Spanish hospitals is unclear, nor is it understood why that clone has not spread to other European countries.

Previous studies had reported the gradual replacement in the clinical setting of multiresistant SCCmec elements—classically found in HA-MRSA isolates—by SCCmec IV, which was initially detected in CA-MRSA isolates and harbors no further resistance elements [17,20,23,28,30,34,35]. The genetic structure of the SCCmec elements analyzed in this study supports that finding, since 98.5% of the isolates in this study harbored SCCmec IV (14.9% of them also harbored specific ccr gene elements of SCCmec type VI). The successful introduction and persistence of the genetically shorter SCCmec IV in clinical settings might have provided MRSA strains with an evolutionary advantage in terms of faster replication times, allowing them to outcompete the previously prevalent HAMRSA clones harboring multiresistant SCCmec types [21].

Further subtyping of SCCmec IV revealed that all of them belonged to subtype IVc/IVe. Previous nationwide studies reported that around 70% of all MRSA isolates harboring SCCmec IV belong to subtype IVa [17,28,35]. However, our findings are in agreement with those published by Argudín et al. [2], in which 88.5% of MRSA isolates carry subtype IVc. Note that the study

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Table 3. Correlation between the genotyping methods used in 70 Staphylococcus aureus isolates (68 MRSA), at the University of Leon Hospital, 2007 and 2008 CC(n)

ST(n)

Allelic profile

CC5(54)

5(24)

1-4-1-4-12-1-10

125(26)

CC8(16)

1-4-1-4-12-1-54

PFGE cluster(n)

SCCmec type(n)

A(18)

IV(18)

B(1)

IV(1)

Sporadica(5)

IV(4); Noneb(1)

A(1)

IV(1)

B(10)

IV(10)

C(9)

IV/VI(9)

Sporadic(6)

IV(5); IV/VI(1)

MRSA clone(n) ST5-MRSA-IV(23); ST5-MSSA(1)

ST125-MRSA-IV(16); ST125-MRSA-IV/VI(10)

15(1)

13-13-1-1-12-11-13

Sporadic(1)

Noneb(1)

ST15-MSSA(1)

228(1)

1-4-1-4-12-24-29

Sporadic(1)

I(1)

ST228-MRSA-I(1)

2755(1)c

314-4-1-4-12-1-10

A(1)

IV(1)

ST2755-MRSA-IV(1)

2756(1)c

1-4-1-4-12-1-320

A(1)

IV(1)

ST2756-MRSA-IV(1)

8(16)

3-3-1-1-4-4-3

D(16)

IV(16)

ST8-MRSA-IV(16)

CC, clonal complex. ST, sequence type. PFGE, pulse-field gel electrophoresis. SCCmec, staphylococcal cassette chromosome mec. MLST, multilocus sequence typing. a Pulsotypes that did not belong to the main clusters were designed as “sporadic.” b Methicillin-susceptible S. aureus did not harbor a SCCmec element. c Novel genotype as determined by MLST.

of Argudín et al. was performed in a hospital located in Asturias, the neighbor region of León. Further research is necessary to clarify whether, in Spain, there is a correlation between SCCmec types and their geographical origin.

The successful establishment of SCCmec IV in the clinical environment has led to a global change in MRSA resistance profiles. Many studies have reported the gradual replacement of the classical multiresistant HA-MRSA clones by non-multiresistant isolates harboring SCCmec IV [1,2,6,25,31,33].

Accordingly, in this work 94% of MRSA isolates were nonmultiresistant, as they were resistant to only one or two antibiotics in addition to β-lactams (RP1 and RP2) (Table 1). This scenario is in agreement with the presence of the prevalent lineage CC5 harboring SCCmec IV, which carries no further resistance elements. Moreover, all the isolates were susceptible to glycopeptides. The increase in gentamicin resistance— from 5.7% in 2007 to 28.6% in 2008 (Table 2)—was not associated with any particular genotype.

The presence of CC5-MRSA strains harboring multiple or composite SCCmec elements has been previously reported [19]. In this study, we found ST125 isolates that carried specific ccr gene elements of both SCCmec types IV and VI (38.5% of ST125 isolates and 14.9% of all isolates harboring SCCmec IV). This clone was recently detected in a third-level hospital

in Valladolid, a neighbor city of León [17]. Nine out of the ten isolates were grouped into cluster C by PFGE, and the remaining one yielded a sporadic pulsotype (Fig. 1). The recovery rate of ST125-MRSA-IV/VI increased from 5.6% in 2007 to 25% in 2008, while the presence of ST125-MRSA-IV decreased from 33.3% in 2007 to 12.4% in 2008 (Fig. 2). As previously observed by Menegotto et al. [17], in our study, 80% of ST125-MRSA-IV/VI isolates but just 25% of ST125MRSA-IV isolates were resistant to erythromycin (RP2).

These findings suggest that the clonal replacement observed might have occurred because of antibiotic selective pressure.

Two novel STs, designated ST-2755 and ST-2756, were detected in this study. Both genotypes were single-locus variants of ST5 belonging to CC5. ST-2755 had one SNP in nucleotide 39 of allele 1 of arcC, and ST-2756 one SNP in nucleotide 256 of allele 10 of yqiL. These STs might have emerged as punctual mutations from ST5-MRSA isolates since both are single-locus variants of ST5 and, to our knowledge, this genotype has not been observed in MSSA strains (or at least there are no S. aureus strains with those genotypes in the MLST database). Both strains were recovered from blood culture samples, were RP2, and were grouped by PFGE into cluster A, along with ST5 isolates. Moreover, neither CAMRSA nor LA-MRSA strains were detected in this study, and

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all 70 isolates tested negative for the presence of PVL genes.

According to previous studies carried out in Spanish clinical settings, it seems that, despite worldwide emergence of CAMRSA strains harboring PVL genes [33], this is still an atypical situation in Spain and is mainly restricted to the introduction of exotic strains from South America [3,4,29].

As expected, molecular typing by PFGE provided better discrimination than MLST (Simpson’s index of diversity of 0.909 vs. 0.701, respectively). We observed two group violations by PFGE (Fig. 1), which has been previously reported since these two typing methods index very different types of variations. PFGE-typing can detect genomic variations with a wide range of causes, and thus identify minor variations among isolates collected within a restricted geographic area (e.g., in one hospital). However, such rapidly evolving sequences are susceptible to homoplasy, which can lead to incorrect phylogenetic inferences. MLST indexes variations in seven housekeeping genes under stabilizing evolution, and is therefore more suitable for population studies.

Continuous monitoring of MRSA clones circulating in clinical settings is necessary to better understand the clonal evolution of successful MRSA lineages and to apply appropriate treatment, control, and prevention strategies aimed at hindering the dissemination of MRSA infections. Our molecular epidemiology study of MRSA isolates in a Spanish health-care institution identified CC5-MRSA-IV strains (ST5-MRSA-IV and ST125-MRSA-IV clones) as the most frequently recovered strains, which confirms the previously reported prevalence of these clones in the Spanish clinical environment.

Acknowledgments. This work was supported by EU 7th Framework Programme under the project PROMISE (project.265877). We thank Hortensia Rodríguez Pollán for her technical assistance, and Dr. Bruno González-Zorn for kindly providing MRSA strain ATCC 49775, used as a positive control for the presence of the PVL genes.

Competing interests. None declared.

References

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RESEARCH ARTICLE International Microbiology (2014) 17:159-164 doi:10.2436/20.1501.01.218. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Significance of tagI and mfd genes in the virulence of non-typeable Haemophilus influenzae Denis A. Spricigo,1 Pilar Cortés,1 David Moranta,2 Jordi Barbé,1 José Antonio Bengoechea,2,3 Montserrat Llagostera1* Department of Genetics and Microbiology, Faculty of Biosciences, Autonomous University of Barcelona, Bellaterra, Spain.

Network Biomedical Research. Respiratory Diseases and Health Research Foundation Balearic Island, Joan March Hospital, Bunyola, Spain. 3Centre for Infection and Immunity, Queens University Belfast. Belfast, UK

1 2

Received 29 April 2014 · Accepted 29 September 2014

Summary. Non-typeable Haemophilus influenzae (NTHi) is an opportunist pathogen well adapted to the human upper respiratory tract and responsible for many respiratory diseases. In the human airway, NTHi is exposed to pollutants, such as alkylating agents, that damage its DNA. In this study, we examined the significance of genes involved in the repair of DNA alkylation damage in NTHi virulence. Two knockout mutants, tagI and mfd, encoding N3methyladenine-DNA glycosylase I and the key protein involved in transcription-coupled repair, respectively, were constructed and their virulence in a BALB/c mice model was examined. This work shows that N3-methyladenine-DNA glycosylase I is constitutively expressed in NTHi and that it is relevant for its virulence. [Int Microbiol 2014; 17(3):159-164] Keywords: Haemophilus influenzae · alkylating agents · virulence · genes tagI and mfd

Introduction Non-typeable Haemophilus influenzae (NTHi) is a commensal gram-negative bacterium well adapted to the human upper respiratory tract [7]. It has been implicated in the etiology of otitis media, conjunctivitis, sinusitis, pneumonia, and chronic bronchitis, and in the progression of chronic obstructive pulmonary disease (COPD) [19]. However, within its human host, this opportunistic pathogen is exposed to high levels of genotoxic stress in the form of airway pollutants. In a study Corresponding author: M. Llagostera Department of Genetics and Microbiology Faculty of Biosciences Autonomous University of Barcelona 08193 Bellaterra, Spain Tel. +34-935812615. Fax 34-935812387 E-mail: Montserrat.Llagostera@uab.cat *

based on proteomic expression profiling of H. influenzae grown in pooled sputum from adults with COPD, both the expression of antioxidant activity and stress responses were shown to be important for NTHi survival in the airways [13].

DNA-damaging agents are ubiquitous. They are generated endogenously during cell metabolism and are present in the environment—in air, water and foods—although generally in low concentrations. For example, tobacco smoke contains a mixture of alkylating agents, some of which act directly (alkyl halides, acrolein, crotonaldehyde, ethylene oxide, propylene oxide, acrylonitrile, and acrylamide), while others act indirectly (requiring metabolic transformation to form reactive species) [15]. Moreover, human airway pollutants such as tobacco smoke damage not only eukaryotic cells but also the DNA of the respiratory tract microbiota.

The repair of DNA alkylation damage in bacterial cells has been mainly studied in Escherichia coli. As in other

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bacteria, E. coli has two specific mechanisms to remove alkyl radicals from its DNA: (i) via the constitutive expression of genes encoding the necessary repair enzymes and (ii) via the alkyl-induced expression of these proteins [16]. This adaptive response to the repair of DNA alkylation damage is regulated by the Ada protein, a positive transcriptional regulator that stimulates the expression of the ada, alkA, alkB, and aidB genes [5,16]. Bacteria also have two additional enzymes involved in the specific repair of DNA alkylation damage: Ogt (O6-meG-DNA methyltransferase) [10] and TagI (N3meADNA glycosylase I) [2]. In addition, two other systems are involved in the repair of DNA alkylation damage: the nucleotide excision repair (NER) [20] and the transcriptioncoupled repair (TCR) [17] systems. The latter system mediates the bulk repair of DNA damage via the Mfd protein, followed by the engagement of NER. The aim of the present work was to determine the significance of tagI and mfd genes involved in the repair of DNA alkylation damage in NTHi virulence. Accordingly, knockout mutants in tagI, specific for DNA alkylation damage,

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and mfd, involved in bulk DNA repair, were constructed and their virulence in a BALB/c mouse model was studied.

Materials and methods Bacteria, media, and growth conditions. Haemophilus influenzae NTHi375, an otitis media isolate [4], was grown on chocolate agar + PolyViteX plates (PVX; BioMerieux), on brain heart infusion (BHI) medium with or without agar supplemented with 10 µg hemin ml–1, and 10 µg NAD ml–1 (sBHI). The cultures were grown at 37 °C for 18 h in an atmosphere of 5 % CO2. Escherichia coli DY380 strain was grown in LB (Luria–Bertani) broth or on agar plates at 37 °C for 18 h. When necessary, 50 µg ampicillin ml–1 and 50 µg spectinomycin ml–1 were added.

Construction of tagI and mfd knockout mutants. The tagI knockout mutant was constructed from strain NTHi375 using a previously described method [18]. Briefly, the entire gene targeted for deletion was PCR-amplified from the genomic DNA of NTHi375 strain (Table 1, Fig. 1A), cloned into pGEM-T (Promega), and electroporated into E. coli DY380.

Strain DY380 harboring the plasmid with the tagI gene was selected by plating onto LB agar plates supplemented with 50 µg ampicillin ml–1. Then, with plasmid pRSM2832 [18] as template, PCR was used to generate an amplicon

Table 1. Oligonucleotide primers used in this study Primer

Sequence (5′-3′)

Primers used to obtain the mutants TagI_F

cggtgtcgcagcaatca

TagI_R

tctgtgaaagccttatgtgaactc

Mfd_F

tacactatgcctcaattttacaca

Mfd_R

acaatgatcgggcttctttttatg

P1-TagI

ggttggcgaacaatctatttatattgattatcatgacaaggaatggggaaagcctgaattcgacagccaaaagctatttgattccggggatccgtcgacc

P2-TagI

aatcatttaaatgatcgtccaccagccccatagattgcataaacgcatagcacgtggtttcgccaataaagacgaaaccatgtaggctggagctgcttcg

P1-Mfd

cattttaaaggaaatgtactgttttcggtggagacggaaggtcgccgagagactttgcttgatttgctttcaccgttaaaattccggggatccgtcgacc

P2-Mfd

tattaagcgttcgaggaattggcgttgccgttagcgtaagaatatcgatattcgcacgaagctgtttgattttctctttttgtaggctggagctgcttcg

Primers used for RT-qPCR assays trpA F

cttcgtgccgttcgttacc

trpA R

tgaccgcactttttccaatagt

Tag F

cgccaaataagctttcgcat

Tag R

cggggctttcgtggatta

recA F

cagtgcggcaacggagtc

recA R

cgcaaaaagcaggaaaaacc

Forward primer, up; reverse primer, dw. Underlined text corresponds to the 80 nucleotides of the 5′ and 3′ ends (H1 and H2) of the NTHi gene to be deleted.

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Fig. 1. Construction of tagI mutant. (A) The PCR-amplified tagI gene from NTHi375 was cloned into pGEMT and electroporated into E. coli DY380. (B) An amplicon containing a cassette with both the rpsL and the spectinomycin resistance genes, flanked by each of the FRT regions, was obtained by PCR of pRSM2832 with primers P1-TagI and P2-TagI (see Table 1). Homology arms H1 and H2 are 80-nucleotide (nt) sequences of the 5′ and 3′ ends of the tagI gene to be deleted, and P1 and P2 are 20-nt sequences of DNA homologous to the 5′ and 3′ ends of the cassette, respectively. (C) The inserted region of the amplicon of panel B in the chromosome of the NTHi375 knockout tagI mutant.

containing a cassette with both the rpsL and the spectinomycin resistance genes, flanked by FRT (FLP recombinase target) sites. In addition, the design of the primers produced an amplicon that contained 80 nucleotides (nt) of the 5′ and 3′ ends of the tagI gene to be deleted, flanking each of the FRT regions (Table 1, Fig. 1B). The amplicon was electroporated into E. coli strain DY380 harboring pGEM-T carrying the tagI gene. After induction of the recombinase genes of strain DY380 by heat shock at 42°C, spectinomycin-resistant clones were isolated by plating the transfectants onto LB agar plates supplemented with 50 µg spectinomycin ml–1. One of these positive clones was chosen for further use after PCR and sequencing to confirm that it harbored a plasmid with the correct insertion. This plasmid was digested with NcoI and NsiI restriction enzymes, and the fragment with the appropriately sized insert was recovered and used to transform NTHi375 strain by the MIV method, as previously described [12], to obtain the desired construct (Fig. 1C). Spectinomycin-resistant clones were isolated by plating onto BHI agar plates supplemented with 200 µg spectinomycin ml-1 followed by incubation at 37°C. One of these clones was isolated for further use after PCR and DNA sequencing to confirm that it contained the desired mutation and that it did not harbor any remnants of the plasmid. The same procedure was used to obtain the mfd mutant. Afterwards, pRSM2947, a temperature-sensitive replicon appropriate for NTHi and harboring both the FLP recombinase under the control of the tet regulatory system and a kanamycin resistance marker, was transformed by electroporation into the knockout mutants, to remove the cassette containing both rpsL and the spectinomycin resistance genes.

NTHi infection BALB/c model. To infect the mice, the bacteria were recovered with 1 ml of PBS from a chocolate-agar plate grown for 16 h, yielding a bacterial suspension of ~5 × 109 colony-forming units (CFU)/ ml. Twenty microliters of bacteria (~107 CFU) were inoculated into the nares of 5- to 7-week-old female BALB/c mice (Harland Iberica). After 48 h of infection, the mice were killed by cervical dislocation and their lungs were rapidly dissected for the determination of bacterial load. The dissected lungs were homogenized on ice in 500 µl of PBS using an Ultra-Turrax TIO basic homogenizer (IKA). Bacteria from the homogenates and from serial dilutions thereof were recovered on chocolate-agar plates. The results are reported as log CFU per gram of tissue. In each case, clones recovered from the mice were confirmed by PCR. The mice were treated in accordance with the Directive of the European Parliament and of the Council on the protection of animals used for scientific purposes (Directive 2010/63/EU) and in agreement with the Bioethical Committee of the University of the Balearic Islands. This study was approved by the Bioethical Committee of the University of the Balearic Islands under authorization number 1748.

Reverse transcription–quantitative real-time PCR. RNA from strain NTHi375 grown in sBHI and treated or not with 1.5 μg N-methylN′-nitro-N-nitrosoguanidine (MNNG) ml–1 for 1 h was extracted using the RNeasy minikit (Qiagen) and DNase treatment (Ambion). Reverse transcription–quantitative real-time PCR (RT-qPCR) was performed in a 20-µl reaction mixture with Lightcycler RNA Master SYBR Green I (Roche) on a

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Lightcycler 480 instrument (LC480; Roche), following the manufacturer’s instructions and using suitable oligonucleotide primer pairs for each gene (Table 1). The relative mRNA concentration obtained from the tag gene was determined according to a standard curve generated by amplifying an internal fragment of the trpA gene, which is not affected by MNNG treatment.

Similarly, the recA gene served as the positive control for the induction of gene expression by MNNG. The expression factor was calculated as the ratio between the mRNA concentrations obtained from genes expressed in MNNG-treated NTHi 375 cells with respect to those from untreated cells.

In silico analysis. To identify the proteins of H. influenzae involved in DNA alkylation repair, E. coli protein sequences implicated in this system were scanned for homologues by using BLASTP [http://blast.ncbi.nlm.nih. gov/Blast.cgi] against the published genomes of H. influenzae strains.

Results and Discussion The present work was designed to determine the significance of DNA alkylation damage repair in NTHi virulence. In silico analysis revealed that the adaptive response to the repair of DNA alkylation damage was missing in H. influenzae because Ada, AlkA, AlkB, and AidB proteins were absent. However, this bacterium contains the genes encoding ogt and tagI as well as the genes involved in the NER and TCR systems.

Based on these results, we studied the importance of N3-meADNA glycosylase I, encoded by the tagI gene, and the Mfd protein involved in transcription-coupled repair. To achieve this, we constructed both tagI and mfd knockout mutants, as detailed in Fig. 1 for the construction of the tagI mutant.

However, rpsL and the spectinomycin resistance genes could not be removed from the knockout mutants. It seems that the plasmid pRSM2947 was not compatible with strain NTHi375 because transformed cells were not obtained even when the cells were incubated at the non-restrictive temperature. Thus, spectinomycin resistance was conserved in the mutants.

Investigation of the virulence of the two mutants in a BALB/c mouse model showed that after infection of the mice, both the mfd mutant and the wild-type strain were recovered at the same concentration; while the recovery of tagI cells was dramatically lower (P = 0.023) (Fig. 2). This effect was not due to differences in growth of the tagI mutant because its growth rate was similar to that of the wild-type strain (Fig. 3).

Based on these results, we considered whether tagI expression was inducible by DNA alkylation damage. Cultures of NTHi375 were treated with a sublethal concentration (1.5

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Statistical analysis. Statistical analyses were performed using oneway analysis of variance (ANOVA) with Bonferroni contrasts. P < 0.05 was considered statistically significant. The analyses were performed using Prism4 for PC (GraphPad Software).

Fig. 2. Bacterial loads of NTHi in the lungs of BALB/c mice after 48 h of infection with the wild-type strain and the mfd and tagI mutants. (Standard deviations are shown.)

μg/ml) of MNNG for 1 h after which the expression of tagI was determined by RT-qPCR. Expression of the recA gene served as a positive control. The results showed that, as in other bacteria [13], the expression of NTHi375 tagI was constitutive because it was not further induced by MNNG treatment, whilst the expression of recA gene was induced by a factor of 3.6 (Fig. 4).

These observations indicated that the 3-methyladenine DNA glycosylase I activity encoded by the tagI gene is crucial for NTHi 375 survival during lung infection. Similar to AlkA, TagI is a monofunctional glycosylase of the base excision repair system; as such, it hydrolytically cleaves the glycosidic bond of alkylated purine bases. However, unlike AlkA, TagI has a very high specificity because it almost exclusively cleaves 3-methyladenine [2,6]. This specificity probably arises from the enzyme’s unique aromatic-residuerich 3-MeA binding pocket and the absence of the catalytic aspartate that is present in all other helix-hairpin-helix family members, including AlkA [6].

Our results contrast with those reported for Salmonella enterica, in which inactivation of the ada, ogt, tag, uvrB, and mfd genes is necessary to decrease bacterial virulence when the cells are orally inoculated in mice [1]. It has been suggested that the extensive alkylation repair system of Salmonella is involved in the survival of Salmonella cells outside the infected animal, enabling them to overcome the potentially massive

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setting of tobacco smoke and NTHi respiratory infections [10], the bacterial TagI protein would thus be critical in repairing DNA damage caused by the alkylating agents in cigarette smoke during the infective process of NTHi. Acknowledgments. We are deeply indebted to Dr. E. Tracy for provide us with E. coli strain DY380 and plasmids pRSM2832, and pRSM2947. This work was supported by grants BFU2008-01078 and 2009SGR1106, from the Ministerio de Ciencia y Innovación and the Generalitat de Catalunya, respectively. CIBERES is an initiative from Instituto de Salud Carlos III. D.A. Spricigo has a predoctoral fellowship supported by CAPES (Coordenação de Aperfeiçoamento de Pessoal de Nível Superior), Brazil.

Competing interests. None declared.

Fig. 3. In vitro growth of NTHi375 wild-type strain () and tagI mutant (□) in sBHI.

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References

Fig. 4. Expression factor of the tagI and recA genes in the NTHi375 strain after treatment with MNNG at 1.5 µg/ml. The expression factor is the ratio of the mRNA concentration of each gene from the treated wild-type strain with respect to the non-treated. The amount of mRNA of each gene was determined by using a standard curve generated by the amplification of an internal fragment of the H. influenzae trpA gene (see Table 1 for primer sequences). Standard deviations of two independent experiments are shown.

DNA injuries induced by alkylating agents present in the environment [1]. By contrast, H. influenzae is a human obligate pathogen well adapted to the human upper respiratory tract, and with a low persistence outside the host [8], which would explain why it does not have the full complement of repair mechanisms needed to repair alkylation-type damage. Consequently, the deletion of a key protein in the repair of alkylation injuries must be more relevant for this species than it is for Salmonella. In this context, the role of N3-meA-DNA glycosylase I in NTHi survival in human airways must be emphasized, because this enzyme catalyzes the specific removal of N3-methyladenine, a mainly lethal insult that blocks DNA replication [3,9]. In the

1. Álvarez G, Campoy S, Spricigo DA, Teixidó L, Cortés P, Barbé J (2010) Relevance of DNA alkylation damage repair systems in Salmonella enterica virulence. J Bacteriol 192:2006-2008 2. Bjelland S, Seeberg E (1987) Purification and characterization of 3-methyladenine DNA glycosylase 1 from Escherichia coli. Nucleic Acids Res 15:2787-2801 3. Boiteux S, Huisman O, Laval J (1984) 3-Methyladenine residues in DNA induce the SOS function SfiA in Escherichia coli. EMBO J 25:25692573 4. Bouchet V, Hood DW, Li J, et al. (2003) Host-derived sialic acid is incorporated into Haemophilus influenzae lipopolysaccharide and is a major virulence factor in experimental otitis media. Proc Natl Acad Sci USA 100:8898-8903 5. Cairns J (1980) Efficiency of the adaptive response of Escherichia coli to alkylating agents. Nature 286:176-178 6. Drohat AC, Kwon K, Krosky DJ, Stivers JT (2002) 3-Methyladenine DNA glycosylase I is an unexpected helix-hairpin-helix superfamily member. Nat Struct Biol 9:659-664 7. Garmendia J, Martí-Lliteras P, Moleres J, Puig C, Bengoechea JA (2012).

Genotypic and phenotypic diversity in the noncapsulated Haemophilus influenzae: adaptation and pathogenesis in the human airways. Int Microbiol 15:157-170 8. Kramer A, Schwebke I, Kampf G (2006). How long do nosocomial pathogens persist on inanimate surfaces? A systematic review. BMC Infect Dis 6:130 9. Larson K, Sahm J, Shenkar R, Strauss B (1985) Methylation-induced blocks to in vitro DNA replication. Mutat Res 150:77-84 10. Margison GP, Cooper DP, Potter PM (1990) The E. coli ogt gene. Mutat Res 233:15-21 11. Martí-Lliteras P, Regueiro V, Morey P, Hood DW, Saus C, Sauleda J, Agustí AGN, Bengoechea JA, Garmendia J (2009) Nontypeable Haemophilus influenzae clearance by alveolar macrophages is impaired by exposure to cigarette smoke. Infect Immun 77:4232-4242 12. Poje G, Redfield RJ (2003) Transformation of Haemophilus influenzae.

Methods Mol Med 71:57-70 13. Qu J, Lesse AJ, Brauer AL, Cao J, Gill SR, Murphy TF (2010) Proteomic expression profiling of Haemophilus influenzae grown in pooled human sputum from adults with chronic obstructive pulmonary disease reveal antioxidant and stress responses. BMC Microbiol 10:162

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14. Riazuddin S, Lindahl T. (1978) Properties of 3-methyladenine-DNA glycosylase from Escherichia coli. Biochemistry 17:2110-2118 15. Scherer G, Urban M, Hagedorn HW, Serafin R, Feng S, Kapur S, Muhammad R, Jin Y, et al. (2010) Determination of methyl-, 2-hydroxyethyl- and 2-cyanoethylmercapturic acids as biomarkers of exposure to alkylating agents in cigarette smoke. J Chromatogr B Analyt Technol Biomed Life Sci 878:2520-2528 16. Sedgwick B (2004) Repairing DNA methylation damage. Nat Rev Mol Cell Biol 5:148-157 17. Selby CP, Sancar A (1994) Mechanisms of transcription-repair coupling and mutation frequency decline. Microbiol Rev 58:317-329

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18. Tracy E, Ye F, Baker BD, Munson RS Jr (2008) Construction of non-polar mutants in Haemophilus influenzae using FLP recombinase technology.

BMC Mol Biol 9:101 19. Van Eldere J, Slack MP, Ladhani S, Cripps AW (2014) Non-typeable Haemophilus influenzae, an under-recognised pathogen. Lancet Infect Dis. doi:10.1016/S1473-3099(14)70734-0 20. Van Houten B, Sancar A (1987) Repair of N-methyl-N’-nitro-Nnitrosoguanidine-induced DNA damage by ABC excinuclease. J Bac­ teriol 169:540-545

RESEARCH ARTICLE International Microbiology (2014) 17:165-174 doi:10.2436/20.1501.01.219. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Endophytic and rhizospheric bacterial communities isolated from the medicinal plants Echinacea purpurea and Echinacea angustifolia Carolina Chiellini,1,2 Isabel Maida,1 Giovanni Emiliani,3 Alessio Mengoni,1 Stefano Mocali,2 Arturo Fabiani,2 Sauro Biffi,4 Valentina Maggini,5 Luigi Gori,5 Alfredo Vannacci,5 Eugenia Gallo,5 Fabio Firenzuoli,5 Renato Fani1* Department of Biology, University of Florence, Florence, Italy. 2Agrobiology and Pedology Research Center, Agricultural Research Council, Florence, Italy. 3Trees and Timber Institute National Research Council, Florence, Italy. 4Botanical Garden, Casola Valsenio, Italy. 5Center for Integrative Medicine, Careggi University Hospital, University of Florence, Florence, Italy 1

Received 23 May 2014 · Accepted 17 September 2014

Summary. In this work we analyzed the composition and structure of cultivable bacterial communities isolated from the stem/leaf and root compartments of two medicinal plants, Echinacea purpurea (L.) Moench and Echinacea angustifolia (DC.) Hell, grown in the same soil, as well as the bacterial community from their rhizospheric soils. Molecular PCR-based techniques were applied to cultivable bacteria isolated from the three compartments of the two plants. The results showed that the two plants and their respective compartments were characterized by different communities, indicating a low degree of strain sharing and a strong selective pressure within plant tissues. Pseudomonas was the most highly represented genus, together with Actino­ bacteria and Bacillus spp. The presence of distinct bacterial communities in different plant species and among compartments of the same plant species could account for the differences in the medicinal properties of the two plants. [Int Microbiol 2014; 17(3):165-174] Keywords: Echinacea purpurea · Echinacea angustifolia · rhizosphere · medicinal plants · endophytes

Introduction Herbal medicine has become a popular approach to the prevention and treatment of several diseases. Species of the genus Echinacea are among the most commonly used medicinal plants. Echinacea purpurea (L.) Moench, and Echinacea an­ gustifolia (DC.) Hell, are currently used in Europe and the Corresponding author: R. Fani Laboratory of Microbial and Molecular Evolution. Department of Biology University of Florence Via Madonna del Piano 6, I-50019 Sesto Fiorentino Florence, Italy Tel. +39-0554574742 E-mail: renato.fani@unifi.it, renato.fani@virgilio.it *

USA to treat the common cold and respiratory infections [1–3, 69]. The alkylamide, alkaloid, and polyacetylene fractions are considered to have immune-modulatory and anti-inflammatory effects [13,43]. The various bioactivities have been traced to the multiple components rather than to the multiple effects of individual chemical compounds present in Echinacea extracts [23]. The chemical diversity of such compounds has made it difficult to determine whether Echinacea extracts are genuinely medicinally effective and the benefits of these products are controversial [35,39]. However, several studies indicate that Echinacea indeed has antiviral, antioxidant, and anti-inflammatory properties, making it a very promising medicinal botanical species [9,29,61].

The chemical composition of the compounds from medicinal plants varies widely depending on the geographic

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provenance of the plant, the botanical species, its genetics and its biotic factors, as well as the specific anatomic part of the plant (e.g., seed, flower, root, leaf), the extraction techniques, etc. [18]. In addition, it is not known whether the compounds of interest in the Echinacea extracts are synthesized solely by the plant or by the microbial endophytes inhabiting the plant and producing bioactive compounds [7]. Advances in studies of the curative properties of medicinal plants have sparked interest in discovering the characteristics of their bacterial communities, in order to clarify whether the curative properties of a medicinal plant are directly or indirectly related to the presence of certain endophytic bacteria [45]. Endophytes are defined as “microorganisms that live for at least part of their life inside the internal tissues of a plant without causing any disease in the host” [68]. They are largely studied for their ability to produce a wide range of natural products, with pharmaceutical, agrochemical, and biotechnological properties of interest [17,27,59,70]. Endophytes are present in most plant species [42,59], both terrestrial and aquatic [6,63,64]. Their beneficial effects on their hosts include growth promotion through interactions in nitrogen metabolism [48] or phosphorus solubilization [57] and siderophore production [16]. The growth-promoting effects of endophytes include the induction of phytohormone biosynthesis, in particular that of indole3-acetic acid [62], and of 1-aminocyclopropane-1-carboxylate deaminase activity [22]. Endophytes control plant pathogens through the production of antimicrobial compounds [49,65] and act as elicitors of systemic resistance in plants [36].

The colonization mechanisms, ecology, function, and plant interactions of endophytes have been widely investigated [12,27,44,46]. These studies have shown that the bacterial endophytes of plants derive from different sources, such as the spermosphere, anthosphere, caulosphere, and phyllosphere [25,27,58,59]. However, most endophytic bacteria are soil-derived [12], as hypothesized by Galippe already in 1887 [20,21]. Endophytes enter plant tissues through cracks in the roots or aerial parts [37] or through rhizosphere soil colonization and become distributed within the whole plant tissues through its vascular system [5]. Thus, endophytes can be detected in roots, stems, and leaves, inside plant reproductive organs and the apoplastic space and, in some cases, also in intracellular spaces (as is the case in symbiotic rhizobial species) [12,56,58].

The root compartment of Echinacea was shown to contain the highest levels of bioactive molecules [67]. The aim of this study was to determine the endophytic and rhizospheric bacterial biodiversity of two medicinal plants, E. purpurea and E. angustifolia, in order to evaluate their core and accessory

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cultivable microbiomes in relation to the roots and aerial parts of these plants. To the best of our knowledge, this work is the first to describe the diversity of the culturable endophytic and rhizospheric bacterial community of medicinal plants.

Materials and methods Rhizospheric soil chemical variables. The main chemical-physical variables of the soil were: pH (H2O) 8.4; total organic carbon, (TOC) 1.6%; N (g/kg); 1.50; C/N, 10.97; CaCO3 total, 18.70%; cation exchange capacity (CEC, cmol[+]/kg), 9.74.

Plant material and soil properties. Five E. purpurea and five E. angustifolia plants were cultivated in the same square basin (160 cm length × 75 cm height), located in an open field of the botanical garden “Giardino delle Erbe”, Casola Valsenio, Ravenna, Italy, and collected in October 2012.

The basin was filled with the same soil as that sampled. The soil was air-dried at room temperature (21ºC), sieved through a 2-mm mesh, and then analyzed for pH, cation-exchange capacity, total CaCO3, TOC, and total N. Isolation of bacterial strains and preparation of cell lysates for DNA amplification. Collected plants were immediately taken to the laboratory. The anatomical part of the plants, i.e., the roots and stems/leaves, were separated and considered as independent samples throughout the experiment. Roots from the five individual plants of each Echinacea species were grouped and pooled, as were the stems/leaves. Two grams of fresh tissue from each pool was surface-sterilized with 1% HClO solution in sterile 50-ml Falcon tubes at room temperature and then washed three times with sterile water to remove the epiphytic bacteria. Aliquots (100 μl) of the last wash were plated in triplicate as sterility controls, which by the end of the experiment had not become contaminated (data not shown). Subsequently, the samples were homogeneously pottered in a sterile mortar with the addition of 2 ml of 0.9% NaCl (Sigma Aldrich, USA). One hundred μl samples of tissue extracts and their different dilutions were plated in triplicate.

Rhizospheric soil (RS) from five plants of each plant species was also analyzed and treated separately at room temperature for 1 h with 20 ml of 10 mM Mg2SO4 in 50-ml sterile Falcon tubes, which allowed detachment of the bacteria from soil particles. After sedimentation, 100-μl samples of the supernatant and different dilutions thereof were plated in triplicate.

Endophytic and rhizospheric bacteria were grown on solid tryptone soya broth (TSB) medium (Biorad, CA, USA) at 30°C for 48 h. The total number of aerobic heterotrophic fast-growing bacteria was expressed as colony-forming units (CFU), which were determined for each sample in triplicate based on an average value of the bacterial titer. From each sample, about 100 colonies were randomly selected and individually plated onto solid TSB Petri dishes.

A collection of 514 bacterial isolates was prepared for both Echinacea sp. plant species by dissolving the freshly isolated and plated colonies in 2-ml deep-well plates with 500 μl of TSB and 500 μl of 50% glycerol (25% final concentration). The plates were stored at –80°C. Random amplified polymorphic DNA (RAPD) analysis. Cell lysates of endophytic and rhizospheric bacterial isolates were prepared by processing 100 μl of each glycerol suspension with thermal lysis (95°C for 10 min), followed by cooling on ice for 5 min.

Random amplification of DNA fragments [65] was carried out in a 25-μl total volume composed of 1× reaction buffer, 300 μM MgCl2, deoxynucleoside triphosphate (200 μM each), 0.5 U of PolyTaq DNA polymerase (all re-

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agents were from Polymed, Florence, Italy), 500 ng of primer 1253 [5′-GTT TCCGCCC-3′] [46], and 2 μl of cell lysate prepared as described above. The reaction mixtures were incubated in a PTC-100 Peltier thermal cycler (MJ Research, Quebec, Canada) at 90°C for 1 min, and 95°C for 90 s followed by 45 cycles at 95°C for 30 s, 36°C for 1 min, and 75°C for 2 min. Finally, the reaction mixtures were incubated at 75°C for 10 min, 60°C for 10 min, and 5°C for 10 min. Reaction products were analyzed by agarose (2% w/v) gel electrophoresis in Tris-acetate EDTA buffer (TAE) containing 0.5 μg ethidium bromide/ml.

stem/leaves compartments of both plant species (4.80 × 103 ± 2.5 × 103 CFU/g in E. purpurea and 3.43 × 103 ± 2.1 × 103 CFU/g in E. angustifolia); the highest bacterial titers were in the root compartment of E. angustifolia (1.34 × 106 ± 9.7 × 105 CFU/g) and the RS compartment of E. purpurea (9.6 × 105 ± 6.4 × 105). Overall, the numbers of CFU/g isolated from the same compartment of the different plants were very similar.

Analysis of RAPD profiles. The genetic similarities in the different samples belonging to the same haplotype group were determined based on the fingerprinting pattern of each RAPD product and pairwise comparisons of the presence/absence of bands using the GelCompar II software (Applied Maths). For each recognized RAPD haplotype, a 16S rRNA gene sequence was obtained via PCR amplification (as described below) for taxonomic attribution of the bacterial isolates. For haplotypes represented by more than one strain, a single bacterial strain was randomly chosen for gene amplification.

RAPD fingerprinting. To determine the degree of genetic variability at the strain level and to analyze the structure of the isolated bacterial communities, 262 isolates were collected from E. angustifolia (83 from S/L, 88 from R, and 91 from RS) and 252 from E. purpurea (81, 89, and 82, respectively). Each of the 514 bacterial isolates was subjected to RAPD fingerprinting as follows: DNA from the lysed cell suspensions was amplified with the 10-mer oligonucleotide 1253 as described in Materials and methods. RAPD amplicons were then analyzed by agarose gel electrophoresis. Each RAPD profile (hereinafter referred to as haplotype) was then compared with all of the others as described in Materials and methods. Thus, from the 514 bacterial isolates 380 different RAPD haplotypes were identified, corresponding to at least 380 bacterial strains. Specifically, the 252 and 262 bacterial isolates from E. purpurea and E. angustifolia yielded 201 and 203 haplotypes, respectively. Among the 514 bacterial strains, 316 (61.5%) had a unique haplotype. Thirty-three haplotypes were composed of two strains each (8.7% of all haplotypes); 16 haplotypes (4.2%) were composed of 3 strains; 12 haplotypes (3.1%) were composed of 4–6 strains. Haplotype 1 comprised 7 different bacterial isolates; haplotype 66, 8 different bacterial isolates; and haplotype 2, 12 different bacterial isolates from both Echinacea species.

In E. purpurea, the RS compartment had the highest number of RAPD haplotypes (80 haplotypes out of 82 bacterial isolates), whereas the root compartment harbored 59 total different haplotypes, and the S/L compartment, 70 haplotypes.

Similarly, the highest number of different haplotypes (84) isolated from E. angustifolia was detected in the RS compartment, whereas the S/L and R compartments yielded 62 and 67 different RAPD haplotypes, respectively.

The distribution of the RAPD haplotypes within the different compartments of the same plant species is shown in Fig. 1. Notably, no strain was shared among the three compartments in either E. purpurea or E. angustifolia, and a very low number of strains was shared between two different compartments. In E. purpurea, three haplotypes were shared between R and RS, four between S/L and RS, and two between S/L and R. In E. angustifolia eight, two and zero haplotypes

PCR amplification and sequencing of 16S rRNA coding genes. PCR amplification of 16S rRNA genes was carried out in 20-μl reactions containing 1× reaction buffer, 150 μM MgCl2, deoxynucleoside triphosphate (250 μM each), 2 U of PolyTaq DNA polymerase (all reagents were from Polymedaly), 0.6 μM of each primer [P0 5′-GAGAGTTTGATCCTGGCTCAG and P6 5′-CTACGGCTACCTTGTTACGA] [22], and 2 μl of each cell lysate. The samples were incubated in a PTC-100 Peltier thermal cycler (MJ Research) under the following conditions: primary 90-s denaturation at 95°C, 30 cycles of 30 s at 95°C, 30 s at 50°C, and 1 min at 72°C, followed by a final extension of 10 min at 72°C. Amplicons were then excised from 0.6% agarose gels and purified using the MinElute gel extraction kit (Qiagen) according to the manufacturer’s instructions. Direct sequencing of the amplified 16S rRNA genes was performed with primer P0 [5′-GAGAGTTTGATCCTGGCTCAG] using an ABI PRISM 310 genetic analyzer (Applied Biosystems) and the chemical dye terminator [59]. Each 16S rRNA gene sequence was submitted to GenBank and assigned an accession number from AJ642225 to AJ642604.

Analysis of 16S rRNA gene sequences. The 16S rRNA gene sequences were analyzed using the “classifier” tool of the Ribosomal Database Project –RDP [10] for taxonomic assignment. All sequences showed a degree of similarity ≥80% with the sequences available in the RDP database (at genus level) and were therefore considered of good quality. Diversity indices were calculated for both plant species and for each compartment using data on the genus-level taxonomic attribution of the RDP database. Abundance data were first normalized by calculating the percentage of each bacterial genus present in the three compartments of the two plant species. Diversity indices were calculated using PAST software version 3.0 [25]. The number of taxa for each plant compartment (S) was calculated together with the Simpson’s diversity index (D), Shannon’s diversity index (H), and evenness (e^H/S).

Results Bacterial counts. Bacteria were extracted from the rhizospheric soil (RS), roots (R), and stems/leaves (S/L), diluted in saline (0.9% NaCl), and plated as described in Materials and methods. The lowest CFU/g values were detected in the

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Fig. 1. Schematic representation of the core, accessory, and unique RAPD haplotypes detected in Echinacea purpurea and E. angustifolia rhizospheric soil (RS), roots (R) and stem/leaves (S/L).

Comparison of the bacterial cultivable communities inhabiting Echinacea purpurea and E. angustifolia. The bacterial communities inhabiting the two plant species were then compared to determine the degree of sharing at the genus level (Fig. 3). The analysis revealed that the two plants shared 14 genera (including all the most represented ones). Again, Pseudomonas was the most highly represented genus in both plant species, with no considerable

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were shared between R and RS, S/L and RS, and S/L and R, respectively. An analysis of the distribution of RAPD in the two Echinacea plant species showed that they shared only 23 haplotypes (about 6%; data not shown). The number of haplotypes common to the two plant species and the different compartments ranged from one (a single haplotype shared by E. purpurea RS and E. angustifolia RS, and by E. purpurea R and E. angustifolia RS) to six (shared by E. purpurea RS and E. angustifolia R compartments).

Analysis of 16S rRNA gene sequences. By assuming that bacterial isolates sharing the same RAPD profile corresponded to the same (or a closely related) strain, the composition of the six bacterial communities was assessed by analyzing the 16S rRNA gene sequence from a representative of each RAPD group. Thus, 16S rRNA genes were amplified from the 380 strains. An amplicon of the expected size was obtained from each one (not shown). The nucleotide sequences of these 380 amplicons were determined and compared with those available in databases. The entire dataset is summarized in Fig. 2. The analysis revealed that: (i) the 380 strains were affiliated with 29 different bacterial genera; (ii) the majority (47.4% of RAPD haplotypes) of the 16S rRNA gene sequences were affiliated with the genus Pseudomonas; (iii) Staphylo­ coccus was the second most highly represented genus (9.5% of RAPD haplotypes); (iv) Microbacterium spp. and Curto­ bacterium spp. accounted for 6.1% and 6.3%, respectively, and Arthrobacter sp. for 4.2% of the strains; (v) 11 bacterial genera accounted for 1.3–2.6% of the strains; and (vi) the remaining 13 genera represented <1%.

Fig. 2. Abundance of the different bacterial genera in all haplotype detected with RAPD analysis of both Echinacea purpurea and E. angustifolia.

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Fig. 3. Distribution of the different bacterial genera in the two medicinal plants analyzed in this study.

differences in their abundances (51.5% in E. angustifolia and 48% in E. purpurea). There was a large difference in abundances of Curtobacterium spp.: 0.8% in E. purpurea and 11.8% in E. angustifolia. Similarly, Pantoea spp. accounted for 0.4% of the genera in E. purpurea and 4.2% of those in E. angustifolia. By contrast, Arthrobacter spp. was much more abundant in E. purpurea than in E. angustifolia (6% vs.

0.8%), as were Flavobacterium spp. (2.8% vs. 0.8%) and Methylobacterium spp. (2.8% vs. 0.4%). E. purpurea hosted a larger number of bacterial genera than E. angustifolia (23 vs.

16), such that some bacterial genera were found only in E. purpurea, albeit at low percentages: Agrococcus spp., Rhi­ zobium spp. and Stenotrophomonas spp. (0.8%), Cupriavidus spp., Shinella spp., and Xanthomonas spp. (0.4%), Frigori­ bacterium spp. (2.8%), Rhodobacter spp. (3.2%), and Sphin­ gomonas spp. (4.0%). Kocuria spp. (0.4%) and Acinetobacter spp. (0.8%) were found only in E. angustifolia.

Comparison of the bacterial communities inhabiting different plant compartments. Figure 4 shows the composition of the bacterial communities isolated from the different compartments of the two Echinacea plant species. The diversity indices are shown in Table 1.

The composition of the cultivable bacterial community inhabiting the S/L compartment of both plants was highly different from that in the other two compartments (RS and R), which were much more similar to each other. The genus Pseu­ domonas was less represented in the S/L compartment than in the R and RS compartments, whereas a high percentage of bacteria associated with the genus Staphylococcus were detected in the S/L compartment of both plant species (25.9% in E. purpurea and 38.6% in E. angustifolia), thus underlining the differences in bacterial composition of this compartment compared with the other two. Differences were also evident concerning the genera Bacillus and Curtobacterium, which

Table 1. Diversity indices calculated from the percentages of the presence of the different bacterial genera in each plant compartment for both plant species Ep S/L

Ep R

Ep RS

Ea S/L

Ea R

Ea RS

Taxa_S

12

6

16

8

7

9

Simpson_D

0.8584

0.4574

0.7

0.6972

0.473

0.4448

Shannon_H

2.174

0.9625

1.785

1.405

1

1.063

Evenness_e^H/S

0.7329

0.4364

0.3724

0.5096

0.3885

0.3216

Ep: Echinacea purpurea; Ea: Echinacea angustifolia; S/L: stem/leaf com­par­tment; R: root compartment; RS: rhizospheric soil compartment.

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Fig. 4. Relative abundances (expressed as percentages) of cultivable bacterial taxa isolated from rhizospheric soil (RS), roots (R), and stem/leaves (S/L) of Echinacea purpurea and E. angustifolia.

were not detected in the root compartment of either of the two plant species; Agrobacterium, which was not identified in the S/L compartment of the two Echinacea species; Rhein足 heimera, which was present only in the rhizosphere of both plants; and Achromobacter, which was present only in the roots of the two plants.

The differences among the culturable bacterial communities were also highlighted by the diversity indices (Table 1).

Overall, E. purpurea compartments were characterized by a higher number of bacterial taxa (genera) than E. angustifolia.

The highest number of taxa were found in the rhizosphere of E. purpurea (16), followed by the stems/leaves of this species.

The lowest number of taxa occupied the roots of both plant species, with just seven different bacterial genera in E. angus足 tifolia and six in E. purpurea.

In E. purpurea, the Simpson diversity index was the highest (0.85) in the S/L compartment, and the lowest in the R compartment (0.46). Analogously, in E. angustifolia, the Simpson index was the highest (0.69) in the S/L, and the lowest (0.47) in the R compartment. The same was true for the Shannon diversity and evenness indices in both plant species.

Considering the same compartment in the two species, the Simpson index values in the root compartments of E. purpurea and E. angustifolia were similar (0.46 and 0.47, respectively), as

ENDOPHYTES AND RHIZOSPHERE OF ECHINACEA

was the Shannon index (0.96 and 1, respectively). In the RS compartment of E. purpurea, Simpson’s and Shannon’s diversity indices were higher than in the RS compartment of E. angustifolia, but the evenness values were very similar (0.37 and 0.32, respectively).

Discussion To the best of our knowledge, this is the first report in which the endophytic and rhizospheric cultivable bacterial communities were enumerated and described in the medicinal plant Echinacea spp. in an open field trial. In the only other available report in which Echinacea sp. endophytes were characterized, the plants were propagated in vitro for 9 months [41].

In a recent study, Pugh et al. [51] enumerated the total bacterial endophytes in E. purpurea roots and aerial samples in a molecular approach that used a PCR-based quantification method. Their study was aimed at determining whether differences in bacterial load correlated with the in vitro macrophage activity of the plant material.

In this work, bacterial endophytes were isolated from the S/L and R compartments of E. purpurea and E. angustifolia, as well as from their RS. We are aware that the approach used in this work has several limitations related to the experimental conditions, which selected for bacteria able to grow on TSB/ TSA within 48 h of incubation at 30°C; however, this bias was negligible considering the aim of this preliminary study exploring the differences within and between the culturable microbial communities inhabiting two plants and their compartments. The effect of different growth conditions should be evaluated in additional experiments aimed at obtaining slowgrowing bacteria and/or bacteria able to grow in different media or at different temperatures.

The yield of culturable heterotrophic fast-growing bacteria in both plant species ranged from 103 CFU/g in the S/L compartments to 105–106 CFU/g in the R and RS compartments; these values are in agreement with those in previously published studies showing greater abundances of plant-associated bacterial populations in R than in S/L [37,40]. The very low CFU/g values detected in the S/L compartment in both plant species is also in agreement with the findings of previous studies [51]. The RAPD data suggested a non-clonal structure and a very high degree of genetic variability at the strain level of the endophytic and rhizospheric bacterial communities from both Echinacea spp. There was also a low level of strain sharing in the different compartments of the same plant and between the same compartment of the two different

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plants. For example, just five strains were shared by the S/L compartments of the two plants and only four haplotypes were shared between their R compartments.

This finding raises the intriguing question of the nature of the (internal?) plant forces driving the selection of different bacterial endophytic strains in the diverse compartments of the same plant, both for E. purpurea and for E. angustifolia.

One explanation may be related to the finding that plants have an innate immune system, with receptors that detect the presence of molecules both inside and on the surface of host cells [34]. Indeed, the selection of bacteria at the rhizospheric level (the rhizosphere effect) is well documented and was shown to be related to plant exudates released in the soil surrounding the roots, which results in the selection of a bacterial biota that is different from the one recovered in bulk soil (for a review see [8]).

Most of the bacterial genera identified through 16S rRNA gene amplification in Echinacea sp. have been detected in other plant species. Ikeda et al. [33] characterized 217 endophytic isolates from the roots of maize; the most highly represented bacterial genera were Pantoea and Bacillus. In the study of Gagne-Bourgue et al. [19], the endophytic components of the aerial parts of the switchgrass (Panicum virgatum L.) were analyzed; the most highly represented bacterial genera were Microbacterium, Curtobacterium, Bacillus, Pseudo­ monas, Pantoea, Sphingomonas, and Serratia. As seen in the diversity indices shown in Table 1, the higher numbers of bacterial genera in the S/L and RS compartments of E. purpurea than E. angustifolia suggests a higher microbial diversity level in the former. By contrast, there were no significant differences in the root compartments.

As expected, some genera present in the RS compartment were also detected within plant tissues, since most bacterial endophytes are presumably derived from soil. Note that the genera Arthrobacter, Staphylococcus, and Methylobacterium were detected in the RS and in the S/L compartments of E. purpurea but not in its R compartments, whereas the genus Stenotrophomonas was detected in the RS and R compartments. Analogously, in the two plant species the genera Curto­ bacterium and Bacillus were detected in the RS and S/L compartments, and the genus Agrobacterium in the RS and R compartments. The genus Microbacterium was present in all E. pur­purea compartments and in the RS and R compartments of E. an­gustifolia.

The distributions of these genera suggest that different endophytic bacterial species are selected not only at the RS level but also within plant tissues, thus indicating selection at the rhizospheric level and, once the bacteria reach plant tissues from the RS, their stabilization as endophytes.

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In the two plant species, the highest values of D, H, and evenness were those of the S/L compartments. This finding, together with the observation that these same compartments had the lowest bacterial titers, led us to hypothesize that, although there were fewer endophytes in the S/L compartments than in the other two plant compartments, they were much more diversified and each taxon (in this case, genus) was represented more or less by an equal number of individuals, as indicated by the evenness values. By contrast, in the roots of both plants, the low evenness values together with the bacterial counts revealed higher bacterial abundances in the roots (4.6 × 105 ± 2.8 × 104 in E. purpurea and 1.3 × 106 ± 9.6 × 105 in E. angustifolia) than in the stem/leaves (4.5 × 103 ± 2.5 × 103 in E. purpurea and 3.4 ×103 ± 2.1 × 103 in E. angustifolia), consistent with the presence of only a few dominant bacterial taxa.

These differences in bacterial numbers and distribution between roots and stem/leaves of Echinacea spp. could be related to the different environmental and nutritional conditions to which the roots and the aerial parts are exposed and/or to anatomical and phytochemical features that, in turn, create specific ecological niches for endophytes. The aerial parts of plants are exposed to fluctuations in temperature, humidity, and UV irradiation and to a different trophism/physiology than roots [29]. Endophytic bacteria are likely selected on the basis of their adaptation strategies and tolerance of the different conditions in the various plant compartments, as reported for several plant species (e.g., [4,13,32,50]). Our analysis of the bacterial genera in the stems/leaves of Echinacea sp., showed Sphingomonas sp. in E. purpurea (12.3%) and Meth­ ylobacterium in E. purpurea and E. angustifolia (7.4% and 1.2%, respectively). Delmotte et al. [15] identified microbial proteins that appear to reflect differential adaptation strategies to the leaf environment of two abundant colonizers belonging to these two genera. This finding is in agreement with the data obtained in our study and could explain the presence of Sphin­ gomonas and Methylobacterium in Echinacea sp., as endophytes adapted to this environment.

Among the compartements of Echinacea, the root compartment is richest in bioactive molecules responsible for the medicinal properties of these plants [67]. The potential of endophytes from medicinal plants to produce anticancer, antibacterial, and antifungal compounds was recently demonstrated [45]. Similarly, a few dominant bacterial genera inhabiting Echinacea sp. roots might be candidates for the synthesis of bioactive compounds and/or used to drive plant metabolism to synthesize these compounds. The dataset obtained with RAPD fingerprinting highlights a very low degree of

CHIELLINI ET AL.

sharing between the two plant species and, especially, between the two rhizospheric soils. Although, as discussed above, different plants select for different rhizospheric microbial communities [38], this result was unexpected given that the two plant species were grown in the same soil, that is, having the same chemical-physical characteristics, and within a few centimeters of each other. This suggests a high specific interaction between the plant roots and the microbial communities residing in soil.

Many rhizospheric bacteria switch from root-surface to endophytic lifestyles [12,58], including species of Bacillus and Pseudomonas. The structures and functions of their lipopeptides of these two genera were recently reviewed [54,55].

Lipopeptides are used by rhizosphere bacteria in antibiotic production and the induction of plant defense mechanisms.

Thus, the dominance of Pseudomonas in the rhizospheres and roots of the two medicinal plants considered in this study could be related to the production by the plants of metabolites having medicinal properties, either directly, as already described for other species [10], or indirectly, through potential plant-growth-promoting properties [30].

In the present study, bacteria belonging to the phylum Ac­ tinobacteria (genera Microbacterium, Frigoribacterium, Cur­ to­bac­terium, Arthrobacter, Agrococcus and Kocuria) accounted for 18.4% of the total endophytes in E. purpurea and 24.1% of those in E. angustifolia, which in both cases is a significant portion of the whole bacterial component. Endophytes belonging to Actinobacteria have been widely studied for their production of secondary metabolites [51,52], which were shown to include those with diverse biological activities, such as antibiotics, antitumor and anti-infection compounds, plant growth promoters, and enzymes [28,52]. This suggests that Actinobacteria together with Pseudomonas and Bacillus would be in large part responsible for the production of the compounds that account for the characteristics of the medicinal plant Echinacea spp. However, additional work should be carried out in order to specifically confirm such a hypothesis.

In summary, in this work, we analyzed the structure and composition of cultivable bacterial communities isolated from the stem/leaves, roots, and rhizospheric soils of two species of medicinal plants, E. purpurea and E. angustifolia, using a combination of PCR-based techniques. The results revealed differences in the microbial communities inhabiting the two plants, despite their growth in the same soil. In addition, different bacterial communities inhabited the different plant compartments. These findings together with the very low degree of strain sharing, raise intriguing question regarding the existence of a (strong) selective pressure that deter-

ENDOPHYTES AND RHIZOSPHERE OF ECHINACEA

mines bacterial composition at the strain level, rather than at a higher taxonomic rank (genus). Although yet to be confirmed experimentally, bacterial endophytes would seem to elicit the synthesis of some if not all of the bioactive molecules produced by the plant, perhaps synthesizing some of these compounds themselves.

Acknowledgements. This work was supported financially by Ente Cassa di Risparmio di Firenze (Project 2013.0657). We are also very grateful to three anonymous referees for their helpful comments that improved the manuscript.

Competing interests. None declared.

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RESEARCH ARTICLE IÄã ÙÄ ã®ÊÄ ½ M® ÙÊ ®Ê½Ê¦ù (2014) 17:175-184 doi:10.2436/20.1501.01.220. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

A glucuronoxylan-specific xylanase from a new Paenibacillus favisporus strain isolated from tropical soil of Brazil Itácio Q.M. Padilha,1 Susana V. Valenzuela,2 Teresa C.S.L. Grisi,1 Pilar Diaz,2 Demetrius A.M. de Araújo,1* Francisco I. Javier Pastor2* 1

Department of Biotechnology, Biotechnology Center, Federal University of Paraíba, João Pessoa, Brazil.

2 Department of Microbiology, Faculty of Biology. University of Barcelona, Barcelona, Spain.

Received 27 May 2014 · Accepted 30 September 2014

Summary. A new xylanolytic strain, Paenibacillus favisporus CC02-N2, was isolated from sugarcane plantation fields in Brazil. The strain had a xylan-degrading system with multiple enzymes, one of which, xylanase Xyn30A, was identified and characterized. The enzyme is a single-domain xylanase belonging to family 30 of the glycosyl hydrolases (GH30). Xyn30A shows high activity on glucuronoxylans, with a Vmax of 267.2 U mg–1, a Km of 4.0 mg/ml, and a kcat of 13,333 min–1 on beechwood xylan, but it does not hydrolyze arabinoxylans. The three-dimensional structure of Xyn30A consists of a common (β/α)8 barrel linked to a side-chain-associated -structure, similar to previously characterized GH30 xylanases. The hydrolysis products from glucuronoxylan were methylglucuronic-acid-substituted xylooligomers (acidic xylooligosaccharides). The enzyme bound to insoluble xylan but not to crystalline cellulose. Our results suggest a specific role for Xyn30A in xylan biodegradation in natural habitats. The enzyme is a good candidate for the production of tailored xylooligosaccharides for use in the food industry and in the biotechnological transformation of biomass. [Int Microbiol 2014; 17(3):175-184] Keywords: Paenibacillus favisporus · xylanase · glycosyl hydrolases GH30

Introduction Xylan is an abundant component of the plant cell wall and the major component of hardwood hemicelluloses. The biodegradation of xylan is a complex process that requires the coordinated activity of several enzymes, including xylanases (EC 3.2.1.8), which play a key role by catalyzing the hydrolysis of internal linkages in the -1,4-xylose backbone of the polymer.

Xylanases are produced by fungi, bacteria, and protozoa

*

Corresponding authors:

FIJ. Pastor. University of Barcelona 08028 Barcelona, Spain. E-mail: fpastor@ub.edu DAM. de Araújo. Federal University of Paraiba João Pessoa, Brazil. E-mail: demetrius@cbiotec.ufpb.br

[10,31] and are currently used in a wide range of industrial applications, such as the food and textile industries, in wastewater management, and in pulp bleaching, all of which exploit the robust activity of xylanases under extreme conditions, including those of industrial processes [10,33]. Xylanases are currently the subject of intense research into the bioconversion of lignocellulosic biomass for use as biofuels and high-added-value products [22,46,47].

Xylanases are grouped into different families according to their amino acid sequence, structural fold, and catalytic mechanism [9]. Most of the xylanases characterized to date belong to glycoside hydrolase (GH) families GH10 and GH11. They hydrolyze different types of xylan, including arabinoxylans, glucuronoxylans, and even algal -1,3--1,4-xylan [3,31]. A few xylanases specific for glucuronoxylans have been characterized. These belong to GH family 30 (GH30) and their

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activity requires methylglucuronic acid (MeGlcA) side chains [36,42,45]; for this reason they are referred to as glucuronoxy­ l­anases. A common feature of their 3D structure is a typical (b/a)8 barrel catalytic domain [34] fused to a side b-structure of nine strands that seems to be required for catalytic activity [42]. Xylanases also occur in the GH8 family [11,13] and a xylanase of family GH5 specific for arabinose-substituted xylan was recently characterized [12]. The cooperation and synergism between xylanases from different families facilitate the degradation of plant xylan in natural habitats and the complete depolymerization of lignocellulosic biomass.

With their great biodiversity, the soils of the Brazilian Atlantic tropical region are a potential source of as yet uncharacterized microorganisms, including lignocellulolytic bacteria belonging to the phyla Proteobacteria, Actinobacteria, and Firmicutes [8]. The diversity of microorganisms present in soils is a notable source of industrial enzymes [20]. With the aim of identifying new xylanases with biotechnological potential, we screened Brazilian tropical soils for xylan-depolymerizing microorganisms and thereby isolated a new bacterial strain, tentatively classified as Paenibacillus favisporus.

The bacterium has a multiple xylanase system resembling those described in members of the genus Paenibacillus [5,21].

In this study, we cloned, purified, and characterized a new xylanase from P. favisporus.

Materials and methods Isolation of the microbial strain. Soil (25 g) from a sugarcane plantation field in Brazil (6.59 S 35.1 W) was used as a screening source. The soil was suspended in 225 ml of Ringer solution containing (g/l): NaCl, 2.250; KCl, 0.105; CaCl2, 0.120; NaHCO3, 0.050. The suspension was mixed by rotation on an orbital shaker (Certomat RM, B. Braun Biotech Intl.) at 200 rpm for 20 min. Suitably diluted samples were spread onto solid medium containing (g/l): beechwood xylan (Sigma-Aldrich, St. Louis, MO), 1.0; NaNO3, 0.5; K2HPO4, 1.0; MgSO4·7H2O, 0.5; FeSO4·7H2O, 0.01; yeast extract, 1.0; and agar, 15.0; at pH 7.0. The plates were incubated at 55 °C for 3 days. Isolates were transferred into xylan-containing agar medium and incubated at 55 °C for 3 days. Microorganisms with xylanolytic activity were detected based on the formation of clear zones around colonies, visualized using the Congo red staining method [38]. Enzyme production was studied in Erlenmeyer flask cultures (100 ml) containing 20 ml of LB medium and (g/l): yeast extract, 5.0; NaCl, 10; tryptone, 10; rice straw, 10.0. The flasks were inoculated with 0.5% of the overnight inoculum and incubated at 37ºC under shaking (200 rpm) for 48 h. Cultures were centrifuged at 10,000 ×g at 4ºC for 20 min, and the cell-free clear supernatant was analyzed in a zymogram assay. Isolate CC02-N2 was chosen for subsequent studies.

Nucleic acid manipulation. Bacterial genomic DNA was extracted with the High Pure PCR product purification kit (Roche) according to the manufacturer’s instructions. Genomic DNA was diluted appropriately and used as a template in polymerase chain reactions (PCRs) with the universal

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bacterial primers 27F and 1525R. The PCRs were run for 25 cycles with the following thermal profile: 94ºC for 1 min, 55ºC for 1 min, and 72ºC for 2 min and final extension for 10 min at 72 °C. The PCR amplicon was sequenced using an ABI BigDye Terminator v3.1 cycle sequencing kit and an ABI 3730XL DNA analyzer (Applied Biosystems, USA).

Degenerate primers deduced from the sequence of the gene encoding xylanase Xyn30D from Paenibacillus barcinonensis [42] allowed amplification of a portion of a xylanase gene from isolate CC02-N2. The complete sequence of the gene, xyn30A, was obtained by gene walking from the DNA amplified fragment using the Genome Walker universal kit (Clontech).

To overexpress the xylanase, xyn30A was PCR amplified (Kapa-HiFi, KAPA Biosystems) with the oligonucleotide primers FW 5′-AACTATGATTC TATCAAAGAGAATGGAG-3′) and BW (5′-GTGGTGGT­GATGGTGATGG CCATGCGCCAATTCACCTACGAA-3′) and cloned into pLATE31 (Thermo Scientific), giving rise to recombinant plasmid pLATE31-Xyn30A, which produced the full length enzyme, containing the signal peptide, linked to a C-terminal His6 tag (Xyn30A). All DNA constructs were verified by sequencing. Sequence homology was analyzed by BLAST [3].

Expression and purification of recombinant proteins in E. coli. Xyn30A was purified from E. coli BL21 Star (DE3) recombinant clones containing plasmid pLATE31-Xyn30A. Exponential-phase cultures (OD600 0.6) were induced with 1 mM isopropyl-b-d-thiogalactopyranoside at 37°C for 3 h. The cells were disrupted in a French press. The recombinant His6-tagged protein was purified from cell extracts by immobilized metal affinity chromatography using 1-ml HisTrapHP columns (GE Healthcare) and elution in 20 mM phosphate buffer (pH 7.0) with a 0–500 mM imidazole gradient. An additional purification step was performed by gel filtration in Superdex 200/10-300 columns of 24 ml (GE Healthcare) on a fast protein liquid chromatography system (ÄKTA FPLC; GE Healthcare). Buffer exchange and protein concentration were performed in Centricon centrifugal filter units of 10-kDa molecular mass cutoff (Millipore).

Enzyme assays. Xylanase activity was assayed by measuring the amount of reducing sugar released from xylan from hardwoods or cereals by the Nelson-Somogyi method [33]. The standard assay was performed at 50°C in phosphate buffer (pH 6.5) for 15 min as described previously [42]. Birchwood, beechwood, and oat spelt xylans and 4-O-methyl-glucuronoxylan were purchased from Sigma-Aldrich. Rye and wheat arabinoxylans were purchased from Megazyme. One unit of enzymatic activity was defined as the amount of enzyme that releases 1 μmol of reducing sugar equivalent per min under the assay conditions described. A standard curve of xylose was used to calculate activity units. The protein concentrations of the samples were determined using the Bradford method [6]. All determinations of enzyme activity were done in triplicate.

The effects of temperature and pH on xylanase activity were evaluated by response surface methodology (RSM) using 22 central composite designs with 5 coded levels leading to 11 sets of experiments, 8 unique combinations, and 3 replications at the central point . The Briton-Robinson buffer, in a pH range between 4.0 and 11.0 [7], and temperatures ranging from 50 to 90ºC were used in the analysis.

The influence of metal ions and chemicals on xylanase activity was determined by incorporating AlCl3, BaCl2, CaCl2, CuSO4, FeSO4, HgCl2, KCl, LiCl2, MgCl2, MnCl2, NaCl, NH4Cl2, or ZnSO4 at a concentration of 10 mM; and EDTA, SDS, Tween 80, Triton X-100 or DMSO at a concentration of 0.5% into the reaction mixture. Relative xylanase activity was determined.

The kinetics of Xyn30A were characterized in terms of Michaelis-Menten kinetic constants (Km and Vmax) by assaying the enzyme activity on beechwood xylan at concentrations of 1.3–40.0 mg/ml under conditions of maximum activity. Enzyme kinetics were analyzed using GraphPad software version 4.0.

Gel electrophoresis and zymograms. Sodium dodecyl sulfate-poly­­ acrylamide gel electrophoresis (SDS-PAGE) was performed in 12% gels, essentially as described [23]. For the detection of xylanase activity, 0.2% birchwood xylan was included in the gels before polymerization, and zymograms were developed as described [41]. The samples were heated at 50ºC for 15 min in sample buffer before their application to the gels. After electrophoresis, the gels were washed in 2.5% triton X-100 for 30 min, incubated in 200 mM phosphate buffer at pH 6.5 for 30 min, and finally incubated at 55ºC for 10 min in the same pre-warmed fresh buffer. In some cases, phosphate buffer at pH 6.5 was replaced by 200 mM acetate buffer pH 5.0, 200 mM Tris-HCl buffer pH 7.0, or 200 mM Tris-glycine buffer pH 9.0. The gels were stained with 0.1% Congo red for 15 min, washed with 1M NaCl until the xylanase bands became visible, immersed in 10% (v/v) acetic acid, and then photographed.

Binding to insoluble polysaccharides. Binding activity to insoluble polysaccharides was assayed as described by Hogg et al. with some modifications [18]. Briefly, 250 μg of purified Xyn30A was mixed with 25 mg of Avicel or insoluble oat spelt xylan in a final volume of 500 μl of 50 mM phosphate buffer (pH 6.5) in 1.5-ml microcentrifuge tubes. The samples were incubated at 4°C for 1 h with gentle orbital mixing and then centrifuged at 18,000 ×g for 20 min. The supernatants, containing unbound protein, were carefully removed. The pellets were washed three times with 400 μl of the same buffer, resuspended in 400 μl of 10% SDS, and heated at 100ºC for 10 min to release bound protein. The samples were then analyzed by SDS-PAGE on 10–15% polyacrylamide gels.

Analysis of the hydrolysis products from xylan and xylo­ oligosaccharides. Thin-layer chromatography (TLC) was performed as previously described [15]. Xyn30A (1.7 μM) was incubated with 1.5% birchwood or beechwood xylan at 65°C in 50 mM phosphate buffer (pH 6.0) for 18 h. The reactions were stopped by heating at 90°C for 15 min. All samples and markers were adjusted to pH 6.0 before loading. For the analysis of xylan hydrolysis products by matrix-assisted laser desorption-ionization time-offlight mass spectrometry (MALDI-TOF MS), 1 μl of the hydrolysates was mixed with 1 μl of matrix solution [10 mg/ml 2,5- dihydroxybenzoic acid dissolved in acetonitrile-water (1:1, vol/vol), 0.1% (wt/vol) trifluoroacetic acid]. One μl of the mixture was spotted onto the MALDI-TOF MS plate and allowed to dry before the analysis. Positive mass spectra were collected with a 4800 Plus MALDI TOF/TOF (ABSciex 2010) spectrometer with an Nd:YAG 200-Hz laser operated at 355 nm.

Sequence analysis. The 16S rRNA gene sequences of related taxa were obtained from GenBank. Phylogenetic trees were constructed using the neighbor-joining method with the program MEGA 5.0 [37]. BLAST searches were performed for DNA or protein sequence analysis, including domain classification. A putative signal peptide was identified through PrediSi [http:// www.predisi.de] [17]. The 3D structure homology models were generated with SwissModel software [http://swissmodel.expasy.org] based on template gtnA, corresponding to XynC from Bacillus subtilis 168. Pymol software (PyMOL Molecular Graphics System, version 1.2r3pre; Schrödinger) was used to visualize the 3D protein models. The ExPASy proteomics server [http://us.expasy.org/tools/protparam.html] was used to analyze the protein physicochemical parameters (ProtParam tool).

Nucleotide sequence accession number. The DNA sequences of the 16S rRNA gene and xyn30A gene of P. favisporus CC02-N2 were submitted to the GenBank database under accession numbers KF442953 and KF442954, respectively.

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Results and Discussion Isolation and characterization of Paenibacillus favisporus CC02-N2. Strain CC02-N2 was isolated from the soil of a sugarcane plantation in Northeastern Brazil and selected for its high hydrolytic activity on xylan. Culture of the strain on rice-straw-supplemented LB for 48 h resulted in the secretion of 6.95 xylanase U/ml, whereas endoglucanase activity on carboxymethyl cellulose was not detectable. The composition of the xylan-degrading system of the strain was studied by zymographic analysis of the culture supernatants. Several xylanase bands, ranging from 20 to 135 kDa, all of them showing activity in the pH range 5.0–9.0, were detected in the gels (Fig. 1). Based on the multiple xylanase system of the strain CC02-N2 and the high production of xylanase activity, devoid of significant activity on cellulose, it was chosen for further study.

The analysis of its 16S rDNA sequence revealed that strain CC02-N2 belonged to the genus Paenibacillus, with high phylogenetic relatedness to Paenibacillus favisporus GMP01 (99.4% 16S rDNA gene identity) [43] and Paenibacillus cineris LMG18439 (99.3% identity) [25], isolated from cow

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Fig. 1. Zymogram analysis of xylanases from strain CC02-N2. Crude supernatants of strain CC02-N2 cultures in 1% rice-straw-supplemented LB. The zymograms were performed at: (1) pH 5.0; (2) pH 7.0; and (3) pH 9.0. (M) position of the mass standards.

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6.8, respectively. Sequence analysis by comparison to proteins contained in the NCBI database showed that the cloned enzyme was a single-domain xylanase belonging to GH30.

The cloned enzyme, named Xyn30A, showed high homology to hypothetical xylanases from Paenibacillus terrae HPL-003 [32] and Paenibacillus polymyxa E681 [21] (84% and 82% identity, respectively) and to the GH30 domains of the well characterized Xyn30D from P. barcinonensis and XynC from B. subtilis (81% and 80% identity, respectively), whereas homology to Erwinia chrysanthemi XynA was lower (39% identity).

Fig. 2. SDS-PAGE analysis of purified Xyn30A. (1) Cell extracts, (2) active fraction from His-Trap chromatography, (3) purified Xyn30A from Superdex 200/10-300 gel filtration columns, (4) zymogram of xylanase activity, (M) position of the mass standards.

feces and Antarctic volcanic soils, respectively. A phylogenetic tree illustrating the relationship of strain CC02-N2 to closely related species was constructed (data not shown). In accordance with its 16S rRNA gene identity to known species, the isolated strain was tentatively classified as Paenibacillus favisporus CC02-N2, although additional analysis will be required for its full taxonomic characterization.

Cloning and sequence analysis of Xyn30A. The above-described results suggested that Paenibacillus favisporus strain CC02-N2 secreted xylanases from different GH families, as reported for Paenibacillus barcinonensis [5,15, 42]. Among these enzymes, a glucuronoxylanase from family GH30 was recently characterized [42]. In light of the novelty of xylanases specific for MeGlcA branched xylan and their synergism with GH11 xylanases in pulp bleaching [14], we searched for these enzymes in P. favisporus CC02-N2. Using degenerate primers deduced from P. barcinonensis Xyn30D, a fragment of the coding region of a xylanase from P. favisporus CC02-N2 was amplified. By gene walking of this DNA fragment, the sequence of a 1,432-bp DNA fragment containing a 1,290-bp open reading frame (ORF) encoding a xylanase of 430 amino acids was determined. This ORF has an N-terminal region of 29 amino acids with the features of a signal peptide. The predicted molecular mass and isoelectric point of the deduced mature protein were 47,972.9 Da and

Characterization of Xyn30A. Xyn30A was cloned into plasmid (pLATE31) under the control of the T7 promoter in E. coli BL21 Star (DE3) to produce a fusion protein of fulllength Xyn30A and a C-terminal His tag. The recombinant protein Xyn30A was purified to homogeneity from cell extracts by two-step chromatography using HisTrapHP and Superdex 200/10-300 columns. The purified Xyn30A was analyzed by SDS-PAGE and zymography, which showed that it had an apparent molecular mass of 47.9 kDa, in accordance with that deduced from the amino acid sequence (Fig. 2).

The substrate specificity of Xyn30A was determined by evaluating the activity of the enzyme on xylans and other poly­saccharides. The enzyme showed high hydrolytic activity on glucuronoxylans. Beechwood xylan was the preferred substrate, yielding a specific activity of 244 xylanase U/mg (Table 1). By contrast, the enzyme did not show detectable activity on arabinoxylans from oat spelt, rye, and wheat, nor did it show activity on any of the other polysaccharides tested, including crystalline or amorphous celluloses. The substrate specificity of Xyn30A was similar to that of the four GH30 glucuronoxylanases characterized to date: E. chrysanthemi Table 1. Substrate specificity of Xyn30A Substrate

Activity (U/mg)

Beechwood xylan

244.0 ± 4.1

Birchwood xylan

202.5 ± 11.4

4-O-methyl-glucuronoxylan

225.1 ± 18.3

Oat spelt xylan

NDa

Wheat arabinoxylan

ND

Rye arabinoxylan

ND

Carboxymethyl cellulose

ND

Avicel

ND

ND, not detected.

a

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Fig. 3. Response surface plot showing the effect of temperature and pH on the xylanase activity of Xyn30A. The dots represent the averages of three replicates.

RSM (Fig. 3). The conditions that led to maximum activity were 65ºC and pH 6.0. Thermostability assays showed that Xyn30A remained highly stable up to 50ºC after 3 h of incubation at pH 6.0; about 60% of the original activity was maintained after incubation under these conditions but at at 60ºC (Fig. 4).

An analysis of the kinetic variables of Xyn30A on beech-

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XynA [19,45], B. subtilis XynC [36], Xyn30B from Bacillus sp. BP-7 [14], and P. barcinonensis Xyn30D [42]. All of these enzymes are specific for glucuronoxylans, as a result of their requirement for MeGlcA branches for catalysis, and are not active on arabinoxylans.

The effect of temperature and pH on xylanase activity and the interaction between the two factors were determined by

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Fig. 4. Effect of temperature and pH on the stability of Xyn30A. (A) The samples were incubated in 50 mM phosphate buffer (pH 6.0) at 50ºC (■), 60ºC (▲) and 70ºC (□), and residual activity after different time intervals was determined. (B) The samples (○) were incubated at 65ºC in buffers at different pH for 3 h, after which residual activity was determined.

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Table 2. Effect of metal ions and other compounds on the activity of Xyn30A Relative activity (%) Control

100 ± 1.4

Al

0.0

3+

Ba2+

93.2 ± 8.9

Ca2+

86.4 ± 13.4

Cu2+

5.8 ± 0.8

Fe

6.9 ± 0.3

2+

0.6 ± 0.2 106.0 ± 1.2

K+

100.9 ± 5.3

Li+ Mg

2+

103.3 ± 0.5

Mn

2+

38.5 ±0.6

Na+

96.5 ± 0.8

NH4+

101.4 ± 3.4

Zn2+

7.7 ± 0.5

EDTA

99.1 ± 4.0

SDS

6.1 ± 3.1

Tween 80

26.6 ± 8.2

Triton X-100

11.5 ± 0.0

DMSO

92.4 ± 1.0

wood xylan showed a Vmax of 267.2 U mg–1 and a Km of 4.0 mg/ml at optimal conditions for activity. Comparison of the kinetic constants of Xyn30A with those reported for GH30 xylanases showed that the Km of Xyn30A was higher than that of XynC from B. subtilis (1.63 mg/ml) but lower than that of Xyn30D from P. barcinonensis (14.72 mg/ml). A larger difference was found among the kcat values of these enzymes. Thus, while B. subtilis XynC and P. barcinonensis Xyn30D had kcat values of 2,635 and 1,510 min–1, respectively, the kcat of Xyn30A was 13,333 min–1, much higher than the values of the GH30 xylanases characterized so far.

The effect of different metal ions and chemicals on Xyn30A activity was also determined (Table 2). The enzyme was completely inhibited by Al3+ and strongly inhibited by Hg2+, Cu2+, Fe2+, Zn2+, Mn2+, Triton X-100, Tween 80, and SDS. Inhibition by Ba2+, Ca2+, Na+, and DMSO was minimal. Li+, NH4+, and EDTA had no effect on enzyme activity, whereas K+ and Mg2+ produced a small stimulating effect. The inhibitory effect of Al3+ on Xyn30A resembles that reported for sev-

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Hg

2+

Fig. 5. Computer modeling of the 3D structure of Xyn30A. The (β/a)8 barrel is shown in cyan and the associated b-domain in yellow. The numbering of the bs strands and amino acids is indicated.

eral amylases, in which aluminum ions were shown to inactivate the enzymes, presumably by binding to catalytic residues [2,29].

All GH30 glucuronoxylanases share a unique overall 3D structure of the catalytic module, which consists of a (β/a)8 barrel fused to a side β-structure of nine strands, referred to as the side-associated β-domain [24,34]. Computer modeling of Xyn30A based on the crystalline structure of B. subtilis XynC showed a similar 3D structure of a (β/a)8 barrel with an associated β-domain (Fig. 5). Valenzuela et al [42] reported that deletion of this side structure abolishes the activity of P. barcinonensis Xyn30D, evidence of the importance of the β-side structure for the catalytic activity of GH30 xylanases. A comparison to sequences of Bacillus subtilis XynC and Erwinia chrysanthemi XynA identified the glutamic acid residues (Glu177 and Glu266) in the catalytic groove of Xyn30A. These residues are thought to provide the catalytic acid/base and nucleophile of GH30 glucuronoxylanases (Fig. 5). The two enzymes also contain a conserved arginine, Arg303 in B. subtilis XynC and Arg293 in E. chrysanthemi XynA, responsible for MeGlcA side-chain recognition in glucuronoxylan, which determines the substrate specificity of these xylanases [35,40]. The conserved arginine was also found in the corresponding location (Arg309) of Xyn30A. Two recently characterized GH30 xylanases from fungal microorganisms differ from the aforementioned enzymes in their activity on both

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Products of xylan degradation by Xyn30A. The hydrolysis products from glucuronoxylans were analyzed by TLC. Beechwood xylan was degraded to a mixture of products of intermediate mobility between linear xylooligosaccharides, indicating that they were MeGlcA-substituted xylooligomers, while xylose and linear oligosaccharides were not found among the hydrolysis products (Fig. 6). To better characterize the products of beechwood xylan hydrolysis, they were also analyzed by MALDI-TOF MS. The mass spectra showed the presence of molecular ions of substituted xylooligomers and their sodium salts, identified as sodium adducts (Table 3). The major ions corresponded to substituted xylooligosaccharides consisting of 3â&#x20AC;&#x201C;8 xylopyranosyl residues and a single MeGlcA residue, in accordance with the results of the TLC analysis. The results also agree with the mode of action of GH30 xylanases, which cleave the bonds at the second position after the MeGlcA branches, giving rise to xyloligosaccharides with a substitution at the penultimate xylose residue from the reducing end [36,45].

Binding to polysaccharides. Binding of Xyn30A to Avicel and insoluble oat spelt xylan was studied as described in Materials and methods. As expected, Xyn30A did not bind to Avicel. Most of the enzyme remained in the supernatants and only a very small amount was detected adsorbed on Avicel. By contrast, the enzyme bound to insoluble oat spelt xylan, as most of the enzyme was found adsorbed on the polymer while only a small amount remained in the supernatants (Fig. 7). Xyn30A is a single-domain enzyme devoid of a carbohydrate binding module (CBM). However, our results clearly showed that the enzyme bound to insoluble xylans, implying a polysaccharide binding ability of its catalytic GH30 domain. Reported studies of xylan binding of P. barcinonensis Xyn30D, a modular GH30 xylanase that includes a CBM35, have shown that not only the whole enzyme and its CBM but also the isolated GH30 domain binds to insoluble xylan [42]. Polysaccharide binding of single-domain xyla-

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Table 3. Oligosaccharides identified by MALDI-TOF MS after the hydrolysis of beechwood xylan by Xyn30A

m/z Xylooligomer

Mra

MeGlcAX2

472

â&#x20AC;&#x201C;

MeGlcAX3

604

627.2

MeGlcAX4

736

759.2

MeGlcAX5

868

891.3

MeGlcAX6

1000

1023.3

MeGlcAX7

1132

1155.4

MeGlcAX8

1264

1287.4

Na

Mr, molecular weight.

a

nases was previously reported for Bacillus circulans BcX, a GH11 xylanase thought to contain a secondary xylan-binding site that overcomes the lack of a CBM [26]. The observed binding ability of the GH30 catalytic module could be attributed to substrate binding site of the catalytic groove, or to the b-side associated domain, which as mentioned above seems to be required for the catalytic activity of GH30 xylanases. In

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glucoronoxylan and arabinoxylan [27,39] Alignment of the respective amino acid sequences showed that these fungal enzymes do not have the conserved arginine residue proposed to recognize glucuronoxylan [39]. In a dendrogram analysis of GH30 xylanases, the fungal xylanases clustered separately from glucuronoxylanases. In fact, according to St John et al.

[34], the fungal enzymes can be placed in a different subgroup of GH30 xylanases, acidic xylanases, which are active on glucuronoxylan and arabinoxylan and show an acidic pH optimum for activity [27,39].

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Fig. 6. Thin-layer chromatography analysis of thte hydrolysis products from beechwood xylan. (1) Control, no digested samples; (2) products of xylan hydrolysis by Xyn30A; (M) size markers of xylose (X), xylobiose (X2), xylotriose (X3), xylotetraose (X4), xylopentaose (X6), and xylohexaose (X6).

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fact crystallographic analysis of XynC from Bacillus subtilis identified ligand coordination to its b-side domain, which has been proposed as a new type of CBM [35].

The enzyme characterized in this study is one of the few glucuronoxylanases of the GH30 family described so far. The enzyme’s lack of activity on arabinoxylans has probably hindered the identification of these enzymes, which may complement the activity of GH10 and GH11 xylanases on lignocellulosic biomass. The recent identification of several GH30 glucuronoxylanases, including P. favisporus Xyn30A, suggests that these enzymes are widespread among gram-positive xylanolytic bacteria [14,42]. Their unique mode of action on glucuronoxylans suggests their use to selectively modify xylan and in the design of tailored oligomers. These MeGlcAsubstituted xylooligosaccharides (acidic XOS) have also been proposed as emerging prebiotics, as they were shown to selectively stimulate the growth of probiotic microorganisms, including Bifidobacterium, which ferment them to short-chain fatty acids [1,30]. Hardwood xylan can provide a source of acidic XOS as food additives and nutraceuticals [16,28]. In those reports, xylan was depolymerized by chemical means, and the resulting XOS was refined to eliminate subproducts such as xylose and other monosaccharides. Enzymatic treatment as an alternative for XOS production could diminish subproduct formation. In addition, GH30 glucuronoxylanases are unique because xylose is not released from xylan, thus avoiding further processing of the hydrolysates. The high kcat of Xyn30A makes this enzyme of particular interest among GH30 xylanases for the production of acidic XOS and for the transformation of biomass. Glucuronoxylanases can be important tools in biorefinery approaches to obtain added-value products from abundant raw materials, such as Eucalyptus

Fig. 7. SDS-PAGE analysis of the binding of Xyn30A to insoluble polysaccharides. Proteins were mixed with Avicel or with the insoluble fraction of oat spelt xylan for 1 h; bound and unbound fractions were separated by centrifugation and analyzed by SDSPAGE. (1) Unbound fraction, (2) wash, (3) fraction absorbed to the polymer, (C) control protein, (M) position of the mass standards.

wood, whose potential based on its high xylan content is usually underestimated. Further research is required to ascertain the role of GH30 glucuronoxylanases on xylan hydrolysis.

Analysis of the function of their distinctive b-side domain in substrate recognition and catalysis will provide insights into its contribution to xylan degradation in natural habitats.

Acknowledgments. This study was supported in part by the Spanish Ministry of Science and Innovation, grant CTQ2010-20238-C03-02, and AGAUR from Generalitat de Catalunya (Autonomous Government of Catalonia), grant 2009 SGR 819. Itácio Padilha was the recipient of an SWE grant from the Science without Borders Program of the Brazilian Council for Research (CNPq), and the Coordination for Higher Education Staff Development (CAPES). Demetrius Araújo is a CNPq fellow.

Competing interests. None declared.

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6. Bradford MM (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72:248-254 7. Britton HTS (1952) Hydrogen ions, 2nd ed. London 8. Bruce T, Martinez IB, Maia Neto O, Vicente ACP, Kruger RH, Thompson FL (2010) Bacterial community diversity in the Brazilian Atlantic forest soils. Microb Ecol 60:840-849 9. Cantarel BL, Coutinho PM, Rancurel C, Bernard T, Lombard V, Henrissat B (2009) The Carbohydrate-Active EnZymes database (CAZy): an expert resource for glycogenomics. Nucleic Acids Res 37:233-238 10. Collins T, Gerday C, Feller G (2005) Xylanases, xylanase families and extremophilic xylanases. FEMS Microbiol Rev 29:3-23 11. Collins T, Meuwis M-A, Stals I, Claeyssens M, Feller G, Gerday C (2002) A novel family 8 xylanase, functional and physicochemical characterization. J Biol Chem 277:35133-35139 12. Correia MAS, Mazumder K, Brás JLA, Firbank SJ, Zhu Y, Lewis RJ, York WS, Fontes CMGA, Gilbert HJ (2011) Structure and function of an arabinoxylan-specific xylanase. J Biol Chem 286:22510-22520 13. Fushinobu S, Hidaka M, Honda Y, Wakagi T, Shoun H, Kitaoka M (2005) Structural basis for the specificity of the reducing end xylose-releasing exo-oligoxylanase from Bacillus halodurans C-125. J Biol Chem 280:17180-17186 14. Gallardo O, Fernández-Fernández M, Valls C, Valenzuela SV, Roncero MB, Vidal T, Díaz P, Pastor FIJ (2010) Characterization of a family GH5 xylanase with activity on neutral oligosaccharides and evaluation as a pulp bleaching aid. Appl Environ Microbiol 76:6290-6294 15. Gallardo O, Pastor FIJ, Polaina J, Diaz P, Łysek R, Vogel P, Isorna P, González B, Sanz-Aparicio J (2010) Structural insights into the specificity of Xyn10B from Paenibacillus barcinonensis and its improved stability by forced protein evolution. J Biol Chem 285:2721-2733 16. Gullón P, González-Muñoz MJ, Gool MP van, Schols HA, Hirsch J, Ebringerová A, Parajó JC (2011) Structural features and properties of soluble products derived from Eucalyptus globulus hemicelluloses. Food Chem 127:1798-1807 17. Hiller K, Grote A, Scheer M, Münch R, Jahn D (2004) PrediSi: prediction of signal peptides and their cleavage positions. Nucleic Acids Res 32:375-379 18. Hogg D, Pell G, Dupree P, Goubet F, Martín-Orúe SM, Armand S, Gilbert HJ (2003) The modular architecture of Cellvibrio japonicus mannanases in glycoside hydrolase families 5 and 26 points to differences in their role in mannan degradation. Biochem J 371:1027-1243 19. Hurlbert JC, Preston JF 3rd (2001) Functional characterization of a novel xylanase from a corn strain of Erwinia chrysanthemi. J Bacteriol 183: 2093-2100 20. Juturu V, Wu JC Microbial xylanases: engineering, production and industrial applications. Biotechnol Adv 30:1219-1227 21. Kim JF, Jeong H, Park S-Y, Kim S-B, Park YK, Choi S-K, Ryu C-M, Hur C-G, Ghim S-Y, Oh TK, Kim JJ, Park CS, Park S-H (2010) Genome sequence of the polymyxin-producing plant-probiotic rhizobacterium Paenibacillus polymyxa E681. J Bacteriol 192:6103-6104 22. Kumar S, Singh SP, Mishra IM, Adhikari DK (2009) Recent advances in production of bioethanol from lignocellulosic biomass. Chem Eng Technol 32:517-526 23. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685 24. Larson SB, Day J, Barba de la Rosa AP, Keen NT, McPherson A (2003) First crystallographic structure of a xylanase from glycoside hydrolase family 5: implications for catalysis. Biochemistry 42:8411-8422

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Volume 17(3) SETEMBER 2014

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Official journal of the Spanish Society for Microbiology Volume 17 · Number 3 · September 2014

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Ariza-Miguel J, Hernández M, FernándezNatal I, Rodríguez-Lázaro D Molecular epidemiology of methicillinresistant Staphylococcus aureus in a university hospital in northwestern Spain

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Spricigo DA, Cortés P, Moranta D, Barbé J, Bengoechea JA, Llagostera M Significance of tagI and mfd genes in the virulence of non-typeable Haemophilus influenzae

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Padilha IQM, Valenzuela SV, Grisi TCSL, Diaz P, de Araújo DAM, Pastor FIJ A glucuronoxylan-specific xylanase from a new Paenibacillus favisporus strain isolated from tropical soil of Brazil

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INTERNATIONAL MICROBIOLOGY www.im.microbios.org

2014 pp 131-184

Manrique-Ramírez P, Galofré-Milà N, Serrano M, Aragon V Identification of a class B acid phosphatase in Haemophilus parasuis

Number 3

López-Martínez G, Borrull A, Poblet M, Rozès N, Cordero-Otero R Metabolomic characterization of yeast cells after dehydration stress

Chiellini C, Maida I, Emiliani G, Mengoni A, Mocali S, Fabiani A, Biffi S, Maggini V, Gori L,Vannacci A, Gallo E, Firenzuoli F, Fani R Endophytic and rhizospheric bacterial communities isolated from the medicinal plants Echinacea purpurea and Echinacea angustifolia

Volume 17

RESEARCH ARTICLES

International Microbiology

INTERNATIONAL MICROBIOLOGY

Volume 17 · Number 3 · September 2014 · ISSN 1139-6709 · e-ISSN 1618-1905

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17(3) 2014 INDEXED IN

Agricultural and Environmental Biotechnology Abstracts; ASFA/Aquatic Sciences & Fisheries Abstracts; BIOSIS; CAB Abstracts; Chemical Abstracts; SCOPUS; Current Contents®/Agriculture, Biology & Environmental Sciences®; EBSCO; EMBASE/Elsevier Bibliographic Databases; Food Science and Technology Abstracts; ICYT/CINDOC; IBECS/BNCS; ISI Alerting Services®; MEDLINE®/Index Medicus®; Latindex; MedBioWorldTM; SciELO-Spain; Science Citation Index Expanded®/SciSearch® September 2014

Official journal of the Spanish Society for Microbiology