International Microbiology

Volume 15 · Number 1 · March 2012 · ISSN 1139-6709

Official journal of the Spanish Society for Microbiology Volume 15 · Number 1 · March 2012

International Microbiology

INTERNATIONAL MICROBIOLOGY

Volume 15 Number 1

RESEARCH ARTICLES

Heindl H, Thiel V, Wiese J, Imhoff JF Bacterial isolates from the bryozoan Membranipora membranacea: influence of culture media on isolation and antimicrobial activity

33

9

Mariscotti JF, Quereda JJ, Pucciarelli MG Contribution of sortase A to the regulation of Listeria monocytogenes LPXTG surface proteins

43

pp 1-54

Chen P, Yan L, Wang Q, Li Y, Li H Surface alteration of realgar (As4S4)

by Acidithiobacillus ferrooxidans

1

www.im.microbios.org

2012

Schinke C, Germani JC Screening Brazilian Macrophomina phaseolina isolates for alkaline lipases and other extracellular hydrolases

García-Maldonado JQ, Bebout BM, Celis LB, López-Cortés A Phylogenetic diversity of methyl-coenzyme M reductase (mcrA) gene and methanogenesis from trimethylamine in hypersaline environments

INTERNATIONAL MICROBIOLOGY

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15(1) 2012 INDEXED IN

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Published Quarterly by • March 2012

Official journal of the Spanish Society for Microbiology

Publication Board Editorial Board Editor-in-chief Carles Pedrós-Alió, Institute of Marine Sciences-CSIC Associate Editors Mercedes Berlanga, University of Barcelona Mercè Piqueras, International Microbiology Wendy Ran, International Microbiology Secretary General Ricard Guerrero, University of Barcelona, Institute for Catalan Studies Adjunct Secretary and Webmaster Nicole Skinner, International Microbiology Managing Coordinator Carmen Chica, International Microbiology Members Teresa Aymerich, University of Girona Susana Campoy, Autonomous University of Barcelona Jesús García-Gil, University of Girona Josep Guarro, Rovira i Virgili University Enrique Herrero, University of Lleida Emili Montesinos, University of Girona José R. Penadés, Institute of Mountain Livestock-CSIC Jordi Vila, University of Barcelona Jordi Urmeneta, University of Barcelona Addresses Editorial Office International Microbiology Poblet, 15 08028 Barcelona, Spain Tel. & Fax +34-933341079 E-mail: int.microbiol@microbios.org www.im.microbios.org Spanish Society for Microbiology Vitruvio, 8 28006 Madrid, Spain Tel. +34-915613381. Fax +34-915613299 E-mail: sem@microbiologia.org www.semicrobiologia.org Publisher Viguera Editores, S.L. Plaza Tetuán, 7 08010 Barcelona, Spain Tel. +34-932478188. Fax +34-932317250 E-mail: info@viguera.com; www.viguera.com © 2012 Spanish Society for Microbiology & Viguera Editores, S.L. Printed in Spain Print ISSN: 1139-6709 Online ISSN: 1618-1095 D.L.: B.23341-2004 With the collaboration of the Institute for Catalan Studies

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Ricardo Amils, Autonomous University of Madrid, Madrid, Spain Albert Bordons, Rovira i Virgili University, Tarragona, Spain Albert Bosch, University of Barcelona, Barcelona, Spain Enrico Cabib, National Institutes of Health, Bethesda, MD, USA Victoriano Campos, Pontificial Catholic University of Valparaíso, Chile Josep Casadesús, University of Seville, Sevilla, Spain Yehuda Cohen, The Hebrew University of Jerusalem, Jerusalem, Israel Rita R. Colwell, Univ. of Maryland & Johns Hopkins University, MD, USA Katerina Demnerova, Inst. of Chem. Technology, Prague, Czech Republic Esteban Domingo, CBM, CSIC-UAM, Cantoblanco, Madrid, Spain Mariano Esteban, Natl. Center for Biotechnol., CSIC, Cantoblanco, Spain M. Luisa García López, University of León, León, 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 Paloma Liras, University of León, León, Spain Ruben López, Center for Biological Research, CSIC, Madrid, Spain Juan M. López Pila, Federal Environ. Agency, Dessau-Roßlau, Germany Michael T. Madigan, Southern Illinois University, Carbondale, IL, USA M. Benjamín Manzanal, University of Oviedo, Oviedo, Spain 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 José Olivares, Experimental Station of Zaidín, CSIC, Granada, Spain Juan A. Ordóñez, Complutense University of Madrid, Madrid, Spain Eduardo Orías, University of California-Santa Barbara, CA, USA José M. Peinado, Complutense University of Madrid, Madrid, Spain J. Claudio Pérez Díaz, Ramón y Cajal Institute Hospital, Madrid, Spain Antonio G. Pisabarro, Public University of Navarra, Pamplona, Spain Carmina Rodríguez, Complutense University of Madrid, Madrid, Spain Manuel de la Rosa, Virgen de las Nieves Hospital, Granada, Spain Tomás A. Ruiz Argüeso, Technical University of Madrid, Spain Hans G. Schlegel, University of Göttingen, Germany James A. Shapiro, University of Chicago, IL, USA John Stolz, Duquesne University, Pittsburgh, PA, USA James Strick, Franklin & Marshall College, Lancaster, PA, USA Jean Swings, Ghent University, Ghent, Belgium Gary A. Toranzos, University of Puerto Rico, San Juan, Puerto Rico Antonio Torres, University of Seville, Sevilla, Spain Josep M. Torres-Rodríguez, Municipal Inst. Medical Research, Barcelona José A. Vázquez-Boland, University of Edinburgh, Edinburgh, UK Antonio Ventosa, University of Seville, 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

CONTENTS INTERNATIONAL MICROBIOLOGY (2012) 15:1-54 ISSN 1139-6709 www.im.microbios.org

Volume 15, Number 1, March 2012

RESEARCH ARTICLES

Schinke C, Germani JC Screening Brazilian Macrophomina phaseolina isolates for alkaline lipases and other extracellular hydrolases

1

Chen P, Yan L, Wang Q, Li Y, Li H Surface alteration of realgar (As4S4) by Acidithiobacillus ferrooxidans

9

Heindl H, Thiel V, Wiese J, Imhoff JF Bacterial isolates from the bryozoan Membranipora membranacea: influence of culture media on isolation and antimicrobial activity

17

García-Maldonado JQ, Bebout BM, Celis LB, López-Cortés A Phylogenetic diversity of methyl-coenzyme M reductase (mcrA) gene and methanogenesis from trimethylamine in hypersaline environments

33

Mariscotti JF, Quereda JJ, Pucciarelli MG Contribution of sortase A to the regulation of Listeria monocytogenes LPXTG surface proteins

43

A1

Spanish Society for Microbiology 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. It’s 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 approximately 1700 members working in different fields of microbiology.

International Microbiology Aims and scope INTERNATIONAL MICROBIOLOGY, the official journal of the SEM, is a peerreviewed, open access journal whose aim is to advance and disseminate information in the fields of basic and applied microbiology among scientists around the world. The journal publishes research articles and complements (short papers dealing with microbiological subjects of broad interest such as editorials, perspectives, book reviews, etc.). A feature that distinguishes it from many other microbiology journals is a broadening of the term “microbiology” to include eukaryotic microorganisms (protists, yeasts, molds), as well as the publication of articles related to the history and sociology of microbiology.

INTERNATIONAL MICROBIOLOGY offers high-quality, internationallybased information, short publication times (< 3 months), complete copy-

editing service, and online open access publication available to any reader prior to distribution of the printed journal.

The journal encourages submissions in the following areas: • Microorganisms (prions, viruses, bacteria, archaea, protists, yeasts, molds) • Microbial biology (taxonomy, genetics, morphology, physiology, ecology, pathogenesis) • Microbial applications (environmental, soil, industrial, food and medical microbiology, biodeterioration, bioremediation, biotechnology) • Critical reviews of new books on microbiology and related sciences are also welcome.

Jounal Impact Factor The 5-Year Journal Impact Factor (VIF) of INTERNATIONAL MICROBIOLOGY is 2.928.

The journal is covered in several leading abstracting and indexing databases, including the following ones: AFSA Marine Biotechnology Abstracts; Biological Abstracts; Biotechnology Research Abstracts; BIOSIS Previews; CAB Abstracts; Chemical Abstracts; Current Contents – Agriculture, Biology & Environmental Sciences; EBSCO; Embase; Food Science and Technology Abstracts; Google Scholar; IEDCYT; IBECS; Latíndex; MedBioWorld; PubMed; SciELO-Spain; Science Citation Index Expanded; Scopus

Cover legends Front cover CENTER. View of a raised microbial mat growing on top of highly sulfidic sediments in an evaporitic flat in Laguna San Ignacio, Baja California Sur, Mexico. The irregular pustular morphology is due to the accumulation of gas bubbles, which in those emanating from the sediment were found to contain methane. Phylogenetic analyses of the samples revealed that the methanogen community was dominated by moderately halophilic members of the genus Methanohalophilus. [See article by García-Maldonado et al., pp. 33-41, this issue.] (Scale ca. 1:30) UPPER LEFT. Particles of human immunodefficiency virus type 1 (HIV1) budding from a lymphoid infected cell. The structural protein Gag oligomerizes in the inner leaflet of the plasma membrane to generate new HIV particles. Immature particles are characterized by their circular outlines, and mature HIV-1 virions by inner dense areas. Micrograph

by M. Teresa Fernández-Figueras, and Julià Blanco, Hospital Trias i Pujol, Badalona, Spain. (Magnification, ca. 60,000×) UPPER RIGHT. Typical position of filaments in a mature colony of Nostoc punctiforme Kützing (Hariot), isolated from a temporarily inundated soil. The thallus is microscopic, gelatinous, and changes during development. N. punctiforme is able to fix nitrogen in heterocysts, distinguished from vegetative barrel-shaped cells by their thickwalls and pale aspect. Isolation and micrograph by Mariona Hernández Mariné, University of Barcelona, Spain. (Magnification, ca. 1000×) LOWER RIGHT. Giemsa-stained promastigotes of Leishmania infantum.

This flagellated form of the protist occurs in the insect vector.

Following inoculation into their human hosts, promastigotes enter macrophages, where they develop into amastigotes (the non-flagellated form) before multiplying. Micrograph by Roser Fisa and Cristina Riera, University of Barcelona, Spain. (Magnification, ca. 2000×) LOWER LEFT. Low-temperature scanning electron micrograph of mycobiont hyphae from the lichen Xanthoria elegans exposed to space conditions in the BIOPAN-5 facility of the European Space Agency. Lichenized fungal and algal cells survived in space after full exposure to massive UV, cosmic radiation and high vacuum. Image by Carmen Ascaso and Asunción de los Ríos (MNCN, CSIC, Madrid).

(Magnification, ca. 1900×)

Back cover Portrait and signature of Antonio Vargas Reyes (1816–1873), Colombian pioneer of medicine and public health. Vargas Reyes belonged to a family linked to the development and progress of medicine in

Colombia throughout the nineteenth century. He was born in Charala in 1816, into wealthy family that, shortly afterwards, lost its fortune, when Colombia became independent (1819) and the family was accused of having backed the Spanish crown. The family moved to Bogota, but 5-year-old Antonio was left in the care of a priest, who was supposed to educate him but instead beat him and kept him illiterate, as well as close to starvation. He was rescued at the age of 12, by his older, married sister, who took him to Bogota, where they joined their widowed mother. He soon recovered the years that were lost with no education, and in 1834 began to study medicine. Vargas Reyes may have been poverty stricken—sometimes he could not even afford shoes, not to mention textbooks—but he was certainly the most brilliant student in his class, and was often bullied by jealous fellow classmates. His situation greatly improved after the Rector of the University appointed him as an assistant in the anatomy classes. After his graduation in 1838, he worked briefly in the northern provinces of Colombia before joining the revolutionary army. Along with Antonio Vargas Vega, he helped their common relative Jorge Vargas Suárez to organize the pox vaccine campaign during the 1840–1841 epidemics. This was the first step in the eventual creation of the Instituto Central de Propagación de la Vacuna (Central Institute for the Dissemination of Vaccination), in 1856. In 1842, Vargas Reyes went to Paris, where he attended courses in various medical specialties but also chemistry and botany. In 1845, after receiving a permit to work as a physician in France, he traveled to England, Italy, and Spain. Upon his return to Colombia in 1847, the country’s President, with the support of influential supporters, paid him an annual salary of 4000 pesos to ensure that he did not leave Colombia. In 1849, when a cholera pandemic reached several towns along the Colombian Atlantic shoreline, Vargas Reyes published a public health monography whose aim was to establish rules for the prevention and treatment of cholera. In the following decades, Vargas Reyes taught various medical subjects and cofounded the first Colombian scientific journals—La Lanceta (AprilOctober, 1852) and La Gaceta Médica (July, 1864–December, 1867), as well as a private School of Medicine (1865) that would be the basis for the Faculty of Medicine of the National University, founded in 1867. In 1872, he went to Europe. He returned in 1873, retiring to Villeta, where he died on 23 August. A physician from the pre-bacteriological era, Vargas Reyes studied fevers and grouped them in families. Although unaware of their microbial origin, he distinguished between fever as a disease symptom and fever as a disease in itself. His efforts towards the establishment of medicine as a profession in Colombia and the foundation of a School of Medicine have been widely recognized.

Front cover and back cover design by MBerlanga & RGuerrero

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RESEARCH ARTICLE INTERNATIONAL MICROBIOLOGY (2012) 15:1-7 DOI: 10.2436/20.1501.01.153 ISSN: 1139-6709 www.im.microbios.org

Screening Brazilian Macrophomina phaseolina isolates for alkaline lipases and other extracellular hydrolases Claudia Schinke,* José C. Germani Department of Raw Materials Production, Faculty of Pharmacy, Federal University of Rio Grande do Sul, Porto Alegre, Brazil Received 27 December 2011 · Accepted 25 February 2012

Summary. Macrophomina phaseolina, phylum Ascomycota, is a phytopathogenic fungus distributed worldwide in hot dry areas. There are few studies on its secreted lipases and none on its colony radial growth rate, an indicator of fungal ability to use nutrients for growth, on media other than potato-dextrose agar. In this study, 13 M. phaseolina isolates collected in different Brazilian regions were screened for fast-growth and the production of hydrolases of industrial interest, especially alkaline lipases. Hydrolase detection and growth rate determination were done on citric pectin, gelatin, casein, soluble starch, and olive oil as substrates. Ten isolates were found to be active on all substrates tested. The most commonly detected enzymes were pectinases, amylases, and lipases. The growth rate on pectin was significantly higher (P < 0.05), while the growth rates on the different media identified CMM 2105, CMM 1091, and PEL as the fastest-growing isolates. The lipase activity of four isolates grown on olive oil was followed for 4 days by measuring the activity in the cultivation broth. The specific lipolytic activity of isolate PEL was significantly higher at 96 h (130 mU mg protein–1). The broth was active at 37 °C, pH 8, indicating the potential utility of the lipases of this isolate in mild alkaline detergents. There was a strong and positive correlation (0.86) between radial growth rate and specific lipolytic activity. [Int Microbiol 2012; 15(1):1-7] Keywords: Macrophomina phaseolina · pectinases · amylases · proteases · lipolytic activity · radial growth rate

Introduction Enzymes are an important group of biological products used in several processes in the food industry and in environmental and industrial biotechnological applications [23]. As biocatalysts, they have many advantages over chemical cata-

*Corresponding author: C. Schinke Laboratório de Tecnologia Bioquímica, Faculdade de Farmácia Universidade Federal do Rio Grande do Sul Av. Ipiranga 2752 sala 707 Porto Alegre, RS, CEP 90610-000, Brazil Tel./Fax +55-5133085354 E-mail: claudia_schinke@yahoo.com.br

lysts: the ability to function under relatively mild conditions of temperature, pH, and pressure; their specificity, and in some cases, their stereoselectivity. In addition, they produce no unwanted by-products [40]. Lipases are of particular interest because of their many applications in oleochemistry, organic synthesis, the detergent industry, and nutrition [30].

Indeed, the single biggest market for enzymes is in detergent formulations [27].

Fungi are excellent sources of enzymes as they produce these biocatalysts in great variety [8,31]. Macrophomina phaseolina (Tassi) Goid. [http://nt.ars-grin.gov/fungaldatabases, accessed Feb. 24, 2012] is a phytopathogenic filamentous fungus belonging to the anamorphic Ascomycota,

2

INT. MICROBIOL. Vol. 15, 2012

Botryosphaeriaceae family [5,10], producing both sclerotia and pycnidia. It is responsible for the plant disease called charcoal rot, which affects both the roots and the stems.

There are also reports of the fungus causing human ocular infection, skin infection in an immunocompromised child, and granuloma in a cat [6,14,37]. Macrophomina phaseolina is widely distributed in tropical regions and specifically in areas subjected to water stress, where it infects hundreds of different hosts [36] and causes severe economic losses [34].

In Brazil, M. phaseolina is found from the Northeastern region, where the climate is mostly hot and dry, to the South, where the humidity is high and temperatures range from 30 °C in the summer to 5 °C in winter. The microorganism penetrates host tissues through mechanical pressure exerted by the spore germ tube and the sclerotia hyphae, as well as through dissolution of the cell wall through processes mediated by secreted enzymes [4].

The plant cell wall is a complex structure of polymers that surrounds the cell. M. phaseolina produces cellulolytic, hemicellulolytic, pectolytic, and proteolytic extracellular enzymes [2], as well as lipases. Their concerted action results in the breakdown of the main polymeric components of the cell wall and the cell membrane. Studies on extracellular enzymes produced by M. phaseolina are few, and the most recent ones focused almost exclusively on cellulases and endoglucanases [1,7,29,41–43]. The colony radial growth rate reflects the ability of the fungus to use a particular substrate for growth, by secreting the necessary enzymes and thus enabling nutrient uptake for fungal metabolism and cell multiplication. Thus far, only one study examined the relative growth rate of this phytopathogen, on potato-dextrose agar [18], whereas radial growth rates on other substrates have not been reported. The lipolytic activity of M. phaseolina, involving one or two isolates, has been described in only a few studies [2,16,28].

The objective of the present work was to screen wild-type M. phaseolina collected in Brazil for fast growing isolates that produce hydrolases of industrial interest, and especially to select those producing alkaline lipases in large amounts.

Materials and methods Equipment and reagents. Reagents and cultivation media were of the purest grade available, bought from Himedia (India), Merck (Germany), Vetec and Nuclear (Brazil). Extra-virgin olive oil was of commercial grade.

A Minisart (Sartorius) filter, porosity 0.2 μm, was used for filter sterilizations. The rotatory shaker was from Oxylab (Brazil), and the spectrophotometer was from Shimadzu UV Mini-1240.

SCHINKE ET AL.

Macrophomina phaseolina isolates. Isolates CMM 527, CMM 932, CMM 979, CMM 1048, CMM 1091, CMM 2100, CMM 2105, collected in the northeastern region of Brazil, were provided by the Phytopathogenic Fungi Culture Collection Prof. Maria Menezes of the Federal Rural University of Pernambuco (UFRPe). Isolates MMBF 564, MMBF 16–98, collected in the northeastern region, and MMBF 808, MMBF 04–10, collected in the southeastern region, were from the Fungi Collection Mário Barreto Figueiredo of the Biological Institute of the Department of Agriculture and Supply of the State of São Paulo (IB-SP). Isolate PEL, collected in the southern region, was obtained from the Phytosanitary Department of the Federal University of Pelotas (UFPel). Isolate AJAM, collected in the southeastern region, was from the Phytopathology Department of the Federal University of Viçosa.

Isolates maintenance. Isolates were cultivated on potato dextrose agar (PDA) at 24±1 °C until colonies covered approximately two-thirds of the area of the Petri dishes. Discs of 0.5 cm in diameter were collected from the actively growing regions of the colonies and kept in sterile distilled water, pH 6.5, at 6–8 °C, as stock for future inoculations.

Production of extracellular hydrolases. Petri dishes containing Pontecorvo’s minimal medium agar [24], pH 6.8, and 0.2 % glucose [33], with the addition of 1 % (w/v) citric pectin, 4 % (w/v) gelatin (sodium nitrate reduced to 3 mM), or 1 % (w/v) soluble starch, was used to detect pectinases, proteases on gelatin, and amylases, respectively. After the incubation period, substrate hydrolysis was detected by covering the plate with 1 % (w/v) hexadecyltrimethylammonium bromide (CTAB) [13] for pectinases, saturated solution of ammonium sulfate for proteases, or 1 % Lugol solution for amylases. A clear halo around the colony against an opaque surrounding, indicated pectin or gelatin hydrolysis. A reddish or yellowish halo around the colony on a dark background indicated starch hydrolysis.

Extracellular proteolytic enzymes on casein were detected with skim milk agar [19]. After incubation of the plates, a transparent halo around the colony against an opaque background indicated casein hydrolysis. Lipases were detected with sterilized rhodamine B agar [39] added of previously filter-sterilized olive oil at a 1 % (v/v) concentration. A yellow-orange color around the colony, detected using 350 nm UV light, indicated fungal production of lipases.

Each hydrolase assay was done in triplicate per isolate. A mycelium disc in the center of each 9-cm Petri dish was inoculated and the plates were then incubated at 30 °C in the dark for variable periods, until colonies covered 60–75% of the plate area.

Colony radial growth rate. The growth rate was determined using the media and the incubation conditions described above, as well as PDA (pH 6.8). All assays were done in triplicate. Colony size was assessed at regular intervals by measuring the colony diameter along two axes at a right angle to the inoculation point, using a Vernier caliper. Measurements were done until the colonies reached the sides of the plate. Radial growth rate on PDA was determined in 9-cm diameter plates with all isolates, and in 20-cm diameter plates with isolates MMBF 04-10, MMBF 808, PEL, and CMM 2105. The radial growth rate (mm/h), expressed as the angle of the linear portion of the regression line, was calculated based on the radius of the colony vs. incubation time.

Lipolytic activity. Erlenmeyer flasks (250 ml) containing 100 ml of a minimal salts broth [9], pH 6.8, and 1 % (v/v) previously filter-sterilized olive oil were inoculated with three mycelium discs of isolates MMBF 0410, CMM 2105, PEL, and MMBF 808, one flask per isolate. The flasks were incubated at 30 °C in a rotary shaker (160 rpm). Every 24 h for 4 days, 5ml samples were collected, filtered through Whatmann paper, and frozen at –17 °C until analysis. The lipolytic activity of the cultivation broths was

LIPASES FROM M. PHASEOLINA

assayed by using 4-nitrophenyl palmitate (pNPP) as substrate [22] in TrisHCl buffer, pH 8, and 15-min incubation at 37 °C (ε = 13,300 M–1 cm–1).

Absorbance was read at 410 nm by using heat-inactivated cultivation broth as blank. One unit (U) of lipolytic activity is defined as the amount of enzyme that liberates 1 μmol of 4-nitrophenol (pNP) per minute per ml of cultivation broth. The protein content of the samples was determined according to Lowry’s method, and the specific lipolytic activity (U mg protein–1) was calculated.

Statistical analysis. Assistat. Statistical Assistance software was used for ANOVA, Tukey test, and Scott-Knott test [http://www. assistat.com/indexp.html].

Results and Discussion Production of extracellular hydrolases. Table 1 shows the hydrolases produced by each isolate. Consistent with previous reports of pectin hydrolysis with enzymatic extracts of M. phaseolina [26], all 13 isolates hydrolyzed pectin. Pectinases were also detected in another study of three isolates [3], one of which was more efficient in causing stem rot. Dhingra et al. [11] demonstrated differentiated pectolytic and cellulolytic activity in vitro and in vivo between a virulent and an avirulent isolate. Maximum pectinase activity was detected at 48 h when this phytopathogen was grown in submerged cultivation [2], with a rapid decrease in activity after 96 h. By contrast, weak pectinase activity was reported for Macrophomina sp. MS 139 [35].

All 13 isolates secreted amylases. Onilude and Oso [21] also obtained amylases from M. phaseolina and used them, either as crude or partially purified preparations, in feed diet to improve the weight gain of broiler chicken. Another study [35], testing for extracellular hydrolases in several fungi, failed to detect amylases in Macrophomina sp. MS 139.

However, according to one report [12] an isolate of this fungus showed good dextrinizing and saccharizing specific activities on carbon sources such as starch, jackfruit seed flour, and rice flour.

In the present study, 11 isolates produced proteases able to hydrolyze gelatin. Proteases active on casein were also frequently detected, except in two (AJAM and MMBF 564) of the 13 isolates. The sparse references on the proteases produced by M. phaseolina also mention variability in their detection. Ahmad et al. [2] examined two strains of this fungus but did not detect proteolysis on casein either in solid medium in Petri dishes or in submerged culture. However, in their study of several fungi, Sohail et al. [35] reported the detection of proteolytic enzymes produced by Macrophomina sp. MS 139 in mineral medium with casein. The authors concluded that proteases are the most common

INT. MICROBIOL. Vol.15, 2012

3

Table 1. Detection of extracellular hydrolases of Macrophomina phaseolina isolates grown on different substrates Peca

Amyb

Prot gelc

Prot casd

Lipe

PEL

+

+

+

+

+

AJAM

+

+

–

–

–

CMM 527

+

+

–

+

+

CMM 932

+

+

+

+

+

CMM 979

+

+

+

+

+

CMM1048

+

+

+

+

+

CMM1091

+

+

+

+

+

CMM2100

+

+

+

+

+

CMM2105

+

+

+

+

+

MMBF564

+

+

+

–

+

MMBF808

+

+

+

+

+

MMBF16-98

+

+

+

+

+

MMBF04-10

+

+

+

+

+

Hydrolases detection: (+) hydrolases detected, (–) negative for substrate hydrolysis on the triplicates. a Pectinases. bAmylases. cProteases on gelatin. dProteases on casein. e Lipases on olive oil.

hydrolases in filamentous fungi. Kakde and Chavan [15] also observed the ability of this fungus to use casein as source of nitrogen.

In this study, all isolates showed lipase activity when induced with olive oil. The exception was isolate AJAM, which showed very restricted growth and no lipolysis even after 6 days of cultivation. Other studies also reported the production of lipolytic enzymes by this pathogen. In an experiment comparing lipase production by M. phaseolina and Phoma nebulosa [28], enzyme production was shown to depend on the culture medium used. In that work, M. phaseolina produced higher amounts of lipases when stimulated by the addition of sesame flour to the medium. Another work [38] examined the M. phaseolina-induced deterioration of peanuts, specifically, the changes in moisture content, fatty acids, and proteins. A decrease in oil content and an increase in free fatty acids was noted, demonstrating the lipolytic action of this fungus. In the above-mentioned study by Ahmad et al. [2], the two M. phaseolina isolates also produced lipases.

Ten of our 13 isolates showed hydrolytic activity on all substrates tested. In contrast to some studies on filamentous fungi, among the hydrolases, proteases were less frequently detected.

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INT. MICROBIOL. Vol. 15, 2012

Table 2. Radial growth rate (RGR)of Macrophomina phaseolina on different substrates RGRa (mm/h)

Substrate Pectin

0.90±0.44 a

Soluble starch

0.60±0.22 b

Gelatin

0.77±0.27 a

Casein

0.55±0.25 b

Olive oil

0.62±0.28 b

a

Mean radial growth rate from triplicates of thirteen isolates on the same medium. Means followed by the same letter do not differ statistically from each other (Scott-Knott test, P < 0.05).

Colony radial growth rate. The radial growth rates (mm/h) of each isolate grown on the different media were calculated from the radius of the colony and the incubation time. The overall radial growth rate achieved on a medium was determined by taking the individual rates of the triplicates of the 13 isolates on the same medium and calculating

their mean (Table 2). The several culture media resulted in two distinct growth rates, showing that M. phaseolina grew significantly better (P < 0.05) on pectin and gelatin. Radial growth rates were quite variable among isolates grown on the same medium, and also varied for the same isolate on the different culture media (Table 3). The linear correlation coefficient (Pearson’s r) of the regression lines remained between 0.99 and 0.83 for all media, except gelatin (r = 0.75).

Although cultivated under the same conditions, isolates of M. phaseolina in the present study showed radial growth rates on PDA well above those determined by Mayek-Pérez et al. [18] with Mexican isolates from different hosts and different regions of the country. In that study, the rates ranged from 0.45 to 0.50 mm/h, and the authors related the heterokaryotic nature of the mycelium of M. phaseolina to the variability of its morphological characteristics, development in vitro, and virulence. Also mentioned as sources of variability were the geographical origin of the isolate, host type, cultivation time, and culture medium employed. Although in our study isolate MMBF 04-10 showed the highest growth rate on PDA, the other isolates grew faster on several media. Of all substrates tested, pectin yielded the highest growth rates,

Table 3. Radial growth rate of Macrophomina phaseolina isolates grown on different substratesa Substrates

Pectin

Soluble starch

Gelatin

Casein

Olive oil

PDA

PEL

0.64±0.09 e

0.68±0.02 b

1.06±0.04 a

0.96±0.02 a

0.98±0.03 a

1.05±0.05 b

AJAM

0.22±0.01 g

0.19±0.05 c

0.28±0.01 d

0.04±0.01 h

0.07±0.01 h

0.06±0.01 i

CMM 527

0.62±0.08 e

0.72±0.04 b

0.72±0.20 b

0.83±0.03 b

0.74±0.01 b

0.61±0.02 f

CMM 932

0.45±0.06 f

0.26±0.18 c

0.35±0.17 d

0.24±0.02 g

0.34±0.02 g

0.45±0.01 g

CMM 979

1.05±0.06 d

0.67±0.02 b

0.98±0.03 a

0.63±0.05 c

0.78±0.01 b

0.74±0.01 d

CMM1048

1.38±0.05 b

0.58±0.01 b

0.84±0.06 b

0.55±0.08 d

0.76±0.01 b

0.49±0.02 g

CMM1091

1.62±0.07 a

0.72±0.02 b

1.06±0.03 a

0.89±0.01 b

0.98±0.05 a

0.61±0.06 f

CMM2100

1.21±0.22 c

0.62±0.04 b

0.91±0.03 a

0.58±0.03 d

0.65±0.03 c

0.41±0.01 g

CMM2105

1.25±0.08 c

1.02±0.01 a

0.98±0.03 a

0.69±0.08 c

1.00±0.01 a

1.02±0.01 b

MMBF564

0.78±0.03 e

0.68±0.11 b

0.78±0.03 b

0.45±0.01 e

0.30±0.01 g

0.69±0.02 e

MMBF808

0.97±0.03 d

0.56±0.03 b

0.57±0.04 c

0.36±0.04 f

0.52±0.02 e

0.85±0.03 c

MMBF16-98

0.26±0.04 g

0.38±0.01 c

0.47±0.09 c

0.47±0.02 e

0.41±0.08 f

0.21±0.06 h

MMBF04-10

1.28±0.10 c

0.60±0.27 b

0.93±0.07 a

0.49±0.03 e

0.59±0.04 d

1.33±0.07 a

a

Mean radial growth rate (mm/h) and standard error of three determinations. Means followed by the same letter do not differ statistically from each other (Scott-Knott test, P < 0.05).

INT. MICROBIOL. Vol.15, 2012

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LIPASES FROM M. PHASEOLINA

Fig. 1. Extracellular lipase production by M. phaseolina isolates PEL (diamonds), CMM 2105 (squares), MMBF 04-10 (triangles), and MMBF 808 (circles) in minimal mineral salts medium with olive oil as inducer. Values represent the means and standard deviation of three determinations.

with isolate CMM 1091 growing significantly faster than the other isolates (1.61 ± 0.06 mm/h, P < 0.05). On starch, isolate CMM 2105 showed a significantly higher growth rate (P < 0.05). Several isolates grew rapidly on gelatin but not on casein, although both are protein substrates; in fact, casein yielded the lowest growth rates among all substrates. Similar results were obtained with the filamentous fungi Batrachochytrium dendrobatidis [25] and Aspergillus sydowii [32], which also showed different growth patterns on gelatin than on casein, with both developing a higher mycelium mass in media containing the latter. On olive oil, isolates PEL, CMM 1091, and CMM 2105 showed significantly faster growth (P < 0.05) than the other isolates.

The variability in both the production of several extracellular hydrolases by M. phaseolina and the fungus’ rate of radial growth on PDA, as described in the present study, confirms the diversity reported by other authors. To our knowledge, ours is the first report on M. phaseolina radial growth rates on media other than PDA. A high growth rate and the production of specific enzymes are features that allow the selection of isolates with specific characteristics. CMM 1091 grew rapidly on all sub-

strates tested for the production of hydrolases, while isolates PEL and CMM 2105 grew quickly on most media, producing the corresponding hydrolase. Thus, CMM 1091, PEL, and CMM 2105 are isolates that produce enzymes of potential industrial interest and therefore merit further research.

Lipolytic activity. To verify the production of lipases by four isolates using olive oil as sole source of carbon, the lipolytic activity of their cultivation broths was tested against 4-nitrophenyl palmitate. Figure 1 shows the specific lipolytic activity (U mg protein–1) of isolates PEL, CMM 2105, MMBF 04-10, and MMBF 808 during 4 days of cultivation.

The activities of three isolates (PEL, CMM 2105, and MMBF 04-10) increased with cultivation time and in each case reached a maximum at 96 h: PEL 130 mU mg protein–1, CMM 2105 110 mU mg protein–1, and MMBF 04-10 80 mU mg protein–1. The activity of the fourth isolate, MMBF 808, was minimal (2 mU mg protein–1 in 96 h). The ANOVA and Tukey test (P < 0.05) of the specific lipolytic activities of PEL, CMM 2105, and MMBF 04-10 indicated that they were significantly different, with PEL showing the highest activity throughout the cultivation period.

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Ahmad et al. [2] cultivated two strains of M. phaseolina in minimal salts medium with olive oil for 96 h, while measuring the lipolytic activity of the broth at regular intervals.

The activity of strain 1 peaked at 24 h and reached a minimum at 96 h, while strain 2 activity, which was only 40 % of that of strain 1, peaked at 24 h and reached a minimum at 48 h.

Kakde and Chavan [16] cultivated an isolate of M. phaseolina for 25 days in submerged cultivation in minimal salts medium containing oil and tested the lipolytic activity of the broth every 5 days for 25 days. Maximum activity was reached on day 25, and half-maximal activity already on day 5. Since no other studies were found using spectrophotometry to determine the lipolytic activity of M. phaseolina, our results cannot be compared with those of other researches. However, it is clear that the incubation time necessary for M. phaseolina to reach peak lipolytic activity in submerged culture is quite variable, depending on culture conditions and the particular isolate.

A comparison of the radial growth rate on olive oil medium with the specific lipolytic activity of the four isolates tested showed that both CMM 2105 and PEL had high growth rates and produced the highest lipase activities. Studies of other fungi have shown that growth rate and lipolytic activity are regulated by the cAMP/PKA (cyclic AMP-dependent protein kinase A) signaling pathway. In a study by Ocampo et al. [20], a mutant of Mucor circinelloides lacking the gene for one of the regulatory subunits of PKA (and thus exhibiting high PKA activity) exhibited a decrease in growth and alterations in germination rates, cell volume, germ tube length, and asexual sporulation. Klose et al. [17] found that cAMP/PKA signaling regulates the morphological growth of Ustilago maydis, whether filamentous or budding, and that higher amounts of lipase are secreted in the presence of triglycerides only by strains showing filamentous growth.

They speculated that cAMP signaling is involved in the ability of the fungus to use oils as carbon source and that the gene(s) encoding the lipase activity is regulated by PKA. Our findings suggest that this is also the case for Macrophomina phaseolina, as the correlation coefficient between radial growth rate on olive oil and lipolytic activity was strong and positive (0.86), indicating that faster filamentous growth was associated with the higher production of lipolytic enzymes.

In summary, Macrophomina phaseolina was shown to produce extracellular pectinases, amylases, proteases, and lipases. The isolates, however, varied in their abilities to produce these enzymes, as some did not produce all the hydrolases tested, and proteases were less commonly detected on both gelatin and casein.

The determination of radial growth rates on different substrates together with the detection of the corresponding extracellular hydrolases identified CMM 1091, CMM 2105, and PEL as fast-growing isolates with great diversity in the production of extracellular hydrolases of industrial interest.

Among the isolates tested, PEL produced the highest lipase activity, and the enzyme was active at 37 °C, pH 8, with potential use in mild alkaline detergents. It is also reasonable to suggest that, as in other fungi, the radial growth rate and lipolytic activity of M. phaseolina are probably regulated by the cAMP/PKA pathway, although this remains to be demonstrated in further studies.

Acknowledgements. The authors appreciate the financial support from CAPES (Coordenação de Aperfeiçoamento de Pessoal de Ensino Superior, Ministry of Education, Brazil) in the form of a scholarship to C. Schinke.

They also thank Igor Villela Marroni, the Federal Rural University of Pernambuco (UFRPE), the Biological Institute of the Department of Agriculture and Food Supply of the State of São Paulo (IB-SP), the Federal University of Pelotas (UFPel), and the Federal University of Viçosa for providing samples of M. phaseolina. Thanks are also extended to Dr. Marco Antonio Z. Ayub and Dr. Adriano Brandelli from the Institute of Food Science and Technology of the Universidade Federal do Rio Grande do Sul (Brazil) for providing reagents and facilities.

Competing interests. None declared.

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26. Radha K (1953) The enzymic activity of Macrophomina phaseoli (Maubl.), Ashby. Proceedings: Plant Sci 38:231-234 27. Rathi P, Saxena RK, Gupta R (2001) A novel alkaline lipase from Burkholderia cepacia for detergent formulation. Process Biochem 37:187-192 28. Reddy AS, Reddy SM (1983) Lipase activity of two seed-borne fungi of sesamum (Sesamum indicum Linn.). Folia Microbiol 28:463-466 29. Roy PK, Roy U, Vora VC (1993) Hydrolysis of wheat bran, rice bran and jute powder by immobilized enzymes from Macrophomina phaseolina. World J Microbiol Biotechnol 9:164-167 30. Saxena RK, Sheoran A, Giri B, Sheba Davidson W (2003) Purification strategies for microbial lipases. J Microbiol Methods 52:1-18 31. Serrat M, Rodríguez O, Camacho M, Vallejo JA, Ageitos JM, Villa TG (2011) Influence of nutritional and environmental factors on ethanol and endopolygalacturonase co-production by Kluyveromyces marxianus CCEBI 2011. Int Microbiol 14:41-49 32. Sharma AK, Sharma V, Saxena J (2011) Production of protease and growth characteristics of Aspergillus sydowii. Nat Sci 9:217-221 33. Silva JH, Monteiro RTR (2000) Degradação de xenobióticos por fungos filamentosos isolados de areia fenólica. R Bras Ciência Solo 24:669-674 (In Portuguese) 34. Smith GS, Carvil ON (1997) Field screening of commercial and experimental soybean cultivars for their reaction to Macrophomina phaseolina. Plant Dis 81:363-368 35. Sohail M, Naseeb S, Sherwani SK, Sultana S, Aftab S, Shahzad S, Ahmad A, Khan SA (2009) Distribution of hydrolytic enzymes among native fungi: Aspergillus the pre-dominant genus of hydrolase producer.

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RESEARCH ARTICLE INTERNATIONAL MICROBIOLOGY (2012) 15:9-15 DOI: 10.2436/20.1501.01.154 ISSN: 1139-6709 www.im.microbios.org

Surface alteration of realgar (As4S4)

by Acidithiobacillus ferrooxidans Peng Chen,1,2 Lei Yan,1,3 Qiang Wang,4 Yang Li,1 Hongyu Li1* 1

Institute of Microbiology, School of Life Sciences, Lanzhou University, Lanzhou, PR China. 2GIBT, Gansu Institute of Business and Technology, Lanzhou, PR China. 3College of Life Science and Technology, Heilongjiang Bayi Agricultural University, Daqing, PR China. 4College of Chemistry and Chemical Engineering, Lanzhou University, Lanzhou, PR China.

Received 6 January 2012 · Accepted 26 February 2012

Summary. The chemical and physical characteristics of realgar (an arsenic sulfide mineral that occurs in several crystalline forms) in the presence of Acidithiobacillus ferrooxidans BY-3 were investigated in this work. Grains of the mineral were incubated for 10, 20, and 30 days with A. ferrooxidans cultured in 9K medium at 30 °C and at 150 rpm agitation.

Abiotic control experiments were conducted in identical solutions. The effect of bioleaching on the surface properties of realgar was characterized by scanning electron microscopy (SEM), energy-dispersive spectroscopy (EDS), inductively coupled plasma atomic emission spectroscope (ICP-AES), X-ray diffraction (XRD), and Raman spectroscopy. SEM and EDS analyses confirmed the ability of A. ferrooxidans to modify surfaces of realgar and to efficiently enhance its dissolution.

ICP-AES showed the dissolution and precipitation of realgar during bioleaching. Based on the XRD pattern and the Raman spectra, the decrease in arsenic in the liquid phase was due to co-precipitation of the mineral with Fe(III) or Fe(III) compounds (e.g., jarosite or goethite). Thus, not only did Fe(III) alter the surface of realgar, but it also promoted its dissolution during bioleaching. [Int Microbiol 2012; 15(1):9-15] Keywords: Acidithiobacillus ferrooxidans · realgar (arsenic sulfide) · bioleaching · Raman spectroscopy · X-ray diffraction

Introduction Realgar is a red, semiconductor, arsenic sulfide mineral that exists in several crystalline forms. It is often found in association with orpiment, another arsenic sulfide. Natural realgar occurs in a low-temperature phase termed α-As4S4. A hightemperature polymorph, β-As4S4, can be obtained by heating realgar above 252 °C. Both forms are based on the same *Corresponding author: Hongyu Li Institute of Microbiology, School of Life Sciences Lanzhou University Tianshui Road No. 222 Lanzhou, 730000, PR China Tel. +86-9318912560. Fax +86-9318912561 E-mail: RedGeneSCI@gmail.com; lihy@lzu.edu.cn

cage-like molecule even though they differ in their molecular packing, which involves two distinct monoclinic lattices [4]. Throughout history, realgar has had many applications in the manufacture of fireworks and leather , in pesticides, and as a medicinal agent [16]. Recently, realgar was reported to be clinically effective in the treatment of various forms of cancer both in vitro and in vivo [3,20,27].

The preparation of realgar is achieved using various traditional and modern methods, such as acid extraction, calcination, membrane dialysis, mechanical milling, cryo-grinding, and the chemical synthesis of quantum dots [1,21].

However, realgar is poorly soluble in aqueous and most organic solvents due to its high intrinsic lattice energy.

Consequently, these technologies are often expensive and

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environmentally deleterious; moreover, their success may be hindered by ineffective dissolution and storage difficulties [7]. Additional problems are the high toxicity and poor bioavailability of realgar, which have seriously limited its use in the clinical setting.

Bioleaching, a long-standing technology in hydrometallurgy, has recently been applied to prepare realgar, with the aim of increasing its bioavailability [7]. In the case of realgar production, the first report of biological precipitation was that of Huber et al. in 2000 [6,12]. Those authors described bioleaching of the mineral by the hyperthermophile Pirobaculum arsenaticum. Under organotrophic conditions and in the presence of thiosulfate and arsenate, P. arsenaticum forms realgar [5]. By taking advantage of an arsenicresistant strain of indigenous Acidithiobacillus ferrooxidans (A. ferrooxidans) BY-3, we applied this bioleaching method to develop a bio-arsenic aqueous solution from coarse realgar and confirmed its in vitro and in vivo anticancer activities [26].

Acidithiobacillus ferrooxidans (formerly Thiobacillus ferrooxidans) is a mesophilic and acidophilic chemolithotrophic bacterium and the most well-studied acidophilic organism [8]. Due to its bioleaching capabilities, it is an important member of microbial consortia involved in the iron cycle. In fact, A. ferrooxidans, Acidiphilium spp., and Leptospirillum spp. were reported to account for 80% of the prokaryotic diversity in the Río Tinto ecosystem (Huelva, Spain), where ferric iron and sulfates are abundant [9,10,19].

Even though arsenic can be toxic to A. ferrooxidans, the bacterium can be adapted to tolerate much higher arsenic concentrations than it does in nature [Mandi C (2003) The effects of arsenic on Thiobacillus ferrooxidans. Columbia University, Master Thesis], with some strains resistant to arsenic at concentrations in the milligram per liter range [23].

Compared to the traditional above- mentioned methods, the bioleaching of realgar produces extraordinary increases in its solubility and bioavailability while decreasing its toxic effects [3].

Previous investigations revealed that ferrous iron and elemental sulfur exert important effects on metal extraction during the bioleaching of realgar, but the chemical and physical changes that take place at the realgar surface have yet to be thoroughly studied [7]. Several analytical investigations of the inorganic alteration process of realgar surfaces have provided information about their oxidation kinetics and the light-induced degradation of realgar as well as electrochemi-

cal effects and the surface properties of realgar nanoparticles in the absence of bacteria [2,14,15,22]. These studies have improved our understanding of realgar alterations in the presence of air, water, acidic, neutral, and other abiotic surroundings. To date, however, surface analytical techniques, such as Raman spectroscopy, have not been applied in the examination of realgar after its reaction with A. ferrooxidans. In a previous study, the reactivity of realgar with iron-oxidizing bacteria was monitored by scanning electron microscopy (SEM) and energy dispersive spectrometric (EDS) [17].

The aim of this study was to observe the chemical and physical changes occurring at powdered realgar surfaces in the presence of the bacterium A. ferrooxidans. The long-term goal is to achieve the necessary modifications of the mineral that will facilitate its effective use in biotechnological and clinical applications. Accordingly, change in the realgar surfaces were analyzed using several complementary techniques: SEM, EDS, powder X-ray diffraction (XRD), and Raman spectroscopy.

Materials and methods Realgar. The investigation was carried out using realgar As4S4 (99.01 % purity) from Shimen County, Hunan Province, China, which was purified by traditional methods according to the Chinese Pharmacopeia [Chinese Pharmacopoeia Committee (2010) Pharmacopoeia of the People’s Republic of China, China People’s Press, Beijing, pp 316]. The raw realgar was crushed to 200 mesh size (approximately 75 ± 10 μm) and then analyzed chemically, using an inductively coupled plasma atomic emission spectroscope (ICP-AES, IRIS Advantage, Thermo Jarrell Ash Corporation, USA), and mineralogically by XRD.

Microorganism and bioleaching experiments. The native A. ferrooxidans strain BY-3 (CCTCC-M203071) was previously isolated from an abandoned copper mine in Baiyin, Gansu Province, China. The bacterium was cultured in 9K medium [18], consisting of (per liter): 3.0 g (NH4)2SO4, 0.1 g KCl, 0.5 g K2HPO4, 0.5 g MgSO4·7H2O, 0.01 g Ca(NO3)2. Bioleaching experiments were performed in 250-ml conical flasks containing 100 ml of 9K medium (with 44.69 g of FeSO4·7H2O per liter) and 0.5 g of realgar at an initial pH of 1.7 (adjusted with sulfuric acid). The flasks were incubated at 30 °C, with shaking at 150 rpm. A sterile flask without bacteria but subjected to identical experimental conditions was included as a negative control.

The experiments lasted for 30 days. All experiments were performed in duplicate at a minimum. The average values are reported.

Analytical procedures. In bioleaching experiments, soluble arsenic concentrations were measured by ICP-AES [7]. Surfaces of realgar before and after bioleaching were coated with gold and observed with a JEOL scanning electron microscope (JSM-5600LV, Tokyo, Japan) operated at 20 kV and a JEOL field emission scanning electron microscope (FESEM; JSM-

ALTERATION OF REALGAR BY A. FERROOXIDANS

6701F, Tokyo, Japan) operated at 5–8 kV and using image slave software for image capture. The ratio of arsenic to sulfur was analyzed by EDS (Thermo Kevex) at a JSM-5600LV workstation. Sample preparation for SEM analysis was previously described [17]. XRD analyses of the powder form of the raw sample and residual bioleaching were conducted on a multipurpose Xray diffraction system (X’Pert-Pro MPD, Philips), using Cu Kα radiation (λ= 0.15406 nm) and operated at 40 kV and 40 mA. The sample preparation method for XRD was previously described [23]. Raman spectroscopy measurements were carried out with a high-resolution Raman spectrometer (Horiba-Jobin, Yvon, HR-800) using a He–Ne laser (λ = 532 nm) as the excitation source.

Results and Discussion

11

Ca (0.01057), Fe (0.04015), Mg (0.00293), Hg (0.00323), K (0.00412), Se (0.00598), Al (0.00662), Cd (0.00004), Zn (0.00092), Cu (0.00013), and Ba (0.00028).

The interactions between realgar powder and cultures of A. ferrooxidans BY-3 in the presence of ferrous iron were analyzed by SEM and EDS, with the results presented in Fig. 1 and Table 1, respectively. Realgar samples subjected to bioleaching by A. ferrooxidans BY-3 for 0 (non-treated), 10, 20, and 30 days, are shown in Fig. 1. During the first 10 days of the experiment, no significant changes in the raw realgar powder were observed (Fig. 1B). However, after 20 days, as shown in Fig. 1C,D, larger cracks or pits appeared on the the mineral surface. Previous investigations suggested that the attachment of A. ferrooxidans to the realgar surface is the key step in the bioleaching process [7].

The combined actions of attached bacteria and free bacteria were determined to increase the rate of realgar dissolu-

Int. Microbiol.

The studied sample mostly consisted of realgar (97 %, w/w), with very small inclusions of arsenolite (3 %, w/w). Chemical analysis of this sample showed that it contained the following elements (expressed as %, w/w): As (68.0), S (31.01),

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Fig. 1. Scanning electron microscopy (SEM) image of the surface of realgar reacted with Acidithiobacillus ferrooxidans. (A) Before leaching; raw realgar powder. (B) After 10 days. (C) After 20 days. (D) After 30 days. Scale bar = 1 μm.

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Table 1. The EDS analysis of the arsenic and sulfur ratio of realgar Sample Before bioleaching

After bioleaching

Time (days) 0

20

30

Element

Weight conc. (%)

Atom conc. (%)

Ratio As/S

Stoichiometric As/S

As

50.44

30.34

0.44

As0.44S0.56

S

49.56

69.66

As

48.95

29.10

0.41

As0.41S0.59

S

51.05

70.90

As

46.85

27.39

0.38

As0.38S0.62

S

53.15

72.61

as described in Table 1. This is in agreement with previous studies showing that the proportion of soluble arsenic gradually increases in the bioleaching process [25]. On day 20, the ratio of arsenic to sulfur decreased to a greater extent in the bioleaching realgar than in the non-treated realgar. Thus, bioleaching not only leads to a more rapid extraction of arsenic, it also causes a relatively severe attack on the realgar surface compared to non-treated realgar. The stoichiometric ratio gradually decreased from 0.41:0.59 on day 20 to 0.38:0.62 on day 30. This decrease in the ratio of arsenic to sulfur was accompanied by an increase in the concentrations

Int. Microbiol.

tion, confirming that direct bacterial action plays a major role in the modification of realgar surfaces.

As evidenced by EDS analysis, the major constituents of bioleached realgar were As, S, Fe, K, Mg, and Ca. This result was consistent with those of the XRD analysis, which revealed that magnetite and jarosite were the major Fe(III) compounds, while they were not found in the non-treated realgar. According to the EDS results, the ratio of arsenic to sulfur was 0.44:0.56 at the beginning of the experiment; 0.41:0.59 on day 20, and 0.38:0.62 on day 30. The ratios on the latter two days were lower than the stoichiometric ratio,

Fig. 2. X-ray diffraction (XRD) of: (A) solid realgar; (B) residual realgar after bioleaching by Acidithiobacillus ferrooxidans cultures in a medium containing ferrous sulfate. (a) realgar, (b) arsenolite, (c) dimorphite, (d) magnetite, (e) jarosite.

ALTERATION OF REALGAR BY A. FERROOXIDANS

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INT. MICROBIOL. Vol.15, 2012

Fig. 3. Raman spectra: (a) realgar powder; (b) residual realgar after bioleaching.

of soluble arsenic in the leach liquor. The results demonstrated that A. ferrooxidans can efficiently enhance the dissolution of realgar.

The changes in arsenic concentration over time in the presence of ferrous iron were also analyzed. In the presence of A. ferrooxidans (121.95 mg/l), the maximum level was reached on day 15 and was 7.4 times higher than in the sterile control (16.44 mg arsenic/l). Afterwards, however, the arsenic content began to decrease such that by day 25 it was even lower than in the sterile control. In order to understand these variations in arsenic concentrations, the XRD patterns and Raman spectra of the solid residues were evaluated.

Figure 2 shows the XRD patterns of the realgar particles before and after bioleaching. The many overlapping peaks were accompanied by a tendency toward gradual amorphization as a consequence of microbial action. All identified peaks belonged to different phases. Based on a comparison with the Inorganic Crystal Structure Database (ICSD) data, two phases in the studied raw realgar sample could be ascribed to this form: As4S4 (97 %) and As2O3 (3 %). In con-

trast, the mineral composition found in the solid residues after bioleaching comprised realgar, dimorphite, magnetite and jarosite, which accounted for 43.0 % (w/w), 23.0 % (w/w), 6.0 % (w/w) and 28.0 % (w/w) of the sample, respectively. These results are consistent with those obtained by SEM and EDS. The most widely used adsorbents are those that are iron-based (e.g., jarosite or magnetite), due to their high arsenic removal efficiency [11]. After 30 days, the presence of jarosite and magnetite was confirmed by the XRD patterns of the solid residues in the bioleaching experiment.

However, jarosite was not found in the residue of the culture without A. ferrooxidans. The above results suggested that arsenic leached from realgar is adsorbed onto Fe(III) or Fe(III) compounds (including jarosite, goethite, magnetite and hematite), and co-precipitated with Fe(III) extracted from jarosite, as described in [24]. Co-precipitation may have further inhibited the dissolution of realgar. Moreover, previous studies have proposed that the adsorption of As(V) and is competitive, or that ionic As species (H2AsO4â&#x20AC;&#x201C;) is selectively adsorbed [24]. Consequently, adjustment of the Fe(III)

14

CHEN ET AL.

INT. MICROBIOL. Vol. 15, 2012

or Fe(III) compounds is of great importance during the bioleaching of realgar, as it significantly decreases the removal of arsenic from the liquid phase.

The Raman spectrum of raw realgar is presented in Fig. 3a, which shows two As-As stretching vibration peaks at 189 cm–1 and 204 cm–1, two As-S stretching vibration peaks at 350 cm–1 and 362 cm–1, and a characteristic As-S-As bending vibration at 274 cm–1. This spectrum did not change significantly over time, confirming that the material was realgar. The Raman spectrum of realgar modified by bioleaching is shown in Fig. 3b. The relative intensity of the As-As stretching vibration peak at 189 cm–1 is increased; in addition, there are two As-S stretching vibration peaks at 270 cm–1 and 351 cm–1 and a characteristic As-S-As bending vibration at 234 cm–1.

According to Kyono [13], the Raman spectrum of para-realgar is characterized by a pair of strong peaks near 230 cm–1 and a grouping of four distinguishable peaks centered at approximately 340 cm–1. These phase characteristics are not seen in the spectrum of Fig. 3b. Instead, there was a strong increase in the dissolution of realgar, which retained its original characteristics after bioleaching without alterations in its physicochemical properties.

This study shows that A. ferrooxidans is able to modify realgar surfaces and to enhance dissolution of the mineral, as detected by SEM and EDS. Additionally, the ICP-AES results indicated that both dissolution and precipitation occurred during the bioleaching of realgar. As determined from XRD and Raman spectral analyses, the bioleaching of realgar by A. ferrooxidans is due to the oxidative properties of ferric iron. Therefore, the addition of Fe(III) and Fe(III) compounds can improve the dissolubility of realgar, in addition to decreasing the arsenic concentration in the liquid phase.

References 1.

2.

3.

4.

5.

6.

7.

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11.

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13.

14.

Acknowledgements. This work was supported by the Technology Program of Gansu Province (grant nos. 2GS064-A43-019-02, 0912TCYA025, 098TTCA013 and 1004TCYA041), the International Cooperation Project (grant no. 0708WCGA150), Open Project of Key Laboratory for Magnetism and Magnetic Materials of the Ministry of Education, Lanzhou University (grant no. LZUMMM2010016), and the Fundamental Research Funds for the Central Universities (grant no. lzujbky2010-36).

Competing interests. None declared.

16.

17.

18.

Baláz P, Fabián M, Pastorek M, Cholujová D, Sedlák J (2009) Mechanochemical preparation and anticancer effect of realgar As4S4 nanoparticles. Mater Lett 63:1542-1544 Baláz P, Nguyen AV, Fabián M, Cholujová D, Pastorek M, Sedlák J, Bujnáková Z (2011) Properties of arsenic sulphide As4S4 nanoparticles prepared by high-energy milling. Powder Tech 211:232-236 Baláz P, Sedlák J (2010) Arsenic in cancer treatment: challenges for application of realgar nanoparticles (a minireview). Toxins 2:1568-1581 Bonazzi P, Bindi L, Muniz-Miranda M, Chelazzi L, Rödl T, Pfitzner A (2011) Light-induced molecular change in HgI2·As4S4: Evidence by single-crystal X-ray diffraction and Raman spectroscopy. Amer Mineral 96:646-653 Bruneel O, Pascault N, Egal M, Bancon-Montigny C, Goñi-Urriza M, Elbaz-Poulichet F, Personné JC, Duran R (2008) Archaeal diversity in a Fe–As rich acid mine drainage at Carnoulès (France). Extremophiles 12:563-571 Chaban B, Ng SYM, Jarrell KF (2006) Archaeal habitats-from the extreme to the ordinary. Can J Microbiol 52:73-116 Chen P, Yan L, Leng FF, Nan WB, Yue XX, Zheng YN, Feng N, Li HY (2011) Bioleaching of realgar by Acidithiobacillus ferrooxidans using ferrous iron and elemental sulfur as the sole and mixed energy sources.

Bioresour Technol 102:3260-3267 Corkhill C Vaughan D (2009) Arsenopyrite oxidation—A review. Appl Geochem 24:2342-2361 De los Ríos A, Valea S, Ascaso C, Davila A, Kastovsky J, McKay CP, Gómez-Silva B, Wierzchos J (2010) Comparative analysis of the microbial communities inhabiting halite evaporites of the Atacama Desert. Int Microbiol 13:79-89 García-Muñoz J, Amils R, Fernández VM, De Lacey AL, Malki M (2011) Electricity generation by microorganisms in the sediment-water interface of an extreme acidic microcosm. Int Microbiol 14:73-81 Giménez J, Martínez M, de Pablo J, Rovira M, Duro L (2007) Arsenic sorption onto natural hematite, magnetite, and goethite. J Hazard Mater 141:575-580 Huber R, Sacher M, Vollmann A, Huber H, Rose D (2000) Respiration of arsenate and selenate by hyperthermophilic archaea. Syst Appl Microbiol 23:305-314 Kyono A (2010) Growth and Raman spectroscopic characterization of As4S4 (II) single crystals. J Cryst Growth 312:3490-3492 Lazaro I, Gonzalez I, Cruz R, MG M (1997) Electrochemical study of orpiment (As2S3) and realgar (As2S2) in acidic medium. J Electrochem Soc 144:4128-4132 Liu J, Lu YF, Wu Q, Goyer RA, Waalkes MP (2008) Mineral arsenicals in traditional medicines: orpiment, realgar, and arsenolite. J Pharmacol Exp Ther 326:363-368 Ozturk S, Aslim B, Suludere Z (2010) Cadmium (II) sequestration characteristics by two isolates of Synechocystis sp. in terms of exopolysaccharide (EPS) production and monomer composition. Bioresour Technol 101:9742-9748 Silverman MP, Lundgren DG (1959) Studies on the chemoautotrophic iron bacterium Thiobacillus ferrooxidans. I. An improved medium and a harvesting procedure for securing high cellular yields. J Bacteriol 77:642-647

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19. Starek M, Kolev KI, Berthiaume L, Yeung CW, Sleep BE, Wolfaardt GM, Hausner M (2011) A flow cell simulating a subsurface rock fracture for investigations of groundwater-derived biofilms. Int Microbiol 14:163-171 20. Wang L, Zhou GB, Liu P, Song JH, Liang Y, Yan XJ, Xu F, Wang BS, Mao JH, Shen ZX, Chen SJ, Chen Z (2008) Dissection of mechanisms of Chinese medicinal formula Realgar-Indigo naturalis as an effective treatment for promyelocytic leukemia. Proc Natl Acad Sci U S A 105:4826-4831 21. Wu J, Shao Y, Liu J, Chen G, Ho PC (2011) The medicinal use of realgar (As4S4) and its recent development as an anticancer agent. J Ethnopharmacol 135:595-602 22. Xu JL, Xu XX, Zhou XY, Zhang P, Zhang CZ (2007) Microscopic raman imaging spectra of realgar and light-induced degradation products in realgar. Spectrosc Spectral Anal 27:577-580 23. Yan L, Yin HH, Zhang S, Leng FF, Nan WB, Li HY (2010) Biosorption of inorganic and organic arsenic from aqueous solution by Acidithiobacillus ferrooxidans BY-3. J Hazard Mater 178:209-217

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24. Yuichi T, Naoki H, Yasumasa K, Masami T (2005) Effect of jarosite on the removal of arsenic ions in sulfuric acid solution. Shigen-to-Sozai 212:597-602 (in Japanese) 25. Zhang JH, Zhang X, Ni YQ, Yang XJ, Li HY (2007) Bioleaching of arsenic from medicinal realgar by pure and mixed cultures. Process Biochem 42:1265-1271 26. Zhang X, Xie QJ, Wang X, Wang B, Li HY (2010) Biological extraction of realgar by Acidithiobacillus ferrooxidans and its in vitro and in vivo antitumor activities. Pharm Biol 48:40-47 27. Zhao QH, Zhang Y, Liu Y, Wang HL, Shen YY, Yang WJ, Wen LP (2010) Anticancer effect of realgar nanoparticles on mouse melanoma skin cancer in vivo via transdermal drug delivery. Med Oncol 27:203-212

RESEARCH ARTICLE INTERNATIONAL MICROBIOLOGY (2012) 15:17-32 DOI: 10.2436/20.1501.01.155 ISSN: 1139-6709 www.im.microbios.org

Bacterial isolates from the bryozoan Membranipora membranacea: influence of culture media on isolation and antimicrobial activity Herwig Heindl, Vera Thiel, Jutta Wiese, Johannes F. Imhoff* Kieler Wirkstoff-Zentrum (KiWiZ) at the Helmholtz-Zentrum für Ozeanforschung, GEOMAR, Kiel, Germany Received 8 January 2012 · Accepted 13 February 2012

Summary. From specimens of the bryozoan Membranipora membranacea collected in the Baltic Sea, bacteria were isolated on four different media, which significantly increased the diversity of the isolated groups. All isolates were classified according to 16S rRNA gene sequence analysis and tested for antimicrobial properties using a panel of five indicator strains and six different media. Each medium featured a unique set of isolated phylotypes, and a phylogenetically diverse collection of isolates was obtained. A total of 96 isolates were assigned to 49 phylotypes and 29 genera. Only one-third of the members of these genera had been isolated previously from comparable sources. The isolates were affiliated with Alpha- and Gammaproteobacteria, Bacilli, and Actinobacteria. A comparable large portion of up to 22 isolates, i.e., 15 phylotypes, probably represent new species. Likewise, 47 isolates (approximately 50%) displayed antibiotic activities, mostly against grampositive indicator strains. Of the active strains, 63.8 % had antibiotic traits only on one or two of the growth media, whereas only 12.7 % inhibited growth on five or all six media. The application of six different media for antimicrobial testing resulted in twice the number of positive hits as obtained with only a single medium. The use of different media for the isolation of bacteria as well as the variation of media considered suitable for the production of antibiotic substances significantly enhanced both the number of isolates obtained and the proportion of antibiotic active cultures. Thus the approach described herein offers an improved strategy in the search for new antibiotic compounds. [Int Microbiol 2012; 15(1):17-32] Keywords: Membranipora membranacea · antimicrobial activity · gene analysis · cultivation media · Baltic Sea

Introduction Surfaces in the marine environment, whether biotic or abiotic, are exposed to colonization by a multitude of organisms.

For example, the encrusting bryozoan Membranipora membranacea and related species populate kelps in temperate *Corresponding author: J.F. Imhoff Kieler Wirkstoff-Zentrum Am Kiel Kanal 44 24106 Kiel, Germany Tel.+49-4316004450. Fax +49-4316004452 E-mail: jimhoff@geomar.de

waters all over the world. The genus Membranipora is a potent colonizer and disperser; its global distribution most likely begain in the North Pacific several million years ago [42]. In the Baltic Sea, a preferred substrate is provided by phyloids of Saccharina latissima (newer synonym of Laminaria saccharina [24]). In turn, bryozoan surfaces are themselves subjected to colonizers and grazers. Like other sessile and colony-forming organisms in the marine environment, bryozoans rely on mechanical and chemical defense strategies [13]. As such, bryozoans and their associated microorganisms might be a source of biologically active substances.

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INT. MICROBIOL. Vol. 15, 2012

Although the phylum Bryozoa contains several thousands of recent species, studies on natural products have focused only on a few of them [43]. Bryozoan metabolites account only for about 1 % of marine natural products, according to the annually reviews of Blunt et al. [2]. Bryostatins are the most prominent compounds [35] extracted and isolated from bryozoans. However, bryostatins from Bugula neritina were show to be produced by the bacterial symbiont “Candidatus Endobugula sertula” [9], which is associated with the bryozoan. The resemblance of natural products originally isolated from marine macroorganisms to those identified from microorganisms has led to the assumption that these compounds are of microbial origin [21,22]. This has been a prominent reason to intensify studies on the production of bioactive compounds from bacteria and fungi associated with marine algae, sponges, other invertebrates and, in this study, the bryozoan Membranipora membranacea.

Host-associated bacteria, in particular those colonizing surfaces and living in biofilm communities, establish complex interactions with other microorganisms and with their hosts. Communication is mediated chemically, and the sum of all factors shapes the composition of microbial covers, which in many cases have been shown to differ considerably from the surrounding environment [11]. Therefore, regarding the discovery of bioactive compounds, marine surface-associated microorganisms represent excellent sources [10,31].

Bryozoan-associated microorganisms have been studied so far using microscopic [30,48], genetic [20], and cultivationbased [16,36] methods, as well as combinations thereof [14].

So far, no report is available on natural products produced by M. membranacea or bacteria associated with this bryozoan [49]. Therefore, the aim of this study was to isolate bacteria from the surface of M. membranacea by varying the culture media and growth conditions and then to analyze the ability of these isolates to produce antibacterial compounds.

Materials and methods Sampling site and sample preparation. Bryozoan samples collected by dredging in the Baltic Sea north of Læsø (Kattegat, coordinates 57° 28.3′ N, 11°10.4′ E, depth 20 m) were identified as Membranipora membranacea growing on phylloids of Saccharina latissima. Three separate bryozoan colonies were cut out, washed with sterile filtered surrounding sea water, and transferred aseptically into sterile tubes containing 50 % (v/v) glycerol and 3 % (w/v) sodium chloride. The tubes were immediately frozen and stored at –18 °C until further treatment. Excess bryozoan samples growing on algae were placed in a closeable beaker containing local sea water (total volume about one liter) and stored at 4 °C until used for media preparation. In addition, 2 l of seawater from the sampling site were collected before dredging, filtered through a 0.2-μm cellulose acetate filter, and added to selected agar media.

HEINDL ET AL.

Culture media. For the isolation of microorganisms four media were prepared: “bryozoan extract medium” (BM), “algal extract medium” (AM), diluted “Reasoner’s 2A medium” (R2Ad), and diluted “Difco all culture medium” (ACd). For antibiotic activity testing six media were prepared: “Väätänen nine salt solution medium” (VNSS), “Pseudoalteromonas specific medium” (PSA), “Reasoner’s 2A medium” (R2A), “Difco all culture medium” (AC), “marine broth medium” (MB), and “tryptic soy broth medium” (TSB). Isolation media were prepared as follows: the excess samples from the beaker were used for the media that resembled the natural habitat (BM and AM). Bryozoans were cut out from the algae and minced. An equivalent weight of 3 % (w/v) saline was added. This material was thoroughly blended with an Ultraturrax-homogenizer (IKA Werke, Germany), frozen at –100 °C, and lyophilized to obtain a “bryozoan extract.” The remaining algae were recombined with the seawater in a beaker, homogenized, frozen, and lyophilized to yield an “algal extract.” Both extracts were dissolved in sea water collected from the sampling site at concentrations of 0.06 % (w/v), yielding BM and AM media. Additionally, R2Ad medium (containing 0.01 % (w/v) Bacto yeast extract, Difco proteose peptone, Difco casamino acids, glucose, soluble starch; 0.006 % (w/v) sodium pyruvate and K2HPO4; and 0.00048 % (w/v) MgSO4), and ACd medium [0.06 % (w/v)], both with 3% (w/v) sea salt (Tropic Marin) were prepared.

Six media for the activity tests were prepared (all percentages are w/v): (i) VNSS medium with 0.1 % peptone from soymeal (Merck), 0.05% yeast extract, 0.05 % glucose, 0.5 % soluble starch, 0.001 % FeSO4·7H2O, 0.001 % NasHPO4·2H2O, 1.7 6% sodium chloride, 0.147 % Na2SO4, 0.008 % NaHCO3, 0.025 % KCl, 0.004 % KBr, 0.187 % MgCl2·6H2O, 0.041 % CaCl2·2H2O, 0.001 % SrCl2·6H2O, and 0.001 % H3BO3) (according to Mården et al. [28]); (ii) PSA medium with 0.2 % peptone from soymeal, 0.2 % yeast extract, 0.1 % glucose, 0.02 % KH2PO4, 0.005 % MgSO4·7H2O, 0.1 % CaCl2·2H2O, 0.01 % KBr, and 1.8 % sea salt (according to Kalinovskaya et al. [19]); (iii) R2A medium; (iv) AC medium were fivefold concentrated compared to isolation media and 3 % sea salt was added; (v) MB medium with 0.5 % peptone, 0.1 % yeast extract, and 3.14 % sea salt), and (vi) TSB medium with 0.3 % tryptic soy broth (Difco) and 2.5 % sodium chloride).

To all media, 1.5 % agar was added for solidification.

Isolation and cultivation of bacteria. For comparison, two different methods of sample preparation were applied. The first two bryozoan samples were crushed with a sterile micropestle, the third was processed with a Precellys 24 lysis & homogenization device with a hard tissue grinding MK28 kit (Bertin Technologies) at 6300 rpm for 20 s. Dilution series with sterile seawater were prepared (10–1 to 10–5) [16] and a 100-μl aliquot of each one was spread on agar plates containing four different media. In addition, pieces of the bryozoan samples were placed on plates with all four media. The plates were incubated at 25 °C in the dark until colonies were visible. These were picked and sub-cultured on MB agar plates. For preservation, pure cultures were suspended in liquid MB medium containing 5 % (v/v) DMSO and stored at –100 °C. Screening for inhibitory activities against indicator organisms. Bacterial isolates were grown on MB agar plates directly from the DMSO stock. Colonies were picked, and suspended in 1 ml sterile 3 % (w/v) saline, and a 15-μl aliquot of each one was pipetted onto agar plates with six different media. After growing for 3–4 days at room temperature (ca. 22 °C), the bacterial colonies were checked for the presence of clearance zones to anticipate false-positive results. The plates were then covered with 5 ml TSB soft agar (with 1 % (w/v) sodium chloride and 0.8 % (w/v) agar) containing one of the following indicator strains: Escherichia coli DSM 498, Bacillus subtilis subsp. spizizenii DSM 347, Staphylococcus lentus DSM 6672, Pseudomonas fluorescens NCIMB 10586, and the yeast Candida glabrata DSM 6425. The presence of inhibition zones was examined the following day as well as on days 3, 7 and 14.

BACTERIA FROM THE BRYOZOAN M. MEMBRANACEA

Amplification, sequencing, and classification of the isolates.

Amplification, sequencing, and phylogenetic analysis of the 16S rRNA gene sequences from the bacterial isolates were carried out as previously described [16]. Isolates were grouped into phylotypes by sequence similarities ≥99.5 %. Genus affiliation was determined using the RDP classifier [46].

If resulting confidence values were <60 % for the classified genus, the affiliation was specified by constructing phylogenetic trees and comparing BLAST results. This was the case for phylotypes 1 (Erwinia), 16 (Roseobacter), and 19 (Ruegeria).

In the case of strain BB77, a 16S rRNA gene clone library was constructed because direct sequencing of the PCR product was not successful.

The PCR product was purified after gel electrophoresis with a MinElute Gel extraction kit (Qiagen, Hilden, Germany) and excision of the band. The purified 16S rRNA gene was cloned into the pCR 2.1-TOPO vector and transformed into One Shot TOP10 chemically competent E. coli cells, using the TOPO TA cloning kit (Invitrogen, Karlsruhe, Germany) according to the manufacturer’s instructions. Correct insertion was checked by PCR with vector binding primers included in the kit. Fourteen clones were chosen for sequencing and classification of the inserted 16S rRNA gene as described above. The 16S rRNA gene sequences were deposited with the EMBL Nucleotide Sequence Database under the accession numbers FR693269 to FR693364.

Cluster analysis. The distribution patterns of phylotypes and antibiotic activities of isolates were compared by cluster analysis using the Bray-Curtis similarity index. Dendrograms were generated with the program PAST, applying the paired group algorithm [15].

Results Isolation of Membranipora membranacea associated bacteria. Four media were used for the isolation of bacteria, and all colonies grown on the agar plates were picked and purified on MB agar. The results are shown in Table 1. Most isolates (60.4 % of all 96 isolates) derived from media inoculated with a piece of the bryozoan; 30.2 % from dilution step 10–1, 8.3 % from step 10–2, and 1.0 % from step 10–4. Most isolates (43.8 % of all isolates) were obtained from ACd medium, fewer isolates resulted from R2Ad medium (34.4 %), BM medium (15.6 %), and AM medium (6.2 %). Portions of 29, 32 and 39 % of the isolates were obtained from the three bryozoan samples.

Phylogenetic affiliation. All isolates were classified phylogenetically based on 16S rRNA gene sequences and grouped into phylotypes according to sequence similarity values of ≥99.5 %. The resulting 49 phylotypes were affiliated with 28 different genera (Table 2). A cluster analysis regarding the presence and absence of phylotypes within the three bryozoan samples revealed low similarity values at ≤0.3, with samples 1 and 2 as the most related (Fig. 1A). This clearly indicated that distinct phylotypes were obtained from each sample, especially from the third bryozoan specimen,

INT. MICROBIOL. Vol.15, 2012

19

which was prepared differently as described above. The media had a significant influence on the types of bacteria isolated and resulted in dissimilar phylotype patterns. 37 phylotypes (75.5 %) were unique to one of the media, i.e., representatives of these phylotypes were not found elsewhere.

Most “unique” phylotypes derived from R2Ad medium (17 of the 23 phylotypes of this medium) followed by ACd medium (11 of 23), BM medium (6 of 12), and AM medium (3 of 6). Accordingly, similarity values for the phylotypes obtained from the different media were low and ranged from 0.1 to 0.3 (Fig. 1B).

The bacterial isolates were affiliated with four classes: Gammaproteobacteria (40 isolates), Alphaproteobacteria (21 isolates), Bacilli (12 isolates), and Actinobacteria (23 isolates). Representatives of these classes were isolated from each bryozoan sample, with the exception of the Alphaproteobacteria, which were not obtained from sample 3 (Fig. 2).

Gammaproteobacteria. The 40 isolates of the Gammaproteobacteria could be grouped into 15 phylotypes and assigned to ten genera: Erwinia, Pseudoalteromonas, Vibrio, Shewanella, Halomonas (4 phylotypes), Marinobacter, Psychrobacter, Microbulbifer (2 phylotypes), Alcanivorax, and Pseudomonas (2 phylotypes) (Fig. 3A; Table 2).

The majority of these bacteria (85%, covering 14 phylotypes) were picked from media inoculated with a piece of the bryozoan, and most isolates (47.5 %, covering 11 phylotypes) were obtained using ACd medium (Table 1).

Bacteria related to Halomonas were isolated from all three bryozoan samples and from all four media. In contrast, all eight isolates assigned to Pseudoalteromonas originated from sample 3 but were picked from three different isolation media (BM, R2Ad, Acd). Representatives of Psychrobacter, Microbulbifer, and Pseudomonas each derived from more than one medium and bryozoan sample. Single isolates were obtained of Vibrio, Shewanella, Marinobacter, Alcanivorax, and Erwinia. The latter (BB49, phylotype 1) could represent a new species, as indicated by 16S similarity values ≤ 97% with validly described species (Table 2). This is below the threshold value of 98.7 % for 16S similarity values proposed

by Stackebrandt and Ebers [44], which indicate genomic uniqueness of novel isolates.

Alphaproteobacteria. Members of 16 phylotypes (21 isolates) were affiliated with the Alphaproteobacteria and assigned to ten genera: Roseobacter, Roseovarius (2 phylotypes), Ruegeria (4 phylotypes), Jannaschia, Paracoccus,

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HEINDL ET AL.

INT. MICROBIOL. Vol. 15, 2012

Table 1. Origin, affiliation, and antimicrobial activity of Membranipora membranacea-associated bacteria Isolation Phylotype

Isolates

Affiliation

Sa

Medium

Activityb Bs

Sl

1

BB49

Erwinia

2

BM

2

BB66a

Pseudoalteromonas

3

BM

2

BB67

Pseudoalteromonas

3

R2Ad

2

BB68/ BB69

Pseudoalteromonas

3

R2Ad

M

2

BB71

Pseudoalteromonas

3

ACd

M

2

BB72b/ BB73/ BB74

Pseudoalteromonas

3

ACd

TM

3

BB8

Vibrio

1

ACd

4

BB86

Shewanella

1

ACd

5

BB12b

Halomonas

1

ACd

5

BB66b

Halomonas

3

BM

A

5

BB85

Halomonas

2

ACd

PRA

5

BB7

Halomonas

1

AM

6

BB10/ BB3

Halomonas

1

ACd

6

BB6

Halomonas

1

AM

6

BB65

Halomonas

3

BM

7

BB9

Halomonas

1

ACd

8

BB11b

Halomonas

1

ACd

8

BB81/ BB82b

Halomonas

1

R2Ad

9

BB15

Marinobacter

1

ACd

10

BB13/ BB14

Psychrobacter

1

R2Ad

10

BB20/ BB83

Psychrobacter

1

ACd

10

BB62

Psychrobacter

3

BM

TV

11

BB44

Microbulbifer

2

R2Ad

MVPRA

12

BB27

Microbulbifer

1

AM

R

12

BB34

Microbulbifer

2

ACd

PR

12

BB48

Microbulbifer

2

ACd

R

13

BB31

Alcanivorax

2

R2Ad

n.t.

14

BB5/ BB82a/ BB79

Pseudomonas

1

R2Ad

14

BB80

Pseudomonas

1

R2Ad

15

BB75a/ BB78

Pseudomonas

3

ACd

16

BB43

Roseobacter

2

R2Ad

R

17

BB22

Roseovarius

1

R2Ad

R

18

BB19

Roseovarius

1

AM

RA

19

BB50b

Ruegeria

2

BM

VRA

20

BB40

Ruegeria

2

R2Ad

MVPRA

Ec

Pf

Cg

T

T

TMVA

T

P

TA

A

A

n.t.

n.t.

n.t.

n.t.

TVA

(Continued on next page)

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21

Table 1. (Continued) Origin, affiliation, and antimicrobial activity of Membranipora membranacea-associated bacteria Isolation Phylotype

Isolates

Affiliation

Sa

Medium

Activityb Bs

21

BB2

Ruegeria

1

ACd

MVA

21

BB29

Ruegeria

2

R2Ad

RA

21

BB45

Ruegeria

2

R2Ad

MVR

22

BB33

Ruegeria

2

ACd

MVRA

23

BB23

Jannaschia

1

R2Ad

24

BB51b

Paracoccus

2

BM

TRA

25

BB54

Anderseniella

2

AM

R

26

BB18

Amorphus

1

AM

27

BB32

Erythrobacter

2

R2Ad

28

BB17

Erythrobacter

1

R2Ad

MV

29

BB1

Sphingopyxis

1

ACd

T

29

BB4/ BB46

Sphingopyxis

1/2

ACd

30

BB24

Sphingopyxis

1

R2Ad

VP

30

BB28

Sphingopyxis

2

ACd

M

31

BB21

Pelagibius

1

R2Ad

MPRA

32

BB58a/ BB58b

Staphylococcus

3

ACd

n.t.

32

BB60

Staphylococcus

3

R2Ad

P

32

BB26

Staphylococcus

1

ACd

33

BB41

Bacillus

2

R2Ad

34

BB50c

Bacillus

2

BM

34

BB51c

Bacillus

2

BM

35

BB42

Bacillus

2

36

BB52

Bacillus

37

BB61

38

Sl

n.t.

P

P

R2Ad

TVPRA

TMVPRA

2

R2Ad

T

PA

Bacillus

3

R2Ad

TP

BB75b

Exiguobacterium

3

ACd

TMPR

38

BB76

Exiguobacterium

3

ACd

M

39

BB36

Mycobacterium

2

ACd

39

BB55/ BB56/ BB57

Mycobacterium

3

ACd

39

BB64

Mycobacterium

3

BM

40

BB35

Mycobacterium

2

ACd

41

BB37

Mycobacterium

2

ACd

42

BB38

Mycobacterium

2

BM

43

BB63

Pseudonocardia

3

R2Ad

Ec

Pf

n.t.

n.t.

Cg

n.t.

VR

TP

(Continued on next page)

22

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INT. MICROBIOL. Vol. 15, 2012

Table 1. (Continued) Origin, affiliation, and antimicrobial activity of Membranipora membranacea-associated bacteria Isolation Phylotype

Isolates

Affiliation

Sa

Activityb

Medium

Bs

Sl

44

BB16

Streptomyces

1

BM

44

BB47a/ BB47b

Streptomyces

2

ACd

44

BB50a/ BB51a

Streptomyces

2

BM

44

BB11a

Streptomyces

1

ACd

TMVPRA

TMVPRA

44

BB84

Streptomyces

1

ACd

TMVPRA

TMVPRA

45

BB12a

Streptomyces

1

ACd

TMVPRA

TMVPRA

46

BB72a

Arthrobacter

3

ACd

47

BB59/ BB70

Arthrobacter

3

R2Ad

48

BB77

Arthrobacter

3

BM

49

BB25/ BB30

Microbacterium

1/2

R2Ad

Ec

Pf

Cg

P

T

Specimen.

Strains tested: Bs, B. subtilis; Sl, S. lentus; Ec, E. coli; Pf, P. fluorescens; Cg, C. glabrata. Media used: T, TSB; M, MB; V, VNSS; P, PSA; R, R2A; A, AC, n.t.: not tested.

a b

Anderseniella, Amorphus, Erythrobacter (2 phylotypes), Sphingopyxis (2 phylotypes), and Pelagibius (Fig. 3B; Table 2). In contrast to the Gammaproteobacteria, these isolates were predominantly picked from R2Ad medium (47.6 %, covering 9 phylotypes) and from the dilution series (71.4 %, covering 12 phylotypes) (Table 1). Except for the isolates affiliated with Sphingopyxis (both phylotypes) and Ruegeria (phylotype 21), all Alphaproteobacteria were single isolates (Table 2). New species were possibly represented by members of 13 phylotypes: Roseobacter (phylotype 16), Roseovarius (phylotype 17), Ruegeria (phylotypes 19 and 21), Jannaschia (phylotype 23), Paracoccus (phylotype 24), Anderseniella (phylotype 25), Amorphus (phylotype 26), Erythrobacter (phylotypes 27 and 28), Sphingopyxis (phylotypes 29 and 30), and Pelagibius (phylotype 31) (Table 2).

Fig. 1. Similarity dendrograms of shared phylotypes within the three Membranipora membranacea specimens (A), the four isolation media (B), as well as the antimicrobial activities expressed by the isolates on different media (C).

Int. Microbiol.

Bacilli. This class [27] was represented by seven phylotypes (12 isolates) affiliated with the genera Staphylococcus, Bacillus (5 phylotypes), and Exiguobacterium (Fig. 3C; Table 2). All but one of the isolates were obtained from plates inoculated with bryozoan samples 2 and 3 (41.7 % and 50 %, covering four and three phylotypes, respectively). ACd and R2Ad media yielded five isolates each. Bacillus and

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23

Table 2. Phylogenetic affiliation (RDP) and nearest type strains (BLAST) Phylotype

Representative

No. of isolates

Affiliation

Nearest type strain

Similarity (%)

Accession no.

1

BB49

1

Erwinia

Erwinia tasmaniensis Citrobacter gilleni

97 97

AM055716 AF025367

2

BB66a

8

Pseudoalteromonas

Pseudoalteromonas aliena

99

AY387858

3

BB8

1

Vibrio

Vibrio tasmaniensis

98

AJ316192

4

BB86

1

Shewanella

Shewanella kaireitica

98

AB094598

5

BB66b

4

Halomonas

Halomonas titanicae

99

FN433898

6

BB65

4

Halomonas

Halomonas boliviensis

99

AY245449

7

BB9

1

Halomonas

Halomonas boliviensis

99

AY245449

8

BB82b

3

Halomonas

Halomonas boliviensis

99

AY245449

9

BB15

1

Marinobacter

Marinobacter algicola

99

AY258110

10

BB13

5

Psychrobacter

Psychrobacter piscatorii

99

AB453700

11

BB44

1

Microbulbifer

Microbulbifer thermotolerans

99

AB124836

12

BB34

3

Microbulbifer

Microbulbifer epialgicus

98

AB266054

13

BB31

1

Alcanivorax

Alcanivorax venustensis

99

AF328762

14

BB82a

4

Pseudomonas

Pseudomonas perfectomarina

100

U65012

15

BB78

2

Pseudomonas

Pseudomonas chloritidismutans

99

AY017341

16

BB43

1

Roseobacter

Leisingera nanhaiensis Seohicola saemankumensis

97 96

FJ232451 EU221274

17

BB22

1

Roseovarius

Roseovarius aestuarii

97

EU156066

18

BB19

1

Roseovarius

Roseovarius aestuarii

99

EU156066

19

BB50b

1

Ruegeria

Ruegeria scottomollicae

96

AM905330

20

BB40

1

Ruegeria

Ruegeria scottomollicae

98

AM905330

21

BB2

3

Ruegeria

Ruegeria atlantica

96

D88526

22

BB33

1

Ruegeria

Ruegeria atlantica

99

D88526

23

BB23

1

Jannaschia

Jannaschia pohangensis

97

DQ643999

24

BB51b

1

Paracoccus

Paracoccus homiensis

97

DQ342239

25

BB54

1

Anderseniella

Anderseniella baltica

97

AM712634

26

BB18

1

Amorphus

Amorphus coralli

95

DQ097300

27

BB32

1

Erythrobacter

Erythrobacter longus

97

AF465835

28

BB17

1

Erythrobacter

Erythrobacter aquimaris

98

AY461441

29

BB1

3

Sphingopyxis

Sphingopyxis litoris

98

DQ781321

30

BB24

2

Sphingopyxis

Sphingopyxis litoris

98

DQ781321

31

BB21

1

Pelagibius

Pelagibius litoralis

92

DQ401091

32

BB58b

4

Staphylococcus

Staphylococcus epidermidis

99

D83363

33

BB41

1

Bacillus

Bacillus hwajinpoensis

98

AF541966 (Continued on next page)

24

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INT. MICROBIOL. Vol. 15, 2012

Table 2. (Continued) Phylogenetic affiliation (RDP) and nearest type strains (BLAST) Phylotype

Representative

Affiliation

34

BB50c

2

Bacillus

Bacillus hwajinpoensis

99

AF541966

35

BB42

1

Bacillus

Bacillus stratosphericus

99

AJ831841

36

BB52

1

Bacillus

Bacillus licheniformis

99

CP000002

37

BB61

1

Bacillus

Bacillus cereus

99

AE016877

38

BB76

2

Exiguobacterium

Exiguobacterium oxidotolerans

99

AB105164

39

BB64

5

Mycobacterium

Mycobacterium frederiksbergense

99

AJ276274

40

BB35

1

Mycobacterium

Mycobacterium aurum

99

X55595

41

BB37

1

Mycobacterium

Mycobacterium aurum

98

X55595

42

BB38

1

Mycobacterium

Mycobacterium komossense

97

X55591

43

BB63

1

Pseudonocardia

Pseudonocardia carboxydivorans

99

EF114314

44

BB51a

7

Streptomyces

Streptomyces griseorubens

99

AB184139

45

BB12a

1

Streptomyces

Streptomyces praecox

99

AB184293

46

BB72a

1

Arthrobacter

Arthrobacter parietis

99

AJ639830

47

BB70

2

Arthrobacter

Arthrobacter tumbae

97

AJ315069

48

BB77

1

Arthrobacter

Arthrobacter agilis

98

X80748

49

BB25

2

Microbacterium

Microbacterium schleiferi

99

Y17237

Exiguobacterium related isolates originated predominantly from agar plates inoculated with a bryozoan piece, while those of Staphylococcus derived from the dilution series (Table 1).

Actinobacteria. The 11 phylotypes (23 isolates) affiliated with this class [45] were assigned to the genera Mycobacterium (4 phylotypes), Streptomyces (2 phylotypes), Ar-

Nearest type strain

Similarity (%)

Accession no.

throbacter (3 phylotypes), Pseudonocardia, and Microbacterium (Fig. 3C; Table 2). The majority of the isolates was obtained from ACd medium (52.2 %, 6 phylotypes). Isolates affiliated with Streptomyces and Arthrobacter originated from media inoculated with a piece of the bryozoan specimens, while all others were obtained from dilution series (Table 1). Isolates belonging to Mycobacterium (phylotype

Int. Microbiol.

No. of isolates

Fig. 2. Relative abundance of isolates from three Membranipora membranacea specimens affiliated with the four bacterial classes observed in this study.

INT. MICROBIOL. Vol.15, 2012

(Continued on next page)

25

Int. Microbiol.

BACTERIA FROM THE BRYOZOAN M. MEMBRANACEA

Fig. 3. Maximum-likelihood tree constructed from 16S rRNA gene sequences showing the phylogenetic relationships of isolates from this study with closely related species and some other selected representatives of the Gammaproteobacteria (A), the Alphaproteobacteria (B) and gram-positive bacteria (C). Nonparametric bootstrapping analysis (100 datasets) was conducted. Values â&#x2030;Ľ 50 are shown. The scale bar indicates the number of substitutions per nucleotide position. The total number of represented sequences is given in square brackets.

42) and Arthrobacter (phylotype 47) might represent new species (Table 2).

Antibiotic activity. Antibiotic activity against at least one indicator strain was shown by 47 out of 93 tested bacteria. The vast majority of the tested isolates inhibited growth of grampositive test strains (45.2 % B. subtilis, 10.8% S. lentus, 7.5 % both), while only a minor part (5.4 %) was active against gram-negative indicator bacteria (1.1 % E. coli, 4.3% P. flu-

orescens) or against the yeast C. glabrata (1.1 %). Three isolates were not analyzed due to insufficient growth on the test media (Table 1).

Activity profiles on different media. The media used for the antibiotic tests had a clear influence on the pattern of antibiotic activities. Antibiotic activities were analyzed on six different media (TSB, MB, VNSS, PSA, R2A, and AC medium). 21 isolates each displayed activity on R2A

INT. MICROBIOL. Vol. 15, 2012

HEINDL ET AL.

(Continued on next page)

Int. Microbiol.

26

Fig. 3. (Continued) Maximum-likelihood tree constructed from 16S rRNA gene sequences showing the phylogenetic relationships of isolates from this study with closely related species and some other selected representatives of the Gammaproteobacteria (A), the Alphaproteobacteria (B) and gram-positive bacteria (C). Non-parametric bootstrapping analysis (100 datasets) was conducted. Values â&#x2030;Ľ 50 are shown. The scale bar indicates the number of substitutions per nucleotide position. The total number of represented sequences is given in square brackets.

and MB media, followed by TSB (20 isolates), AC (19 isolates), PSA (18 isolates), and VNSS (15 isolates). The majority (63.8 % of active isolates) inhibited growth of indicator

strains on a single medium or on two media. Only six isolates (12.8 % of active isolates) showed antibacterial activities on five or all six media (Table 1). A corresponding cluster analy-

INT. MICROBIOL. Vol.15, 2012

27

Int. Microbiol.

BACTERIA FROM THE BRYOZOAN M. MEMBRANACEA

Fig. 3. (Continued) Maximum-likelihood tree constructed from 16S rRNA gene sequences showing the phylogenetic relationships of isolates from this study with closely related species and some other selected representatives of the Gammaproteobacteria (A), the Alphaproteobacteria (B) and gram-positive bacteria (C). Non-parametric bootstrapping analysis (100 datasets) was conducted. Values â&#x2030;Ľ 50 are shown. The scale bar indicates the number of substitutions per nucleotide position. The total number of represented sequences is given in square brackets.

sis revealed similar activity patterns on R2A and AC as well as on MB and VNSS media, whereas patterns on TSB and PSA media were clearly different from those on the other media (Fig. 1C).

Activity profiles according to phylogenetic affiliation. While the Gammaproteobacteria were mainly active on TSB and MB media, most of the Alphaproteobacteria were active on R2A and AC media, and most of the Bacilli and Actinobacteria were active on PSA and TSB

plates. The number of isolates active on the different media and their affiliation with the four bacterial classes are shown in Fig. 4.

Gammaproteobacteria. Almost 50 % of the Gammaproteobacteria (19 of 39 tested strains) displayed antimicrobial properties. Antibiosis was mostly directed against B. subtilis (78.9 % of active isolates), followed by S. lentus (15.8 %) and P. fluorescens (10.5 %). Antibiotically active isolates were affiliated with Erwinia (phylotype 1), Shewanella (phylotype 4),

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HEINDL ET AL.

Int. Microbiol.

28

Fig. 4. Number of bioactive isolates on different growth media, (A) absolute counts, (B) percentage of active isolates within each bacterial class.

Marinobacter (phylotype 9), Pseudoalteromonas (phylotype 2; 7 out of 8 isolates), Halomonas (phylotype 5; 3 of 4), Psychrobacter (phylotype 10; 1 of 5), Microbulbifer (phylotypes 11 and 12; all isolates), and Pseudomonas (phylotype 14; 1 of 4). All Microbulbifer affiliated isolates were active on R2A medium, while most Pseudoalteromonas isolates displayed activity on MB agar plates. Whereas activity against B. subtilis was shown by almost all active strains, bacteria of phylotype 5 (assigned to Halomonas) were active against S. lentus and P. fluorescens instead. Only single isolates, Microbulbifer sp. BB44 (phylotype 11) on AC medium and Pseudoalteromonas sp BB67 (phylotype 2) on TSB medium, were active against E. coli or C. glabrata, respectively. No activity was exhibited by phylotypes 3 (Vibrio), 6, 7, 8 (all Halomonas), and 15 (Pseudomonas).

Alphaproteobacteria. Among the Alphaproteobacteria, > 75 % (16 out of 21 strains) were antimicrobially active. Activity was directed exclusively against B. subtilis. Active isolates were members of Roseobacter (phylotype 16), Roseovarius (phylotypes 17 and 18), Ruegeria (phylotypes 19 to 22), Paracoccus (phylotype

25), Erythrobacter (phylotype 28; 1 of 2 isolates), Sphingopyxis (phylotype 29; 1 of 3 isolates; and phylotype 30), and Pelagibius (phylotype 31). Most of the active strains (68.8 %) showed antimicrobial activities on R2A medium, followed by AC (50.0 %) medium. Especially isolates of the Roseobacter-clade, which was represented by phylotypes 16 to 23, displayed activity (90 % of these isolates) preferentially on R2A medium (88.9 % of the active isolates). The Jannaschia-related strain BB23 (phylotype 23) was the only non-active member of the Roseobacter-clade. In addition, strains of phylotypes 26 (Amorphus) and 27 (Erythrobacter) displayed no activity, nor did two strains of phylotype 29 (Sphingopyxis).

Bacilli. Antibiotic activity of isolates related to Bacilli was directed against B. subtilis (7 strains) or both B. subtilis and S. lentus (3 strains, all Bacillus, phylotypes 34, 35, and 36).

One of these isolates (BB42, phylotype 35) additionally showed activity against P. fluorescens and also expressed antimicrobial activities on all media. All Exiguobacterium affiliated strains (phylotype 38) were active against B. sub-

BACTERIA FROM THE BRYOZOAN M. MEMBRANACEA

tilis. One representative each of phylotypes 32 (Staphylococcus) and 34 (Bacillus) as well as phylotype 33 (Bacillus) did not impede the growth of any indicator strain.

Actinobacteria. Only five out of 23 Actinobacteria related isolates inhibited the growth of indicator strains. Four strains each were active against B. subtilis and S. lentus.

Three of them (phylotypes 44 and 45, both Streptomyces) were active against both gram-positive test strains on all six media. Isolate BB84 (phylotype 44) was additionally active against P. fluorescens. The Mycobacterium-related strain of phylotype 41 showed anti-S. lentus activity. No antibiotic active representatives were found in phylotypes 39, 40, and 42 (all Mycobacterium), 43 (Pseudonocardia), 46 to 48 (all Arthrobacter), and 49 (Microbacterium).

Discussion Phylogenetic affiliation of isolates. A phylogenetically diverse collection of isolates was obtained during this study from three specimens of Membranipora membranacea using different isolation media. Each medium featured a rather unique set of isolated phylotypes. This resulted in a highly diverse array of 96 isolates assigned to 49 phylotypes and 29 genera. Only one-third of the members of these genera had been isolated previously from comparable sources.

Three of these genera (Shewanella, Pseudoalteromonas, and Pseudomonas) were isolated in all three comparable studies on bryozoans from the North Sea and the Baltic Sea, as well as from Baltic Sea S. latissima samples [16,36,47]. Other genera (Bacillus, Arthrobacter, Vibrio, Psychrobacter, Ruegeria, Staphylococcus, Streptomyces) were also found, but not consistently, in all three studies.

The use of four different isolation media was undoubtedly an important factor in our success in obtaining such a diverse collection of bacteria. The high number of phylotypes that were exclusively found on one medium (75.5 %) and the resulting large differences in the phylotypes obtained from the different media (Fig. 1B) resulted in a “unique” set of isolates obtained from each medium. This observation correlates with the fact that different bacteria differ in their needs on nutrients, growth factors, salt composition, trace elements, etc., which cannot be covered by a single medium. Nonetheless, clear media preferences could not be narrowed down to a specific group of phylotypes or genera, as all media yielded phylogenetically diverse isolates. However, a certain bias of ACd medium towards the isolation of Gammaproteobacteria and

INT. MICROBIOL. Vol.15, 2012

29

Actinobacteria, as well as of R2Ad medium towards Alphaproteobacteria and Bacilli was noted.

Note that fewer isolates were obtained from media that contained algal or bryozoan extract (BM and AM media), approximately one third of the isolates that grew on the other media (R2Ad and ACd). This may be related to the compounds that originate from bryozoans or algae, which might be involved in the chemical defense mechanisms of these sessile organisms [13,34] and, as such, inhibit bacterial growth. In particular, marine algae are known as producers of a variety of active metabolites that prevent biofouling of their own surfaces [1]. Some of these natural products may be stable enough to express growth inhibiting properties even after autoclaving or long-term storage. Unsaturated fatty acids have been identified as antibacterial active agents from brown algae with activities especially directed against grampositive bacteria, and they maintain their antibiotic properties even if stored at room temperature for several years [40].

This fact correlates well with our finding that the fewest isolates were obtained from “algal extract medium” (AM) and that no gram-positive bacteria were obtained from this medium. Thus, detrimental effects of inhibitory compounds in this medium might have outbalanced the usually beneficial impact of habitat water demonstrated in previous studies [12,29].

As attempts to isolate bacteria from bryozoans are still very scarce compared to other marine sources, bryozoans provide a good source for the search for new bacteria and new antibiotic compounds. Validly described type strains that were originally isolated from bryozoans include Tenacibaculum adriaticum (Flavobacteria), Marinobacter bryozoorum (Gammaproteobacteria), and Paracoccus seriniphilus (Alphaproteobacteria) [17,37,39]. Single isolates of the latter two genera were also obtained in the present study. Quite significant was the finding that 15 out of 49 phylotypes of this study represented new species (some even new genera), applying the phylogenetic relationship according to Stackebrandt and Ebers [44]. Most significant was the high number of Alphaproteobacteria with 16S rRNA gene sequence similarities of or below 97 % to known species. Moreover, half of the phylotypes (16 to 23) of Alphaproteobacteria isolated in this study were affiliated with members of the Roseobacter lineage, which represents typical marine bacteria [6] and is abundant in bacterial communities associated, e.g., with algal blooms, biofilms, and cephalopods.

Note that the three bryozoan specimens yielded a similar amount of exclusive phylotypes: 38 phylotypes (77.6 %) originated from single samples exclusively, which resulted

30

INT. MICROBIOL. Vol. 15, 2012

in clear differences between the samples (Fig. 1A). Similar to the influence of the isolation media, this resulted in “unique” collections of bacteria obtained from each specimen. A previous cultivation-based study on the microbial diversity with samples of the North Sea bryozoan Flustra foliacea yielded similar results: although a great array of different isolation media was used and the same procedures were applied to all specimens, such that the distribution of bacterial taxons was highly divergent. Indeed, not a single genus could be found on all three samples [36]. Another culture-independent study on bacterial communities of bryozoans in the North Sea demonstrated species-specific associations for three of the four bryozoan species (Aspidelectra melolontha, Electra monostachys, and E. pilosa). In contrast, a site-dependent influence was observed in Conopeum reticulum specimens [20].

Antimicrobial activity. A large proportion, almost 50%, of the bacteria isolated from Membranipora membranacea revealed antibiotic activity, predominately against gram-positive test strains. This result is similar to those obtained in our previous study on the antibiotic activities of bryozoan-associated bacteria with the same indicator organisms [16]. However, isolates from the present work, with a few exceptions (Vibrio, Shewanella, Pseudoalteromonas, Pseudomonas), were affiliated with different genera and included also gram-positive representatives. Activity against the gram-positive bacteria Bacillus subtilis and Staphylococcus lentus could be advantageous on surfaces in situ, as members of both genera were also isolated from the bryozoan specimens.

The pattern of antibiotic activities was quite variable and strain-specific, phylotype-specific as well as genus-specific activity patterns were observed. All isolates of the genera Microbulbifer, Roseovarius, Ruegeria, and Exiguobacterium showed consistent genus-specific activity profiles. Moreover, this antibiosis was expressed on the same media, each with single exceptions of Ruegeria-affiliated strains (Table 1). In contrast, only some of the isolates related to Pseudoalteromonas, Psychrobacter, Pseudomonas, Sphingopyxis, Bacillus, Staphylococcus, or Streptomyces inhibited target organisms in a strain-specific pattern.

Note that strain-specific activities will more likely be detected if larger subsets of isolates of the considered group are included, such as those related to Pseudoalteromonas and Streptomyces in this study [25,26]. In addition, growth conditions and media are important factors for the production of

HEINDL ET AL.

bioactive compounds and should be considered in all studies on antibiosis and antibiotic activity of microorganisms. In this study, the influence of test media on antibiotic traits reflected this dependency of the bacterial isolates on a “suitable” environment. Most activities were expressed on one or two media only (63.8 %), whereas a minor fraction of the isolates produced growth inhibitory compounds on all or five media (12.8 %).

The activation of secondary metabolite pathways, which remain silent under standard laboratory conditions, is a feasible way to access new natural products in microorganisms [4,33]. Five isolates related to Sphingopyxis expressed activities against B. subtilis in different media or did not show activity on any of those used (Table 1). Altogether, the use of six different media resulted in a twofold increase in the discovery of antibiotic active bacteria compared to the results obtained with a single medium.

Microorganisms belong to the prominent producers of natural products in the marine environment. Among the best studied genera in terms of published metabolites are Streptomyces, Alteromonas, Bacillus, Vibrio, Pseudomonas, Actinomyces, and Pseudoalteromonas, all of which were also isolated from M. membranacea in this work. Other genera found in this study, such as Microbacterium, Marinobacter, Halomonas, Ruegeria, and Erythrobacter, have also contributed to published marine natural products but to a lesser extent [23]. However, only some of these compounds have been reported as antimicrobially active. For example, in the case of Pseudoalteromonas and Pseudomonas some secondary metabolites display antibiotic properties [3,18]. Spongeand ascidian-associated Microbulbifer strains produce variations of parabens [32,38]. As far as Alphaproteobacteria are concerned, only a few members of this class are known to produce antimicrobial metabolites. Among them are representatives of the Roseobacter clade, producing thiotropocin and its precursor tropodithietic acid [5,7,8]. Finally, well-documented producers of antimicrobially active compounds, such as Streptomyces strains, can be a source of novel compounds, although only few of the Actinobacteria isolated in this study were antibiotically active.

Recently, the production of the antibacterial compound mayamycin by a marine Streptomyces related strain reported to be induced by variation of culture conditions [41], which further supports the requirement for varying the culture conditions to find new antibiotic compounds. Novel species or known taxa of this work with as yet unknown antimicrobial properties, such as members of the genera Roseobacter,

BACTERIA FROM THE BRYOZOAN M. MEMBRANACEA

Roseovarius, Ruegeria, Paracoccus, Anderseniella, Erythrobacter, Sphingopyxis and Pelagibius, are candidates to be studied more intensively with regard to the production of new antimicrobial compounds.

Acknowledgements. This study was supported by the Ministry of Science, Economic Affairs and Transport of the State of Schleswig-Holstein (Germany) within the framework of the “Future Program of Economy,” which is co-financed by EFRE.

Competing interests. None declared.

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RESEARCH ARTICLE INTERNATIONAL MICROBIOLOGY (2012) 15:33-41 DOI: 10.2436/20.1501.01.155 ISSN: 1139-6709 www.im.microbios.org

Phylogenetic diversity of methyl-coenzyme M reductase (mcrA) gene and methanogenesis from trimethylamine in hypersaline environments José Q. García-Maldonado,1 Brad M. Bebout,2 Lourdes B. Celis,3 Alejandro López-Cortés1* 1

Laboratory of Molecular Microbial Ecology, Northwestern Center for Biological Research (CIBNOR), La Paz, Mexico.

Exobiology Branch, Ames Research Center, National Aeronautics and Space Administration, Moffett Field, CA, USA. 3 Applied Geosciences Division, Scientific and Technological Research Institute of San Luis Potosi (IPICYT), San Luis Potosi, Mexico

2

Received 13 January 2012 · Accepted 28 February 2012

Summary. Methanogens have been reported in complex microbial communities from hypersaline environments, but little is known about their phylogenetic diversity. In this work, methane concentrations in environmental gas samples were determined while methane production rates were measured in microcosm experiments with competitive and non-competitive substrates. In addition, the phylogenetic diversity of methanogens in microbial mats from two geographical locations was analyzed: the well studied Guerrero Negro hypersaline ecosystem, and a site not previously investigated, namely Laguna San Ignacio, Baja California Sur, Mexico. Methanogenesis in these microbial mats was suspected based on the detection of methane (in the range of 0.00086 to 3.204 %) in environmental gas samples. Microcosm experiments confirmed methane production by the mats and demonstrated that it was promoted only by non-competitive substrates (trimethylamine and methanol), suggesting that methylotrophy is the main characteristic process by which these hypersaline microbial mats produce methane. Phylogenetic analysis of amino acid sequences of the methyl coenzyme-M reductase (mcrA) gene from natural and manipulated samples revealed various methylotrophic methanogens belonging exclusively to the family Methanosarcinaceae. Moderately halophilic microorganisms of the genus Methanohalophilus were predominant (>60 % of mcrA sequences retrieved). Slightly halophilic and marine microorganisms of the genera Methanococcoides and Methanolobus, respectively, were also identified, but in lower abundances. [Int Microbiol 2012; 15(1):33-41] Keywords: Methanosarcinaceae · hypersaline environments · microbial mats · trimethylamine · gene mcrA

Introduction Methane-producing anaerobes (methanogens) were first identified by Woese and Fox in 1977 [45] as being phyloge*Corresponding author: A. López-Cortés Centro de Investigaciones Biológicas del Noroeste (CIBNOR) Mar Bermejo 195, Colonia Playa Palo de Santa Rita La Paz, B.C.S. 23096, México Tel. +52-6121238484. Fax +52-6121253625 E-mail: alopez04@cibnor.mx

netically distinct from all other cell types, and they are the founding members of the Archaea Domain [11].

Methanobacteria comprise a large and diverse Class whose members are the main constituents of the Kingdom Euryarchaeota [46]. The five Orders recognized thus far (Methanobacteriales, Methanococcales, Methanomicrobiales, Methanosarcinales, and Methanopyrales) have distinctive characteristics [11,24,44]; a novel Order of methanogens, Methanocellales, was proposed recently and is currently represented by a single strain [23,36]. Metha-

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nogenic archaea are vital for the anaerobic microbial degradation of organic waste; the resultant production of methane, a potent greenhouse gas, is valuable as a non-fossil fuel [29].

Methanogenic archaea obtain energy for growth by the oxidation of a limited number of substrates, with the concomitant production of methane gas. In freshwater aquatic environments, methanogens are quantitatively extremely important terminal oxidizers of organic matter. In marine and hypersaline environments, characterized by the presence of high concentrations of sulfate, sulfate-reducing bacteria, not methanogenic archaea, are the primary mediators of terminal anaerobic mineralization. Sulfate reducers [6] utilizing sulfate as terminal electron acceptor outcompete methanogens for CO2/H2, formate, and acetate. A limited number of other compounds such as methanol, monomethylamine (MMA), dimethylamine (DMA), trimethylamine (TMA), and dimethylsulfide, and some alcohols, such as isopropanol, isobutanol, cyclopentanol, and ethanol, are also substrates for some methanogens [44]. Of these, MMA, DMA, TMA and dimethylsulfide are unavailable to sulfate reducers, and so are referred to as “non-competitive substrates.” The relative lack of information on methanogens from hypersaline environments may stem, in part, from a belief that significant rates of methane production are unlikely to occur when sulfate concentrations exceed many tens of millimoles per liter (> 30 mM), as is commonly found in the brine of these environments [15].

Although sulfate-reducing organisms dominate anaerobic carbon consumption in marine microbial mats, methanogens persist and their activities vary both vertically and temporally in the mat system in response to non-competitive substrates such as TMA [32]. All methanogens that have been isolated to date from hypersaline environments use TMA as a catabolic substrate [44], and all the TMA-degrading methanogens from marine and hypersaline environments belong to the family Methanosarcinaceae.

Culture-dependent and culture-independent techniques targeting 16S rRNA and methyl coenzyme M reductase (mcrA) genes have been used to assess the phylogenetic diversity of methanogen assemblages [20,28,29,41].

Methane production has been studied most extensively in microbial mats from the salterns of Exportadora de Sal in Baja California Sur, Mexico. Incubation of the surface layers of microbial mats obtained near the photic zone predominantly yield Methanolobus spp., while Methanococcoides has been preferentially recovered from incubations of unconsolidated sediments underlying the mat. Methanohalophilus sequences in low abundances have been retrieved from sam-

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ples of 20- to 60-mm depth [33]. Clone libraries from microbial mats maintained in field-like conditions in a long-term greenhouse study consist exclusively of sequences related to methylotrophic members of the genus Methanolobus.

Increases in pore water methane concentrations under conditions of low sulfate (from 50 to 0 mM), in mats maintained for more than one year in that same greenhouse study, were associated with an increase in the abundance of putative hydrogenotrophic mcrA sequences related to Methanogenium [37].

Profoundly distinct vertical microenvironments at millimetric and micrometric scales have been recognized in microbial mats [4,6,34]. Differences in microbial community structure have also been observed in the horizontal direction, as a result of either different adaptations to gradients of salinity [5,10,35,38], thereby favoring marine and halophilic methanogens, or by the association of methanogen assemblages with different types of minerals. These observations suggest heterogeneity and the existence of three-dimensional microniches. The study of microbial community structure from as yet unexamined sites could broaden our understanding of the composition of the microbial community.

As such, the aim of the present work was to further our knowledge of methanogenesis in hypersaline environments.

To this end, we analyzed several features of both the wellstudied Guerrero Negro hypersaline ecosystem and a location not previously investigated, namely, Laguna San Ignacio, Baja California Sur, Mexico. The locations share a physiographic setting and climate [16]. Methane concentrations in environmental gas samples were investigated, together with the methane production rates obtained in microcosm experiments using hypersaline microbial mats samples amended with competitive and non-competitive substrates. Finally, the phylogenetic diversity of the methanogenic community was determined.

Materials and methods Site description and sample collection. Samples were collected from two locations along the Pacific coast in the state of Baja California Sur, Mexico in March 2009. Location 1, with one site, corresponded to a concentrating pond of Exportadora de Sal (ESSA), a solar salt works located near Guerrero Negro (28°N, 114°W). Location 2, with two studied sites (26° 50′ N, 113° 10′ W), was the evaporitic flats at Laguna San Ignacio (LSI), a natural hypersaline ecosystem. Three types of mats were studied: (i) thick (10 mm), soft, well-laminated, green-black microbial mats found in Area 1 of ESSA (site ESSA-A1); (ii) thick (7 mm), leathery textured, pustular, non-laminated, black microbial mats found at site H7 of LSI (site LSI-H7); and (iii) thick (8 mm), smooth, laminated, orange-pink microbial mats found at site H8 of LSI (site LSI-H8). All three microbial mats were growing on top of

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highly sulfidic sediments. Mat cores (1 cm deep × 1 cm diameter) were collected and stored in liquid nitrogen for molecular analysis. Five core samples 1 cm deep × 1cm of diameter (~10–20 g) from LSI sites were placed into 38ml serum vials, and water samples were saved for further microcosm experiments. Larger cores (8 cm deep × 8 cm diameter) were collected and transported at room temperature for subsequent measurements of sulfate content in the sediments.

Gas bubbles were collected by perturbing the mat and sediments and trapping the released bubbles in a capped, inverted funnel at flooded sites (ESSA-A1). From desiccated sites (LSI-H7 and LSI-H8), gas samples were collected directly from large (2–10 cm) bubble structures overlain by a raised microbial mat (sometimes called pustular mat) using a hypodermic needle fitted to a 5-ml syringe. All bubbles were then transferred with a syringe to an evacuated serum vial for quantification of the methane concentration by gas chromatography.

Microcosm experiment. Ten ml of deoxygenated (N2-purged) brine from the site was added to the serum vials with the mat samples to make a slurry. The vials were capped with blue butyl rubber stoppers and aluminum crimps, and the headspace was flushed with N2 to remove any O2. The slurries from the LSI sites were amended with TMA (15 mM), methanol (20 mM), formate (20 mM), acetate (20 mM), H2/CO2 (80/20 %), and sodium molybdate (2 mM). The sodium molybdate was added to specifically inhibit sulfate reduction in these samples. TMA (1 mM) was tested in slurries of ESSA-A1. Samples of ESSA-A1 were incubated at room temperature (ca.

25 °C) for 48 h, while LSI samples were incubated at 30 °C for 33 days. Gas samples were collected at different time points to monitor the methane concentration in the headspace, which allowed the specific production rates per gram of mat sample (nmol g–1 day–1) to be calculated. At the end of the experiment, vials of LSI were maintained at room temperature (25 °C) for 1 year and were subsequently re-activated with specific medium for Methanohalophilus DSM 525, using TMA (15 mM) and methanol (20 mM) as substrates, with an incubation at 30 °C for one week. Vials with DSM 525 medium that were previously enriched with TMA, and methanol were incubated again with the same substrates at the same concentration. Meanwhile, the vials used for methanogenesis inhibition were further incubated with methanol (data not shown); these new incubations were used for further molecular analyses.

Methane and sulfate determinations. Gas samples collected from all incubations and from environmental bubbles were used to measure the evolved methane and methane concentrations, respectively, by gas chromatography with a flame ionization detector (Shimadzu GC-14 A, Kyoto, Japan) equipped with a 2-m Porapak N column held at 40 °C [3]. Sulfate content in the sediments was analyzed by turbidimetry in an automated spectrophotometer (QuickChem series 8000 FIAS, Lachat Instruments, Hach, Loveland, CO, USA), using Morgan solution as extractant and BaCl2 for the precipitation [7].

DNA extraction. DNA was extracted from natural and enriched samples (PowerBiofilm DNA isolation kit 24000-50, Mo Bio Laboratories, Carlsbad,

CA, USA). Extraction was performed according to the manufacturer’s protocol, starting from 0.1 g samples and with 45-s bead-beating at speed 5 for cell lysis. DNA was observed in an agarose gel.

PCR of mcrA gene. PCRs were carried out in a final volume of 25 μl containing 2 μl of undiluted template DNA, 1 μl of each primer (10mM), and 12.5 μl of GoTaq master mix (Promega M7122, Madison, WI, USA). The mcrA gene was amplified using the primers developed by Luton et al. [29].

Amplification consisted of the following steps: 95 °C for 1 min, 35 cycles at 94 °C for 30 s each, 55–54.5 °C (decreasing 0.1 °C for the first five cycles) for 30 s each, 72 °C for 1 min, and a final elongation for 5 min at 72 °C. Denaturing gradient gel electrophoresis (DGGE). PCR products from DNA extracted from slurries in microcosm experiment with DSM 525 medium were separated using a modification of a previously published protocol [26]. The gels were stained with ethidium bromide and archived with a UV photograph documentor (BioDoc-It® Imaging System, GelDoc-It TS300, UVP, Upland, CA, USA). Representative bands were excised with a sterile scalpel and DNA was eluted in ultra clean, pure water overnight at 4 °C. DNA was re-amplified and sequenced by a commercial service.

Clone libraries. Fresh PCR products from natural samples from ESSAA1 and LSI-H8 were purified with an extraction kit (QIAquick gel extraction kit 28704, Qiagen, Valencia, CA, USA). One–2 μl of the purified PCR product was ligated into a cloning vector (pCR4-TOPO), which was then used to chemically transform TOP10 Escherichia coli competent cells according to the manufacturer’s instructions (Invitrogen, Carlsbad, CA, USA). White colonies were inoculated into 125 μl LB broth amended with 8 % (v/v) glycerol and carbenicillin (100 mg/ml), then incubated overnight at 37 °C. The inserts were verified by PCR using the M13F and M13R primers. Positive colonies were shipped at room temperature (ca. 23 °C) to a commercial firm for sequencing (Sequetech, Mountain View, CA, USA).

Phylogenetic analysis and nucleotide sequence accession numbers. All sequences were quality-filtered, trimmed, translated into protein sequences, and aligned with a custom database of methyl-coenzyme M reductase alpha protein sequences using bioinformatics software (Geneious 5.3, Biomatters, Auckland, NZ). Inferred amino acid sequences were clustered at 97 % similarity using CD-HIT [22]; representative sequences were queried against the NCBI non-redundant peptide sequence databases [2] to identify closest protein matches for phylogenetic analysis and tree building. Taxonomic assignment of the sequences was based on comparisons with sequences in the gene databases [1] with > 97 % similarity (Table 1).

Phylogenetic analyses were performed on the aligned amino acid sequences using maximum-parsimony and neighbor-joining evolutionary models in PAUP* version 4 (Sinauer Pub., Sunderland, MA, USA). The robustness of inferred tree topologies was evaluated by 1000 bootstrap resamplings of the data. Two topologies from the different analyses were similar and the presented tree was based on a neighbor-joining analysis, with

Table 1. Taxonomic assignment and accession numbers of all the sequences obtained in this study Genes

Techniques

35

Accession numbers

Taxonomic assignment/number of sequences

mcrA

DGGE

JF836061 – JF836067

Methanolobus/5 Methanococcoides/2

mcrA

Clone libraries

HQ131850 - HQ131869

Methanohalophilus/17 Methanolobus/3

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Table 2. Salinity in water samples, methane concentration in environmental gas samples and sulfate concentration in sediment samples from studied sites Site

Geoposition

Salinity (ppt)

Methane (%)

Sulfate (g/kg)

LSI-H7

26°47.998´N 113°07.649´W

~93

0.00086 (± 0.0003)

5.35 (± 1.36)

LSI-H8

26°45.223´N 113°07.406´W

>100

0.01293 (± 0.02)

8.36 (± 0.86)

ESSA-Area1

27°36.01´N 113°53.46´W

~50

3.204 (± 0.37)

4.12 (± 0.15)

bootstrap support of branch nodes only when supported by the two models.

All sequences determined in this study are available in GenBank (accession nos. HQ131850 to HQ131869 and JF836061 to JF836067) (Table 1).

Results and Discussion Detection and quantification of methane in environmental gas samples. Methane gas was detected from environmental samples from all field sites sampled (two evaporitic flats at Laguna San Ignacio and from area 1 of Exportadora de Sal at Laguna Ojo de Liebre).

Methane concentrations ranged from 0.00086 to 3.204 %.

Methane production has been observed in a wide variety of hypersaline environments [32], including stratified microbial mats and endoevaporites [8,13,15,39]. Similar results were previously reported for soft microbial mats from hypersaline environments near the field sites described here (Tazaz et al., personal communication).

Site ESSA-A1 had the highest concentrations of methane and the lowest concentration of sulfate in sediments. In contrast, sites LSI-H7 and LSI-H8 had the lowest concentrations of methane and the highest concentration of sulfate (Table 2).

These results are consistent with the hypothesis that methanogenesis is attenuated by sulfate-reducing bacteria at non-limiting sulfate concentrations [27]. Considering that methanogenic archaea are abundant only in habitats where electron acceptors such as SO42– are limiting, the cell densities of methanogens in the samples analyzed may have been low, since the concentrations of sulfate in sediments ranged from 4.12 to 8.36 g/kg. Nevertheless, methanogenesis is also constrained by ecological interactions (both stimulatory and competitive) and/or physicochemical environmental factors that act at biochemical or bioenergetic levels. In addition to physicochemical “extremes” (mainly temperature, salinity, and pH), other factors affect the environmental distribution of methanogens, which is constrained to a great extent by energy availability, the environmental distributions of oxy-

gen (biochemical inhibition), and the seawater anion sulfate content (competitive effects that act at a bioenergetic level) [27]. More exhaustive sampling will be needed to establish which physicochemical factors (salinity, temperature, pH, moisture, water activity and nutrients) have the greatest influence on microorganism distributions in different natural environments [9].

An alternative explanation for the low concentrations of methane, which will not be considered further here, is the aerobic and anaerobic oxidation of methane, which has been reported in microbial mats in salt marshes [4].

Production of methane from samples incubated with substrates. To further explore the metabolic processes giving rise to the methane detected at our field sites, microcosm incubations were conducted. Determination of the use of substrates in methanogenesis is of great relevance for explaining trophic relationships and the level of activity in nature. As an example, only members of the Order Methanosarcinales use methylamines as catabolic substrates. These organisms also use H2 + CO2, while the other four Orders only contain hydrogenotrophic or acetoclastic members.

Although the production of CH4 within the upper layer (0–20 mm) of hypersaline microbial mats has been previously correlated with the cyanobacterial production of H2 [13], microcosm experiments with different substrates showed that methanogenesis was not stimulated by the addition of hydrogen; rather, only TMA stimulated methane production in samples from ESSA-A1 and H7 and H8 of LSI (Fig. 1).

These results are congruent with previous studies that have recorded methane production in media designed to enrich methylotrophic methanogens on TMA using sulfate-rich samples from hypersaline microbial mats in the Napoli mud volcano [19]. Methanol additions also resulted in an increase in methane production in samples from LSI. Substrates utilized by hydrogenotrophic and acetoclastic methanogens (H2/CO2, formate and acetate) did not increase

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Fig. 1. Methane production rates from microbial mat samples from Laguna San Ignacio (sites H7 and H8) and the site ESSA-A1 in Guerrero Negro amended with site field water and non-competitive and competitive substrates under anoxic conditions. Samples of ESSA-A1 were incubated at room temperature (ca. 25 °C) for 48 h, while LSI samples were incubated at 30 °C for 33 days. Error bars indicate one standard deviation about the mean of two replicate samples.

methane production in the LSI samples (Fig. 1), confirming previous studies in which acetate, hydrogen, or methionine additions did not stimulate methanogenesis in freshly collected marsh sediments from intertidal sediments [43].

Acetoclastic and hydrogenotrophic methanogens, with their lower energetic yields, are therefore more susceptible than methylotrophic methanogenesis, which further explains the predominance of methylotrophic methanogens in hypersaline environments [30].

Our understanding of how methanogenesis is coupled to energy conservation has proceeded more slowly. As for all respirers, energy conservation is fundamentally chemiosmotic. A methyl transfer step plays a central role in most methanogenic pathways and directly drives the export of sodium ions. Other components of the energy conservation apparatus appear to differ in the methylotrophic and hydrogenotrophic methanogens. Methylotrophic metha-

nogens have cytochromes and a proton-translocating electron transport chain, which they use to conserve energy in the last, exergonic step in methanogenesis. Hydrogenotrophic methanogens, however, lack these components, and it is not clear how these organisms achieve a net positive gain in energy conservation, because the first step in methanogenesis from CO2 is endergonic [21]. A proposed mechanism involving electron bifurcation, in which exergonic electron flow directly drives endergonic electron flow, could resolve this conundrum [42].

To assess whether the activity of sulfate-reducing bacteria attenuated hydrogenotrophic methanogenesis at non-limiting sulfate concentrations, sulfate reduction was inhibited in microcosm experiments. Thus, the addition of sodium molybdate resulted in a 76â&#x20AC;&#x201C;78 % decrease in hydrogen sulfide production in the microcosm (data not shown), indicating a sharp decrease in sulfate reduction rates. This decrease

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Fig. 2. Phylogenetic tree based on comparison of inferred amino acid sequences of mcrA gene from methanogenic euryarchaeota from natural (clusters AD) and manipulated (clusters E, F) samples of microbial mats from hypersaline environments. Branch nodes supported by phylogenetic analysis (bootstrap values >95 % by both maximum parsimony; MP and neighbor-joining; NJ analyses) are indicated by filled circles. Open circles indicate >75 % bootstrap support by either MP or NJ analysis, while branch nodes without circles were not resolved (bootstrap value <75 %). Bootstraped trees were generated with 1000 resamplings. The tree is rooted using an environmental sequence related to anaerobic methanogenic-oxidizing archaea group 1 (ANME-1) as the outgroup. Abundance of each phylotype cluster (97 % identity) detected is stated in parentheses. The bar represents 0.2 changes per amino acid.

did not, however, increase methane production (Fig. 1), which might be explained by the low abundances of methanogens capable of using H2/CO2 or acetate as substrate in those samples. Methane production in the unamended samples best represented natural conditions, which are probably suboptimal for methylotrophic methanogens due to the low concentrations of methylamines (Kelley et al., personal communication).

Phylogenetic diversity of methanogens. Methanogens are frequently studied without cultivation, owing to a generally good correspondence between phylogeny and phenotype, which is less typical in other groups [30]. The phylogenetic diversity of methanogens was different for all the studied sites. Clone libraries of mcrA from natural ESSA-A1 samples consisted exclusively of sequences related to methy-

METHANOGENESIS IN HYPERSALINE ENVIRONMENTS

lotrophs of the Order Methanosarcinales, including moderately halophilic and marine members of the genera Methanohalophilus (8 sequences from cluster A and 3 sequences from cluster C) and Methanolobus (3 sequences from cluster B). Sequences closely related to members of the genera Methanohalophilus were also retrieved from natural samples from LSI-H8 (3 sequences from cluster A and 3 sequences from cluster D) (Fig. 2). This confirms earlier reports that the methanogenic community in hypersaline environments is dominated by methylotrophic methanogens [44]. The mcrA sequences retrieved from DGGE bands from LSI-H7 samples incubated with methanol and TMA were similar to sequences of uncultured Euryarchaeota that phylogenetic analyses showed to be distantly related to slightly halophilic and marine organisms of the genera Methanococcoides (2 sequences from cluster F) and Methanolobus (5 sequences from cluster E) (Fig. 2). However, we were not able to detect hydrogenotrophic or acetoclastic methanogens using the described molecular approaches, presumably due to the low abundances of methanogens that utilize the pathway involving CO2 reduction and acetoclastic reaction. Members of the genus Methanococcoides have been frequently found in anoxic marine sediments [17], but have also been reported for incubations of surface layers of microbial mats from hypersaline environments [33]. Enrichment cultures have also shown the presence of viable methylotrophic Methanococcoides in shallow sediment layers from hypersaline microbial mats in the Napoli mud volcano [19], suggesting that this genus is common in hypersaline environments.

Enrichments made it possible to assess the differential effects of environmental factors imposed on mixed microbial populations, as well as to select organisms capable of attacking or degrading particular substrates or of thriving under unusual conditions [18]. The retrieval, from enriched samples, of sequences that were not found in natural samples showed the importance of applying different approaches to the characterization of methanogenic community composition.

There is uncertainty about the phenotype of uncultivated organisms giving rise to sequences that cluster within the Euryarchaeota but outside of known methanogens [30].

Recent studies, however, have shown the importance of studying uncultured microorganisms, because novel microbes can be detected with molecular data [12]. Examples of this are the recently described novel major lineage of Nanohaloarchaea, from hypersaline microbial communities [31], and the report of a new candidate division, MSBL1, which branchs deeply within the Euryarchaeota, from

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extremely halophilic microbial communities in anaerobic sediments from a solar saltern [25].

Although mcrA sequences of the same genera of methylotrophic methanogens found in our samples have been reported from other hypersaline environments [33,37,40], the relative abundances of retrieved sequences were different at the field sites reported here. Previous studies at the site ESSA-A4 reported that Methanolobus and Methanococcoides sequences were the most abundant while Methanohalophilus-like sequences were retrieved in lower abundances [33,37]. In our study, > 60% of the sequences (17 sequences) were related to Methanohalophilus. These results show the importance of studying the microbial community composition from different hypersaline environments even though many identical phylogenetic groups are detected in sediments independent of their geographic location [17]. In contrast to the hypothesis that microbial diversity in hypersaline environments is essentially the same at different geographical locations, there can apparently be great heterogeneity in phylogenetic diversity between sites that share physiographic setting and climate.

We can conclude that geochemical and molecular evidence confirm the presence of methanogenesis in these hypersaline environments. The detection of methane suggests that physicochemical extreme conditions in hypersaline environments should not prevent methanogenesis. The retrieval of mcrA sequences showed that the methanogen community was dominated by moderately halophilic organisms of the genus Methanohalophilus (more than 60 of mcrA sequences retrieved). Nevertheless, slightly halophilic and marine organisms of the genera Methanococcoides and Methanolobus, respectively, were identified at lower abundances. These results suggest that the community composition of methanogens differs even in similar ecosystems. All sequences were related exclusively to methylotrophic members of the Family Methanosarcinaceae.

Acknowledgements. We thank Ira Fogel and Manuel Trasvi単a of CIBNOR for editorial improvements and sulfate determinations, respectively.

This work was supported by CONACYT grant 105969-2008-2012, CIBNOR grant PC0.18-2011 to A.L.C, and a grant from the NASA Exobiology Program to B.M.B. A grant from the NASA-Planetary Biology Internship Program 2009 at the Marine Biological Laboratory, Woods Hole, MA was received by J.Q.G.M.; he also has a CONACYT doctoral fellowship. We are grateful to Exportadora de Sal, S.A. de C.V. for access to the Guerrero Negro field site. We are thankful for the field and laboratory assistance of Jeff Chanton, Cheryl Kelley, Jennifer Poole, Amanda Tazaz, Angela Detweiler, and Natalia Trabal.

Competing interests. None declared.

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RESEARCH ARTICLE INTERNATIONAL MICROBIOLOGY (2012) 15:43-51 DOI: 10.2436/20.1501.01.157 ISSN: 1139-6709 www.im.microbios.org

Contribution of sortase A to the regulation of Listeria monocytogenes LPXTG surface proteins Javier F. Mariscotti,1¶ Juan J. Quereda,1¶ M. Graciela Pucciarelli1,2* 1

Spanish National Center of Biotecnology, Spanish National Research Council (CSIC), Madrid, Spain. 2Department of Molecular Biology and Center for Molecular Biology Severo Ochoa, Autonomous University of Madrid, Spain Received 21 January 2012 · Accepted 19 February 2012

Summary. Gram-positive bacteria of the genus Listeria contain many surface proteins covalently bound to the peptidoglycan. In the pathogenic species Listeria monocytogenes, some of these surface proteins mediate adhesion and entry into host cells. Specialized enzymes called sortases anchor these proteins to the cell wall by a mechanism involving processing and covalent linkage to the peptidoglycan. How bacteria coordinate the production of sortases and their respective protein substrates is currently unknown. The present work investigated whether the functional status of the sortase influences the level at which its cognate substrates are produced. The relative amounts of surface proteins containing an LPXTG sorting motif recognized by sortase A (StrA) were determined in isogenic wild-type and ΔsrtA strains of L. monocytogenes. The possibility of regulation at the transcriptional level was also examined. The results showed that the absence of SrtA did not affect the expression of any of the genes encoding LPXTG proteins. However, marked differences were found at the protein level for some substrates depending on the presence/absence of SrtA. In addition to the known “mis-sorting” of some LPXTG proteins caused by the absence of SrtA, the total amount of certain LPXTG protein species was lower in the ΔsrtA mutant. These data suggested that the rate of synthesis and/or the stability of a subset of LPXTG proteins could be regulated post-transcriptionally depending on the functionality of SrtA. For some LPXTG proteins, the absence of SrtA resulted in only a partial loss of the protein that remained bound to the peptidoglycan, thus providing support for additional modes of cell-wall association in some members of the LPXTG surface protein family. [Int Microbiol 2012; 15(1):43-51] Keywords: Listeria monocytogenes · sortases · peptidoglycan · surface proteins · covalent anchoring · bacterial regulation

Introduction Gram-positive bacteria have a thick cell wall consisting of a multilayered peptidoglycan [26]. This backbone is decorated

*Corresponding author: M.G. Pucciarelli Departamento de Biología Molecular / Centro de Biología Molecular Severo Ochoa Universidad Autónoma de Madrid 28049 Cantoblanco (Madrid), Spain Tel. +34-915854551. Fax +34-915854506 E-mail: mg.pucciarelli@uam.es ¶

Equal contributors

with teichoic (TA) and lipoteichoic acids (LTA) that are either directly bound to the peptidoglycan or tethered to the membrane, respectively. Besides these structures, the bacterial cell wall acts as a platform for the anchorage of a variety of surface proteins, many of which play essential physiological roles, for example, in nutrient acquisition, quorum sensing, biofilm formation, and host interactions [21]. The mechanisms of protein association with the cell wall are diverse and include non-covalent protein-peptidoglycan interactions via peptidoglycan-binding domains such as LysM and WxL; direct protein-TA and protein-LTA interactions; and covalent anchoring of the protein to the peptidoglycan lattice [1,21].

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In the latter case, the covalent association between the protein and the cell wall occurs upon recognition of the protein substrate by specialized enzymes called sortases [10,21].

Enzymatic cleavage of a C-terminal sorting motif in turn leads to the covalent anchorage of the protein to the peptidoglycan via a transpeptidation reaction. The first sorting motif, LPXTG, was identified in protein A of Staphylococcus aureus. Since then, distinct sortases with different specificities for different sorting motifs have been described [11,18, 24,30].

Genome sequencing studies have revealed that bacteria of the genus Listeria contain the largest family of genes encoding predicted surface proteins recognized by sortases. More than 40 genes of this class have been annotated in every Listeria genome sequenced to date [5,6,15,16]. Listeria contains two sortases, SrtA and SrtB, which recognize LPXTG and NPKSS/NAKTN sorting motifs, respectively [2,3, 20,27]. In the case of the pathogenic species L. monocytogenes, SrtA was reported to be essential for virulence [3].

This phenotype is linked to the major role in virulence played by distinct surface proteins containing the LPXTG sorting motif. Examples of these proteins are internalin A (InlA), Vip, InlJ and LapB, which promote bacterial adhesion and invasion of eukaryotic cells and modulate host immune responses [7,12,25,28,29]. By contrast, no evidence of a requirement for alternative sortase B in L. monocytogenes virulence has been found, and SrtB recognizes only the surface proteins Lmo2185 (SvpA) and Lmo2186 (SvpB) [2].

Our previous proteomic studies revealed that the absence of SrtA resulted in a cell-wall proteome devoid of the 13 LPXTG surface proteins that were detected by this technique in the wild-type strain [27]. This study was performed with peptidoglycan material obtained under harsh purification conditions, i.e., extensive boiling of the cell-wall extract in solutions containing high percentage of ionic detergents (4 % SDS). This methodology could have impacted, at least to some extent, the ability to detect LPXTG surface proteins that, in the absence of SrtA, were retained in the cell wall by non-covalent associations with other proteins or cell-wall components. These hypothetical cell-wall–protein associations might naturally play additional roles in the fine-tuning of protein function. A similar diversity of associations for a single surface protein with the cell wall was recently demonstrated for the L. monocytogenes ActA protein, involved in actin-tail polymerization in intracellular bacteria [13]. Thus, in addition to being tethered to the membrane by its C-terminal hydrophobic domain, ActA strongly associates with the cell wall in bacteria that proliferate within epithelial cells

[13,14]. Variable associations of surface proteins with the cell wall have also been reported for members of the same protein family. Specifically, L. monocytogenes surface proteins covalently bound to the peptidoglycan by SrtA, such as InlA, InlH and InlJ, or by alternative sortase SrtB, such as Lmo2185 (SvpA), were recently shown to preferentially locate in distinct regions of the bacterial surface [4]. Another aspect, poorly understood, is to what extent, if any, sortases contribute to regulating the production of surface proteins that they recognize for anchoring to peptidoglycan. Consistent with what is known for essentially every enzymatic process, a tight co-regulation of the relative amount of the sorting enzyme and its substrates can be expected. Based on these considerations, in the present study the effect of a lack of L. monocytogenes SrtA on the expression of LPXTG surface proteins was examined. In addition, we analyzed whether the absence of SrtA results in changes in both the relative amounts and the subcellular distribution of these proteins.

Materials and methods Bacterial strains and growth conditions. The Listeria monocytogenes serotype 1/2a strain used in this study was EGDe; its genome has been sequenced [15]. The isogenic mutant derivative was the previously described ΔsrtA BUG 1777 [3]. Both were grown at 37 ºC to exponential phase (OD600 = 0.2) in brain-heart infusion (BHI) medium with shaking (150 rpm). For experiments involving stationary-phase regulation, the bacteria were grown at 37 ºC in BHI medium under static, non-shaking conditions.

In these stationary-phase conditions, bacteria grown display high invasion rates when exposed to cultured eukaryotic cell lines [13].

Bacterial fractionation and Western blot analysis. Fractions containing cytosolic, membrane, cell-wall, and secreted proteins of L. monocytogenes grown at 37 ºC in BHI media were obtained as described previously [27], except that the pellets were incubated in lysis buffer for 4 h instead of 1 h. The supernatants, corresponding to the cell-wall fractions, were filtered and precipitated on ice in 15 % TCA for 1 h. Proteins present in the cell-wall fraction were recovered by centrifugation at 29,000 ×g for 20 min at 4 ºC . The pellet was washed in cold acetone and centrifuged again under the same conditions. A volume of the cell-wall fraction was analyzed

by SDS-PAGE followed by Western blotting using polyclonal rabbit immune sera recognizing the LPXTG proteins Lmo0130, Lmo0159, Lmo0160, Lmo0263 (InlH), Lmo0610, and Lmo0880, and mouse monoclonal antibody anti-Lmo0434 (InlA). Other proteins analyzed in these fractions as controls were the StrB substrate SvpA and the peptidoglycan hydrolase Iap (P60). Polyclonal rabbit antibody recognizing P60 was a gift of Dr. Andreas Bubert (University of Würzburg, Germany). Mouse monoclonal anti-Lmo0434 (InlA) and polyclonal rabbit anti-SvpA were a gift of Prof.

Pascale Cossart (Institut Pasteur, Paris, France).

RNA preparation and RT-PCR assays. RNA was purified in three independent experiments from bacteria grown in 10 mL of BHI at exponential phase (OD600 = 0.2). Total RNA from bacterial pellets was extracted

REGULATION OF L. MONOCYTOGENES SURFACE PROTEINS

using the TRIzol reagent method (Invitrogen) [31]. RNA was treated with DNase I for 30 min at 37 ºC (Turbo DNA-free kit, Ambion/Applied Biosystems). RNA integrity and concentration were assessed by agaroseTAE electrophoresis and absorbance at 260 nm, respectively. RT-PCR was performed using a one-step RT-PCR kit (Qiagen). Briefly, RNA samples were diluted to 25 ng/μl and used in the RT-PCR, which was carried out in a final volume of 25 μl consisting of 5 μl buffer (5×), 1 μl of dNTP (10 mM), 3 μl each of the forward and reverse primers (5 mM), 1 μl of RT-PCR enzyme mix, 40 ng of RNA, and RNase-free water. RT-PCR cycling conditions were as follows: 50 ºC for 35 min and 95 ºC for 15 min, followed by 30 cycles of 94 ºC for 30 s, 55 ºC for 30 s, and 72 ºC for 40 s and then an extra elongation step at 72 ºC for 10 min. Oligonucleotides used in these RTPCR assays were designed using the software Primer Express v3.0 (Applied Biosystems) and are listed in Table 1.

Results and Discussion The expression of L. monocytogenes genes encoding LPXTG surface proteins is unaltered in a strain lacking sortase A. Sortases anchor surface proteins to the peptidoglycan of gram-positive bacteria once most of the protein substrate has passed through the membrane and is further retained there by its C-terminal region rich in hydrophobic amino acids [21]. At this stage, the sorting motif of the substrate protein is exposed on the outer leaflet of the membrane for recognition and cleavage by the sortase, leading to the covalent anchoring of the protein substrate to the peptidoglycan. Considering that the encounter between enzyme and substrate must be finely modulated spatially, temporally, and stoichiometrically, we examined whether the relative amounts of the sortase and its substrates were, to some extent, co-regulated. This type of regulation would prevent the potentially detrimental accumulation of surface proteins in their precursor form, i.e., tethered to the membrane, when either the level or the activity of SrtA was not optimal. This hypothesis was tested using the intracellular bacterial pathogen L. monocytogenes, which contains a large family of surface proteins bearing the LPXTG motif, recognized by StrA. Our first goal was to determine whether the expression of any of the genes encoding thesetranscriptionally regulated any of the genes encoding these LPXTG surface proteins could be modulated by StrA. Thus, the expression of the respective genes was analyzed by RT-PCR using total RNA purified from exponential cultures of the L. monocytogenes isogenic strains EGDe (wild-type) and BUG1777 (ΔsrtA). The genome of the EGDe strain has been sequenced and is known to contain 41 LPXTG-encoding genes [15]. As expected, the relative expression levels of these genes in the wild-type strain varied,

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with some genes more highly expressed than others (Fig. 1).

However, there were no major differences in the relative amounts of their respective transcripts between the wild-type and ΔsrtA isogenic strains (Fig. 1). The only differences between the wild-type and srtA strains were in the relative levels of the lmo0263 (inlH), lmo0880, lmo1289 and lmo1666 (lapB) transcripts (Fig. 1).

Further expression analyses by quantitative real-time PCR (qRT-PCR), however, ruled out the differential expression of any of these four genes (data not shown). Altogether, these data led us to conclude that the presence of SrtA on the L. monocytogenes cell surface does not, per se, constitute a cellular signal to modulate the expression of the genes encoding its protein substrates. Rather, as sugggested in previous reports [7,9,28,29], regulation appears to be governed mostly at the level of regulators that modulate the expression of a specific set of genes encoding the LPXTG surface proteins.

Representative examples of these genes are lmo0434 (inlA), regulated by PrfA and SigB [23], and lmo0263 (inlH), lmo0610, lmo0880 and lmo2085, regulated by SigB [9,17,31].

The lack of SrtA does not completely abolish the strong attachment of certain LPXTG surface proteins to cell-wall peptidoglycan. Since SrtA does not seem to regulate the expression of any of the 41 genes encoding the protein substrates of this sorting enzyme, we asked whether post-transcriptional regulatory mechanisms relied on the functional status of the sortase. To answer this question, it was necessary to quantify the relative levels of LPXTG proteins in the absence/presence of SrtA. Antibodies for all LPXTG surface proteins encoded in the genome of L. monocytogenes strain EGDe were generated, with the exception of anti-InlA and antiInlJ sera, available from other sources. While all the immune sera recognized the recombinant protein produced for immunization purposes, only some of these sera readily recognized native LPXTG proteins in cell-wall extracts of L. monocytogenes (data not shown). In most cases, the latter consisted of those LPXTG proteins encoded by highly expressed genes (Fig. 1 and data not shown), such as InlA, Lmo0130, Lmo0160, Lmo0263 (InlH) and Lmo0610. In the other cases, differences were observed between the RT-PCR and Western-blot data, perhaps reflecting the existence of post-transcriptional regulatory mechanisms. Alternatively, and irrespective of the amount of protein produced, a distinct topology of the protein within the cell wall could render it either more resistant or labile to extraction from the peptidoglycan lattice.

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Table 1. Oligonucleotides used in this studya Oligonucleotide

Forward sequence (5′→3′)

Reverse sequence (5′→3′)

iap

AAAGCAACTATCGCGGCTAC

TCTTGAACAGAAACACCGTA

lmo2026

CAGCCTCCACTTCATGGATTG

TCACCATGGGAAACTCCTGTAA

lmo2396

CCAGTTACCGGCCTAATTAAAGATT

TGGTATGCAGGATTGGACCAT

inlJ

GCAGCAACGAATGATGTTATTGATA

GCTAAGATCAAGGGTGGTAATGTTGT

lmo0171

ATCATCCAGTGAAGAAGCTGCAA

GTGAGCTCATTGTTGTTCACATTG

lmo0732

ACCAAAAACGGTCGACGTAAA

AGGTATTTGCCCAACCAAACG

lmo0175

TACGGACTTACAGGTTTCTGTTGGT

GCTTTAACAACTGGTGCGCTAA

lmo0327

CAATAACTGCCTTGCGAGTAACA

TGGAACTTTCACTAAGCGGTTATTATTT

inlI

CTTGGTTATTTAGCGCCTTTTGA

CCTGTTGCAGCCGCATTT

lmo0835

CAGATGGAGCGGAGATTGCT

TATGCAAGCCATCTATGCTAATGTC

lmo1413

GGCAATGCAGCTTATGTGCTT

AGTTTCACCGAGTTTTCCGGATA

lmo2178

GACTATGGATTCTCTTTTACCTGGTGAT

TTCCCTGAAAATGCAAAGTTCA

lmo2179

TTGAATCTGGTCAGACGGTAAACTAT

CTGCGCTTAATTCACCTGTTACTACT

inlA

AATATTAGTATTTGGCAGCGGAGTATG

GGCGTTATGTCCGTAAGTTGATT

lmo0130

GCGACAACTGACAATGCTATCC

GCCTTCTGCGACTGTGTTTG

lmo0159

CGATCTCGCTTTTGACGTGAA

TTACTAGCATCTACCCAACCATATTTGA

lmo0160

AGGGCAATTAACAGGCGATAAC

GACGCATCCAACACGTATCCT

inlG

AGTAGATAAAATGCCGGCTACGA

TTTCCGGGAGCTGTTTGAGA

inlH

CAGTAGCGCCAACGAAGGAT

GCGTCGCTGTTCCTAGCAA

inlE

GGTGAGCTTATTGCACCGGATA

TGCGTCATACCATCCATCGA

lmo0320

CCGTTTATCATCCAAGTGGCTAT

GCGCATCCAATTGTTGTTGA

lmo0331

GCCACCGTAACAAGCAGCTTAC

ACCTTGGCTTCGTGCATCTAG

inlF

ACCGAACTTGAAGTGGTCTTTACC

ATTTGCCCGGTTGTGAAGTC

lmo0435

GACTTTGTGGATGTTAGTGCGAAT

TGCTACGTCCTCCCCGTTT

lmo0463

GAGTTCAAGCTAGTCAAACAGTGGTT

CCAAATAAGGACGAGCACTAAGC

lmo0514

TGCTGCAGGACTCAAAGCAA

TGTCCACTGTCGCTTGTAGTCA

lmo0550

GCGTCGCCATTTTATGTGAA

ATGAAACCAACCCCTAGTAAAACAA

lmo0610

GTTTAAAAGCAACGCCAACACA

GTGGCGTCGGAGGTTCATT

lmo0627

TCACAAACCCAGTTGACATTCC

AATTTGCCGCGCCAGAA

lmo0725

TGCTGTTTTCATTTCACTTGGATT

GCAAGCGCGACAAGTACCA

lmo0801

AGATCCTGCCATGGCTAATGAA

TTAGCTAAATCAGCCACTGGTGAT

lmo0842

AACGGATGTGTCAGTGGGAGTATA

CCAGCCACGCTTAAGAAGGT

lmo0880

TAAAGTGCGAGTGGCGTATGA

TTTACCGGAAATGCGATTGG

lmo1115

ACGAAAGCTGGAGAAGAGGCTAA

AGAGAACGGAATAGGGCGTGTA

lmo1136

GCAACATCAACTACACTCGAGACA

AATTGTTACGGGCAAAGCGTAT

lmo1289

CTGAAATCAAAGCCACAACCAA

TAGGATTTCTTGAGTTTGAATGTTGAAC

lmo1290

ACAGGCATTGAATATGCTCACAATA

TGGCATTGTTTTAAGTGGCATAA

lmo1666

TTATGACTTCGACTGCTGACGAAA

TCGGACTTCCAACATCAACTACA

lmo1799

GATGATGGTAGCGCCTGTCA

CCGCCAATTCCTAAATAAGCAA

lmo2085

GTATTCAGCAAGATAGCGAAGAACCT

TCGTCGTTATTCCCGCATCTA

lmo2576

TCGTCGTTTACAATTGGTCGAT

CGCTGATTGGGAAAACGATT

lmo2714

CCGGCAGATGAAAACTTTGG

GTCACCTGTGCTTGGCAAATC

b

Oligonucleotides were designed using the Primer Express v3.0 software (Applied Biosystems).

Oligonucleotide sequences taken from the study of Cabanes et al. [7].

a b

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Fig. 1. As seen in real-time PCR (RT-PCR) assays, there was no difference between EGDe (wild-type) and BUG1777 (ΔsrtA) isogenic strains with respect to the expression of the 41 Listeria monocytogenes genes encoding LPXTG surface proteins. Bacteria were grown to exponential phase (OD = 0.2) in BHI medium at 37 ºC. The iap gene, encoding the peptidoglycan hydrolase P60, which is not recognized by SrtA, was included as control. Note that, despite the variability in expression between some genes encoding LPXTG proteins, there were no differences among the two strains tested.

Among the LPXTG proteins better detected by our immune sera were Lmo0130, Lmo0159, Lmo0160, Lmo0263 (InlH), Lmo0610 and Lmo0880. Therefore, the relative amounts of these six LPXTG proteins and Lmo0434 (InlA) were examined in the cell-wall and extracellular-medium fractions of exponentially growing isogenic wild-type and ΔsrtA strains. The cell-wall-associated protein Iap (also known as P60), which binds to peptidoglycan non-covalently by its LysM domain, and the SrtB substrate Lmo2185 (SvpA) were included as controls of surface protein not recognized by SrtA. In the ΔsrtA strain, the lack of SrtA resulted in a significant decrease in LPXTG protein levels in the cell wall and a concomitant increase in the amount of protein released into the extracellular medium (Fig. 2A). As expected, the mis-sorting of the LPXTG proteins by the ΔsrtA mutant was not observed for the P60 and SvpA proteins, which do not depend on SrtA

for their cell-wall association (Fig. 2A). Instead, consistent with the mode of association reported for these two proteins, P60 was mostly secreted into the medium while SvpA was retained in the cell wall by the covalent linkage mediated by SrtB (Fig. 2A). Of interest, the levels of some the LPXTG proteins examined, such as Lmo0610 and Lmo0880, were lower in the ΔsrtA mutant than in the wild-type strain, when considering both the cell-wall and the extracellular fractions (Fig. 2A). Slight decreases in the amounts of two other LPXTG proteins, Lmo0159 and Lmo0160, were also noted while the remaining two LPXTG proteins examined, Lmo0263 (InlH) and Lmo0434 (InlA), were detected in similar amounts irrespective of the presence/absence of SrtA (Fig. 2A).

Of note, some of these LPXTG proteins were detected in the Western immunoassays as double or multiple bands.

However, they were found to be specific when compared

INT. MICROBIOL. Vol. 15, 2012

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Fig. 2. (A) Western-blot assays showing the relative levels of distinct LPXTG proteins (Lmo0130, Lmo0159, Lmo0160, Lmo0263 -InlH-, Lmo0434 -InlA-, Lmo0610, and Lmo0880) in the cell wall (CW) and extracellular medium (‘Secreted’) fractions of L. monocytogenes EGDe (wild-type) and BUG1777 (ΔsrtA) strains grown to exponential phase (OD600 = 0.2) in BHI medium at 37 ºC. Non-SrtA substrates, such as the peptidoglycan hydrolase Iap (P60) and the SrtB substrate Lmo2185 (SvpA), were run as controls. (B) Western assays showing the distribution of the LPXTG proteins Lmo0130 and Lmo0160 in subcellular fractions of bacteria grown as in panel A. Cyt: cytosol; Mb: membrane; Sec.: secreted.

with mutants defective in these LPXTG proteins (data not shown). This multiplicity of bands for sortase substrates in both peptidoglycan and secreted fractions has been previously reported [2] and might reflect the attachment of muropeptides to proteins extracted from the cell wall by muramidase digestion. In these two fractions (cell wall and secreted), differences in protein mobility in the gel are not likely to be due to unprocessed precursor forms of these proteins, as they would be almost certainly retained at the membrane.

To determine whether the LPXTG proteins detected at lower levels in the cell wall and extracellular fractions of the ΔsrtA mutant were produced at normal levels but accumulated in the membrane or cytosol, these two subcellular fractions were examined for the representative cases of Lmo0130 and Lmo0160. As shown in Fig. 2B, Lm0130 was not detect-

ed in either the cytosol or the membrane fraction of wild-type and ΔsrtA strains. These data suggested that Lmo0130 could be subjected to post-transcriptional regulation, with the involvement of SrtA. By contrast, Lmo0160 was detected in the cytosol and membrane fractions of the ΔsrtA mutant (Fig. 2B). Considering that Lmo0160 was also detected in the cellwall and extracellular medium fractions and that its total amount was indistinguishable between wild-type and ΔsrtA strains (Fig. 2B), the production of this protein might have been constitutive. Nonetheless, it remains to be explained why a portion of Lmo0160 molecules remained bound to the peptidoglycan in the absence of SrtA. This behavior, shared by other LPXTG proteins such as Lmo0263 (InlH) and Lmo0434 (InlA) (Fig. 2A), suggests that the presence of additional modules of these proteins favored the retention of

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Fig. 3. (A) Western-blot assays depicting the relative levels of the LPXTG proteins Lmo0130, Lmo0159, Lmo0160, Lmo0434 -InlA-, and Lmo0610 in the cell wall (CW) and extracellular medium ('Secreted') fractions of L. monocytogenes EGDe (wild-type) and BUG1777 (ΔsrtA) strains grown to stationary phase (OD600 = 1.0) in BHI medium at 37 ºC under non-shaking conditions. Non-SrtA substrates, such as the peptidoglycan hydrolase Iap (P60) and the SrtB substrate Lmo2185 (SvpA), were run as controls. (B) Western blots showing the distribution of the SigB-regulated LPXTG proteins Lmo0263 (InlH) and Lmo0880 in subcellular fractions prepared from wild-type and ΔsrtA bacteria grown as in panel A. In the anti-InlH immunoblot, the asterisk denotes an unspecific band not related to the LPXTG protein. Cyt: cytosol; Mb: membrane; Sec.: secreted.

the protein in the cell wall. The existence of additional peptidoglycan-binding domains in LPXTG surface proteins is, to our knowledge, unreported. Note that in-silico analyses have demonstrated that the L. monocytogenes Lmo0880 harbors a LysM domain [1], as do other surface proteins not recognized by sortases, such as P60. Our subcellular fractionation studies showed, however, that Lmo0880 was not extensively retained in the ΔsrtA mutant. This result is consistent with the low percentage of P60—a surface protein that includes two LysM domains—able to withstand the harsh peptidoglycan purification method, involving extensive boiling in SDS-containing solutions (Fig. 2A) [26]. Overall, our observations point to as-yet-unidentified modes of association—and thus independent of the action of SrtA—between peptidoglycan and certain LPXTG surface proteins. These associations would presumably also rely on domain(s) other than LysM and perhaps contained in LPXTG proteins, such as Lmo0263 (InlH), Lmo0434 (InlA) and Lmo0160. That some of these proteins might be able to form macromolecular complexes is also an interesting possibility.

Growth arrest also regulates the production of certain L. monocytogenes LPXTG proteins in an SrtA-dependent manner. Previous studies have shown that certain L. monocytogenes LPXTG proteins, namely, Lmo0263 (InlH), Lmo0610, Lmo0880, and Lmo2085, are regulated by the alternative sigma factor SigB [17,19,25].

SigB is also known to control cellular responses to diverse stresses, such as high osmolarity and acidic pH, to govern bacterial adaptation to conditions of nutrient limitation, and to modulate the expression of other virulence genes (reviewed in [22]). Our proteomic studies showed that many LPXTG proteins are up-regulated when L. monocytogenes is grown to stationary phase [8,13]. Based on these observations, we analyzed whether the up-regulation of these LPXTG proteins in growth-arrested cells requires a functional sortase enzyme. Thus, the distribution of different LPXTG proteins was examined in cell-wall and extracellular-medium extracts of L. monocytogenes wild-type and ΔsrtA strains grown to stationary phase. In a first set of experiments, the distribution of Lmo0130, Lmo0159, Lmo0160, Lmo0610,

50

MARISCOTTI ET AL.

INT. MICROBIOL. Vol. 15, 2012

and Lmo0434 (InlA) was examined. In all cases, the relative amount of these LPXTG proteins anchored to the peptidoglycan was lower in the ΔsrtA mutant (Fig. 3A).

The fact that most of these LPXTG proteins were not detected in the extracellular medium suggested that the lack of SrtA blocked their up-regulation during the transit of L. monocytogenes to stationary phase. Taken together, these data support a model in which the lack of SrtA affects the production of LPXTG proteins, at least those tested, when the cells reach stationary phase. This effect was especially evident for Lmo0434 (InlA), since during exponential phase it was detected at similar levels in wild-type and ΔsrtA strains (see Fig. 2A). Control experiments with the SigB-regulated LPXTG proteins Lmo0263 (InlH) and Lmo0880 demonstrated that their marked decrease in the ΔsrtA mutant grown to stationary phase was not due to an accumulation in the cytosol or the membrane (Fig. 3B). In addition, non-SrtA substrates, such as P60 and Lmo2185 (SvpA), were detected both in the cell wall and, especially, in the extracellular fractions (Fig. 3A). This finding excludes the possibility of a general degradation of LPXTG proteins in the extracellular medium by proteases secreted during stationary phase.

Overall, our findings provide support for a novel mechanism of regulation of LPXTG surface proteins, in which SrtA seems to contribute mostly when the bacteria reach stationary phase. This mechanism in essence acts as a negative form of regulation of LPXTG proteins when StrA, which anchors these substrate proteins to cell-wall peptidoglycan, is functionally altered. Future studies will need to address the exact regulatory elements involved and the mechanism(s) underlying their regulatory activities in the distinct phases of bacterial growth.

3.

4.

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

Acknowledgements. We thank Francisco García-del Portillo for critical reading of the manuscript, and Hélène Bierne and Pascale Cossart (Institut Pasteur, Paris, France) for the L. monocytogenes ΔsrtA strain. This study was funded by grant BIO2010-18962 of the Spanish Ministry of Economy and Competitiveness to M.G.P.

18.

19.

Competing interests. None declared.

20.

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Volume 15 · Number 1 · March 2012 · ISSN 1139-6709

Official journal of the Spanish Society for Microbiology Volume 15 · Number 1 · March 2012

International Microbiology

INTERNATIONAL MICROBIOLOGY

Volume 15 Number 1

RESEARCH ARTICLES

Heindl H, Thiel V, Wiese J, Imhoff JF Bacterial isolates from the bryozoan Membranipora membranacea: influence of culture media on isolation and antimicrobial activity

33

9

Mariscotti JF, Quereda JJ, Pucciarelli MG Contribution of sortase A to the regulation of Listeria monocytogenes LPXTG surface proteins

43

pp 1-54

Chen P, Yan L, Wang Q, Li Y, Li H Surface alteration of realgar (As4S4)

by Acidithiobacillus ferrooxidans

1

www.im.microbios.org

2012

Schinke C, Germani JC Screening Brazilian Macrophomina phaseolina isolates for alkaline lipases and other extracellular hydrolases

García-Maldonado JQ, Bebout BM, Celis LB, López-Cortés A Phylogenetic diversity of methyl-coenzyme M reductase (mcrA) gene and methanogenesis from trimethylamine in hypersaline environments

INTERNATIONAL MICROBIOLOGY

17

15(1) 2012 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/Elservier Bibliographic Databases; Food Science and Technology Abstracts; ICYT/CINDOC; IBECS/BNCS; ISI Alerting Services®; MEDLINE®/Index Medicus®; Latíndex; MedBioWorldTM; SciELO-Spain; Science Citation Index Expanded®/SciSearch®

Published Quarterly by • March 2012

Official journal of the Spanish Society for Microbiology