International Microbiology

Volume 16 路 Number 1 路 March 2013 路 ISSN 1139-6709 路 e-ISSN 1618-1905

INTERNATIONAL MICROBIOLOGY www.im.microbios.org

16(1) 2013

Official journal of the Spanish Society for Microbiology

INTERNATIONAL MICROBIOLOGY Publication Board

Editorial Board

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

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

Associate Editors Mercedes Berlanga, University of Barcelona Mercè Piqueras, Catalan Association for Science Communication Wendy Ran, International Microbiology Secretary General Jordi Mas-Castellà, International Microbiology Webmaster Nicole Skinner, Institute for Catalan Studies Jordi Urmeneta, University of Barcelona Managing Coordinator Carmen Chica, International Microbiology Members Josefa Antón, University of Alicante Susana Campoy, Autonomous University of Barcelona Ramón Díaz, CIB-CSIC, Madrid Josep Guarro, University Rovira Virgili Enrique Herrero, University of Lleida Emili Montesinos, University of Girona José R. Penadés, Institute of Mountain Livestock-CSIC, Castellon Jordi Vila, 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 (electronic version) Institute for Catalan Studies Carme, 47 08001 Barcelona, Spain Tel. +34-932701620. Fax +34-932701180 E-mail: int.microbiol@microbios.org © 2013 Spanish Society for Microbiology. Printed in Spain ISSN (Print): 1139-6709 e-ISSN (electronic): 1618-1095 D.L.: B.23341-2004 With the collaboration of the Institute for Catalan Studies

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CONTENTS International Microbiology (2013) 16:1-68 ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

Volume 16, Number 1, March 2013

RESEARCH REVIEW

Tortajada M, da Silva LF, Prieto MA Second-generation functionalized medium-chain-length polyhydroxyalkanoates: the gateway to high-value bioplastic applications

1

RESEARCH ARTICLES

Marsh SE, Poulsen M, Gorosito NB, Pinto-Tomás A, Masiulionis VE, Currie CR Association between Pseudonocardia symbionts and Atta leaf-cutting ants suggested by improved isolation methods

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López AC, Minnaard J, Pérez PF, Alippi AM In vitro interaction between Bacillus megaterium strains and Caco-2 cells

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Hashimoto W, Miyamoto Y, Yamamoto M, Yoneyama F, Murata K A novel bleb-dependent polysaccharide export system in nitrogen-fixing Azotobacter vinelandii subjected to low nitrogen gas levels

35

Wróbel B, Filippini M, Piwowarczyk J, Kędra M, Kuliński K, Middelboe M Low virus to prokaryote ratios in the cold: benthic viruses and prokaryotes in a subpolar marine ecosystem (Hornsund, Svalbard)

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Rajhi H, Conthe M, Puyol D, Díaz E, Sanz JL Dark fermentation: isolation and characterization of hydrogen-producing strains from sludges

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BOOK REVIEWS

63

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INTERNATIONAL MICROBIOLOGY 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. Its main objectives are to foster basic and applied microbiology, promote international relations, bring together the many professionals working in this science, and contribute to the dissemination of science in general and microbiology in particular, among society. It is an interdisciplinary society, with about 1800 members working in different fields of microbiology.

International Microbiology Aims and scope International Microbiology, the official journal of the SEM, is a peer-reviewed, 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, internationally-based information, short publication times (<3 months), complete copy-editing service, and online open access publication available 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.

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

The journal is covered in several leading abstracting and indexing databases, including the following ones: 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. Atta colombica, a leaf-cutter ant from Costa Rica. Atta spp. cut leaves that carry to their nests to support fungal gardens that feed the colony. They associate with various symbiotic microorganisms, including Actinobacteria, from which they obtain antibiotics that protect the ants against infection by potentially virulent microfungal parasites. This impressive photograph was taken by Jennifer Theobald [www.deepgreenphoto.com] during a field trip in 2011. (Magnification, ca. 0.1×) (© J. Theobald) Upper left. Particles of the turnip mosaic virus, a Potyvirus that infects mainly cruciferous plants and is one of the most damaging viruses in plant crops. It causes chlorotic ringspots in young leaves; as the leaf ages, yellow or brownish spots surrounbded by circular or irregular necrotic rings appear. Micrograph by Fernando Ponz, CBGP, UPM-INIA, Madrid. (Magnification, ca. 100,000×) Upper right. Transmission electron micrograph of a group of cells of the polyhydroxyalcanoate-producing Halomonas venusta MAT-28.

The cells are growing as a microcolony in an artificial biofilm of alginate beads. In this situation, cells maintain a metabolic state equivalent to that of the planktonic culture. Micrograph by M. Berlanga, Faculty of Pharmacy, University of Barcelona, and Carmen López, CCiT, University of Barcelona. [See article by Berlanga M., et al., Int Microbiol (2012) 15:191-199.] (Magnification, ca. 6,000×) Lower left. Scanning electron micrograph of Minorisa minuta, a bacterivorous protist described in 2012. With an average size of 1.4 mm, it is one of the smallest bacterial grazers known to date. It has a worldwide distribution and can account for 5 % of heterotrophic protists in coastal waters. Micrograph by Javier del Campo, Institute for Marine Sciences, CSIC, Barcelona, Spain. [For more information, see article by del Campo J, et al., ISME J (2013) 7:351-358] (Magnification, ca.

40,000×) Lower right. Scanning electron micrograph of Saksenaea vasiformis (Saksenaeaceae, Mucorales, Mucoromycotina) sporangiophores isolated from human tissue. This microorganism is a filamentous fungus with characteristic flask-shaped sporangia that can cause severe human infections in both immunocompromised and immunocompetent hosts. Micrograph by José F. Cano, University Rovira Virgili, ReusTarragona, Spain. (Magnification, ca. 1,000×)

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Back cover Portrait of Rafael Rangel (1877–1909), born in Betijoque, Venezuela. Rangel is considered to be the founder of Venezuelan parasitology and biological analytics. He was born on April 25, 1877, to a single mother, Maria Teresa Estrada, who died when he was sixmonths-old. His father, who had since married, claimed him, and his wife raised the child as if he was her own son. Rangel graduated from high school in Maracaibo and went to Caracas to study medicine. He was a brilliant student, but after his second year at university he abandoned his studies. Despite his failure to obtain an academic degree, he received good laboratory training and followed courses in bacteriology at the Pasteur Institute in Caracas.

Over time, he became an expert in the use of the microtome and microscope, the staining of microorganisms, the preparation of culture media, and the inoculation of pathogens into laboratory animals. In 1902, he was appointed director of the new Laboratory of Histology and Bacteriology at the Vargas Hospital in Caracas.

Even without a medical degree, he assisted students in their doctoral theses and carried out research in various fields. In 1908, when he was at the peak of his career, several cases of bubonic plague were reported in La Guaira, Venezuela. Rangel was commissioned to deal with this devastating disease. His first quick tests consistently yielded negative results. However, as more cases were reported, Rangel renewed his efforts, initiating a campaign to fight the plague that involved strong measures, including temporarily closing the harbor and ordering the burning of four buildings in which rats, the vectors of the pathogen, were known to dwell. The epidemic was soon controlled and Rangel was rewarded for his work. But there were also people envious of his success who blamed him for not having recognized the disease until it had widely spread, and for having spent large sums of money to fight it. These accusations, as well as the denial of a fellowship that would have allowed him to travel to Europe to complete his medical training, caused Rangel to sink into a deep depression. Sadly, he killed himself, by drinking mercury cyanide, on August 20, 1909. On the centenary of his birth, in 1977, Venezuela paid homage to Rangel and moved his remains to the National Pantheon. In 1999, he vas posthumously invested Doctor Honoris Causa by the Fermín Toro University at Barquisimeto, Venezuela.

Front cover and back cover design by MBerlanda & RGuerrero

RESEARCH REVIEW International Microbiology (2013) 16:1-15 doi:10.2436/20.1501.01.175 ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

Second-generation functionalized mediumchain-length polyhydroxyalkanoates: the gateway to high-value bioplastic applications Marta Tortajada,1 Luiziana Ferreira da Silva,2 María Auxiliadora Prieto3* 1 Microbial Biotechnology Department, BIOPOLIS SL, Valencia, Spain. 2Laboratory of Bioproducts, Institute of Medical Sciences, University of São Paulo, São Paulo, Brazil. 3Polymer Biotechnology Group, Biological Research Center (CIB), CSIC, Madrid, Spain

Received 7 February 2013 · Accepted 7 March 2013

Summary. Polyhydroxyalkanoates (PHAs) are biodegradable biocompatible polyesters, which accumulate as granules

in the cytoplasm of many bacteria under unbalanced growth conditions. Medium-chain-length PHAs (mcl-PHAs), characterized by C6-C14 branched monomer chains and typically produced by Pseudomonas species, are promising thermoelastomers, as they can be further modified by introducing functional groups in the side chains. Functionalized PHAs are obtained either by feeding structurally related substrates processed through the β-oxidation pathway, or using specific strains able to transform sugars or glycerol into unsaturated PHA by de novo fatty-acid biosynthesis. Functionalized mclPHAs provide modified mechanical and thermal properties, and consequently have new processing requirements and highly diverse potential applications in emergent fields such as biomedicine. However, process development and sample availability are limited due to the toxicity of some precursors and still low productivity, which hinder investigation. Conversely, improved mutant strains designed through systems biology approaches and cofeeding with low-cost substrates may contribute to the widespread application of these biopolymers. This review focuses on recent developments in the production of functionalized mcl-PHAs, placing particular emphasis on strain and bioprocess design for cost-effective production. [Int Microbiol 2013; 16(1):1-15] Keywords: Pseudomonas spp. · polyhydroxyalkanoates (PHAs) · medium-chain-length (mcl)-PHAs · functionalization

of polymers · metabolism · low-cost substrates

The relevance of functionalized mediumchain-length-PHAs From an industrial standpoint, polyhydroxyalkanoates (PHAs) are biopolyesters attracting extensive interest as *Corresponding author: M.A. Prieto Environmental Biology Department Biological Research Center (CIB)-CSIC Ramiro de Maeztu, 9 28040 Madrid, Spain Tel. +34-918373112. Fax +34-915360432 Email: auxi@cib.csic.es

technical-grade polymers due to their singular set of properties: (i) substitution potential for industrial thermoplastics such as polypropylene, polyethylene, polyvinylchloride and polyethylene terephthalate, (ii) biodegradability both in aerobic and anaerobic conditions, including aquatic environments, (iii) bio-based, renewable origin, (iv) biocompatibility with cells and tissues and (vi) structural diversity [4]. This last characteristic is critical to define potential applications, since the specific chemical monomer composition and molecular structure will determine the biological, thermal and mechanical properties of the resulting polymer.

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Fig. 1. Schematic representation of the chemical structure of PHAs. (A) PHAs are generally composed of 3-hydroxy fatty acids, where the pendant group (shown as R) varies from methyl (C1) to undecyl (C11). C3 carbon atom is asymmetric leading to pure R enantiomers as monomers. Thus, PHA monomers can be useful synthons in the pharmaceutical industry [9]. The best-known PHAs are PHB (R = methyl), P(HB-co-HV) (R = methyl or ethyl), and P(HO-co-HH) (R = pentyl or propyl). Second generation mcl-PHAs carry functionalized substitutions at pedant group.

(B) Schematic representation of the chemical structure of the family of PHACOS, the acetylthioester-functionalized mcl-PHA [18]. OH-Alk 3-hydroxyalkanoate, OH-6ATH 3-hydroxy-6-acetylthiohexanoate, OH4ATB 3-hydroxy-4-acetylthiobutanoate.

In this respect, PHAs have been classified attending to various criteria (Fig. 1): (i) their monomer size as shortchain-length PHAs (scl-PHAs), with C4-C5 monomers and medium-chain-length PHAs (mcl-PHAs) with C6-C14 monomers, (ii) their functional substituents found in the radical chain (such as double bonds or aromatic groups), and (iii) the structure of the polymer (formed by homogenous, random or block copolymers [46]). Although other added-value PHAs have been described, including in vivo protein-anchored, bioactive PHAs for biomedical applications [22,,49], this review will focus on functionalized mcl-PHAs as promising and versatile candidates for high added-value applications.

Short-chain-length-PHAs (scl-) such as poly-3-hydroxybutyrate (PHB) and its copolymers with poly-3-hydroxy-

valerate (PHB-co-HV) are being produced on a commercial scale and have extensive application in packaging, moulding, fibres and other commodities [4]. Currently, mcl-PHAs are considered to be promising candidates for special bioplastic applications due to properties derived from their longer side-chains and altered crystalline structure, such as elasticity, hydrophobicity, low oxygen permeability, water resistance, and biodegradability. They can be moulded and processed into compostable packaging and resorbable materials for medical applications, and have also been used as food coatings, pressure-sensitive adhesives, paint binders and biodegradable rubbers [73].

Furthermore, unconventional mcl-PHAs bearing different functional moieties in their side chains can be produced through different biotechnological strategies, which will be reviewed in detail in the following sections of this article.

These reactive groups enable tuning of the physical and chemical properties of the polymer, and they are also potential targets for post-biosynthetic modifications (Fig. 2).

For example, the higher the molar fraction of unsaturated constituents in the monomers, the lower the resulting melting and glass transition temperatures due to crystallization inhibition by unsaturated side chains. Double bonds are also easily attacked in chemical reactions, allowing the polymer properties to be diversified even more [25]. A number of treatments have been described as responsible for crosslinking of unsaturated PHAs, namely electronbeam irradiation, UV-irradiation or even autoxidation and in some cases these PHAs are transformed into rubbers [3].

Chemical epoxidation of the pendant vinyl groups has also been applied to decrease melting temperature and increase glass transition temperature [48].

Thus, controlling monomer composition provides the chance to establish functionalized PHAs as tailor-made polymers for different applications. However, this scenario can be only achieved once we have gained a better understanding of how the incorporation of different monomers into PHA is controlled and how material properties are influenced by PHA composition.

Biochemical pathways for mediumchain-length-PHA synthesis Chemically, PHAs are branched hetero-oxo-polyesters composed of (R)-3-hydroxy-alkanoic acid monomers (Fig. 1). Polyhydroxyalkanoates are synthesized as storage polymers by some Archaea and by a wide range of gram-

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Medium-chain-length PHAs

Fig. 2. Diagram of the key challenges for the production of added-value bacterial polyesters.

positive and gram-negative bacterial species, both in aerobic and anaerobic environments, although only a few strains produce PHAs at industrially significant rates.

These biopolymers are accumulated as hydrophobic inclusions (PHA granules) (Fig. 3) in the bacterial cytoplasm, generally as a response to unbalanced growth conditions, such as inorganic nutrient limitations in the presence of excess carbon source [56].

A large array of bacterial species can produce PHB; however, mcl-PHAs are mainly, though not exclusively, produced by fluorescent Pseudomonads such as Pseudomonas putida [54] (Fig. 3). This division arises on account of differences in the in vivo substrate specificity of the PHA polymerase or synthase, the enzyme responsible for the assembly of PHA monomeric precursors, (R)-3-hydroxyacyl-CoAs, together with the evolved specialization of peripheral metabolic pathways and regulatory networks

in each species. To date, the most widely studied and reference mcl-PHA producers are P. putida KT2440 (and its rifampicin-resistant mutant, KT24) and P. oleovorans GPo1 (ATCC 29347), reclassified and referred to herein as P. putida GPo1 [50,70].

In PHB producers, such as the paradigmatic Ralstonia eutropha H16 strain, the main enzymes involved in PHB synthesis are encoded in a gene cluster expressing: (i) a 3-ketothiolase, which condenses two acetyl-CoA molecules into acetoacetyl-CoA, (ii) a NADPH-dependent acetoacetyl-CoA reductase, which stereo-selectively reduces acetoacetyl-CoA to (R)-3-hydroxybutyryl-CoA, and (iii) a PHB synthase that finally converts (R)-3-hydroxybutyrylCoA into PHB releasing free CoA [56]. Depending on the capacity of each strain to metabolize either simple or complex sugars, fats or oils into acetyl-CoA, PHB will be produced out of all, or only some of these substrates. Con-

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versely, Pseudomonads have a much more complex metabolism (Fig. 4). The pha cluster is formed in mcl-PHA producers by the genes encoding two synthases (PhaC1 and PhaC2), a depolymerase (PhaZ) [11] and other PHArelated proteins, including phasins, involved in formation, maintenance and segregation of PHA granules (PhaF and PhaI) [19]. The substrates for PHA synthesis, (R)-3-hydroxyacyl-CoAs, are supplied by two central pathways, β-oxidation pathway and de novo fatty acid synthesis, fed by fatty acids or other non-PHA-related substrates such as carbohydrate intermediates, respectively. In addition, these central pathways are supplemented by strain-dependent peripheral routes that transform non-conventional precursors into PHA. Structurally related substrates, such as fatty acids, are processed by Pseudomonads through β-oxidation cycle [39]. The resulting acyl-CoAs are sequentially oxidized into enoyl-CoA, (S)-3-hydroxyacyl-CoA and (R)-3-ketoacyl-CoA. All of these intermediates may then be converted into (R)-3-hydroxyacyl-CoA by a stereospecific transenoyl-CoA hydratase (PhaJ), an epimerase, or a specific (R)-ketoacyl-CoA reductase (FabG), respectively [61].

Pseudomonas putida KT2440, in particular, has a large array of β-oxidation enzymes [43]. Concerning the fatty acids β-oxidation protein complex (FadAB), two sets of fadAB genes have been described in the strain P. putida KT24 [5,47]: fadB and fadA (PP_2136 and PP_2137), and fadBx and fadAx (PP_2214 and PP_2215). The former set would appear to play a role in fatty acid degradation, since although fadB and fadA deletion mutants do not show completely blocked β-oxidation, they produce PHA with a higher content of longer chain monomers, possibly due to their defective β-oxidation pathway [5,18,47].

Non-fatty acid precursors can be oxidized to acetylCoA and channelled into PHA by the de novo synthesis pathway, via (R)-3-hydroxyacyl-acyl carrier protein (ACP) intermediates. In this process, malonyl-CoA and its precursor, acetyl-CoA, are activated by transacylation to acylcarrier protein (ACP). Malonyl and acyl-ACP derivatives

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Fig. 3. (A) Transmission electronic microscopy (TEM) image of mclPHA-producing cells of Pseudomonas putida KT2442. (B) TEM image of a mcl-PHA granule isolated from P. putida KT2442. Granules are composed of mcl-PHA (shown as whitest core of the granule) coated by a monolayer of phospholipids and granule-associated proteins (gray layer at the surface of the granule). The most abundant proteins in the surface granules are phasins. Polymerases and intracelluar depolymerases are also associated to granules. (C) mcl-PHA film isolated from P. putida KT2440 and octanoic acid as growth and mclPHA precursor.

Medium-chain-length PHAs

are condensed by ketoacyl-ACP synthetase, reduced, losing a ketone group, dehydrated, and saturated to the corresponding (R)-3-hydroxyacyl-ACP chain (Fig. 4), which may be further elongated in two-carbon growing chains.

Acyl-ACP intermediates can then be re-transformed into (R)-3-hydroxyacyl-CoAs by a specific transacylase, PhaG, present in most Pseudomonas. Only P. putida GPo1 and P. fragi are unable to synthesize mcl-PHA out of non-fatty acid substrates, such as gluconate, apparently due to deficiencies in PhaG transcription [29,55]. Unsaturated monomers such as 3-hydroxy-5-dodecenoate and 3-hydroxy7-tetradecenoate are also generated in this process [30], by fatty acid de novo synthesis that introduces double bonds into acyl-CoA or acyl-ACP intermediates, a regulatory mechanism of membrane fatty acid composition that alters membrane fluidity in response to changes in ambient temperature [14]. Tables 1, 2 and 3 revise the precursors described in the literature to produced mcl-PHA with substitutions at the side chain.

Functionalized PHAs through β-oxidation pathway Structurally related carbon sources, such as alkanoic and alkenoic acids are incorporated to PHA by mcl-PHA producers through the β-oxidation pathway without being completely oxidized to acetyl-CoA [54]. The resulting PHAs are generally copolymers, a consequence of the sequential degradation of acetyl-CoA units (e.g., poly[3-hydroxy-octanoic-co-3-hydroxy-hexanoic]), at around 95 mol% of 3-hydroxy-octanoic acid and 5 mol% of 3-hydroxy-hexanoic is produced when P. putida is fed with octanoic acid).

Linear and branched n-alkanes and alkenes can also be channelled to polymer accumulation, when processed by the alkane oxidation pathway encoded on the octane (OCT) plasmid [70]. Fatty acids however are usually preferred for bioprocessing as they overcome the limitations derived from two-phase fermentation systems required for hydrocarbons and avoid risk of explosion [67].

The structure of the precursor, including unsaturations, is recovered in the resulting mcl-PHA. Early reports by Lageveen and co-workers [34] describe the generation of PHA containing 3-hydroxy-5-hexenoate, 3-hydroxy-7-octenoate, 3-hydroxy-8-nonenoate and 3-hydroxy-9-decenoate when octene, nonene or decene were supplied. Nonterminal unsaturations were also successfully introduced.

The molar fraction of unsaturated monomers depends on

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both the carbon source supplied and the metabolic capabilities of the bacterial strain. For example, whereas P. putida GPo1 fed with octene produces PHA with 50 mol% 3-hydroxyalkenoate content [53], when 10-undecenoate is used, practically all monomers are unsaturated.

The molar content of unsaturated monomers can then be controlled by co-feeding nonanoic acid, as it correlates with the amount of 10-undecenoate. When P. putida was fed with mixtures of octanoic acid and undecenoic acid, the monomeric composition seemed to depend on the growth rate both in batch and chemostat cultures, but also correlated linearly with the fatty acids fed [24].

This strategy can be extended to a large variety of organic compounds in order to generate functionalized PHAs [62]. Depending on their chemical structure and the specific bacterial species, the compounds will support both cell growth and PHA accumulation, growth alone, or will support neither cell growth nor PHA accumulation [31].

Simultaneous feeding strategies based on co-metabolism can be applied to address the last two cases, as it has been shown that precursors that do not enable PHA production or growth can be processed when accompanied by an efficiently processed substrate, such as octanoic or undecenoic acid [24]. The first examples of these strategies include copolyesters obtained from mixtures of octanoic acid and methylalkanoic acids such as 7-methyloctanoate, which contained both the unsubstituted and the methyl-branched 3-hydroxyoctanoate and 3-hydroxyhexanoate units. Although no polymer was formed when P. putida GPo1 was grown on pure 5- and 6-methyloctanoates, the presence of methyl-branched units in the polymer obtained by co-metabolism was detected when 5- and 6-methyloctanoate were fed as mixture with octanoate [37]. There was a direct correlation between the variation in ratios of PHA repeating units and feeding mixture composition.

This concept has been exploited to produce a plethora of tailor-designed mcl-PHAs, with highly diverse structures that include acetylthioester, acetoxy, alkoxy, amino, cyano, cyclohexyl, epoxy, halogenated, hydroxy or propylthiol groups (Tables 1 and 2). Note that besides the precise functional moeity introduced, a larger variety of PHA compositions can be generated in most cases, with varying molar amounts of functional groups, by altering the ratio of co-substrates.

Together with unsaturations, functional groups prone to chemical modifications have been introduced, mainly in the canonical P. putida GPo1 and KT24 strains, such as thio, bromine, chlorine and fluorine radicals, cyano and

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Table 1. Precursors used in the literature to produce functionalized mcl-PHA (branched alkyl, cyclohexyl, halogenated) % Mol functional groups

% PHA

Pseudomonas strains

Ref.

Citronellol

>99.0

27.2

P. citronellolis ATCC 13674

[1-T1]

Alkylhydroxyoctanoates

5.0

16.0–22.0

P. putida GPo1

[2-T1]

Methyloctanoates

3.0–96.4

7.3–19.2

P. putida GPo1

[3-T1, 4-T1]

Cyclohexylbutyric acid

>99.0

21.0

P. cichorii YN2

[5-T1]

Cyclohexylvaleric/butytic acid

13.2–100.0

2.0–31.0

P. putida GPo1

[6-T1, 7-T1]

Alkenes (C7-C9)

45.0–55.0

5.0–13.4

P. putida GPo1

[8-T1]

Undecenoic acid

27.1–100.0

1.8–37.4

P. putida KCTC 2407

[9-T1]

Undecenoic acid

5.0–99.0

17.0–34.0

P. putida GPo1

[10-T1,11-T1, 12-T1]

Hydroxyoctenoic acids

63.5-81.6

10.3-12.5

P. putida GPo1

[13-T1]

Dicarboxylic acids (C4-C10)

4.7-11.7

7.3-14.4

P. citronellolis ATCC 13674

[14-T1]

Undecynoic acid

32.0-100.0

N/A

P. putida GPo1

[15-T1]

Undecynoic acid

22.0-100.0

7.5-22.5

P. putida KCTC 2407

[15-T1]

Bromoalkanoic acids (C6-C11)

3.7-38.0

4.5-38.2

P. putida GPo1

[16-T1, 17-T1]

Chlorooctane

69.0

5.0-19.0

P. putida GPo1

[18-T1]

Fluorohexanoic/nonanoic acids

1.9-8.8

N/A

P. putida GPo1

[19-T1]

Fluorohexanoic/nonanoic acids

1.0-17.3

N/A

P. putida KT2440

[19-T1]

Fluorophenoxyundecanoic acid

>99.0

8.5-13.9

P. putida 27N01

[20-T1]

Precursor

Group at mcl-PHA side chain: Branched alkyl

Group at mcl-PHA side chain: Cyclohexyl

Unsaturated

Group at mcl-PHA side chain: Halogens

[1-T1] Choi MH, Yoon SC (1994) Appl Environ Microbiol 60:3245-3254 · [2-T1] Scholz C, et al. (1994) Macromolecules 27:6358-6362 · [3-T1] Fritzsche K, et al. (1990) Int J Biol Macromol 12:92-101 · [4-T1] Lenz RW, et al. (1992) FEMS Microbiol Lett 103:207-214 · [5-T1] Honma T, et al. (2004) J Environ Biotechnol 4:49-55 · [6-T1] Andújar M, et al. (1997) Macromolecules 30:1611-1615 · [7-T1] Kim DY, et al. (2001) Int J Biol Macromol 29:145-150 · [8-T1] Lageveen RG, et al. (1988) App Environ Microbiol 54:2924-2932 · [9-T1] Kim DY, et al. (2000) Int J Biol Macromol 28:23-29 · [10-T1] Kim YB, et al. (1995) J Polym Sci Part A: Polym Chem 33:1367-1374 · [11-T1] Park WH, et al. (1998) Macromolecules 31:1480-1486 · [12-T1] Sparks J, Scholz C (2008) Biomacromolecules 9:2091-2096 · [13-T1] Fritzsche K, et al. (1990) Int J Biol Macromol 12:85-91 · [14-T1] Choi MH, Yoon SC (1994) Appl Environ Microbiol 60:3245-3254 · [15-T1] Kim DY, et al. (1998) Macromolecules 31:4760-4763 · [16-T1] Kim YB, et al. (1992) Macromolecules 25:1852-1857 · [17-T1] Lenz RW, et al. (1992) FEMS Microbiol Lett 103:207-214 · [18-T1] Doi Y, Abe C (1990) Macromolecules 23:3705-3707 · [19-T1] Kim O, et al. (1996) Macromolecules 29:4572-4581 · [20-T1] Takagi Y, et al. (2004) Eur Polym J 40:1551-1557

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7

Table 2. Precursors used in the literature to produce functionalized mcl-PHA (acetoxy, ester, alkoxy, epoxy, thio, cyano, nitro) Precursor

Pseudomonas strains

% Mol functional groups

% PHA

Ref.

3.3–10.3

19.0–26.0

P. putida GPo1

[1-T2]

Alkylheptanoate

2.5–60.0

0.27–6.9

P. putida GPo1

[2-T2]

Alkxyhexanoic/octanoic/undecanoic acids

31–100.0

3.0–15.0

P. putida GPo1

[3-T2]

10-epoxyundecanoic acid

25.0–75.0

3.0–18.0

P. putida GPo1

[4-T2]

C7-C12 alkenes

4.2–20.0

25.0–35.0

P. cichorii YN2

[5-T2]

63.0

63.0

P. stutzeri 1317

[6-T2]

Acetylthiohexanoic acid

16.5–78.5

5.0–37.0

P. putida KT2442, KT24FadB

[7-T2]

Propylthiohexanoic acid

14.5–17.5

24.0–43.0

Ralstonia eutropha DSM541

[8-T2]

6.02% w/w S

N/A

P. putida KT2440

[8-T2]

Methylsulfanylphenoxyvaleric acid

12.2–35.6

10.9–21.4

P. cichorii H45, YN2

[9-T2]

Methylsulfanylphenoxyvaleric acid

18.4

9.9

P. jessenii P161

[9-T2]

Thiophenoxyundecanoic acid

>99.0

9.6–19.5

P. putida 27N01

[10-T2]

17.0–32.0

19.5–36.2

P. putida GPo1

[11-T2]

Cyanophenoxyhexanoic acid

0-2.2

N/A

P. putida GPo1

[13-T2]

Cyanophenoxyhexanoic acid

0.0–34.0

2.8–12.0

P. putida KT2440

[12-T2, 13-T2]

Nitrophenoxyhexanoic acid

4.2–5.1

N/A

P. putida GPo1

[13-T2]

Nitrophenoxyhexanoic acid

1.0–4.8

N/A

P. putida KT2440

[13-T2]

Dinitrophenylvaleric acid

1.2–6.9

15.0–40.0

P. putida GPo1

[14-T2]

Group at mcl-PHA side chain: Acetoxy Octanone , octylacetate Ester, alkoxy, epoxy

Soybean oil Group at mcl-PHA side chain: Thio, sulfanyl

Propylthioundecanoic acid

Group at mcl-PHA side chain: Cyano, nitro Cyanoundecanoic acid

[1-T2] Jung K, et al. (2000) Macromolecules 33:8571-8575 · [2-T2] Scholz C, et al. (1994) Macromolecules 27:2886-2889 · [3-T2] Kim DY, et al. (2003) J Microbiol Biotechnol 13:632-635 · [4-T2] Bear MM, et al. (1997) React Funct Polym 34:65-77 · [5-T2] Imamura T, et al. (2001) Int J Biol Macromol 29:295-301 · [6-T2] He W, et al. (1998) FEMS Microbiol Lett 169:45-49 · [7-T2] Escapa IF, et al. (2011) App Microbiol Biotechnol 89:1583-1598 · [8-T2] Ewering C, et al. (2002) Microbiology 148:1397-1406 · [9-T2] Kenmoku T, et al. (2002) EP1275727 B1. Priority date: 10-07-2002 · [10-T2] Takagi Y, et al. (1999) Macromolecules 32:8315-8318 · [11-T2] Lenz RW, et al. (1992) FEMS Microbiol Lett 103:207-214 · [12-T2] Gross RA, Kim O-Y, et al. (1996) Polym Int 39:205-213 · [13-T2] Kim O, et al. (1995) Can J Microbiol 41:32-43 · [14-T2] Aróstegui SM, et al. (1999) Macromolecules 32:2889-2895

epoxy groups (Table 2). The introduction of such groups also modifies the thermal properties, and thus the process-

ing requirements of the resulting polymers, enabling higher melting and lower glass-transition temperatures, or modi-

8

Int. Microbiol. Vol. 16, 2013

fied biological activities. For example, different PHA compositions bearing acetylthioester groups in the side chain have been produced using P. putida KT24. These novel mcl-PHAs (PHACOS) can be obtained using decanoic acid as an inducer of growth and PHA synthesis, and 6-acetylthiohexanoic acid as PHA precursor, in one or twostage cultivation strategies. The composition of PHACOS includes 6-acetylthio-3-hydroxyhexanoic acid (OH-6ATH) and the shorter derivative 4-acetylthio-3-hydroxybutanoic acid (Fig. 1). These polymers have tunable thermal properties as a result of different glass transition temperatures.

Use of the derived strain KTFadB, mutated in the fadB gene from the β-oxidation pathway, gives rise to PHACOS overproduction and the polymer contains mainly OH6ATH units [18] [Escapa IF, Morales V, García JL, Prieto MA (2012) Synthesis of polyhydroxyalkanoates (PHAS) with thioester groups in the side chain. International Patent WO 2012/038572].

Many reports have been devoted to the accumulation of aromatic radicals in P. putida strains (Table 3), in order to mimic widespread synthetic aromatic polyesters such as polyethyleneterephtalate (PET). Processing of aromatic substrates into PHA seems to depend both on the chemical structure of the precursors and the bacterial strain.

Homopolymers with 100 mol% content of aromatic moieties, random copolymers or a blend of both have been produced. Bacterial isolates that are able to transform aromatic precursor into alkyl-PHAs have also been reported [72].

Hydrophilized PHAs bearing alkoxy, acetoxy or hydroxy groups are also of great interest, as they show enhanced solubility and biocompatibility [26].

Although diverse fermentation strategies based on cometabolism have been extensively applied to produce functionalized mcl-PHAs, few works have been published connecting these findings to the molecular basis driving the carbon metabolism in Pseudomonas [10,20]. This is partially caused by the complex regulation of mcl-PHA metabolism, which is controlled: (i) at the enzymatic level, by cofactor inhibition and metabolite availability [17], (ii) at the transcriptional level, by specific and global transcriptional regulatory factors [9,10], and (iii) at the translational level, driven by global post-transcriptional regulators.

Recently, it has been shown that synthesis and degradation of mcl-PHA consist of a synchronous cycle in P. putida KT2440, as PHA synthase and depolymerase are simultaneously produced ensuring the PHA turnover. In P. putida KT2440, simultaneous production of PhaC (C1 and C2)

tortajada et al.

and PhaZ enzymes is controlled by the protein PhaD [10], one of the few activators of the TetR-like family of transcriptional regulators. The phaD gene controls its own transcription and that of phaIF operon coding for phasins.

Mutagenesis analyses and 3D structural models suggest that PhaD behaves as a carbon-source-dependent activator of the pha cluster, possibly induced by CoA intermediates of β-oxidation. The pha cluster would only be indirectly induced by fatty acids, such as octanoic acid, the true inducer being CoA derivatives of β-oxidation. This hypothesis supports co-metabolism effects and also the lower activation of PHA production on non-structurally related carbon sources, such as glucose, for which PHA cycle is only driven on basal activities of the pha promoters. However, differences in activation mechanisms are expected among the different mcl-PHA-producing strains because phylogenetically related strains, such as Pseudomonas corrugata and Pseudomonas mediterranea, bear vast differences in nucleotide sequences in the intergenic regions of pha cluster [66]. In addition to the intrinsic interest in PHAs due to their wide ranging biological, mechanical and thermal properties, much effort is still required to unravel the molecular and biochemical underpinnings of tuned PHA production.

De novo synthesis for functionalized PHAs Although most mcl-PHA intermediates are obtained through fatty-acid β-oxidation, non-related carbon sources such as acetate, ethanol, fructose, glucose, gluconate or glycerol are also channelled to PHA by the de novo fatty acid pathway in Pseudomonas species such as P. putida or P. aeruginosa [68]. The main monomer found in these mclPHAs is 3-hydroxydecanoate, but unsaturated 3-hydroxy5-dodecenoate and 3-hydroxy-7-tetradecenoate are also found [30]. The molar fraction of unsaturated monomers usually ranges from around 5 to 10 mol% depending on the strain [60,64], and it can be increased by reducing culture temperature [30]. This is consistent with the isolation of a psychrotrophic P. fluorescens strain able to accumulate mcl-PHA with up to 35 mol% of 3-hydroxy-5-dodecenoate content [36].

The use of carbohydrate-related sources to partially unsaturated PHA side chains is advantageous in terms of substrate cost and invariant monomer composition although PHA yields are generally lower in comparison to fatty acid

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9

Table 3. Precursors used in the literature to produce functionalized mcl-PHA (aromatics) Precursor

% Mol functional groups

% PHA

Pseudomonas strains

Ref.

Group at mcl-PHA side chain: Aromatics (benzoyl, methylphenoxy, phenoxy, phenyl) Benzoylalkanoic acids (C4–C8)

8.3–79.8

3.0–41.0

P. cichorii YN2

[1-T3]

Methylphenoxyalkanoic acids (C6, C8)

40.0–65.0

0.8–13.0

P. putida KCTC 2407

[2-T3]

Methylphenoxyalkanoic acids (C6, C8) (PVA, NA)

68.0–100.0

N/A

P. putida GPo1

[3-T3]

Methylphenoxyalkanoic acids (C6, C8)

24.0–100.0

0.8–23.7

P. putida KCTC 2407

[3-T3]

Phenoxyundecanoic acid

>99.0

>19.0

P. putida GPo1

[4-T3]

Phenoxyalkanoic acids (C6,C8,C11)

100.0

10.0–11.0

P. putida GPo1

[5-T3]

Phenoxyundecanoic acid

12.0–100.0

13.1–46.8

P. putida BM01

[6-T3]

Phenylvaleric acid

13.7–19.5

18.0–56.0

P. putida BM01

[7-T3]

Phenylvaleric acid

>99.0

N/A

P. putida GPo1

[8-T3]

3.0–64.0

0.03–23.0

P. putida GPo1

[9-T3]

Phenylalkanoic acids (C4-C8)

>95.0

22.0–42.0

P. jessenii C8

[10-T3]

Phenylalkanoic acids (C4-C8)

>95.0

8.0–36.0

P. putida S12, CA-1, H4, F6, D5

[10-T3]

Phenylalkanoic acids (C6-C11)

>99.0

10.0–87.0

P. putida U fadA-, ΔFadBA-PhaZ

Phenylvaleric acid

>99.0

25.0

P. putida GPo1

[14-T3]

Phenylvaleric acid

12.6–40.6

15.1–39.2

P. putida GPo1

[15-T3]

Phenyl, tolylvaleric/octanoic acids

[11-T3, 12T3, 13-T3]

[1-T3] Honma T, et al. (2004) J Environ Biotechnol 4:49-55 · [2-T3] Kim DY, et al. (2000) Int J Biol Macromol 28:23-29 · [3-T3] Kim YB, et al. (1999) Macromolecules 32:6058-6064 · [4-T3] Ritter H, Spee von AG (1994) Macromol Chem Phys 195:1665-1672 · [5-T3] Kim YB, et al. (1996) Macromolecules 29:3432-3435 · [6-T3] Song JJ, Yoon SC (1996) App Environ Microbiol 62:536-544 · [7-T3] Song JJ, et al. (2001) J Microbiol Biotechnol 11:435-442 · [8-T3] Curley JM, et al. (1996) Int J Biol Macromol 19:29-34 · [9-T3] Curley JM, et al. (1996) Macromolecules 29:1762-1766 · [10-T3] Tobin KM, O’Connor KE (2005) FEMS Microbiol Lett 253:111-118 · [11-T3] Abraham GA, Get al. (2001) Biomacromolecules 2:562-567 · [12-T3] García B, et al. (1999) J Biol Chem 274:29228-292241 · [13-T3] Olivera ER, et al. (2001) Mol Microbiol 39:863-874 · [14-T3] Fritzsche K, et al. (1990) Macromol Chem Phys 191:1957-1965 · [15-T3] Kim YB, et al. (1991) Macromolecules 24:5256-5260

substrates. Nevertheless, the wide metabolic versatility of Pseudomonads, which do not prefer glucose over alternative carbon sources, can be exploited to generate functionalized PHAs out of a variety of substrates. Pseudomonads lack the main glycolytic enzyme, phosphofructokinase, but instead transform glucose into 6-phosphogluconate (6PG) through three convergent pathways. Then 6PG is metabolized by the Entner-Doudoroff (ED) enzymes into pyruvate

to finally yield acetyl-CoA, which may be channelled into citric acid cycle or PHA cycle through de novo fatty acid synthesis (Fig. 4) [50].

The expression of the main metabolic steps of the phosphorylative branch of carbohydrate metabolism and ED pathways in P. putida KT2440 is tightly regulated by transcriptional repressors such as HexR, specifically induced by 2-keto-3-deoxy-6-P-gluconate (KDPG). Modified tran-

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tortajada et al.

Int Microbiol

10

Fig. 4. (A) Metabolic pathways involved in PHA biosynthesis of Pseudomonads. In Pseudomonas putida synthesis and degradation of PHA were found to operate as a continuous cycle with 3-hydroxy-fatty acids released from PHA granules by PhaZ depolymerase and activated to 3-hydroxyacylCoAs by ACS1 acyl synthetase with the concomitant consumption of one ATP molecule. These activated monomers will be either metabolized via fatty acid degradation or re-incorporated into PHA by PHA synthase. The specific PHA metabolic pathways are interconnected with the main central pathways that converge in acetyl-CoA. (B) Genetic organization of pha cluster in P. putida. The open arrows indicate the directions of gene transcription. Phasins PhaI and PhaF encoding genes are shown in black. Enzymes-encoding genes are shown in blue; phaC1 and phaC2 genes (light blue) code for two synthases and are separated by the phaZ gene (dark blue) coding for an intracellular depolymerase. phaD gene (yellow) encodes a transcriptional regulator.

scription of HexR has been shown to improve the efficiency of the use of alternative carbon sources, such as xylose, in engineered P. putida S12 strain, balancing ED and pentose phosphate pathway activities [41].

Besides sugars, glycerol has also been considered to be an advantageous raw material for PHA production due to its availability as a by-product of the biodiesel industry [21]. In Pseudomonas strains, glycerol is converted into glycerol3-phosphate and then to dihydroxyacetone phosphate, which is catabolized by a branch of the ED pathway [16,63]. Glyc-

erol use is hindered by a prolonged lag phase, caused by the transcriptional repressor GlpR. However, GlpR repression can be overcome by adding small amounts of fatty acids as cofeeders, fully eliminating lag-phase in P. putida KT2440.

Knock-out GlpR mutants also result in faster consumption of glycerol, together with improved PHA accumulation, possibly as a consequence of the larger availability of intermediates generated by de novo fatty acid synthesis [16].

Hence, to improve the utilization of non-structurally related carbon sources in bacterial PHA production re-

Medium-chain-length PHAs

quires knowledge of the metabolic steps involved in their catabolism. It is also necessary to study their relationship with the central metabolic pathways, possibly through the cellular levels of key metabolic intermediates that may be directed to growth, energy generation or PHA synthesis.

In-depth knowledge of the PHA cycle is therefore essential to coordinate global metabolism with resource availability in PHA producing microorganisms [9,17].

Functionalized PHAs from non-food raw materials In addition to high-added value applications for functionalized PHAs, cost-efficient exploitation of these polymers can be approached through the use of different agro-industrial by-products [33]. Raw materials that do not compete or interfere with food supply are preferred in line with global sustainability strategies and biorefinery concepts.

The ability of different bacteria to use plant oils or animal fats to produce PHA has been widely described [7,23,27,40,65]. The same concept of controlling PHA composition in functionalized PHAs by feeding mixtures of carbon sources can be readily translated to unsaturated plant or animal derived fatty acids.

Accordingly, by using oleic or linoleic acid, which are constituents of a large number of plant oils, the monomers 3-hydroxy-6-dodecenoate, 3-hydroxy-5-tetradecenoate and 3-hydroxy-5,8-tetradecenoate have been incorporated into the PHA produced by P. putida GPo1 [12]. Also 10-undecenoic acid can be derived from inexpensive castor oil, and used to produce PHA containing 3-hydroxy-10-undecenoate, 3-hydroxy-8-nonenoate and 3-hydroxy-6-heptenoate in this strain [32]. Ashby and co-workers [2] evaluated the production of mcl-PHA by Pseudomonas resinovorans from glucose, soybean and coconut oils. Culture on mixtures of these substrates led to monomer composition, and thermal and mechanical properties that were intermediate to those of PHAs obtained from pure glucose or plant oils. Also mixtures of linoleic and oleic acids as well as different plant oils were supplied to P. putida IPT046 and P. aeruginosa IPT171 to evaluate the contribution of unsaturated fatty acids to the insertion of unsaturated monomers into the polymer [64]. A non-linear relationship between the molar fractions of 3-hydroxy-6-dodecenoate detected in PHA and the linoleic acid supplied was observed, which is compatible with the ability of biosynthesis system saturation to channel intermediates of β-oxidation to PHA synthesis.

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11

Some bacterial species have also been shown to produce functionalized mcl-PHA from non-related carbon sources. P. aeruginosa 44T1 cultured in Euphorbia and castor oils produced mcl-PHA containing epoxy groups, besides other constituents normally found in PHA produced from fatty acids. Additional hydroxyl groups not involved in ester bond and the unsaturated monomer 6-hydroxy-3-dodecenoate have also been detected [15]. Epoxy group formation from soybean oil and 1-alkenes has been also reported for Pseudomonas stutzeri and Pseudomonas cichorii [28]. Concomitant lipase-catalysed self-epoxidation of the unsaturated precursors may be hypothesized as the underlying mechanism.

Triglycerides present in plant oils and animal fats can also be used for mcl-PHA production although the ability to convert these substrates is associated to lipase production and therefore much more restricted [1,7,66]. Recombinant expression of lipase genes has been used to improve triglyceride use [66]; however, functional expression of lipase in P. putida GPo1 did not enable growth on soybean oil (JGC Gómez, personal communication).

In relation to sugars, the use of an engineered strain of P. putida KT2440 for PHA production out of xylose has been reported recently [35]. The hemicellulose derivate, second in abundance after glucose and important for the production of second-generation bioethanol [13], can be used to support growth whereas mcl-PHA production is sustained by fatty acid addition in a sequential feeding strategy. Although only 20% w/w PHA content was obtained, the scheme may possibly be extended to combine cheap hemicellulosic hydrolysate with more expensive precursors to produce functionalized mcl-PHAs.

Perspectives and challenges Research into functionalized mcl-PHAs spans the last 25 years; nonetheless, opportunities remain open for the enhancement both of microbial strains and bioprocesses to produce optimal mcl-PHA compositions. In silico genomescale analysis of P. putida KT2440 has unveiled several isoenzyme-coding genes involved in hydroxy-acyl-CoA generation [43], which supports the well known high metabolic diversity of this strain, enabling it to incorporate different monomers into biopolyester. A higher fraction of long-chain-length monomers was produced when the main set of genes fadA and fadB was knocked-out [45,47]. This could be due to lower efficiency of the alternative

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β-oxidation routes on mcl-fatty acid precursors. In fact, when fadAB mutant cells were cultured with conventional aliphatic precursors, such as decanoic acid, a higher proportion of 3-hydroxydecanoic acid monomer was detected when compared to that of the wild type. However, this effect was less evident when unsaturated fatty acids, such as 10-undecenoic acid, were used as carbon sources [18]. It would be interesting to ascertain if this could be ascribed to a kinetic effect or to the substrate specificity of the active β-oxidation routes. The possibility of controlling PHA monomer composition has been demonstrated by the inactivation of various selected sets of those fad genes, and a collection of homopolymers has been produced [6,38,69, 71]. An interesting approach would be to verify the effect of those mutations on the incorporation of unsaturated monomers into the polymer when produced not only from pure fatty acids, but also from fats and oils. Unsaturated monomers have also been detected in scl-PHA producers, such as Rhodospirillum rubrum or Burkholderia sp. [57].

Although some evidence would suggest the existence of two PHA synthases in Burkholderia [58,59], the metabolic pathway supplying 3-hydroxy-4-pentenoic acid monomers remains unknown, thus hindering the performance of processes with controlled insertion of this monomer in the PHA. The growing availability of omics data and increased understanding of model strains have facilitated bottom-up approaches to design specialized strains [17,44]. By combining transcriptomic, proteomic and metabolomic measurements under well-controlled nutrient limitations, Poblete and co-workers [52] have reported the global multi-omics analysis of the P. putida KT2440 response to various nutrient limitations. Such studies provide a tremendous amount of knowledge that will be of great assistance in metabolic engineering design, thereby enhancing and diversifying mcl-PHA production. Furthermore, in-depth modelling and computational analysis of both metabolic versatility and PHA biosynthesis pathways represent a valuable tool for the design and production of less common and/or new functional PHA monomers. One example is the use of a detailed genome-scale model of P. putida (iJN746) to identify in silico a large set of non-glycolytic substrates, such as aromatic compounds, which are highly suitable for the production of different PHA monomers [44]. Very recently, a systems metabolic engineering approach has been reported, driven by in silico modelling to tailor P. putida for mclPHA synthesis on glucose [51]. Using the physiological properties of the parent wild type as constraints, elemen-

tortajada et al.

tary flux mode analysis of a large-scale model of P. putida metabolism was used to predict genetic targets for strain engineering. Among a set of priority-ranked targets, glucose dehydrogenase (encoded by gcd) was predicted as the most promising deletion target. This study illustrates the power of computational prediction to tailor microbial strains for the enhanced biosynthesis of added-value compounds. Moreover, the control of unsaturated content when mcl-PHA is produced from carbohydrates and related carbon sources will depend on the selection of strains showing enzymatic systems with a greater ability to channel such monomers into PHA precursors. In this respect, directed evolution strategies should be applied to modify enzymes involved in directing intermediates from fatty acid biosynthesis to PHA biosynthesis.

With regard to low-cost substrates, unsaturated fatty acid constituents present in oils or fats offer an excellent opportunity to insert functional groups in PHA. However, fats and oils are mixtures of different fatty acids that imply an even more complex metabolic network for tailoring polymers. This represents a challenge to mcl-PHA production and will depend on acquiring comprehensive knowledge of the factors involved in fatty acid and PHA metabolism. To reduce PHA production costs, crops could be used as a raw material source, though preferably those not competing with food sources. However, the greatest challenge will be the use of waste and biowaste, mostly because of their substrate and contaminant contents. Some such tested wastes from fats, oils and fatty acids include residues from food processing, such as waste frying oil [7] and residues from oil processing, such as oil sludge [23]. Considering carbohydrates and related carbon sources, a number of biowastes have been tested as substrates to produce PHA, namely glycerol, rice chaff, coconut oil cake, cotton seed cake, wafer residue, citrus pulp waste. Glycerol is an important carbon source, since it is the main residue from biodiesel production [8,21]. Although some of those residues may have the potential to generate unsaturated monomers, PHA composition has not been reported for most of them.

Last but not least, it is essential to define which PHA compositions are the best. This should be done by performing processing and technological assays to further characterize emerging strategies. To achieve this goal requires precise and robust manufacturing processes, to scale-up and provide samples to consolidate the design of tailormade PHAs. Data correlating the type and amount of precursors used with the molar fraction of monomers detected in the polymer, and the changes in physical and chemical

Medium-chain-length PHAs

properties of the resulting PHA could be used in the future to establish mathematical models. The different variables involved in these processes could be defined in order to provide a robust basis for directed optimization strategies to produce tuned functionalized mcl-PHAs. Acknowledgements. This work was supported by the Ibero-American Programme for Science, Technology, and Development (CYTED) and by the Spanish Ministry of Economy and Competitiveness (BIO201021049). The authors are members of the CYTED network 310rt0393.

Competing interests. None declared.

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RESEARCH ARTICLE International Microbiology (2013) 16:17-25 doi:10.2436/20.1501.01.176 ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

Association between Pseudonocardia symbionts and Atta leaf-cutting ants suggested by improved isolation methods Sarah E. Marsh,1¶,2§ Michael Poulsen,1¶,3§ Norma B. Gorosito,4¶,5§ Adrián Pinto-Tomás,1¶,6§,7§ Virginia E. Masiulionis,4¶,8§ Cameron R. Currie1,9* Dept. of Bacteriology, Univ. of Wisconsin-Madison, Madison, USA. 2Inst. de Biologie, École Normale Supérieure, Paris, France.

Section for Ecol. & Evol., Dept. of Biology, Univ. of Copenhagen, Copenhagen, Denmark. 4Center of Studies and Research, Natl.

Univ. of Quilmes, Bernal, Argentina. 5Dept. of Plant Production, Univ. of Buenos Aires, Argentina. 6Dept. of Biochemistry, School of Medicine, Univ. of Costa Rica, San José, Costa Rica. 7Center of Research in Microscopic Structures, Univ. of Costa Rica, San José, Costa Rica. 8Inst. of Biosciences, UNESP, Sao Paulo, Center of Studies of Social Insects, Rio Claro, Brazil. 9US Dept. of Energy Great Lakes Bioenergy Research Center, Univ. of Wisconsin-Madison, USA 1

3

Received 18 October 2012 · Accepted 22 March 2013 Summary. Fungus-growing ants associate with multiple symbiotic microbes, including Actinobacteria for production of antibiotics. The best studied of these bacteria are within the genus Pseudonocardia, which in most fungus-growing ants are conspicuously visible on the external cuticle of workers. However, given that fungus-growing ants in the genus Atta do not carry visible Actinobacteria on their cuticle, it is unclear if this genus engages in the symbiosis with Pseudonocardia. Here we explore whether improving culturing techniques can allow for successful isolation of Pseudonocardia from Atta cephalotes leaf-cutting ants. We obtained Pseudonocardia from 9 of 11 isolation method/colony component combinations from all 5 colonies intensively sampled.

The most efficient technique was bead-beating workers in phosphate buffer solution, then plating the suspension on carboxymethylcellulose medium. Placing these strains in a fungus-growing ant-associated Pseudonocardia phylogeny revealed that while some strains grouped with clades of Pseudonocardia associated with other genera of fungus-growing ants, a large portion of the isolates fell into two novel phylogenetic clades previously not identified from this ant-microbe symbiosis. Our findings suggest that Pseudonocardia may be associated with Atta fungus-growing ants, potentially internalized, and that localizing the symbiont and exploring its role is necessary to shed further light on the association. [Int Microbiol 2013; 16(1):17-25] Keywords: Pseudonocardia · Actinobacteria · symbiosis · mutualism · Atta leaf-cutter ants

Introduction The fungus-growing ant-microbe symbiosis originated approximately 50 million years ago [28]. Fungus-growing ants farm a specialized fungal cultivar that serves as the ants’ primary food source [15]. These fungal gardens host potentially virulent Corresponding author: Cameron R. Currie Department of Bacteriology 1550 Linden Drive, 6155 Microbial Sciences Building University of Wisconsin-Madison, Madison WI 53706, USA Tel. +1-6082658034. Fax +1-6082629865 E-mail: currie@bact.wisc.edu ¶ Affiliation when the work was conducted. §Current address. *

microfungal parasites in the genus Escovopsis, which consume the fungal mutualist [8,27]. To help suppress Escovopsis, the ants associate with Actinobacteria, which provides protection through antibiotic production. The Actinobacteria genus Pseudonocardia has been shown to be consistently present in the association, where it provides defensive antibiotic compounds [4,7,10,23,26]. In addition, other Actinobacteria genera have also been isolated or detected using cultureindependent methods [2,14,17], and have been suggested to also contribute to Escovopsis suppression [2,14]. The degree of specificity and functional role of these additional Actinobacteria remains to be fully established [cf. 4,5].

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The ant-Pseudonocardia association has been documented across most of the phylogenetic diversity of fungus-growing ants (tribe Attini), including the paleo-attine genera Apterostigma, Mycocepurus and Myrmicocrypta (lower attines), and the neo-attine genera Cyphomyrmex, Mycetosoritis, Mycetarotes (lower attines), Trachymyrmex and Acromyrmex (higher attines) [4,9,10]. Pseudonocardia is typically maintained on modified structures on the ant cuticle, and most genera of fungus-growing ants appear to provide the bacteria with nourishment through glandular secretions [9]. In at least two genera, Sericomyrmex and Atta, the abundance of Actinobacteria on the cuticle of the ants is markedly reduced or completely absent [9], possibly because alternative defenses play a more prominent role in controlling Escovopsis [11,13,32]. Despite the absence of visible Actinobacteria symbionts on the cuticle of Atta workers, several strains of Pseudonocardia that occur in the same clades as symbionts isolated from the other leaf-cutting ant genus Acromyrmex have been obtained (clades IV and VI in [4]).

Co-occurrence of Pseudonocardia in these phylogenetic clades suggests that Atta and Acromyrmex maintain the same Pseudonocardia species. However, consistent isolations of Pseudo-nocardia from Atta have proven difficult. This difficulty, in addition to the lack of visible growth of Actinobacteria on workers, makes it uncertain whether Atta persistently harbors symbiotic Pseudonocardia and even suggests that the Actinobacteria symbiont might have been lost in this ant genus.

However, given that microbial symbionts are often fastidious, attempts to improve culturing techniques might determine whether Pseudonocardia is consistently present in Atta. Here we explore whether different isolation techniques can reliably acquire isolates of Pseudonocardia from Atta. We conducted targeted isolations for Actinobacteria in association with Atta cephalotes applying four methods, including bead-beating, scraping, and washing with and without sonication, to different colony components. To improve our understanding of where Pseudonocardia might be located within Atta cephalotes colonies, we applied methods to material from both young and mature fungus garden, callow and mature workers, as well as dissected ant infrabuccal pockets.

Materials and methods Ant colonies. Most isolation attempts were done using five Atta cephalotes colonies collected in Gamboa, Panama in 2003 and 2005. These colonies (Pan03MB, Pan03BB, CC031208-10, AL050513-21, AL050513-22) had been housed at the University of Wisconsin-Madison for between 2 and 7 years at the time of isolation. Colonies were maintained at 24 °C in the dark to mimic underground conditions, with overhead lights only illuminated periodically.

Ant colonies were housed in plastic containers, with each colony kept in a large

marsh et al.

outer container (28 cm-high × 40 cm-wide × 56 cm-long) accommodating one smaller plastic container for refuse material (dump) and one to five smaller plastic containers (ranging from 3.0 cm-high by 7.5 cm-wide × 7.5 cm-long to 11 cm-high by 19.5 cm-long by 12.5 cm-wide) enclosing the colony fungus gardens. Each container had a 1 cm diameter hole drilled to allow ants to move in and out. Mineral oil was regularly applied to the top 4 cm of each outer container to prevent the ants from escaping. The ants were provisioned with maple (Acer sp.) and oak (Quercus sp.) leaves (frozen in the winter months and fresh during summer) three times per week. Leaves were supplemented with oatmeal, rice and cornmeal. Wet cotton balls were applied to the outer box to increase humidity. After initially determining what methods of isolation were most likely to be successful (see below), we increased the sampling to other laboratory colonies (A. cephalotes JS090511-01 from Panama; AP061022-01 from Costa Rica; EC090907-05 and MP090907-15 from Peru; A. colombica MP010708 from Panama; Atta sp. EC090827-04 and UP08 from Peru).

Isolations. To compare the yield and efficiency of various Actinobacteria isolation techniques, we tested 4 different methods on 1 to 5 colony components, using 1 or 2 types of media and material from five A. cephalotes colonies including callow and media workers, infrabuccal pockets dissected from media workers, as well as young and mature garden. The isolation methods included washing, scraping, homogenization, sonication, and beadbeating (see below and Table 1 for details). We replicated each method five times for each of five colonies, for a minimum of 25 Petri plates per method employed. In total, we conducted targeted isolations on 375 plates. For methods generating liquid inocula, plates were allowed to dry before being wrapped with parafilm.

Inocula were plated onto either chitin or carboxymethylcellulose (CMC) minimal nutrient media. Chitin medium is selective for Actinobacteria and we made it by autoclaving 15 g agar, 3 g chitin, 0.575 g K2HPO4, 0.375 g MgSO4·7H2O, 0.275 g KH2PO4, 7.5 mg FeSO4·7H2O, 0.75 mg MnCl2·4H2O, 0.75 mg ZnSO4·7H2O in 750 ml H2O for 30 min. On this medium, Actinobacteria appear white and faintly fuzzy, and grow flat on the surface. CMC media consisted of carboxymethylcellulose (5 g/l) and agar (15 g/l) as the sole nutrient sources. Once bacteria were observed on minimal media, individual cultures were transferred to richer yeast malt extract agar (YMEA, 4 g yeast extract, 10 g malt extract, 4 g dextrose, 20 g agar, 1-liter H2O). Antifungals (20 ml/l nystatin and 0.05 g/l cycloheximide) were added to media [3,33].

Actinobacteria derived from washes were obtained by aseptically placing workers in 1 ml of autoclaved Milli-Q water inside an autoclave-sterilized 1.5 ml microcentrifuge tube, which was vortexed for 30 s. One hundred µl of this suspension were plated onto each of five plates. Using this washing technique, we sampled medium-sized workers (three per replicate), callow workers (two per replicate, identified by their lighter color), dissected infrabuccal pockets, and small pieces of top (youngest) or middle (older) garden material. Media workers and middle fungus garden were sonicated for 45 s prior to plating.

‘Scraping’ isolations were done by rubbing the surface of workers with a sterilized bent micro-spatula under a dissecting microscope. This method has been employed to isolate visible Pseudonocardia from the cuticle of workers from other fungus-growing ant genera [3]. Bacteria inocula from the tool surface were transferred to chitin plates. This method was conducted on the cuticle of medium-sized workers. Infrabuccal pocket material was collected by dissecting out pockets, then using a sterile isolation loop to spread the pocket content directly onto chitin plates [18].

Bead-beating (Mini-Beadbeater-96, BioSpec products [http://www. biospec.com]) was performed on single medium-sized ants placed in 1.5-ml microcentrifuge tubes containing 500 µl phosphate buffer solution (PBS). We used PBS for bead-beating following Medina-Rivera (2008 Masters thesis, University of Puerto Rico-Mayaguez), who successfully used this solution to isolate Actinobacteria from a non-fungus-growing ant, Odontomachus ruginodis. Ants were bead-beaten for two and a half min, after which 100 µl of the suspension were pipetted onto either chitin or CMC plates.

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Pseudonocardia isolation from Atta ants

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Table 1. Actinobacteria and Pseudonocardia isolated from Atta cephalotes colonies Total Actinobacteria

Wash

Wash & Sonication

Scraping

Bead-beating

Pseudonocardia only

Liquid

Media

Colony component

#

CFUs Mean ± SE

Success rate

CFUs Mean ± SE

Success rate

H20

Chitin

Media callow worker

2

0±0

0

0±0

0

Media adult worker

3

16.4 ± 10.1

80 %

6.0 ± 2.8

60 %

Worker infrabuccal pocket

1

48.6 ± 17.4

100 %

0±0

0

Young garden

2

1.0 ± 0.6

40 %

0.4 ± 0.4

20 %

Mature garden

2

1.2 ± 0.5

60 %

0.4 ± 0.4

20 %

Media adult worker

3

11.2 ± 7.7

60 %

0.8 ± 0.4

20 %

Mature garden

2

32.4 ± 27.9

40 %

0.2 ± 0.2

20 %

Media adult worker

1

16.2 ± 11.6

100 %

0.4 ± 0.2

40 %

Media worker infrabuccal pocket

1

32.2 ± 11.3

80 %

0.4 ± 0.2

40 %

Chitin

Media adult worker

1

40.1 ± 20.4

100 %

4.3 ± 3.3

40 %

CMC

Media adult worker

1

35.8 ± 24.9

100 %

7.4 ± 5.1

80 %

H20

–

PBS

Chitin

Chitin

An overview of the four Actinobacteria isolation methods (coupled with colony component variation) and their results. Techniques fall into four main categories: washing, washing with sonication, scraping, and beat-beating. Within each technique the liquid used (water or PBS) and growth media used (chitin or CMC) are specified. The number (of individuals or pieces) and type of colony components are also specified including whole callow and media workers, infrabuccal pockets isolated from media workers, as well as young and mature garden. The number of each type of CFU counts and standard errors are reported per plate per colony, based on an average of five plates from each of five colonies, for the total Actinobacteria as well as for Pseudonocardia specifically. Isolation success rate of Actinobacteria and Pseudonocardia represent the proportion of the colonies yielding at least one CFU of total Actinobacteria and Pseudonocardia in particular in five isolation attempts per colony.

To evaluate whether Actinobacteria could have been introduced to the colonies on lab-provisioned leaves, we isolated Actinobacteria from leaf material.

These isolations targeted both bacteria from leaf surfaces and endophytic Actinobacteria from surface-sterilized leaves. Two to three leaves were collected from each of two oak and two maple trees routinely used to feed the ants in Madison, WI, for a total of five leaves per species and 10 leaves in total. Leaves were placed in sterile tubes and transported back to the lab. We cut leaves into 0.5-cm2 pieces and performed four types of targeted isolations. One set of untreated leaves was blotted directly onto chitin plates to isolate external bacteria. A second set was homogenized for 20 s with a sterile plastic pestle in 1.5-ml Eppendorf tubes containing 500 µl autoclaved Milli-Q water. One hundred µl of liquid were then plated onto chitin plates, spread and allowed to dry before wrapping with parafilm.

A third set of 0.5-cm2 pieces were surface sterilized by submersion in 70 % ethanol for 2 min. Pieces were then submerged in 1 % bleach containing 0.001 % Tween 20 for 1 minute. Leaves were subsequently rinsed in 1 ml autoclaved Milli-Q water for at least 10 s. The bleach wash was repeated three times. Finally, these surfacesterilized pieces were homogenized and plated as described above. A fourth set of 0.5-cm2 pieces were surface sterilized and blotted directly onto chitin plates to assess the efficiency of surface sterilization.

For all isolation attempts performed, plates were incubated for 3–5 weeks at room temperature (ca. 22 °C) before bacterial colonies with Actinobacteria morphology (filamentous, dry and spiky or dusty in appearance) were picked and transferred to YMEA. We allowed colonies to grow for 1–4 weeks, and subsequently confirmed morphology-based species identifications for all Pseudonocardia-like colonies using 16S rDNA sequencing (see below). To compare and quantify the efficiency of Actinobacteria and Pseudonocardia isolation, we counted the number of colony forming units (CFUs) produced on plates. We further calculated the proportion of colonies from which we were able to successfully isolate Actinobacteria and Pseudonocardia for each method. Additional Pseudonocardia isolated from lab colonies were included in tables and the phylogenetic analysis but not in the method efficacy analysis.

DNA sequencing and phylogenetic analysis. Once bacteria were obtained in axenic culture, we extracted DNA using a standard cetyltrimethylammonium bromide (CTAB) protocol [3,25]. DNA was quantified using a NanoDrop photospectrometer (Thermo-Fisher) and diluted to 50 ng/µl in trisethylenediaminetetraacetic acid (TE) buffer. We then amplified 16S rRNA using universal primers (27f 5′-AGAGTTTGATCMTGGCTCAG-3′ and 1492r

20

Int. Microbiol. Vol. 16, 2013

5′-TACGGYTACCTTGTTACGACTT-3′, [17b]) and nuclear Elongation Factor-Tu (EF-Tu 5′-CACGACAAGTCCCGAACCT-3′ and 5′-AGTTG TTGAAGAACGGGGTG-3′ [5]). Amplicons were sequenced using the dideoxyterminator method (Big Dye terminator mix; Applied Biosystems, Foster City, CA, USA) on an ABI 3730xl DNA Analyzer at the University of Wisconsin Biotechnology Center (Madison, WI, USA) following a cleaning using the CleanSeq magnetic bead protocol (Agencourt Biosciences, Beverly, MA, USA).

We edited sequences using Sequencher 4.5 (Gene Codes Corporation) to assemble the forward and reverse sequences, then reconciled differences and removed automated calling errors as required. In cases of uncertainty, samples were re-sequenced. To identify each isolate to the genus level, we used BLASTn nucleotide sequences against the non-redundant nucleotide collection using megablast [20]. We screened all 16S sequences for possible chimeras using Bellerophon [16] through the Greengenes database with the default parameters. We found no chimeric sequences.

To generate a phylogeny for all Pseudonocardia isolates, we aligned sequences from the EF-Tu gene using Clustal X [http://www.clustal.org/], checked the alignment in MacClade 4.07 [19], and thereafter used Mega 5.0 [30] to generate a maximum likelihood phylogeny (Tamura-Nei assuming uniform rates and using all sites, sequences were truncated such that all were the same length) and bootstrapped 1000 times. The analysis included 38 strains previously isolated and sequenced from other colonies of fungus-growing ants (14 from leaf-cutting ants: 5 in genus Atta, and 9 in the genus Acromyrmex) as well as 16 species of free-living (non-ant-associated) Pseudonocardia. We rooted the tree using two species of Streptomyces. All generated sequences are available at GenBank under accession numbers JQ731834–85.

Results and Discussion We were consistently able to isolate Actinobacteria from Atta cephalotes colonies using the different methods employed including washing, washing with sonication, scraping, and bead-beating (Table 1). On average, across all isolation methodcolony component combinations for the five ant colonies tested, we obtained 20.6±6.3 (Mean ± SE) Actinobacterial CFUs per plate. We also isolated Pseudonocardia from all Atta colonies, with an average of 2.86 ± 1.47 CFUs of Pseudonocardia per plate across all isolation methods and the five ant colonies sampled. 16S sequencing of these Actinobacteria revealed some phylogenetic diversity: seven Streptomyces, two Nocardioides, and seventeen Pseudonocardia isolates (Table 2).

Best-BLAST hits for the Nocardioides and Streptomyces isolated here correspond to strains obtained from a wide variety of habitats, including soil, ocean and other environments. While some insect-associated samples have similar 16S rRNA genes sequences (in one case, up to 100 % identity with a single termite gut-associated bacterial sequence that shares an equivalent level of identity with 57 other environmentally associated sequences), none of the strains found here had an identical 16S rRNA gene sequence to those previously reported from leaf-

marsh et al.

cutting ants [e.g., 2,14,29]. The phylogenetic distance between leaf-cutting ant-associated strains of Nocardioides and Streptomyces found here suggests a lack of specificity, and that the obtained isolates could be allochthonous.

Bead-beating was the most successful method for Actinobacteria isolation, generating an average of 40.1 ± 24.0 (Mean ± SE) CFUs per plate on chitin medium and 35.8 ± 24.9 CFUs per plate on CMC medium. Washing also resulted in large numbers of Actinobacteria, including an average of 16.4 ± 10.1 CFUs from washed workers. The number of Actinobacterial CFUs isolated for each of the 11 method/colony component combinations was significantly different from one another by a Kruskal-Wallis rank sum test (df =10, p = 0.008). Bead-beating, worker scraping, and infrabuccal pocket washes all resulted in the isolation of Actinobacteria, while worker washes and infrabuccal pocket scrapes were less efficient, yielding Actinobacteria from only 4 of the 5 colonies. The most efficient methods for Pseudonocardia isolation involved direct processing of worker ants, while fungus garden isolations were less successful.

Isolates of Pseudonocardia were never obtained from leaves provided to the colonies, despite this bacterial genus including endophytic species [6,22]. Collectively, the findings suggest that Atta colonies harbor Pseudonocardia and that the bacterium is associated with colony workers. This is congruent with the finding that the majority of the diversity of fungus-growing ants maintains Actinobacteria on the cuticle [cf. 9].

Previous work has had limited success isolating Pseudonocardia using maceration and vortexing methods, but Pseudonocardia had previously been obtained from five Atta colonies [4,33]. For Pseudonocardia isolations, nine of our eleven isolation method/colony component combinations resulted in at least one isolate (Table 1). Bead-beating of workers was the most efficient method for obtaining Pseudonocardia, yielding 7.4 ± 5.1 and 4.3 ± 3.3 CFUs per plate on CMC and chitin, respectively. Bead-beating isolated significantly more CFUs of Pseudonocardia than the other methods tested here by the Wilcoxon Rank sum test (W = 127, p-value = 0.016). This method was not successful across all colonies, recovering Pseudonocardia from only 4 out of 5 colonies tested, however bead-beating represents a substantial improvement over previously employed techniques. Worker washing was the only other method that resulted in substantial Pseudonocardia CFUs, with an average of 6 ± 2.8 per plate. A comparison of CFUs isolated by this method against all others pooled by the Wilcoxon Rank sum test approached, but did not reach, statistical significance (W = 68, P = 0.056). We were able to obtain Pseudonocardia from 10 additional lab colonies applying these methods (see Table 2, Fig. 1).

Panama Panama Panama Panama Panama Panama Panama

A. cep

A. cep

A. cep

A. cep

A. cep

A. cep

A. cep

CC031208-10(A)

CC031208-10(B)

MB03(C)

AL050513-22(B)

CC031208-10

MB03(C)

MB03(A)

worker wash/sonicate-H2O/Cih

worker wash/sonicate-H2O/Chi

worker wash/sonicate-H2O/Chi

worker wash, worker bead-beat-PBS/CMC

worker wash-H2O/Chi

worker wash-H2O/Chi

worker wash-H2O/Chi

worker wash & wash/sonicate-H2O/Chi

worker scrape-Chi

worker infrabuccal pocket wash-H2O/Chi

worker infrabuccal pocket scrape-Chi

worker bead-beat-PBS/CMC

worker bead-beat-PBS/CMC

worker bead-beat-PBS/Chi & PBS/CMC

worker bead-beat-PBS/Chi & PBS/CMC

worker bead-beat-PBS/Chi & PBS/CMC

worker bead-beat-PBS/Chi

worker bead-beat-PBS/Chi

worker bead-beat-H2O/Chi

worker bead-beat-H2O/Chi

worker bead-beat-H2O/Chi

worker e beat-H2O/Chi

new garden wash-H2O/Chi

homogenize-H2O/Chi

homogenize-H2O/Chi

garden wash-H2O/Chi

fungal garden wash/sonicate-H2O/Chi

fungal garden wash-H2O/Chi

blot-Chi

blot-Chi

blot-Chi

blot-Chi

Method

JQ731863

Streptomyces sp. JW1 (EU906929.1)

JQ731846

Pseudonocardia sp. TFS 1235 (EF216360.1) Pseudonocardia sp. YIM 45505 (NR_043742.1)

JQ731839 JQ731841 JQ731852 JQ731850

Nocardioides albus (AF004997.1) Streptomyces sp. 10213 (FJ262955.1) Streptomyces sp. Ank245 (HQ662223.1)

JQ731856

JQ731843 Pseudonocardia sp. BMWB1 (FJ948119.2)

Pseudonocardia sp. GB7 (EF451805.1)

Pseudonocardiaceae isolate SR 244a (X87314.1)

JQ731842

JQ731853 Pseudonocardia sp. GB7 (EF451805.1)

JQ731834 Nocardioides albus (AF004997.1)

JQ731835

Streptomyces sp. RS-2011-I28 (HE617241.1) Streptomyces sp. JS520 (JQ288109.1)

JQ731847

JQ731849

JQ731844

JQ731854

JQ731848

JQ731837

JQ731858

Streptomyces sp. 172624 (EF550508.1)

Pseudonocardia sp. BMWB1 (FJ948119.2)

Pseudonocardia sp. MVT7 (EU931094.1)

Pseudonocardia sp. BMWB1 (FJ948119.2)

Pseudonocardia sp. BMWB1 (FJ948119.2)

Pseudonocardia sp. BMWB1 (FJ948119.2)

Pseudonocardia sp. JS020524-10_B1F6 (EU928990.1)

JQ731857

JQ731845

Pseudonocardia sp. YIM 45505 (NR_043742.1)

Pseudonocardia sp. FXJ3.021 (JN683673.1)

JQ731838 JQ731840

Pseudonocardia sp. YIM 45505 (NR_043742.1)

JQ731851

JQ731862

Streptomyces sp. 163005 (GU263862.1) Actinobacterium ZXY009 (JN049458.1)

JQ731836 JQ731855

Streptomyces drozdowiczii strain GYB24 (JQ342930.1)

JQ768367

Pseudonocardia sp. YIM 45505 (NR_043742.1) Pseudonocardia sp. CC011205-08 (EU928998.1)

JQ731864

JQ731861

JQ731860

JQ731859

GenBank #

Streptomyces halstedii strain G8A-5 16S (CP002993.1)

Streptomyces owasiensis, strain: NBRC 13832 (AB184515.1)

Streptomyces sp. I08A-01824 (GU550598.1)

Streptomyces glauciniger strain FXJ14 (AY314782.1)

Closest 16S BLAST hit (GenBank Accession number)

Table of 16S rDNA sequences acquired from Atta sp. colonies and leaf material. Abbreviations are A. cep: Atta cephalotes; A. col: Atta colombia; Acer: Acer sp.; Q. pal: Quercus palustris; Chi: Chitin; CMC: carboxymethylcellulose; CR: Costa Rica. In the far let column, (A), (B) and (C) indicate different isolates from the same colony.

Panama Panama

A. cep

A. cep

AL050513-21(A)

Panama

A. cep

AL050513-21(B)

MB03

Panama Panama

A. cep

A. cep

BB03(B)

BB03

Panama Peru

A. cep

A. cep

Panama

A. cep

BB03(A)

MB03(A)

Panama

A. cep

AL050513-21

EC090827-04

Peru

Peru

A. cep

MP090907-15 Panama

Peru

A. cep

EC090907-05

Atta sp.

CR

A. cep

AP061022-01(A)

A. cep

Panama

A. cep

AL050513-22(A)

UP08

Panama

A. cep

MB03(B)

JS090511-01

US US

Panama

A. cep

MB03(B)

Acer

Panama

A. cep

AL050513-21(C)

Acer

Panama

A. col

MP010708F-1

N/A (A)

US

Acer

N/A (C)

N/A (B)

US US

Q. pal

Q. pal

N/A (B)

US

Q. pal

N/A (A)

N/A (C)

Location

Taxon

Colony Code (Isolate)

Table 2. Actinobacteria sequences

Pseudonocardia isolation from Atta ants

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marsh et al.

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Fig. 1. Elongation factor-Tu phylogeny of Pseudonocardia isolates from Atta and Acromyrmex generated in this study, along with other symbionts from across the diversity of fungus-growing ants. Pseudonocardia strains are labeled with the colony code and species of the ant from which they were isolated. Grey boxes highlight strains isolated as a part of this study, with strains isolated from colonies housed in the laboratory long-term in bold. Some Atta-associated strains of Pseudonocardia isolated in this study fall into clades with other leaf-cutting ant-associated Pseudonocardia (clades IV and VI, cf. 4); however, this study also identified two previously undescribed clades of ant-associated Pseudonocardia (indicated with circle and square).

Pseudonocardia isolation from Atta ants

Overall, this study isolated 20 additional strains of Pseudonocardia (4 confirmed by 16S rDNA gene sequencing, 3 confirmed by Ef-Tu gene sequencing and 13 confirmed by both) from a total of 15 ant colonies from three Atta species.

Because 16S rRNA genes alone provide limited species-level phylogenetic resolution, to further elucidate potential specificity we generated a phylogeny of the Pseudonocardia isolates based on EF-Tu (Fig. 1). Three Pseudonocardia isolates grouped with isolates from Acromyrmex leaf-cutting ants (one in clade IV, isolated by bead-beating with PBS onto chitin, and two in clade VI isolated by bead-beating with PBS onto chitin and CMC, Fig. 1). We also recovered 13 Pseudonocardia strains from two clades that had not previously been identified in associations with fungus-growing ants, which might represent hitherto unknown diversity within the symbiosis (Fig. 1, clades labeled with circle and square, respectively).

A concurrent study employing bead-beating with Acromyrmex ants identified sequences mapping to these novel clades from three Acromyrmex leaf-cutter ants (unpublished data, sequences included in Fig. 1; Acromyrmex echinatior colonies AP061104-01, AP061105-02, and AP061106-01 from Costa Rica). The clustering of these ant-associated strains suggests a specific association between Atta and Pseudonocardia.

Given the different isolation methods (e.g., bead-beating, washing with sonication) it is possible that the Pseudonocardia originated from inside the ants, and not on the external surface or in structures associated with cuticle. Additional isolations and phylogenetic studies will be needed to substantiate the presence of these novel genotypes and further explore the extent and role of this relationship.

We did not isolate Pseudonocardia from oak and maple leaves blotted on chitin media, but did obtain CFUs of other Actinobacteria: an average of 0.4 ± 0.28 (mean ± SE) (0.5 ± 0.47 from oak, and 0.3 ± 0.28 from maple). Isolations by homogenization resulted in an average of 0 CFUs of Actinobacteria from oak, and 0.8 ± 0.76 from maple. Sequencing of the strains recovered from these isolations revealed only Streptomyces. No Actinobacteria colonies were isolated on CMC medium. The Actinobacteria isolation rate from leaves was therefore low overall, with only a single leaf out of the ten resulting in CFUs. This indicates that ant-associated Pseudonocardia do not appear to be present on provisioned leaf material, suggesting their presence in the symbiosis, but we cannot rule out alternative sources of introduction of the bacteria to ant workers.

Our limited isolation success from certain colony components concurs with the culture-dependent findings by Mueller and colleagues [21]. Both studies found little to no Pseudonocardia

Int. Microbiol. Vol. 16, 2013

23

from the majority of Atta colony components, including fungal gardens and the ants’ infrabuccal pockets. Further, Pseudonocardia has not been detected in a recent community metagenomic study of Atta cephalotes garden [1], suggesting that it is unlikely to reside in this component of the system. Neither Mueller et al.

[21], Aylward et al. [1], nor this study found convincing evidence of garden or other environmental sources for Pseudonocardia in the Atta leaf-cutting ant symbiosis. We found the highest isolation rates directly from whole ants, suggesting that if the association of Pseudonocardia is maintained in Atta, it is specifically with ant workers, but not in the infrabuccal pocket.

Atta colonies include large work forces of highly polymorphic ants, in addition to age-dependent division of labor [15]. In other genera of attine ants, Pseudonocardia are present on a variety of cuticular structures [9], and might be the symbiont is present on inaccessible parts of the ant cuticle. It is furthermore conceivable that only some worker castes associate with Pseudo-nocardia and employ its antifungal compound(s), as is the case in the less polymorphic genus Acromyrmex, where mainly major workers carry Pseudonocardia [7,24]. Collectively, this suggests that ageor caste dependent within-colony composition of the bacterial symbiont population could make it harder to obtain without considerable targeted effort to screen across a wide range of ages and castes.

It is well known that many symbiotic microbes are extremely difficult to culture in isolation, and even when these microbes can grow in culture, they are often extremely fastidious, requiring a specific combination of growth conditions and nutrients normally provisioned by the host to meet their physiological needs [12]. Thus, one possible reason for previous attempts to isolate Pseudonocardia from Atta having had limited success is not an absence of the symbionts in these hosts, but rather that the strains associated with Atta are more fastidious. A considerable effort allowed us to isolate strains of Pseudonocardia from 15 Atta colonies; this suggests the presence of Pseudonocardia in the association, but the importance in defense remains to be established. Currie and Stuart [11] have shown that Atta ants employ fungus grooming at increased levels in response to pathogens, and FernándezMarín and colleagues [13] went on to demonstrate that Atta ants increase the levels of metapleural gland grooming behavior when challenged with Escovopsis. These findings indicate that Atta ants may have a low need to utilize Pseudonocardia for defense. Consequently, although our work supports that there might still be an association between Pseudonocardia and Atta, further work is needed to understand the role and impact of the bacteria on Atta colonies.

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Acknowledgements. We thank Michael Sullivan for assistance with bacterial isolations, X. Yu from the University of Wisconsin’s College of Agriculture and Life Sciences Statistical Consulting for statistical advice. We thank Alexis Barber and Jonathan Klassen for comments on this manuscript.

This work was supported by NSF CAREER Award DEB-747002 to CRC, and Lundbeckfonden to MP. We acknowledge the Organization for Tropical Studies (OTS) and the Ministry of Environment and Energy of Costa Rica, and the National Authority for the Environment of Panama for facilitating the research and granting collecting permits.

marsh et al.

13. Fernández-Marín H, Zimmerman JK, Nash DR, Boomsma JJ, Wcislo WT (2009) Reduced biological control and enhanced chemical pest management in the evolution of fungus farming in ants. Proc R Soc B 276:2263-2269 14. Haeder S, Wirth R, Herz H, Spiteller D (2009) Candicidin-producing Streptomyces support leaf-cutting ants to protect their fungus garden against the pathogenic fungus Escovopsis. Proc Natl Acad Sci USA 106:4742-4746 15. Hölldobler B, Wilson EO (1990) The Ants. Belknap Press of Harvard, Cambridge, MA, USA 16. Huber T, Faulkner G, Hugenholtz P (2004) Bellerophon: a program to

Competing interests. None declared.

detect chimeric sequences in multiple sequence alignments. Bioinformatics 20:2317-2319 17. Kost C, Lakatos T, Böttcher I, Arendholz W-R, Redenbach M, Wirth R

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(2007) Non-specific association between filamentous bacteria and fungus-growing ants. Naturwissenschaften 94:821-828 17b. Lane DJ (1991) 16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M (eds) Nucleic acid techniques in bacterial systematics. Wiley, Chichester, UK, pp 115-175 18. Little AEF, Murakami T, Mueller UG, Currie CR (2006) Defending against parasites: fungus-growing ants combine specialized behaviours and microbial symbionts to protect their fungus gardens. Biol Lett 2:12-16 19. Maddison DR, Maddison WP (2005) MacClade 4 release version 4.07 for OSX. Sinauer Associates, Inc., Sunderland, MA, USA 20. Morgulis A, Coulouris G, Raytselis Y, Madden TL, Agarwala R, Schaffer AA (2008) Database indexing for production MegaBLAST searches.

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RESEARCH ARTICLE International Microbiology (2013) 16:27-33 doi: 10.2436/20.1501.01.177 ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

In vitro interaction between Bacillus megaterium strains and Caco-2 cells Ana C. López,1 Jessica Minnaard,2 Pablo F. Pérez,2,3 Adriana M. Alippi1* Bacteriology Unit, Phytopathology Research Center, Faculty of Agricultural and Forestry Sciences, National University of La Plata, La Plata, Argentina.2Center of Research and Development in Food Cryotechnology (CONICET), La Plata, Argentina.

3 Chair of Microbiology, Faculty of Exact Sciences, National University of La Plata, La Plata, Argentina

1

Received 3 January 2013 · Accepted 3 March 2013

Summary. To further our understanding of the virulence potential of Bacillus megaterium strains, cell association and

invasion assays were conducted in vitro by infecting human enterocytes (Caco-2 cells) with 53 strains of this bacterium isolated from honey. Two series of experiments were performed: (i) necrosis and cell detachment assays with the supernatants of bacterial culture filtrates from 16-h cultures and (ii) adhesion/invasion assays in which cultured enterocytes incubated with bacteria from 3-h cultures were resuspended in Dulbecco’s modified Eagle’s medium and chloramphenicol. The detachment of Caco-2 cells was evaluated by staining the cells with crystal violet. Necrosis was assessed by fluorescence microscopy of cells labeled with propidium iodide. Association (adhesion plus invasion) was determined by plate counts and invasion in an aminoglycoside protection assay. The results showed that spent culture supernatants detached and necrotized Caco-2 cells in a strain-dependent manner. Seven out of 53 B. megaterium filtered culture supernatants caused complete cell detachment. Suspensions of these same bacterial strains adhered and invaded enterocytes in 2-h infection experiments. To our knowledge, this is the first report on the interaction between B. megaterium and intestinal epithelial Caco-2 cells. [Int Microbiol 2013; 16(1):27-33] Keywords: Bacillus megaterium · Caco-2 cells · adhesion · cellular necrosis · spent culture supernatant · honey

Introduction Due to its natural properties and to industrial control measures, commercially sold honey has minimal microbial contamination. Nevertheless, it may contain yeasts and spore-forming bacteria [41]. Although vegetative forms of disease-causing bacteria have not been found in honey, bacterial spores, when present, can remain viable for long periods without losing their infectivity.

Corresponding author: A.M. Alippi Unidad de Bacteriología. Centro de Investigaciones de Fitopatología (CIDEFI)-CIC. Facultad de Ciencias Agrarias y Forestales Universidad Nacional de La Plata Calle 60 y 119 S/N c.c. 31 1900 La Plata, Argentina Tel. + 54-2214236758. Fax +54-2214252346 E-mail: alippi@biol.unlp.edu.ar; adrianaalippi@gmail.com

*

Several studies have shown that Bacillus and Paenibacillus species, including Bacillus cereus, B. thuringiensis, B. licheniformis, B. pumilus, B. megaterium, B. coagulans, P. alvei, and P. larvae [2,3,11,17,35,45], are commonly detected in honey. While P. larvae and P. alvei are associated with different honeybee diseases, B. subtilis, B. pumilus, B. cereus, B. licheniformis, B. circulans, and B. megaterium are predominant in the digestive tracts of larvae and honey bees [3,41]. Bacillus cereus and B. megaterium are also common in soils, dust, and flower surfaces. In Argentinian honeys, the most prevalent spore-forming species are P. larvae, P. alvei, B. cereus, B. pumilus, and B. megaterium [2,3,21,22].

Bacillus megaterium is an aerobic, gram-positive, spore-forming bacterium that in addition to honey has been isolated from various food sources, i.e., shellfish, raw meat,

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rice, corn, and soymilk [1–3,18,31]. The antibiotic-like compounds produced by B. megaterium include megacins and other inhibitory substances [4,28,32]. To date, B. megaterium has not been associated with any human disease, but its pathogenic potential is evidenced by the presence in its genome of several toxin-encoding genes, i.e., bceT (enterotoxin-T), hblC, hblA, hblD (HBL complex), and nheA, nheB, nheC (NHE complex) [23,34,38]. In addition, the production of a novel heat-stable toxin by B. megaterium strain F98/3079 has been reported [43]. Of particular relevance to this study is the detection of both hemolytic and coagulase activities in 39 B. megaterium isolates obtained from honey samples of different origins [22].

The pathogenic potential of bacteria and the biological activity of bacterial toxins have been studied in vitro in a variety of cell culture systems [14]. As a cell line with differentiation markers characteristic of mature intestinal epithelial cells, enterocyte-like Caco-2 cells (ATCC HTB-37) are a well-recognized in vitro model system.

These markers include functional tight junctions, microvilli, and a number of brush border-associated enzymes and transporters, i.e., peptidases, esterases, P-glycoproteins, and uptake transporters for amino acids, bile acids, and carboxylic acids [36,39]. Consequently, Caco-2 cells have been used to mimic bacteria-host interactions in the intestinal tract [6,7,12,16,25,26,33]. They were chosen in our study of the virulence potential of B. megaterium strains isolated from honey because the intestinal epithelium is rapidly exposed to potential foodborne pathogens. Although interactions between Bacillus spp. and cultured cells, e.g., Caco-2, human epithelial type 2 cells (Hep-2), and cell lines derived from cervical cancer (HeLa), have been investigated [26,37,38], to our knowledge, ours is the first study to focus on B. megaterium.

Materials and methods Bacterial strains, media and culture conditions. Fifty-three strains of B. megaterium were evaluated. Except for B. megaterium NRRL B-939, all strains were isolated from honey (49 from Argentina [Bm1–Bm21 and Bm25–Bm52], 2 from Brazil [Bm23 and Bm24], and 1 from France [Bm22], as previously described [22]). The bacteria were stored at –80 °C in tryptone soy broth with 20 % (v/v) glycerol as cryoprotectant. Prior to the experiments, each strain was cultured in nutrient broth (NB) under constant agitation at 32 °C for 16 h. Afterwards, they were inoculated (2 % v/v) in 5 ml of NB and incubated under constant agitation at 32 °C for 3 h or 16 h, according to the assay conditions. Bacteria were harvested by centrifugation (900 ×g for 10 min).

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Culture of Caco-2 cells. The human colon cancer cell line Caco-2 was routinely grown in Dulbecco’s modified Eagle’s medium (DMEM) (25 mM glucose) (Life Technologies, Carlsbad, CA, USA), supplemented with 15 % (v/v) heat-inactivated (56 ºC, 30 min) fetal calf serum (FCS, PAA Laboratories, Pasching, Austria), 12 IU penicillin/ml-12 μg streptomycin/ml (Life Technologies), and 1 % (v/v) non-essential amino acids (Life Technologies). Monolayers were prepared in 24-well tissue culture plates (Greiner Bio One, Frickenhausen, Germany) by seeding 7 × 104 cells per well. Experiments and cell maintenance were carried out in an atmosphere of 5 % CO2 at 37 °C. Assays were performed with differentiated cells (late post-confluence, 15 days of culture).

Detachment of Caco-2 cells. Detachment of enterocyte-like cells was studied as reported previously [25]. Briefly, filter-sterilized (0.45-µm pore size) spent culture supernatants (SCSs) were obtained from 16-h bacterial cultures in NB. Differentiated Caco-2 monolayers were incubated with 0.5 ml of serially diluted SCSs in NB (ratios of SCSs/total volume ranged from 0/0.5 to 0.5/0.5 per well) at 37 °C for 1 h. The cells were washed twice with phosphate buffered saline (PBS) (pH 7.2), fixed at room temperature with 2 % (v/v) formaldehyde in PBS for 1 min, and washed again with PBS. Afterwards, the cells were stained by incubating them with 500 µl of a crystal violet solution (0.13 % [w/v] crystal violet, 5 % [v/v] ethanol, and 2 % [v/v] formaldehyde in PBS) at room temperature for 20 min. After a PBS wash to remove excess stain, the samples were treated with freshly prepared 50 % (v/v) ethanol at room temperature for 1 h. Absorbance was measured in a microplate reader at 620 nm (BioTek, Winooski, Vermont, USA). All experiments were performed in triplicate. The percentage of cell detachment was calculated as follows: cell detachment % = 100 × (Ac – As)/Ac, where Ac is the A620 of control cells and As the A620 of sample cells.

Necrosis. Necrosis was assessed according to a modification of a previously published method [47], using, short incubation periods to prevent cell detachment (37 °C, 20 min). After their incubation with SCSs from 16-h bacterial cultures in NB, the cells were washed twice with binding buffer containing 25 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 125 mM NaCl, 2.5 mM CaCl2, (pH 7.2), and 0.2 % (p/v) gelatin. One µg of propidium iodide in 100 µl of binding buffer was then added to each well, and the plates were placed on ice for 15 min.

The samples were subsequently mounted in 50 % (v/v) glycerol in PBS and analyzed by conventional fluorescence microscopy using a Leica DMLB microscope coupled to a Leica DC 100 camera (Leica Microscopy Systems, Heerbrugg, Switzerland).

Cell association assays. Bacterial cultures (37 °C, 3 h) were centrifuged and the pellets were suspended in 1 ml of DMEM containing 100 μg chloramphenicol/ml. As reported by Minnaard et al. [26], under these conditions the bacteria remain viable but do not grow. The bacterial concentration was adjusted to 108 colony-forming units (CFU)/ml. The cell monolayers were washed twice with PBS, treated with the bacterial suspensions (multiplicity of infection = 100 bacteria per cell), and incubated in an atmosphere of 5 % CO2 at 37 °C for 2 h. To evaluate cell association (adhering plus invading bacteria), the monolayers were exhaustively washed with PBS and then incubated with 1 ml of distilled water per well, which lysed only the eukaryotic cells. The bacteria present in the lysate were counted by plating appropriate dilutions on nutrient agar and incubating the samples at 32 °C for 16 h.

Cell invasion assays. Enterocyte invasion was assessed by the aminoglycoside protection assay as previously reported [13], using gentamicin to kill non-internalized bacteria. Briefly, after infection, monolayers washed three times with PBS were treated with 1 ml of gentamicin

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(100 µg/ml in PBS) per well, and incubated at 37 °C for 1 h. The monolayers were then washed twice with PBS, lysed in 1 ml of distilled water, and maintained at 37 °C for 1 h. Serial dilutions of the suspensions were plated on nutrient agar as described above. To assess the presence of spores, aliquots of each sample were heat treated in a block heater (Nova Etica, Vargem Grande Paulista, SP, Brazil) at 60 °C for 30 min, and appropriate dilutions were plated onto nutrient agar. Strains were considered to be invasive only when the number of survivors after gentamicin treatment was significantly higher than the number after thermal treatment [26]. All infection assays were performed in FCS-free medium.

PCR analysis. Strains that detached enterocyte-like cells were screened by polymerase chain reaction (PCR) for the presence of enterotoxic genes (entFM, entS, and piplC) and sph (sphingomyelinase) gene according to the procedures described by Ghelardi and co-workers [10].

DNA was amplified in a thermal cycler (Eppendorff Mastercycler, Eppendorf AG, Hamburg, Germany) using the same PCR primers and conditions as previously reported [10]. PCR products were resolved in 1.6 % agarose gels in 0.5× TBE buffer, stained with GelRed (genBiotech SRL, Buenos Aires, Argentina), and visualized with a UV transilluminator (UVP, Upland, California, USA). Gel images were digitalized and photographed using a digital image capture gel documentation system (Digi Doc-it, UVP, v. 1.1.25, Upland, California, USA).

Results Detachment of Caco-2 cells. Exploratory experiments analyzing the effect of different B. megaterium SCSs on fully differentiated Caco-2 cells showed that the filtered culture supernatants of seven out of the 53 bacterial strains tested caused complete cell detachment, with no evidence of biological activity in the remaining 46 strains (data not shown). The SCSs of these strains, i.e., Bm1, Bm8, Bm9, Bm10, Bm29, Bm30 and Bm51, all isolated from Argentinian honeys, were used for further studies.

Differences in the biological activity of the seven SCSs at low doses were observed. At a SCS/total volume ratio of 0.2/0.5, the biological activities of strains Bm8, Bm29, Bm30 and Bm51 were higher than those of strains Bm1, Bm9 and Bm10 (P < 0.05). For all of the strains under study, undiluted supernatants (ratios 0.5/0.5) caused the total detachment of Caco-2 cell monolayers (Fig. 1).

Necrosis. Cell necrosis was evaluated with SCSs prepared from 16-h cultures of strains Bm1, Bm8, Bm10, Bm29,

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Statistical analysis. Microsoft Office Excel (version 2003) was used for the statistical analysis. The results of the experiments were compared by means of two-tailed Student's t-test, with P < 0.05 considered statistically significant.

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Fig. 1. Detachment of Caco-2 cells after their co-incubation (1 h at 37 °C) with different concentrations of filtered culture supernatants of Bacillus megaterium strains: Bm1 (closed triangles), Bm8 (open squares), Bm9 (open circles), Bm10 (closed circles), Bm30 (closed squares), Bm29 and Bm51 (open triangles; the same symbol is used for Bm29 and Bm51 strains because they induced the same responses). All symbols overlap at 0 and 100 % cell detachment. A relative concentration of supernatant equal to 1 indicates undiluted supernatants. Spent cultures were diluted in NB.

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Fig. 2. Fluorescence microscopy of Caco-2 cells co-incubated at 37 °C for 20 min with filtered culture supernatants of strains Bm1 (B), Bm8 (C), Bm10 (D), Bm29 (E), and Bm30 (F). The control was co-incubated with NB medium only (A). Cells were labeled with propidium iodide (PI). Red cells indicate the nuclei of necrotic cells (40×). All micrographs are representative of three independent experiments conducted with successive cell passages.

and Bm30 (Fig. 2). When briefly (20 min) incubated with the cell monolayers, none of these strains induced detachment. Instead, high ratios of necrotic cells were obtained (Fig. 2B–D). By contrast, only a few necrotic cells were seen in untreated control monolayers (Fig. 2A).

Cell association and invasion. Infection assays were conducted only with strains whose supernatants detached Caco-2 cells. Strains Bm1, Bm9 and Bm10 associated with the monolayers at levels significantly higher than strains Bm29, Bm51 and Bm8 (P < 0.05) (Fig. 3A). Strain Bm30 was not included in these assays because it was gentamicin resistant and thus inappropriate for the assessment of invasion in the aminoglycoside protection assay.

As seen in Fig. 3B, strains Bm1 and Bm10 were the most invasive (P < 0.05). The number of bacteria penetrating the cells was 1 × 104 CFU/ml for strain Bm1 and 6 × 103 CFU/ml for strain Bm10. For strains Bm8, Bm9 and Bm29, about 4 × 103 CFU/ml invaded the Caco-2 cells while strain Bm51 was not internalized at all (Fig. 3B). Note that the experiments were performed with 3-h cultures to minimize the presence of sporulated microorganisms. For all six strains evaluated, the number of survivors was significantly (P < 0.05)

higher after gentamicin treatment than after thermal treatment (60 °C, 30 min).

PCR analysis. Strains Bm8, Bm9, Bm10, and Bm51 were positive for all four enterotoxic genes tested, i.e., sph, entFM, entS, and piplC, as determined by PCR. Strain Bm29 was positive for entS, sph and piplC, and strain Bm1 for entS and piplC (data not shown).

Discussion Bacteria belonging to the B. cereus group have been associated with outbreaks of foodborne illness and other diseases [42]. Their pathogenicity can be related to the production of several toxins [5,8–10,15,20,40,42]. In this work, we focused on B. megaterium, since genes encoding biologically active molecules have also been found in strains of the bacterium isolated in Argentinian honeys [23].

In our study, high doses of SCSs from seven strains of B. megaterium completely detached monolayers of cultured enterocytes and led to cell necrosis. Furthermore, six of

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Fig. 3. Cell-association (A) and cell-invasion (B) assays of six different Bacillus megaterium strains with fully differentiated Caco-2 cells. Values Âą standard deviations are from a representative experiment with three replicates.

those detachment-inducing strains were also capable of cell invasion (one strain could not be tested, see above).

These findings demonstrated that the induction of cell detachment by extracellular factors secreted by B. megaterium is not a general property of this species, which is in agreement with findings published for B. cereus [25,27]. From the dose-response curves obtained in cell detachment assays with B. cereus SCSs, Minnaard and co-workers defined three types of strains: (i) high detaching (HD) strains detach cells even at low doses; (ii) low detaching (LD) strains detach cells at high doses; and (iii) non-detaching (ND) strains [27]. Accordingly, in the present study of B. megaterium, two HD (strains Bm 29 and Bm 51) and five LD (strains Bm 1, Bm 8, Bm 9, Bm 10 and Bm 30) strains were identified, whereas the remaining 46 strains were ND. The biological activity of the SCSs from the seven HD and LD strains may depend on secreted extracellular factors. The SCSs of strains Bm9 and Bm51 resulted in cell detachment within a few minutes, indicative of their stronger biological activity than the SCSs of the other five isolates, and could therefore not be assayed for their ability to cause cell necrosis.

All B. megaterium strains that led to enterocyte detachment had been previously reported to have hemolytic and coagulase activities [22,23]. Note that B. cereus, B. thuringiensis and B. megaterium share several sequences encoding virulence factors [10,23]. Besides the enterotoxic genes sph, entFM, entS, and piplC detected in this work,

sequences of the HBL complex, the nheC gene from the NHE complex, and bceT and cytK genes had been previously detected in the LD strains Bm8 and Bm9 [23]. HD strains Bm29 and Bm51 were positive for the enterotoxic genes sph, entS, and piplC and sph, entFM, entS, and piplC, respectively, while LD Bm10 was positive for all four enterotoxic genes, and LD Bm1 was positive for only two of the genes tested. By contrast, Bm30 was negative for all the virulence genes tested here and in a previous study [23].

Adhesion is a key event in the pathogenicity of many bacteria. Adhered bacteria either remain on the cell surface (e.g., diffusely adherent E. coli [DAEC]; enteropathogenic E. coli [EPEC]) or subsequently invade the cell (e.g., enteroinvasive E. coli [EIEC]; Shigella spp.; Yersinia spp.; Listeria spp.) [6,24,29,44,46].

The values obtained for B. megaterium in the association and invasion assays were in the range of those previously reported for B. cereus [26]. Note that, while the LD strains Bm8 and Bm29 had the lowest association values, their internalization efficiencies (radio of invading to associated bacteria) were relatively high (0.16 and 0.06, respectively).

Bacteria that adhere to eukaryotic cells can trigger a biological response by interacting with host cell surface receptors linked to signaling pathways [13]. In vitro, this ability depends on both the differentiation status of the host cells, which determines receptor expression, and the

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growth phase at which the bacteria were harvested [19].

We found that cultured human enterocytes infected with B. megaterium suspensions prepared from 3-h cultures led to cell detachment (data not shown). These findings are in agreement with those reported for B. cereus infection of cultured human enterocytes [26]. In addition, we were able to show that cell detachment following infection with bacterial vegetative cells was inhibited when the assays were performed in the presence of chloramphenicol, suggesting that protein synthesis was necessary for a maximal biological effect.

The ability of B. megaterium to attach to and subsequently invade enterocytes may reflect its pathogenic potential. This conclusion supports a previous study showing that B. megaterium strains isolated from infant formula are not only able to adhere to and invade epithelial cells but are also positive for the presence of enterotoxins [38].

To our knowledge, this is the first report that demonstrates strain-dependent biological effects following the infection of cultured human enterocytes by B. megaterium isolates, and the biological activity of spent culture supernatants. Our results are consistent with the multifactorial character of B. megaterium virulence. Although further studies are necessary to completely unravel the pathogenic potential of this species, our work provides the first step in the study of the interaction between B. megaterium and its host in the context of intestinal infections.

Acknowledgements. This research was partially supported by the Agencia Nacional de Promoción Científica y Tecnológica (ANPCyT), CONICET and UNLP, Argentina. ACL; JM and PFP are members of the Scientific Research Career of Consejo Nacional de Investigaciones Científicas y Técnicas CONICET (CCT La Plata, Argentina) and AMA is a member of the Scientific Research Career of Comisión de Investigaciones Científicas de la Provincia de Buenos Aires (CIC), Argentina.

Competing interest. None declared.

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RESEARCH ARTICLE International Microbiology (2013) 16:35-44 DOI: 10.2436/20.1501.01.xxx ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

A novel bleb-dependent polysaccharide export system in nitrogen-fixing Azotobacter vinelandii subjected to low nitrogen gas levels Wataru Hashimoto, Yukiko Miyamoto, Mayumi Yamamoto, Fuminori Yoneyama, Kousaku Murata* Laboratory of Basic and Applied Molecular Biotechnology, Graduate School of Agriculture, Kyoto University, Kyoto, Japan Received 21 January 2013 · Accepted 20 March 2013

Summary. The alginate biofilm-producing bacterium Azotobacter vinelandii aerobically fixes nitrogen by oxygensensitive nitrogenases. Here we investigated the bacterial response to nitrogen/oxygen gas mixtures. A. vinelandii cells were cultured in nitrogen-free minimal media containing gas mixtures differing in their ratios of nitrogen and oxygen.

The bacteria did not grow at oxygen concentrations >75 % but grew well in the presence of 5 % nitrogen/25 % oxygen.

Growth of wild-type and alginate-deficient strains when cultured with 50 % oxygen did not differ substantially, indicating that alginate is not required for the protection of nitrogenases from oxygen damage. In response to decreasing nitrogen levels, A. vinelandii produced greater amounts of alginate, accompanied by the formation of blebs on the cell surface. The encystment of vegetative cells occurred in tandem with the release of blebs and the development of a multilayered exine.

Immunoelectron microscopy using anti alginate-antibody revealed that the blebs contained alginate molecules. By contrast, alginate-deficient mutants could not form blebs. Taken together, our data provide evidence for a novel blebdependent polysaccharide export system in A. vinelandii that is activated in response to low nitrogen gas levels. [Int Microbiol 2013; 16(1):35-44] Keywords: Azotobacter vinelandii · alginate · nitrogen stress · outer membrane vesicles · bleb-dependent export

Introduction Azotobacter vinelandii is a member of the Gammaproteobacteria that under oxic conditions fixes atmospheric nitrogen by converting it into ammonia [8]. Nitrogen fixation requires the expression of molybdenum-, vanadium-, Corresponding author: K. Murata Laboratory of Basic and Applied Molecular Biotechnology Graduate School of Agriculture Kyoto University Gokasho, Uji, Kyoto 611-0011, Japan Tel. +81-774383766. Fax +81-774383767 E-mail: kmurata@kais.kyoto-u.ac.jp

*

and iron-nitrogenases [30] and large amounts of ATP, which in turn implies a role for aerobic respiration. Since the three nitrogenases are sensitive to oxygen, bacterial cells protect them from oxygen damage through the removal of intracellular oxygen by an unknown mechanism and by cell-surface repression of the influx of oxygen from the outside. In the latter, two possible mechanisms have been suggested: respiratory protection and the development of an alginate biofilm that acts as a barrier [2,24].

Cell-surface proteins are responsible for respiratory protection by consuming oxygen as the final electron acceptor [22]. Bacterial genome sequencing and biochemical experimental data have revealed that four NADH-

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ubiquinone oxidoreductases, five terminal oxidases, and two ATP synthases confer respiratory protection [28].

Their expression is positively or negatively controlled by the oxygen-responsive transcriptional regulator CydR. Alginate is a linear polysaccharide consisting of two monosaccharides, b-d-mannuronate and its C-5 epimer al-guluronate [6]. It protects nitrogenases from oxygen damage by inhibiting oxygen uptake under phosphate-limited conditions [24]. A. vinelandii cells produce alginate as a component of the extracellular biofilm. In addition to A. vinelandii, pseudomonads such as Pseudomonas aeruginosa and P. syringae secrete alginate biofilms [6,14]. In both azotobacters and pseudomonads the synthesis and secretion of alginate is mediated by algA, algD, algE, algF, algG, algI, algJ, algK, algL, algX, alg8, and alg44. These genes are assembled into a single cluster and their expression is closely regulated by several factors, such as the sigma factor AlgU [6,23].

In the case of A. vinelandii, the molecular weight, production level, and mannuronate/guluronate ratio of alginate vary depending on the dissolved oxygen tension [21].

Diverse mannuronate C-5-epimerases specific for A. vinelandii cells have been implicated in the increase in guluronate that, together with divalent cations, is essential for the formation of an alginate barrier [29]. GDP-mannose 6-dehydrogenase (AlgD) catalyzes the conversion of GDPmannose to GDP-mannuronic acid and plays a crucial role in alginate synthesis [3], which occurs during the stationary phase of the growth cycle. The expression of algD and thus alginate formation are controlled by a two-component system consisting of GacS-GacA, which regulates the transcription level of the stationary phase-specific sigma factor RpoS [7]. Thus, the disruption of algD either indirectly, by alterations in one or both of these components, or directly, e.g., by alterations in the gene itself, prohibits alginate production by A. vinelandii [3].

The bacterial response to oxygen has been well studied with respect to the protection of nitrogenases from oxygen damage and during phosphate limitation [2,7,18,21,23], whereas little is known about the bacterial response to disturbances in nitrogen levels under normal phosphate conditions. In this study, the effects of nitrogen/oxygen levels on bacterial growth, alginate production, and cell structure were examined in wild-type A. vinelandii strain ATCC 12837 cells and in the mutant, which is deficient in alginate production through the insertion of a tetracycline-resistance gene into algD.

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Materials and methods Bacterial growth. Azotobacter vinelandii strain ATCC 12837

(NBRC 13581 purchased from the National Institute of Technology and Evaluation, Japan) was aerobically precultured at 30 ºC in nitrogen-free modified Burk’s (MB) medium [20] consisting of 20 mg sucrose/ml, 0.2 mg NaCl/ml, 0.05 mg CaSO4·2H2O/ml, 0.2 mg MgSO4·7H2O/ml, 2.9 mg Na2MoO4·2H2O/ml, 27 mg FeSO4·7H2O/ml, 0.66 mg K2HPO4/ml, and 0.16 mg KH2PO4/ml. The bacterial cells were grown at 30 ºC with shaking at 123 rpm in 50 ml of MB medium in 500-ml airtight flasks, except during the 5-min injection of the filter-sterilized gas mixture at a pressure of 0.05 MPa every 8 h, as described below (Fig. 1A). Gas injection was evaluated using a single-channel oxygen meter (Fibox 3, Presens) to determine the dissolved oxygen concentration (Fig. 1B).

Cultivation under various gas-mixture conditions. The gas mixtures were purchased from Imamura Oxygen, Kyoto, Japan. The involvement of alginate in nitrogen fixation was examined in wild-type A. vinelandii strain ATCC 12837 and in mutant cells carrying a disrupted algD gene. Both were grown on nitrogen (yeast extract)-free or supplemented MB agar plates under atmospheric air conditions (approximately 78 % nitrogen, 21 % oxygen, and trace amounts of other gases). Bacterial growth in response to the various gas mixtures was determined by cultivating the wild-type and mutant cells in MB liquid media in the presence of gas mixtures containing various ratios of nitrogen and oxygen (Fig. 1) as follows: The cells were cultivated under hermetic conditions. Every 8 h, they were exposed for 5 min to artificial air containing 79 % nitrogen and 21 % oxygen (N79/O21), N100, 50 % nitrogen/25 % oxygen/25% helium (N50/O25), N50/O50, N25/O75, or O100.

Plasmid construction. The algD gene [3] was cloned from the genomic DNA of A. vinelandii cells using the DNeasy blood & tissue kit (Qiagen, Tokyo, Japan). To construct the algD disruptant, algD (1.3 kb) was PCR-amplified using KOD-Plus polymerase (Toyobo, Osaka, Japan), the A. vinelandii genomic DNA as template, and two synthetic oligonucleotide primers (forward, 5′-GGGAATTCTTTTCGGACTGG GCTATGTAGG-3′; reverse, 5′-GGGAATTCTACCAGCAGATGCCTTC TGC-3′) (Hokkaido System Science, Sapporo, Japan) containing an EcoRI site at their 5′ ends. After amplification, the PCR product was ligated using Ligation High (Toyobo) with HincII-digested pUC119 (Takara Bio, Otsu, Japan). The resultant plasmid was designated pUC119-algD. The tetracycline-resistance gene (1.9 kb) was PCRamplified using KOD-Plus polymerase, the plasmid pACYC184 (Nippon Gene, Tokyo, Japan) as template, and two synthetic oligonucleotide primers (forward, 5′-GGATTCTCATGTTTGACAGCTTATCATCGA-3′; reverse, 5′-CCCTACCGGACAGCGGTGCGGACTGTTGTA-3′). The amplified fragment was ligated into AatI-digested pUC119-algD using Ligation High, taking advantage of the unique AatI restriction site present in the middle of algD. The resultant plasmid was designated pUC119algD::Tetr. After digestion of the plasmid with EcoRI, the fragment containing algD::Tetr was isolated and ligated with the EcoRI-digested plasmid pKTY320 [13] containing the ampicillin-resistance gene, yielding pKTY320-algD::Tetr. E. coli strain DH5a cells were transformed with pKTY320-algD::Tetr. The correct sequence of the resultant plasmids was confirmed by DNA sequencing [27]. DNA manipulations were performed as described previously [26].

Transformation of Azotobacter vinelandii. A. vinelandii cells were transformed according to the method of Page and Sadoff [19].

Briefly, the cells were oxically grown at 30 ºC in MB medium with

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Fig. 1. Azotobacter vinelandii culture system. (A) Injection of the gas mixture into the culture flask.

(B) Dissolved oxygen in the medium of cultures exposed to N100, N75/O25, N50/O50, N25/O75, or O100 was measured using an oxygen meter. The asterisk indicates the dissolved oxygen in medium exposed to atmospheric air.

10 mg glucose /ml (instead of 20 mg sucrose/ml) until they reached exponential phase (OD at 600 nm, 0.5). Fifty microliters of the A. vinelandii cell culture was then transferred to sterilized filter paper placed on an antibiotic-free MB plate solidified with agar and 5 ml (1.5 mg) of plasmid pKTY320-algD::Tetr was added. After 24 h of incubation at 30 °C, the bacterial cells on the filter paper were recovered in MB buffer consisting of 0.05 mg CaSO4·2H2O/ml, 0.2 mg MgSO4·7H2O/ml, 2.9 mg Na2MoO4·2H2O/ml, 27 mg FeSO4·7H2O/ml, 0.66 mg K2HPO4/ml, and 0.16 mg KH2PO4/ml and inoculated onto a tetracycline-containing MB plate, followed by incubation at 30 ºC for 24 h. Tetracycline-resistant and ampicillin-sensitive colonies were isolated and designated as disruptant cells. Gene disruption in the constructed strain was confirmed by genomic PCR and DNA sequencing.

Alginate assay. Alginate was extracted from the bacterial culture (300 ml) with 30 ml of 1 M NaCl and 12 ml of 0.5 M ethylenediaminetetraacetic acid, as described previously [24]. Bacterial cells were removed by centrifugation at 15,000 ×g at 4 °C for 5 min and the resultant supernatant (280 ml) was mixed with 900 ml of ice-cold isopropanol at 4 °C for 10 min. The precipitated alginates were collected in a centrifugation step (15,000 ×g at 4 °C for 5 min), washed with ice-cold 70 % ethanol, and dried at 25 °C under vacuum. The dried alginates were then dissolved at 4 °C for 24 h in 1 ml of distilled water. Alginate content was determined by measuring the increase in absorbance at 235 nm, indicating the detection of a double bond in the alginate degradation products formed in a reaction catalyzed by Sphingomonas sp. strain A1 alginate lyase A1-I, purified from recombinant Escherichia coli cells, as previously described [31]. Briefly, 50 ml of 1 M Tris-HCl (pH 7.0) was added

to the diluted alginate solution (1 ml) and the absorbance was measured (preAbs235). A 500-ml aliquot of the alginate solution was then incubated at 37 °C for 3 h with 5 ml of 18.8 mg A1-I/ml, and the absorbance was again measured (postAbs235). A calibration curve was obtained by measuring the increase (postAbs235 – preAbs235) in the absorbance at 235 nm, using sodium alginate from the kelp Eisenia bicyclis (Nacalai Tesque, Kyoto, Japan) as the standard.

Electron microscopy. A 1-ml aliquot of A. vinelandii cells grown in MB medium under various mixed gases conditions until stationary phase was pre-treated with an equal volume of fixative [4 % paraformaldehyde and 4 % glutaraldehyde in 30 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (pH 7.4), containing 100 mM NaCl and 2 mM CaCl2 (HEPES)] at 25 °C for 1 h. After centrifugation, the bacterial cells were fixed at 4 °C for 24 h with 2 % glutaraldehyde in HEPES and washed at 4 °C with HEPES. Thin sections were prepared and analyzed at Tokai Electron Microscopy Analysis Co. (Nagoya, Japan) as follows: The fixed cells were treated at 4 °C for 2 h with 2 % OsO4 in HEPES, dehydrated in an ethanol series (30 %, 50 %, 70 %, 90 %, and 100 %, 10 min each), treated twice with propylene oxide at 25 °C for 20 min, and then embedded at 60 °C for 48 h in Quetol-812 (Nisshin EM, Tokyo, Japan). The resultant samples were cut with a diamond knife on a 2088 Ultratome V (LKB, Bromma, Sweden) into sections of 65–70 nm thicknesses, stained at 25 °C with 2 % uranyl acetate for 15 min followed by lead citrate for 3 min, and examined using a JEM-1200EX transmission electron microscope (JEOL, Tokyo, Japan) at 80 kV. For immunoelectron microscopy, the bacterial cells were fixed at 4 °C for 1 h by mixing the culture (1 ml) with an equal volume of fixative

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(4 % paraformaldehyde and 0.05 % glutaraldehyde in 0.1 M phosphate buffer [pH 7.4]). After centrifugation, the bacterial cells were washed with 0.1 M phosphate buffer (pH 7.4), dehydrated at 4 °C in an ethanol series (50–70 %, 15 min each), and then embedded at 50 °C for 24 h in LR-White resin (London Resin Co., London, UK). The resultant samples were cut with a diamond knife on a 2088 Ultrotome V (LKB) into 70-nm thick sections. After blocking at 25 °C for 30 min with 1% bovine serum albumin and 1.5 % goat serum in phosphate saline, the sections were incubated at 25 °C overnight with anti-alginate antibodies [4] (kindly supplied by Dr. Kazuo Okuda, Kochi University, Japan), washed with phosphate saline containing 1 % bovine serum albumin, and then incubated at 25 °C for 1 h with 10 nm colloidal gold-conjugated goat antirabbit IgG (BBI solutions, Cardiff, UK). The immunolabeled sections were washed with phosphate saline, fixed at 25 °C for 5 min with 2 % glutaraldehyde in 0.1 M phosphate buffer (pH 7.4), and stained at 25 °C with 2 % uranyl acetate for 15 min and lead citrate for 3 min. They were

Fig. 2. Azotobacter vinelandii exposed to a gas mixture.

The bacterial cells were grown on plates containing modified Burk’s medium (MB) and exposed to various gas mixtures. (A) Atmospheric air. (B) N75/O25. (C) N50/O50. (D) N25/O75. WT and MT indicate the wild-type parental strain and alginate-deficient mutant, respectively. Plates on the left contained no nitrogen source, those on the right contained yeast extract.

then examined using a JEM-1200EX transmission electron microscope (JEOL) at 80 kV.

Results Alginate unessential for protection from oxygen damage. Azotobacter vinelandii cells produce alginate at the stationary growth phase [7] but not during exponential growth [11]. Wild-type cells at the stationary growth phase under atmospheric air conditions produced 0.215 mg alginate/ml, as measured in the culture broth, whereas alginate production by mutant cells carry-

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Fig. 3. Azotobacter vinelandii growth profile. (A) Bacteria grown in nitrogen-free MB medium and exposed to various gas mixtures. Closed circles, wild-type strain under atmospheric air conditions; open circles, alginate-deficient mutant under atmospheric air conditions; closed squares, wild-type strain exposed to N75/O25; open squares, mutant exposed to N75/O25; closed triangles, wild-type strain exposed to N50/O50; open triangles, mutant exposed to N50/O50. (B) Gas stress of cells in the late-exponential growth phase. Closed circles, wild-type strain under atmospheric air conditions; open circles, alginate-deficient mutant under atmospheric air conditions; closed triangles, wild-type strain exposed to N50/O50; open triangles, mutant exposed to N50/O50; closed rhombus, wild-type strain exposed to N25/O75; open rhombus, mutant exposed to N25/O75. (C) The effect of nitrogen levels on bacterial growth. Wild-type cells were grown under various mixed gases conditions. Closed circles, N75/O25; closed squares, N50/O25; closed triangles, N5/O25.

ing the disrupted algD gene was negligible. Mutant cells formed nonmucoid colonies on nitrogen-free MB medium solidified with agar, in contrast to the mucoid colonies of wild-type cells (Fig. 2). However, the growth of wild-type vs. mutant cells did not differ (Fig. 3A), indicating that algD disruption had no influence on either bacterial growth or nitrogen fixation.

Both the wild-type and the mutant cells were cultivated in MB medium under atmospheric air conditions, N79/ O21, N75/O25, N50/O50, N100, or O100. In medium exposed for 5 min to a gas mixture of increasing oxygen content, the dissolved oxygen concentration increased accordingly (Fig. 1B). The growth of wild-type cells under atmospheric air conditions was essentially the same as that of cells exposed for 5 min at 8 h intervals to N79/O21. As expected, wild-type and mutant cells exposed to N100 or O100 did not grow in nitrogen-free medium since A. vinelandii is an aerobic, nitrogen-fixing bacterium. Increased oxygen levels (21 %, 25 %, and 50 %) were accompanied by a decrease in growth of both the wild-type and the mutant cells (Fig. 3A). Similar growth patterns were observed in the plate cultures exposed to the gas mixtures and the mucoid phenotype of each strain did not change in re-

sponse to the different gas ratios tested (Fig. 2). Wild-type and mutant cells were also cultivated aerobically up to the exponential growth phase in the MB medium and subsequently grown under atmospheric air conditions, N25/ O50, and N25/O75 (Fig. 3B). The poor growth of either culture under N25/O75 conditions was consistent with a lack of protection from oxygen damage rather than an insufficiency of nitrogen gas. Therefore, in general, there was little difference in growth between wild-type and mutant cells exposed to the different gas mixtures.

For stationary phase cultures of both strains, growth under high oxygen conditions (25 %) yielded amounts of alginate larger than those obtained under atmospheric air conditions (atmospheric air: 0.215 mg/ml; N50/O25: 0.646 mg/ml). However, the alginate content was higher under N50/O25 than under N50/O50 (0.381 mg/ml). The higher alginate content in the culture broth under N50/O25 conditions in comparison with that of N50/O50 was dependent on the growth of the bacterial cultures.

Response to nitrogen level. The growth of bacterial cells exposed to N75/O25 or N50/O25 was comparable to that of cultures under atmospheric air conditions (Fig. 3A,C).

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Note that A. vinelandii cells were able to grow even when exposed to N5/O25, a slight delay was observed in the exponential growth phase in comparision with bacterial cells exposed to N75/O25 and N50/O25.

Alginate production seemed to be dependent on the nitrogen level, with lower nitrogen ratios associated with increased alginate synthesis (N75/O25: 0.220 mg/ml; N50/ O25: 0.646 mg/ml; N5/O25: 0.793 mg/ml). Under atmo-

Fig. 4. Bleb formation on the cell surface of Azotobacter vinelandii.

Wild-type cells were grown to stationary phase in the presence of various gas mixtures. Thin sections were prepared from fixed cells and observed by electron microscopy.

(A) Atmospheric air. (B) N50/O25.

(Bâ&#x20AC;˛) Magnification of the boxed region in (B). (C) N5/O25. (D) N50/ O50. (E) N25/O50.

spheric air conditions, A. vinelandii formed rod-shaped cells and produced polyhydroxybutyrate (PHB), as detected by electron microscopy of thin sections (Fig. 4A). In cells incubated under low nitrogen (N50/O25 and N5/O25) (Fig. 4B,C) or high oxygen (N25/O50) (Fig. 4D,E) conditions, a large number of blebs formed on the cell surface whereas fewer blebs were observed on bacterial cells grown under atmospheric air (Fig. 4A).

Fig. 5. Bleb-dependent alginate export in Azotobacter vinelandii. Wild-type (A, D, E1, and E2) and alginate-deficient mutant (B and C) cells were grown to stationary phase in the presence of various gas mixtures. Thin sections were prepared from fixed cells and observed by electron microscopy. The sections were further stained with anti-alginate antibodies for immunogold electron microscopy (D, E1, and E2). (A) Wild-type cyst formation (N25/ O50). (A′) Magnification of the boxed region in (A). (B) Mutant cell (atmospheric air). (C) Mutant cell (N50/O50). (D) Wild-type cell (atmospheric air). (E1 and E2) Wild-type cell (N5/O25). Thin arrows indicate alginate contained in blebs; the thick arrow points to bleb fusion at the interface of neighboring cells.

In addition, some cells exposed to N25/O50 accumulated higher amounts of PHB in the cytoplasm and produced extracellular multilayered biofilms, indicating the conversion of vegetative cells to the cyst form (Fig. 5A).

A novel alginate secretion system involving the formation of cell-surface blebs. A. vinelandii cells incubated under nitrogen concentrations lower than the atmospheric concentration formed blebs (Fig.

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4B–E) and the alginate concentration in the culture broth increased. In cells exposed to N25/O50, bleb formation was followed by the formation of an extracellular multilayered biofilm (Fig. 5A′), suggesting that the blebs contained alginate polysaccharides. The mutant cells did not form blebs when incubated in various gas mixtures, including atmospheric air and N50/O50 (Fig. 5B, C). Immunoelectron microscopy was carried out to directly examine the pres-

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ence of alginate in the blebs of wild-type cells exposed to N5/O25. The accumulation of gold particles in the blebs (Fig. 5E1 and 5E2, arrows) confirmed that they contained alginate. In wild-type cells grown under atmospheric air conditions, the polysaccharide was distributed on the cell surface (Fig. 5D).

Discussion In this study, we investigated the responses of A. vinelandii cells to incubation with gas mixtures differing in their percentages of nitrogen and oxygen. Specifically, cell growth, the concentration of alginate in the culture broth, and cell structure at the stationary growth phase in response to several different gas conditions were examined in order to better understand the bacterial response to nitrogen levels higher or lower than atmospheric conditions.

Alginate biofilms are thought to protect bacterial cells from oxygen damage at limited phosphate levels [24].

However, even at high levels of oxygen (N50/O50 and N25/O50) and normal phosphate levels, both the wild-type and the alginate-deficient mutant cells were able to grow quite well, suggesting that alginate does not contribute to the protection of nitrogenases from oxygen damage.

Azotobacter vinelandii cells exposed to high oxygen levels (50 %) were round rather than rod-shaped (Fig. 4).

In the presence of low nitrogen/high oxygen, vegetative cells formed cysts by synthesizing extracellular multilayered biofilms. Together with an increase in respiration at the exponential growth phase, encystment at the stationary growth phase is an important mechanism by which bacteria protect themselves from excess oxygen influx at high oxygen concentrations.

In A. vinelandii cells, several stress factors, such as dessication [10], cause a shift from vegetative cells to alginate-coated cysts [25]. Similarly, changes in the gas composition from atmospheric conditions, e.g., to low nitrogen/high oxygen, also trigger cyst formation in A. vinelandii. In our study, alginate polysaccharide accumulated in the blebs formed on the cell surface during encystment.

Release of the blebs from the cell surface was accompanied by the formation of multilayered biofilms that surrounded the bacterial cells. This system of direct alginate export in A. vinelandii through cell-surface alterations can be regarded as the counterpart of the one allowing alginate uptake in Sphingomonas sp. strain A1, which we characterized in a previous study [1]. This gram-negative sphingo-

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monad accumulates external alginate in mouth-like pits that form on the cell surface, allowing the transport of polysaccharide across the outer and inner membranes into the cytoplasm by a periplasmic binding-protein-dependent ATP-binding cassette (ABC) transporter [17]. Subsequently, alginate is degraded into monosaccharides by four alginate lyases [9]. These unusual cell-surface alterations in these two species of bacteria may be related to the unique rheological properties (e.g., gel-sol transition and/or chelator) of alginate.

In gram-negative bacteria, macromolecules such as DNA and proteins are secreted through the formation of cell-surface blebs or membrane vesicles [16], but to our knowledge, a similar bleb-dependent system that mediates polysaccharide secretion has not been described. Following the initial report in the early 1970s of bleb formation during encystment [10], there have been no further investigations into this process, and the components of the blebs have not been determined. This may have been due to the inability to obtain stable blebs in cultured bacteria. In our study, the regulation of gas levels under the culture conditions described here resulted in stable blebs and thus insights into their contents.

Haemophilus influenza shows a natural competence for DNA and transfers DNA to recipient cells through its release from cell-surface blebs [12]. As an example of protein secretion through bleb formation, the cell-surface blebs of P. aeruginosa contain autolysin, the release of which results in the killing of other bacterial cells [15].

Analogous to the “transformasomes”, i.e., the DNAcontaining bleb-like structures on the cell surface of H. influenzae [12], we have named the novel alginate export system of A. vinelandii as a “transalgisome.”

Gram-negative bacteria often release outer membrane blebs or vesicles 0.5–1 nm in diameter into the culture medium. These blebs typically contain enzymes or signaling molecules that are transferred to other bacterial cells when the vesicles fuse to the outer membrane of the recipient cells, providing a mechanism for prokaryotic cell–cell communication [16]. This has been observed in Bacillus subtilis, which transfers cytoplasmic molecules to neighboring B. subtilis but also to Staphylococcus aureus, and E. coli cells, through intercellular nanotubes formed on the cell surface [5]. A similar process may occur in A. vinelandii cells, in which blebs were found to mutually fuse at the interface of neighboring cells (Fig. 5E2, thick arrow). Note that bleb fusion provides a means of cell–cell communication, e.g., in the conversion of a population of vegetative

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cells to cysts.

In conclusion, the ability of A. vinelandii cells grown at normal phosphate concentrations to fix nitrogen indicated that alginate did not contribute to the protection of oxygensensitive nitrogenases from oxygen damage. Gas stress, such as induced by low nitrogen/high oxygen levels, triggered bleb formation on the surface of these bacterial cells.

During encystment, alginate contained in membrane vesicles was secreted via cell-surface blebs, resulting in aggregation of the polysaccharide, which constituted the exine of cysts. To our knowledge, this is the first report of a bacterial bleb-dependent polysaccharide export system.

Acknowledgements. Anti-alginate antibodies were kindly provided by Dr. Kazuo Okuda, Kochi University, Japan. We thank Dr. Akihito Ochiai and Ms. Rinko Kabata for their excellent technical assistance. This work was supported, in part, by the Targeted Proteins Research Program (W.H.) from the Ministry of Education, Culture, Sports, Science and Technology of Japan and by the program (K.M.) for the Promotion of Basic Research Activities for Innovative Bioscience (PROBRAIN) in Japan.

Competing interests: None declared.

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J Bacteriol 178:2479-2488 16. Mashburn-Warren LM, Whiteley M (2006) Special delivery: vesicle trafficking in prokaryotes. Mol Microbiol 61:839-846 17. Mishima Y, Momma K, Hashimoto W, Mikami B, Murata K (2003) Crystal structure of AlgQ2, a macromolecule (alginate)-binding protein of Sphingomonas sp. A1, complexed with an alginate tetrasaccharide at 1.6-Å resolution. J Biol Chem 278:6552-6559 18. Oelze J (2000) Respiratory protection of nitrogenase in Azotobacter species: is a widely held hypothesis unequivocally supported by experimental evidence? FEMS Microbiol Rev 24:321-333 19. Page WJ, Sadoff HL (1976) Physiological factors affecting transformation of Azotobacter vinelandii. J Bacteriol 125:1080-1087 20. Peña C, Campos N, Galindo E (1997) Changes in alginate molecular mass distributions, broth viscosity and morphology of Azotobacter vinelandii cultured in shake flasks. Appl Microbiol Biotechnol 48:510-515 21. Peña C, Trujillo-Roldán MA, Galindo E (2000) Influence of dissolved oxygen tension and agitation speed on alginate production and its molecular weight in cultures of Azotobacter vinelandii.

Enzyme Microb Technol 27:390-398 22. Poole RK, Hill S (1997) Respiratory protection of nitrogenase activity in Azotobacter vinelandii–roles of the terminal oxidases. Biosci Rep 17: 303-317 23. Remminghorst U, Rehm BH (2006) Bacterial alginates: from biosynthesis to applications. Biotechnol Lett 28:1701-1712 24. Sabra W, Zeng AP, Lünsdorf H, Deckwer WD (2000) Effect of oxygen on formation and structure of Azotobacter vinelandii alginate and its role in protecting nitrogenase. Appl Environ Microbiol 66:4037-4044 25. Sadoff HL (1975) Encystment and germination in Azotobacter vinelandii. Bacteriol Rev 39:516-539 26. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA 27. Sanger F, Nicklen S, Coulson AR (1977) DNA sequencing with chain-terminating inhibitors. Proc Natl Acad Sci USA 74:5463-5467 28. Setubal JC, dos Santos P, Goldman BS, et al. (2009) Genome se-

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quence of Azotobacter vinelandii, an obligate aerobe specialized to support diverse anaerobic metabolic processes. J Bacteriol 191:4534-4545 29. Steigedal M, Sletta H, Moreno S, Maerk M, Christensen BE, Bjerkan T, Ellingsen TE, EspĂŹn G, ErtesvĂĽg H, Valla S (2008) The Azotobacter vinelandii AlgE mannuronan C-5-epimerase family is essential for the in vivo control of alginate monomer composition and for functional cyst formation. Environ Microbiol 10:1760-1770 30. Walmsley J, Toukdarian A, Kennedy C (1994) The role of regulatory

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RESEARCH ARTICLE International Microbiology (2013) 16:45-52 doi: 10.2436/20.1501.01.179 ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

Low virus to prokaryote ratios in the cold: benthic viruses and prokaryotes in a subpolar marine ecosystem (Hornsund, Svalbard) Borys Wróbel,1* Manuela Filippini,2 Joanna Piwowarczyk,1 Monika Kędra,1,3 Karol Kuliński,1 Mathias Middelboe4 1 Institute of Oceanology, Polish Academy of Sciences, Sopot, Poland. 2Institute of Evolutionary Biology and Environmental Studies, University of Zurich, Switzerland. 3Chesapeake Biological Laboratory Center for Environmental Science, University of Maryland, USA. 4 Marine Biological Section, University of Copenhagen, Helsingor, Denmark

Received 1 February 2013 · Accepted 1 March 2013

Summary. The density and spatial distribution of benthic viruses and prokaryotes in relation to biotic and abiotic

factors were investigated in sediment cores collected in Hornsund, a permanently cold fjord on the West coast of Svalbard, Norway. The cores were obtained from the mouth of the fjord to the central basin, along a longitudinal transect. The results of our analyses showed lower densities of viruses (0.2 × 108 to 5.4 × 108 virus-like particles/g) and lower virus-toprokaryote ratios (0.2–0.6, with the exception of the uppermost layer in the central basin, where the ratio was about 1.2) at the study site than generally found in the temperate areas, despite the relatively high organic matter content in subpolar sediments. Variations in benthic viral and prokaryote abundances along gradients of particle sedimentation rates, phytopigment concentrations, and macrobenthic species composition together suggested the influence of particle sedimentation and macrobenthic bioturbation on the abundance and spatial distribution of prokaryotes and viruses in cold habitats. [Int Microbiol 2013; 16(1):45-52] Keywords: viriobenthos · bacteriobenthos · marine sediment · subpolar ecosystems · stable isotopes · Svalbard

archipelago (Norway)

Introduction Marine benthic viruses are one of the most diverse and abundant components of the global ecosystem. Metagenomic analyses have shown the presence of at least 104 viral genotypes per kilogram of sediment [5], which is between

Corresponding author: B. Wróbel Institute of Oceanology Polish Academy of Sciences Powstańców Warszawy 55 81-712 Sopot, Poland Tel. +48-587311767. Fax +48-585512130 Email: bwrobel@iopan.gda.pl

*

10- and 100-fold higher than estimated for the water column [6]. Other studies have shown that in most marine areas, viral densities are around 107–1010 g sediment–1 and thus 10- to 1000-fold higher in sediments than in the overlaying water column [8–10,15]. Virus-to-prokaryote ratios (VPRs) in the sediments range over four orders of magnitude, from close to 0.1 in oligotrophic and deep-sea sediments to >100 in eutrophic estuarine ecosystems [8], with viral abundance and activity correlated positively with both benthic prokaryote activity [25,40–42] and prokaryote abundance [23].

The high density and activity of prokaryotes and small distances between prokaryotic cells in the sediment is thus expected to result in a higher frequency of virus-prokaryote encounter [8] and consequently a high rate of viral pro-

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duction [40,48]. Deposition of viruses (attached to sinking particles) from the water column could also contribute to viral counts in the sediments. Biotic and abiotic factors affecting the properties of the sediment, such as sedimentation, bioturbation and irrigation processes at particular sites or regions might affect viral counts directly (by influencing delivery of viruses, viral decay, the chance for virus-host cell encounter, and the rate of viral propagation) or indirectly, influencing prokaryote growth and turnover.

Viral lysis can be responsible for as much as 50 % of mortality of prokaryotes in the sediment [37]. One effect of viral lysis is the acceleration of the transformation of particulate to dissolved organic matter. Instead of being directed to higher trophic levels through protozoan grazing on prokaryotes, organic matter released through lysis is assumed to be rapidly metabolized by the prokaryotic community, providing up to 38 % of its carbon demands [8,48].

In order to elucidate the role of viruses in controlling prokaryote mortality and biogeochemical cycling in different benthic environments, it is necessary to investigate the abundance and distribution of viruses and prokaryotes in diverse geographical regions. Only a few studies have thus far investigated the distribution of microbes along trophic gradients. For example, Hewson et al. [30], investigated the vertical and horizontal distribution of viruses and prokaryotes along a trophic gradient in subtropical estuaries (Australia). In another study [10], the top sediment layer was collected along a trophic gradient from oligotrophic Eastern to more productive Western Mediterranean.

Similarly, the distribution of benthic viruses and prokaryotes was investigated along productivity gradients in the Eastern Mediterranean [13] and the Chilean upwelling zone [40]. Generally, these studies have found a decrease in the viral abundance at the surface of the sediment with decreasing organic matter supply and prokaryote activity along transects from shallow coastal sediments towards deeper open ocean environments.

In this paper we present the first analysis, as far as we know, of the vertical and horizontal abundance of viruses in marine sediments collected along a trophic gradient at high latitudes. We designed our sampling strategy to capture an ecologically important gradient in productivity, sedimentation and bioturbation activity in the sub-arctic fjord Hornsund, Svalbard. We aimed to identify the biotic and abiotic factors that may influence the vertical and horizontal distribution of microbes and VPR in the subpolar benthic ecosystem.

Wróbel et al.

Materials and methods Study site and sampling. Hornsund is a 30-km long and 12- to 15-km wide fjord. It is located on the west coast of Spitsbergen (between 74º and 81º N, and 10º and 35º E), an island within the Svalbard archipelago in Northern Europe. Hornsund is the southernmost fjord of the island. The fjord is exposed to cold Arctic water transported by the Sorkapp current as well as inflows of relatively warmer Atlantic water of the West Spitsbergen current. A sea ice cover occurs between December and June, and floating sea ice from the Barents Sea from May to July [26].

About 70 % of Hornsund’s catchment area is covered by glaciers. The fjord sediment consists of glacimarine mud [26]. Sedimentation is dominated by melt-water processes, which release organic and mineral particles to the water column. In the central basin, high sedimentation rates (about 1 cm/yr) increase water turbidity, reducing the euphotic zone to a depth of 5–10 m (J. Wiktor and M. Darecki, personal communication) and limiting primary production. In the mouth of the fjord, where sedimentation rates are much lower (about 0.1 cm/yr [26]), the depth of the euphotic zone is 20–55 m (J. Wiktor and M. Darecki, personal communication) and primary production is correspondingly higher than in the central basin, with estimates comparable to those for regions without ice coverage at lower latitudes (up to 120 g C m–2 yr–1) [17,45]. Due to its pristine nature and limited anthropogenic stressors, Hornsund was selected as a European Marine Biodiversity Site (EMBS) and an All Taxa Biodiversity Inventory site (ATBI) [53].

To analyse the benthic communities in Hornsund, we used the published data collected from 89 stations located throughout the fjord between 2002 and 2007 [32]. The species composition of benthic fauna inhabiting soft sediments in Svalbard fjords has been shown to be stable on time scales of 5 years and more [32,33,46]. Arctic benthic species are predominantly sessile or undertake at most local migrations [3,52], tend to be relatively long lived (2–5 years and longer), and exhibit spatial and temporal patterns that reveal major environmental features and gradients [1,14,27,32,33,44].

We collected sediment cores at 3 locations in August 2010 (HornWest, HornMid, HornEast; Table 1, Fig. 1) along the central axis of Hornsund using a GEMAX gravity corer for soft sediments (Geological Survey of Finland, Espoo, Finland), which allows for undisturbed sampling of soft muds. The cores were cut in 1-cm-thick slices onboard and homogenized manually. Small subsamples (about 1–2 g) were packed into plastic bags, sealed, and stored at –80 °C until further analysis (about 3 months).

Virus and prokaryotes counting. The procedure for virus counting was adapted from Danovaro and Middelboe [12]. Briefly, approximately 1 g of the thawed sediment was mixed with 5 ml of ice-cold 2.5 % glutaraldehyde in 5 mM Na4P2O7 (pH of the solution: 8.2). After a 15-min incubation on ice, the mixture was sonicated (3 cycles: 30 s of 200 J at 20 kHz, 30-s intervals with manual shaking) and then diluted to 50 ml with deionized water. Depending on the viral counts, between 20 and 35 μl of the diluted sample was filtered onto 0.02-µm Anodisc filters (Whatman). The filters were then stained with SYBR Gold (Invitrogen) for 10 min in the dark and rinsed with 0.02-µm-filtered sterile water. For each filter, a minimum of 250 prokaryotic cells and viruses (differentiated from each other by their dimensions) were counted in at least 15 fields. Replicate measurements in selected samples showed that the average abundance was determined with an error of <10 %. Similar precision was obtained before in the determination of abundance of viruses and prokaryotes using the same procedure [25,41].

Viruses and prokaryotes in subpolar sediments

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Table 1. Basic characteristic of the sediment at the sampling site. The values in parentheses show the range (lowest and highest values) determined for five slices in each core Factor

HornWest

HornMid

HornEast

Sampling date

12 Aug 2010

12 Aug 2010

11 Aug 2010

Location

76º56.54′ N, 15º18.80′ E

76º57.67′ N, 15º41.26′ E

76º59.62′ N, 15º56.54′ E

Water depth (m)

171 m

135 m

135 m

Near-bottom salinity (PSU)

34.8

34.8

34.7

Near-bottom temp. (ºC)

1.9

1.9

1.3

Water content (%)

(42.5 to 54.7)

(43.5 to 58.1)

(39.8 to 56.1)

Sand fraction (%)

(14.8 to 22.0)

(9.2 to 18.3)

(6.7 to 19.4)

Silt fraction (%)

(78.0 to 85.2)

(81.7 to 90.8)

(80.6 to 93.3)

Mean (φ)

(5.45 to 5.59)

(5.55 to 6.14)

(5.89 to 6.18)

Sorting (σφ)

(1.52 to 1.67)

(1.46 to 1.77)

(1.44 to 1.76)

TC (%)

(2.5 to 2.6)

(2.4 to 2.9)

(2.3 to 2.4)

TOC (%)

(1.7 to 1.9)

(1.5 to 1.6)

(1.3 to 1.4)

N (%)

(0.19 to 0.22)

(0.16 to 0.19)

(0.13 to 0.14)

P (%)

(0.13 to 0.15)

(0.12 to 0.16)

(0.08 to 0.10)

TOC/N

(10 to 11)

(10 to 12)

(11 to 13)

TOC/P

(31 to 38)

(25 to 35)

(24 to 44)

N/P

(2.9 to 3.8)

(2.2 to 3.5)

(2.2 to 3.4)

δ CTC (‰)

(–17.6 to –17.1)

(–16.0 to –14.5)

(–14.6 to –14.4)

δ13CTOC (‰)

(–25.0 to –23.1)

(–24.8 to –24.3)

(–25.4 to –24.9)

δ15N (‰)

(2.1 to 4.9)

(3.1 to 5.2)

(2.6 to 3.4)

13

Sediment composition. Another sediment subsample was used to determine water content, measured as weight loss after drying at 60 °C for 24 h. This treatment removes interstitial water but not water bound in hydrates. Grain size composition (range: 0.01–2000 μm) was determined using a Malvern instrument Mastersizer 2000 equipped with a Hydro MU sample dispersion unit combined with a laser diffraction system.

Laser granulometry data were analyzed using GRADISTAT [4]. The concentrations of chlorophyll a and phaeophytin were determined fluorometrically after extraction of the pigments from freeze-dried sediment subsamples with 90 % acetone in the dark at 4 oC for 24 h [20].

Total carbon (determined as δ13CTC; TC), total organic carbon (δ13CTOC; TOC), and total nitrogen (with δ15N; N) were measured in subsamples using an elemental analyzer (Flash EA 1112 series coupled with the isotopic ratio mass spectrometer IRMS Delta V Advantage, Thermo Electron, Bremen, Germany). The freeze-dried sediment was weighted in two silver capsules (with 1-μg accuracy). One capsule was used to determine TC, δ13CTC, N, and δ15N. The other capsule was used to measure TOC and δ13CTOC by first soaking the sample portion in 2 M HCl in order to remove inorganic carbon species, followed by drying at 60 °C for 24 h.

Quality control standards (EA instruments calibration and recovery tests), purchased from Thermo Electron, were: acetanilide (C = 71.09 %, H = 6.71 %, N = 10.36 %, O = 11.84 %), atropine (C = 70.56 %, H = 8.01 %, N = 4.84 %, O = 16.59 %), and cyclohexanone 2–4 dinitrophenyl hydrazone (C = 51.79 %, H = 5.07 %, N = 20.14 %, O = 23.00 %). Additionally, certified reference materials (HEKAtech GmbH, Wegberg, Germany), consisting of environmental samples (including marine sediments) with an established concentration of analyzed chemical elements, were used to assess the accuracy of the analytical method. The average recovery for the standard and certified reference materials was 99.2 % for TC, 99.3 % for TOC, and 98.8 % for N; standard deviations were less than 0.5 %, 0.5 %, and 0.7 % for TC, TOC, and N, respectively. The results obtained for δ13CTC, δ13CTOC, and δ15N are given in the conventional delta notation versus Pee Dee Belemnite for δ13CTC and δ13CTOC and versus air for δ15N. Pure CO2 and N2 calibrated against an IAEA (International Atomic Energy Agency) standard (CO-8 and USGS40 for δ13C and N-1 and USGS40 for δ15N) were used to calculate stable isotope ratios. Standard deviations for replicate samples were less than 0.14 ‰, 0.12 ‰, and 0.17 ‰ for δ13CTC, δ13CTOC, and δ15, respectively. To measure phosphorus

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Wróbel et al.

Int Microbiol

48

Fig. 1. Sampling area and station locations at Hornsund fjord (Svalbard, Norway).

(P) concentrations, dry sediment samples (100 mg) in glass tubes were first mineralized in a temperature gradient from 100 to 300 °C with 1.6 ml of 10 N acid mixture (HClO4 and H2SO4, 1:3.85 molar ratio). The mineralized samples were diluted with Milli Q water and filtered through MN GF5 glass-fiber filters (Macherey Nagel, Düren, Germany). The filtrates were transferred into 100-ml flasks and neutralized with NaOH in the presence of phenolphthalein. The flasks were then filled up to 100 ml with Milli Q water, 2 ml of 0.02 M (NH4)6Mo7O24 (in water), and 0.5 ml of 0.11 M SnCl2 (in glycerine). After 10–12 min, P concentrations were measured colorimetrically. The estimated relative standard deviation of the measurement was <2.1 %.

Statistical analysis. The R package [http://www.r-project.org/] was used for the statistical analysis of the data, including a linear model for regression analysis and a one-tailed t-test to confirm the null hypothesis that measurements in the uppermost layer were based on a normal distribution estimated using the remaining measurements for the same core. The level of significance throughout the analysis was 0.05.

Results and Discussion The sediments collected at the three locations had similar general properties, consisting mostly of silt with an admixture of sand (Table 1). The sediment was poorly sorted,

with sorting values (σφ) characteristic of sediments originating from glacier material (Table 1). The arithmetic mean in the φ scale for all slices was >2 φ, which indicates that the sediments were transported as suspension. The TC, TOC, N, and P contents were relatively high, as observed before [24] and there was little variation between slices or sites. In general, the concentrations were higher in the outer fjord than towards the east. This likely reflects the combined higher production and greater sedimentation of organic matter in the west and supports previous observations of higher inorganic matter sedimentation in the central basin, resulting in a “dilution effect” of organic matter by mineral particles [26].

Based on the carbon/nitrogen molar ratios (around 10 in all samples, Table 1), the organic matter is mostly of marine origin and relatively freshly deposited. The slight increase in the TOC/N ratios towards the east (up to 13 at HornEast) suggests, however, the increased contribution of terrestrial organic matter in the bulk of sedimentary organic material along the fjord’s latitudinal axis [38,39]. This is supported by the δ13CTOC values, which decreased towards

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Viruses and prokaryotes in subpolar sediments

Fig. 2. Vertical distribution of viral and prokaryote abundance and concentrations of chlorophyll a and phaeophytin in Hornsund cores (Svalbard, Norway).

the east, from –23.1 ‰ at HornWest to –25.4 ‰ at HornEast (Table 1) [16,18,19,47,49–51]. A latitudinal trend was observed also for inorganic carbon concentrations (the difference between TC and TOC; Table 1): the highest values (1.1 %) occurred at HornEast and the lowest (0.7 %) at HornWest. The δ13CTC values also increased towards the east, consistent with an increasing input of terrestrial carbonates (e.g., from limestone formations [28]). By contrast, there were no readily apparent latitudinal trends for δ15N and TOC/P, which suggests similarities in the food web structure and in processes affecting P preservation in sediments, respectively [31,54].

While TC, TOC, N, and P showed little vertical variation, chlorophyll a and pheophytin concentrations varied

both vertically and horizontally. The concentration of chlorophyll a was higher in the uppermost layer at HornWest than at other stations, reflecting the fact that HornWest lies in the region with the highest primary production (J. Wiktor and M. Darecki, personal communication).We hypothesize that organic matter delivered to the sediment in the central basin originates in part from the outer fjord and that the phytoplankton undergoes a more pronounced degradation during delivery. At HornWest and HornEast the chlorophyll a concentrations were significantly higher in the uppermost layer than in the lower sediment layers (onesided t-test, P = 0.0029, and P = 0.0013, respectively, with the null hypothesis that measurements of all the layers have the same distribution). At all stations, pheopigment

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concentrations were higher in the uppermost layer than in deeper sediment layers (HornWest: P = 0.033, HornMid: P = 0.010, HornEast: P = 0.011). The HornEast data could be fit to a linear model relating pigment concentrations with sediment depth (chlorophyll a: P = 0.0018, phaeophytin: P = 0.0063). These observations suggest that following its depostion, organic matter undergoes further degradation. High concentration of chlorophyll a at 4.5 cm sediment depth in the core collected at HornMid may correspond to the burial of the organic matter produced during the extremely warm 2005–2006 period in the European Arctic [7], quickly overlaid with the sediment that is deposited at about 1 cm/yr in this region [26].

In general, the prokaryote and viral counts determined in Hornsund (Fig. 2) were low (around 108 cells/viruses g–1) but still within the range reported for marine sediments [8]. Note that the VPRs in this study were very low (0.2– 0.6, with the exception of the uppermost layer at HornMid, Fig. 2). The results of our linear regression analysis of the relationship between prokaryote and viral density indicated that the data from all cores could be described using a linear model (P = 0.015). For all stations, prokaryote density was significantly higher in the uppermost layer than deeper in the sediment (one-sided t-test, HornWest: P = 0.034, HornMid: P = 0.0022, HornEast: P = 0.025), as was the case for viruses (HornWest: P = 0.012, HornMid: P = 0.000022, HornEast: P = 0.023). In the uppermost layer at HornMid, the VPR significantly differed from the ratio determined in the other layers from this core (P = 0.000082) and indeed from the ratios of all the other samples (P = 7.405 × 10–14).

Prokaryotes and viruses in surface sediments originate either from production within the sediments or from sinking particles that act as vectors for the vertical transport of pelagic microorganisms. Previous studies suggested that the inherent production of viruses is the dominant source of benthic viruses in coastal sediments [29,40] and that higher benthic viral production corresponds to higher benthic productivity and thus to the trophic state of the sediment [40]. In Hornsund, however, viral and prokaryote abundances were uncoupled from the sediment trophic state. In other words, the higher viral and prokaryote abundances in the surface sediments at HornMid than in the more productive HornWest did not correspond with the relatively low sediment chlorophyll a concentrations and the low chlorophyll/pheophytin ratios at HornMid. We speculate that the high abundance of prokaryotes and viruses in the central basin reflects the 10-fold higher sedi-

Wróbel et al.

mentation of particles which may scavenge viruses and prokaryotes from the water column and contribute significantly to their abundance in the surface sediment at this station. This hypothesis is supported by the rapid decrease in viral and prokaryote abundances with sediment depth at HornMid. This decrease suggests that the imported prokaryote populations are not maintained in the sediment due to limited substrate availability and prokaryote growth, and rapidly decay.

The relatively low VPR in the subpolar sediments might stem from very low viral production or from physicochemical conditions in these environments that enhance viral decay. Viral production depends on the growth and turnover rates of prokaryote cells [55], which are affected by abiotic and biotic factors. Conditions in which prokaryotes grow slowly, e.g., low temperature or nutrient limitation, may limit viral production and lysogeny, and favour pseudolysogeny and chronic infection [8,11].

Long-term data on the composition of the benthic communities in Hornsund [32] indicate a pronounced difference between stations of the outer fjord (close to HornWest) and central basin (HornMid and HornEast), with higher species diversity and a higher degree of bioturbation in outer fjord sediments. By contrast, the total abundance of infauna in the sediment is similar for the outer fjord and the central basin (about 600 individuals 0.1 m–2, the data here and further are from the 2007 sampling campaign), but the abundance of subsurface deposit feeders (burrowers) in the outer fjord sediments is about two times higher than in those of the central basin (over 300 ind. 0.1 m–2 vs. about 150 ind.

0.1 m–2, respectively). Outer fjord sediments are dominated

by Leitoscoloplos mammosus and Cossura longocirrata (15 % and 8.5 % of total infaunal abundance, respectively), which are non-selective deposit-feeders that burrow freely through the sediment [21].

A decrease in viral abundances with sediment depth has been reported for a number of temperate and subtropical sediments [10,13,30,41,42]. This decline generally reflects the high prokaryote activity in the benthic surface layers in response to the input of organic matter from the water column. The higher degree of bioturbation in outer fjord sediments might be responsible for the more even depth distribution of viruses in our HornWest core (Fig. 2) than in the cores we collected in the central basin, in which viral and prokaryote densities clearly decreased with sediment depth. Although further analysis of sediments in other fjords is necessary, we propose that differences in the distribution of viruses and prokaryotes in the Hornsund sedi-

Viruses and prokaryotes in subpolar sediments

ments reflect functional differences in the benthic invertebrate community along the west-east axis, i.e., a decrease in the abundance of subsurface burrowers and a corresponding increase in the abundance of mobile surface deposit feeders (from about 100 ind. 0.1 m–2 in the outer fjord to over 300 at stations close to HornEast). According to our hypothesis, the higher activity of burrowers in the outer fjord (HornWest) results in the delivery of viral particles and prokaryotes to lower sediment layers. Less bioturbation at HornMid is supported by the presence of a chlorophyll-rich layer at 4.5 cm, perhaps corresponding to the warm 2005–2006 period. Finally, at HornEast, still more degraded organic matter and presumably even lower bioturbation rates account for the almost continuous decrease in prokaryote and viral counts and in pigment concentrations with sediment depth.

Higher abundances of prokaryotes than of viruses (VPR <1.0) were previously recorded in oligotrophic marine sediments in the temperate zone [11,37] and in inland sediments [2,22,35–37]. In nutrient-poor deep-sea sediments, low rates of prokaryotic metabolism were suggested as an explanation for the low virus-bacteria ratios [11]. We found virus-bacteria ratios similar to those of deep sea sediments, despite the higher organic carbon contents. The low VPRs in polar environments might be indicative of prokaryotic activity that is too low (e.g., with temperature as a limiting factor for prokaryotic metabolism) to maintain a high level of virus production [34,43]. Nonetheless, viruses may still be important in controlling prokaryote populations in subpolar sediments. In fact, there is evidence of high virus-induced prokaryote mortality (>40 %) even in oligothropic sediments, where the VPR is below 1.0 [37]. Further research will contribute to our ability to quantify the role of virus-induced prokaryote mortality in high-latitude sediments.

Acknowledgements. This study was funded by the Polish Ministry of Science and Higher Education (AOD/W6/2008 and AODP/09/2010/0 to BW) and a grant from the Danish Council for Independent Research to MM. We are grateful to Mirosław Darecki, Ilona Goszczko, Witold Szczuciński, Joanna Przytarska, Jan Marcin Węsławski, Józef Wiktor, and the crew of the r/v Oceania for their help during sampling, laboratory procedures, and/or discussion of the results.

Competing interest: None declared

References 1. Ambrose WGJr, Renaud PE (1995) Benthic response to water column productivity patterns: evidence for benthic-pelagic coupling in the Northeast Water Polynya. J Geophys Res 100:4411-4421

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Limnol Oceanogr 57:1732-1742 44. Piepenburg D, von Dorrien CF, Schmid MK, Chernova NV, Neyelov AV, Gutt J, Rachor E, Saldanha L (1996) Megabenthic communities in the waters around Svalbard. Polar Biol 16:431-446 45. Piwosz K, Walkusz W, Hapter R, Wieczorek P, Hop H, Wiktor J (2009) Comparison of productivity and phytoplankton in a warm (Kongsfjorden) and a cold (Hornsund) Spitsebergen fjord in midsummer 2002. Polar Biol 32:549-559 46. Renaud PE, Wlodarska-Kowalczuk M, Trannum H, Holte B, Weslawski JM, Cochrane S, Dahle S, Gulliksen B (2007) Multidecadal stability of benthic community structure in a high-Arctic glacial fjord (van Mijenfjord, Spitsbergen). Polar Biol 30:295–305 47. Rullkötter J (2006) Organic matter: The driving force for early diagenesis. In: Schulz HD, Zabel M (eds) Marine geochemistry. Springer-Verlag, Berlin, Germany, pp 125-206 48. Siem-Jørgensen M, Glud RN, Middelboe M (2008) Viral dynamics in a coastal sediment: Seasonal pattern, controlling factors and relations to the benthic-pelagic coupling. Mar Biol Res 4:165-179 49. Struck U, Emeis K-C, Voss M, Christiansen C, Kunzendorf H (2000) Records of Baltic Sea eutrophication in δ13C and δ15N of sedimentary organic matter. Mar Geol 164:157-171 50. Voss M, Emeis K-C, Hille S, Neumann T, Dippner JW (2005) Nitrogen cycle of the Baltic Sea from an isotopic perspective. Global Biogeochem Cycles 19:GB3001 51. Voss M, Larsen B, Leivuori M, Vallius H (2000) Stable isotope signals of eutrophication in coastal Baltic Sea sediments. J Mar Systems 25:287-298 52. Warwick RM (1993) Environmental impact studies on marine communities: pragmatical considerations. Aust J Ecol 18:63-80 53. Warwick RM, Emblow CS, Féral J-P, Hummel H, van Avesaath P, Heip CHR (2003) European marine biodiversity research sites: report of the European Concerted Action BIOMARE. Netherlands Institute of Ecology–Center for Estuarine and Marine Ecology, Yerseke, the Netherlands, 135 pp 54. Westman P, Borgendahl J, Bianchi TS, Chen N (2003) Probable causes for cyanobacterial expansion in the Baltic Sea: role of anoxia and phosphorus retention. Estuaries 26:680-689 55. Wommack KE, Colwell RR (2000) Virioplankton: viruses in aquatic ecosystems. Microbiol Mol Biol Rev 64:69-114

RESEARCH ARTICLE International Microbiology (2013) 16:53-62 doi:10.2436/20.1501.01.180 ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

Dark fermentation: isolation and characterization of hydrogen-producing strains from sludges Haifa Rajhi,1 Mónica Conthe,1 Daniel Puyol,2§ Emiliano Díaz,3 José Luis Sanz1* Department of Molecular Biology, Autonomous University of Madrid, Madrid. Spain. 2Chemical Engineering Section, Autonomous University of Madrid, Madrid, Spain. 3Mygen Laboratory, Cantoblanco, Spain

1

Received 15 December 2012 · Accepted 15 February 2013 Summary. To improve bacterial hydrogen production, ten hydrogen-producing strains belonging to Clostridium spp.

were isolated from various sludges under low vacuum. Hydrogenogenesis by dark fermentation in batch cultures of these strains was optimal at about 35 ºC and an initial pH of 6.5, which for all strains gradually dropped to ca. pH 4 during the fermentation. Clostridium roseum H5 and C. diolis RT2 had the highest hydrogen yields per total substrate (120 ml H2/g initial COD). Substrate consumption alone by C. beijerinckii UAM and C. diolis RT2 reached 573 and 475 ml H2/g consumed COD, respectively. Butyric acid fermentation was predominant, with butyrate and acetate as the major by-products and propionate, ethanol, and lactate as secondary metabolites. The acetate:butyrate ratios and fermentation pathways varied depending on the strains and environmental conditions. Hydrogenogenesis was studied in greater detail in C. saccharobutylicum H1. In butyric acid fermentation by this representative strain, acetoacetate was detected as an intermediate metabolite. Hydrogenogenesis was also analyzed in an enrichment culture, which behaved similarly to the axenic cultures. [Int Microbiol 2013; 16(1):53-62]

Keywords: Clostridium spp. · hydrogen production · dark fermentation · pH and temperature optimization · fermentation pathways · kinetic glucose degradation

Introduction Environmental pollution resulting from the use of fossil fuels together with the world’s increasing energy demand have stimulated the search for renewable and environmentally friendly sources of energy. Hydrogen (H2) is a promising alternative energy carrier: it is renewable, with a high Corresponding author: J.L. Sanz Departamento de Biología Molecular Universidad Autónoma de Madrid 28049 Madrid, Spain Tel. +34-914974303. Fax +34-914978300 E-mail: joseluis.sanz@uam.es

*

Current address: Department of Chemical and Environmental Engineering, University of Arizona, Tucson, AZ, USA

§

energy yield. It is also clean, producing only water when it burns, unlike other renewable energy sources such as methane [33]. Furthermore, its use in hydrogen-fuel cells makes it a promising alternative to hydrocarbons to power land vehicles. Hydrogen is currently obtained by thermal cracking (reformation) of natural gas and the electrolysis of water, both of which are energy-consuming processes.

In contrast to these physico-chemical methods, biological processes using renewable sources for H2 production, i.e., photo-fermentation or dark fermentation, are economical and environmentally friendly [6]. Photo-fermentation requires sunlight and is performed by algae, cyanobacteria, and anaerobic photosynthetic bacteria. However, despite its high yields of H2 , photo-fermentation has several drawbacks, such as the need for light energy and thus the diffi-

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culty of designing efficient photo-reactors [8]. Dark fermentation involves the conversion of organic substrates to H2 by anaerobic microorganisms, including different species of Clostridium, syntrophobacteria, and enteric bacteria such as Enterobacter. The advantages of dark fermentation over other processes include better process economy because of lower energy requirements, process simplicity, higher rates of hydrogen production, and the use of inexpensive wastes as raw materials.

Clostridium species are the most significant microorganisms in anaerobic H2 dark fermentation [12,32,36].

These bacteria ferment sugars or proteins to yield organic acids (mainly acetate, propionate, butyrate, and lactate), alcohols (ethanol), carbon dioxide, and molecular H2 (also referred to as biohydrogen, specifiying its biological origin).

The production of biohydrogen by dark fermentation (hydrogenogenesis) is limited by the fact that the oxidation of reduced pyridine nucleotides and ferredoxin associated with H2 formation is inhibited by high partial pressures of H2 (PH2) [2,13]. Moreover, at high PH2,, the metabolism of Clostridium switches from acid formation (acidogenesis) to solvent formation (solventogenesis) such that H2 is no longer released [35]. Consequently, for efficient hydrogenogenesis, the prevention of H2 accumulation (i.e., high PH2) in the headspace and liquid medium of the dark fermentation system is of utmost importance. Most studies, disregarding this fact, have focused on systems in which the H2 produced is allowed to accumulate. However, in an increasing number of studies continuous H2 removal was shown to significantly increase hydrogen yields. For example, experiments in which the fermenters are flushed with dinitrogen gas have increased H2 yields by 1.5- to 1.7fold [17,27]. A recent study reported an impressive 11-fold increase in H2 yield during dark fermentation, achieved by the continuous removal of H2 from the medium using a fuel-type electrochemical cell [2]. Apart from maintaining low PH2, other factors must be optimized for efficient dark fermentation, particularly the inoculum [15,22], the temperature [20,34], the pH [14,18,34], and the substrate, all of which are critical in defining end-product formation and thus H2 production.

In this study, we developed an innovative approach— applying a low vacuum in the headspace of the fermentation system, thus maintaining low PH2 and preventing product inhibition—for the isolation of H2-producing strains from various sources. As part of our efforts to optimize the system, ten hydrogen-producing Clostridium strains were isolated. Data on the effect of pH and temperature on H2

rajhi et al.

production by the isolated strains and an enrichment culture are presented and discussed. Additionally, we analyzed intermediary products of H2-producing fermentations and conclude this work with a discussion of the respective metabolic pathways and reaction kinetics. Our results provide valuable information for the optimization of dark fermentation and the achievement of efficient hydrogen production.

Materials and methods Sources of microorganisms. The following sources were used as inocula for the isolation of hydrogen-producing bacteria: (i) granular sludge from a full-scale upflow anaerobic sludge bed (UASB) reactor treating wastewater from a brewery (Mahou SA, Alovera, Spain); (ii) sludge from an anaerobic digester treating municipal solid wastes (Madrid, Spain); (iii) activated sludge from an aerobic domestic wastewater treatment plant (Universidad Autónoma de Madrid [UAM], Cantoblanco, Madrid, Spain); (iv) sediments from an acidic river (Río Tinto, Huelva, Spain).

Culture media. The bacteria were cultured in synthetic (MR) medium containing 280 mg NH4Cl/l, 328 mg K2HPO4/l, 100 mg MgSO4/l, 500 mg NaHCO3/l, and 2 g sucrose/l, 1 g meat extract/l, 500 mg yeast extract/l as carbon source, and 1 ml of micronutrient solution [31]. The initial pH was adjusted to 7.5. Solid media were prepared by adding 15 g bacteriological agar/l to MR medium.

Preparation of enrichment cultures and isolation of axenic cultures. Prior to isolating the strains, we prepared enrichment cultures under low vacuum in 120-ml glass serum bottles containing 60 ml of medium. Anoxic conditions were produced by flushing the headspace with N2:CO2 (80:20) and adding l-cysteine (40 mg/l). The medium was inoculated with 1g of homogenized sludge, and then incubated under partial vacuum at 30 °C for 30 days. The nutrients were renewed weekly. Cultures producing significant amounts of H2 were inoculated onto MR agar plates and incubated at 30 °C for 72 h. Single colonies were selected and grown in MR in order to establish pure cultures of the H2-producing strains (see Phylogenetic analysis, next). An enrichment culture (EC), obtained from granular sludge after 5 months of incubation under the conditions described above and with periodically renewed medium was monitored along with the axenic cultures.

Phylogenetic analysis. Single colonies from the MR agar plates were inoculated into 25-ml serum bottles containing 5 ml of MR and incubated at 37 °C overnight. Cells from bottles with high H2 yields were collected for sequencing. One ml of liquid culture was centrifuged (10,000 rpm, 15 min) and the pellet was washed with 1 ml of phosphate buffered saline, centrifuged (10,000 rpm, 15 min), and re-suspended in 100 µl of distilled water. The suspension was heated at 94 °C for 10 min and centrifuged (15,000 rpm, 5 min). Five µl of the resulting supernatant was PCR-amplified using the primers 27f and 1492r [21]. The amplicons were assembled and the consensus sequence corrected manually for errors using DNA Baser 3.0. The sequences were compared to those in the 2011 GenBank database of NCBI [http://www.ncbi.nlm.nih.gov/] using the Basic Local Alignment Search Tool (BLAST) algorithm and the 2011 version of the tool Classifier of the Ribosomal Database Project [http://rdp.cme.msu.edu/].

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Dark fermentation

Optimization of pH and temperature. Tests for the optimization of pH and temperature were conducted in 120-ml serum bottles containing 20 ml of MR. Each bottle was inoculated to an initial optical density (OD660) of 0.001 with one of the axenic cultures grown to exponential phase. The conditions were those described in “Culture media” except for the variable tested (pH or temperature). The tested pH values were 5, 5.5, 6.5, and 7.5.

The temperature optimization experiment was carried out at 25, 30, 35, and 40 °C. All tests were done in triplicate. Hydrogen production, OD, and pH were analyzed after 14, 24, and 48 h.

Metabolic and kinetic studies. Studies of the metabolic pathways and growth kinetics of the strains and enrichment culture were carried out using glucose (4 g/l) as the sole carbon source, added to the mineral medium used in MR medium. Batch tests were followed over 2 days until glucose depletion. The main organic metabolites as well as pH, H2, CO2, and total suspended solids (TSS) were monitored periodically. All tests were done in triplicate.

Analytical methods. Hydrogen and CO2 were quantified by gas chromatography using a Bruker 450-GC coupled with a thermal conductivity detector (TCD) and a Varian CP2056 0.6 m ×1/8′′ Ultimetal Cromosorb GHP 100–120 mesh column in bypass. The temperature of the injection, TCD chambers, and oven was maintained at 150, 200, and 50 °C, respectively. Nitrogen was used as the carrier gas in the column at a flow rate of 25 ml/min. Volatile fatty acids (VFAs) were quantified by high-performance liquid chromatography coupled with a refraction in-

dex detector (HPLC-RID) (Varian Prostar 350 RID, Agilent) using a sulfonated polystyrene resin in the protonated form (67H type) as the stationary phase (Varian Metacarb 67H 300 mm) and sulfuric acid (0.25 mM in milliQ water) as the mobile phase at a flow rate of 0.8 ml/min. The column temperature was set at 65 °C. Non-common metabolites were identified by HPLC coupled to a diode array detector (Varian DAD 330) and a triple quadrupole ion trap mass spectrometry detector (HPLCDAD-MS) with positive and negative electrospray ionization (ESI) and atmospheric-pressure chemical ionization (APCI) (Varian 1200L). The same 67H column described above served as the stationary phase and 0.5 M formic acid as the mobile phase at a flow rate of 0.6 ml/min. The column temperature was set at 65 ºC. All the other analyses were done according to standard methods [3].

Results and Discussion Isolation of hydrogen-producing strains. Ten H2-producing strains were isolated from the four sources listed in Materials and methods: three (H1, H5, H17) from the UASB reactor, four (R4, R6, R12, R14) from the anaerobic digester, one from the activated sludge of a domestic wastewater treatment plant (UAM), and two from sedi-

Table 1. Hydrogen-producing strains isolated from different sources Query length

Species of Clostridium with higher homology

Source*

Similarity (%)

H1

1170

C. saccharobutylicum C. acetobutylicum C. saccharoperbutylacetonicum

1

99

H5

1293

C. roseum C. beijerinckii

1

100

H17

1180

C. butyricum

1

100

R4

1093

C. butyricum

2

99

R6

1095

C. butyricum

2

96

R12

1328

Clostridium sp.

2

96

R14

1335

Clostridium sp.

2

96

RT1

1342

C. beijerinckii

4

100

RT2

1337

C. diolis C. beijerinckii C. acetobutylicum

4

100

UAM

1371

C. beijerinckii C. diolis C. acetobutylicum

3

100

Strain isolated

55

*1: Anaerobic granular sludge from a UASB reactor treating brewery wastewater.

2: Sludge from an anaerobic digester of municipal solid wastes.

3: Activated sludge from an aerobic domestic waste treatment plant.

4: Anaerobic sediments from a river.

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Fig. 1. Effect of pH (A) and temperature (B) on hydrogen production by representative strains. The results are presented as percentages relative to maximum hydrogen production by each strain.

ments from the Río Tinto (RT1, RT2). All of the isolates belonged to the genus Clostridium (Table 1). The 16S rRNA gene sequences have been deposited in the GenBank database under accession number JX575125–JX575134.

The four sludge samples studied proved to be appropriate sources of hydrogen-producing bacteria. Indeed, diverse clostridia and syntrophobacteria have repeatedly been isolated from granular and other kinds of sludge, accounting for their frequent use as inocula in studies focusing on hydrogen production [10,12,22]. The sludge and enrichment cultures from which the strains were isolated were submitted to low vacuum, which allowed continuous removal of the gaseous products from the headspace, thus keeping the PH2 low. This innovative method boosts the growth of H2-producing microorganisms by avoiding the product inhibition caused by high PH2.

Although hydrogen-producing bacteria are diverse [16,18], the medium used in these experiments, rich in sugars and proteins, may have selected Clostridium spp., which are typical H2 producers associated with the fermentation of carbohydrates [4,12,37] and proteins [30] to acetate, butyrate, carbon dioxide, and organic solvents [29].

Effect of pH and temperature. Figure 1A shows the production of H2 at different initial pH settings. In most cases, pH 6.5 was optimal for H2 production. Strains H1

(not shown in the figure) and H5 (no significant differences between pH 5.5 and 6.5), R12 (optimal pH of 5.5), and H17 (pH 7.5) were the exceptions to this general trend. In several reports, pH 5.5–6.5 was reported to be the optimal range for H2 production [7,19,26]. Indeed, in at least two studies evaluating the effect of pH on fermentative biohydrogen production by isolated Clostridium species H2 production rates and yields were shown to peak at an initial pH 6 [11,24]. However, other studies have not found that the initial pH profoundly affects H2 production [9], which concurs with our results for Clostridium R12 and for the enrichment culture. In our system, the initial optimal pH for C. butyricum H17 (7.5) was close to that reported for C. butyricum ATCC 19398, C. acetobutyricum M12, and C. tyrobutyricum FYa102, all of which produced maximum amounts of hydrogen at pH 7.2 [25].

The effect of temperature on hydrogen production is shown in Fig. 1B. With the exception of strains H17 (optimum production at 25 °C) and R12 (30 °C), the highest H2 production was recorded at 35 °C. Note that the effect of temperature in the range studied was limited for some strains (H17), yet dramatic for others (R4, R12). The highest temperature tested (40 ºC) had a negative effect on the growth of all ten strains, and especially on the growth of isolates from the municipal solid waste digester and the enrichment culture. Differences in the behavior of hydro-

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Fig. 2. Growth curve (dot), hydrogen production (dash), and pH trend (solid) for strains H1, H5, and R12 during batch fermentation at 30 °C and pH 6.5 during the first 48 h.

gen-producing strains due to temperature had been described previously [1,10,23].

General trends and hydrogen yields during batch fermentation. Figure 2 shows the time-course of hydrogen production by three of the isolated strains growing in MR as well as the pH and OD of the respective cultures. In most cases (i.e. H5, R12) H2 production mirrored the growth curve (determined by OD) of each strain, and was accompanied by a decrease in the pH of the culture due to the accumulation of acids. In some case (i.e., for strain RT2), the apparent growth rate is low over the first 24 h, while the H2 production rate is high. From this moment on, probably due to the low pH, the cell number increases but stop producing H2. A summary of the results obtained in the fermentation studies is provided in Table 2.

The hydrogenogenesis potential ranged from 120 ml/[g of initial chemical oxygen demand (CODi)] for C. roseum H5 and C. diolis RT2 to nearly 70 ml/g CODi for C. butyricum H17 and R4. H2 production levels of 140 ml/g starch by C. pasteurianum [26], 150 ml/g sucrose by Clostridium sp. [34], 134 ml/g molasses by a mixed culture [26], and almost 180 ml H2 per g maltose [39] and g lactose [11,39] have been reported, indicating that yields are lower when complex wastes are used as substrates. In other

studies, much lower amounts of H2 were achieved: 2.7 ml H2/g COD of food waste by C. beijerinckii KCTC [19] and 9.1 ml H2/g of raw sludge by raw sludge seed inocula [5].

Some authors consider only the COD consumed when calculating H2 yields. Tang et al. [34], for example, reported a yield of 319 ml H2/g COD of cattle waste consumed. The yields obtained in our study were mostly higher than 300 ml H2/g COD consumed, or even above 500 ml H2/g COD consumed for C. diolis RT2, C. beijerinckii UAM, and the enrichment culture (Table 2). It is not easy, however, to compare hydrogen production in different experiments because dark fermentation is influenced by many factors, including the type of inoculum, the pH, the temperature, and the nature of the substrate. The high hydrogen production rates obtained in our experiments can perhaps be ascribed to the use of the complex MR medium, which might have promoted the growth of both carbohydrate and protein fermenters.

Fermentation products. The decrease in pH observed during the batch experiments may have resulted from the accumulation of VFAs during acid fermentation by the strains. The major by-products accumulated were the VFAs butyrate and acetate (Table 3). When grown on MR medium, strains H1, H5, RT1, RT2, and UAM mainly

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Table 2. pH, OD, COD, and hydrogen produced at the end of experiments for the isolates and the enrichment culture Strains of Clostridium

Final pHa

Final OD

H2 (ml)b

H2 (ml/g CODi)c

H2 (ml/g COD removed)

C. saccharobutylicum H1

4.39 ± 0.04

0.75 ± 0.05

7.3 ± 3.0

76.4 ± 31.6

251.7

C. roseum H5

4.16 ± 0.02

1.09 ± 0.02

11.5 ± 0.5

120.3 ± 5.2

396.6

C. butyricum H17

4.35 ± 0.01

0.23 ± 0.09

6.4 ± 0.2

68.0 ± 2.1

300.9

C. butyricum R4

4.28 ± 0.06

0.66 ± 0.03

6.4 ± 0.1

69.0 ± 1.0

336.7

C. butyricum R6

4.00 ± 0.01

0.70 ± 0.04

7.6 ± 0.6

79.5 ± 6.3

263.9

Clostridium sp. R12

3.98 ± 0.05

0.67 ± 0.05

4.2 ± 0.1

51.3 ±1.0

175.0

Clostridium sp. R14

4.02 ± 0.03

0.66 ± 0.05

6.8 ± 0.2

77.4 ± 2.1

345.8

C. beijerinckii RT1

4.23 ± 0.02

0.52 ± 0.04

9.3 ± 0.7

99.4 ± 7.3

341.6

C. diolis RT2

4.12 ± 0.02

0.88 ± 0.05

11.2 ± 0.1

120.3 ± 1.0

475.2

C. beijerinckii UAM

4.17± 0.01

0.88 ± 0.03

7.7 ± 0.1

90.0 ± 1.0

573.3

Enrichment culture

4.46 ± 0.04

1.02 ± 0.01

10.5 ± 1.0

114.1 ± 10.1

500.0

Initial pH: 6.5.

Serum bottles containing 20 ml of medium. c Initial COD varied from 4.6 to 4.78 g/l. a b

carried out butyrate fermentation. The presence of alcoholic and propionic fermentation can be inferred for strains H1 and H5 cultured on meat extract as a carbon source and glucose, respectively, based on the production of acetate, propionate, and ethanol as final products. Moreover, in strain H1, the alcoholic fermentation pathway was apparently used for glucose fermentation. In strains R12 and R14, lactate was the second most abundant fermentation by-product (data not shown), suggesting heterolactic or mixed-acid fermentation.

The effect of pH and temperature on the fermentation end-products was also studied in enrichment culture. In general, both had limited effects, except at an incubation temperature of 40 °C. At this temperature, H2 was not released, sucrose was not fully consumed, and metabolism shifted to lactic acid fermentation (data not shown). With respect to pH, the butyrate:acetate ratio increased from 1.8 to 4.6 at an initial pH range of 7.5–5.5, with the amounts of minor products (ethanol and propionate) remaining constant (data not shown).

Metabolic and kinetic studies. Among our isolates, Clostridium saccharobutylicum H1 was selected as a

representative strain and its metabolism and kinetics were studied in cultures containing 4 g glucose/l. Figure 3 shows the evolution of glucose and its main degradation products over time (Fig. 3A) and the evolution of pH, glucose, and biomass concentration (expressed as volatile suspended solids [VSS]) (Fig. 3B). Glucose evolution is presented in both figures to allow comparisons of glucose consumption and biomass production. Both the CO2 and the H2 values (mmol) were corrected for pressure. The results showed that butyrate fermentation was the predominant process involved in glucose fermentation, with 0.1 mmol of butyrate and 0.06 mmol of acetate produced during the fermentation of 0.48 mmol of glucose.

Trace concentrations of lactate were detected during the first hours of the experiment, suggesting the appearance of residual lactic fermentation. However, lactate was consumed during the course of the experiment, consistent with the anaerobic oxidation of lactate to acetate. The accumulation of VFAs led to a drop in pH to around 4 (Fig. 3B). Similar amounts of H2 and CO2 (approximately 2 mmol/mmol glucose consumed) were produced, which is in accordance with the maximum H2/glucose ratio reported thus far [28].

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Table 3. Concentrations of fermentation end products (expressed in mM) Strain

Medium

Acetate

Butyrate

Ethanol

Glucose

Lactate

Propionate

Sucrose

H1

Complex medium MR

7.6

13

2.7

2

–

–

–

H1

With 11.1 mM glucose

4.9

–

18.2

1.6

–

–

H5

Complex medium MR

4.2

18.8

–

1.4

–

0.7

–

H5

With 2g/l meat extract

7.2

1.7

4.6

–

–

3.3

–

RT1

Complex medium MR

11

14

–

1.2

–

1.4

–

UAM

Complex medium MR

5.5

13.4

–

–

1.9

2.2

6

Glucose consumption was linked to biomass growth, which indicated the use of a high fraction of glucose by the strains during anabolic processes. Both glucose consumption and biomass growth can be modeled by the following equation system, simplified from the general Monod model: dG = –k · X 1 Yx/s dt

(1)

dG dX = –Yx/s · dt dt

(2)

Int Microbiol

where k1 is the pseudo-first order kinetics constant (liter per h per mmol glucose), YX/S is the biomass yield factor (mg VSS/

mmol glucose), and X and G are the biomass (mg VSS/l) and glucose concentrations (mM), respectively. Equations (1) and (2) were integrated by the Episode numerical method for Stiff systems, with the initial conditions t = 16, X = 57.8, and G = 31.75, by means of a non-linear least squares minimization of the error, using a simplex algorithm followed by a Powell minimization algorithm (Micromath Scientist 3.0). Fitting glucose and biomass concentrations to this model resulted in the following values for the fitting parameters: k1 = 0.00130 ± 0.00007 liter per h per mmol glucose, Y = 13.26 ± 0.32 mg VSS produced per mmol glucose consumed, and R2 = 0.997.

The fitting curves are depicted in Fig. 3B. The model accurately describes both biomass growth and glucose consump-

Fig. 3 (A) Time-evolution of glucose (closed triangles), hydrogen (closed squares), CO2 (closed circles), acetate (open triangles) and butyrate (open circles). (B) Time evolution of pH (closed circles), glucose (open squares), and biomass expressed as VSS (closed squares) during the metabolic and kinetic studies. Continuous lines are model fittings.

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rajhi et al.

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Fig. 4. Representative chromatograms of samples from the metabolic study of strain H1. (A) HPLC-RID chromatogram at 18 h. (B) HPLCDAD chromatogram at 24 h. (C) HPLC-MS profile of peak at 14.7 min from the HPLC-DAD chromatogram. (D) HPLC-MS chromatogram of the 102 g/mol fraction of the HPLC-DAD chromatogram, showing positive (up) and negative (down) ionization of the sample.

tion. Additionally, from these data, a duplication period of 1.87 h could be estimated.

Considering the approximate composition of biomass to be C4H7O2N [38], which corresponds to approximately 50 % molar/molar C and 31 % m/m O, the C and O mass balances were not closed at the end of the experiment. This suggests the presence of other metabolites not considered in the glucose fermentation processes. The HPLC-RID chromatogram at 18 h is presented in Fig. 4. A peak at 11.5 min was detected, indicating that only one other metabolite was produced during glucose fermentation, in addition to those already mentioned. The sample was analyzed by HPLC-DAD-MS at 24 h (Fig. 4B). The same distribution of metabolites was observed, with the detection of an unknown peak at 14.7 min. This peak was analyzed individu-

ally by MS, which showed that the predominant molecular mass was 102 g mol–1 (Fig. 4C). The ionic profiles indicated that the compound is negatively ionizable, because no cations were detected at the cationic 103 g mol–1 analysis whereas a significant peak was detected at the anionic 101 g mol–1 analysis (Fig. 4D). Carbon and oxygen mass balances were carried out for all of the samples, and an average C to O ratio of 1.25 ± 0.05 mol C per mol O was calculated for the non-closed mass balance. This result combined with the molecular mass determined in the HPLC-MS analysis (102 g mol–1) suggested that the unknown compound has four C atoms and three O atoms. We therefore propose that the unknown metabolite corresponds to acetoacetic acid (C4H6O3), an intermediate in butyric acid fermentation that usually combines with CoA before

Dark fermentation

butyric acid production. This compound was previously detected in a butyric acid fermentation under pH conditions unfavorable for butyric acid production [13]. Microorganisms adopt the strategy of excreting acetoacetate during butyric acid production when growth conditions again become favorable [13].

Acknowledgements. The present work was supported by grants from the Spanish Ministry of Science and Innovation (MICCIN) to J. L. Sanz (PNICDIT, CTM2009-10521) and J.L. Sanz and E. Diaz (INNPACTO, IPT-310000-2010-35). H. Rajhi thanks the Spanish Institute of International Cooperation for his fellowship. M. Conthe thanks the Autonomous University of Madrid for her grant for research initiation. D. Puyol thanks the Spanish Ministry of Education, the Spanish Fulbright Commission, and the American Council for International Exchange of Scholars for his Fulbright scholarship.

Competing interests. None declared.

References 1. Alalayah WM, Kalil MS, Kadhum AAH, Jahim JM, Alauj NM (2009) Effect of environmental parameters on hydrogen production using Clostridium saccharoperbutylacetonicum N1-4 (ATCC 13564). Am J Environ Sci 1:80-86 2. Ananyev GM Skizim NJ, Dismukes GC (2012) Enhancing biological hydrogen production from cyanobacteria by removal of excreted products J Biotechnol 162:97-104 3. APHA AWWA WPCF (1998) Standard methods for examination of water and wastewater, 20th ed, Washington, DC, USA 4. Cai G, Jin B, Saint C, Monis P (2010) Metabolic flux analysis of hydrogen production network by Clostridium butyricum W5: Effect of pH and glucose concentrations. Int J Hydrogen Energy 35:6681-6690 5. Cai M, Liu J, Wei Y (2004) Enhanced biohydrogen production from sewage sludge with alkaline pretreatment. Environ Sci Technol 38:3195-3202 6. Chang JS, Lee KS, Lin PJ (2002) Biohydrogen production with fixed-bed bioreactors. Int J Hydrogen Energy 27:1167-1174 7. Chen WM, Tseng ZJ, Lee KS, Chang JS (2005) Fermentative hydrogen production with Clostridium butyricum CGS5 isolated from anaerobic sewage sludge. Int J Hydrogen Energy 30:1063-1070 8. Chong ML, Sabaratnam V, Shirai Y, Hassan MA (2009) Biohydrogen production from biomass and industrial wastes by dark fermentation. Int J Hydrogen Energy 34:3277-3287 9. Fan Y, Li C, Lay JJ, Hou H, Zhang G (2004) Optimization of initial substrate and pH levels for germination of sporing hydrogenproducing anaerobes in cow dung compost. Bioresour Technol 91:189-193 10. Fang HHP, Zhang T, Liu H (2002) Microbial diversity of a mesophilic hydrogen-producing sludge. Appl Microbiol Biotechnol 58:112-118 11. Ferchichi M, Crabbe E, Gil GH, Hintz W, Almadidy A (2005) Influence of initial pH on hydrogen production from cheese whey. J Biotechnol 120:402-409

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12. Hu B, Chen S (2007) Pretreatment of methanogenic granules for immobilized hydrogen fermentation. Int J Hydrogen Energy 32:3266-3273 13. Jones DT (1986) Acetone-butanol fermentation revisited. Microbiol Rev 50:484-524 14. Jun YS, Yu SH, Ryu KG, Lee TJ (2008) Kinetic study of pH effects on biological hydrogen production by a mixed culture. J Microbiol Biotechnol 18:1130-1135 15. Kalogo Y, Bagley DM (2008) Fermentative hydrogen production using biosolids pellets as the inocule source. Bioresour Technol 99:540-546 16. Kanso S, Dasri K, Tingthong S, Watanapokasin RY (2011) Diversity of cultivable hydrogen-producing bacteria isolated from agricultural soils waste water sludge and cow dung. Int J Hydrogen Energy 36:8735-8742 17. Karlsson A, Vallin L, Ejlertsson J (2008) Effects of temperature hydraulic retention time and hydrogen extraction rate on hydrogen production from the fermentation of food industry residues and manure.

Int J Hydrogen Energy 33:953-962 18. Khanna N, Kotay SM, Gilbert JJ, Das D (2011) Improvement of biohydrogen production by Enterobacter cloacae IIT-BT 08 under regulated pH. J Biotechnol 152:9-15 19. Kim KJ, Nhat L, Chun YN, Kim SW (2008) Hydrogen production conditions from food waste by dark fermentation with Clostridium beijerinckii KCTC 1785. Biotechnol Bioprocess Eng 13:499-504 20. Koskinen PEP, Beck SR, Örlygsson J, Puhakka JA (2008) Ethanol and hydrogen production by two thermophilic, anaerobic bacteria isolated from Icelandic geothermal areas. Biotechnol Bioeng 101:679-690 21. Lane D J (1991)16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M (eds) Nucleic acid techniques in bacterial systematic. John Wiley, New York, USA, pp 115-175 22. Lee KS, Wu JF, Lo YS, Lo YC, Lin PJ, Chang JS (2004) Anaerobic hydrogen production with an efficient carrier-induced granular sludge bed bioreactor. Biotechnol Bioeng 87:648-657 23. Lin CY, Chang RC (2004) Fermentative hydrogen production at ambient temperature. Int J Hydrogen Energy 29:715-720 24. Liu IC, Whang LM, Ren WJ, Lin PY (2011) The effect of pH on the production of biohydrogen by clostridia: thermodynamic and metabolic considerations. Int J Hydrogen Energy 36:439-449 25. Liu G, Shen J (2004) Effects of culture and medium conditions on hydrogen production from starch using anaerobic bacteria. J Biosci Bioeng 4:251-256 26. Logan BE, Oh SE, Kim IS, van Ginkel SV (2002) Biological hydrogen production measured in batch anaerobic respirometers. Environ Sci Technol 36:2530-2535 27. Mizuno O, Dinsdale R, Hawkes FR, Hawkes DL, Noike T (2000) Enhancement of hydrogen production from glucose by nitrogen gas sparging. Bioresour Technol 73:59-65 28. Morimoto M, Atsuko M, Atif AAY, Ngan MA, Fakhru’l-Razi A, Iyuke SE, Bakir AM (2004) Biological production of hydrogen from glucose by natural anaerobic microflora. Int J Hydrogen Energy 29:709-713 29. Rainey FA, Hollen BJ, Small A (2009) Genus I Clostridium. In: Vos P, Garrity GM, Jones D, Krieg NR, Ludwig W, Rainey FA, Schleifer KH, Whitman WB (eds) Bergey’s Manual of Systematic Bacteriology 2nd ed, vol 3: The Firmicutes. Springer, Dordrecht, The Netherlands, pp 738-828 30. Reddy MV, Chandrasekhar K, Mohan SV (2011) Influence of carbohydrates and proteins concentration on fermentative hydrogen pro-

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duction using canteen based waste under acidophilic microenvironment. J Biotechnol 155:387-395 31. Sanz JL, RodrĂ­guez N, Amils R (1997) Effect of chlorinated aliphatic hydrocarbons on the acetoclastic methanogenic activity of granular sludge. Appl Microbiol Biotechnol 47:324-328 32. Sciarria TP, Romano S, Correnti A (2010) Use of mixed substrate composed of organic solid wastes and energetic crop plants (Jerusalem artichoke) for the combined production of hydrogen ethanol and methane in a two-stage fermentation process. J Biotechnol 150 (suppl 1):S141-S152 33. Sivaramakrishna D, Sreekanth D, Himabindu V, Anjaneyulu Y (2009) Biological hydrogen production from probiotic wastewater as substrate by selectively enriched anaerobic mixed microflora.

Renew Energy 34:937-940 34. Tang GL, Huang J, Sun ZJ, Tang QQ, Yan CH, Liu GQ (2008) Biohydrogen production from cattle wastewater by enriched anaerobic mixed consortia: influence of fermentation temperature and pH. J Biosci Bioeng 106:80-87

rajhi et al.

35. Valdez-VĂĄzquez I, Poggi-Varaldo HM (2009) Hydrogen production by fermentative consortia. Renew Sust Energy Rev 13:1000-1103 36. Wang X, Hoefel D, Saint CP, Monis PT, Jin B (2007)The isolation and microbial community analysis of hydrogen producing bacteria from activated sludge. J Appl Microbiol 103:1415-1423 37. Xu JF, Ren NQ, Wang AJ, Qiu J. Zhao QL, Feng YJ, Liu BF (2010) Cell growth and hydrogen production on the mixture of xylose and glucose using a novel strain of Clostridium sp. HR-1 isolated from cow dung compost. Int J Hydrogen Energy 35:13467-13474 38. Zeng AP, Biebl H, Schlieker H, Deckwer WD (1993) Pathway analysis of glycerol fermentation by Klebsiella pneumoniae: Regulation of reducing equivalent balance and product formation. Enzyme Microb Technol 15:770-779 39. Zhao P, Fan SQ, Tian L, Pan CM, Fan YT, Hou HW (2010) Hydrogen production characteristics from dark fermentation of maltose by an isolated strain F.P 01. Int J Hydrogen Energy 35:7189-7193

BOOK REVIEWS International Microbiology (2013) 16:63-68 ISSN 1139-6709 www.im.microbios.org

INTERNATIONAL MICROBIOLOGY

Molecular typing in bacterial infections Ivano de Flippis, Marian L. McKee (eds) Series Editor: Vassil St. Georgiev 2013, Humana Press 482 pp, 16 × 24 cm Price: 176.75 € (hardcover); 142.79 € (eBook) ISBN: 978-1-162703-184-4

New molecular tools taking advantage of DNA-based approaches are highly relevant to epidemiological studies and they have expanded rapidly over the past 20 years.

Molecular typing in bacterial infections, edited by de Filippis and McKee, is structured into seven parts: (I) General consideration of microorganism typing methods, (II) gastrointestinal pathogens, (III) oral and respiratory pathogens, (IV) urogenital pathogens, (V) vector-borne pathogens, (VI) pathogens causing hospital-associated infections, and (VII) emerging and re-emerging pathogens.

Each part discusses the most important microorganisms in each group and the chapters highlight the most frequently used methods for the typing of each of these pathogens.

However, the chapters are not unified by a regular format; for example, while some have an introduction and conclusion or concluding remarks, others do not.

In part I, A.E. Seitz and D.R. Prevots define molecular epidemiology and describe its current applications. In part II, Chapter 2, L.M. Teixeira and V.L.C. Merquior discuss the different methods used in the molecular typing of Enterococcus, remarking on the absence of a single definitive typing method for this group of bacteria. In Chapter 3, T. J. Ward presents the different phenotypic and genotypic methods used to investigate the epidemiological relationships among Listeria monocytogenes strains. He compares the key features of pulse-field gel electrophoresis (PFGE), multilocus sequence typing (MLST), multilocus geno-typing (MLGT), and multiple-locus variable-number tandem repeat analysis (MLVA). Although PFGE remains the gold

standard for the discrimination of L. monocytogenes strains, further development of DNA-sequence-based subtyping will probably provide the best combination of discriminatory power, epidemiological utility, and efficiency.

In Chapter 4, S.L. Foley, A.M. Lynne, and R. Nayak review the different typing methods that comprise restriction-based methods, DNA-amplification-based methods, and DNA sequencing-based methods to discriminate among strains from the most frequently isolated species of Enterobacteriaceae. Here as well, PFGE is the gold standard; however, MLST can be used to gain a better appreciation of the genetic diversity of the population of isolates being examined. A report on molecular methods to type Vibrio cholera is provided by T. Ramamurthy, A.K. Mukhopadhyay, R.K. Nandy, and G. Balakrish in Chapter 5.

These authors described the most frequently used typing methods but also include a discussion of genetic elements, such as plasmids, insertion sequences, and integrons, as the basis of potential phylogenetic typing systems. This is the first chapter in the book that evaluates the whole genome approach as a powerful tool to understand the origin and relationship of pandemic clones. In Chapters 6 and 7, the typing of two anaerobes (C. difficile and Bacteroidetes) is introduced. A plethora of different methods have been used to type C. difficile because of its clinical importance, with PFGE and MLVA having the highest discriminatory power.

Part III covers oral and respiratory pathogens. L. McGee and B. Beall, in Chapter 8, analyze the three major streptococci (S. pyogenes, S. agalactiae, and S. pneumonia).

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Int. Microbiol. Vol. 16, 2013

They note that MLST is the most discriminatory tool but also describe the underlying concept of eBURST and its application in exploring patterns of evolutionary descent.

In Chapter 9, specific typing of Streptococcus mutans is presented. Chapter 10 deals with metagenomic analysis of the oral microbiome and the specific typing of periodontal pathogens. In the authorsâ&#x20AC;&#x2122; view, typing is more informative for therapeutics than for epidemiology. Typing of non-tuberculous mycobacteria is the focus of Chapter 11. The next two chapters address the typing of Neisseria meningitides (Chapter 12), and Haemophilus influenza (Chapter 13); in both cases, MLST is currently the most commonly used method. Chapters 14â&#x20AC;&#x201C;18 cover Moraxella, Legionella pneumophila, mycoplasma and ureaplasma, Corynebacterium diphteriae, and Burkholderia. Overall, the content in these chapters is very heterogeneous; for instance, only seven pages are devoted to L. pneumophila while 53 pages are devoted to mycoplasma and ureaplasma. In the chapters on Moraxella and L. pneumophila, matrix-assisted laser desorption-ionization time-of-light mass spectrometry (MALDI-ToF MS) as a potential typing tool is discussed.

Part IV covers Treponema (Chapter 19) and the family Chlamydiaceae (Chapter 20). PCR-based restriction fragment length polymorphism (RFLP) analysis of specific genes is one of the most common techniques used to type these microorganisms. Part V focuses on vector-borne

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pathogens, i.e., Borrelia (Chapter 21) and Erysipelotrix (Chapter 22). MLST and PCR-sequencing are viewed as future typing techniques for these microorganisms. Parts VI and VII discuss major nosocomial microorganisms: staphylococci (Chapter 23), Pseudomonas (Chapter 24), and Acinetobacter baumannii (Chapter 25). In these microorganisms, two routes of investigation can be distinguished: local hospital outbreaks and international population studies, with PFGE and MLST as the most appropriate typing techniques, respectively.

Familiarity with the different methods for the correct typing of microorganisms is important for many fields of medical and microbiological research. This book should be required reading not just for microbiologists, epidemiologists, and infectious disease specialists but also for university students of infectious diseases and clinical microbiology, who while increasingly trained in molecular epidemiology may have a poor understanding of the techniques used in this field.

Jordi Vila University of Barcelona jvila@ub.edu

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Int. Microbiol. Vol. 16, 2013

65

Halophiles and hypersaline environments.

Current research and future trends.

Antonio Ventosa, Aharon Oren, Yanhe Ma (eds) 2010. Springer-Verlag, Berlin, Germany 387 pp, 16 × 25 cm Price: 176.79 € (hard cover); 142.79 € (eBook) ISBN 978-3-642-20197-4 (hard cover) ISBN 978-3-642-20198-1 (eBook)

Although we often ignore their presence, colonies of halophiles can be readily recognized by the bright red color they confer on marine salterns and crystallizer ponds world-wide. They include populations of halophilic Archaea (Halobacteriales), Bacteria (Salinibacter), and Eukarya (Dunaliella salina) [p.173]. Salt (NaCl) has been generally considered hostile to most forms of life. Thus, for centuries, in addition to its use as a ubiquitous food seasoning, it has long been employed as a food preservative. However, halophilic and halotolerant microorganisms can contaminate salt-preserved food, consistent with their colonization of natural saline environments.

Halophiles and hypersaline environments originated at the international meeting “Halophiles 2010”, held in Beijing, China, in July 2010. Edited by Antonio Ventosa (University of Sevilla), Aharon Oren (The Hebrew University of Jerusalem), and Yanhe Ma (Chinese Academy of Sciences), the book comprises 20 chapters whose authors, like the editors, are experts in the field. Three of these 20 chapters are devoted to historical aspects of halophiles research.

Chapter 1 (coauthored by the editors) is a very moving text dedicated to the memory of Prof. Helge Larsen (1922– 2005), one of the founders of halophilic microbiology and the greatest expert of his time on halophilic microorganisms. Chapter 2, by A. Oren, looks back on the history of microbiology, focusing on the work of Baas Becking (1895–1963), whose first studies on the salt lakes in California were done in the 1920s and laid the foundations for halophilic microbiology. Oren is also the author of the book’s last chapter, a historical overview of the symposia

on halophilic microorganisms, which started in 1978 in Rehovot, Israel. The remaining chapters of Halophiles and hypersaline environments inform the reader about the state of the art of the field of halophiles, thoroughly discussing the various lines of research.

Chapters 3–6 focus on the diversity that can be found in hypersaline environments, as evaluated by molecular (metagenomics, lipidomics) and culture-dependent methods. Several taxonomic aspects of certain halophilic microorganisms, including those in the Family Halomonaceae as well as Salinibacter ruber, a typical representative of the Bacteroidetes that shares many properties with Haloarchaea.

Baas Becking [see Quispel A. 1998. Lourens G.M. Baas Becking (1895–1963), inspirator for many microbiologists. Int. Microbiol. 1:69-72] was probably the first to realize that microorganisms living at high salt concentration could employ multiple survival strategies, and he focused his research on the adaptation of halopiles to life at extremes of salinity, pH, etc. Since then, there have been major studies of halophilic and halotolerant Bacteria and Archaea, as well as the eukaryotic alga Dunaliella salina, the only eukaryote able to adapt to these extreme conditions. Chapter 7 describes the molecular mechanism of adaptation to high salt concentration by the extremely halotolerant black yeast Hortaea werneckii. Halophilic and halotolerant fungi use polyols such as glycerol, erythritol, arabitol, and mannitol as osmotic solutes and retain low salt concentrations in their cytoplasm.

The first studies on organisms living in extreme environments date back to the late nineteenth century. However,

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it was not until the 1970s that viruses in halophilic environments were first described, and their role therein remains largely unexplored. Only recently has their participation in biogeochemical cycles and the genetic plasticity of their hosts been investigated. The morphology of viral particles in saturated brines has also been studied directly by electron microscopy of crystallizer samples, and new viral architectures have been found. It has become clear that NaClsaturated brines are among the planktonic systems with the highest number of virus-like-particles (VLPs), ranging from 7.3×107 VLP/ml in crystallized ponds to 2×109 VLP/ml in the Dead Sea. The isolation and characterization of viruses that contain a lipid envelope surrounding either a singlestranded or double-stranded DNA genome has shown that viral diversity in hypersaline environments is much larger than previously assumed.

In salt lakes, haloviruses generally outnumber cells by 10- to 100-fold. The term halovirus refers to viruses that infect halophiles, the hosts being either bacterial or archaeal species. Phages have been obtained as pure cultures from haloarchaea for many years (Halobacterium, Natrialba sp., Haloarcula sp., Haloferax sp., and Halorubrum sp.) and some of them have been sequenced. Chapters 8 and 9 deal with the diversity of halophilic viruses in salterns and in the Great Salt Lake (Utah, USA), respectively.

Chapters 10–17 describe the physiology and molecular biology of halophiles, including different mechanism of translation, the glycosylation of proteins, protein insertion, and the transport of proteins across lipid membranes.

Among the interesting physiological properties of halophiles is gas vesicle formation, by Halobacterium salinarum, and the nature and function of pigments such as carotenoids and xanthorhopopsin. The novel anaerobic halophilic alkali thermophiles are discussed as well. Halophiles exposed to multiple stressors have developed several unique adaptive mechanisms to control membrane permeability, intracellular osmotic balance, and the stability of the cell wall and intracellular proteins. Anaerobic halophilic, alkaliphilic, thermophilic bacteria have been isolated from the Wadi an-Natrun, in the Egyptian Sahara. Two new species,

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designated Natranaerobius jonesii and Natranaerobius grantii, are currently being characterized. The discovery of microorganisms capable of living in a combination of high salinity, pH, and temperature has extended our concept of the limits of life.

Thus far, halophiles have not been extensively exploited in biotechnological processes, with notable exceptions including the production of b-carotene by Dunaliella, bacteriorhodopsin by Halobacterium, and ectoine by Halomonas. However, as discussed in Chapters 18 and 19, because of their unique physiological traits halophilic microorganisms hold promise for other biotechnological applications. Their high stability at extreme conditions, such as high salt concentrations, low water availability, and high temperatures, make halophilic enzymes, including cellulases, lipases, amylases, and restriction enzymes, of particular interest in industrial and biotechnological applications, e.g., bioremediation and biofuel production.

The hypersaline environments of saltern pond brines and natural salt lakes have been traditionally overlooked by ecologists because they are considered to be relatively simple ecosystems with low diversity and high community densities. Nonetheless, they offer fascinating models to study fundamental questions of microbial biodiversity, selection, biogeography, and evolution. The book Halophiles and hypersaline environments provides an excellent overview of the current understanding of all aspects of the microbial world that lives in areas of high salt concentrations.

Together with an earlier book on “salt-loving” microorganisms, Halophilic microorganisms, also edited by Antonio Ventosa (Springer, 2004; see review in Int. Microbiol.

[2004] 7:233), this book provides a useful reference for researchers, students, and anyone interested in knowing more about these unique microorganisms.

Ricardo Guerrero University of Barcelona rguerrero@iec.cat

Int. Microbiol. Vol. 16, 2013

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67

Horizontal gene transfer in microorganisms M. Pilar Francino (ed) 2012, Caister Acad Press, Norfolk, UK 202 pp, 16 × 24 cm Price: £159 ISBN: 978-1-908230-10-2

Horizontal gene transfer (HGT) is a major force in microbial evolution and a great source of genetic innovation in prokaryotes. The first evidence of HGT was reported in 1928 by Frederick Griffith, who noted that bacteria (pneumococci) are capable of transferring genetic information (virulence determinants) through a process known as transformation. Later, the identification of gene transfer mediated by plasmids (conjugation) and viruses (bacteriophages, transduction) and the recognition of transposable elements further improved our knowledge of gene flux and the importance of mobile genetic elements. The book Horizontal gene transfer in microorganisms provides state of the art information on this fascinating topic. The nine chapters are written by several experts in the field, including the book’s editor M. Pilar Francino.

The flow of genes between different species is a form of genetic variation whose implications have yet to be fully appreciated. Strains of the same bacterial species frequently differ in up to 30–35% of their total genes in the genome, resulting in organisms with contrasting phenotypes and/or ecological strategies (e.g., highly invasive pathogens vs. commensal strains of Escherichia coli). Mobile genetic elements, such as transposons, phages and plasmids, account for most of the differences in gene content among closely related strains. This genomic flexibility contributes to the ability of bacteria to adapt to varying environmental challenges and thus ensure survival. The persistence of new genes depends on whether they are replicated as parts of integrated genomes. Genomes and genes evolve in response to natural selection at the organismal

level, as they regulate gene interactions and permanence (Chapter 1).

In prokaryotes, usually around 80% of the genome encodes proteins. Bacterial chromosomes also seem to be organized into regions with different gene expression levels.

Functionally related genes are usually transcribed in operons or superoperons. Genomics and genome sequencing studies have confirmed that the position of genes in the chromosome is not random but the result of selection.

However, despite these common organizing principles, prokaryotic genomes are extremely diverse. Bacterial genome sequencing has become so easy and accessible that the genomes of multiple strains of an increasing number of individual species have been and will be rapidly determined. These data sets provide for in-depth analysis of intra-species diversity.

The pan-genome is the sum of the core genome, i.e., those genes common to all strains, and the dispensable genome. The latter group of genes may confer adaptive advantages under certain environmental conditions, such as in the presence of antibiotics, xenobiotics, and other compounds. These genes also confer the important characteristics that allow the colonization of new ecological niches governed by biotic factors such as symbiotic and pathogenic relationships. Pan-genome analyses are aimed at interpreting the sequence data among widespread lateral gene flow. A comparison of 20 strains of Escherichia coli showed that the core genome consists of only 2000 genes, less than half the average genome size in a typical strain.

Modifications of gene repertoires can strongly affect cellular

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Int. Microbiol. Vol. 16, 2013

metabolism and morphology, and such changes can provide prokaryotes with the opportunities for adaptation to new ecological niches (Chapter 2). The mammalian-adapted bacterium Bartonella and plant-adapted rhizobia are discussed as examples in Chapter 3.

Horizontal gene transfer operates by a variety of mechanisms, relies on different ecological circumstances, and results in a wide range of fitness effects on the recipient organisms. Chapter 4 explores the barriers that limit HGT among microorganisms, such as DNA restriction. Exogenous DNA that survives restriction must be integrated into the bacterial host chromosome and then expressed in order to persist in the recipient cell. Transferred genes may continue evolving in different genomic and ecological environments. Chapter 5 analyzes the process of sequence divergence of transferred genes at different evolutionary time-scales in the Xanthomonadales, a group of plant pathogens belonging to Gammaproteobacteria. The genomes of the Xanthomonadales are the result of multiple HGT events from other Proteobacteria to the ancestral genome of the group. Chapter 6 discusses the environmental regulation of HGT, as opposed to genetic regulation at the cellular level as explained in Chapter 4. HGT has been studied in natural ecosystems, including soil, water, and the gut microbiota. Conditions in the gut are favorable for HGT because of the constant body temperature and inflow of nutrients, which support high population densities and vast diversity. Bacterial responses to host stress hormones can facilitate conjugative gene transfer between enteric bacteria. In in-vitro experiments, the physiological concentration of noradrenaline was shown to stimulate the transfer of a conjugative plasmid from a clinical strain of Salmonella sp. to an E. coli recipient. These mediators of host stress may affect HGT within the in vivo environment as well.

Plasmids, bacteriophage elements, transposons, and insertion sequences are mobile genetic elements because they can move between microbial cells. The total of all the mobile genetic elements in a cell is referred to as the mobilome. Despite the ability of plasmids to transfer across dif-

BOOK REVIEWS

ferent taxonomic groups of bacteria, it is unclear which factors are responsible for plasmid continuance among bacterial populations. Chapter 7 describes the different hypotheses explaining the maintenance of plasmids.

Several approaches can be used to detect mobile genetic elements. In Chapter 8, four different experimental methodologies are described: sequencing studies, sequence-based screening, capture studies (hybridization with magnetic beads), and functional studies. The appearance and dissemination of antibiotic-resistance in bacterial pathogens have stimulated investigations of the genetic aspects of the different phenomena associated with resistance development, such as gene pickup, heterologous expression, HGT, and mutation. Chapter 9 focuses on HGT and recombination in the evolution of antibiotic resistance genes. Essentially any mobile genetic element found in bacteria can acquire resistance genes and promote their transmission; the type of element involved varies with the genus of the pathogen, but plasmid-mediated transmission is far and away the most common mechanism of HGT. Moreover, the existing processes of gene acquisition, transfer, modification, and expression are expanding and their presence in the modern biosphere is accelerating in response to anthropogenic influences.

Clearly, prokaryotes, are under different forms of selective pressures with respect to DNA exchange than most multicellular organisms. The theory of prokaryotic evolution needs further work as it must take into account the population genetics and molecular evolution of organisms that have no traditional species boundaries but share genes across considerable evolutionary distance. Horizontal gene transfer in microorganisms is an up-to-date and necessary review of the most topical aspects of HGT.

Mercedes Berlanga University of Barcelona mberlanga@ub.edu

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Volume 16(1) MARCH 2013

Divisió Alimentària del IRTA-Centre de Tecnologia de la Carn. Generalitat de Catalunya. Monells (Girona) / Biblioteca Montilivi. Facultat de Ciències. Universitat de Girona / Área de Microbiología. Departamento de Ciencias de la Salud.

Universidad de Jaén / Microbiologia. Departament de Ciències Mèdiques Bàsiques. Facultat de Medicina. Universitat de Lleida / Laboratorio de Microbiología Aplicada. Centro de Biología Molecular. Universidad Autónoma de Madrid-CSIC. Cantoblanco (Madrid) / Laboratorio de Patógenos Bacterianos Intracelulares. Centro Nacional de BiotecnologíaCSIC. Cantoblanco (Madrid) / Grupo de Investigación de Bioingeniería y Materiales (BIO-MAT). Escuela Técnica Superior de Ingenieros Industriales. Universidad Politécnica de Madrid / Biblioteca. Centro de Investigaciones Biológicas, CSIC. Madrid / Merck Sharp & Dohme de España, Madrid / Departamento de Microbiología. Facultad de Ciencias.

Universidad de Málaga / Grupo de Fisiología Microbiana.

Depto. de Genética y Microbiología. Universidad de Murcia.

Espinardo (Murcia) / Library. Department of Geosciences.

University of Massachusetts-Amherst. USA / Biblioteca de Ciencias. Universidad de Navarra. Pamplona / Grupo de Genética y Microbiología. Departamento de Producción Agraria. Universidad Pública de Navarra. Pamplona / Microbiología Ambiental. Departamento de Biología. Universidad de Puerto Rico. Río Piedras. Puerto Rico / Biblioteca General.

Universidad San Francisco de Quito. Ecuador / Biblioteca.

Facultat de Medicina. Universitat Rovira Virgili. Reus / Instituto de Microbiología Bioquímica-Departamento de Microbiología y Genética. CSIC-Universidad de Salamanca / Departamento de Microbiología y Parasitología. Universidad de Santiago de Compostela. Santiago / Laboratorio de Referencia de E. coli (LREC). Facultad de Veterinaria.

Universidad de Santiago de Compostela. Lugo / Departamento de Genética. Universidad de Sevilla / Tecnología de los Alimentos. Facultad de Ciencias. Universidad de Vigo / General Library. Marine Biological Laboratory. Woods Hole, Massachusetts, USA.

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INTERNATIONAL MICROBIOLOGY Official journal of the Spanish Society for Microbiology Volume 16 · Number 1 · March 2013

RESEARCH REVIEW

Tortajada M, da Silva LF, Prieto MA Second-generation functionalized mediumchain-length polyhydroxyalkanoates: the gateway to high-value bioplastic applications

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RESEARCH ARTICLES

Marsh SE, Poulsen M, Gorosito NB, PintoTomás A, Masiulionis VE, Currie CR Association between Pseudonocardia symbionts and Atta leaf-cutting ants suggested by improved isolation methods

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López AC, Minnaard J, Pérez PF, Alippi AM In vitro interaction between Bacillus megaterium strains and Caco-2 cells

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Hashimoto W, Miyamoto Y, Yamamoto M, Yoneyama F, Murata K A novel bleb-dependent polysaccharide export system in nitrogen-fixing Azotobacter vinelandii subjected to low nitrogen gas levels

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Wróbel B, Filippini M, Piwowarczyk J, Kędra M, Kuliński K, Middelboe M Low virus to prokaryote ratios in the cold: benthic viruses and prokaryotes in a subpolar marine ecosystem (Hornsund, Svalbard)

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Rajhi H, Conthe M, Puyol D, Díaz E, Sanz JL Dark fermentation: isolation and characterization of hydrogen-producing strains from sludges

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BOOK REVIEWS

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