International Microbiology

Volume 17 路 Number 2 路 June 2014 路 ISSN 1139-6709 路 e-ISSN 1618-1905

INTERNATIONAL MICROBIOLOGY www.im.microbios.org

17(2) 2014

Official journal of the Spanish Society for Microbiology

Publication Board

Editorial Board

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

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

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

Addresses Editorial Office International Microbiology C/ Poblet, 15 08028 Barcelona, Spain Tel. & Fax +34-933341079 E-mail: int.microbiol@microbios.org www.im.microbios.org Spanish Society for Microbiology C/ Rodríguez San Pedro, 2 #210 28015 Madrid, Spain Tel. +34-915613381. Fax +34-915613299 E-mail: sem@microbiologia.org www.semicrobiologia.org Publisher (electronic version) Institute for Catalan Studies Carme, 47 08001 Barcelona, Spain Tel. +34-932701620. Fax +34-932701180 E-mail: int.microbiol@microbios.org © 2014 Spanish Society for Microbiology & Institute for Catalan Studies.

Printed in Spain ISSN (print): 1139-6709 e-ISSN (electronic): 1618-1095 D.L.: B.23341-2004

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

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CONTENTS International Microbiology (2014) 17:65-130 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

Volume 17, Number 2, June 2014 RESEARCH REVIEW

Romero D, Kolter R Functional amyloids in bacteria

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RESEARCH ARTICLES

López-García MT, Rioseras B, Yagüe P, Álvarez JR, Manteca A Cell immobilization of Streptomyces coelicolor: effect on differentiation and actinorhodin production

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Bonaterra A, Badosa E, Rezzonico F, Duffy B, Montesinos E Phenotypic comparison of clinical and plant-beneficial strains of Pantoea agglomerans

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Becerra A, Rivas M, García-Ferris C, Lazcano A, Peretó J A phylogenetic approach to the early evolution of autotrophy: the case of the reverse TCA and the reductive acetyl-CoA pathways Carrasco P, Pérez-Cobas AE, van de Pol C, Baixeras J, Moya A, Latorre A Succession of the gut microbiota in the cockroach Blattella germanica Castro N, Toranzo AE, Magariños B A multiplex PCR for the simultaneous detection of Tenacibaculum maritimum and Edwardsiella tarda in aquaculture PERSPECTIVES

Report NIH-CDC BOOK REVIEWS

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111 119 129

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

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

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

Electron microscopy micrograph showing the intimate contact between a mature dendritic cell exposed to HIV-1 (top) and a CD4+ T cell (bottom). HIV-1 particles captured by the mature dendritic cell are polarized to the cell-to-cell contact area, favoring CD4+ T cell infection throughout the formation of an infectious synapse. Micrograph by M. Teresa Fernández-Figueras and Núria Izquierdo-Useros, Pathology Department at HUGTIP and AIDS Research Institute IrsiCaixa, Barcelona, Spain. (Magnification, ca. 20,000×) Center. The Maya Hero Twins are the central figures of a narrative included in the document called Popol Vuh (translated as Book of the Council, Book of the Community, Book of the People, and Sacred Book), the oldest Maya myth known, the creation account of the Guatemalan Quiché Mayan people. The twins are Hunahpu and Xbalanque; they represent the two forces in Creation, the Light and the Dark, the Good and the Bad. Without these opposing forces, life would be static and slow to a halt. Both contrasting forces are needed in order to provide the energy and the counterbalance that keeps life in motion. (Comments and illustration provided by Susana Osorio de Mrozek.) See article on the early evolution of authotrophy by A. Becerra et al., pp 91-97, this issue.

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

32,000×)

late orders of the phylum Axostylata, specifically Tri­cho­monadida, Hypermastigida, and Oxymonadida. Photograph (dark-field microscopy) by Rubén Duro (Center for Microbiological Research, CIM, Barcelona). See covers of Int. Microbiol. vol.

14 (2011) and article by R. Guerrero, L. Margulis, M. Berlanga, Int. Microbiol. 15(2013):133-143.

(Magnification, ca. 1,500×) Lower right. Scanning electron micrograph of a 24-h mixed biofilm containing Candida albicans hyphae and blastoconidia of Candida glabrata.

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

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

Back cover: Pioneers in Microbiology Vicente Izquierdo (1850–1926), Chile Portrait of Vicente Izquierdo Sanfuentes (1850–1926), one of the founding fathers of microbiology in Chile. He was born in Santiago in 1850, to Vicente Izquierdo Urmeneta and Ana Sanfuentes Torres.

Although he had always been interested in the natural world, he was very fond of his mother and, in accordance with her wishes, studied law. After his graduation from law school in 1872, he pursued his scientific vocation and enrolled in the School of Medicine, from which he graduated in March 1875. At that time, the Chilean Government was granting stays in Europe to the best young scientists, which allowed Izquierdo to spend almost five years in prestigious laboratories in Leipzig, Vienna, and Strasbourg—the latter was then a German city. But in 1879, he and his fellow countrymen had to return to Chile, which had become involved in the “Pacific War” against Bolivia and Peru. Young physicians trained in Europe were very well acquainted with the work of the British surgeon Joseph Lister (1827–1912). Thus, Izquierdo was able to introduce Lister’s antiseptic and aseptic methods in his hospital in Santiago, where he treated wounded soldiers.

By the 21st-century concept of microbiology, Izquierdo would not be considered a true microbiologist. In fact, the Chair that he held at the University of Chile, in Santiago, was that of Histology and Entomology.

However, the focus of his work and research included topics related to

microbiology and to the epidemiology of infectious diseases. As a member of the Chilean Parliament, in 1886 he helped to launch a public health law aimed at preventing the spread of the cholera pandemic in Chile. In addition, he headed the Junta de la vacuna (Board of Vaccination), which directed the virtual eradication of smallpox from the country.

In 1883, Izquierdo published his first work on experimental bacteriology, a study of the tuberculosis bacillus, which just one year earlier had been described

by Robert Koch. In 1885, he published a study on the “Peruvian wart,” asserting that this common name was incorrect because the lesion was not characterized by papillary hypertrophy but by an inflammatory infiltration of the interstitial tissues. In the same year, the young Peruvian physician Daniel Alcides Carrion [(1857–1885), who at that time was still a student of medicine; see back cover and page A2 of Int Microbiol 12(3-4), 2009] would discover that it was in fact an infectious disease. The causative microorganism was later identified to be the bacterium known today as Bartonella bacilliformis.

A major work by Izquierdo in microbiology was the 228-page Ensayo sobre los Protozoos de aguas dulces de Chile (Essay on the freshwater Protozoa of Chile), published in 1906 as an addendum to the Annales of the University. When he was 63, he suffered from retinal detachment, a disease for which an effective treatment had yet to be discovered. Not being able to read or use the microscope, Izquierdo resigned his university Chair in Histology in 1913. Despite his visual handicap, he did not abandon his scientific activity but instead devoted his time to entomology, his youngness’ interest. In the early 1920s, he described the principle of pheromones, compounds that were finally recognized and named around 30 years later. Izquierdo died in 1926.

Front cover and back cover design by MBerlanga & RGuerrero

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RESEARCH REVIEW International Microbiology (2014) 17:65-73 doi:10.2436/20.1501.01.208 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

Functional amyloids in bacteria Diego Romero,1*§ Roberto Kolter2§ Institute of Subtropical and Mediterranean Hortofruticulture “La Mayora”- CSIC, and Department of Microbiology, University of Malaga, Malaga, Spain.

2 Department of Microbiology and Immunobiology, Harvard Medical School, Boston, MA, USA

1

Received 12 June 2014 · Accepted 28 June 2014

Summary. The term amyloidosis is used to refer to a family of pathologies altering the homeostasis of human organs.

Despite having a name that alludes to starch content, the amyloid accumulations are made up of proteins that polymerize as long and rigid fibers. Amyloid proteins vary widely with respect to their amino acid sequences but they share similarities in their quaternary structure; the amyloid fibers are enriched in β-sheets arranged perpendicular to the axis of the fiber. This structural feature provides great robustness, remarkable stability, and insolubility. In addition, amyloid proteins specifically stain with certain dyes such as Congo red and thioflavin-T. The aggregation into amyloid fibers, however, it is not restricted to pathogenic processes, rather it seems to be widely distributed among proteins and polypeptides. Amyloid fibers are present in insects, fungi and bacteria, and they are important in maintaining the homeostasis of the organism. Such findings have motivated the use of the term “functional amyloid” to differentiate these amyloid proteins from their toxic siblings. This review focuses on systems that have evolved in bacteria that control the expression and assembly of amyloid proteins on cell surfaces, such that the robustness of amyloid proteins are used towards a beneficial end. [Int Microbiol 2014; 17(2):65-73] Keywords: Bacillus subtilis · bacterial biofilms · extracellular matrix · TasA amyloid-like fibers

Amyloids in history The term amyloidosis is used to refer to a family of pathologies altering the homeostasis of human organs. Written descrip­tions of what most likely were amyloidoses dates back to the late 17th century. An autopsy report describing a spleen full of white stones can be considered the first description of amyloidosis, which is now known as “sago spleen” to describe the Corresponding author: D. Romero Departamento de Microbiología. Facultad de Ciencias Universidad de Málaga-Instituto de Hortofruticultura Subtropical y Mediterránea “La Mayora”(IHSM-UMA-CSIC) Bulevar Louis Pasteur-Campus Universitario de Teatinos, s/n 29071 Málaga, Spain Tel. +34-952134274 E-mail: diego_romero@uma.es *

§

Both authors are equal contributors

starch-like granules that grow in the organ [20]. The term amyloid, meaning resembling starch, was first used two centuries later. The German chemist Rudolph Virchow discovered that the corporea amylacea of the nervous system stained with Congo red in a similar way as did cellulose and starch [41]. There was a long-standing controversy on the chemical nature of such amyloid plaques with some maintaining that they were made of starch (thus amyloid) and others arguing that they were more akin to lard [20]. It was not until the 20th century that chemical analyses revealed that such accumulations consisted of protein. By then, however, the medical term had gained a stronghold and to date these proteins are referred to as amyloids, despite their having no starch content whatsoever. However, the early descriptions of amyloid proteins already revealed a peculiarity; under the electron microscope amyloids appeared as long and rigid fibers [20].

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ROMERO, KOLTER.

Table 1. Amyloid proteins 1A. Pathogenic amyloids Name

Precursor

Disease

Ref.

AA

Serum amyloid

Systemic amyloidosis

[10]

Aβ

β-Protein (APP)

Alzheimer’s disease

[10]

AIAPP

Islet amyloid polypeptide

(Type 2 diabetes and insuloma)

[10]

α-Synuclein

α-Synuclein

Parkinson’s disease

[10]

Species

Protein

Function

Humans

Pmel17

Elimination of toxic intermediates during melanin synthesis

[19]

Fungi

Hydrophobins

Formation of fungal coat

[56]

Escherichia coli, Salmonella enteritidis

Curli

Interaction with host, biofilm formation

[9,15]

Pseudomonas sp.

FapC

Biofilm formation

[12]

Streptomyces coelicolor

Chaplins

Formation of aerial structures

[13]

Bacillus subtilis

TasA

Biofilm formation

[31]

Klebsiella pneumoniae

Microcin E492

Antimicrobial

[40]

Staphylococcus aureus

PSM

Biofilm formation

[37]

Streptococcus mutans

Adhesin P1

Biofilm formation

[29]

Mycobacterium tuberculosis

MTP

Host interaction

[1]

Xanthomonas axonopodis

Harpin

Virulence factor, multicellularity

[28,39]

1B. Functional amyloids

The intense research on amyloid proteins has shown that even though they vary widely with respect to their amino acid sequences, they share similarities in their quaternary structure.

Amyloid protein fibers are enriched in ���������������������� �������������������� -sheets arranged perpendicular to the axis of the fiber [50]. This structure provides great robustness, which is the defining feature of amyloid protein. Amyloids have remarkable stability, insolubility and specifically stain with certain dyes such as Congo red and thioflavin-T. The fact that amyloid fibers had always been associated with human pathologies led to the perception that the amyloid state was due to an erratic processing or misfolding of soluble and functional proteins [44]. However, the aggregation into amyloid fibers seems to be widely distributed among proteins and polypeptides, and in some cases, these amyloid fibers are important in maintaining the homeostasis of the organism [10,23]. Two outstanding examples of amyloids non-related to pathologies in humans are: The protein pMel17, which form amyloid fibers to eliminate intermediate aggregates that may be toxic to the organism, and proteins or peptides hormones of the

human endocrine system, which are efficiently stored in secretory granules in an amyloid-state, thus contributing to the normal physiology of cells [19,23]. Amyloid fibers are also present in insects, fungi and in bacteria, and they participate in protection, interaction with surfaces, and detoxification (Table 1).

Prions, another group of proteins with propensity to fold into amyloid fibers but with the astonishing ability of self-propagating, are mostly known for their pathological implications, although exceptions to this rule are arising [27,49]. This happens to be with the translation regulation protein CPEB (cytoplasmic polyadenylation element binding protein) in the mollusk Aplysia. As other prions, this protein possesses the ability to acquire different functional conformations, and it appears that the prion dominant form contributes to stabilizing the long-term stimulated synapses in memory storage [42]. Another example of a non-pathological prion is the protein HET-s in the fungus Podospora anserine [26,55], which in the prion state seems to contribute to build a physical barrier that impede the transfer of deleterious elements between genetically in-

compatible strains [26]. All these findings have motivated the use of the term “functional amyloid” to differentiate these amyloid proteins from their toxic siblings [14]. In this review we focus on systems that have evolved in bacteria that control the expression and assembly of amyloid proteins on cell surfaces, such that the robustness of amyloid proteins are used towards a beneficial end.

Functional amyloids in bacteria One of the first reports of functional amyloid proteins in bacteria was a study of the curli pili of an uropathogenic strain of Escherichia coli. These pili were initially described as fibronectin-binding organelles on the surface of cells [30]. Subsequently, Chapman and collaborators demonstrated that the curli fibers that emerged from the surfaces of E. coli cells had the same physical properties as the well-studied amyloid proteins responsible for human amyloidoses, e.g., staining with specific dyes (Fig.1) [9]. Escherichia coli uses curli amyloidlike fibers for a variety of physiological functions. Among these are interactions with host tissues, biofilm formation, and evasion of the immune system [4]. This first description of a bacterial functional amyloid opened the possibility that amy-

Fig. 1. Top row: Escherichia coli and Bacillus subtilis colonies stain with the amyloid specific dye Congo red. The pictures were taken after 72 h of growth in MSgg (minimal salts glycerol glutamate) agar (B. subtilis) or YESCA agar (E. coli) supplemented with Congo red. Bottom row: Transmission electron micrographs of curli amyloid fibers in E. coli (left) and TasA amyloid-like fibers in B. subtilis (right). Bars equal 200 nm.

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loid proteins could be present in other bacterial species. Indeed, Larsen and collaborators carried out an immunolabeling study with diverse bacterial species and suggested that amyloid proteins were present as constituents of bacterial biofilms in a variety of environmental samples [22]. However, there is still just a limited number of examples where the direct implication of amyloids in biofilm formation has been demonstrated: Tafi, the curli homolog in Salmonella, FapC in many Pseudomonas species, TasA in Bacillus subtilis, and the recently found phenol soluble modulins (PSMs) in Staphylococcus aureus and the adhesin protein P1 in Streptococcus mutans in dental plaque biofilms [12,15,29,31,37].

Amyloid proteins may be utilized to perform other physio­ l­ogical processes in bacteria. For example, chaplins of Streptomyces coelicolor not only serve to interact with surfaces, but they also facilitate the rising of the aerial hyphae [5,11]. In Mycobacterium tuberculosis, amyloid pili (MTP) are necessary to interact with the host during pathogenesis [1]. Another example of an amyloid having a role during pathogenesis involves the harpins, important virulence factors of Xanthomonas axonopodis and other plant pathogenic bacteria. The harpin Hpag of Xanthomonas has been demonstrated to form amyloid fibers in vitro, and to be related to the hypersensitive response (HR) caused in the host and most recently shown to

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BACTERIAL AMYLOIDS

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participate in the multicellular behavior of this bacterial species [28,39]. In this case, it was demonstrated that tetrameric oligomers were the main protein species found in hypersensitive responses. These findings contribute to the debate on the toxicity of amyloids. Historically, fibers have been considered as the etiological agents of amyloidosis. However, intermediate aggregates have a high propensity to insert into biological membranes and induce structural destabilization [16,51].

Thus, it is now thought that the intermediate aggregates, rather that the fibers themselves, are responsible for disease development [51].

Other studies demonstrate that, besides their role in developmental programs or interaction with the host cells, amyloids can be exploited as a detoxifying system. An interesting example of how the toxicity of amyloids can be modulated in self-benefit is that of the toxin microcin E492 produced by Klebsiella pneumoniae [40]. This toxin is a small peptide that oligomerizes into the cytoplasmic membranes of Enterobactericeae, causing pores that result in cell death [21]. The toxicity of microcin E492 was observed to vary depending on the state of growth of the producing cells; it was higher during exponential growth and decreased progressively with the aging of the culture. Not that the reduction of toxicity was associated to the transition of the toxin from an oligomeric state to further polymerization into amyloid fibers. This observation led to the conclusion that the toxic oligomers of microcin E492 resemble the intermediate aggregates of the protein A��������� β�������� associated with Alzheimer’s disease [2,24].

The inclusion bodies (IBs) that form during heterologous expression of proteins in bacteria are also an example of proteins with an amyloid conformation. The IBs have been historically considered the bottleneck that reduces the yield of overexpressed soluble and functional proteins. However, recent research demonstrates that peptides can be recovered from the IBs and still retain functionality. Besides the biological role of IBs as scavengers of putative toxic molecules for the bacterial cells, they offer an exciting new way for the efficient and controlled delivery of drugs in chemotherapy [52,53].

Amyloidogenesis as a sophisticated process of protein aggregation The discovery that amyloid proteins can carry out important physiological functions in bacteria has forced the re-evaluation of the concept that amyloidogenesis is always an erratic process of protein aggregation. As we describe in this section,

ROMERO, KOLTER.

rather than being an uncontrolled process, it is now clear that in bacteria, and probably in other organisms, sophisticated machineries have evolved that direct the polymerization of these amyloid fibers outside the cell, thus avoiding toxicity yet providing the structural robustness to produce very stable organelles [36].

Curli: a paradigm for bacterial amyloidogenesis.

Amyloid proteins have the intrinsic propensity to polymerize from the native monomeric state to the ordered and insoluble amyloid fibers [50]. During the formation of the fibers, amyloids go through diverse stages of aggregation with variable biochemical and morphological features. The kinetics of amyloid polymerization follow a typical sigmoidal curve with an initial lag phase, followed by an exponential phase of growth and a final plateau, where the fibers saturate and do not grow any further [25]. What allows the members of this family of proteins to form fibers? The study of curli in E. coli has clarified much of the molecular mechanisms that direct amyloidogenesis. The first clear difference with pathogenic amyloids is that curli formation is a highly regulated process. The protein products of two divergent operons are directly involved in the formation of curli fibers: csgABC and csgDEFG. The products of csgA and csgB are the main components of the fibers and the other proteins participate in regulating gene expression or control the proper secretion and polymerization of the fibers. CsgD regulates the expression of csgABC. CsgG forms a pore-like structure in the outer membrane, and allows the translocation of CsgA and CsgB from the periplasm to the outside of the cell. GsgF helps expose CsgB to the surfaces, and CsgE is thought to facilitate the access of CsgA to the secretion complex formed by CsgG. Finally, CsgC, which also has oxidoreductase activity, is thought to control the formation of the CsgG pore-like structure. The fibrillation of curli follows a nucleation-polymerization model, which means that the mixture of single mutants csgA and csgB complement each other extracellularly.

This is a fascinating result indicating that there is no need to produce both subunits in the same cell. The detailed analysis of the curlin subunits CsgA and CsgB demonstrated the existence of an amyloidogenic core, composed of four imperfect repeats within the proteins’ amino acid sequence. Thus, when CsgB encounters CsgA, it induces a conformational change towards the amyloidogenic state making its polymerization into fibers possible. The reaction between the two subunits is mediated by CsgC [4]. All this knowledge on curli biogenesis has served to establish an elegant E. coli cell-based methodology to evaluate the potential amyloidogenic proper-

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Int Microbiol

BACTERIAL AMYLOIDS

Fig. 2. Fibers of TasA purified from Bacillus subtilis as seen in electron microscopy and negative staining. Bar equals 100 nm.

ties of proteins [43]. In this study, human or yeast amyloid proteins were shown to be targeted to the envelop of E. coli cells via the curli system, where they propagated fibers with amyloid properties.

How conserved is the mechanism of polymerization among bacterial amyloids is a question that needs intensive investigation. In the closely related bacterium Salmonella typhimurium, homologs to each curli gene have been identified and thus the polymerization is hypothesized to follow a similar scheme [58]. Consistent with the above idea, a recent study demonstrated that homologs of CsgA found in E. coli and S. typhimurium, among other Gram-negative bacteria, can cross seed fiber formation in vitro. As stated by the authors, this observation leads to the idea that these heterogeneous curli fibers may be produced in mixed-species biofilms in natural settings [57]. In the case of the amyloid protein FapC of Pseudomonas species, although no similarity in sequence with curli genes is observed, the presence of genes that could play similar roles to each component of the machinery dedicated to curli in E. coli has been demonstrated [12]. Studies on other bacterial filaments described as pilli and fimbriae have demonstrated differences in the specific way that they are formed in Gram-positive and Gram-negative bacteria which should come as no surprise given the dramatic differences in the cell envelopes of these two general classes of bacteria [18].

Amyloid fibers in Gram-positive bacteria: TasA in Bacillus subtilis. A good model for the study of amyloid proteins of Gram-positive bacteria is TasA in B. subtilis.

The story of TasA is quite intriguing. TasA was first described in the late 1990s, by two separate groups, as a protein that was both secreted into the medium during stationary phase and as a constituent of the spores [38,46]. The absence of TasA did not affect the viability of the spores but the spores appeared morphologically altered. In addition, Stover and Driks [46] reported an intriguing result for TasA; when overexpressed in E. coli, it had broad-spectrum of antimicrobial activity. The name of TasA reflects the findings of this protein as a translocation-dependent antimicrobial spore component. Later, TasA was shown to be a major component of the extracellular matrix of B. subtilis biofilms and also required for the formation of biofilms [6]. This functionality was further described as being related to the amyloid-like nature of TasA (Fig. 1) [31].

Note that although not demonstrated or even predicted, the observation of TasA’s antimicrobial activity when produced in E. coli pointed towards one of the putative features of TasA as an amyloid protein: when produced in large quantities and in the absence of the additional elements necessary for the assembly in fibers, TasA may form toxic aggregates that like other amyloids, may cause cell death [46].

As introduced earlier, it can be proposed that B. subtilis uses TasA to produce amyloid fibers outside the cell for two purposes: (i) to detoxify the possible accumulation of toxic aggregates of this protein in the cytoplasm, and (ii) to form the protein-scaffold that supports the assembly of the extracellular matrix, the network of molecules necessary for the formation of bacterial biofilms. Thus, the formation of such fibers needs to be highly regulated. There is a complex regula-

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Fig. 3. The formation of biofilms of Bacillus subilis and TasA amyloid-like fibers in B. subtilis depend on the proteins TasA and TapA. (A) Top row: Highly wrinkly colony of wild type and featureless colony of tasA or tapA mutants after 72 h of growth at 30 ºC in MSgg agar plates. Bottom row: wrinkly pellicles of wild type and absence of pellicle of tasA and tapA mutants grown in MSgg broth for 24 h at 30 ºC. (B) Transmission electron microscopy and coimmunolabeling with anti-TapA (15 nm gold particles) and anti-TasA (10 nm gold particles) antibodies of TasA fibers in B. subtilis biofilms. Bar equals 100 nm.

tory network that controls and connects the expression of TasA with other bacterial factors [54]. To summarize, the master regulator SinR directly represses the expression of tasA until conditions are propitious for biofilm development [17]. One operon containing three genes, tapA-sipW-tasA, is necessary for the formation of the amyloid fibers [31]. This is markedly different from the chaplin amyloid fibers in the Gram-positive bacterium Streptomyces coelicolor, where eight chaplin genes have been described [13]. Streptomyces coelicolor shows a complex developmental program and it is thought that the diversity of chaplins play important roles in the different stages of the S. coelicolor life cycle [5].

TasA is the main component of the B. subtilis amyloid fibers.

When purified directly from B. subtilis, TasA retains amyloid properties, such as self-aggregation into insoluble fibers (Fig. 2). TasA fibers can be depolymerized with aggressive acid treatments and then they can be observed to repolymerize. Such a property has been useful to study the kinetics of polymerization of TasA, which reflects a dynamic transition through different stages of aggregation. In agreement with its amyloid nature, some of these intermediate aggregates, but not the fiber nor the monomers, reacts with an antibody that specifically detects the toxic aggregates of the amyloid protein Aβ associated with Alzheimer’s disease [31]. However, TasA is not purified from B. subtilis in its monomeric form.

Rather, we obtain a homogenous suspension of stable oligo-

mers, which could aggregate as different morphotypes depending on the physical properties of the medium: (i) fibers form on hydrophobic surfaces but (ii) plaques form under acidic conditions. This is an appealing discovery that could reflect a way in which bacteria can adapt to different habitats, modifying the state of aggregation of an external protein and probably the final arrangement of the extracellular matrix depending on environmental cues such as pH and surface hydrophobicity [8].

Bacterial cell surfaces are hydrophobic, and we have seen that this physical feature promotes the formation of fibers [8].

However, additional factors must be dedicated to increase the efficiency of the fiber polymerization process outside the cells. To build the curli fibers, E. coli uses CsgA and the nucleator protein CsgB [3]. As stated above, tasA is part of a three gene operon that encodes another protein, TapA, which is also necessary for the assembly of biofilms (Fig. 3A) [6,34].

The failure of a tapA deletion mutant to form biofilms has been related to the absence of TasA fibers and even attachment of TasA to cell surfaces [34]. Note that, differently from what was reported for curli, TapA and TasA have to be produced in the same cell in order to form the fibers, a process similar to the formation of pili in B. cereus [7].

Immunolocalization studies proved that TapA localizes, as foci, at certain regions of the cell wall, where presumably it directs the polymerization of TasA into fibers. In addition,

Fig. 4. A feature of TasA amyoid-like fibers is the binding of the amyloid specific fluorescent dye thioflavin T (structure on top). Thioflavin T fluorescence is used to follow the kinetics of polymerization of amyloid proteins. The increase of the fluorescence intensity indicates an increase in the β-sheet content of the protein, typical in amyloid fibers. Purified TasA follows a typical amyloid kinetics of polymerization (Ñ), and the addition of TapA accelerates the polymerization of TasA into amyloid-like fibers (¡).

TapA appears distributed along the TasA fibers (Fig. 3B), although at much less abundance (the ratio of TasA:TapA is 100:1) [34]. Based on these findings, we proposed that TapA is an accessory protein that promotes the efficient polymerization of TasA at the cell envelope and contributes to the organization of the growing fibers. In addition, the localization of TapA at the base of the TasA fibers, and its close association to the cell wall led to the idea that TapA is the connector of the fibers to the cell envelope [34].

Consistent with this conclusion, the absence of TapA causes a decrease in the amount of detectable extracellular TasA, which in addition appears as small and disorganized fibrils that are disconnected from the cell envelope. The name TapA was coined to refer to these functions: TasA anchoring and assembly protein [34].

In a recent publication we showed that indeed TapA accelerates the polymerization of TasA in vitro (Fig. 4) [35].

How it is that TapA facilitates the polymerization of TasA into amyloid fibers is still unclear. What we do know is that a mutant with a tapA allele lacking an eight-amino-acid sequence in the N-terminal half of the protein failed to form biofilms and failed to promote TasA fiber formation [35]. By comparison with curli in E. coli, we propose that this domain in the N-terminal half of TapA can either promote the processing of TasA or induce a conformational change of TasA toward its amyloidogenic state.

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The knowledge of the polymerization of pili in other Gram-positive bacteria can be compared and contrasted with TasA fiber formation. The formation of pili in B. cereus needs two subunits, BcpA and BcpB. The major subunit, BcpA is processed by a specific sortase, and the product is linked to the minor subunit BcpB [7]. However, a hypothetical processing of TasA and TapA would have to involve other proteins than the putative sortases identified in B. subtilis because mutants in those sortases had no defect in biofilm formation [34].

The anchoring of TasA fibers mediated by TapA could be thought to be similar to other pili in Gram-positive bacteria.

However, two lines of evidence discard this hypothesis: (i) The C-terminal region of TapA does not contain a canonical sorting signal for anchoring to the peptidoglycan, and (ii) the deletion of the two putative sortases, the enzymes that recognize the sorting signals, did not alter biofilm formation [34].

These findings open the possibility that there exist diverse mechanisms of attachment of fibers to the cell wall of Grampositive bacteria.

Finally, the formation of TasA fibers depends on a third protein, SipW, also encoded in the same operon. This is a signal peptidase that processes TasA and TapA to their mature form before sorting to the cell envelope [45,46]. SipW is an atypical signal peptidase, given that it resembles more those found in the endoplasmic reticulum of eukaryotic cells than typical bacterial signal peptidases [48]. An additional role for

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SipW has been recently proposed. A version of this protein impaired in protease activity was shown to stimulate the expression of the tapA-sipW-tasA operon during the formation of biofilms on solid surfaces, thus introducing a new level of control and complexity to the formation of amyloids [47].

Final notes The studies of bacterial amyloids have revolutionized our perception of this family of proteins: from their pathological implications and erratic processing to the possibility that they sometimes arise as the result of a well-defined program that has evolved to maintain the homeostasis of the organism and to interact with the environment [36]. To the increasing number of functional amyloids, we have added a new member, the TasA amyloid-like fiber in the Gram-positive bacterium B. subtilis. A number of reasons make the TasA amyloid-like fibers an attractive model for the molecular study of amyloidogenesis. (i) B. subtilis is a model microbe for the study of developmental programs such as biofilm formation and sporulation, two features mediating the interaction of the bacterium with its environment. The study of TasA fiber formation will contribute not only to our understanding of the mechanisms of amyloid fiber formation, but also its implications in bacterial adaptation to the habitat. The fact that TasA is purified to homogeneity directly from B. subtilis offers the outstanding capability to investigate which other factors, physical-chemical or intrinsic to bacterial cells, are involved in fiber formation. (ii) Amyloid fibers represent interesting targets for the search of new therapies that contribute to the fight of bacterial biofilms and the biochemical problems they may cause [32]. We have previously demonstrated that B. subtilis biofilms represent a formidable tool for the search of molecules that can be exploited in two directions: anti-biofilm and anti-amyloid [33].

Acknowledgements. The authors thank Hera Vlamakis for critical reading and suggestions during the writing of the manuscript. DR is funded by the program Ramón y Cajal (RyC-2011-080605) and grant AGL-201231968 from the Plan Nacional de I+D+I from Ministerio de Economía y Competitividad (Spain) and co-financed by FEDER funds (European Union).

Competing interest. None declared.

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RESEARCH ARTICLE International Microbiology (2014) 17:75-80 doi:10.2436/20.1501.01.209 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

Cell immobilization of Streptomyces coelicolor : effect on differentiation and actinorhodin production María Teresa López-García,1 Beatriz Rioseras,1 Paula Yagüe,1 José Ramón Álvarez,2 Ángel Manteca1* Microbiology Section, Department of Functional Biology and IUOPA, School of Medicine, University of Oviedo, Spain.

2 Department of Chemical Engineering and Environmental Technology, University of Oviedo, Spain

1

Received 9 January 2014 · Accepted 24 April 2014

Summary. Streptomycetes are mycelium-forming bacteria that produce two thirds of the clinically relevant secondary metabolites. Despite the fact that secondary metabolite production is activated at specific developmental stages of the Streptomyces spp. life cycle, different streptomycetes show different behaviors, and fermentation conditions need to be optimized for each specific strain and secondary metabolite. Cell-encapsulation constitutes an interesting alternative to classical fermentations, which was demonstrated to be useful in Streptomyces, but development under these conditions remained unexplored. In this work, the influence of cell-encapsulation in hyphae differentiation and actinorhodin production was explored in the model Streptomyces coelicolor strain. Encapsulation led to a delay in growth and to a reduction of mycelium density and cell death.

The high proportion of viable hyphae duplicated extracellular actinorhodin production in the encapsulated cultures with respect to the non-encapsulated ones. [Int Microbiol 2014; 17(2):75-80] Keywords: Streptomyces coelicolor · encapsulation · differentiation · antibiotics · cell death

Introduction Streptomycetes are Gram-positive, environmental soil bac­ teria that play important roles in the mineralization of organic matter. Streptomyces species are extremely important in biotechnology, given that approximately two thirds of all clinical antibiotics and several bioactive compounds are synthesized by members of this genus [25]. Streptomycetes are mycelial microorganisms with complex developmental cycles that include programmed cell death and sporulation

Corresponding author: A. Manteca Microbiology Section. Department of Functional Biology and IUOPA School of Medicine, University of Oviedo 33006 Oviedo, Spain Tel. +34-985103555 E-mail: mantecaangel@uniovi.es *

[7,32–34]. A young, compartmentalized vegetative mycelium (MI) differentiates into a multinucleated mycelium (MII); MII corresponds to the differentiated stage expressing genes and proteins related to secondary metabolism and sporulation [7,32–34].

Most processes for secondary metabolite production are performed in liquid cultures, conditions in which most streptomycetes do not sporulate, but in which differentiation remains fundamental for secondary metabolite production [19,24]. Different streptomycetes have different behaviors in liquid cultures. Some of them, such as S. coelicolor or S. griseus, form large pellets and clumps [13], while others, such as S. clavuligerus, grow in dispersion [23], and some species such as S. venezuelae [11], can even sporulate. Fermentation conditions need to be optimized for each specific strain and secondary metabolite, and alternatives to the classical fermentations in liquid and solid cultures are highly demanded

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in industry in order to improve the production of secondary metabolites produced in small amounts, or to activate the production of cryptic clusters [3,9,17,20,26].

Cell-encapsulation constitutes one fermentation alternative which was demonstrated to be useful in bacterial and fungal fermentations [22,30], but whose potential applications have only just begun to be explored in Streptomyces. Cellimmobilization was demonstrated to be useful in the production of cephamycin C [8] or clavulanic acid [14,27]

by S. clavuligerus; neomycin by S. marinensis [1,28], or S. fradiae [16]; and α-galactosidase by S. griseoloalbus [2]; to name just a few examples. However, the influence of encapsulation in Streptomyces hyphae differentiation has remained unexplored.

The main objective of this work was to analyze Streptomyces differentiation growing into alginate capsules, focusing on their potential industrial applications. Streptomyces coelicolor, one of the best-characterized Streptomyces strains [6], was used as a model. S. coelicolor produces various secondary metabolites, including undecylprodigiosines and actinorhodines, two colored antibiotics (red and blue respectively) whose production can be easily visualized [5].

In addition, γ-actinorhodin is secreted to the extracellular medium [5] and can be easily recovered from the extracellular medium between the mycelial capsules.

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in the CaCl2 solution for one hour, before being washed with 0.9 % NaCl solution to remove any excess CaCl2. As discussed below, modification of these standard encapsulation conditions did not significantly change the development of Streptomyces.

Streptomyces coelicolor encapsulated and non-encap­ sulated cultures. In the case of encapsulated cultures, 10 ml of mycelial capsules (7.5 ml intra-capsule volume, 2.5 ml of dead volume) were used to inoculate 90 ml of R5A sucrose-free culture medium. In the case of the control non-encapsulated cultures, the original pre-culture (72 h cultures, inoculated with spores) was processed as reported for the encapsulated cultures (see above), but was resuspended in fresh R5A sucrose-free medium without alginate; 7.5 ml were used to inoculate 92.5 ml of R5A sucrose-free culture medium. Flasks of 500 ml, were incubated at 30 ºC and 200 rpm in both cases. Two biological replicates were performed for each condition.

Streptomyces sampling throughout the fermentations.

Samples (one milliliter) were collected at different developmental time points and centrifuged at 7740 ×g for 10 min at 4 ºC. Supernatants were collected and constituted the extracellular samples. The pellets were resuspended in 1ml of 0.5M NaOH, boiled for 10 min, and cellular debris was removed by centrifugation (at 7740 ×g for 15 min at 4 ºC), in order to obtain the intra­ cellular samples.

Actinorhodin quantification. Actinorhodin was quantified spectro­ photometrically according to Bystrykh et al. [5]. Sodium alginate interfered with spectrophotometric measurements, and an extraction step was necessary to eliminate the alginate. Supernatants from 1ml of cultures (extracellular samples) were extracted twice with 1 volume of ethyl acetate containing 1 % formic acid. Extracts were vacuum-dried, and resuspended in 1 ml of NaOH 1 N. Actinorhodin was quantified spectrophotometrically with a UV/visible spectrophotometer (Shimadzu, Model UV-1240), applying the linear Beer– Lambert relationship to estimate concentration (ε640 = 25,320).

Protein quantification. Protein was quantified using the Bradford method [4] with bovine serum albumin (Sigma) as the standard.

Materials and methods Strain, media and pre-culture conditions. The strain used in this work was S. coelicolor M145. Spores were harvested from 7 days SFM plates [12] grown at 30 ºC, and stored in 30 % glycerol for long-term preservation at –80 ºC. Pre-cultures for Streptomyces immobilization were performed in liquid medium as previously reported [19]: freshly prepared spores were used to inoculate 100 ml sucrose-free R5A medium [10] at a final concentration of 107 spores/ml. Flasks were incubated at 30 ºC and 200 rpm for 72 h.

Streptomyces immobilization in sodium alginate. Sodium alginate (Sigma, A0682) was prepared in sucrose-free R5A medium to a final concentration of 2 %. The solution was heated at 45 ºC under agitation, until alginate was completely dissolved. The alginate solution could not be autoclaved [15], but was boiled for 5 min which was enough to allow sterilization. Alginate stock solution was cooled to room temperature before use.

All of the encapsulation steps were performed at room temperature under sterile conditions. With the exception of the alginate solution (see above), all of the solutions and pumping tubes were autoclaved. Mycelium from 100 ml of the pre-culture (72 h, inoculated with spores) was harvested by centrifugation (7900 ×g), washed with fresh sucrose-free R5A medium, and resuspended in 10 ml of sucrose-free R5A, which were further diluted with other 90 ml of sucrose-free R5A-2 % alginate (final volume of 100 ml, alginate concentration of 1.8 %). This mixture was pumped using a peristaltic pump through six hypodermic syringes with 0.9 mm diameter and 40 mm length, and dropped into sterile 2.5 % CaCl2 solution to solidify the alginate and form the capsules (3 mm average diameter). Capsules were hardened

Viability staining. Culture samples were obtained and processed for microscopy at different incubation times. In the case of non-encapsulated control cultures, samples were stained directly, while in the encapsulated cultures, capsules were cut manually into slices with a scalpel. Cells were stained with a cell-impermeable nucleic acid stain (propidium iodide, PI) in order to detect the dying cell population and with SYTO 9 green fluorescent nucleic acid stain (LIVE/DEAD Bac- Light Bacterial Viability Kit, Invitrogen, L-13152) to detect viable cells. The SYTO 9 green fluorescent stain labels all of the cells, i.e. those with intact membranes, as well as those with damaged membranes. In contrast, PI only penetrates bacteria with damaged membranes, decreasing SYTO 9 stain fluorescence when both dyes are present. Thus, in the presence of both stains, bacteria with intact cell membranes appear to fluoresce green, whereas bacteria with damaged membranes appear red. After incubating them for at least 1 minute in the dark, the samples were examined under a Leica TCS-SP2-AOBS confocal laser-scanning microscope at a wavelength of either 488 nm or 568 nm excitation and 530 nm (green) or 630 nm (red) emission, respectively (optical sections about 0.2 µm). Images were mixed using Leica Confocal Software.

In some cases, samples were also examined in the differential interference contrast mode, which is available using the same equipment.

Unstained samples were used as controls to determine the minimum photomultiplier tube (PMT) gain necessary to detect autofluorescence in the confocal microscope. The interference of green autofluorescence [31] was negligible when compared to the SYTO9 green fluorescence and did not interfere appreciably with the green fluorescent fluorochromes. Tenconi et al. [29] have recently demonstrated the existence of red autofluorescence associated with undecylprodigiosin that displays an excitation-emission spectrum similar to PI. Under the experimental conditions used in this work,

CELL IMMOBILIZATION OF S. COELICOLOR

red autofluorescence was significantly weaker than PI fluorescence and the minimum PMT gain necessary to observe it was 860 volts (using the 63x objective), which is 60 % more than the PMT gain used to observe PI fluorescence (535 volts under the 63x objective) (data not shown). Despite this, red autofluorescence was not negligible, and some of the red fluorescent background detected at later time points in the centers of mycelial pellets and capsules may have been derived from undecylprodigiosin.

More than 30 images were analyzed for each developmental time point using a minimum of three independent cultures.

Results and discussion

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used in laboratory flasks [19]. Liquid cultures were inoculated with spores; they were grown until differentiation of the second multinucleated antibiotic-producing mycelium (72 h) [19], and were then split into two: the first sample was inoculated directly into fresh liquid medium (non-encapsulated control culture) (Fig. 1), and the second was encapsulated prior to inoculation in fresh liquid medium (encapsulated culture) (Fig. 2). (See Methods for details.)

Development in the non-encapsulated control cultures was comparable to that previously observed in liquid cultures inoculated with spores [19]. Viable hyphae resumed growth, increasing the diameter of mycelial pellets from 480 µm to 600 µm (compare Fig. 1F with Fig. 1J), and there was a high proportion of dead hyphae in the center of mycelial pellets at any time (red staining in Fig. 1F–J). Hyphae showed the appearance of the second multinucleated mycelia (they lacked the discontinuities characterizing MI hyphae) [19] (Fig. 1K–O). The most important differences with respect to liquid cultures inoculated

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Confocal laser-scanning fluorescence micros­ copy (CLSM) analysis of Streptomyces coeli­co­ lor encapsulated and non-encapsulated li­quid cultures. In order to facilitate a comparison of Streptomyces differentiation in encapsulated liquid cultures with differentiation previously reported in non-encapsulated cultures, the culture conditions used were similar to those previously

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Fig. 1. Development of Streptomyces coelicolor M145 control non-encapsulated cultures. Upper pannels: macroscopic view of cultures. Undecylprodigiosin (red) and actinorhodin (blue) productions are visible. Lower panels: CLSM microscopy analysis. (A–E) interference contrast mode images; (F–O) fluorescence images (SYTO9/PI staining). Developmental time points are indicated.

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Fig. 2. Development of Streptomyces coelicolor M145 encapsulated cultures. Upper panels, macroscopic view of the capsules. Undecylprodigiosin (red) and actinorhodin (blue) productions are visible. Lower panels, CLSM microscopy analysis. (A–E) interference contrast mode images; (F–O) fluorescence images (SYTO9/PI staining). Developmental time points are indicated. Arrows indicate the border of the capsules.

with spores were: first, most of the mycelial growth was observed in the center of the pellets in the form of spots of viable hyphae (green staining in Fig. 1I) growing between dead hyphae (red staining in Fig. 1 and Fig. 1I); and second, mycelial pellets were much more fragile and appeared highly fragmented (Fig. 1B–E). This instability is probably due to the aging of the pellets, and the subsequent disintegration of dead hyphae which formed an important part of the pellet structure that had been formed in the original cultures used for inoculation [19].

Growth in the encapsulated cultures showed important differences with respect to the non-encapsulated ones. Mycelial pellets were completely destroyed during the encapsulation, and viable hyphae were dispersed around the entire capsule (green spots in Fig. 2F) at the beginning of the culture. The extension of cell death was lower, and the presence of dead hyphae only was significant at 0 h, derived from the original inocula (Fig. 1K), and at late time points (86 h) (red staining in Fig. 2O). Note that, despite the fact that the mycelia were growing embedded in a solid surface (alginate), there was no sporulation. As in

the case of the control non-encapsulated cultures, viable hypha showed the appearance of the second multinucleated mycelia (Fig. 2K–O). In summary, the most spectacular difference between encapsulated and non-encapsulated cultures was, that capsules were fully of viable second multinucleated mycelia (Fig. 2F–J), while in the case of the non-encapsulated cultures, a high proportion of hyphae in the center of the mycelial pellets were dead (Fig. 1F–J) [19]. As discussed below, this interesting result might be related to the growth of the hyphae inside the alginate matrix which maintain them at lower densities than in the non-encapsulated cultures, reducing stresses related to low nutritional/oxygen levels.

Growth and extracellular actinorhodin produc­ tion of Streptomyces coelicolor M145 in encap­ sulated and non-encapsulated liquid cultures.

Growth rate was 1.5-fold lower in the encapsulated cultures (0.0077 mg ml–1 h–1) than in the non-encapsulated ones (0.0117 mg ml–1 h–1) (Fig. 3A), and maximum growth was also lower

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Fig. 3. Time-course of growth (A) and γ-actinorhodin production (B) in encapsulated and non-encapsulated cultures. Values are the average ±SD from two biological replicates. Growth rates (slopes) in (A) to non-encapsulated: 0.0117 mg ml–1 h–1; to encapsulated: 0.0077 mg ml–1 h–1.

in the encapsulated cultures (0.47 mg of protein per ml) than in the non-encapsulated cultures (0.62 mg of protein/ml) (Fig. 3A). The slow growth observed in encapsulated cultures might be a consequence of the difficulty for hyphae to grow inside the alginate matrix, and it might be the reason for the low levels of cell death observed in these conditions, since low cell densities might reduce the stresses triggering cell death [18].

Second mycelium hyphae (MII) produced antibiotics in the encapsulated and non-encapsulated cultures from the beginning of the experiment (red and blue colors in the macroscopic views shown in Figs. 1 and 2). Mycelium encapsulation has obvious advantages for the production of extracellular secondary metabolites, but not in the case of intracellular ones, since the recovery of intracellular compounds from encapsulated cultures is more complicated than from non-encapsulated ones. For this reason, extra­cellular levels of γ-actinorhodin [5] were analyzed (Fig. 3B). The production of γ-actinorhodin was slower in the encapsulated cultures (starting at 48 h) than in the non-encapsulated ones (starting at 38 h) (Fig. 3B). However, the maximum levels of production were duplicated in the encapsulated cultures (Fig. 3B).

These data correlate with growth curves and microscopic observations: encapsulation led to a slower rate of mycelial growth (Fig. 3A), and to larger proportion of viable MII antibiotic producing hyphae (compare the proportions of green and red stained hyphae in Figs. 1J and 2J) which were producing more antibiotics than the non-encapsulated cultures (Fig. 3B).

Different encapsulation conditions (concentrations of alginate and CaCl2, time of capsule hardening in CaCl2) were tested, but they did not significantly modify the development of S. coelicolor described above (data not shown). Inoculation density is one of the most well-known fermentation variables, that is usually conducive to modifications in growth and production [19,24]. Different inoculations modify the final actinorhodin production levels and growth kinetics, but not the general development described in this work in which cell death was reduced and antibiotic production increased in the encapsulated cultures with respect to the non-encapsulated ones (data not shown).

Results overview and conclusions. One important drawback of Streptomyces fermentations is the need to optimize culture conditions in order to obtain the most optimal equilibrium between differentiation, cell death and secondary metabolite production [19,21,24]. Cell-encap­sulation cons­ titutes an alternative to classical fermentations in liquid or solid cultures, which, in S. coeli­color, was demonstrated to maximize the proportion of viable antibiotic producing hyphae (MII), reporting the maximum levels of actinorhodin production even with lower levels of total biomass. In addition, cell immobilization has intrinsic advantages facilitating the separation between culture medium and cells, and recycling the productive biomass. Further work will be necessary to evaluate the applicability of cell immobilization in order to improve and/or activate secondary metabolite production in other streptomycetes.

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Acknowledgments. We wish to thank the European Research Council (ERC Starting Grant; Strp-differentiation 280304) for financial support, Beatriz Gutiérrez Magán (Universidad de Oviedo, Departamento de Biología Funcional, Área de Microbiología) for laboratory assistance, students of Biotechnology (Universidad de Oviedo, Class of 2013) for their collaboration in the preliminary experimentation, and Proof-Reading-Service.com for proofreading the text.

Conflit of interests. None declared.

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Biochem Eng J 39:131-136 15. Leo WJ, McLoughlin AJ, Malone DM (1990) Effects of sterilization treatments on some properties of alginate solutions and gels. Biotechnol Prog 6:51-53 16. Machado I, Teixeira JA, Rodríguez-Couto S (2013) Semi-solid-state fermentation: a promising alternative for neomycin production by the actinomycete Streptomyces fradiae. J Biotechnol 165:195-200

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17. Manivasagan P, Venkatesan J, Sivakumar K, Kim SK (2014) Pharma­ ceutically active secondary metabolites of marine actinobacteria. Micro­ biol Res 169:262-278 18. Manteca A, Fernandez M, Sanchez J (2006) Cytological and biochemical evidence for an early cell dismantling event in surface cultures of Streptomyces antibioticus. Res Microbiol 157:143-152 19. Manteca A, Álvarez R, Salazar N, Yagüe P, Sánchez J (2008) Mycelium differentiation and antibiotic production in submerged cultures of Streptomyces coelicolor. Appl Environ Microbiol 74:3877-3886 20. Marinelli F (2009) Antibiotics and Streptomyces: The future of antibiotic discovery. Microbiol Today: February Issue:21-23 21. Olmos E, Mehmood N, Haj Husein L, Goergen JL, Fick M, Delaunay S (2013) Effects of bioreactor hydrodynamics on the physiology of Streptomyces. Bioproc Biosyst Eng 36:259-272 22. Pham-Hoang BN, Romero-Guido C, Phan-Thi H, Waché Y (2013) Encapsulation in a natural, preformed, multi-component and complex capsule: yeast cells. Appl Microbiol Biotechnol 97:6635-6645 23. Pinto LS, Vieira LM, Pons MN, Fonseca MM, Menezes JC (2004) Morphology and viability analysis of Streptomyces clavuligerus in industrial cultivation systems. Bioprocess Biosyst Eng 26:177-184 24. Rioseras B, López-García MT, Yagüe P, Sánchez J, Manteca A (2014) Mycelium differentiation and development of Streptomyces coelicolor in lab-scale bioreactors: Programmed cell death, differentiation, and lysis are closely linked to undecylprodigiosin and actinorhodin production.

Bioresource Technol 151:191-198 25. Ruiz B, Chávez A, Forero A, García-Huante Y, Romero A, Sánchez M, Rocha D, Sánchez B, Rodríguez-Sanoja R, Sánchez S, Langley E (2010) Production of microbial secondary metabolites: Regulation by the carbon source. Crit Rev Microbiol 36:146-167 26. Sánchez J, Yagüe P, Manteca A (2012) New insights in Streptomyces fermentations. Ferment Technol 1:2 27. Saudagar PS, Shaligram NS, Singhal RS (2008) Immobilization of Streptomyces clavuligerus on loofah sponge for the production of clavulanic acid. Bioresource Technol 99:2250-2253 28. Srinivasulu B, Prakasham RS, Jetty A, Srinivas S, Ellaiah P, Ramakrishna SV (2002) Neomycin production with free and immobilized cells of Streptomyces marinensis in an airlift reactor. Process Biochemistry 38:593-598 29. Tenconi E, Guichard P, Motte P, Matagne A, Rigali S (2013) Use of red autofluorescence for monitoring prodiginine biosynthesis. J Microbiol Methods 93:138-143 30. Westman JO, Ylitervo P, Franzén CJ, Taherzadeh MJ (2012) Effects of encapsulation of microorganisms on product formation during microbial fermentations. Appl Microbiol Biotechnol 96:1441-1454 31. Willemse J, van Wezel GP (2009) Imaging of Streptomyces coelicolor A3(2) with reduced autofluorescence reveals a novel stage of FtsZ localization. PLoS One 4:e4242 32. Yagüe P, López-García MT, Rioseras B, Sánchez J, Manteca A (2013) Pre-sporulation stages of Streptomyces differentiation: state-of-the-art and future perspectives. FEMS Microbiol Lett 342:79-88 33. Yagüe P, Rodríguez-García A, López-García MT, Martín JF, Rioseras B, Sánchez J, Manteca A (2013) Transcriptomic analysis of Streptomyces coelicolor differentiation in solid sporulating cultures: first compartmentalized and second multinucleated mycelia have different and distinctive transcriptomes. PLoS One 8:e60665 34. Yagüe P, Rodríguez-García A, López-García MT, Rioseras B, Martín JF, Sánchez J, Manteca A (2014) Transcriptomic analysis of liquid nonsporulating Streptomyces coelicolor cultures demonstrates the existence of a complex differentiation comparable to that occurring in solid sporulating cultures. PLoS One 9:e86296

RESEARCH ARTICLE International Microbiology (2014) 17:81-90 doi:10.2436/20.1501.01.210 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

Phenotypic comparison of clinical and plantbeneficial strains of Pantoea agglomerans Anna Bonaterra,1* Esther Badosa,1 Fabio Rezzonico,2,3 Brion Duffy,2,3 Emilio Montesinos1 1 Institute of Food and Agricultural Technology, University of Girona, Girona, Spain. 2Agroscope Changins-Wädenswil ACW, Division of Plant Protection, Wädenswil, Switzerland. 3Zurich University of Applied Sciences ZHAW, LSFM-IUNR, Environmental Genomics and Systems Biology Research Group, Wädenswil, Switzerland

Received 12 March 2014 · Accepted 4 June 2014

Summary. Certain strains of Pantoea are used as biocontrol agents for the suppression of plant diseases. However, their commercial registration is hampered in some countries because of biosafety concerns. This study compares clinical and plantbeneficial strains of P. agglomerans and related species using a phenotypic analysis approach in which plant-beneficial effects, adverse effects in nematode models, and toxicity were evaluated. Plant-beneficial effects were determined as the inhibition of apple fruit infection by Penicillium expansum and apple flower infection by Erwinia amylovora. Clinical strains had no general inhibitory activity against infection by the fungal or bacterial plant pathogens, as only one clinical strain inhibited P. expansum and three inhibited E. amylovora. By contrast, all biocontrol strains showed activity against at least one of the phytopathogens, and three strains were active against both. The adverse effects in animals were evaluated in the plant-parasitic nematode Meloidogyne javanica and the bacterial-feeding nematode Caenorhabditis elegans. Both models indicated adverse effects of the two clinical strains but not of any of the plant-beneficial strains. Toxicity was evaluated by means of hemolytic activity in blood, and genotoxicity with the Ames test. None of the strains, whether clinical or plant-beneficial, showed any evidence of toxicity. [Int Microbiol 2014; 17(2):81-90] Keywords: Pantoea agglomerans · Erwinia amylovora · Meloidogyne javanica · Penicillium expansum · Caenorhabditis elegans · biocontrol · biosafety · toxicity · hemolytic activity · Ames test

Introduction Pantoea agglomerans (formerly known as Enterobacter agglomerans, Erwinia herbicola, or Erwinia milletiae) is a Gram­-­negative enterobacterium that has been subjected to numerous taxonomic rearrangements, grouping strains of diCorresponding author: A. Bonaterra Institute of Food and Agricultural Technology University of Girona Campus Montilivi 17071 Girona, Spain E-mail: anna.bonaterra@udg.edu *

verse ecological origin [3,9,10,18,20,22,33,34]. P. agglome­ rans is a ubiquitous epiphytic bacterium found on a widerange of plant species. It is also frequently isolated from animal, aquatic, and soil environments [1,24,25,35]. Certain strains of P. agglomerans and P. vagans (formerly included in P. agglome­rans) isolated from plant environments are among the most beneficial biological control agents for the suppression of plant diseases caused by phytopathogenic bacteria and fungi [7,27,28,30,41,53,56,57]. Several such strains have been developed as active ingredients of microbial biopesticides registered as plant protection products against fire blight caused by Erwinia amylovora [38].

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Clinical reports have implicated P. agglomerans as an opportunistic human pathogen, but these have typically been descriptive, indicate polymicrobial isolations, and lack verification of pathogenicity or demonstrate a role for the bacte­ rium in disease [4,14,32,55]. Inaccurate identification of many clinical strains [48] has further exaggerated the association of P. agglomerans with human infections. Although sometimes reported as a plant pathogen [14,23,47], only strains belonging to the pathovars gypsophilae and betae have been shown to be truly phytopathogenic, as defined by Koch’s postulates [13,37]. Plasmids with virulence factors (e.g., type III secretion system genes) that are responsible for phytopathogenicity [37] are carried by both strains but not by other plant and clinical strains [48].

Regulatory decisions for the registration of biopesticides as alternatives to chemical/antibiotic plant protection pro­ ducts rely upon available data [39]. Currently, P. agglomerans is classified in Europe as a biosafety level 2 species, precluding its consideration for beneficial applications (EU Directive 2000/54/EC). However, only a few studies have compared beneficial and clinical strains, in contrast to the many studies of other bacterial species, e.g., Serratia marcescens [62], Burkholderia cepacia, [5,42,59], and Pseudomonas aeruginosa [16].

Recent reports of comparative molecular and biochemical analyses found no clear distinction between clinical and plant P. agglomerans sensu stricto strains. Clinical and biocontrol strains clustered together according to standard microbiological, metabolic, or biochemical characteristics [48], pattern polymorphisms of total or partial genomic DNA (fAFLPs, ITS, and ERIC/REP-DNA), single-locus sequence analyses

[45,48], or DNA-DNA hybridization [8]. Clinical and biocontrol strains also showed no difference in their ability to colonize soybean roots or embryonated chicken eggs [58]. Registration in the USA and Canada of plant strains C9-1 (Blight Ban C9-1, NuFarm) and E325 (BloomTime, Verdesian Life Sciences) and in New Zealand of P10c (BlossomBless, GroChem New Zealand Ltd.) demonstrated a lack of animal pathogenicity, allergenicity, and toxicity; however these data are proprietary and do not extend to strain comparisons.

The aim of our study was to complement genotypic analyses with phenotypic comparisons between Pantoea clinical strains and plant-beneficial strains. The strains were compared based on hemolysis, genotoxicity and nematode infectivity. Differential adaptation to plant habitats was evaluated using biocontrol models against fungal and bacterial phytopathogens.

Materials and methods Pantoea agglomerans strains and growth conditions.

Twelve P. agglomerans strains and P. vagans C9-1 were used, including clinical isolates and plant epiphytes previously described to have beneficial activity as biocontrol agents against plant diseases (Tables 1 and 2). The clinical isolates included the type strain ATCC 27155 (syn. LMG 1286) as well as other strains from research or culture collections. Plant-beneficial bacteria were those typical of most biocontrol strains that are either commercial or have been the subject of considerable research. All strains were previously identified as P. agglomerans based upon the results of biochemical tests. According to 16S rRNA gene sequences, the strains belonged to P. agglomerans (including P. vagans strain C9-1) [48], except for EM13cb and EM17cb, which are closely related to P. agglomerans but are not members of this species. Based on Phoenix analysis, they are P. agglomerans but along with 16S rRNA sequencing both gyrB analysis and MALDI-TOF mass spectrometry

Table 1. Relevant characteristics of strains of plant-beneficial Pantoea agglomerans and P. vagans used in the present work Strain

Plant host/material

Country of origin

Target disease

Toxicology CFU/kga

CPA-2

apple fruit surface

Spain

postharvest rot

4.3 × 1011

nd + + +

EPS125

pear fruit surface

Spain

postharvest rot

>10

nd + + +

C9-1

apple stem

USA

fire blight

5 × 1011

+ + – –

P10c

apple flower

New Zealand

fire blight

nr

nd + + +

Eh252

Malus pumila

USA

fire blight

nr

+ + – –

Eh318

apple stem

USA

fire blight

nr

nd + + +

Eh1087

apple flower

New Zealand

fire blight

nr

nd + + +

10

Identification as P. agglomerans (Phoenix, 16S rRNA, gyrB, MALDI-TOF MS)b

Toxicology, acute oral toxicity on mammals; nr, not reported; strain CPA-2 in ref. [41]; strain EPS125 in ref. [7]; strain C9-1 in US-EPA Code 006470 [www.epa.gov]. b According to ref. [48,49]. +, positive result; –, negative result; nd, not done. a

COMPARISON OF STRAINS OF P. AGGLOMERANS

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Table 2. Relevant characteristics of strains of Pantoea agglomerans of clinical origin used in the present work Strain

Country of origin

Material of origin

Nature of the infection

Identification as P. agglomerans (Phoenix, 16S rRNA, gyrB, MALDI-TOF MS)b

ATCC 27155 (LMG 1286)

Zimbabwe

knee

laceration

+ + + +

CIP A181

France

blood

bacteremia

+ + + +

VA21971

Switzerland

wound

infected wound

+ + + +

EM13cba

Spain

blood

bacteremia

+ – – –

EM17cb

Spain

blood

bacteremia

+ – – –

EM22cb

Spain

blood

bacteremia

+ + + +

a

Based on Phoenix these strains are P. agglomerans but 16S rRNA, gyrB and MALDI-TOF mass spectrometry do not support that identification; instead they belong to Pantoea conspicua for EM13cb and Pantoea anthophila for EM17cb. b According to ref. [22,48,49]. +, positive result; –, negative result. a

place them close to but nonetheless distinct from P. agglomerans. Rather, based on the binary data, EM13cb is Pantoea conspicua and EM17cb Pantoea anthophila. Sequence analysis of housekeeping genes including gyrB confirmed the identity of P. agglomerans strains and assigned C9-1 to P. vagans. Strains for this study were chosen to be representative of different environments of isolation (human/plant, tissue, country) and of the variability in relation to biochemical properties (Phoenix, MALDI-TOF mass spectrometry) [49] and gyrB sequences [48]. Strains were recovered from cultures preserved at –80 ºC and cultured overnight on Luria-Bertani (LB) agar at 25 ºC. Colonies were scraped from the agar surface and suspended in sterile distilled water. The cell culture was adjusted to a cell density corresponding to 1 × 108 CFU/ml. Appropriate concentrations were prepared by dilution with sterile distilled H2O. Biocontrol of blue mould, a postharvest decay caused by Penicillium expansum. Penicillium expansum EPS46 [21] was used for the blue mould infection inhibition assay and maintained on potato dextrose agar (PDA) at 4 ºC. Conidia were collected from 7-day PDA cultures incubated at 25 ºC in darkness. The colonies were scraped with a moist cotton swab and resuspended in distilled H2O containing 0.5 % Tween 80. Spore concentration was adjusted to 104 spores/ml using a hemocytometer, and a fresh suspension was used for each trial. Apples (Malus × domestica L. ‘Golden Delicious’) were surface-disinfected for 1 min by immersion in dilute sodium hypochlorite (1 % active chlorine), washed twice in sterile H2O, and air-dried prior to use. Fruit were wounded with a flame-sterilized 3-mm diameter cork-borer to a uniform depth of 5-mm at three equidistant points around the middle and then placed on polystyrene mats in plastic incubation boxes. Each wound was first inoculated with a 50 µl of a bacterial suspension of a Pantoea strain (1 × 108 CFU/ml) and incubated for 24 h at 20 °C in sealed boxes with high humidity (ca. 100 % RH). Each wound was then inoculated with 20 ml of a P. expansum (1 × 104 conidia/ml) suspension and incubated for 5 days. Non-treated controls consisted of fruit inoculated with water or with the pathogen alone. Treatments consisted of three replicates with three pieces of fruit each (three wounds) and arranged in a completely randomized design with two independent trials. Disease incidence for each replicate was determined as the percentage of wounds infected 5 days after inoculation.

Biocontrol of fire blight caused by Erwinia amylovora. Biocontrol activity against E. amylovora EPS101 was evaluated using apple flowers, following the methods of Cabrefiga and Montesinos [11]. Newly

opened pear flowers were collected from an experimental orchard at the Mas Badia Agricultural Experiment Station (Girona, Spain). Individual flowers were placed with the cut peduncle submerged in 1 ml of a 10 % sucrose solution in a single 1.5-mL Eppendorf plastic tube [45]. Tubes containing flowers were supported in tube racks placed in incubation boxes. Flower hypanthia were treated with 20 ml of a Pantoea suspension (1 × 108 CFU/ml) and incubated overnight at 20 ºC in sealed boxes with high relative humidity (ca. 100 % RH). The flowers were then inoculated with 10 ml of a 1 × 107 CFU/ml suspension of E. amylovora deposited on the hypanthia and incubated for 5 days as described above. Non-treated controls consisted of water or pathogen-alone treatments. Treatments consisted of three replicates with eight flowers, and were arranged in a completely randomized design with two trials. Disease severity was evaluated 5 days after inoculation with a severity scale from 0 to 3, in which 0 indicated no symptoms; 1, partial hypanthia necrosis; 2, total hypanthia necrosis; and 3, necrosis progression through the peduncle. Data analysis included calculation of the mean disease severity for each replicate and maximum severity observed within an experiment, as described previously [6].

Adverse effects on nematodes. Toxicity of the Pantoea strains to the plant-parasitic nematode Meloidogyne javanica and pathogenicity to the bacterial-feeding nematode Caenorhabditis elegans were studied.

A population of M. javanica was maintained on the root system of tomato plants (Lycopersicon esculentum L. [‘Rio Grande’] in the greenhouse by periodic transfer to new plant material every 2-3 months. Prior to each trial, M. javanica eggs were collected from root galls following Cobb’s method [50]. The volume of suspension collected was measured and the egg concentration was determined using a counting chamber at 40× magnification (Olympic Equine Products, Issaquah, WA, USA). M. javanica egg suspensions were disinfected by adding sodium hypochlorite (3 % active chlorine) to the suspension and gently agitating for 20 min. Eggs were collected on a sterile 500-mesh sieve and washed with sterile distilled H2O to remove residual sodium hypochlorite. Eggs were aerated at 20 ºC for 48 h to induce hatching; stage J2 juveniles were collected and the concentration adjusted.

Bacterial cell suspensions and cell-free culture supernatants were prepared from 48-h P. agglomerans liquid LB cultures centrifuged at 10,000 ×g for 15 min. Cell pellets were resuspended in glucose minimal medium (GMM) containing (per liter) 5 g of glucose, 1 g NH4Cl, 3 g KH2PO4, 2.4 g Na2HPO4, 0.5 g NaCl, and 0.2 g MgSO4 at pH 7. The concentration was adjusted to 1 × 108 CFU/ml.

Culture supernatants (bacterial culture extracts) were separated from the pel-

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lets and filtered through a 0.22-µm pore size filter membrane. Toxicity was assayed by placing 100 µl of the M. javanica J2 suspension (2000 juveniles/ ml) in 30-ml sterile tubes, and adding either 100 µl of bacterial suspension in 1.8 ml of sterile GMM or 1.9 ml of cell-free culture supernatant. Negative control treatments consisted of J2 M. javanica incubated with only GMM. A positive control treatment was included, consisting of the chemical nematicide Vydate P (100 mg/l, Du Pont Ibérica, Barcelona, Spain). The tubes were sealed with Parafilm and incubated for 24 h at 25 ºC in darkness. Toxicity was determined as the concentration of viable J2 nematodes by observation at 40× magnification. Dead individuals were determined based on immobility and the presence of straight rigid bodies. Treatments consisted of three replicate tubes arranged in a completely randomized design. Two independent trials were conducted.

Caenorhabditis elegans SS104 was maintained on nematode growth medium (NGM) prepared as follows: 3 g of NaCl, 17 g of agar, and 2.5 g of peptone were added to 975 ml of distilled H2O. The solution was autoclaved and then cooled to 55 °C. One ml of 1 M CaCl2, 1 ml of 5 mg cholesterol/ml (prepared in ethanol), 1 ml of 1 M MgSO4, and 25 ml of 1 M KPO buffer (108.3 g KH2PO4, 25.6 g K2HPO4, H2O to 1 liter, pH 6). Escherichia coli OP50 was used as the food source. Strain SS104 is a temperature-sensitive mutant unable to reproduce when incubated at 25 ºC, which ensured a constant number of nematodes during the assay. The nematode age distribution was synchronized before the experiment by a bleaching procedure as previously described [43]. Briefly, the surface of agar plates containing eggs was washed with 5 ml sterile M9 buffer, centrifuged at 1500 ×g for 2 min, and the pellet suspended in 4.5 ml of bleaching solution (0.5 ml water, 2.5 ml sodium hydroxide 1 M, and 4 ml sodium hypochlorite ~4 %). The tube was gently mixed intermittently for approximately 5 min to kill all nematode forms except eggs. The reaction was stopped by centrifugation for 1 min at 1500 ×g.

The pellet was washed three times with M9, resuspended, and then incubated for 14 h at 25 ºC with gentle agitation. Synchronized nematodes (L1 larval stage) were placed on NGM agar plates with E. coli OP50 and incubated at 25 ºC for 2 day to obtain stage L4 adult nematodes, that were recovered from the agar plates as described above. Bacterial cell suspensions of the Pantoea strains, E. coli OP50, and Salmonella enterica CECT 4595 (ATCC 14028), used as positive controls in this test, were plated on NGM and grown for 24 h at the appropriate temperature. For the survival assay, collected worms were transferred to fresh lawn plates (150 per plate) of the bacteria (treatments).

Each assay was carried out in triplicate. For the assay with cell-free culture supernatants of the bacterial strains, cultures were grown for 48 h in LB broth at 28 ºC, except S. enterica CECT 4594, which was incubated at 37 ºC. The cultures were centrifuged at 10,000 ×g for 10 min and the supernatants were filter sterilized through a 0.22-µm pore size filter membrane. Four hundred µl of the effluent was dispensed into 24-well plates containing 50 µl of an E. coli OP50 suspension in M9 buffer and 50 µl of synchronized L4 nematodes (approximately 100 individuals). Worm mortality was scored over 7 days for bacterial cell suspension and 5 days for cell-free culture supernatants by observation at 40× magnification. Dead individuals were determined based on immobility and the presence of straight rigid bodies. The assay was carried out in triplicate. Sodium azide at 750 µM was used as a positive control.

Hemolytic activity. Hemolytic activity was evaluated in solid and liquid media. Streptococcus pyogenes ATCC 19615 and melittin were used as positive controls. In solid medium, hemolytic activity was scored by the presence of a clear halo around bacterial colonies plated in triplicate at the adequate dilutions on blood agar base (Oxoid Limited, Basingstoke, UK) with either 5 % (v/v) sheep or horse erythrocytes (Oxoid Limited) after incubation at 28 ºC for 48 h. In liquid medium, both cells suspended in Ringer’s and cellfree culture supernatants obtained after centrifugation at 10,000 × g for 10 min of 48-h cultures in LB broth were used. Hemolysis was evaluated by determining hemoglobin release from erythrocyte suspensions of sheep or horse blood (5 % v/v, Oxoid) [2]. Blood was centrifuged at 6000 ×g for 5 min,

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washed three times with Tris buffer (10 mM Tris, 150 mM NaCl, pH 7.2), and diluted ten-fold. Red blood cell aliquots (65 ml) were mixed with 65 ml of bacterial cell suspension (5 × 109 CFU/ml) or cell-free culture supernatants in triplicate in a 96-well reaction plate and incubated with continuous shaking for 1 h at 37 ºC. After the incubation, the tubes were centrifuged at 3500 ×g for 10 min. Supernatant aliquots (80 ml) were transferred to 100-well microplates (Oy Growth Curves Ab, Helsinki, Finland) and diluted with 80 ml of deionized H2O (Milli-Q, Millipore, Billerica, MA, USA). Hemolysis was measured as the absorbance at 540 nm using a Bioscreen C plate reader (Oy Growth Curves Ab). Complete hemolysis was determined in Tris buffer amended with melittin (100 mM) (Sigma-Aldrich, Madrid, Spain) as a positive control.

Genotoxicity. Prior to the Ames test, the culture supernatants were assayed to ensure that they were not cytotoxic to the Salmonella strains (survival above 50 %) [32,31]. The bacterial reverse mutation test (Ames test) was performed as described in the Test Guideline 471 (OECD) [19] using two bacterial strains of Salmonella typhimurium as reference strains (TA98, to detect frameshift mutations and TA100, for base-pair substitutions), without metabolic activation. Cell-free Pantoea culture supernatants were prepared as described above for the hemolytic activity test. Overnight cultures of two strains of Salmonella were prepared in LB broth. Genotoxicity was assayed by mixing 0.1 ml of Pantoea cell-free supernatants with 3 ml of overlay agar, 0.2 ml of histidine-biotin solution (0.5 mM), and 0.2 ml of S. enterica suspension at a concentration of 5 × 108 CFU/ml and plated in minimal medium.

Sodium azide at 1.25 and 2.5 µM was used as the mutagenic agent for strain TA100 and 2-nitrofluorene at 2.5 and 5.0 µM as the mutagenic agent for strain TA98. The assay was performed in triplicate. Genotoxicity was scored as positive when the ratio of induced to natural revertants was ≥2.

Statistical analysis. The significance of the effect of treatments was determined using a one-way analysis of variance. Means were separated using the Waller-Duncan test at P < 0.05. The analysis was performed with the GLM procedure of the PC-SAS (SAS Institute, Cary, NC, USA).

Results and Discussion Recent genotypic and biochemical analyses have provided limited discrimination of P. agglomerans biocontrol and clinical strains [8,15,48,58], whereas here we used phenotypic comparisons to obtain a greater degree of differentiation.

Thus, plant-beneficial strains could be clearly distinguished from clinical strains on the basis of antagonistic activity against plant pathogens. We found no evidence for the toxicity of any of the biocontrol strains, and some evidence for the pathogenicity of the two clinical strains using nematode models. These findings support the previously reported lack of genetic virulence determinants in clinical and biocontrol strains [48].

All seven plant-beneficial strains significantly suppressed at least one of the two plant diseases in assays with the commercial strains (C9-1 and P10c), and Eh252, which showed significant biocontrol activity in both assays. Only two plantbeneficial strains (Eh318 and Eh1087), both originally selected

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Fig. 1. Biological control of blue mould caused by Penicillium expansum on â&#x20AC;&#x2DC;Golden Deliciousâ&#x20AC;&#x2122; apples by clinical (black bars) and plant-beneficial (white bars) strains of Pantoea spp. Values are the mean of three replicates, each consisting of three fruits, each with three wounds. Error bars represent the 95 % confidence interval of the mean. Bars for blue mould rot incidence labeled with the same letter do not differ significantly (P < 0.05) according to the Waller-Duncan test.

2 and EPS125) failed to suppress the bacterial pathogen E. amylovora in flowers (Fig. 2), while all of the fire blight biocontrol agents were effective. The inhibitory activity observed

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as fire blight biocontrol agents, failed to suppress the fungal pathogen P. expansum (Fig. 1). Similarly, only the two strains originally selected as postharvest rot biocontrol agents (CPA-

Fig. 2. Biological control of fire blight on pear flower by Erwinia amylovora by clinical (black bars) and plant-beneficial (white bars) strains of Pantoea spp. Values are the mean of three replicates consisting of eight flowers. Error bars represent the 95 % confidence interval of the mean. Bars for fire blight severity labeled with the same letter do not differ significantly (P < 0. 05) according to the Waller-Duncan test.

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for the biocontrol strains is not surprising since they were previously selected in large screenings of environmental isolates specifically on the basis of their superior biocontrol activity against target pathogens [38]. By contrast, the six clinical Pantoea strains were generally ineffective or had only a weak effect in either the postharvest assay or the flower assay. Four out of the six clinical strains inhibited E. amylovora infections in flowers, although generally to a lesser extent than the biocontrol strains (Fig. 2).

Our results provide no indication of toxicity or pathogenicity in either nematode models for plant-beneficial strains and only some evidence for the two clinical strains (ATCC 27155 and EM22cb). Cells of most Pantoea strains had no effect on the survival of the plant-parasitic nematode M. javanica, with only the clinical strain EM22cb killing J2 juveniles to a degree even close to that of the nematicide control (Fig. 3). The pathogenicity of this strain to the bacterial-feeding nematode C. elegans was similar to that of Salmonella ATCC 4594 (Fig. 4). Except for cells of P. agglomerans ATCC 27155, which were also effective in killing C. elegans, the

Fig. 3. Effect of cells (upper panel) and cell-free culture supernatants (lower panel) of clinical (black bars) and plant-beneficial (white bars) strains of Pantoea on the survival of the plant parasitic nematode Meloidogyne javanica (J2 stage juveniles) compared to the chemical nematicide Vydate P (dashed bars). Error bars represent the 95 % confidence interval of the mean. Bars for survival labeled with the same letter do not differ significantly (P < 0.05) according to the Waller-Duncan test.

remaining strains did not have significant effects on nematode survival in either of the models (Figs. 3 and 4). Cell-free culture supernatants from clinical or biocontrol strains also had no effect on C. elegans survival (data not shown) whereas those from most of the biocontrol strains had inhibitory effects on M. javanica. Note that only the cell-free culture super足 natants of the clinical strain ATCC 27155, whose cells showed pathogenicity to C. elegans, also had adverse effects on M. javanica. Therefore, we conclude that the clinical strains EM22cb and ATCC 27155 exhibited adverse effects in both of the nematode models studied.

Nematode assays are widely accepted as simple models to study bacterial virulence mechanisms [12,36,61] and to identify toxic compounds [40]. Although C. elegans is most often used, Meloidogyne is also sensitive to infection and killing by a wide-range of bacteria, including other Enterobacteriaceae such as Enterobacter cloacae [17,51]. Production of the antibacterial compounds that contribute to the biocontrol activity of some P. agglomerans strains [28,29,44] could be a factor in the adverse effects of the cell-free culture supernatants on

plant-parasitic nematode killing, observed in our study.

Strains Eh252, Eh318, and P. vagans C9-1 produce the antibiotic pantocin A, and strain Eh318 also produce the antibiotic pantocin B [51,52,53,57,60]. The pantocin A biosyn-

Fig. 5. Hemolytic activity of clinical (left bars) and plant-beneficial (right bars) strains of Pantoea compared to a reference hemolytic Streptococcus pyogenes strain. The assay was performed using cells and cell-free culture supernatants of Pantoea strains together with sheep blood (upper panel) and horse blood (lower panel). Error bars of hemolysis represent the 95 % confidence interval of the mean.

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Fig. 4. Survival of Caenorhabditis elegans SS104 (adult stage) temperaturesterile mutants in the presence of cells of clinical strains of Pantoea: ATCC 27155 (up closed triangle), EM13cb (down closed triangle), EM17cb (closed circle), EM22cb (closed square), CIP A181 (closed diamond), VA21971 (closed hexagon), plant-beneficial strains CPA-2 (up open triangle), EPS125 (down open triangle), C9-1 (open circle), P10c (open square), Eh252 (open diamond), and Eh1087 (open hexagon) compared to the E. coli OP50 feeding strain (circle with Ă&#x17D;) and to Salmonella 4594 (Ă&#x17D;), pathogenic to mammals.

Error bars indicated the minimum significant difference according to the Waller-Duncan test.

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thetic genes paaABC are present in those plant-beneficial strains, but absent in the clinical strain EM22cb [48]. Another antimicrobial peptide, herbicolin I, is produced by the plantbeneficial strain C9-1 [27,29,51,52]. Phenazine is reportedly

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Fig. 6. Genotoxicity of cell-free culture supernatants of Pantoea strains (Ames test) compared to mutagenic agents. Mutant strains of Salmonella TA100 and TA98 were used as indicators. Sodium azide at 1.25 (LC) and 2.5 (HC) µM was used as the mutagenic agent to strain TA100 and 2-nitrofluorene at 2.5 (LC) and 5.0 (HC) µM as the mutagenic agent to strain TA98.

produced by the plant-beneficial strain Eh1087 [30], which also had adverse effects on M. javanica survival. The type III secretion system (T3SS) gene hrcN, an important factor in bacterial virulence against eukaryotes [26,46], is present in clinical strain VA2197, but absent in clinical strain EM22cb, the most lethal strain in the nematode assays [48]. It should be noted, however, that the T3SS present in P. agglomerans is phylogenetically more similar to the one present in biocontrol strains of Pseudomonas spp. than to the Inv/Spa system present in animal pathogens.

The data obtained in the present work with both nematode models are in agreement with the reported lethal oral dose of the biocontrol strains P. vagans C9-1 and P. agglomerans E325, CPA-2, and EPS125, which exceeded 108–1010 CFU/kg animal body weight in Sprague-Dawley CD rats [7,41, EPA Federal Registers 71:54928-54933 and 71:24590-24596]. Accordingly, the USA Environmental Protection Agency (EPA) has registered two commercial biopesticides containing strains C9-1 and E325 on the basis of several toxicological tests, labeling these strains as toxicity category IV (i.e., “practically non-toxic”).

None of the plant-beneficial or clinical Pantoea strains had significant hemolytic activity in red blood cell assays conducted in liquid or agar culture, compared to the S. pyogenes

ATCC 19915 positive control. In blood agar, a clear halo was observed and the percentage of hemolysis in the liquid assays for the supernatants was around 60������������������������� –������������������������ 65 % of the melittin hemolysis for S. pyogenes using sheep and horse blood (Fig. 5).

None of the cell-free culture supernatants, from either clinical or plant-beneficial Pantoea strains, showed genotoxicity to strains TA98 and TA100 in the Ames test (Fig. 6).

In conclusion, clinical Pantoea strains are indistinguishable from plant-beneficial strains on the basis of hemolytic or genotoxicity tests. However, the plant models permit differentiation of plant-beneficial from clinical strains. While the nematode models provided some proof of the adverse effects of the two clinical strains (ATCC 27155, EM22cb), negative effects were not observed with the plant-beneficial strains P. agglomerans and P. vagans C9-1. Finally, there was no evidence suggesting the toxicity of any of the plant-beneficial or clinical strains tested.

Acknowledgements. Funding was provided by Spain MINECO (AGL2009-13255-c02-01 and AGL-2012-39880-C02-01), FEDER of the European Union, the Catalonia Government (CIRIT 2009SGR00812), the Swiss Federal Secretariat for Education and Research (SBF C06.0069), the Swiss Federal Office of the Environment, and the Swiss Federal Office of Agriculture (FOAG Fire Blight Control Project). This work was conducted within the European Science Foundation funded research networks COST Action 873 and COST Action 874.

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Competing interests. None declared.

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RESEARCH ARTICLE International Microbiology (2014) 17:91-97 doi:10.2436/20.1501.01.211 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

A phylogenetic approach to the early evolution of autotrophy: the case of the reverse TCA and the reductive acetyl-CoA pathways Arturo Becerra,1* Mario Rivas,1 Carlos García-Ferris,2 Antonio Lazcano,1 Juli Peretó2 School of Sciences, National Autonomous University of Mexico, Mexico DF, Mexico.

Department of Biochemistry and Molecular Biology and Cavanilles Institute of Biodiversity and Evolutionary Biology, University of Valencia, Valencia, Spain 1

2

Received 20 March 2014 · Accepted 4 June 2014 Summary. In recent decades, a number of hypotheses on the autotrophic origin of life have been presented. These proposals invoke the emergence of reaction networks leading from CO or CO2 to the organic molecules required for life. It has also been suggested that the last (universal) common ancestor (LCA or LUCA) of all extant cell lineages was a chemolitho-autotrophic thermophilic anaerobe. The antiquity of some carbon fixation pathways, the phylogenetic basal distribution of some autotrophic organisms, and the catalytic properties of iron-sulfur minerals have been advanced in support of these ideas. Here we critically examine the phylogenetic distribution and evolution of enzymes that are essential for two of the most ancient autotrophic means of metabolism: the reductive tricarboxylic acid (rTCA) cycle and the reductive acetyl-CoA pathway. Phylogenetic analysis of citryl-CoA synthetase and of citryl-CoA lyase, key enzymatic components of the rTCA cycle, and of CO dehydrogenase/acetylCoA synthase, a key enzyme in the reductive acetyl-CoA pathway, revealed that all three enzymes have undergone major lateral transfer events and therefore cannot be used as proof of the LCA’s metabolic abilities nor as evidence of an autotrophic origin of life. [Int Microbiol 2014; 17(2):91-97] Keywords: autotrophic pathways · reverse Krebs cycle · Wood–Ljungdahl pathway · origin of life · last common ancestor (LCA, LUCA)

Introduction Although a heterotrophic origin of life based on the prebiotic synthesis and accumulation of organic compounds is supported by several major lines of evidence [2], other, competing Corresponding author: A. Becerra Facultad de Ciencias Universidad Nacional Autónoma de México Ciudad Universitaria 04510 México DF, México email: abb@ciencias.unam.mx *

alternatives that advocate an autotrophic emergence of living systems have been suggested as well [20,28,30,34]. An autotrophic origin of life, i.e., the hypothesis that the first organisms fed on CO2 as sole carbon source, was proposed in the 19th century. However, in recent decades this proposal has been reassessed, based on biochemical analyses and new geochemical data. Both sources invoke the emergence of reaction networks leading from CO or CO2 to the organic molecules required for life [23]. One of the most well-articulated proposals was that of Wächtershäuser [34], who argued that life began without genetic information and with the appearance of

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a prebiotic autocatalytic reductive tricarboxylic acid (rTCA) cycle (also called the reductive citric acid cycle, reverse Krebs cycle, or Arnon cycle), which is assumed to be originally based on the formation of the highly insoluble mineral pyrite in sulfur-rich hydrothermal environments. The fact that this mode of carbon fixation is found in the most deeply divergent bacteria, i.e., Aquificales, has been used as evidence of its primitive character [35]. The geochemical emergence of a primitive version of the reductive acetyl-CoA pathway (or Wood–Ljungdahl pathway) associated with the FeS-rich mineral boundaries of alkaline hydrothermal vents was proposed

by Russell and Hall [20,28]. Accordingly, it has also been suggested that the last common ancestor (LCA) of all extant cell lineages was a chemolitho-autotrophic thermophilic anaerobe [19,31,36].

Nonetheless, the hypothesis that the first organisms on Earth could fix carbon is far from proven, since the ultimate nature of the first life form is unknown. Phylogenetic analysis using comparative genomics offers clues to the nature of the LCA and could provide evidence of its ability to fix carbon.

Even so, the attributes of the first living entities are unknown and a cladistic approach to the origin of life is not feasible, given that all possible intermediate organisms that may have once existed have long since vanished [2,17]. Moreover, it is not possible to extend an investigation beyond the threshold that corresponds to a period of cellular evolution in which protein biosynthesis was already in operation, i.e., the RNA/ protein world [3].

With these caveats, a few clues about the early evolution of autotrophy have been acquired, by searching in modern genomes. Extant beings are able to fix carbon in at least six different ways, namely: the reductive pentose phosphate cycle (Calvin–Benson cycle), the rTCA cycle, the reductive acetylCoA pathway, the dicarboxylate/4-hydroxybutyrate cycle, the 3-hydroxypropionate/4-hydroxybutyrate cycle, and the 3-hydroxybutyrate bicycle [13,14]. Here we critically examine the distribution and phylogenetic evolution of enzymes that are essential for the purportedly most ancient autotrophic pathways, i.e., the rTCA cycle and the reductive acetyl-CoA pathway [6, 22]. We analyzed sequences from citryl-CoA synthetase and citryl-CoA lyase, two enzymes participating in the rTCA cycle, and CO dehydrogenase/acetyl-CoA synthase, from the reductive acetyl-CoA pathway. Our results show that the early evolution of autotrophy was not free from horizontal gene transfer. Thus, these pathways cannot be invoked as proof of the LCA’s metabolic abilities, nor as evidence of an autotrophic origin of life in volcanic environments rich in transition-metal sulfides.

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Materials and methods Sequences and genomes. The query sequences citryl-CoA synthetase (hth:HTH_1737), citryl-CoA lyase (hth:HTH_0311), acetyl-CoA synthase (mta:Moth_1202), and CO dehydrogenase subunits (mta:Moth_1203) were retrieved, respectively, from the completely sequenced Hydrogenobacter thermophilus and Moorella thermoacetica genomes available in the Kyoto Encyclopedia of Genes and Genomes (KEGG) [16]. Reports on biochemical information were collected from KEGG, BRENDA [29], MetaCyc [7], and PDB [5] databases.

Search for homologous genes. Searches for homologous proteins were carried out by comparing the query sequences to the genomes database in the KEGG, using BLAST searches of its platform [http://www.genome.jp/ tools/blast/]. The BLASTP cutoff value for homologous identification was set at e ≤ 1 ×10–7 and an identity ≥34 %.

Phylogenetic analysis. The amino acids sequences of each enzyme were aligned using MUSCLE 3 [12] software with default parameters.

Neighbor-joining (500 bootstrap replications, maximum composite likelihood distance estimation, and uniform rates among sites) and maximumlikelihood (model Kimura-2P plus gamma distribution with invariant sites, selected according to the Bayesian information criterion, 500 bootstrap replications) phylogenetic trees were constructed using MEGA5 software [33].

The root of the trees was placed using the midpoint method.

Results and Discussion The rTCA cycle can be defined as the (oxidative) Krebs cycle running in reverse (Fig. 1A). While the Krebs cycle is a central pathway in many organisms and is used to oxidize acetylCoA to CO2 and to generate intermediates for biosynthesis, the rTCA cycle allows the inverse process, i.e., the biosynthesis of acetyl-CoA from two molecules of CO2 [14]. Following its discovery in the anaerobic green sulphur photosynthetic bacterium Chlorobium limicola, this autotrophic pathway has been detected in strict anaerobic (and microaerobic) bacteria, such as some members of Aquificales [13]. Many of the enzymes involved in the rTCA and Krebs pathways are the same, with the exception of the key enzymes that allow the cycle to run in reverse, namely: 2-oxoglutarate synthase (2-oxoglutarate:ferredoxin oxidoreductase), fumarate reductase, and the citrate-cleaving enzymes [13,14].

The key step of the rTCA cycle is the ATP-dependent cleavage of citrate into acetyl-CoA and oxaloacetate [14]. In different species, this essential reaction is catalyzed by ATP citrate lyase (ACL; EC 2.3.3.8) in one step or by citryl-CoA synthetase (CCS; EC 6.2.1.18) and citryl-CoA lyase (CCL; EC 4.1.3.34) in two steps. Sequence analysis supports the origin of ACL through the fusion of the gene encoding CCS and CCL. Hence, ACL is a derived sequence and our study focused on

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Fig. 1. Reaction schemes of (A) the rTCA cycle and (B) the Wood–Ljungdahl pathway.

its parental enzymes (CCS/CCL). Furthermore, a common ancestry has been demonstrated between CCS and succinylCoA synthetase (SCS) and between CCL and citrate synthase (CS), both of which participate in the Krebs cycle [1].

A feasible evolutionary history of the reaction was proposed

by Aoshima [1], who suggested that CCS and CCL activities would be the plesiomorphic state of SCS and CS, respectively.

This idea is congruous with the hypothesis of an earliest appearance of a primitive version of the rTCA cycle, as proposed by Wächtershäuser [34]. However, our phylogenetic analysis based on the primary structure of the enzymes does not support Aoshima’s hypothesis [1]; instead, the results indicate a horizontal gene transfer between archaea and hyperthermophilic bacteria. As shown in Fig. 2A, the CCS clade from hyperthermophilic bacteria does not appear as a sister group of the SCS bacterial clade. Similar results were obtained with the phylogenetic tree of CCL and CS (Fig. 2B).

The reductive acetyl-CoA pathway was discovered and described in acetogenic bacteria by the laboratories of Wood, Ljungdahl, Thauer, and others [4,13]. This pathway is a linear

metabolic route in which two molecules of CO2 (or CO2 and CO) are combined directly to form acetyl-CoA (Fig. 1B). The pathway can be divided into two branches: the methyl or “eastern” branch, in which CO2 is sequentially reduced to a cofactor-bound methyl residue, and the carbonyl or “western” branch, in which another molecule of CO2 is reduced to an enzyme-bound carbonyl residue [14,25]. The key enzyme in this pathway is CO dehydrogenase/acetyl-CoA synthase (CODH/ACS; EC 1.2.7.4/2.3.1.169), which has a metallic cluster integrated by iron, sulfur, nickel, zinc, and/or copper [10], although the most active form seems to be Ni-Ni-[4Fe4S] [32]. CODH/ACS is a bifunctional catalyst that reduces CO2 to carbon monoxide, forming the carbonyl group of acetyl-CoA, and catalyzes the synthesis of acetyl-CoA [10,24].

This enzyme is found both in anaerobic archaea and in chemoautotrophic bacteria [4,13,14].

Based on its catalytic activity and metabolic function, the acetyl-CoA synthase of CODH/ACS is a class I enzyme.

Since this class is thought to be older than class II [18], CODH/ACS was the focus of our study. Moreover, it has been

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Fig. 2. Maximum-likelihood tree (bootstrap 500 replicates) from the amino acid sequences of (A) citryl-CoA synthetase and succinyl-CoA synthetase and (B) citryl-CoA lyase and citrate synthase. The colors of the clades describe the function and taxonomy of the OTUs. The root of the trees was placed using the midpoint method.

proposed as one of the oldest enzymes of life, responsible for the ability of early organisms to live in CO2-rich atmospheres [11].

CODH/ACS is a α2β2 heterotetramer that catalyzes two different reactions using seven metalloclusters. The β-subunit is involved in the CODH activity that generates CO from CO2, and the α subunit in the ACS activity that synthesizes acetylCoA [9]. The subunits were found to differ in their phylogenetic distribution (Fig. 3). Also, there were more β-subunits and the topology of the archaeal subunit was more intricate.

These findings can be explained in part by considering CODH as occurring within a large group of bacteria that includes carboxydotrophic bacteria, species of anaerobic acetogenic bacteria, sulfate-reducing bacteria and archaea, phototrophic bacteria, hydrogenogenic bacteria, and methanogenic archaea [8]. Moreover, CODH is present in a larger number of organisms as a monofunctional enzyme than as a bifunctional activity together with the ACS subunit [15]. The topology of the

α-subunit tree is defined by two distinct major clades, one containing the majority of archaeal subunits and another that includes all bacteria. However, a well-supported branch of four methanoarchaea appears as the root of the bacterial clade, suggesting horizontal gene transfer from Archaea to Bacteria (Fig. 3A).

Furthermore, CODH/ACS is an enzymatic complex that requires Ni-S-Fe clusters to transfer electrons, i.e., the 2Ni– [4Fe–4S] cluster (A-cluster) of the α-subunit and the [Ni– 4Fe–5S] cluster (C-cluster) of the β-subunit. Both CODH and ACS are highly specialized enzymes that require specific chaperone systems for their assembly [21,27]. The role of the Fe–S clusters in catalysis is to funnel electrons onto an assembly line that opens and closes in order to catalyze the condensation of CO with a methyl moiety and then “pump out” acetyl-CoA [9]. This complex molecular machinery implies that the activity of CODH/ACS is an evolutionary innovation,

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Fig. 3. Maximum-likelihood tree (bootstrap 500 replicates) from the amino acid sequences of CO dehydrogenase/acetyl-CoA synthase: (A) the α-subunit ACS (EC 2.3.1.169) and (B) the β-subunit of CODH (EC 1.2.7.4; 1.2.99.2). The colors of the clades describe the taxonomy of the OTUs. The root of the trees was placed using the midpoint method.

involving the repurposing of a large number of pre-existing components. It also implies that CODH/ACS was not present at the very early stages of the evolution of life, but was preceded by a long evolutionary history. This possibility is supported by the phylogenetic distribution of the subunits, which suggests that the enzyme appeared after the divergence of the ancestral LCA/ LUCA population.

Our phylogenetic analysis of the citrate-cleaving enzymes participating in the rTCA cycle argue against the hypothesis of Aoshima [1], because these enzymes are not the plesiomorphic version of their homologous counterparts in the oxidative Krebs cycle. Instead, if the key enzymes that allow the Krebs cycle to run in reverse derive from older versions, then it is reasonable to propose that the enzymes required for the oxidative citric acid cycle were present on Earth earlier than those of the rTCA cycle. Phylogenetic analysis of the fundamental citrate-cleaving enzymes, CCS and CCL, do not sup-

port the presence of a complete rTCA cycle in the LCA, since both were acquired by horizontal gene transfer, from archaeal homologs participating in Krebs-cycle-like activities to bacteria. Moreover, the autotrophic growth of the strictly anaerobic Thermoproteales can be explained in full by the dicar­ boxylate/4-hydroxybutyrate cycle rather than by the rTCA cycle, as was initially proposed [4,13,26]. In summary, current evidence strongly suggests that the rTCA cycle is an idiosyncratic pathway of the bacterial domain that evolved after the LCA. Similar conclusions can be derived from the phylogenetic analysis of the subunits of the bifunctional enzyme CODH/ ACS, an essential component of the Wood–Ljungdahl pathway. Despite its wide distribution and diverse activities, there is no evidence that the β-subunit of CODH was present in the LCA. The same holds true for the α-subunit (ACS), which was clearly acquired from methanoarchaea. The bifunctional

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CO dehydrogenase/acetyl-CoA synthase must have emerged after its components and could have not been present during the LCA’s epoch. Since both subunits are part of the Cdh-A-E complex of methanogens, then the CODH/ACS enzymes of the extant reductive acetyl-CoA pathway must have derived from those utilized by methanogens.

It is of course possible that the LCA was an autotrophic organism. However, this hypothesis is not supported by our results on the key enzymes from the oldest CO2 fixation pathways. More studies, including a better phylogenetic representation of the most basal lineages in both prokaryotic domains, are needed to reveal the metabolic abilities of the LCA, and especially to determine whether it was a heterotrophic or an autotrophic organism.

As noted herein and discussed elsewhere [3,17], the origin of life is not amenable to phylogenetic analysis, since molecular cladistics and comparative genomics cannot be extended beyond a threshold that corresponds to a period of cellular evolution in which protein biosynthesis was already in operation. Accordingly, the genome distribution of enzymes from the rTCA cycle and the reductive acetyl-CoA pathway or even the presence of these metabolic routes is not evidence for an autotrophic origin of life.

Acknowledgements. Financial support of CONACYT (100199) to AB and (50520-Q) to AL is gratefully acknowledged. This work was completed during a sabbatical leave of absence of AB, with support from the DGAPAUNAM and the University of Valencia, where he enjoyed the hospitality of Amparo Latorre at the Institut Cavanilles (Valencia, Spain). CGF and JP acknowledge the financial support of MINECO (Grant ref. BFU2012-39816C02-01). We acknowledge the technical support of Sara Islas-Graciano and Ricardo Hernández-Morales.

Competing interests. None declared.

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RESEARCH ARTICLE International Microbiology (2014) 17:99-109 doi:10.2436/20.1501.01.212 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

Succession of the gut microbiota in the cockroach Blattella germanica Purificación Carrasco,1 Ana Elena Pérez-Cobas,2 Claudia van de Pol,1 Joaquín Baixeras,1 Andrés Moya,1,2§ Amparo Latorre1,2§* Institute of Biodiversity and Evolutionary Biology, University of Valencia, Valencia, Spain. 2Genomics and Health Area, Foundation for the Health and Biomedical of the Community of Valencia, Valencia, Spain

1

Received 14 March 2014 · Accepted 10 June 2014

Summary. The cockroach gut harbors a wide variety of microorganisms that, among other functions, collaborate in digestion and act as a barrier against pathogen colonization. Blattabacterium, a primary endosymbiont, lives in the fat body inside bacteriocytes and plays an important role in nitrogen recycling. Little is known about the mode of acquisition of gut bacteria or their ecological succession throughout the insect life cycle. Here we report on the bacterial taxa isolated from different developmental instars of the cockroach Blattella germanica. The bacterial load in the gut increased two orders of magnitude from the first to the second nymphal stage, coinciding with the incorporation of the majority of bacterial taxa, but remained similar thereafter. Pyrosequencing of the hypervariable regions V1–V3 of the 16S rRNA genes showed that the microbial composition differed significantly between adults and nymphs. Specifically, a succession was observed in which Fusobacterium accumulated with aging, while Bacteroides decreased. Blattabacterium was the only symbiont found in the ootheca, which makes the vertical transmission of gut bacteria an unlikely mode of acquisition. Scanning electron microscopy disclosed a rich bacterial biofilm in third instar nymphs, while filamentous structures were found exclusively in adults. [Int Microbiol 2014; 17(2):99-109]

Keywords: Blattella germanica · cockroach gut microbiota · 16S rRNA gene · endosymbionts · ecological succession

Introduction Many insects harbor gut microbial communities that actively interact with their hosts at several different levels [18]. Gut microbes are involved in many processes, such as the digestion of recalcitrant plant polymers, the provision of nutrients, the stimulation of midgut self-renewal, the diet-dependent

Corresponding author: A. Latorre Institut Cavanilles de Biodiversitat i Biologia Evolutiva Universitat de València 46071 València, Spain Tel. +34-963543649. Fax +34-963543480 E-mail: amparo.latorre@uv.es *

Equal contributors

§

duration of developmental stages, resistance to parasite invasion, and host fitness under different environmental regimes [14]. Additionally, some insects have established mutualistic symbiotic associations with intracellular bacteria, which play a metabolic role by providing their hosts with new metabolic pathways that produce nutrients otherwise lacking in their restricted diets [5]. These intracellular symbionts are vertically transmitted from parent to progeny, as revealed by the congruent phylogenies between hosts and endosymbionts [20]. Cockroaches, one of the first insects in which intracellular bodies presumed to be symbionts were recognized [7], harbor the obligate endosymbiont Blattabacterium cuenoti (referred to hereinafter as Blattabacterium) in bacteriocytes, which are specialized cells in the fat body that are required for host fitness and fertility. Genome sequencing revealed that Blattabacterium

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plays a role in host nitrogen metabolism and in the synthesis of essential amino acids [25,36]. In Blatella germanica, Blattabacterium enters the oocyte plasma membrane during ovarian development, prior to chorionogenesis [21]. Once inside, it participates in yolk utilization by vitellophages, thus contributing to embryonic development and providing an added advantage to this symbiotic relationship. Phylogenetic co-cladogenesis between Blattabacterium strains and their corresponding hosts indicates that the initial infection occurred in a common ancestor of cockroaches and termites [31].

However, the endosymbiont was lost in all termite lineages, except in the lower termite Mastotermes darwiniensis [4].

Key changes during the independent evolution of termites include, beside the loss of Blattabacterium, a shift from an omnivorous to a wood diet, the acquisition of specialized hindgut microbiota, and a sophisticated social behavior. The absence of Blattabacterium in nearly all termites suggests that its nutrient-provisioning role was replaced by gut microbes.

In fact, termite gut microbes help fix nitrogen, degrade lig­no­ cellulose, and produce nutrients [48].

In the omnivorous cockroach Periplaneta americana, growth is retarded after the elimination of gut anaerobes by the antimicrobial agent metronidazole, proof of the essential role played by gut microbiota in host physiology [8]. Recent research has focused on the relative contributions of gutresident and intracellular symbionts to host metabolism, both in termites and in cockroaches [45].

Several metagenomic studies have been carried out on the gut bacterial community in cockroaches. Comparison of the phylogenetic relationships of symbiotic bacteria in the xylophagous cockroach Cryptocercus punctulatus with those in lower termites showed a partial coincidence with host phylogeny [6]. However, in the omnivorous cockroach P. americana, a large proportion of sequences proved to be more closely related to environmental sequences than to those of other symbionts represented in current databases [35].

Researchers have also addressed the mode of microbiota acquisition, finding that the gut microbiota of cockroaches and termites is acquired through food or feces. For instance, in C. punctulatus and M. darwiniensis, the intergenerational transfer of hindgut microbiota occurs via proctodeal trophallaxis [27]. Vertical transmission by fecal contamination during oviposition has also been demonstrated, e.g., in stink­bugs of the family Plataspidae, where a specific gut bacterium (Candidatus Ishikawaella capsulata) is vertically transmitted via a symbiont capsule that is laid on the eggs [23]. In B. germanica, descriptions of the transmission mechanism of gut microbiota and the dynamics of ecological

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succession during the developmental stages are lacking. The corresponding scenarios are complex considering that the anterior and posterior cuticles of the intestinal tract (foregut and hindgut) are renewed during each of the 5–6 moultings (males and females, respectively). How they are recolonized after moulting is unknown, although it has been speculated that the mechanism involves either a reservoir of microbiota in the gut or incoming bacteria from the environment.

In this work, we examined the bacterial load and changes in microbial diversity both in the ootheca (embryos) and in the gut of B. germanica in each of the five nymphal instars, as well as in adult males. Our results shed light on the mode of gut microbiota acquisition throughout cockroach development.

Materials and methods Insect rearing. The experimental B. germanica colony originated from a stable laboratory population (started 30 years ago) housed by Xavier Bellés at the “Institut de Biologia Evolutiva”, Barcelona, Spain. Culture chambers were adjusted to 26 ± 1 ºC, 70 % humidity, and a photoperiod of 12D:12L. The insects were bred in lunchboxes with aeration and fed on autoclaved dog food composed of cereals, meat and animal by-products (25 % meat), vegetable origin by-products (2 % beet pulp), oils and fats (poultry fat, source of ω3), minerals, and yeasts. The additives (per kg) were: 11,000 IU vitamin A, 825 IU vitamin D3, 66 mg vitamin E, 55 mg Fe, 1.4 mg iodine, 0.3 mg cobalt, 6 mg copper, 28 mg manganese, 45 mg zinc, and 0.1 mg selenium.

Antioxidants, preservatives, and dyes were also present. In summary, the analytical components were: 24 % protein, 10 % fat, 2.5 %; gross fiber, 8.5 % inorganic matter, and 10 % moisture. Water was supplied ad libitum. The lunchboxes were renewed weekly. A cohort of 40 individuals maximum was maintained per box.

Nymphal instar and adult stage morphological deter­ mination. The nymphal instar (n) was identified by measuring head width, as previously described [42], whereas adults were easily identified by their wings. Accordingly, the specimens were classified as n1, n2, n3, n4, n5 and adults (male), corresponding to individuals 2, 11, 15, 22, 34 and 68 days after hatching, respectively.

Gut dissection and ootheca sampling. Two specimens from the same brood were selected as biological replicates for each of the developmental stages. The insects were anesthetized under a stream of CO2 and placed dorsally on a paraffin plate. After removal of its head, the cockroach was pinned with minute entomological pins through the prothorax and last abdominal segments. The legs were coxally cut. Dissections were carried out under a stereomicroscope using fine forceps (Wild M8, Lawton, GmbH & Co., KG, Fridingen, Germany) and spring scissors. In the tiny young nymphs, the gut was simply stretched through the anus. In the remaining cases, the body cavity was exposed through an incision in the tergal area and the gut removed from the anus to the level of the metathorax. Residual fat body tissue was removed, placed in a tube, and frozen in liquid nitrogen for further studies. The samples were stored at –80 ºC. Ootheca contents were extracted by drilling the external surface with a pipette tip and sucking out the embryo-containing fluid. Surface samples were routinely tested for bacteria

by PCR, with negative results (data not shown).

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The guts and ootheca contents were ground manually with a glass rod in cell lysis buffer (JETFLEX genomic DNA purification kit, Genomed, Löhne, Germany) and digested with proteinase K overnight.

Bacterial load determination. Absolute quantification of bacterial 16S rRNA gene copies was carried out by quantitative PCR (qPCR) using the universal bacterial 16S rRNA gene primers 8F 5′-AGAGTTTGATCCTGG CTCAG-3′ and 338R 5′-TGCTGCCTCCCGTAG GAGT-3′ [46]. The HOT FIREPol EvaGreen (Solis Biodyne, Tartu, Estonia) qPCR Mix Plus kit was used together with Roche Light Cycler 2.1 thermocycler. The thermal profile was: 95 ºC for 15 min followed by 40 cycles of 95 ºC for 10 s, 55 ºC for 10 s, and 72 ºC for 18 s. Standard curves were constructed with purified and photometrically quantified amplicons (103–107 copies, or molecules), to interpolate sample crossing points (Cp). The standards were aliquoted and stored at –80 ºC. The equation of the standard curve was lg (copy number) = –3.58Cp + 36.016, R2 = 0.999. Four measurements per stage were used to statistically test the change in bacterial load from one stage to the next. The Wilcoxon signed-rank test, implemented in R software [51], was applied to compare the sequential stages; comparisons with a p-value less than 0.05 were considered significant.

Bacterial 16S rRNA gene amplification and pyrose­ quencing. The V1–V3 variable regions of the bacterial 16S rRNA genes were PCR-amplified using the universal primers E8F (5′-TAG AGT TTGATCMTGGCTCAG-3′) linked to the adaptor CCATCTCATCCCTGC GTGTCTCCGACTCAG and 530R (5′-TTGCTGCCTCCCGTAGGAGT-3′) and the sample-specific multiplex identifier (MID) for pyro­sequencing. PCR was carried out in a total volume of 50 μl. Each reaction contained the TAKARA reagent (0.25 μl of 5 U/μl Ex Taq HS, 1 μl of 2.5 mM dNTP mixture, 5 μl of 10× Ex Taq buffer), 0.2 μl each of the forward and reverse primers (stock 10 μM), and 50 ng of template DNA. The GeneAmp PCR system 9700 thermocycler reaction conditions were: initial denaturation at 94 ºC for 5 min; 25 cycles of denaturation at 94 ºC for 30 s, annealing at 55 ºC for 30 s, and elongation at 72 ºC for 1 min; and a final elongation at 72 ºC for 10 min. To avoid PCR bias, we chose the lowest DNA template quantity and the fewest possible PCR amplification cycles [1,49]. The integrity and quantity of the amplicons were checked by agarose gel (1.4 %) electrophoresis. To precipitate and purify the DNA, 2 μl NaOAc and 40 μl cold 95 % ethanol were added, and the DNA was resuspended in 10 μl water.

The Qubit dsDNA BR assay kit was used with the Qubit 2.0 fluorometer to accurately quantify the purified amplicons, which were pooled in equimolar amounts for pyrosequencing. Samples were sequenced in a next-generation 454 pyrosequencer (Genome Sequencer FLX system, Roche) available at the FISABIO-Salud Pública, Valencia.

Processing and taxonomic assignment of 16S rRNA reads.

An initial quality trimming and MID sorting process was performed at the Ribosomal Database Project website [15] in the pyrosequencing pipeline.

Primers, MIDs, and sequences with a Phred quality score less than 20 (Q20) and short length (<250 pb) were removed. Taxonomical annotation was done by aligning the sequences against a subset from the SILVA SSU NR 111 database (ssu_clsref_111_rc1), comprising 200,000 sequences. To do so, we used SINA software [33], first fixing the annotation of the bacterial taxa at the genus level and then classifying the non-annotated sequences at the closest possible level (family, order, class or phylum), with the prefix “uc” indicating that classification at the lowest level (genus) was not possible.

Biodiversity and statistical analysis. The composition and structure of the gut microbiota in each sample were characterized using 16S

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rRNA gene amplicons. Structure was analyzed by estimating two diversity variables, i.e., the expected number of taxa and the Shannon diversity index [39], and two richness estimators i.e., the Chao1 [12] and the abundancebased coverage estimator (ACE) [13]. All diversity variables were calculated after re-sampling (with the same number of sequences per sample, to avoid sequencing effort differences) using the multiple_rarefactions.py and alpha_ diversity.py scripts of the QIIME program [11]. The Wilcoxon signed-rank test was used as above to statistically compare the mean ranks of the Shanon index and the Chao1 and ACE richness estimators between nymphs and adults. Rarefaction curves were generated to estimate the number of expected taxa at the different stages [22], using the Vegan Community Ecology Package [30].

A canonical correspondence analysis (CCA) showed sample variation in terms of taxon abundance and its relationship to host stage. A multivariate ANOVA based on dissimilarity tests (Adonis) was used to test the effect of the variable “developmental stage” in explaining the observed variation in the data. These analyses were also run with Vegan software.

A regression analysis was performed to identify those taxa showing a statistically significant trend over time, considering the sampling time (days).

In addition, statistically significant differences between the relative abundances of taxa in n3 nymphal instars vs. adult specimens were determined. These two analyses were performed with the ShotgunFunctionalizeR R package [24]. The function test GeneFamilies.regression was used for the regression analysis, while the test GeneFamilies.dircomp was used for multi-sample comparison.

Both tests are based on the Poisson model.

Clustering analysis. A cluster analysis was used to study inter-sample similarity in taxon composition. The pvclust R software [41] was used to calculate the uncertainty of the hierarchical clusters using bootstrap resam­ pling techniques. The approximate unbiased (AU) p-value with 10,000 replicates was chosen to calculate the probability of each cluster.

Scanning electron microscopy (SEM). The guts of third instar nymphs and adults were inspected by SEM. The study required the use of fixed fresh material from between three and ten specimens of B. germanica.

Before dissection, the cockroaches were starved for three days to obtain a clear gut surface. The specimens were anesthetized by CO2 in a killing jar.

A drop of fixative (paraformaldehyde 2 %/glutaraldehyde 2.5 %) was gently injected through the thorax into the body cavity using a hypodermal syringe (30G needle). The whole specimen was then immersed in the same fixative.

General dissection procedures were modified from standard methods for anatomical preparation for optical microscopy. Dissection and cleaning were carried out under a MZ9.5 Leica stereomicroscope, with the insects placed in 30-mm glass embryo dishes and by using spring micro-scissors, Dumont (Fine Science Tools, GmbH, Heidelberg, Gernany) forceps (number 5), and fine-model brushes (5/0) . After fixation, the abdomen was dissected in insect Ringer solution. Fragments of the foregut, midgut, and hindgut were cleaned and longitudinally opened, then fixed again in the same fixative. Fragments of digestive content and nematodes were removed. Each gut piece was placed inside a microporous specimen capsule (30-μm pore size, Ted Pella, Redding, CA, USA) immersed in absolute ethanol, and then subjected to critical point drying in an Autosamdri 814 critical point dryer (Tousimis, Rockville, MD, USA). The fragments obtained were arranged on SEM stubs using the silverconducting paint (TAAB, Berks, England) and examined under a Hitachi S-4100 scanning electron microscope. Images were edited with Photoshop CS3.

Sequence data deposition. All sequences obtained in this study were submitted to the European Bioinformatics Institute (EBI), EMBL Nucleotide Sequence Submissions Database ID: ERP002663.

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Results Time-course changes in the bacterial load. To estimate the bacterial load per gut we determined the bacterial 16S rRNA gene copy number by qPCR, using total DNA extracted from B. germanica instars n1â&#x20AC;&#x201C;n5 and male adults (2, 11, 15, 22, 34, and 68 days after hatching). As shown in Fig. 1, bacterial load increased from stage n1 to stage n2 but then remained relatively constant through the following moultings, until the adult stage. However, despite some variations between stages, only the change from n1 to n2 was significant (Wilcoxon signed-rank test, p = 0.03).

16S rRNA gene pyrosequencing. From ootheca and the guts of B. germanica, 52,562 sequences belonging to a 700-bp PCR amplicon of bacterial 16S rRNA gene were obtained. As shown in Fig. 2, seven phyla were detected at the highest taxonomic level: Bacteroidetes, Deferribacteres, Firmicutes, Fusobacteria, Planctomycetes, Proteobacteria, and Synergistetes. At the family level, there were 22 different taxa, while at the lowest taxonomic level discriminated by this technique (genus), 18 bacterial genera were identified, 16 of which were Gram-negative and 13 anaerobic. Note that, in the ootheca (embryos), only Blattabacterium and unclassified Blattabacteriaceae, most likely Blattabacterium as well, were detected. In the remaining stages, except n1, Blattabacterium was present in all samples only at low abundance. Dissection of the gut without contamination by traceable amounts of

Fig. 1. Time-course changes in gut bacterial load throughout the development of Blattella germanica assessed by qPCR. Bacterial abundance was inferred from the copy number of bacterial 16S RNA genes in total DNA extracted from cockroach guts. Four determinations were carried out at each stage.

fat body is not technically feasible. Thus, the recovery of Blattabacterium sequences in the extracts from gut tissue was almost certainly an artefact of the method. In n1 nymphs, removal of the fat body from the gut tissue is practically impossible, such that the amount of Blattabacterium recovered may be extraordinarily high (Fig. 2). The dramatic differences observed in ootheca and n1 instar samples with respect to Blattabacterium as well as the poor representation of the remaining taxa recommend the exclusion of these stages in further analyses.

Microbial diversity. Figure 3 shows the rarefaction curves for each stage of cockroach development, including the ootheca. While both the ootheca and the n1 samples were expected to have the lowest number of taxa, the number in instar n1 was higher than that in the ootheca samples.

This indicated that the first colonizers of the gut ecosystem appeared at this stage, even though the relative abundance of Blattabacterium was still high. The remaining nymphs and adults had similar numbers of expected taxa (>40). The rarefaction curves also showed that the most abundant taxa in the cockroach intestinal ecosystem were sequenced, although a higher number of reads would have been required to reach the plateau in all samples.

Estimates of bacterial diversity (expected number of taxa and based on the Shannon index) and richness (Chao1 and ACE) in the gut of nymphal (instars n2â&#x20AC;&#x201C;n5) and adult stages (Table 1) after data re-sampling showed similar numbers of expected taxa and a similar Shannon index for all nymphs

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Fig. 2. Bar plot of the relative abundance in bacterial taxa at each of the developmental stages of Blattella germanica, including Blattabacterium reads. The y-axis represents the proportion of pyrosequencing reads belonging to each taxon (genus and unclassified levels “uc”). Taxa with an abundance <1 % are included in “others.”

and adults (range: 37.04–49.74 and 3.59–4.33, respectively).

Thus, despite compositional differences between samples, they had the same degree of homogeneity. The richness values were also similar between nymphs and adults. According to Wilcoxon signed-rank tests comparing the indexes between nymphs (n2, n3, n4, n5) and adults, the differences were not significant (p = 0.71; 1.0; 0.71 and 0.89 for expected N, Shannon, Chao1, and ACE, respectively).

Variations in bacterial composition in Blatella germanica. A cluster analysis testing sample similarity according to taxa abundance distribution (Fig. 4) yielded two defined clusters, grouping nymphal samples in one and adult samples in the other. In addition, only adults and n3 samples clustered together. To evaluate sample pattern variation in taxon abundance and host developmental stage, an ANOVA was run to define the effect of “stage”, formed by stages

n2–n5 and the adult stage. The difference between the two proved significant (p = 0.02). A CCA then provided further information on differences in taxon abundance according to developmental stage. As shown in Fig. 5, the first axis explained 20 % of the overall variability, separating the n3 nymphal stage and the adults from all the other stages. The second axis explained 19.5 % of the variability and separated adult samples from nymphs. Both statistical approaches indicated that part of the variation in gut bacterial composition in B. germanica is stage-related, and that n3 nymphs and adults differ from the other stages.

To determine whether, during development, the number of taxa significantly accumulated or decreased, we carried out a regression analysis based on a Poisson model. Time served as the independent variable, with samples taken 11, 15, 22, 34, and 68 days after hatching, corresponding to n2–n5 (nymphal stages and adult). The results showed a total of 23 significant

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taxaâ&#x20AC;&#x201D;14 increasing and nine decreasing in abundance between days 11 and 68 and thus depict the microbial ecological succession taking place inside the cockroach gut. Figure 6 shows the dynamics of these significant taxa, excluding those with a level of relative abundance below 0.50 % at all stages. The largest increases over time were in Fusobacterium, which was most predominant in adults, and the unclassified Deltaproteobacteria, Christensenellaceae, Erysipelotrichaceae and Desulfobacteraceae (Fig. 6A). Taxa that decreased over time were Bacteroides, Parabacteroides, Thalassospira, and the unclassified Lachnospiraceae (Fig. 6B).

As early as stage n2, 86 % of the total taxa were already present, concomitant with the burst in bacterial abundance per gut (see Fig. 1). Since the n3 nymphal stage and the adult stages

Fig. 3. Rarefaction curves of the sequencing reads for each developmental sample showing the maximum expected number of taxa.

had the highest within-stage homogeneity (Fig. 4) and the highest heterogeneity between developmental stages (Fig. 5), and given their behavior and physiology (see Discussion), we carried out a comparative analysis to identify which taxa are particularly enriched at each of these stages. At the adult stage, Fusobacterium, uc_Bacteroidetes, uc_Porphyromonadaceae, Dysgonomonas, uc_Deltaproteobacteria, Pseudomonas, and Mucispirillum, were the most abundant, whereas at the nymphal stage, Bacteroides, Enterococcus, uc_Lachnospiraceae, Escherichia-Shigella, Para­ bacteroides, and Odoribacter were more abundant.

Scanning electron microscopy. For the same reasons, we chose a nymphal stage (n3) and the adult stage to visualize the gut microbiota by SEM (Fig. 7). Microbial

Table1. Number of expected taxa (N) and Shannon Chao 1 and ACE indices calculated from the re-sampled data set Sample

N (SD)

Shannon (SD)

Chao1 (SD)

ACE (SD)

n2A

45.00 (1.57)

4.21 (0.02)

55.33 (8.11)

59.72 (7.15)

n2B

44.61 (2.65)

4.10 (0.03)

57.61 (13.25)

58.05 (8.76)

n3A

39.45 (1.78)

3.86 (0.03)

49.76 (10.30)

49.00 (5.79)

n3B

39.33 (1.80)

3.59 (0.03)

45.91 (6.45)

47.02 (5.04)

n4A

44.87 (1.26)

4.29 (0.02)

49.51 (4.99)

51.69 (4.08)

n4B

43.25 (1.59)

4.15 (0.02)

50.14 (7.37)

48.66 (3.98)

n5A

37.04 (1.83)

3.95 (0.03)

48.91 (11.88)

43.66 (4.91)

n5B

49.74 (2.16)

4.33 (0.03)

60.01 (10.65)

57.38 (5.50)

Adult A

42.98 (0.15)

4.12 (0.00)

46.52 (0.51)

47.85 (0.41)

Adult B

44.88 (1.59)

4.13 (0.03)

50.93 (6.55)

50.05 (3.55)

SD, standard deviation.

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colonization was not detected on the foregut or midgut surface, as previously reported for P. americana [9]. However, a dense bacterial biofilm formed by rosettes of rods was observed on the hindgut cuticle in both stages. Filamentous structures were present exclusively in adult cockroaches.

Discussion Cockroaches harbor a rich microbiota in the gut, but how it is acquired after hatching and how it develops are unclear. Our

Fig. 5. Canonical correspondence analysis (CCA) of Blattella germanica gut microbiota. The developmental stages are represented as five vectors (n2, n3, n4, n5, and adult). The axes represent the percentage of the corresponding total variance explained. The closeness of the points is an indicator of similarity.

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Fig. 4. Clustering analysis based on taxon composition. Bootstrap values appear above each cluster and the order of the clustering (edge #) below.

Height indicates the average distance between the corresponding clusters.

qPCR results showed that in B. germanica the bacterial load per gut increased by about two orders of magnitude between hatching and the n2 instar stage, when 86 % of the total detected bacterial taxa appeared. In the other stages, the bacterial load remained more or less constant, with no significant differences between them. This finding is in accordance with those reported for other insects, such as Zonocerus variegatus, whose culturable bacterial load increases 80-fold between the first nymphal instar and the adult [2]. Similarly, the bacterial load of Frankliniella occidentalis increases by four orders of magnitude between the first instar and adult

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Fig. 6. Temporal trend of taxa that show a statistically significant change during the instar and adult developmental stages. (A) Increasing over time; (B) decreasing over time.

stage [47]. Nevertheless, the evenness of the gut bacterial yield between n3 and adult stages is intriguing, considering that the gut cuticle of the hindgut is renewed during moulting and given the changes in the microbial composition detected in this work. A microbial reservoir in the midgut (not shed during successive moults) could spread to newly formed tissues, keeping the bacterial population size constant. This is the case in the wax moth Galleria mellonella, in which a small microbial reservoir persists in the gut [10].

Bacterial composition underwent important temporal changes during development, with some taxa showing significant changes in abundance. Blattabacterium was the only bacteria detected in embryos inside the ootheca, consistent with a lack of vertical transmission of the gut microbiota; instead, horizontal transmission of behaviors such as trophallaxis, coprophagy or body/ootheca licking seems probable. Further experiments with antimicrobial agents and analysis of the gut microbiota of offspring are underway in our laboratory to ascertain whether bacterial taxa eliminated by antibiotics can be restored via environmental sources.

Overall, the gut content among most nymphs is similar but it differs from that of adults, as shown by CCA. This similarity suggests that at these stages the gut habitat is suitable for a specific fraction of early-colonizing microorganisms. Habitats

with similar conditions tend to select for similar microbial communities [16]. When the source (reservoir) diversity is low, similar communities are observed [17], as in the case of the nymphal guts after each moulting. Additionally, some species depend on the presence of other microbial community members [18]. However, given that some of the replicates did not cluster together, it may be that the establishment of the microbiota after each moulting event is influenced by stochastic factors, with the final composition determined by the order of the microorganismsâ&#x20AC;&#x2122; arrival. Similar results were obtained with P. americana [37]. In adults, the gut microbiota is no longer disturbed by moulting, which enables ecological succession to continue until an ecosystem equilibrium is reached. This scenario probably explains the differences in microbial composition between nymphs and adults.

Cluster analysis also revealed that samples belonging to the adult and n3 nymphal instar grouped together, although their bacterial compositions were very different. Strikingly, the bacterial assembly of n3 samples differed not only from those of adults but also from those of all other nymphal stages.

These differences could be related to the physiological state of the host at that particular developmental stage, corresponding to the social/behavioral transitions of the third instar stage [28]. Stage-specific traits in B. germanica, e.g., the specific

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Fig. 7. SEM images of the luminal surface of the hind gut of Blattella germanica. (A,B) adult; (C,D) n3 instar nymph. Scale bars: 50 µm (A,C) and 5 µm (B,D).

activity of acetylcholinesterase, whose levels have been shown to decrease in the n3 instar, have been described [34].

The most abundant phyla in all stages, except embryos and stage n1, were Bacteroidetes (60 %), Firmicutes (30 %), and Proteobacteria (10 %). These phyla are also predominant in the gut of the cockroaches S. lateralis [38] and C. punctulatus [6].

Among the 18 genera identified (abundance >1 %), the presence of five genera of aerobic bacteria, Massilia, Porphyrobacter, Pseudomonas, Sphingomonas, and Thalassospira, was note­ worthy. Among the taxa identified at the family level, Lachnospiraceae produces acetate, propionate, and butyrate, three potential host energy sources. This bacterial family is also present in other insects, such as termites and locusts, and in omnivorous animals, including mammals [44]. The family Erysipelotrichaceae comprises four different genera that in a study in mice responded differently to diet and host health [50]. Other cases of a host-related relationship between bacterial family and host have been reported, such as the overrepresentation of Peptostreptococcaceae in children living with pets [3]. Moreover, Ruminococcaceae, together with Lachnospiraceae, are the predominant autochthonous families in both human and mouse colon [29].

Changes in relative abundances were observed for Fuso­ bac­terium and Bacteroides, which, respectively, accu­mulated and decreased during the insect life cycle (Fig. 6). Both are

anaerobic Gram-negative bacteria. Fusobacterium varium, for instance, is an integral constituent of the human gut microbiota, and, unlike many gut microorganisms, is capable of fermenting both amino acids and glucose [32]; by contrast, simple sugars are not the main energy source of Bacteroides.

In fact, a large part of the Bacteroides proteome includes proteins that hydrolyze polysaccharides. The ability to convert complex polysaccharides into simpler (usable) compounds might allow Bacteroides to be more competitive at early stages in cockroach development. We propose that, as Bacteroides initially predominates and degrades complex nutrients, the monosacharides and amino acids released would allow Fusobacterium to proliferate and progressively accumulate.

SEM images showed a higher abundance of segmented filaments in the adult than in the n3 instar, which suggests an increase in microbial community complexity over time.

Some groups would take longer to colonize the ecosystem, ruling out their presence in nymphs because of the successive renewals of the cockroach fore and hindgut cuticle. The name Arthromitus has been applied collectively to conspicuous filamentous bacteria found in the hindguts of termites and other arthropods. A recent and meticulous study in the termite Reticulitermes definitively clarified the monophyletic origin of Arthromitus within the family Lachnospiraceae [43].

Segmented filamentous bacteria (SFB) from vertebrates

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form a distinct lineage within the family Clostridiaceae.

Arthromitus lives in the hindgut of termites and cockroaches such as Blaberus giganteus [19], C. punctulatus, Blatta orien­ talis, S. lateralis [43], and Gromphodorhina portentosa [26].

P. americana reportedly harbors gut microbiota with a similar morphology [9].

Among the genera detected at the n5 stage that persist in the adult were Anaerofustis, Cetobacterium, Enterobacter, and Hydrogenoanaerobacterium. The latter consists of thin rods that are 14.5-µm long and usually form pairs [40] and is thus a good bacterial candidate for some of the structures observed in the SEM images, although these could also be attributed to filamentous fungi. It is known that fungi colonize the B. germanica gut [37] and some of the morphologies observed are compatible with those of Trichomycetes, which are obligate fungal dwellers in the guts of insects, crustaceans, and millipedes [26]. Methods such as FISH or metagenome analysis are necessary to elucidate the sources of these different structures in adults.

In summary, bacterial community composition in the B. ger­ manica gut differs between nymphal instars and adults.

Specifically, the former has an additional abundant genus (Bacteroides) that decreases in abundance when the insect moults as it becomes an adult, after which there is a progressive accumulation of the genus Fusobacterium. The main increase both in bacterial quantity and diversity takes place after the first moult, after which bacterial load and richness remain constant. According to our results, it is more likely that the gut microbiota is horizontally transferred via fecal contents, rather than vertically via transfer to the egg.

Acknowledgements. This work was supported by projects BFU201239816-CO2-01, co-financed by FEDER funds, and Prometeo/2009/092 from the Ministerio de Economía y Competividad, Spain, and the Generalitat Valenciana, Spain, to AL and AM, respectively. We thank M.D. Piulachs, M. Porcar, J. Pedrola and R. Patiño-Navarrete for their experimental advice.

SEM observation was made at the Electron Microscopy Services of the University of Valencia.

Competing interests. None declared

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Bioinformatics 28:1823-1829 34. Qian K, Wei X, Zeng X, Liu T, Gao X (2010) Stage-dependent tolerance of the German cockroach, Blattella germanica for dichlorvos and propoxur. J Insect Sci Online 10:201 35. Sabree ZL, Huang CY, Arakawa G, Tokuda G, Lo N, Watanabe H, Moran NA (2012) Genome shrinkage and loss of nutrient-providing potential in the obligate symbiont of the primitive termite Mastotermes darwiniensis.

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RESEARCH ARTICLE International Microbiology (2014) 17:111-117 doi:10.2436/20.1501.01.213 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

A multiplex PCR for the simultaneous detection of Tenacibaculum maritimum and Edwardsiella tarda in aquaculture Nuria Castro,* Alicia E. Toranzo, Beatriz Magariños Department of Microbiology and Parasitology, Faculty of Biology/CIBUS, University of Santiago de Compostela, Santiago de Compostela, Spain Received 12 February 2014 · Accepted 6 June 2014

Summary. A specific and sensitive multiplex PCR (mPCR) method was developed as a useful tool for the simultaneous detection of two important flatfish pathogens in marine aquaculture, Tenacibaculum maritimum and Edwardsiella tarda. In fish tissues, the average detection limit for these mPCR-amplified organisms was 2 × 105 ± 0.2 CFU/g and 4 × 105 ± 0.3 CFU/g, respectively. These values are similar or even lower than those previously obtained using the corresponding single PCR. Moreover, mPCR did not produce any nonspecific amplification products when tested against 36 taxonomically related and unrelated strains belonging to 33 different bacterial species. Large amounts of DNA from one of the target bacterial species in the presence of low amounts from the other did not have a significant effect on the amplification sensitivity of the latter. [Int Microbiol 2014; 17(2):111-117] Keywords: Tenacibaculum maritimum · Edwardsiella tarda · multiplex PCR · fish pathology · aquaculture

Introduction Fish diseases, especially those caused by Gram-negative bac­teria, are a serious problem in aquaculture. At present, tena­cibaculosis [2,6] and edwardsiellosis [5,8], caused by Tenacibaculum maritimum and Edwardsiella tarda, respect­ ively, are two important bacterial diseases affecting a wide range of cultured fish species, including flatfish [2,8].

Tenacibaculum maritimum [16] is the causative agent of gliding bacterial disease (or tenacibaculosis) and it infects Corresponding author: N. Castro Departamento de Microbiología y Parasitología Facultad de Biología/CIBUS Universidad de Santiago de Compostela 15782 Santiago de Compostela, Spain Tel. +34-981563100 E-mail: nuria.castro@usc.es *

a wide variety of valuable marine fish species, such as turbot (Scophthalmus maximus), sole (Solea solea, Solea senegalensis), gilthead seabream (Sparus aurata), and salmon (Salmo salar) [2]. The traditional culture-based method for the detection of this pathogen requires several days to weeks before results are obtained. In addition, one of the problems in the study of T. maritimum is the difficulty of distinguishing it from other phenotypically similar and phylogenetically related species, particularly those of the genera Flavobacterium and Cytophaga [4,16]. Therefore, in 2004 Avendaño-Herrera et al. [2] evaluated the specificity and sensitivity of two PCR methods previously described by Toyama et al. [19] and Bader and Shotts [3] in the identification of T. maritimum strains. They found that the former method allowed the accurate detection of T. maritimum in diagnostic pathology as well as in epidemiological studies of gliding bacterial disease of diseased and carrier marine fish.

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In the last decade, E. tarda has become an important bac­ terial pathogen in aquaculture. In addition, the bacterium is associated with septicemia and fatal infections in other animals, such as reptiles, birds, amphibians, marine mammals, and humans, and is thus a possible source of zoonoses [8].

Several attempts to develop methods for the rapid and accurate diagnosis of edwardsiellosis have been made, including PCRbased methods. In 2010, we published an evaluation of the specificity and sensitivity of four PCR primer pairs previously described for the detection of E. tarda [7]. Of these, a PCR protocol employing the gene etfD (which encodes the upstream region of the fimbrial gene) [15] was shown to be the most rapid and sensitive method for the accurate detection of E. tarda in infected fish.

Although simultaneous detection of several pathogens with a multiplex PCR (mPCR) has been widely applied to the detection of multiple viruses and bacteria in clinical specimens, this approach has not been widely used in the detection of fish pathogens [1,12,13,17]. In this work, we developed a mPCR for the rapid and economical simultaneous detection of T. maritimum and E. tarda, and in aquaculture.

Materials and methods Bacterial strains and growth conditions. Forty-seven strains were used to evaluate the mPCR method, including six E. tarda strains and five T. maritimum strains isolated from different hosts and origins and 36 isolates of other taxonomically related and unrelated species (belonging to 33 different bacterial species) (Table 1). Reference strains of E. tarda and T. maritimum were included as positive controls. The identity of each isolate was confirmed employing biochemical tests [18] and, in some cases, using PCR-based analysis and/or serological assays.

For all experiments, the strains were routinely grown on either tryptone soy agar supplemented with 1 % (w/v) sodium chloride (TSA-1, Pronadisa, Spain), marine agar (MA; Difco, USA), or Flexibacter maritimum medium (FMM) [14] as appropriate for each strain. All strains were incubated at 25 °C for 24–72 h. Stock cultures were stored at –70 °C in Cryo-Bille tubes (AES Laboratory, France).

DNA extraction from bacterial cultures. Chromosomal DNA was extracted from pure bacterial using Insta-Gene Matrix (Bio-Rad, Spain), following the manufacturer’s recommendations. The DNA was resuspended in a final volume of 200 μl of Insta-Gene Matrix. The concentration was determined spectrophotometrically at 260 nm and adjusted with sterile distilled water to a concentration of 10 ± 3 ng/μl. The DNA was stored at –30 °C until used for PCRs. All experiments were carried out with DNA obtained in three different extractions of each bacterial strain.

DNA amplification. All PCR amplifications were performed using Ready-To-Go PCR beads (GE Healthcare) according to the manufacturer’s instructions. The species-specific primer pairs described by Toyama et al.

[19] and Sakai et al. [15] were used for the mPCR and were synthesized by Sigma-Genosys.

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One μl of each DNA solution and 1 μl of each primer (100 μM for E. tarda and 2 μM for T. maritimum) were used in the amplification reactions.

Reaction mixtures (25 μl) were amplified in two different thermal cyclers: the T Gradient Termocicler (Biometra) and the Mastercycler Personal (Eppendorf). The PCR annealing temperatures tested ranged from 45 to 55 °C. Both the intensity of the amplicons for each targeted DNA and the absence of nonspecific bands were considered in the selection of optimal mPCR conditions. The cycling protocol was one cycle of 94 °C for 2 min, 35 cycles of 95 °C for 2 min, 45 °C for 1 min 30 s, and 72 °C for 2 min, and a final elongation at 72 °C for 7 min. Negative controls, consisting of the same reaction mixture but with sterile distilled water instead of template DNA, were included in each batch of PCRs. The reproducibility of the results was assessed by repetition of the amplifications in three independent PCR assays. As a positive control, the universal primers pA (5′-AGA GTT TGA TCC TGG CTC AG-3′) and pH (5′-AAG GAG GTG ATC CAG CCG CA-3′ [9] were used to detect 16S rDNA in all strains.

Analysis of PCR products. Ten μl of the PCR products were separated on a 1 % (w/v) agarose gel for 60 min at 100 V in 1× TAE (0.04 M Tris, 1 mM EDTA, pH 8.0) and visualized by staining the gels with 0.06 μg ethidium bromide (Bio-Rad)/ ml. The bands were photographed under UV light and computer digitized (Gel Doc 100, Bio-Rad). A 50- to 1500-bp ladder (Fast Ruler low range DNA ladder, Fermentas, Spain) served as a molecular mass marker. The presence of a single product of the appropriate size and identical to that from the respective reference strains was considered as a positive result.

Specificity and sensitivity from bacterial cultures. The specificity of the mPCR was evaluated using genomic DNA extracted from strains belonging to different bacterial genera and species commonly found in flatfish infections (Table 1). To determine the analytic sensitivity of the mPCR, separate pure bacterial suspensions of the T. maritimum NCIMB2154 and E. tarda ACC35.1 strains were prepared to contain 109 cells/ml (McFarland scale 4) and were then ten-fold diluted in 0.85 % NaCl sterile saline to yield a dilution series containing 108 to 101 cells/ml. One hundred µl of each dilution was cultured on TSA-1 for E. tarda and on FMM for T. maritimum; the plates were incubated at 25 ºC. After the incubation, the colonies were counted and the bacterial concentrations of the stock cultures were calculated as CFU/ml.

Known bacterial concentrations of different dilutions of both fish pathogens were mixed and the DNA was extracted using Insta-Gene Matrix, as previously described. The detection limits were determined by the presence or absence of the specific PCR products on agarose gels.

Applicability to fish tissues. The sensitivity of the mPCR in fish tissues was determined using DNA extracted from in-vitro-spiked spleen, kidney, and liver obtained from healthy sole and turbot (weight: 10–12 g).

Both fish specimens were previously analyzed by bacteriological standard methods [18] to confirm the absence of pathogens that could interfere in the experiments, as described by Castro et al. [7] Each tissue sample (mean weight: 0.1–0.2 g) was homogenized in 100 μl of PBS by repeated pipetting. Each fish sample was seeded with 100 μl of the above-described bacterial dilutions and homogenized. After incubation of the samples at 25 °C for 1 h, genomic DNA was extracted with the Easy-DNA kit (Invitrogen) by following the manufacturer’s recommendations. As negative controls, DNA from fish samples “seeded” with PBS were extracted in the same manner. For mPCR, 1 μl of the purified DNA was added as the template.

Detection limits were determined based on the presence or absence of PCR products in agarose gels.

In addition, to test the effect on the mPCR of a large amount of DNA from one pathogen in the presence of a small amount of DNA from the other, sensitivity was also determined using spiked kidney tissue from sole with different relative amounts of T. maritimum and E. tarda. Here, 100 μl of a suspension containing ca. 107 CFU/ml and prepared from one or bacterial

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Table 1. Strains used in this work Species

Strain

Source

Edwardsiella tarda

ACC35.1

Scophthalmus maximus

Edwardsiella tarda

HL1.1

Scophthalmus maximus

Edwardsiella tarda

RM288.1

Scophthalmus maximus

Edwardsiella tarda

ACR419.1

Solea senegalensis

Edwardsiella tarda

CECT489T

Humans

Edwardsiella tarda

NCIMB2034

Fish specie

Tenacibaculum maritimum

ACC13.1

Solea senegalensis

Tenacibaculum maritimum

PC503.1

Solea senegalensis

Tenacibaculum maritimum

PC424.1

Scophthalmus maximus

Tenacibaculum maritimum

NCIMB2154T

Pagrus major

Tenacibaculum maritimum

IEO19.1

Solea senegalensis

Edwardsiella ictaluri

CECT 885

Ictalurus punctatus

Hafnia alvei

05/1403

Oncorhynchus mykiss

Yersinia ruckeri

SAG 4.1

Oncorhynchus mykiss

Yersinia ruckeri

252/05

Oncorhynchus mykiss

Yersinia ruckeri

01 1651

Oncorhynchus mykiss

Escherichia coli

FV9980

Humans

Enterobacter cloacae

TM 83/03

Scophthalmus maximus

Enterobacter aerogenes

RPM 799.1

Scophthalmus maximus

Tenacibaculum aestuarii

JCM13491T

Tenacibaculum ovolyticum

NBRC 15947

Tenacibaculum gallaicum

DSM 18841T

Seawater

Tenacibaculum discolor

DSM18842T

Solea senegalensis

Tenacibaculum litoreum

JCM13039T

Tidal flat sediment

Tenacibaculum soleae

CECT7292

Solea senegalensis

Tenacibaculum amylolyticum

NBRC 16310T

Avrainvillea riukiuensisÂ

Tenacibaculum mesophilum

NBRC 16307T

Halichondria okadaiÂ

Tenacibaculum lutimaris

DSM16505T

Tidal flat

Tenacibaculum dicentrarchi

CECT 7612

Dicentrarchus labrax

Flavobacterium psychrophilum

PT41

Oncorhynchus mykiss

Pseudomonas fluorescens

ATCC 13525T

Pre-filter tanks

Pseudomonas aeruginosa

ATCC 27853

Oncorhynchus mykiss

Aeromonas salmonicida ssp. salmonicida

ACR 215

Aeromonas media

ATCC 33907

Aeromonas hydrophila

80A1

Oncorhynchus mykiss

Aeromonas caviae

1.25

Humans

Vibrio ordalii

NCIMB 2167T

Oncorhynchus kisutch

Vibrio splendidus

ATCC33125

Marine fish

Vibrio pelagius

ATCC 25916T

Seawater

Vibrio pelagius

NCIMB 1900>T

Seawater

Vibrio tubiashii

EX 1

Crassostrea gigas

Vibrio harveyi

ATCC 14126

Vibrio vulnificus

ATCC 29307

Humans

Vibrio nereis

ATCC 25917T

Seawater

Vibrio anguillarum

R82

Scophthalmus maximus

Photobacterium damselae ssp. damselae

RG-91

Scophthalmus maximus

Photobacterium damselae ssp. piscicida

DI-21

Sparus aurata

T

Tidal flat sediment T

T

T

Hippoglossus hippoglossus

Scophthalmus maximus T

T

T

Fish farm effluent

Talorchestia sp.

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species (E. tarda or T. maritimum) was mixed with the same volume of one of the serial dilutions (from 108 to 10 cells/ml) prepared from the other species.

DNA extraction and the determination of detection limits were performed as described above.

Experimental and natural fish infections. To determine the applicability of the mPCR protocol in infected fish, batches of five turbot and five sole (average weigh 10–12 g) were inoculated with 0.1 ml of a suspension of E. tarda (ACC35.1 isolate) and/or T. maritimum (NCIMB2154T strain) at a concentration of 103 and 105 CFU/ml, respectively. Three batches of fish were inoculated with E. tarda (strain ACC35.1), three batches with T. mari­ timum (strain NCIMB2154T), and three batches with both strains. As the negative control, three batches of fish were “infected” with sterile PBS and maintained under the same conditions as the experimentally infected fish.

The fish were maintained in 50-liter aquaria with continuous aeration and a water temperature of 17 ± 1 ºC. Five days post-infection, the kidneys were collected from all turbot and sole fish and DNA was extracted as previously described. Classical bacteriological analysis by standard plate culture was also performed.

To validate the mPCR in diseased fish held in natural conditions, 30 turbot specimens ranging in weight from 50 to 100 g and showing disease symptoms were tested. The fish were collected from a rearing facility with previous natural outbreaks of edwardsiellosis and tenacibaculosis. The same number of apparently healthy fish sent to our laboratory for routine analysis were tested using the mPCR assay. Kidney samples were analyzed as previously described. Conditions for DNA extraction, PCR amplification, and PCR product visualization were the same as described above. In parallel, classical bacteriological analyses were performed to confirm the presence of E. tarda and/or T. maritimum.

Results

products) using the two primer pairs was 45 °C. The specificity of the method was evaluated using a DNA mixture prepared from the two target pathogens, which yielded amplification products of 1088 bp and 445 bp for T. maritimum and E. tarda, respectively. Non-specific amplifications were not observed using DNA from other taxonomically and/or ecologically related bacteria. The reproducibility of the PCR was demonstrated in that the same results were obtained in at least three independent PCR assays and using two different thermal cyclers. As expected, with the universal primers pA/pH a PCR product of the predicted size (1501 bp) was generated in all strains tested.

Sensitivity of mPCR assay. To assess the sensitivity of the method, the mPCR was performed using DNA extracted from the bacterial serial dilutions. The expected 1088-bp and 445-bp PCR products were obtained in samples containing as few as 200 and 4 cells of T. maritimum and E. tarda, respectively, per PCR tube (2 × 105 ± 0.2 and 4 × 103 ± 0.3 CFU/ml; Fig. 1).

The the mPCR protocol was also tested using DNA templates from fish tissues seeded with different concentrations of the two pathogens. The results demonstrated the presence of these bacteria in kidney, liver, and spleen. The detection limits in these assays were the same, regardless of the type of tissue: 2 × 105 ± 0.2 CFU/g for T. maritimum and 4 × 105 ± 0.3 CFU/g for E. tarda (200 and 400 cells per PCR tube, respectively; Fig. 2). Moreover, large amounts of E. tarda or T. maritimum (107 CFU/ml) had no effect on the detection limit of either pathogen; the values were of the same order of magnitude as described above (105 CFU/g; data not shown).

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Specificity of the mPCR assay. The mPCR method was optimized for the simultaneous detection of two bacteria, T. maritimum and E. tarda, using specific primer pairs. The annealing temperature that produced the best amplification (in terms of band intensity and the absence of non-specific

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Fig. 1. Sensitivity of the mPCR protocol as determined using purified DNA from serial dilutions of mixed cultures of Edwardsiella tarda ACC35.1 and Tenacibaculum maritimum NCIMB2154 strains. Lanes 1 and 11: Fast Ruler low range DNA ladder (50–1500 pb); lanes 2–10: dilutions ranging from 109 to 101 cells/ml from E. tarda and T. maritimum. Numbers on the left indicate the specific amplified product in bp.

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Fig. 2. Sensitivity of the mPCR protocol using purified DNA from serial dilutions of kidney samples seeded with Edwardsiella tarda ACC35.1 and Tenacibaculum maritimum NCMIB 2154. Lanes 1 and 11: Fast Ruler low range DNA ladder (50–1500 pb); lanes 2–10: dilutions ranging from 109 to 101 cells/ml (lane 8) from E. tarda and T. maritimum. Numbers on the left indicate the specific amplified product in bp.

Fig. 3. Results obtained in the detection of experimentally infected fish by mPCR. Lanes 1 and 6: Fast Ruler low range DNA ladder (50– 1500 pb); lane 2: fish infected simultaneously with Edwardsiella tarda and Tenacibaculum maritimum; lane 3: fish infected with T. maritimum only; lane 4: fish infected with E. tarda only; lane 5: negative control. Numbers on the left indicate the specific amplified product in bp.

(445 bp) was not observed. The mPCR results were confirmed by the isolation of filamentous colonies in FMM plates, which were identified as T. maritimum by classical biochemical methods as well as by specific PCR [2]. Consistent with the mPCR results, E. tarda was not detected, neither by growth in TSA-1 nor by specific PCR [7]. Also, in apparently healthy turbot neither of the pathogens could be isolated and the PCR amplifications were negative.

Discussion Currently, aquaculture is one of the fastest growing food production systems in the world. In Europe, both turbot,

Int Microbiol

Experimental and natural fish infections. The mPCR was also applied to kidney samples from experimentally infected fish. Using this technique, E. tarda and T. maritimum were detected even in fish that had not yet developed the disease. In fact, after 5 days, none of the killed turbot or sole showed any clinical symptoms and colonies from either of the two pathogens were not detected when the internal organs were plated on agar plates. By contrast, the mPCR was able to detect one or both pathogens (depending on the inoculation trials) in all experimentally infected fish (Fig. 3).

As expected, negative controls produced no amplifications.

Among the samples obtained from naturally infected fish, T. maritimum was detected in kidney samples, producing the specific 1088 bp product, whereas the E. tarda specific band

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the most commonly produced flatfish, and Senegalese sole (Solea senegalensis), because of its relatively fast growth and highly appreciated meat, are commercially important species.

Under intensive aquaculture conditions, healthy-looking fish without clinical signs may carry pathogens, posing a serious risk for the spread of diseases among fish populations. The bacterial diseases tenacibaculosis (Tenacibaculum sp.) and edwardsiellosis (E. tarda) may result in high mortalities in turbot and sole. The rapid detection of these pathogens would allow for their effective control. PCR-based methods are one of the best tools for the diagnosis of these and other fish bacterial diseases because of their specificity, sensitivity, and rapid performance. In this study, we optimized an mPCR protocol for the simultaneous diagnosis of tenacibaculosis and edwardsiellosis in flatfish.

The detection limits obtained from cultures of T. maritimum and E. tarda in the present study, in which the two PCR primer pairs were simultaneously used in a unique PCR protocol, were similar or even lower than those previously obtained by our group [2,7]. In those studies, the detection limit for E. tarda in mixed bacterial cultures was of the order of 105 CFU/ ml [7], while in the current work the sensitivity was of the order of 2 log-units. These values are within the same order of magnitude or even lower than those reported by other authors for E. tarda detection [10,11]. For T. maritimum, the only published molecular diagnostic method is the PCR previously evaluated by our group, in which the sensitivity was between 1.6 × 104 and 1.1 × 105 CFU/ml [2]. Therefore, our newly developed mPCR provides a powerful tool for the accurate detection of E. tarda and T. maritimum from bacterial cultures.

The detection limits obtained from DNA extracted from fish tissues were approximately 105 CFU/g for both pathogens.

These values were comparable to those previously reported [2,7] for the individual PCR protocols of T. maritimum and E. tarda [2,7] and to those previously described in other studies of the simultaneous detection of important fish pathogens [1, 12,13,17]. mPCR is generally thought to be less sensitive than single PCR because of competition for reaction reagents, especially if the assays differ in their amplification efficiencies or one or more of the target organisms is present in high numbers [17]. However, we found that large amounts of DNA from one of the two fish pathogens did not significantly alter the amplification sensitivity of DNA from the other pathogen, as the detection limits were within the same order of magnitude as those obtained previously [2,7]. Finally, the applicability of this technique was also demonstrated in experimentally and naturally infected fish. The speed, simplicity, sensitivity, and specificity of the mPCR developed

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in this study and the importance of these target pathogens in marine aquaculture make this protocol a very useful tool for the early and simultaneous detection of T. mari­timum and E. tarda in fish cultures.

Acknowledgments. The present research was funded in part by European Project Maximus FP7-SME-2011-286200 and grant AGL201231049 from Ministerio de Economía y Competividad (Spain).

Competing interests. None declared

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J. Fish Dis 29:541-547 6. Castro N, Magariños B, Núñez S, Toranzo AE (2007) Reassessment of the Tenacibaculum maritimum serotypes causing mortalities in cultured marine fish. Bull Eur Assoc Fish Pathol 27:229-233 7. Castro N, Toranzo AE, Núñez S, Osorio CR, Magariños B (2010) Evaluation of four polymerase chain reaction primer pairs for the detection of Edwardsiella tarda in turbot. Dis Aquat Org 90:55-61 8. Castro N, Magariños B, Núñez S, Barja JL, Toranzo AE (2011) Pathogenic potential of Edwardsiella tarda strains isolated from turbot.

Fish Pathol 46:27-30 9. Edwards U, Rogall T, Blocker H, Emde M, Bottger EC (1989) Isolation and direct complete nucleotide determination of entire genes.

Characterization of a gene coding for 16S ribosomal RNA. Nucleic Acids Res 17:7843-7853 10. Lan J, Zhang XH, Wang Y, Chen J, Han Y (2008) Isolation of an unusual strain of Edwardsiella tarda from turbot and establish a PCR detection technique with the gyrB gene. J Appl Microbiol 105:644-651 11. Li GY, Mo ZL, Li J, Xiao P, Hao B, Guo YH (2013) Development of a multiplex PCR for the identification of pathogenic Edwardsiella tarda and application to edwardsiellosis diagnostics. J Fish Dis 36:151-157 12. Mata AI, Gibello A, Casamayor A, Blanco MM, Domínguez L, Fernández-Garayzábal JF (2004) Multiplex PCR assay for detection of bacterial pathogens associated with warm-water streptococcosis in fish.

Appl Environ Microbiol 70:3183-3187 13. Osorio CR, Toranzo AE, Romalde JL, Barja JL (2000) Multiplex PCR assay for ureC and 16S rRNA genes clearly discriminates between both subspecies of Photobacterium damselae. Dis Aquat Org 40:177-183

MULTIPLEX PCR FOR T. MARITIMUM AND E. TARDA

14. Pazos F, Santos Y, Macias AR, Nu単ez S, Toranzo AE (1996) Evaluation of media for the successful culture of Flexibacter maritimus. J Fish Dis 19:193-197 15. Sakai T, Iida T, Osatomi K, Kanai K (2007) Detection of type 1 fimbrial genes in fish pathogenic and non-pathogenic Edwardsiella tarda strains

by PCR. Fish Pathol 42:115-117 16. Suzuki M, Nakagawa Y, Harayama S, Yamamoto S (2001) Phylogenetic analysis and taxonomic study of marine Cytophaga-like bacteria: proposal for Tenacibaculum gen. nov. with Tenacibaculum maritimum comb. nov. and Tenacibaculum ovolyticum comb. nov., and description of Tenacibaculum mesophilum sp. nov. and Tenacibaculum amylolyticum sp. nov. Int J Syst Evol Microbiol 51:1639-1652

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17. Tapia-Cammas D, Ya単ez A, Arancibia G, Toranzo AE, Avenda単o-Herrera R (2011) Multiplex PCR for the detection of Piscirickettsia salmonis, Vibrio anguillarum, Aeromonas salmonicida and Streptococcus phocae in Chilean marine farms. Dis Aquat Org 97:135-142 18. Thoesen JC (ed) (1994) Suggested procedures for the detection and identification of certain finfish and shellfish pathogens. Fish Health Section, American Fisheries Society, 4th ed, version 1. Bethesda, MD, USA 19. Toyama T, Kita-Tsukamoto K, Wakabayashi H (1996) Identification of Flexibacter maritimus, Flavobacterium branchiophilum and Cytophaga columnaris by PCR targeted 16S Ribosomal DNA. Fish Pathol 31:25-31

PERSPECTIVES International Microbiology (2014) 17:119-129 doi:10.2436/20.1501.01.214 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

CDC Report on the Potential Exposure to Anthrax* .

Centers for Disease Control and Prevention, 1600 Clifton Road, Atlanta, GA, USA

Executive Summary The Centers for Disease Control and Prevention (CDC) conducted an internal review of an incident that involved an unintentional release of potentially viable anthrax within its Roybal Campus, in Atlanta, Georgia. On June 5, 2014, a laboratory scientist in the Bioterrorism Rapid Response and Advanced Technology (BRRAT) laboratory prepared extracts from a panel of eight bacterial select agents, including Bacillus anthracis (B. anthracis), under biosafety level (BSL) 3 containment conditions. These samples were being prepared for analysis using matrix-assisted laser desorption/ionization timeof-flight (MALDI-TOF) mass spectrometry, a technology that can be used for rapid bacterial species identification.

What Happened This protein extraction procedure was being evaluated as part of a preliminary assessment of whether MALDI-TOF mass spectrometry could provide a faster way to detect anthrax compared to conventional methods and could be utilized by emergency response laboratories. After chemical treatment for 10 minutes and extraction, the samples were checked for

sterility by plating portions of them on bacterial growth media. When no growth was observed on sterility plates after 24 hours, the remaining samples, which had been held in the chemical solution for 24 hours, were moved to CDC BSL-2 laboratories. On June 13, 2014, a laboratory scientist in the BRRAT laboratory BSL-3 lab observed unexpected growth on the anthrax sterility plate. While the specimens plated on this plate had only been treated for 10 minutes as opposed to the 24 hours of treatment of specimens sent outside of the BSL-3 lab, this nonetheless indicated that the B. anthracis sample extract may not have been sterile when transferred to BSL-2 laboratories.

Why the Incident Happened The overriding factor contributing to this incident was the lack of an approved, written study plan reviewed by senior staff or scientific leadership to ensure that the research design was appropriate and met all laboratory safety requirements.

Several additional factors contributed to the incident: • Use of unapproved sterilization techniques • Transfer of material not confirmed to be inactive

*NOTE: On July 21, 2014, Timothy J. Donohue, President of the American Society for Microbiology, and Ronald M. Atlas, Chair of the Public and Scientific Affairs Board of the ASM, sent to all ASM members a letter [http://www.asm.org/index.php/public-policy/93-policy/93014-biosafety-7-14] concerning the recent events at the Centers for Disease Control and Prevention (CDC) and the National Institutes of Health (NIH), which are documented in the CDC Report issued on July 11, 2014, that International Microbiology reproduces here (the original can be accessed through the following link [http://www.cdc.gov/od/ science/integrity/docs/Final_Anthrax_Report.pdf]). On July 31, Donahue and Atlas sent a second letter [http://www.asm.org/index.php/clinical-microbiologyupdate/137-policy/documents/statements-and-testimony/93024-durc-7-31-14] stating that “The ASM has a long history of supporting and informing public policy that is based on the essential principle of ensuring protection of public health and safety without unduly encumbering legitimate fundamental scientific research, clinical and diagnostic testing for the treatment and prevention of infectious diseases. The ASM has and continues to have a position that microbiological research must be done safely and in accordance with regulations governing the proper conduct of that research.” They also referred to the research for the treatment, control and prevention of infectious diseases, which is crucial and require the utilization of pathogenic material as well as laboratory safety and security measures. To provide microbiologists with background information on this topic, Donohue and Atlas informed that “The ASM has also prepared a history of issues and policy related to biological research with select agents and toxins, high containment laboratories, dual use research of concern and gainof-function (GOF) research. This document is posted on the ASM website at [http://www.asm.org/images/PSAB/History-SelectAgents.pdf].”

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• Use of pathogenic B. anthracis when non-pathogenic strains would have been appropriate for this experiment • Inadequate knowledge of the peer-reviewed literature • Lack of a standard operating procedure or process on inactivation and transfer to cover all procedures done with select agents in the BRRAT laboratory.

What Has CDC Done Since the Incident Occurred CDC’s initial response to the incident focused on ensuring that any potentially exposed staff were assessed and, if appropriate, provided preventive treatment to reduce the risk of illness if exposure had occurred. CDC also ceased operations of the BRRAT laboratory pending investigation, decontaminated potentially affected laboratory spaces, undertook research to refine understanding of potential exposures and optimize preventive treatment, and conducted a review of the event to identify key recommendations.

To evaluate potential risk, research studies were conducted at a CDC laboratory and at an external laboratory to evaluate the extent to which the chemical treatment used by the BRRAT laboratory inactivated B. anthracis. Two preparations were evaluated: vegetative cells and a high concentration of B. anthracis spores. Results indicated that this treatment was effective at inactivating vegetative cells of B. anthracis under the conditions tested. The treatment was also effective at inactivating a high percentage of, but not all B. anthracis spores from the concentrated spore preparation.

A moratorium is being put into effect on July 11, 2014, on any biological material leaving any CDC BSL-3 or BSL-4 laboratory in order to allow sufficient time to put adequate improvement measures in place.

What’s Next Since the incident, CDC has put in place multiple steps to reduce the risk of a similar event happening in the future. Key recommendations will address the root causes of this incident and provide redundant safeguards across the agency, these include: • The BRRAT laboratory has been closed since June 16, 2014, and will remain closed as it relates to work with any select agent until certain specific actions are taken • Appropriate personnel action will be taken with respect to individuals who contributed to or were in a position to prevent this incident • Protocols for inactivation and transfer of virulent pathogens throughout CDC laboratories will be reviewed

CDC REPORT.

• CDC will establish a CDC-wide single point of accountability for laboratory safety • CDC will establish an external advisory committee to provide ongoing advice and direction for laboratory safety • CDC response to future internal incidents will be improved by rapid establishment of an incident command structure • Broader implications for the use of select agents, across the United States will be examined.

This was a serious event that should not have happened.

Though it now appears that the risk to any individual was either non-existent or very small, the issues raised by this event are important. CDC has concrete actions underway now to change processes that allowed this to happen, and we will do everything possible to prevent a future occurrence such as this in any CDC laboratory, and to apply the lessons learned to other laboratories across the United States.

Background This report reviews circumstances leading to June 2014 incident in which CDC staff members were potentially exposed to viable Bacillus anthracis. The incident occurred after B. anthracis extract was transferred from CDC’s Bioterrorism Rapid Response and Advanced Technology (BRRAT) biosafety level (BSL) 3 laboratory to BSL-2 laboratories without proper assurance that the extract did not contain viable cells or spores.

This is not the first time an event of this nature has occurred at CDC, nor the first time it occurred from the BRRAT laboratory. At the time of this writing, CDC is aware of four other such incidents in the past decade. In a prior incident in 2006, CDC’s BRRAT laboratory transferred vials of anthrax DNA to the Lawrence Livermore National Laboratory (LLNL) and a private laboratory. The BRRAT laboratory believed that they had inactivated the samples, but upon receipt and testing of the samples at LLNL, viable B. anthracis was detected. The BRRAT laboratory implemented new quality assurance procedures to ensure non-viability of DNA preparations of select agents and developed policies that require the signature of the laboratory’s principal investigator prior to shipping or transferring DNA derived from bacterial select agents. These procedures were not followed in the current incident, which did not specifically involve preparation of DNA for transfer. Also in 2006, DNA preparations shipped from another CDC laboratory were found to contain live Clostridium botulinum due to the use of inadequate inactivation procedu-

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res. In 2009, newly available test methods showed that a strain of Brucella, thought to have been an attenuated vaccine strain and previously shipped to LRN laboratories as early as 2001, was not the vaccine strain. The vaccine strain is not considered to be a select agent, while the strain that was actually shipped is a select agent.

As this report was being finalized, CDC leadership was made aware that earlier this year a culture of low-pathogenic avian influenza was unintentionally cross-contaminated at a CDC influenza laboratory with a highly pathogenic H5N1 strain of influenza and shipped to a BSL-3, select-agent laboratory operated by the United States Department of Agriculture (USDA). The CDC influenza laboratory where this incident occurred is now closed and will not reopen until adequate improvements are put in place. Although CDC is continuing to investigate and review this matter, Attachment A provides current information on the incident and the agency’s response.

Effective, validated inactivation protocols for B. anthracis have been published. Cultures of B. anthracis cells and spores can be completely inactivated through established protocols using heat (e.g., boiling for 10 minutes or autoclaving for 15 minutes), irradiation (1 million rad), or various chemical treatments (e.g., hydrogen peroxide, peracetic acid, formalin, or gaseous ethylene oxide). In general, longer treatment times and/or higher concentrations are required for inactivation of spores compared to inactivation of viable cells. Solutions can also be sterilized by filtration, through a 0.1 micron filter, to remove viable cells and spores.

Space decontamination can be achieved through one of two approved liquid decontamination methods and one vapor method. A solution of freshly made dilution of household bleach (10% bleach by volume), pH adjusted to 7.0 with acetic acid, is recognized by the Environmental Protection Agency (EPA) to kill B. anthracis spores with a minimum contact time of 10 minutes. The EPA also registered the use of Spor-Klenz® (STERIS®) as a sterilant, as a 1:99 water dilution of the concentration is effective as a sporocide with a minimum contact time of 30 minutes. Vapor phase hydrogen peroxide is also available at CDC as a room disinfectant.

Laboratories CDC laboratories conduct research that is critical to better detect, respond to, and prevent disease and bioterrorism. Research done in CDC laboratories helps identify better ways to detect these infectious agents rapidly. The Laboratory Response Network (LRN) is a network of laboratories that can respond to biological and chemical threats and other public

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health emergencies. It includes state and local public health, veterinary, military, and international labs. The BRRAT laboratory provides technical and scientific support for the approximately 150 laboratories in the LRN. The BRRAT laboratory contains both BSL-3 and BSL-2 labs and was established in 1999 in accordance with Presidential Decision Directive 39, which outlined national anti-terrorism policies and assigned specific missions to federal departments and agencies (http:// www.bt.cdc.gov/lrn/). The BRRAT laboratory provides quality assurance for the specialized reagents used in the LRN and has performed studies with the goal of improving the performance and reliability of tests used to detect biological threat agents. Bacillus anthracis is of particular concern because it can and has been used as a weapon.

Two CDC laboratories received the extracts prepared by the BRATT laboratory BSL-3 laboratory: the Bacterial Special Pathogens Branch laboratory (BSPB laboratory); and the Biotechnology Core Facility Branch (BCFB laboratory).

Methods Used in Reviewing this Incident A CDC team of scientists and leaders interviewed laboratory scientists involved directly with the incident and others who had specific knowledge of the incident and of immediate response activities. Each interview consisted of a standardized set of questions, as well as specific questions based on an individual’s role and responsibilities. Standard operating procedures (SOPs), protocols, and training records were also reviewed.

Description of the Event The BRRAT laboratory was evaluating matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry, which can identify bacteria by bacterial protein “fingerprints.” It is faster and less expensive than conventional species-identification methods, which require culture of organisms on selective bacterial media or extraction and characterization of bacterial nucleic acids. The project was a collaboration among the BRRAT, BSPB, and BCFB laboratories. The researchers intended to use the data collected to submit a joint proposal to CDC’s Office of Public Health Preparedness and Response to fund further evaluation of the MALDI-TOF method because MALDI-TOF is increasingly being used by clinical and hospital laboratories for infectious disease diagnostics.

On June 2, 2014, the BRRAT laboratory supervisor contacted a subject matter expert who had successfully used this technology to identify three pathogenic species of Brucella.

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In response to the BRRAT laboratory supervisor’s request for assistance and advice, a BSPB laboratory supervisor offered to share the methodology, results, and inactivated bacterial preparations used by the BSPB laboratory in their work with Brucella. The BSPB laboratory had modified the MALDITOF equipment manufacturer’s sample preparation protocol to optimize the results for bacterial protein sample extractions of Brucella. In this extraction procedure, each organism is treated with ethanol, then with 70% formic acid for 10 minutes, followed by the addition of 100% acetonitrile, and then is incubated at room temperature. The method used by the BSPB laboratory also incorporated a sterility check of the extract after 10 minutes of incubation in the extraction solution. Specifically, an aliquot of the extract was spread on an agar plate, incubated for 48 hours, and then examined for growth. If no growth was visible, the extract was considered to be sterile and could be safely transferred from the BSL-3 laboratory to a BSL-2 laboratory for processing for use in the MALDI-TOF equipment.

The BSPB laboratory protocol did not call for filtration of the bacterial extract prior to transfer from the BSL-3 laboratory because it had been determined that the extraction procedure inactivated the three species of Brucella tested. It is important to note that, unlike B. anthracis, Brucella does not form spores. Bacterial spores are relatively resistant to harsh conditions, such as the chemicals used in this extraction procedure, and are more difficult to kill than vegetative cells. As a result, additional procedures (e.g., filtration) can be used when working with spore-forming bacteria, such as B. anthracis, to ensure specimens are rendered non-viable.

The BRRAT laboratory supervisor instructed a laboratory scientist to obtain the written protocol for sample preparation from BSPB laboratory. The BSPB laboratory provided a sample preparation protocol, which did not include a viability SOP. The supervisor requested that virulent strains of eight select agents, including B. anthracis, be used for the initial experiment. On June 5, 2014, the laboratory scientist followed the modified protocol to prepare eight individual organism extracts for use in the MALDI-TOF. Another scientist in the BRRAT laboratory raised the question of whether filtration of the extracts might affect the MALDI-TOF results. To answer this question the laboratory scientist split each extract into two aliquots and filtered one aliquot through a 0.1 micron filter. After a 10 minute incubation period, filtered and unfiltered extracts were then plated onto agar and incubated for 24 hours to check the extracts for sterility. The decision to incubate for 24 hours, rather than 48 hours (as recommended by the BSPB staff member) was made by the first laboratory

CDC REPORT.

scientist based on the individual’s own understanding of information conveyed by the laboratory scientist in the BSPB laboratory during a telephone discussion of the protocol.

All work was performed in a biological safety cabinet in the BRRAT BSL-3 laboratory with both BRRAT laboratory scientists present. The first laboratory scientist was primarily involved in performing the extraction, and the second was there to observe and learn the procedure. Both were jointly involved in filtering material, plating onto media, and reading sterility plates at 24 hours. After 24 hours of incubation, they observed no growth on any of the 16 sterility plates that had been prepared after 10 minutes of formic acid treatment. The first laboratory scientist planned to autoclave the plates, then discard them; however, the individual had difficulty opening the autoclave door. As a result, the plates were returned to the incubator and left for 7 additional days.

The first laboratory scientist moved the extracts from the BRRAT laboratory BSL-3 lab to an adjoining BSL-2 laboratory that is also part of the BRRAT laboratory. At this point, the protein extracts had been held in the formic acid/acetonitrile solution for 24 hours. The first laboratory scientist then continued with the process of preparing the material for analysis by MALDI-TOF, and then moved preparations or aliquots of the protein extracts made from the BRRAT’s BSL-2 laboratory to the BSPB and BCFB laboratories on three separate days: June 6, June 11, and/or June 12, 2014.

On June 13, 2014, the second BRRAT laboratory scientist removed the sterility testing plates after 8 days in the BSL-3 incubator for autoclaving and disposal and discovered growth on the sterility plate that had been plated with unfilter­e d B. anthracis. The growth was confirmed as B. anthracis by real-time polymerase chain reaction using the LRN B. anthracis identification assays. It is not known at what point after the initial 24 hour incubation period that growth occurred. If the plates had been autoclaved after 24 hours, as planned, the event would not have been discovered.

The incident was immediately reported to the CDC Select Agent Program Responsible Official within CDC’s Environment, Safety and Health Compliance Office (ESHCO) and DSAT. CDC personnel decontaminated the affected rooms using the liquid decontamination methods described above (see Background). Laboratory floors, benchtops, equipment, and other affected areas (e.g., room door handles) were decontaminated as part of this process. Two potentially affected refrigerators were moved to a secure BSL-3 facility and decontaminated using vapor phase hydrogen peroxide. Rooms will remain closed until the procedures have been validated as EPA compliant by an external safety expert.

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After the incident was discovered, two laboratory studies were undertaken to determine if the formic acid and acetonitrile treatment was effective at inactivating laboratory specimens of B. anthracis: one at CDC and one at an independent LRN laboratory at the Michigan Department of Community Health (MDCH). The CDC study evaluated the effect of treatment exposure times of 10 minutes in formic acid and after 6 hours and 24 hours in formic acid/acetonitrile on B. anthracis vegetative cells. In addition, the CDC study evaluated treatment exposure times of 10 minutes in formic acid and 24 hours in formic acid/acetonitrile using high-concentrations of B. anthracis spores. Cultures from treated cells and spores were monitored daily for viability for up to 8 days post-treatment. The MDCH study independently evaluated the efficacy of the formic acid/acetonitrile treatment on B. anthracis vegetative cells. This study used samples that were taken at three different time points: immediately on addition of the formic acid and subsequently at 1 hour and 24 hours post-treatment.

The MDCH cultures were monitored for up to 8 days for viability.

Findings from both the CDC internal study and the MDCH indicate that the formic acid and formic acid/acetonitrile treatment were effective at inactivating vegetative cells of B. anthracis. No viable material was recovered from formic acid and formic acid/acetonitrile treated cells. These findings were consistent for the 8-day study duration. The formic acid and formic acid/acetonitrile treatments were effective at inactivating a high percentage, but not all, B. anthracis spores. From a starting suspension of 50,000 B. anthracis spores (500,000 per milliliter), which had been treated for 24 hours with the extraction process, there were a total of four colony forming units of growth in the 8-day study period.

Based on review of all aspects of the incident, it appears that while exposure of staff to viable B. anthracis was not impossible, it is extremely unlikely that this occurred. All or the great majority of B. anthracis cells and spores in the sample would have been inactivated by the 24-hour treatment (versus the 10 minute sample which grew anthrax at some point between day 2 and day 8 of incubation).

Findings Incident-related Findings The overriding factor contributing to this incident was the lack of an approved, written study plan reviewed by CDC senior staff, such as laboratory, branch, or division scien-

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tific leadership, to ensure that the research design was appropriate and met all laboratory safety requirements. The first BRRAT laboratory scientist was trained to work in the BSL-3 environment, including training in pathogen-specific procedures for the work normally performed. However, the individual had not performed this specific procedure with pathogenic select agents (the procedure was new to the laboratory) and should not have been instructed to proceed without submitting a complete protocol for review and approval. Further, a written protocol to certify the sterility of material to be transferred to BSL-2 laboratories was not in place, and the BSL-2 laboratories did not have an SOP that required receipt of written certification of non-viability for transfers prior to acceptance of microbiologic material. There was also inadequate supervisory oversight of a relatively new laboratory scientist performing a new experiment with virulent strains.

The first laboratory scientist also assumed that the protocol was appropriate for B. anthracis. It appears that there was incomplete communication between the two BRRAT laboratory scientists and the BSPB laboratory scientist about what was planned by the BRRAT laboratory and what had previously been done by the BSPB laboratory. The procedure used by the BSPB laboratory for Brucella species did not include a filtration step because the BSPB laboratory determined it was not necessary for extracts of Brucella based on the sterility testing they had done on extract material of three species of Brucella. Since B. anthracis forms spores that are more resistant to inactivation by chemicals than vegetative cells, the BRRAT laboratory scientistâ&#x20AC;&#x2122;s assumption that the same treatment would apply to B. anthracis was incorrect.

The BRRAT laboratory scientist did not plan to filter extracts because it was not part of the BSPB laboratory protocol.

The BRRAT laboratory scientist was aware that all DNA preparations of B. anthracis were filtered before leaving the BSL-3 laboratory, but assumed that it was not necessary for MALDI-TOF preparations because a filtration step was not included in the protocol. The BRRAT laboratory scientist had no previous experience transferring select agent-derived materials, other than transferring DNA preparations, from BSL-3 to BSL-2 laboratories. The BRRAT laboratoryâ&#x20AC;&#x2122;s SOP for assuring sterility was specific for DNA preparations, and SOPs for other materials do not appear to have been in place. The SOP for DNA preparations (with which the first BRRAT laboratory scientist was familiar) indicated that sterility check plates for B. anthracis should be held for 24-48 hours.

It is not clear that waiting 48 hours rather than 24 hours to transfer the extracts would have prevented this incident. The bacterial cells or spores were damaged by the extraction pro-

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cedure, and the direct plating of the extract carried over chemicals which could have inhibited growth. Acceptable practice would have been to utilize validated methods to confirm sterility.

The following actions contributed to the incident: 1. Use of unapproved sterilization techniques: Staff in the BRRAT laboratory used sample preparation techniques for protein extraction from the manufacturer of the MALDI-TOF equipment, modified by the BSPB laboratory for non-spore forming bacteria (Brucella species) to sterilize B. anthracis, a spore-forming bacterium. A laboratory scientist modified the methods from the BSPB laboratory to include comparing filtration versus non-filtration in preparing 16 plates (half filtered and half not filtered). This modification was done to assess any effects on the MALDITOF results, not to assure sterility. The incubation period was also shortened from 48 hours to 24 hours.

2. Transfer of material not confirmed to be inactive: After 24 hours without observing growth on the sterility plates, the BRRAT laboratory scientist moved the extracts from the BRRAT laboratory BSL-3 laboratory to an adjoining BSL-2 laboratory, and then continued with the process of preparing the material for analysis by MALDI-TOF. The BRRAT laboratory scientist then moved the extracted materials from the BRRAT laboratoryâ&#x20AC;&#x2122;s BSL-2 laboratory to the BCFB and BSPB laboratories on three separate days: June 6, June 11, and/or June 12, 2014. There is a lack of written procedures which had been validated to reliably ensure that organisms were no longer viable prior to removing microbiological material from BSL-3 containment.

3. Use of pathogenic B. anthracis when non-pathogenic strains would have been appropriate for this experiment: The BRRAT laboratory supervisor instructed the laboratory scientist to use virulent strains because of the possibility that avirulent strains might not yield the same MALDI-TOF profile. However, the instrument manufacturer states that the system identifies bacteria to only the species level and would not distinguish strains of the same species. The use of avirulent strains to develop protocols would have been appropriate, particularly when conducting a pilot study.

4. Inadequate knowledge of the peer-reviewed literature by the BRRAT laboratory supervisor and scientist who

CDC REPORT.

performed the extraction: A review of the literature would have found that filtration has been recommended for inactivation of B. anthracis. There are at least two peerreviewed publications on preparation methods for MALDI-TOF work with pathogenic bacteria, including B. anthracis (Drevinek et al. Letters in Applied Microbiology 2012;55:40-46; and Lasch, et al. Analytical Chemistry 2008;80:2026-2034). While the chemicals used to process the samples differ in the two publications, both required filtration of B. anthracis material with a 0.1 micron filter to remove spores. Drevinek et al. (2012) concluded that the formic acid method (as used by the BRRAT laboratory) did not sterilize B. anthracis; they also used centrifugal filtration to remove viable particles (including spores) from B. anthracis preparations.

5. Lack of a standard operating procedure or process to document inactivation in writing in the BRRAT laboratory: With correct SOPs in place that are adhered to by staff, microbiological material would have been successfully inactivated prior to transfer to a lower containment laboratory (either intra- or inter-facility) and a record of non-viability would have been provided to the receiving laboratory; also, a written record of non-viability would have been provided prior to receipt and utilization of the microbiological materials in the BSL-2 laboratories.

Response-related Findings On June 13, 2014, two CDC staff members went to the emergency department at Emory University where they were assessed; neither presented with symptoms related to anthrax.

Staff were assessed based on their risk of potential exposure that could lead to inhalational anthrax. The number of potentially exposed staff evolved as understanding of the laboratory events unfolded. Additional potentially exposed individuals were identified through supervisor discussions with individuals believed to have handled or been in proximity to the B. anthracis material. The process of identification was slowed by multiple factors, including the evolving nature of understanding of the event. Technology 10 resources such as card key readers and security video were utilized to expand the pool of potential exposures, but this was not an immediate step in the response. Even with the use of available data, several factors made the identification process difficult, including the practice of authorized staff piggy backing (obtaining entrance to a secured area by following a colleague rather than by having all individuals swipe their own card key as should

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be done) and incomplete or inaccurate information collected from laboratory scientists reporting their path of travel with the material between labs. Protocols were not in place for the rapid identification of potentially exposed staff, possibly delaying the use of available data sources including card key readers, visitor logs, and security video logs.

Immediate and comprehensive actions were taken to identify the potentially affected laboratory rooms as well as the individuals that were or may have been in, or traveled through, these areas during the time period of possible exposure. After ascertaining the precise events that took place in the laboratories and characterizing people’s possible exposure was difficult and evolving, there were serious reservations on the part of some staff members of the affected laboratories and others about broad communication until sufficient information was gathered and verified. In retrospect, it is clear that broad communications should have occurred earlier in the process, even if more complete information was not yet available. CDC scientists who worked near the impacted laboratories commented that they first learned of the event by witnessing CDC closing and/or decontaminating laboratories rather than through direct communication regarding the ongoing event. In addition, there were inconsistencies in the decontamination practices used after the incident, which made it difficult to ensure proper methods were used. Individuals also reported the CDC clinic was overwhelmed at times during the response.

The nature of this incident required involvement of many parties from across CDC. While the roles of the responders were generally clear and appropriate actions were taken, there was no clear overall lead for the incident in the first week.

This resulted in uncertainty regarding who was responsible for making decisions and taking action.

As of July 10, 2014, no staff members are believed to have become ill with anthrax.

Actions Already Underway and Plans for the Future A moratorium was initiated July 11, 2014, on any biological material leaving any CDC BSL-3 or BSL-4 laboratory in order to allow sufficient time to put adequate improvement measures in place. In addition, CDC has already begun steps to protect staff and prevent similar incidents in the future. Key actions are planned to address the root causes of this incident.

The recommendations focus on specific actions that provide redundant safeguards across the agency.

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These actions and recommendations relate to • The BRRAT laboratory • Inactivation and transfer procedures of virulent pathogens throughout CDC laboratories • Broader improvements in biosafety in laboratories throughout CDC • CDC response to internal incidents • Broader implications for the use of select agents, including for CDC’s regulatory functions through CDC’s Division of Select Agents and Toxins.

The BRRAT Laboratory 1. The laboratory has been closed since June 16, 2014, and will remain closed as it relates to work with any select agent. This action was reinforced by USDA’s Animal and Plant Health Inspection Services (APHIS). Laboratory scientists do not have access to select agents, which have been placed in storage-only mode. The unit will remain closed with respect to select agents until the following is completed: a. An assessment and appropriate follow-up actions for all BRRAT laboratory staff to determine level of skills, training, supervision, knowledge, and expertise at all levels of the organization b. The establishment of clear, proven procedures that have been communicated to all staff for inactivation and non-viability testing of all types of materials that may be produced by the laboratories (i.e., not limited to nucleic acid preparations from one specific laboratory) and documentation of these processes c. Resolution of all findings included in this report and in the APHIS investigation report 2. Appropriate personnel action will be taken with respect to individuals who contributed to or were in a position to prevent this incident.

Inactivation and Transfer Procedures of Virulent Pathogens throughout CDC Laboratories 3. All inactivation procedures for laboratories working with select agents and other dangerous pathogens are being carefully reviewed and will be updated as needed. This includes, but is not limited to, any inactivation performed in conjunction with MALDI-TOF testing. CDC will notify the MALDI-TOF manufacturer and the Food and Drug Administration (FDA) of this event and encourage the de-

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velopment of informational materials that are clearer regarding appropriate inactivation procedures for all types of pathogens. All CDC laboratories that handle select agents and other dangerous pathogens will be confirmed to have written, validated, and verified procedures to assure materials are non-viable before being removed from containment and to assure the provision of written documentation of non-viability, including the method used, for intra- and inter-facility transfers. These procedures will include requirements that all transferring laboratories confirm non-viability by proven, effective methods before material leaves the containment laboratory and provide documentation to accompany the transfer and that the receiving laboratory confirm the materials are not viable.

When new procedures, techniques, or manufacturer methods are being considered, they must first be reviewed and evaluated through a formal process to assess their risk and incorporate them into standard CDC policies, procedures, and practices prior to implementation.

Laboratories across CDC 4. CDC will establish a lead laboratory science position to be the CDC-wide single point of accountability for laboratory safety. The creation of a single point of accountability does not reduce the responsibility of people at every level of the organization, including center, division, and branch directors, chiefs, supervisors, and all laboratory scientists to strengthen the culture of safety. This position will: a. Establish and enforce agency-wide policies that require formal review and approval of new select agent research or program protocols and provide oversight for ongoing research and program projects (e.g., yearly reviews). b. Create effective and redundant systems and controls for protocols and procedures including, but not limited to, inactivation and access to laboratories (e.g., â&#x20AC;&#x153;piggybackingâ&#x20AC;? and visitor access). c. Ensure adherence to laboratory quality and safety protocols (e.g., quality assurance that biological material is non-viable before it is shipped from CDC select agent laboratories). These protocols will be transferred to new staff whenever there is a turnover in select agent laboratories, especially when there is a new principal investigator. d. Review and monitor the implementation of training policies and procedures for new and existing staff.

5. Use an approach that identifies the points in any project where potential mistakes would have the most serious conse-

CDC REPORT.

quences that provides specific actions to avoid these mistakes. Examples of these critical points and associated preventive actions include requiring protocols to be reviewed by supervisors before they are implemented, having standard and clear procedures to inactivate infectious agents and specify how they will be transferred to other labs, having formal incident response plans in place, controlling laboratory access, and instituting regular review of laboratory processes to ensure proper safety, quality management, and compliances with Select Agent Regulations. a. Identify ways to decrease the risk of an event such as this happening again, which may include fewer laboratories working with select agents and/or a decrease in the number of pathogenic strains being studied and/or a decrease in the number of staff members working with these agents. b. Promote the use of non-pathogenic organisms in research and training activities, whenever possible. c. Accelerate the ongoing implementation of laboratory quality management systems (QMS) throughout CDC laboratories. Over the past 5 years, CDC has begun implementing a QMS for infectious disease laboratories which includes document controls such as protocol archives and approval records as an integral part. Initial adoption of QMS has focused on the laboratories with clinical diagnostic responsibilities and has greatly enhanced their safety and efficiency. Expansion into nonclinical laboratories has been ongoing and will now be accelerated as a high priority, with QMS becoming an integral part of CDC laboratory management practice.

6. CDC will establish an external advisory committee to provide ongoing advice and direction for laboratory quality and safety. It is likely this advisory committee will be established under the Federal Advisory Committee Act (FACA).

Response Efforts 7. CDC will initiate an incident command structure early in any response to an incident at CDC when an event is suspected that the incident is significant or not well understood. CDC may also leverage the assets of CDCâ&#x20AC;&#x2122;s Emergency Operations Center to help coordinate the event response under the incident commander. This does not necessarily mean activating the EOC for such a purpose, but use of the EOC facility, staff, tools, and other resources as well as coordination within CDC offices could be beneficial. Under this structure, CDC can ensure proactive and

EXPOSURE TO ANTRAX

frequent communication with staff, media, and the public.

This structure will also allow for quick access to CDC staff with unique expertise to provide surge capacity (including nurses and physicians to staff the CDC clinic), as needed.

Broader Implications for the Use of Select Agents 8. Lessons based on this incident that will be considered for broader implications. CDCâ&#x20AC;&#x2122;s DSAT program will incorporate findings and recommendations into nationwide regulatory activities to provide stronger safeguards for laboratories across the United States. For example, in its review of biosafety plans with regulated entities, DSAT will em­ phasize the importance of having proven inactivation protocols and utilizing testing for inactivated preparations prior to distribution.

Conclusion Potential exposure of CDC laboratory scientists to anthrax occurred as a result of a series of failures of one laboratory (the CDC BRRAT laboratory) to ensure that B. anthracis specimens had been inactivated before transferring them to other laboratories at CDC. This same laboratory had inadvertently transferred viable B. anthracis on a previous occasion in 2006. Review of the procedures and practices that allowed this event to occur identified: failures of policy, training, scientific knowledge, supervision, and judgment on the part of

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this laboratory. In addition, there was a lack of adequate agency-wide policies and procedures to ensure biosafety, both for decontamination of select agents and other virulent organisms as well as for biosafety more broadly. Further, biosafety policies and procedures adopted in the past were not always adhered to in the present. Response to the incident should have been better organized from the outset.

Review of the incident suggests that it is highly unlikely, but not impossible, that staff members were exposed to viable B. anthracis. None of the potentially exposed workers has become ill with anthrax. Nonetheless, this was a serious and unacceptable incident which should never have happened. A moratorium is being put into effect on July 11, 2014, on any biological material leaving any CDC BSL-3 or BSL-4 laboratory in order to allow sufficient time to put adequate improvement measures in place. Five key steps are being taken immediately: suspension of activities of this individual laboratory pending full review and remediation of all procedures and practices; agency-wide verification of adequate inactivation procedures; strengthening of biosafety agency-wide with appointment of a single point of accountability and through an external group of experts to review and advise CDC; improvement of management of internal incidents with use of an incident management system; and use of lessons learned from this incident to strengthen CDCâ&#x20AC;&#x2122;s regulatory function with regard to select agents.

Given both the critical nature of investigations to enable CDC to improve our ability to detect and respond to naturally occurring and man-made events with select agents and the paramount responsibility of ensuring the safety of CDC staff members when they do this work, CDC leadership, including the CDC Director, will track the rapid and effective implementation of these plans.

BOOK REVIEWS International Microbiology (2014) 17:129-130 ISSN 1139-6709, e-ISSN 1618-1095 www.im.microbios.org

Antimicrobial compounds.

Current strategies and new alternatives T.G Villa, P. Veiga-Crespo (eds.)

2014. Springer, Heidelberg, Germany 316 pp, 16 x 24 cm Price: € 145.59 ISBN 978-3-642-40443-6

Diseases have notably shaped the course of human history, especially when an unknown infection has “attacked” a population for the first time. Of the 1922 species of infectious agents recorded in databases, 632 are bacteria, 329 are fungi, 499 are helminths, 145 are protists, and 317 are viruses and prions. Until the 1930s, the treatment of infectious illnesses was mostly palliative; in most cases, physicians could only hope that their patients were strong enough to overcome the infection on their own. During the past 70 years, however, the morbidity and mortality associated with many communicable infectious diseases have significantly decreased in Western countries, largely because of the use of antibiotics and the implementation of well-planned vaccination strategies.

Nonetheless, infectious diseases remain the second leading cause of death globally, responsible for more than 9 million deaths (16.2 %) per year. Infections of the lower respiratory tract, diarrheal diseases, HIV/AIDS, and tuberculosis are among the top ten causes of death worldwide.

The book Antimicrobial compounds. Current strategies and new alternatives describes the state of the art of antimicrobial research, including non-antibiotic therapeutic strategies in the fight against infectious diseases. The book contains eleven chapters written by several experts in the field, including its editors T.G. Villa and P. Veiga-Crespo.

The emergence of resistance in microbial strains to the current armamentarium of antibiotics is a major threat to public health worldwide. This problem has been made even worse given the slow development of novel antibiotics: since the early 1960s, only four new classes of antibiotics have been introduced. Indeed, the global antibiotics market is still domi-

nated by antibiotic classes discovered half a century ago.

Since then, most “new” antibiotics have been chemically tailored derivatives of these well-worn scaffolds.

In the genomic era, the availability of various genomicsbased platforms including whole-genome sequencing, genotyping, gene expression profiling, and cloning, has resulted in new approaches to the increased production of known antibiotics and, perhaps more importantly, to the discovery of novel antimicrobial agents and targets (Ch. 1, 2, 5, 7, 9, 11). Most antibiotics come from soil actinomycetes, reflecting the historical bias of pharmaceutical screening programs toward these “easily” collected and culturable bacteria. Research into underexplored ecological habitats, such as marine ecosystems or insect symbiosis, have revealed new bacterial taxa that synthesize compounds with biocidal properties. Genome sequencing of known actinomycetes has revealed that these bacteria generally harbor >25 gene clusters encoding secondary metabolites. Because only one to four natural products are known from a typical bacterium under various culture conditions, researchers may thus far have discovered only 10 % of the natural products of screened strains and just 1 % of the molecules of global microbial producers. Genomics also faci­ li­tates the optimization of antibiotic biosynthetic processes in large-scale, cost-effective drug production, either by enhan­cing the flux of the desired bacterial pathways or by removing competitive biochemical pathways in the host.

The search for healing agents in plants can be traced to the ancient world. Ever since the very beginning of civilization, people have made use of plants to obtain relief from pain, heal injuries, and even to cure diseases. Plants are constantly under

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attack by microbial pathogens. As part of their defensive arsenal, they synthetize antimicrobial peptides, such as “defensins” and “thionins,” as well as other antimicrobial metabolites, including alkaloids, flavonoids, and essential oils (Ch. 3, 4).

Essential oils are aromatic and volatile liquids whose main components are alcohols, aldehydes, lactones, and phenols.

They are formed in specialized cells within the plant’s stem or leaves and concentrated in its bark, leaves, or fruits. Essential oils have numerous properties, such as spasmolytic, immunomodulatory, psychotropic, expectorative, antibacterial, and antioxidative activities. Due to the broad range of antimicrobial and other effects, plants that produce essential oils have been used for medicinal purposes for over thousands of years. Today, essential oils are being investigated for the treatment of fungal-, bacterial-, and viral-based infectious diseases.

Extensive research efforts have recently been aimed at the development of novel antimicrobial compounds. Among them, antimicrobial peptides, commonly isolated from several organisms, have been considered for use as antimicrobial drugs (Ch. 4, 10). Antimicrobial peptides vary in their amino acid compositions and sizes (ranging from less than 5 to over 100 amino acid residues) and commonly have cationic and amphipathic properties. About 2300 antimicrobial peptides have been reported in the Antimicrobial Peptide Database and more than 100 peptide-based drugs are currently available, while another 500–600 candidates are in pre-clinical testing.

Along with their antimicrobial activity, some antimicrobial peptides have immunomodulatory and antitumoral activities.

Besides their isolation from natural organisms, antimicrobial peptides might be improved or created using computational tools. With these methodologies thousands of novel molecules can be generated, but they require high-through in vitro and in vivo validation, and therefore the parallel development of rapid assays.

The term “enzybiotics” was coined in 2001. It refers specifically to the antibacterial potential of bacteriophage enzy­

BOOK REVIEWS

mes actively produced during the phage lytic cycle and capable of degrading the bacterial cell wall. Recently, it was suggested that the term enzybiotics should refer to all enzymes displaying antibacterial and/or antifungal activity. In the literature, enzybiotics are also called “lytic enzymes” or “peptidoglycan hydrolases”, as bacterial cell-wall peptidoglycans are their major targets.

Phage-based therapies consist of the administration of phage lytic enzymes or/and whole phages (Ch. 8). The recent dramatic increase in antibiotic resistance has stimulated renewed and intense interest in the therapeutic use of phages.

Félix D’Herelle (one of the co-discoverers of bacteriophages) used phage therapy to treat bacterial dysentery in Paris as early as 1918, although for decades thereafter many scientists remained skeptical regarding his apparent success. Today, phages are already used as food preservatives; for example, the FDA-approved utilization of phages in cheese to control Listeria monocytogenes. Phages release their progeny from bacterial cells via two different pathways. Filamentous phages are continuously extruded from bacterial cells, without killing them, whereas non-filamentous phages induce host-cell lysis via highly evolved peptidoglycan hydrolases that quickly destroy the cell wall of the host bacterium.

Antimicrobial compounds is an exceptional review and update of information on alternative therapeutic solutions to the increasing problem of bacterial antibiotic resistance. This book is of great value to investigators working in the field of antimicrobial research and to students of several biomedical disciplines interested in deepening their knowledge of microbiology.

Mercedes Berlanga University of Barcelona mberlanga@ub.edu

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Volume 17(2) JUNE 2014

Acknowledgement of Institutional Subscriptions International Microbiology staunchly supports the policy of open access (Open Access Initiative, see Int Microbiol 7:157161). Thus, the journal recognizes the help received from the many institutions and centers that pay for a subscription—in spite of the possibility to download complete and current issues of the journal free of charge. We would therefore like to thank those entities. Their generous contribution, together with the efforts of the many individuals involved in preparing each issue of International Microbiology, makes publication of the journal possible and plays an important role in improving and expanding the field of microbiology in the world. Some of those institutions and centers are: Area de Microbiología. Departamento de Biología Aplicada.

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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 Biotecnología-CSIC. Cantoblanco (Madrid) / Grupo de Investigación de Bioingeniería y Materiales (BIOMAT). 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 MassachusettsAmherst. 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 17 · Number 2 · June 2014 RESEARCH REVIEW

Romero D, Kolter R Functional amyloids in bacteria

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RESEARCH ARTICLES

López-García MT, Rioseras B, Yagüe P, Álvarez JR, Manteca A Cell immobilization of Streptomyces coelicolor: effect on differentiation and actinorhodin production

Carrasco P, Pérez-Cobas AE, van de Pol C, Baixeras J, Moya A, Latorre A Succession of the gut microbiota in the cockroach Blattella germanica Castro N, Toranzo AE, Magariños B A multiplex PCR for the simultaneous detection of Tenacibaculum maritimum and Edwardsiella tarda in aquaculture

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Bonaterra A, Badosa E, Rezzonico F, Duffy B, Montesinos E Phenotypic comparison of clinical and plantbeneficial strains of Pantoea agglomerans

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Becerra A, Rivas M, García-Ferris C, Lazcano A, Peretó J A phylogenetic approach to the early evolution of autotrophy: the case of the reverse TCA and the reductive acetyl-CoA pathways

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PERSPECTIVES

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