International Microbiology

Publication Board

Editorial Board

Editor-in-Chief José Berenguer (Madrid), Autonomous University of Madrid

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

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

Addresses Editorial Office International Microbiology C/ Poblet, 15 08028 Barcelona, Spain Tel. & Fax +34-933341079 E-mail: int.microbiol@microbios.org www.im.microbios.org Spanish Society for Microbiology C/ Rodríguez San Pedro, 2 #210 28015 Madrid, Spain Tel. +34-915613381. Fax +34-915613299 E-mail: secretaria.sem@microbiologia.org www.semicrobiologia.org Institute for Catalan Studies C/ Carme, 47 08001 Barcelona, Spain Tel. +34-932701620. Fax +34-932701180 E-mail: int.microbiol@microbios.org © 2017 Spanish Society for Microbiology, Madrid, & Institute for Catalan Studies, Barcelona. Printed in Spain ISSN (print): 1139-6709 e-ISSN (electronic): 1618-1095 D.L.: B.23341-2004

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

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CONTENTS International Microbiology (2017) 20:1-54 ISSN (print): 1139-6709. e-ISSN: 1618-1095 www.im.microbios.org

Volume 20, Number 1, March 2017

RESEARCH REVIEW

Labella AM, Arahal DR, Castro D, Lemos ML, Borrego JJ Revisiting the genus Photobacterium: taxonomy, ecology and pathogenesis

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RESEARCH ARTICLES

Köchling T, Sanz JL, Galdino L, Florencio L, Kato MT Impact of pollution on the microbial diversity of a tropical river in an urbanized region of northeastern Brazil

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Karaduman A, Ozaslan M, Kilic IH, Bayil-Oguzkan S, Kurt BS, Erdogan N Identification by using MALDI-TOF mass spectrometry of lactic acid bacteria isolated from non-commercial yogurts in southern Anatolia, Turkey

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Núñez-Díaz JA, Balebona MC, Alcaide EM, Zorrilla I, Moriñigo MA Insights into the fecal microbiota of captive Iberian lynx (Lynx pardinus)

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Regmi S, Choi YS, Choi YH, Kim YK, Cho SS, Yoo JC, Suh J-W Antimicrobial peptide from Bacillus subtilis CSB138: characterization, killing kinetics, and synergistic potency

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PIONEERS IN MICROBIOLOGY: Paulina Beregoff (1902–1989), Colombia

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Journal Citations Reports 5-year Impact Factor of International Microbiology is 2,17.

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. Transmission electron micrograph of the marine phage H1 negatively stained. Phage was isolated from a strain of Pseudoalteromonas sp. from the Blanes Bay Microbial Observatory station (BBMO), a surface coastal site in the NW Mediterranean. Photo by Elena Lara, Marine Sciences Institute (ICM-CSIC) (Magnification, 330,000×)

Center. Landscape of the Doñana National Park (Huelva, Spain), one of the Iberian territories where the population of the Iberian lynx (Lynx pardinus) have been able to historically survive and from where this species seems to initiate its expansion to a wider range of the Iberian Peninsula geography. [See article by Núñez-Díaz et al. pp 31-41, this issue, on the fecal microbioma of the captive lynx.]

Lower right. Transmission electron micrograph of the plasmolized yeasts (“lias”) of Saccharomyces cerevisiae from the elaboration of sparkling wines according to the “cava” method after second fermentation in closed bottles. Photo

by Montserrat Riu and Rebeca Tudela, Faculty of Pharmacy and Food Sciences, University of Barcelona. (Magnification, 10,000×)

Upper right. Transmission electron micrograph of Sphingobacterium detergens during the process of cellular division. The bacterium was isolated from a soil sample from the Azorean Islands and was selected for its ability to reduce the surface tension of the culture medium. Photo by Ana M. Marqués and César Burgos-Díaz, Faculty of Pharmacy and Food Sciences, University of Barcelona. (Magnification, 10,000×) Lower left. Dark field micrograph of several individuals of the ciliate Vorticella sp. detached from its peduncles. Note the big and active macronuleus with the shape of a long and bluish band. Photo by Rubén Duro, Center for Microbiological Research, Barcelona. (Mag­­ni­fication, 1000×)

Back cover: Pioneers in Microbiology Paulina Beregoff (1902–1989), Colombia Paulina Beregoff was the first woman to obtain a degree in medicine in Colombia.

She was born in 1902 in Kiev—by then a city of the Russian Empire—, in an aristocratic family of Jewish descent. Due to the political situation in her country, she was educated in the United States, where, in 1921, she graduated in Bacteriology and Parasitology and Pharmacy and Chemistry at the University of Pennsylvania. She started working at the laboratory of Pathology of that university and became a member of the Rivas Bacteriological Society of the University of Pennsylvania. In 1922, the Dean of the School of Medicine of the University of Cartagena, Colombia, asked the University of Pennsylvania for an expert in tropical diseases, including yellow fever. This disease was a great concern in Cartagena due to the high mortality rates it caused and because of the implications on the image of the city, which was a major commercial and harbor center. The University needed a qualified advisor that could also train local physicians, and the University of Pennsylvania chose Beregoff for that task. Once in Cartagena, she had to identify an epidemic outbreak that had been causing many fatalities, mostly among indigenous peoples living in the Magdalena River shores. Colombian phys­ icians were not familiar with symptoms and causal agents of diseases such as yellow fever, typhoid fever and malaria, but thought that the epidemic outbreak could be due to one of them. Beregoff sent samples of cultures

from corpses of people killed by the disease to be analyzed at the University of Pennsylvania. The disease turned out to be fiebre tifomalárica and not simply malaria, as they first had considered. Beregoff thought that the infection depended mostly on the deficiencies or resistance of the immune system and proposed that physicians should work to prevent the disease. Once she had achieved her task, she intended to go back to Philadelphia to study medicine at Temple University, but she was asked to remain in Cartagena, where she could also study medicine. In 1922 she enrolled at the University of Cartagena under special conditions. Due to her previous studies and qualification, she could be waived the first two years of the studies of medicine. She set up the first laboratories of bacteriology and parasitology in Cartagena, with microscopes and other equipment donated by the University of Pennsylvania.

Her thesis director recognized her great contribution, she having been able to differentiate the various species of Laveran’s haematozoa, to observe the treponema causing yaws, to find the Piroplasma Donovani, the parasite of KalaAzar (visceral leishmaniasis) in the blood, and having been the first to isolate the “typhoid bacillus”, confirming thus the presence of typhoid fever in town.

She could also to properly perform the Wassermann technique on syphilis.

The fact that she was a foreign woman and the she had had some privileges in her medicine studies was criticized by some people. In 1933 she married bacteriologist Arthur Stanley Gillow and they moved to Canada. Since then she signed her publications as Pauline Beregoff-Gillow. After her husband’s death, in 1964, she returned to Colombia and dedicated his husband’s legacy to set up a foundation under his name that should work on preventive medicine. She died on September 20, 1989 and left her fortune to the foundation.

Front cover and back cover design by MBerlanga & RGuerrero

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RESEARCH REVIEW International Microbiology 20(1): 1-10 (2017) doi:10.2436/20.1501.01.280. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Revisiting the genus Photobacterium: taxonomy, ecology and pathogenesis Alejandro M. Labella,1 David R. Arahal,2 Dolores Castro,1 Manuel L. Lemos,3 Juan J. Borrego1* Universidad de Málaga, Andalucía Tech, Departamento de Microbiología, Campus de Teatinos, Málaga, Spain.

Departamento de Microbiología y Ecología, Universitat de Valencia, Burjassot, Spain. 3Departamento de Microbiología y Parasitología, Instituto de Acuicultura, Universidade de Santiago de Compostela, Santiago de Compostela, Spain 1

2

Received 1 March 2017 · Accepted 30 March 2017 Summary. The genus Photobacterium, one of the eight genera included in the family Vibrionaceae, contains 27 species with valid names and it has received attention because of the bioluminescence and pathogenesis mechanisms that some of its species exhibit. However, the taxonomy and phylogeny of this genus are not completely elucidated; for example, P. logei and P. fischeri are now considered members of the genus Aliivibrio, and previously were included in the genus Vibrio. In addition, P. damselae subsp. piscicida was formed as a new combination for former Vibrio damsela and Pasteurella piscicida.

Moreover, P. damselae subsp. damselae is an earlier heterotypic synonym of P. histaminum. To avoid these incovenences draft and complete genomic sequences of members of Photobacterium are increasingly becoming available and their use is now routine for many research laboratories to address diverse goals: species delineation with overall genomic indexes, phylogenetic analyses, comparative genomics, and phenotypic inference. The habitats and isolation source of the Photobacterium species include seawater, sea sediments, saline lake waters, and a variety of marine organisms with which the photobacteria establish different relationships, from symbiosis to pathogenic interactions. Several species of this genus contain bioluminescent strains in symbiosis with marine fish and cephalopods; in addition, other species enhance its growth at pressures above 1 atmosphere, by means of several high-pressure adaptation mechanisms and for this, they may be considered as piezophilic (former barophilic) bacteria. Until now, only P. jeanii, P. rosenbergii, P. sanctipauli, and the two subspecies of P. damselae have been reported as responsible agents of several pathologies on animal hosts, such as corals, sponges, fish and homeothermic animals. In this review we have revised and updated the taxonomy, ecology and pathogenicity of several members of this genus. [Int Microbiol 20(1): 1-10 (2017)] Keywords: Photobacterium · taxonomy · symbiosis · pathogenesis · virulence factors

Taxonomic and phylogenetic perspectives The genus Photobacterium has a long standing in micro­ biology, having received attention for more than one century *

Corresponding author: Juan J. Borrego E-mail: jjborrego@uma.es

by the bioluminescence that some of its species exhibit. Indeed, etymologically it means light producing bacterium. To date, it contains 27 species with valid names (Table 1). The historical development of the taxonomy of this genus is relatively easy to follow. The type species, Photobacterium phosphoreum, was included in the Approved Lists of Bacterial Names [79] together with P. angustum, P. (Aliivibrio) fischeri and P. leiognathi. The only species described in the

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Int. Microbiol. Vol. 20, 2017

following decade, P. logei, is now considered a member of the genus Aliivibrio and so is P. fischeri [64]. In turn, P. damselae [80] was formed as a new combination for former Vibrio damsela and Pasteurella piscicida. This species is the only one for which subspecies have been proposed so far with the publication of P. damselae subsp. piscicida [27].

Moreover, P. damselae subsp. damselae is an earlier heterotypic synonym of P. histaminum [37,60]. In the last two decades the pace of descriptions has intensified with the proposal of 20 novel species and two new combinations with valid names (Table 1), which gives an average of about two new species names per year. According to minute 17 of the Subcommittee meetings on the taxonomy of Aeromonadaceae, Vibrionaceae and related organisms held in Istanbul, Turkey, in 2008 [31], the type strain of P. aplysiae is not available and a neotype strain has not been proposed to date.

At the time of validation [79], the description of the genus was the one given in the 8th edition of Bergey’s Manual. Although an emendation has never been formally proposed it has been revised and updated recently [84].

Phylogeny. Photobacterium is one of the nine genera contained in the family Vibrionaceae (order “Vibrionales”, class Gammaproteobacteria). It is also the largest one after the type genus Vibrio. Following a practice that is common and more developed for Vibrio spp. [64,73], several authors have established different clades within the genus Photobacterium [6,73,86]. Clades are usually named after the older species name, referring to its validation date, regardless of the position of that strain into the clade. Currently, four clades have been described in the genus Photobacterium: Damselae (P. damselae subsp. damselae and P. damselae subsp. piscicida), Phosphoreum (P. angustum, P. aquimaris, P. iliopiscarium, P. kishitanii, P. leiognathi, P. phosphoreum and P. piscicola), Profundum (P. aestuarii, P. aplysiae, P. frigidiphilum, P. indicum, P. lipolyticum, P. profundum, P. sanguinicancri and P. swingsii), and Rosenbergii (P. aphoticum, P. ganghwense, P. halotolerans, P. jeanii, P. lutimaris, P. marinum, P. rosenbergii). But the clustering of P. aquae, P. gaetbulicola, P. galatheae, P. panuliri, and P. sanctipauli has not been elucidated yet. It has to be noted that this classification into clades has no standard in nomenclature although it can make more amenable the study of large genera by grouping together lines of descents. However, this achievement requires the application of robust molecular approaches and large sets of strains (not

LABELLA ET AL.

just type strains). A comprehensive study meeting both requisites is still pending to the best of our knowledge but at least it is optimistic to see that most recent species descriptions include phylogenetic analysis using alternative genes [48,56,71,83] or MLSA schemes [7,10,26,29,45,50,92]. This means that at least for some genes there are sequences available in public repositories for most (ideally all) the type strains and even for a number of additional isolates of some of them.

The genes more frequently employed to perform phylogenetic studies within the genus Photobacterium are recA (protein RecA, recombinase A), rpoA (RNA polymerase α subunit), gyrB (DNA gyrase subunit B), pyrH (uridylate kinase, uridine monophosphate kinase), gapA (glyceraldehyde 3-phosphate dehydrogenase, NAD-dependent glyceraldehyde-3-phosphate dehydrogenase), ftsZ (cell division protein FstZ), topA (DNA topoisomerase I), and mreB (rod shapedetermining protein MreB).

At the same time, draft and complete genomic sequences of members of the genus Photobacterium are increasingly becoming available and their use is becoming routine for many research laboratories to address diverse goals: species delineation with overall genomic indexes, phylogenetic analyses, comparative genomics, and phenotypic inference [2,29,50].

At the time of writing the present review (1 March, 2017), a search at the Assemblies database in NCBI [http://www.ncbi. nlm.nih.gov/] shows that there are 67 results for Photobacterium, 15 of which are from strains flagged as type material. A more careful examination reveals that two of these can be considered redundant entries (they are from two equivalent designations of the same strain, the type strain of P. damselae, sequenced in different laboratories) and another one is from “P. marinum” that has not been validated to date. Since there are 27 species in the genus, the resulting 13 genomic sequences represent about half of them. Thus the gap to be filled to give full coverage to the type strains of the genus in terms of availability of their genomic sequences is not too large and we can anticipate it might be reached soon. Although most of these genomic sequences are assembled into contigs or scaffolds, there are two completed, P. gaetbulicola Gung47T and P. profundum SS9.

One advantage of having large data sets of genomes is that they can be explored to search for the most suitable single gene phylogenetic marker. This objective has been addressed at the family level by Machado and Gram [49] who concluded that the fur (ferric uptake regulator Fur) gene was suitable for

Int. Microbiol. Vol. 20, 2017

GENUS PHOTOBACTERIUM

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Table 1. Species, habitats and geographic sources of Photobacterium species Species

Habitats

Geographic sources

P. aestuarii

Tidal flat sediment

Yeongam Bay (R. Korea)

[46]

P. angustum

Seawater

North Pacific Ocean (20º30’N 157º30’E)

[79]

P. aphoticum

Seawater

Malvarrosa beach, Valencia (Spain)

[48]

P. aplysiae

Eggs of sea hare (Aplysia kurodai)

Mogiyeo (R. Korea)

[75]

P. aquae

Malabar grouper (Epinephelus malabaricus) in mariculture system

Tianjin (China)

[45]

P. aquimaris

Seawater

Sagami Bay (Japan)

[92]

P. damselae

Damselfish (Chromis punctipinnis) skin ulcer

California (USA)

P. frigidiphilum

Deep-sea sediments (1450 m)

Edison Seamount (western Pacific Ocean)

[74]

P. gaetbulicola

Tidal flat

Gungharbour (R. Korea)

[36]

P. galatheae

Mussel

Solomon Sea (Solomon Islands)

[50]

P. ganghwense

Seawater

Ganghwa Island (R. Korea)

[63]

P. halotolerans

Water from a subterranean saline lake

Lake Martel, Mallorca (Spain)

[71]

P. iliopiscarium

Intestines of fish (herring, coal fish, cod and salmon) living in cold seawater

Norway

[84]

P. indicum

Marine mud (400 m depth)

Indian Ocean

[32]

P. jeanii

Healthy corals (Palythoa caribaeorum, Phyllogorgia dilatata and Merulina ampliata)

Brazil and Australia

[10]

P. kishitanii

Light organs and skin of several marine fish species

Japan, Cape Verde, Hawaii, Florida, South Africa

[7]

P. leiognathi

Light organ of teleostean fish (Leiognathus)

Gulf of Thailand (Thailand)

[66]

P. lipolyticum

Intertidal sediment

Yellow Sea (R. Korea)

[91]

P. lutimaris

Tidal flat sediment

Saemankum (R. Korea)

[33]

P. panuliri

Eggs of spiny lobster (Panulirus penicillatus)

Andaman Sea (India)

[13]

P. phosphoreum

Skin of marine animals, intestines of marine fish, luminous organs, seawater

Hawaii (USA), Japan and other locations

[79]

P. piscicola

Skin and intestine of marine fish, spoiled packed cod

North Sea (Holland), Denmark, Aberdeen Bay (UK)

[26]

P. profundum

Deep-sea sediment (5110 m)

RyukyuTrench (24º15.23’N 126º47.30’E)

[58]

P. rosenbergii

Tissue and water extracts of coral species

Magnetic Island (Australia)

[83]

P. sanctipauli

Coral (Madracis decactis)

St. Peter & St. Paul Archipelago (Brazil)

[56]

P. sanguinicancri

Crab (Maja brachydactyla) haemolymph, mussels (Mytilus edulis)

Spain, Netherlands

[29]

P. swingsii

Pacific oysters (Crassostrea gigas), crab (Maja brachydactyla) haemolymph

Mexico, Spain

[28]

a

Reference

[47,80]

Additional strains are reported in Smith et al. [80] from human puncture wound, diseased shark, diseased turtle, diseased fish, aquarium seawater and fish surface.

that purpose and even developed a PCR method to be used for the amplification and sequencing of the gene. Phylogenetic analysis can also be a method to elucidate horizontal gene transfer as it was performed in the study by Urbanczyk et al.

[85], who assessed the incidence of interspecies transfer of the lux genes (luxCDABEG), which encode proteins involved in luminescence and concluded that horizontal transfer of the lux genes in nature is rare and that horizontal acquisition of the lux genes apparently has not contributed to speciation in recipient taxa.

Ecology of Photobacterium The members of the genus Photobacterium thrive worldwide in oceans and show substantial ecophysiological diversity including free-living, symbiotic, piezophilic, and parasitic life styles. The habitats and isolation source of these species include seawater, sea sediments, saline lake waters, and a variety of marine organisms with which the photobacteria establish different relationships, from symbiotic ones, such as commensalism or mutualism, to pathogenic interactions.

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Int. Microbiol. Vol. 20, 2017

Generally, in the marine environment (seawater and sediment), the species of Photobacterium are free-life forms, but they may colonize several animal surfaces developing neutral or negative relationships with the host. These nonspecific or pathogenic associations contrast with the highly specific, mutually beneficial association of certain Photobacterium species in bioluminescent symbiosis with aquatic animals [17].

There is not a clear discrimination between the Photobacterium species regarding to their relationship with the isolation source or habitat (Table 1). Thus, most of the nonluminous photobacteria (lack of lux operon genes) have been isolated from marine waters or sediments, but several strains of these species have been described in association with diseased or healthy corals, zoanthids, sea hares, mollusks, crabs and fish [28,29,39,45,69,75,83]. Nevertheless, strains of luminous Photobacterium species harbouring genes for luminescence (lux CDABEG) [19], such as P. kishitanii, P. leiognathi, P. phosphoreum and P. piscicola, have also been isolated from squids, corals and fish [6,26,34]. Therefore, the luminescence production property is not a key ability of this bacterial group to the specific colonization of none habitats, excepting the light-organs of squids and fish.

Photobacteria as symbiotic of light-organs.

Several species of this genus contain bioluminescent strains including P. angustum, P. aquimaris, P. damselae, P. ganghwense, P. kishitanii, P. leiognathi, P. phosphoreum, and P. piscicola. From them, P. kishitanii and P. leiognathi establish bioluminescent symbiosis with marine fish, squid and octopus [57]. These associations are typically highly specific at the animal family-bacterial species level; P. leiognathi with families Leiognathidae, Acropomatidae and Apogonidae (Perciformes), and Moridae (Gadiformes) [21,34,82,88]; and P. kishitanii with the fish families Chloropthslmidae (Acilopiformes), Macrouridae, Sleindachneriidae and Moridae (Gadiformes), Trachichthyidae (Beryciformes), Opisthoprectidae (Gemeriformes) and Acropomatidae (Perciformes) [6,20].

The animals accumulate dense populations of luminous bacteria in gland-like tissue complex called light organs [24], providing them with nutrients and oxygen for their growth and light production. The bacterial light in symbiotic animals is associated with sex-specific signalling, predator avoidance, locating or attracting prey, to name a few [82,86]. Symbiotic luminous bacteria have not an obligatory dependency of the host for their reproduction [23], but it seems that exist certain specificity between the symbiotic fish and the luminous Pho-

LABELLA ET AL.

tobacterium species. The animals that establish a relationship with P. leiognathi as light-organ symbionts tend to be found in shallower waters, whereas the fish that are symbiotic with P. kishitanii are usually found in deeper waters [23,34]. This apparent specificity, which presumably would have a genetic basis, is believed to result from the host animal selecting its species of symbiotic bacteria and preventing that other bacteria could colonize its light organs. Several authors have proposed that the bioluminescent symbiosis might involve coevolutionary interactions [21,86], due to the animal dependence of the bacterial light, its specialized anatomical adaptations for harbouring bacteria, and the host family-bacterial species specificity.

Although bioluminescent associations appear to be highly specific, in some cases two Photobacterium species may be present within individual light organs of fish [23,34], representing a phenomenon named cosymbiosis. Furthermore, different species of the same fish family sometimes harbour different Photobacterium species or even bacteria belonging to other bacterial genera, like Aliivibrio or Vibrio [23,24]. In addition, distinct strains of a single species may be present with individual light organs of both adult and larval fish [22,24].

This species- and strain-level variation demonstrates the lack of strict specificity in bioluminescent symbiosis.

Bioluminescent symbioses of fish and squid with luminous bacteria apparently do not exhibit codivergence (cospeciation), since phylogenies for host and their symbiotic bacteria present no meaningful topological congruence [23,34]. The patterns of symbiont-host affiliation in bioluminescent symbioses observed from nature therefore are not likely to have arisen through coevolutionary interactions.

However, the absence of nonluminous bacteria in light-organs of fish and squid indicates that some kind of selection must be operative, like the environmental congruence [30].

The congruence between the environmental distribution of a predominant species of luminous bacteria and the fish developmental stage at which its light-organ is receptive to bacterial colonization, determines which bacterial species and strains establish the symbiosis [23,34]. Some environmental factors, such as the temperature, influence the abundance of the different species of luminous bacteria in the marine environment. Thus, lower temperatures found in deeper waters favour the prevalence of psychrotropic species like P. kishitanii, which is the main light-organ symbiont in these waters. On the contrary, warmer waters favour the growth of mesophilic Photobacterium species, such as P. leiognathi

GENUS PHOTOBACTERIUM

being fish larvae in these waters more receptive to acquire these bacteria as light-organ symbionts.

In short, bioluminescent symbioses, therefore, differ from endosymbiotic associations, which are mutually obligate relationships in which the symbiotic bacteria are housed intracellularly and are transferred maternally.

Symbiotic luminous bacteria are housed extracellularly, and in most cases they are known not be obligately dependent on the host for their reproduction. Unlike obligate intracellular bacteria, the symbiotic luminous bacteria colonize a variety of other marine habitats, including intestinal tracts, skin, and body fluids of marine animals, sediments, and seawater, where they coexist and compete with many other kinds of microorganisms. A second major difference with endosymbiotic associations is that symbiotic luminous bacteria are acquired from the environment with each new generation of the host instead of being transferred vertically through the maternal inheritance mechanisms. Another major difference between bioluminescent symbiosis and endosymbiosis is that luminous bacteria and their host animals show no evidence of cospeciation. Endosymbiosis is generally assumed to involve coevolutionary interactions, that is, reciprocal genetic changes in host and symbiont that result from the obligate and mutual dependence of each partner on the other. Detailed molecular phylogenies of bacterially luminous fish and squids, however, are very different from the phylogenies of their symbiotic light-organ bacteria [18].

This lack of host-symbiont phylogenetic congruence demonstrates that the evolutionary divergence of symbiotic luminous bacteria has occurred independently of the evolutionary divergence of their host animals.

Bioluminescent symbioses appear to represent a paradigm of symbiosis that differs fundamentally from associations involving obligate, intracellularly transferred symbionts. While fish and squids are dependent ecologically on luminous bacteria, the bacteria are not obligately dependent on their bioluminescent hosts. The evolutionary adaptations for bioluminescent symbiosis, for example presence of light organs, accessory tissues for controlling, diffusing, and shaping the emission of light, and behaviour associated with light emission, all are borne by the animal. No genetic adaptations have been identified in the bacteria that are necessary for and specific to their existence in light organs compared to the other habitats they colonize. Therefore, luminous bacteria seem to be opportunistic colonizers, able to persist in animal light-

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organs as well as in a variety of other habitats to which they are adapted.

Other question unanswered is regarding to the benefit of luminescence for the non-symbiotic photobacteria. This question has not been elucidated fully, but several explanations have been arisen. One of the most commonly cited explanations is that the bioluminescence increases the propagation and dispersal of bacteria by attracting fish or other marine animals to consume luminous material. This hypothesis based mostly on the prevalence of luminous bacteria in fish gut has not been demonstrated experimentally. Nevertheless, Zarubin et al. [93] established that zooplankton that contacts and feeds on P. leiognathi starts to glow, and the glowing individuals are highly vulnerable to predation by nocturnal fish. Glowing photobacteria are transferred to the intestines of fish and zooplankton, when they survive digestion and gain effective means for growth and dispersal. The use of bioluminescence, therefore, appears to be highly beneficial for marine bacteria, especially in oligotrophic areas of the deep sea.

Deep-sea sediments as habitats of Photobacterium species. Members of the genus Photobacterium are common inhabitants of marine waters sediments, including P. aestuarii, “P. atrarenae”, P. frigidiphilum, P. gaetbulicola, P. indicum, P. lipolyticum, P. lutimaris, “P. marinum”, P. phosphoreum, and P. profundum. From them, P. frigidiphilum, P. phosphoreum, and P. profundum may be considered as piezophilic (former barophilic) bacteria, because these species enhance its growth at pressures above 1 atmosphere, by mean of several high-pressure adaptation mechanisms [9,74].

The adaptative traits include those related to growth, macromolecules and storage lipids, membrane and soluble proteins, the respiratory-chain compounds, replication, transcription and traslation [9,54,90]. These species are the only ones known to produce a long-chain polyunsaturated fatty acid (PUFA), the eicosapentaenoic acid (EPA) [58]. Recently, Le Bihan et al. [42] analysed the proteome of P. profundum under different pressure regimes, and obtained altered modes of protein function in that conditions. The authors identified differentially expressed proteins involved in high pressure adaptation; thus, proteins belonging to the glycolysis/gluconeogenesis pathway were up-regulated at high pressure, whilts several proteins involved in the oxidative phosphorylation pathway were up-regulated at atmospheric pressure. In addition, the expression of some proteins involved in nutrient transport or assimilation was also directly regulated by pressure.

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Pathogenesis of Photobacterium Some species of this genus, including P. rosenbergii, P. jeanii, P. sanctipauli, and the two subspecies of P. damselae, have been reported to produce several pathologies on animal hosts, such as corals, sponges, fish, and homeothermic animals [10,56,69,83]. Unfortunately, little is known on the pathogenesis mechanisms of P. rosenbergii and P. sanctipauli that cause the coral bleaching and further dead of the corals [83]; however, both P. damselae subspecies have received a great attention as emerging pathogens for many aquatic organisms, including fish, mollusks and crustaceans, and even for humans [41,55,69,72,89].

Photobacterium damselae subsp. damselae (Pdd) is a normal inhabitant of seawater, marine sediments, seaweeds and marine animals [41,76], and prefers warm water conditions (20–30 °C). This microorganism is considered a primary pathogen of several species of wild- and cultured-fish causing wound infections and hemorrhagic septicaemia. It is also an opportunistic human pathogen, causing necrotizing fasciitis [69]. The other subspecies, P. damselae subsp. piscicida (Pdp), is the causal agent of fish photobacteriosis, a serious bacterial disease affecting different economically important cultured marine fish species [72].

Virulence factors of P. damselae. The main bacterial iron-uptake systems include the production of iron-sequestering compounds named siderophores as well as the use of heme group as iron source. Siderophores are chemically diverse low-molecular-weight iron chelators that can effectively solubilize iron or remove it from other chelators and transport it into the cell through the corresponding membrane receptor proteins [43]. Some bacteria not only produce their own siderophores, but also express receptors capable of transport xenosiderophores produced by other organisms [11]. Pdp and Pdd are able to acquire iron from hemin and hemoglobin as unique iron sources in vitro [43]. Their heme uptake systems are encoded by a gene cluster formed by 10 genes [67].

This heme uptake system includes a TonB-dependent outer membrane receptor to transport the heme group into the periplasm, a periplasmic binding protein, and an ATP-binding cassette (ABC) to drive heme across the cytoplasmic membrane [4,67]. It is also known that in Pdp, the acquisition of iron from its host is efficiently achieved by means of the synthesis of the siderophore piscibactin [81], and its transport into the cell through the outer membrane receptor FrpA [62].

LABELLA ET AL.

The synthesis and transport are encoded by a pathogenicity island, which is part of the transmissible plasmid pPHDP70.

It has been demonstrated that this plasmid greatly contributes to the virulence of Pdp for fish, and that it can be horizontally transmitted to other marine bacteria [62]. It has also been reported that Pdd expresses several high-molecular-weight outer membrane proteins under iron limitation conditions [69], and that some strains likely produce the siderophore vibrioferrin [65], although other virulent strains lack this system, being its contribution to virulence yet uncertain. The presence of these or other iron uptake mechanisms in other species of Photobacterium is unknown, although some of the iron-uptake related genes reported in both P. damselae subspecies are present in other species genomes. The role of these mechanisms in non-pathogenic species is uncertain.

Bacterial extracellular products (ECP) containing phospholipase, cytotoxic, and hemolytic activities may account for the damage to infected cells, the consequent release of the microorganisms, and the invasion of adjacent cells [25]. ECP of P. damselae strains were shown to be lethal for different fish species and for fish and homeothermic cell lines [40]. Recently, Vences et al. [87] have demonstrated that phospholipase and collagenase activities contributed to virulence of Pdd. It is well known the existence of a close relationship between the ability of a microorganism to provoke diseases and the production of bacterial toxins. In the case of Pdd, several heat-labile cytolytic toxins have been reported, one of them named damselysin (Dly), a phospholipase-D active against sphingomyelin, presented strong hemolytic activity [38]. It has also been demonstrated that presence of gene dly is not a pre-requisite for the hemolytic activity and for the pathogenicity of Pdd, since dly-negative strains possess virulence potential for animals, and also show toxicity for homeotherm and poikilotherm cell lines [40,61]. Rivas et al. [68] identified and characterized a 150 kb plasmid, pPHDD1, which contains the genes for both Dly and HlyApl, being the lastest a small pore-forming toxin (PFT) with hemolysin activity, named phobalysin [70]. The mutation of both dly and hlyApl genes in a pPHDD1-harbouring strain renders the strain non-virulent for fish, and only slightly virulent for mice, and the hemolytic phenotype on sheep blood agar of a dly and hlyApl double mutant resembles that of naturally plasmidless strains [68,69]. Thus, pPHDD1-harbouring isolates of Pdd produce three different hemolysins, each of them individually prove to be sufficient to cause death in mice. Each hemolysin contributes to virulence in a different degree, although only

GENUS PHOTOBACTERIUM

the Dly-producing strains caused death in fish, demonstrating the importance of the plasmid for the virulence of this bacterium for fish. Despite the importance of pPHDD1, many Pdd virulent strains are plasmidless. The hemolytic activity exhibited by these strains is due to hemolysin PhlyC, encoded by the chromosome-harbored hlyAch gene [69,87], which contributes to virulence for fish [87].

In Pdp a key pathogenicity factor is an exotoxin, a plasmid-encoded apoptosis-inducing protein of 56 kDa (AIP56), responsible for apoptogenic activity against fish macrophages and neutrophils [16]. The AIP56 toxin is a zinc metalloprotease involved in binding and internalization into the cytosol of target cells [77], and acts inducing the activation of caspases 8, 9 and 3, the loss of mitochondrial membrane potential, the release of cytochrome c into the cytosol, and the overproduction of ROS, which suggest that the exotoxin activates both extrinsic and intrinsic apoptotic pathways [12]. Through the activation of the cell death process involving macrophages and neutrophils, the pathogen is able to subvert the immune defenses of the host and to produce infectious disease.

Little is known on the adherent properties to cells of Pdd, although Khouadja et al. [35] established that this subspecies possess the ability to adhere to fish mucus. On the contrary, Pdp is adherent mainly for fish cells [53], and the adherence is heat-sensitive, but it is not affected by proteases or by treating the bacteria with antisera raised against its LPS [53]. Nevertheless, the precise nature of the mechanism responsible for adherence and interaction with host cell receptors and virulence factors contributing to the invasion of fish nonphagocytic cells is still unknown [1,3].

Pdp is considered weakly to moderately invasive to several poikilothermic cell lines. LĂłpez-Doriga et al. [44] showed that the uptake of Pdp by EPC cells is time and bacterial-concentration dependant. These authors have been suggested that internalization of this microorganism

by EPC cells is receptor-ligand mediated (zipper mechanism). Pdp isolates also show the ability to spread to adjacent cells from initially infected cells, forming plaques of dead cells [53]. Similar to that previously reported for other Gram-negative pathogenic bacteria, invasion by Pdp can be inhibited by cytochalasin D, indicating that actin and microfilament-dependent mechanisms are required for bacterial internalization [53].

Virulent Pdp strains are serum resistant and can grow in fresh fish serum, whereas non-virulent strains are sensitive to serum killing and their growth is totally inhibited in

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fresh serum [3]. The inhibitory effect of the serum on the non-virulent strains, however, is totally lost if the complement is inactivated by heating at 56 °C for 1 h [51]. Serum resistance is also associated with capsule production, since capsulated strains prevent formation of C3 convertase (C3bBb) by failing to bind serum protein B, or by a higher affinity for serum protein H than for B. Therefore, capsulated strains evade more efficiently the bacteriolytic activity of fresh serum [53]. Pdp capsule formation depends on growth conditions; thus, cells grown under ironlimited conditions or old-cultures had a significantly reduced amount of capsular material [14]. Studies that describe the contribution of bacterial capsules to adhesion and invasion of host cells are contradictory [44,52]. The presence of a capsule prevents the opsonization by C3b, and bacteria will not efficiently be engulfed by fish macrophages [5]. Furthermore, the capsule plays an important role in lethality of Pdp to fish, as non-virulent strains induced for capsule expression resulted in a reduction of LD50 values [52].

The ability of Pdp to avoid phagocytosis and thus to cause disease, may be explained by the induction of extensive apoptosis on macrophages and neutrophils present in Pdp-infected foci, resulting in lysis of these leukocytes by post-apoptotic secondary necrosis [15]. There are contradictory results on the interaction of Pdp with phagocytes; whereas intact bacteria within phagocytes have been observed in vivo [59], suggesting that Pdp may survive inside macrophages, other in vitro studies indicate that fish macrophages are able to kill the bacteria by means of activation of the respiratory burst or an iron-SOD activity [5,8,78].

Future perspectives In this review, we have described some aspects of the genus Photobacterium, including taxonomy, phylogeny, ecology and pathological mechanisms. There is still a lack of understanding of several features encoded by Photobacterium genomes, such as the novel genes involved in the adaptation to specific habitats, the study of new metabolic pathways and their involved genes, and other cellular functions and metabolites produced by these microorganisms. Moreover, the ability of several species of this genus to produce polyunsaturated fatty acids, cold-adapted

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enzymes and antimicrobial compounds constitutes new ways of investigation for a potential biotechnological application of these products in the future.

Competing interests: None declared.

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damselae subsp. damselae reveals metabolic adaptations to iron levels changes and novel potential virulence factors. Vet Microbiol. 201:257264 Reichelt JL, Baumann P (1975) Photobacterium mandapamensis Hendrie et al., a later subjective synonym of Photobacterium leiognathi Boisvert et al. Int J Syst EvolMicrobiol 25:208-209 Rio SJ, Osorio CR, Lemos ML (2005) Heme uptake genes in human and fish isolates of Photobacterium damselae: existence of hutA pseudogenes. Arch Microbiol 183:347-358 Rivas AJ, Balado M, Lemos ML, Osorio CR (2011) The Photobacterium damselae subsp. damselae hemolysins damselysin and HlyA are encoded within a new virulence plasmid. Infect Immun 79:4617-4627 Rivas AJ, Lemos ML, Osorio CR (2013) Photobacterium damselae subsp. damselae, a bacterium pathogenic for marine animals and humans.

Front Microbiol 4:283-289 Rivas AJ, von Hoven G, Neukirch C, Meyenburg M, Qin Q, Füser S, Boller K, Lemos ML, Osorio CR, Husmann (2015) Phobalysin, a small ß-pore forming toxin of Photobacterium damselae subsp. damselae. Infect Immun 83:4335-4348 Rivas R, García-Fraile P, Mateos PF, Martínez-Molina E, Velasquez E (2006) Photobacterium halotolerans sp. nov., isolated from Lake Martel in Spain. Int J Syst Evol Microbiol 56:1067-1071 Romalde JL (2002) Photobacterium damselae subsp. piscicida: an integrated view of a bacterial fish pathogen. Int Microbiol 5:3-9 Sawabe T, Ogura Y, Matsumura Y, Feng G, Rohul Amin AKM, Mino S, Nakagawa S, Sawabe T, Kumar R, Fukui Y, Satomi M, Matsushima R, Thompson FL, Gómez-Gil B, Christen R, Maruyama F, Kurokawa K, Hayashi T (2013) Updating the Vibrio clades defined by multilocus sequence phylogeny: Proposal of eight new clades, and the description of Vibrio tritonius sp. nov. Front Microbiol 4:1-14 Seo HJ, Bae SS, Lee JH, Kim SJ (2005a) Photobacterium frigidiphilum sp. nov., a psychrophilic, lipolytic bacterium isolated from deep-sea sediments of Edison Seamount. Int J Syst Evol Microbiol 55:1661-1666 Seo HJ, Bae SS, Yang SH, Lee JH, Kim SJ (2005b) Photobacterium aplysiae sp. nov., a lipolytic marine bacterium isolated from eggs of the sea hare Aplysia kurodai. Int J Syst Evol Microbiol 55:2293-2296 Serracca L, Ercolini C, Rossini I, Battistini R, Giorgi I, Prearo M (2011) Occurrence of both subspecies of Photobacterium damselae in mullets collected in the river Magra (Italy). Can J Microbiol 57:437-440 Silva DS, Pereira LMG, Moreira AR, Ferreira da Silva F, Brito RM, Faria TQ, Zornetta I, Montecucco C, Oliveira P, Azevedo JE, Pereira PJB, Marcelo-Ribeiro S, do Vale A, dos Santos NMS (2013) The apoptogenic toxin AIP56 is a metalloprotease A-B toxin that cleaves NF-kappab P65. PLoS Pathog 9:e1003128 Skarmeta AM, Bandin I, Santos Y, Toranzo AE (1995) In vitro killing of Pasteurella piscicida by fish macrophages. Dis Aquat Org 23:51-57 Skerman VBD, McGowan V, Sneath PHA (1980) Approved lists of bacterial names. Int J Syst Bacteriol 30:225-420 Smith SK, Sutton DC, Fuerst JA, Reichelt JL (1991) Evaluation of the genus Listonella and reassignment of Listonella damsela (Love et al.)

MacDonell and Colwell to the genus Photobacterium as Photobacterium damsela comb. nov. with an emended description. Int J Syst Bacteriol 41:529-534

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81. Souto A, Montaos MA, Rivas AJ, Balado M, Osorio CR, Rodríguez J, Lemos ML, Jiménez C (2012) Structure and biosynthetic assembly of Piscibactin, a siderophore from Photobacterium damselae subsp. piscicida, predicted from genome analysis. Eur J Org Chem 29:5693-5700 82. Sparks JS, Dunlap PV, Smith WL (2005) Evolution and diversification of a sexually dimorphic luminescent system in ponyfish (Teleostei: Leiognathidae), including diagnoses for two new genera. Cladistics 21:305327 83. Thompson FL, Thompson CC, Naser S, Hoste B, Vandemeulebroecke K, Munn C, Bourne D, Swings J (2005) Photobacterium rosenbergii sp. nov. and Enterovibrio coralii sp. nov., vibrios associated with coral bleaching. Int J Syst Evol Microbiol 55:913-917 84. Thyssen A, Ollevier F (2015) Photobacterium. In: Bergey's Manual of Systematics of Archaea and Bacteria, pp. 1-11. Published on line. DOI: 10.1002/9781118960608.gbm01076 85. Urbanczyk H, Ast JC, Kaeding AJ, Oliver JD, Dunlap PV (2008) Phylogenetic analysis of the incidence of lux gene horizontal transfer in Vibrionaceae. J Bacteriol 190:3494-3504 86. Urbanczyk H, Ast JC, Dunlap PV (2011) Phylogeny, genomics, and simbiosis of Photobacterium. FEMS Microbiol Rev 35:324-342 87. Vences A, Rivas AJ, Lemos ML, Husmann M, Osorio CR (2017) Chromosome-encoded hemolysin, phospholipase and collagenase, contribute to virulence for fish in plasmidless isolates of Photobacterium damselae subsp. damselae. Appl Environ Microbiol 83:e00401-17 88. Wada M, Kamiya A, Uchiyama N, Yoshizawa S, Kita-Tsukamoto K, Ikejima K, Yu R, Imada C, Karatani H, Mizuno N, Suzuki Y, Nishida M, Kogure K (2006) LuxA gene of light organ symbionts of the bioluminescent fish Acropoma japonicum (Acropomatidae) and Siphamia versicolor (Apogonidae) forms a lineage closely related to that of Photobacterium leiognathi ssp. mandapamensis. FEMS Microbiol Lett 260:186-192 89. Yamane K, Asato J, Kawade N, Takahashi H, Kimura B, Arakawa Y (2004) Two cases of fatal necrotizing fasciitis caused by Photobacterium damsela in Japan. J Clin Microbiol 42:1370-1372 90. Yayanos AA (1995) Microbiology to 10,500 meters in the deep-sea. Annu Rev Microbiol 49:777-805 91. Yoon JH, Lee JK, Kim YO, Oh TK (2005) Photobacterium lipolyticum sp. nov., a bacterium with lipolytic activity isolated from the Yellow Sea in Korea. Int J Syst Evol Microbiol 55:335-339 92. Yoshizawa S, Wada M, Kita-Tsukamoto K, Yokota A, Kogure K (2009) Photobacterium aquimaris sp. nov., a luminous marine bacterium isolated from seawater. Int J Syst Evol Microbiol 59:1438-1442 93. Zarubin M, Belkin S, Ionescu M, Genin A (2012) Bacterial luminescence as a lure for marine zooplankton and fish. Proc Natl Acad Sci USA 109:853-857

RESEARCH ARTICLE International Microbiology 20(1): 11-24 (2017) doi:10.2436/20.1501.01.281. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Impact of pollution on the microbial diversity of a tropical river in an urbanized region of northeastern Brazil Thorsten Köchling,1* José Luis Sanz,2 Luiz Galdino,1 Lourdinha Florencio,1 Mario T. Kato1 Laboratory of Environmental Sanitation, Department of Civil Engineering, Federal University of Pernambuco, Recife PE, Brazil, 2Department of Molecular Biology, Autonomous University of Madrid, Cantoblanco, Spain

1

Received 25 January 2017 · Accepted 15 March 2017

Summary. Rivers are important ecosystems that are integrated into biogeochemical cycles and constitute an essential resource for numerous human uses. However, the assessment of the biological diversity and composition of microbial communities found in rivers remains incomplete, partly due to methodological constraints which are only recently being resolved with the advent of next generation sequencing (NGS) techniques. Using 454-pyrosequencing of the 16S gene, the present study analyzed the microbial diversity of the planktonic and sediment populations in a tropical river in northeastern Brazil that is exposed to severe pollution. Six water and six sediment samples were analysed. The dominant bacterial phyla in both sediment and water were the Proteobacteria, followed by Bacteroidetes and Actinobacteria in the water column and by Chloroflexi and Acidobacteria in the sediment. Biological diversity appeared to be greatly decreased by environmental pollution, whereas the microbial community structure was variable across the analyzed transect. Moreover, a narrow relationship between industrial and urban sources of contamination and the bacterial genera detected at these sites has been observed. A variety of potentially pathogenic bacteria was detected, including Klebsiella, Treponema, Faecalibacterium and Enterococcus, indicating that the river might pose a substantial risk to public health. [Int Microbiol 20(1): 11-24 (2017)] Keywords: environmental pollution · river plankton microbiota · biodiversity

Introduction Rivers and other freshwater habitats play an important role in biogeochemical processes, in which numerous microorganisms, such as bacteria and protists, are integrated into com-

*

Corresponding author: T. Köchling E-mail: th.kochling@gmail.com

plex physiological networks. These networks participate in carbon, nitrogen and other biogeochemical cycles, as well as driving primary production through photosynthetic activity, in particular. Lotic and other freshwater systems also provide important resources to human populations in the form of water for consumption and crop irrigation, as well as transportation routes and areas for recreational activity. Ongoing urbanization and industrialization is often well in advance of environmental protection measures, which leads to formerly

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pristine freshwater systems becoming polluted by industrial on point sources and agricultural and domestic nonpoint sources [2,51]. The contamination of rivers and streams can have a dramatic impact on ecological functioning and poses a serious threat to the health of the population in the drainage area of the disturbed fluvial ecosystems.

As of today, ecological surveys are still needed to assess the structure and composition of microbial populations in lotic ecosystems and to identify composition patterns on a global scale, as well as the possible drivers behind adaptive responses in the microbiota to changing environmental conditions, such as anthropogenic pollution. Previous attempts to assess the community structure and composition of river ecosystems were limited due to the selective bias introduced by the available methodology (culture based techniques) or the limited coverage offered by molecular techniques, such as DGGE (Denaturant Gradient Gel Electrophoresis) or gene library construction, followed by Sanger sequencing [4,13,17,30,45,50,54]. The recent development of economically accessible high-throughput sequencing techniques has provided a far more complete inventory of the microbial communities present in lotic ecosystems, with sampling depths orders of magnitudes higher than those used in previous studies.

By using these new techniques, an emerging set of studies of geographically distinct river ecosystems has expanded our knowledge of lotic [7,9,22,42] and sediment [23] communities, as well as their physiological functions. The microbial communities of rivers in temperate and tropical climates show underlying similarities and differential characteristics. In most studies, Proteobacteria, Actinobacteria and Bacteroidetes are among the most abundant phylogenetic groups encountered, while others, such as Firmicutes and Cyanobacteria, are present in a variable abundance range and are apparently dependent on certain environmental and physicochemical parameters.

The Northeast of Brazil possesses a vast network of freshwater habitats, including many rivers that provide essential resources for domestic, agricultural and recreational purposes. These ecosystems are often put under environmental stress by the introduction of untreated industrial and domestic wastewater and sludge. The Jaboatão River in the vicinity of Recife (state capital of Pernambuco) is an example of an aquatic ecosystem that is exposed to ongoing contamination by industrial and domestic sources in a watershed that is continuously being urbanized and industrialized. The river flows

KÖCHLING ET AL.

through several municipalities where the population uses its water for household purposes, consumption and bathing. The high degree of pollution in the Jaboatão River has direct implications on the public health, given the alarmingly high numbers of hospitalizations caused by waterborne pathogens in the area [19]. Along the river’s course, two paper mills discharge their effluents containing a large number of highly toxic compounds, many of which do not occur naturally [44].

The aim of the present study was to increase knowledge about lotic biodiversity by studying the microbiota of both the sediment and the water column of a tropical river in a progressively urbanized region in Brazil. The survey was performed along a transect from its headwaters in a rural location to areas of dense human population, distributed over six sampling points. This research also sought to determine possible alterations to the biological diversity and community structure resulting from the massive contamination caused by domestic and industrial activities along the river’s course.

Materials and methods Sampling sites. The Jaboatão River is located in the state of Pernambuco in northeastern Brazil. It has a watershed area of 413 km2 and receives input from six communities: Vitória de Santo Antão; Cabo de Santo Agostinho; Moreno; São Lourenço da Mata; Jaboatão dos Guararapes and Recife. The 75 km long river has its source in the vicinity of Vitoria de Santo Antão and flows into the Atlantic Ocean at Barra de Jangada beach, near Recife.

Sediment (S) and water (W) samples were taken on the 29th of May 2013 from six sampling points along the course of the Jaboatão River (Fig. 1). The accumulative precipitation on that day, measured in the town of Jaboatão, was 87 mm [1], being representative of a week with rainfall (the values for the days preceding the sampling date were 120.5 mm, 8.4 mm, 60.3 mm and 51.2 mm). The Jaboatão River is subject to a significant level of anthropogenic contamination throughout its course. In the year 2001 it was exposed to a potential pollutant load of 6679, 25,435 and 35,226 kg of Biochemical Oxygen Demand (BOD) per day by means of industrial, agricultural and domestic activities, respectively [12]. The first three locations are subject to diffuse source agricultural pollution while the other three sites receive untreated wastewater from both domestic and industrial sources. Samples from site S1/W1 were taken near the river’s source. Samples from site S2/W2 were taken from a water collection dam in an area used for the cultivation of sugar cane. Site S3/W3 was located upstream of the community of Moreno in a forested area. S4/W4 samples were taken from Moreno, a densely populated area where the river is polluted by the discharge of untreated domestic and industrial wastewater. A paper mill is located in this city. The S5/W5 site was situated near the biggest settlement along the river, Jaboatão dos Guararapes, downstream from another paper factory. Site S6/W6 was situated in the area of the Jaboatão municipality, close to and upstream of a major cereal processing food company. The sediment samples were retrieved at a depth between 4 and 8 cm, avoiding the upper layer of the sediment that is in contact with the water column. The water samples were retrieved at a depth of approxi-

MICROBIAL DIVERSITY OF A TROPICAL RIVER

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Fig. 1. Area map of the Jaboatão River’s watershed, showing the sampling points (P).

mately 10 cm from the water surface. The values for the river depth at the sampling points were approximately 10 cm (S1/W1), 100 cm (S2/W2, S3/ W3, S6/W6) and 40–50 cm (S4/W4, S5/W5). All samples were taken at distance of 40–50 cm from the river bank. To minimize sampling errors, three sub-samples were collected for each sampling point (both water and sediment) and analyzed as a composite sample.

Physicochemical and bacteriological analyses. The water hardness, salinity and chemical oxygen demand (COD) were measured using standard methods [24]. The water temperature, pH, dissolved oxygen (DO), electrical conductivity (EC) and oxidation reduction potential (ORP) were measured on-site with a Hach HQ40d multi-probemeter (Hach, Loveland, CO, United States). The ORP was also measured in the sediment immediately prior to sampling. Therefore the probe was inserted approximately 6 cm into the sediment. For the other analyses, the samples were stored in sterile airtight containers on ice and processed upon arrival in the laboratory. The total organic carbon (TOC) and inorganic carbon (IC) content were analyzed using a Shimadzu TOC-V CSH analyzer (Shimadzu, Kyoto, Japan). Ion concentrations were determined with a Dionex ICS-1100 (cations) or an ICS2100 (anions) chromatograph (Thermo Scientific, Sunnyvale, CA, United States). The Escherichia coli plate counts of the water samples were determined by filtering different dilutions of the water samples and incubating the filters on Chromocult coliform agar (Cat.# 1.10426, Merck, Darmstadt, Germany) at 37 °C for 24 h. Colony forming units (cfu) of E. coli were determined according to the manufacturer’s instructions.

DNA extraction and PCR. 300 ml of each water sample, or 0.5 g of each well-mixed sediment sample, were subjected to the extraction method with the Powersoil DNA extraction kit (MO BIO, Carlsbad, CA, USA). The water samples were filtered onto a 0.22 µm pore size cellulose ester mem-

brane (Fmaia, São Paulo, Brazil) prior to extraction, while the sediment samples were processed directly, following the manufacturer’s instructions.

The primer set 926F [28] / 1392R [29] was used for the pyrosequencing analysis, partially amplifying the small subunit ribosomal RNA in bacteria and archaea (16S rRNA). PCR was performed with 80 ng of template DNA, 1.25 u Invitrogen Platinum Taq DNA polymerase (Life Technologies, São Paulo, Brazil), 1x reaction buffer, 2 mM Mg2+, 0.2 mM dNTPs (New England Biolabs, Ipswich, MA, USA) and 0.5 μm of each primer, in a reaction volume of 50 μl. Five min at 94 °C initial denaturation were followed by 28 cycles of 30 s at 94 °C, annealing for 30 s at 55 °C, and 30 s at 68 °C of template extension. A final extension step of 10 min at 68 °C ended the program.

Three-replicate PCR products were pooled and purified with the Invitrogen Purelink system (Life Technologies, São Paulo, Brazil). DNA concentrations were measured on a Nanodrop 2000 spectrophotometer (Thermo Scientific, Wilmington, DE, USA) and 454 pyrosequencing was performed by Macrogen (Seoul, South Korea) on a GS-FLX system (454 Life Sciences/ Roche, Branford, CT, USA).

The present study was registered with the National Center for Biotechnology Information (NCBI, Bethesda, MD, USA) under the BioProject identifier PRJNA276740. The data set containing the sequence reads was deposited in the BioSamples database, accessible under the ID numbers SAMN03380194 and SAMN03460198 through SAMN03460208.

Phylogenetic and statistical data analysis. The pyrosequencing raw data were prepared for analysis using SFF Tools v2.6 (Roche 454) and homopolymer errors were corrected with Acacia v1.52 [6]. OTU clustering at a similarity threshold of 97% was performed using UPARSE v6.1.544 [14], excluding singleton OTUs, due to the probability that they were artifacts caused by sequencing errors in homopolymeric stretches. Chimeric sequences were detected and removed using UCHIME v4.2.52 [15]. Assignment of

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Table 1. Physico-chemical measurements of samples taken from the water column, with exception of *ORP, sed (redox potential, sediment sample). All values in mg/l if not indicated otherwise W1

W2

W3

W4

W5

W6

T (°C)

24.9

25.6

26.2

28.1

28.3

27.9

pH

7.3

7.2

6.9

7.3

7.3

6.8

EC (µS/cm]

123.0

107.1

108.9

174.2

250.0

293.0

Salinity [PSU]

0.06

0.05

0.05

0.08

0.11

0.13

DO

8.1

8.0

6.2

6.7

5.9

0.7

ORP, water [mV]

242.0

221.9

105.9

77.7

42.8

84.8

ORP, sed.* [mV]

255.6

–230.0

–116.6

–110.0

–284.1

–206.6

IC

4.0

5.2

6.6

10.8

15.5

22.9

TOC

5.0

5.3

7.3

14.9

11.4

7.6

Nitrate

2.6

1.4

0.8

1.5

3.4

0.5

Chloride

15.4

13.6

13.5

21.5

24.3

25.9

Sulfate

14.5

7.3

4.4

8.1

21.0

26.4

Sodium

13.8

12.5

13.6

19.3

22.0

18.7

Potassium

3.6

3.1

2.5

3.8

5.9

6.4

Magnesium

2.5

2.1

2.2

2.9

3.3

3.8

Calcium

3.7

3.1

2.8

7.3

14.0

22.7

Hardness [mg/l CaCO3]

17.8

14.9

14.7

28.3

46.2

69.7

[COD [mg/l O2]

13.4

14.1

19.6

39.9

30.4

20.4]

E. coli (cfu/100 ml)

16,375±7421

1512±778

4180±2574

60,875±18,268

48,067±8833

19,200±3506

the taxonomic affiliation of the high-quality reads was then performed in Qiime v1.7 [8] applying the RDP classifier v2.6 [49]. To analyze the most abundant bacterial genera in the data set, another OTU matrix was created by assigning all the OTUs that could be determined at genus level (confidence level of ≥ 50 % via the RDP classifier) to their respective genera and obtaining total abundance values.

The calculation of biological diversity indices was performed using the R software environment v3.0 [39], including the vegan v2.0 and GUniFrac v1.0 libraries [11,35]. To allow for the comparison of samples of different size, the OTU matrix was first rarefied to the lowest common sequencing depth obtained in the present study (sample W1, 19791 reads). Several diversity indices were then computed and presented as effective numbers of species (ENS), which is a normalization of diversity measures that permits the quantitative comparison of results between samples and different surveys [27]. The diversity indices used here differ in their sensitivity towards distinctly abundant groups of OTUs. The Shannon-Wiener index takes into consideration even the presence of rare species to contribute to biodiversity, the Berger-Parker metric calculates diversity exclusively based on the proportion of the most abundant OTU and the Simpson index gives a somewhat balanced view, albeit with more weight to the common or more dominant species/ OTUs.

Results Physicochemical and bacteriological analyses.

All of the water samples were turbid with a pH ranging from slightly acidic to slightly basic (Table 1). The total organic

carbon load reached a peak at sampling point 4, which was the first sampling site in an urbanized area along the river’s course with confirmed industrial pollution. ORP values for the sediment samples were all negative, with the exception of the first sampling point near the river’s origin. The water redox potential was positive in all cases, although it was lower in the last three samples, where higher domestic and industrial contamination was recorded. The DO concentration in the water column was by far the lowest in the last sample, with a value close to zero, thus qualifying it as an anaerobic environment, while the other stations ranged from 6 to 8 mg O2/l. Ion concentrations, as well as EC and salinity, correspond with normal values found in freshwaters. They are an index of the quality of the Jaboatão’s waters. The values of IC and hardness run in parallel, increasing from sampling site W3 to W6, showing an increase in the inorganic carbon dissolved. TOC, as well as COD, serve as an index of the contamination the river’s water is exposed to by human activities, showing a peak at site W4 (an urbanized and industrialized area). Concentrations of E. coli covered a wide range from 1500 to 61,000 CFU/100 ml. The highest reads were detected in the last three samples, with a sharp peak at site W4 in the community of Moreno. These values dramatically exceeded the

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Fig. 2. Rarefaction curves for the OTUs of the sediment and water samples.

limit (statistical threshold value) established by the US Environmental Protection Agency, for partial and full body contact, of 410 CFU/ 100 ml [16].

Microbial community structure. After quality filtering and chimera removal, a total of 358,355 high quality sequences remained, which represented 60.2% of the raw data reads. These were then classified into 19,033 OTUs. Table 2 displays the distribution of the bacterial reads and OTUs across the individual samples. While the number of reads obtained for the water samples was similar to those of the sediment, the number of OTUs detected decreased drastically in the last three water samples (W4, W5 and W6).

The OTU coverage of the communities by the sequencing approach was assessed via the Chao 1 estimator. When compared with the numbers of different OTUs detected, the Chao 1 metric indicated a more complete census for the sediment samples (82−90%) than for the water samples (72−79%).

However, the rarefaction analysis shows an asymptotic curve shape at higher sample sizes in all cases (Fig. 2), indicating a valid sampling effort.

The biological diversity of the samples was assessed by a series of descriptors (Table 2). In the present data set, the sediment samples appeared to be distinctly more diverse than the water samples, with values by an order of magnitude higher,

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when considering the Shannon-Wiener, Simpson and BergerParker indices, transformed into ENS [27]. These indices take into account both the OTU richness and the equality of the distribution of individuals throughout the OTUs (evenness).

The reciprocal Berger-Parker index, a simple measure based on the single most abundant OTU of the community, confirmed a more dominant structure for the water and a more even structure for the sediment-derived communities.

The log-abundance plots of the sediment and water samples (Fig. 3) showed two trends in the structures of the corresponding communities. Firstly, the histograms of the sediment communities are approaching a bell shape, which indicates a more uniform distribution of species abundance where relatively few OTUs are very rare, a large number of OTUs are moderately abundant and only a small number of OTUs are very abundant. In contrast, in the water-derived samples, the communities include a higher proportion of rare OTUs and fewer OTUs that contain many members. This indicates a more stable and diversified community structure in the sediment samples, corroborating the observations of the rarefaction analysis and the values of the diversity metrics, in particular the Pielou and Berger-Parker indices. Secondly, the histograms of the water samples W4, W5 and W6 exhibited a flat and tailing shape, when compared with samples W1, W2 and W3, indicating a decrease in rare OTUs and the presence of a small number of highly dominant OTUs with many representatives (sequences).

Microbial community composition. A total of 28 different officially described bacterial phyla, and four candidate divisions, were detected in the sequence set in both the sediment and water samples.

Proteobacteria are commonly found in surveys of river and freshwater microbial communities and were the most dominant phylum in both sediment and water samples in the present study (Fig. 4). While they covered between 52% and 82% of the total number of reads in the water communities, the sediment samples generally exhibited lower abundance values for this group (35−40%) and were more evenly populated by a higher number of different phyla.

While less abundant in the river sediment, members of the class Betaproteobacteria clearly dominated the water-derived communities (35−72% of the sequences) and were by far the most dominant phylogenetic group, followed by the alphaand gamma- classes. Betaproteobacteria affiliated reads were mainly composed of members of the orders Burkholderiales

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Table 2. Descriptors of OTU richness, coverage and alpha-diversity of the sediment and water samples. Shannon-, Simpson- and Berger-Parker indices are transformed to effective numbers of OTUs Number of reads

Number of OTUs

Chao1

Shannon-Wiener [exp(H’)]

Simpson (1/D)

Berger-Parker (1/d)

Pielou evenness (H’/ ln S)

W1

19791

1928

2447

424

100

20.6

0.80

W2

22110

1186

1659

32

4.3

2.1

0.49

W3

31107

1531

2125

73

19.9

8.4

0.60

W4

30101

880

1171

66

22.1

8.4

0.63

W5

31398

738

996

54

18.0

6.2

0.62

W6

30737

920

1230

54

14.3

4.6

0.60

S1

35150

1958

2145

661

256

33.5

0.87

S2

30115

2154

2497

715

273

46.6

0.87

S3

32220

2618

2911

1021

468

75.0

0.89

S4

32040

1395

1698

260

86

16.6

0.78

S5

34355

1630

1917

327

104

20.3

0.80

S6

29241

2095

2347

714

271

40.3

0.87

and Rhodocyclales. Curvibacter and Polynucleobacter were the two most abundant betaproteobacterial genera detected in this survey.

The most common proteobacterial representatives in sediment samples were Beta- (Sphaerotilus, Azonexus genera) and Deltaproteobacteria. The sediments exhibited lower redox potential than the water column. All sediment samples were reductive except S1. These conditions allow a variety of anaerobic bacteria to grow, of which the most abundant phylogenetic group was the Deltaproteobacteria. Within this class, the most common orders were Myxococcales, Syntrophobacterales, Desulfuromonadales and Desulfobacterales.

Besides the predominating Proteobacteria, high proportions of Bacteroidetes were encountered in the Jaboatão River (up to 24% in the contaminated samples). The most abundant orders were Flavobacteriales, Bacteroidales, Sphingobacteriales and the incertae sedis Prolixibacter. Unsurprisingly, due

to the metabolic traits of the classes involved, the most abundant genus affiliated to the Bacteroidetes was different in the sediment and water samples. In the sediment samples, Meniscus, a strictly anaerobic fermenter affiliated to the order Bacteroidales, was common in samples S3 through S6. The most abundant genus in the water column was Flavobacterium, a strict aerobe from the Flavobacteriales order, which was detected in all samples.

The phyla Chloroflexi and Acidobacteria were detected almost exclusively in the sediment samples. The most abundant orders of the phylum Chloroflexi were Anaerolineales, Caldilineales, Dehalococcoidales and Ktedonobacterales. The predominant genus encountered was Bellilinea, a filamentous anaerobic bacterium found in high numbers in all sediment samples. In the case of the Acidobacteria, many members of the Gp subdivision were found, especially Gp1, Gp3, Gp6 and Bryobacter, a recently discovered member of the Gp3 subgroup.

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Fig. 3. Semilogarithmic Preston plots of the OTU abundance distribution of a) the sediment and b) the water samples.

bacteria) and Chlorophyta (green algae). The diverse group of Cyanobacteria and chloroplasts contributed from 0.07% to 2% of the total reads in water and sediment samples.

A subset of 29 bacterial OTUs was present in all the water and sediment samples (data not shown). In terms of bacterial

Fig. 4. Distribution of the relative numerical abundance values for the OTUs detected across bacterial phyla and classes of Proteobacteria.

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Members of six of the Gp subgroups were encountered in the 40 most abundant genera of the sediment samples, suggesting their functional significance in this type of environment.

As indicators of bacterial phototrophic activity, the most abundant microorganisms were Rhodobacter (Alphaproteo-

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richness, this core set represented only 0.4% of the OTUs detected. However, the number of reads that belonged to these 29 OTUs accounted for 16% of the entire 16S rDNA sequence library in the present study. All but two of the 29 core OTUs were affiliated to the Proteobacteria. The remaining two belonged to the Acidobacteria and Firmicutes. The most abundant members of the core population were affiliated to the order Burkholderiales (Betaproteobacteria, 21 to 88% of the core set), represented by the genera Curvibacter, Limnohabitans, Sphaerotilus and Variovorax. The second largest fraction of the core OTU set was affiliated to the order Rhodocyclales (Betaproteobacteria) and contributed between 5 and 34% of the core OTUs. In order to identify the most abundant bacterial genera, the taxonomic assignments of the RDP classifier were filtered so that only OTUs that could be identified at a minimum confidence level of 50% remained. All OTUs pertaining to the same genus were joined and their read abundances were totaled to form a new genus-specific OTU. This new set of taxonomic units covered 53% of the original sequence database.

Taking into consideration the 40 most abundant genus OTUs, the structure of the water microbial communities was skewed towards the class Betaproteobacteria (Fig. 5, lower part of the heatmap). In contrast, the most abundant OTUs in the sediment communities were more evenly distributed across the different phyla. Unsurprisingly, most of the sequences found in the water samples were phylogenetically affiliated to genera that are typically found in aquatic ecosystems, including fresh, polluted and wastewater habitats. From a metabolic point of view, approximately 80% of the 50 most abundant genera were chemoorganotrophs, with a strictly respiratory metabolism that uses oxygen as an electron acceptor, although there are a number of genera that can use nitrate as an alternative electron acceptor. The remaining 20% were strict or facultative anaerobes, the majority of which exhibited a fermentative metabolism, although certain genera use an anaerobic respiratory system.

Discussion The present study shows how environmental pollution affects the biological diversity of microbial communities in a tropical freshwater ecosystem. Differences in community composition between distinct sampling sites were observed, as were possible functions of subgroups of the resident microbiota.

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The sediment samples, for example, contained many OTUs that were not present in the water column, while the latter shared most of its bacterial species with the sediment. A possible explanation for this distribution is that river sediment could act as a reservoir that is rich in bacterial species, releasing microorganisms into the water column via resuspension at the sediment-water interface [20,38].

When comparing the microbial communities in the different samples along the river’s course, the choice of the descriptive metrics was important in terms of determining differences in alpha diversity. While the Pielou index, for example, showed relatively small differences in diversity between the different sets of samples, the Berger-Parker index detected strong variations between the water and the sediment communities. While both descriptors measure the evenness of a community, the Berger-Parker index was probably the more sensitive descriptor as, unlike the Pielou index, it is not constrained to values between zero and one. However, both metrics revealed that dominance increased and evenness decreased in the water-derived microbial communities when compared to the sediment samples. The evolution of the values for the Shannon-Wiener and the Simpson indices are similar along the river’s course. The Shannon-Wiener index estimates the effective numbers of OTUs to be about three to four times higher than the Simpson index due to its inclusion of rare species. The Simpson metric does not consider these species to be contributing to biodiversity to the same degree. The sharp difference in the diversity metrics between sediment and water, with the exception of headwater-derived sediment, shows that, in contrast to the sediment, the water column is not a stable environment.

Due to its movement from the source to the sink of the river the water column at any given point is constantly changing its composition of bacterial species. In addition, factors like dilution by precipitation have a greater impact on the water communities than on those less exposed in the sediment.

The most basic of the diversity indices, the number of different OTUs, showed a marked decrease in the last three water samples (W4–W6). This decrease is also reflected by the values of the Shannon- and the Simpson metrics. This drop in species richness possibly indicates that the high degree of environmental pollution in the most urbanized and industrialized sites along the sampling transect had a profound impact on the microbial communities, exhibiting a strong drive for a smaller number of different species that are resilient enough to grow under more stressful conditions. The Chao1 richness

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estimator predicts lower numbers of species/OTUs for these samples due to their flattened log-abundance curves (Fig. 3), which represent communities with a lower number of rare species. The samples with high numbers of rare OTUs represent the inverse case. Here further sampling will more likely result in the detection of new OTUs. According to the River Continuum Concept by Vannote et al. [48], which describes a river mostly as a longitudinal continuum, alpha diversity (see Table 2) would likely increase along the river’s course, due to the increasing size of the microbial metacommunity, which continuously collects new species downstream. The opposite result was found in the present study for the water communities: diversity and evenness decreased sharply after the first sampling site at the headwaters of the river. The sediment communities were inconsistent in this regard and no pattern could be observed. The most probable explanation for the decline in biodiversity in the water samples might be a) the increase of selective pressure on the microorganisms through the extensive input of mainly organic contaminants and b) the influence of the river’s layout as a dendritic network, which probably leads to the recruitment of new and readily-adapted species at the confluences with smaller streams. At these sites Vannote’s model does not entirely apply, as the communities are modified by the inclusion of new species via mechanisms of resuspension from the sediment and transport along the lotic “conveyor belt” [5].

The overall relative abundance values for members of the Betaproteobacteria in the Jaboatão River were comparable to those detected in the Guyandotte River (USA) [42], the Changjiang River (China) [43], the Dongjiang River (China) [31] and several rivers in the Arctic [13]. The most abundantly present betaproteobacterial orders in the Jaboatão River were the Burkholderiales and the Rhodocyclales. Several members of the Burkholderiales are known for their capabilities in the degradation of even very recalcitrant organic compounds [37], suggesting a high degree of resilience against environmental pollution and a purposeful presence in contaminated freshwaters. They have also been previously encountered in river ecosystems [13,18,50], suggesting that Burkholderiales play a fundamental role in freshwater ecosystems. The Rhodocyclales contain many members that play an important role in the bioremediation of anthropogenic compounds and in biological reactor systems, such as wastewater treatment plants [34]. Among the most abundant representatives of the Rhodocyclales encountered in the present study were Zoogloea and Dechloromonas. These genera are capable

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of degrading a variety of complex organic pollutants [10,32].

Further bacterial phyla were detected at high relative abundance values, as in the case of Bacteroidetes-related OTUs, which coincides with findings from the studies of the Zenne River in Belgium [22] and the Athabasca River in Canada [53], both of which are highly contaminated with organic matter. Also, members of the phylum Actinobacteria are often found in lotic freshwater systems [22, 42,53]. In the present study, the highest abundance values for members of the other Gram-positive phylum, the Firmicutes, were observed at the most contaminated sites (S5, W4 and W5). The Firmicutes are well-known for having many members that are able to degrade even very recalcitrant organic compounds. Interestingly, Firmicutes-related OTUs were also detected to be abundant in the water and sediment samples W1 and S1 near the river’s source, which apparently also suffered from fecal contamination, based on the E. coli counts and the presence of Faecalibacterium. The samples W4 and W5 exhibited the highest E. coli concentrations. The probable source of these peaks in fecal contamination was the discharge of untreated domestic wastewater from the towns of Moreno (W4) and Jaboatão (W5), whereas the pollution of the sampling site W1 was probably caused by small settlements and small-scale farming activities in the area. Possibly because of the proximity to the river’s source the water at the S1/W1 site was more stationary and shallow (about 10 cm deep at a river width of 1.5 m). The combination of a relatively smaller water volume in comparison to the other sections of the river and a slow turnover of the water column might have contributed to the accumulation of fecal microorganisms at this site.

Members of the phototrophic Cyanobacteria exhibited low abundance values for the community composition of water and sediment samples. Higher proportions of these autotrophs were expected to be found in a fluvial system exposed to sunlight in a tropical region. The resuspension of sediment particles in the water column and the high input of allochthonous material through agricultural, domestic and industrial activities along the river’s course might be the causes of the relatively low abundance of phototrophic bacteria in the present study. All of the sampling sites had a turbid river bed. The location directly downstream from the paper mill at S4 was also partially covered by foam. This correlates with the findings of a study of a lotic ecosystem in the USA [42], which compared a largely autochthonous stream with a smaller tributary river characterized by a higher proportion of allochthonous material and elevated turbidity. While the Ohio River

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Fig. 5. Heatmap representing the 40 most abundant OTUs of the a) water column and b) sediment-derived samples and their affiliations at genus level. A new OTU set was generated for this analysis, including reads that could be identified with a minimum confidence of 50% by the RDP classifier.

was abundant in Cyanobacteria, related sequences accounted for less than 1% of the data set in the tributary Guyandotte River.

A variety of bacterial genera in the samples have been associated with the degradation and breakdown of xenobiotic compounds. Dehalogenimonas (phylum Chloroflexi) can reductively dehalogenate polychlorinated aliphatic alkanes and has been isolated from chlorinated solvent-contaminated groundwater. Dechloromonas (class Betaproteobacteria) exhibits a strictly respiratory metabolism, using chlorate, perchlorate and O2 as electron acceptors and organic matter (acetate, propionate, butyrate, fumarate or succinate) as an electron donor. Syntrophus (Deltaproteobacteria) grows in syn-

trophic association with H2-utilizing microorganisms and oxidizes substrates (crotonate, aromatic compounds, or fatty acids) to acetate. Syntrophus-affiliated sequences have been detected in anoxic sediments and sewage sludge. In the present study, members of this genus were most abundant in the sediment sample S5 (2.3% of all reads).The presence of these microorganisms is a possible indication that the JaboatĂŁo River is exposed to a variety of xenobiotic compounds that are generated by industrial activity, including the two paper factories.

In the sediment-derived samples, the 50 most abundant genera ranged from 35−48% coverage of the sequences that were reliably assigned down to the genus level and were evenly distributed between aerobic and anaerobic microorganisms,

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many of them isolated from sediments, water or anaerobic granular sludge. This fact seems to contradict the negative redox potential measured in the sediments (with the exception of S1). Since dissolved oxygen levels were relatively high throughout the river (from 6–8 mg/l, with the exception of W6: 0.7 mg/l), it can be hypothesized that, due to the heterogeneity of the river bed, there were microniches in the sediments in which aerobic bacteria could survive [40]. The high abundance of Geobacter and Bellilinea was common in the sediment samples, as well as several members of the Acidobacteria related uncultured Gp cluster. The low oxygen concentration at W6 is representative for this section of the river. Analyses performed in earlier campaigns in the years 2007 and 2008 showed a comparable mean value of 1.5 mg/l (data not shown). The low oxygen levels at this site were probably caused by the proximity to densely populated areas and an industrial park. Furthermore, an uncontrolled waste dump existed in the vicinity which continues to release its leachate to the surrounding area, including the Jaboatão River.

Nine out of the 50 most abundant genera detected in the present study were present in both water and sediment. However, with the exception of Geobacter, they were predominant in the water samples (Fig. 5). Curvibacter, Sphaerotilus, Piscinibacter and Zoogloea are Betaproteobacteria typically present in fresh or polluted waters. Sphaerotilus lives, attached to submerged plants or stones, in slowly running freshwater that is heavily contaminated with sewage or wastewater. Interestingly, strains of S. natans have previously been isolated from paper mill slime [36]. In the present study, Sphaerotilus-related OTUs exhibited a single peak in abundance in the samples S5 and W5, close to the outlet of a large paper factory, while the abundance values were relatively low in the other samples.

Geobacter was the dominant genus and one of the most abundant microorganisms in the sediment samples. It is known for its unique metabolic properties and ability to anaerobically degrade organic compounds, such as hydrocarbons. Geobacter oxidizes organic compounds, with Fe(III) serving as the sole electron acceptor.

The abundance and diversity of genera affiliated to the Chloroflexi in the sediments is striking.

Non-photosynthetic members of this phylum are usually found in anaerobic environments, both natural as well as engineered systems (such as anaerobic wastewater reactors in which they can be a co-dominant part of the population). For this reason, it has been suggested that they must play an im-

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portant role in the degradation of organic matter in such environments [41,52]. Several genera affiliated to the Chloroflexi (e.g., Bellilinea, Anaerolinea and Caldilinea) are moderate thermophiles. Leptolinea was isolated from the granular sludge of a UASB reactor treating wastewater from a sugarprocessing plant [52].

In our data set, a subgroup comprised of only 29 different OTUs, but with high abundance values, was present in all of the analyzed samples. This structural feature of the microbial communities could indicate how a very small fraction of the bacterial biodiversity could act as a ubiquitous backbone of the freshwater ecosystem of the Jaboatão River. The bacterial species of the core OTU set, most of which were affiliated to the Proteobacteria, might possibly be the most resilient towards the anthropogenic disturbances along the river’s course and its varying physicochemical conditions.

OTUs affiliated to several possibly pathogenic bacterial species were detected. The highest abundance values for Escherichia/Shigella, Enterococcus, Cloacibacillus, Klebsiella and Coprococcus were found in the samples from W4 or W5, while Faecalibacterium was most abundant at W1. Enterococcus is a common part of the human intestinal biota and is a potentially disease-causing bacterium. Strains of Cloacibacillus (phylum Synergistetes) have been isolated from the intestinal tract of pigs and from anaerobic digester sludge [21,33]. Members of the Synergistetes are often found in insalubrious samples and should therefore be considered as potentially harmful to humans. The genus Klebsiella is ubiquitous in nature and can occur inhuman and animal intestinal content, water, sewage and soils. It has also been recovered from aquatic environments that receive wastewater. The genus can be associated with a variety of infections, from bacterial pneumonia to nosocomial urinary tract infections. Treponema (Spirochaetes) affiliated species were also detected in significant proportions in several sediment samples, with a peak at S3 and a relative abundance of 2.5% of all reads. Treponema pallidum carateum causes Pinta disease, an infectious skin condition that is commonly transmitted in tropical regions. A number of members of the Bacteroidetes are known as pathogens for animals (Flavobacterium) or as indicators of fecal contamination (Bacteroides). The latter were mainly found in the water samples W1, W4 and W5, which coincided with the highest plate counts for E. coli, the most widely used fecal indicator organism.

Two genera associated with pathogenicity and industrial activity were detected at relatively high abundance values:

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Cloacibacterium (phylum Bacteroidetes) OTU counts peaked in samples W4 and W5, with relative abundance values of 9 and 16%, respectively, while Aeromonas (class Gammaproteobacteria) affiliated sequences accounted for 12% of the reads in sample W5. Cloacibacterium species have been isolated from untreated human wastewater [3]. Aeromonas is known as an opportunistic pathogen, although it is also ubiquitous in a wide variety of pristine aquatic ecosystems and is therefore unlikely to be a valid indicator for the human contamination of freshwater bodies [46]. However, both Cloacibacterium and Aeromonas have been associated with the Kraft pulping process used in paper production. Interestingly, a paper mill was located in the vicinity of the sampling point at both sites where these genera exhibited their peak read abundance values. Cloacibacterium has been isolated from paper mill pulp [47] and Aeromonas has been successfully used for the treatment of lignin rich paper mill liquor [25].

Besides their potential to act as a facultative pathogen in humans and as a pathogen in the fishery industry, Aeromonas species were found to increase in abundance and survive near the discharge point of a paper mill in the Albemarle Sound in North Carolina, USA [26]. This example shows how locallydefined sources of industrial pollution could influence the structure of the Jaboatão water communities and increase the pathogenic capacity of the water body. While the biological diversity sharply decreased at the sampling sites in the vicinity of the paper mills, a small subpopulation of resilient and adapted species prevailed and dominated the bacterial communities. At W5, the combined relative abundance values for Cloacibacterium and Aeromonas accounted for 28% of the sequence reads, while at the sites that were further away from the paper production facilities, their abundance proportions were well below one percent. Our data set contained a variety of potentially dangerous bacteria for public health, which would otherwise have gone unnoticed if relying on culturedependent methods alone. The parsing of high-throughput 16S rDNA libraries of samples that are a risk to public health presents a possible strategy to assess the threat level, including microorganisms in the analysis that are unknown or not suspected to be present in the sample.

The present study shows how a massive level of anthropogenic contamination is capable of altering the microbial community structure and negatively affecting the biodiversity of a river ecosystem, particularly the water column. In this regard, the combination of several monometric descriptors such as the Shannon-Wiener diversity, the Berger-Parker index and

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OTU richness, as well as multidimensional approaches, such as the recording of log-abundance profiles for microbial communities, proved to be useful in terms of assessing the degree of disturbance in polluted rivers. Industrial on point contamination, which in this case involved the discharge of untreated domestic and paper mill wastewater and sludge, resulted in a decrease of bacterial diversity in water samples.

Pyrosequencing of the 16S rRNA gene is one feasible method of analyzing the taxonomic composition of river microbial communities. The rarefaction curves showed that increasing the sample size, within reasonable limits, would lead to the inclusion of the entire population of the ecosystem under study in the domain Bacteria.

The taxonomic assignment of NGS sequence tag libraries also confirmed fecal contamination, providing information on present pathogenic bacteria down to the genus level, most of which would not have been detected using culture-dependent methods.

Acknowledgments. We thank the Brazilian agency FACEPE (Fundação de Amparo à Ciência e Tecnologia do Estado de Pernambuco) and the CNPq (Conselho Nacional de Desenvolvimento Científico e Tecnológico) for financial support (Grant APQ - 1065-3.07/10). J. L. Sanz received a fellowship as a visiting professor from the Federal University of Pernambuco (UFPE), Brazil (D.O.U. no. 74, p. 102, 18th April, 2013). We also thank CENAPAD-PE (National Center for High Performance Computing, Pernambuco) for providing the computation resources for the treatment of the data sets.

Competing interests. None declared.

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49. Wang Q, Garrity GM, Tiedje JM, Cole JR (2007) Naive bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl Environ Microbiol 73:5261-5267 50. Winter C, Hein T, Kavka G, Mach RL, Farnleitner AH (2006) Longitudinal changes in the bacterial community composition of the Danube River: a whole-river approach. Appl Environ Microbiol 73:421-431 51. Wu Q, Leung JY, Tam NF, Peng Y, Zhou S, Li Q, Geng X, Miao S (2016) Contamination and distribution of heavy metals, polybrominated diphenyl ethers and alternative halogenated flame retardants in a pristine mangrove. Mar Pollut Bull 103:344-348 52. Yamada T, Sekiguchi Y, Hanada S, Imachi H, Ohashi A, Harada H, Kamagata Y (2006) Anaerolinea thermolimosa sp. nov., Levilinea saccharolytica gen. nov., sp. nov. and Leptolinea tardivitalis gen. nov., sp. nov., novel filamentous anaerobes, and description of the new classes Anaerolineae classis nov. and Caldilineae classis nov. in the bacterial phylum Chloroflexi. Int J Syst Evol Microbiol 56:1331-1340 53. Yergeau E, Lawrence JR, Sanschagrin S, Waiser MJ, Korber DR, Greer CW (2012) Next-Generation Sequencing of microbial communities in the Athabasca River and its tributaries in relation to oil sands mining activities. Appl Environ Microbiol 78:7626-7637 54. Zhang M, Yu N, Chen L, Jiang C, Tao Y, Zhang T, Chen J, Xue D (2012) Structure and seasonal dynamics of bacterial communities in three urban rivers in China. Aquat Sci 74:113-120

RESEARCH ARTICLE International Microbiology 20(1): 25-30 (2017) doi:10.2436/20.1501.01.282. ISSN (print): 1139-6709. e-ISSN: 1618-1095

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Identification by using MALDI-TOF mass spectrometry of lactic acid bacteria isolated from non-commercial yogurts in southern Anatolia, Turkey Ayse Karaduman,1* Mehmet Ozaslan,1 Ibrahim H. Kilic,1 Sibel Bayil-Oguzkan,2 Bekir S. Kurt,1 Nese Erdogan1 Department of Biology, Faculty of Art and Science, University of Gaziantep, Gaziantep, Turkey.

Department of Medical Services and Techniques, Health Services, University of Gaziantep, Gaziantep, Turkey 1

2

Received 27 January 2017¡ Accepted 15 March 2017

Summary. Yogurt is a dairy product obtained by bacterial fermentation of milk. Commercial yogurts are produced using standard starters while, in the production of non-commercial yogurt, the microbiota is quite different since yogurts are used as starter for years. To determine the final characteristics of the fermented product it is necessary to know the biochemical properties of the starter cultures, such as acidity, aroma and flavor. This can only be achieved by identifying and characterizing the bacteria in starter cultures. In our study, 208 non-commercial yogurt samples were collected from 9 different locations in Anatolia, southern Turkey. Their pH and lactic acid bacteria profiles were analyzed. Isolated bacteria were identified by MALDI-TOF MS (matrixassisted laser sesorption-ionization time-of-flight, mass spectrometry), which is a fast and reliable method for identification of bacterial isolates compared to classical laboratory methods. In this study, 41% of the isolates were identified by using this method, which is 99.9% and 34.0% confidence. The isolates contained two genera (Enterococcus and Lactobacillus) and four species.

Afterwards, the four lactic acid bacteria were characterized physiologically and biochemically and we found that they differed from lactic acid bacteria used in commercial yogurt production. [Int Microbiol 20(1): 25-30 (2017)] Keywords: yogurt starters ¡ lactic acid bacteria (LAB) ¡ southern Anatolia (Turkey)

Introduction Yogurt is a dairy product obtained by the bacterial fermentation of milk. Lactic acid, which is produced via the fermentation of the milk sugar lactose, is the compound that confers

Corresponding author: A. Karaduman E-mail: ayse.karaduman1@hotmail.com *

part of the typical textural and sensory characteristics of yogurt and acts on milk protein [2]. Yogurt, which contains high concentrations of lactic acid bacteria (LAB), has been described to have several potential health benefits: it can increase lactose tolerance, balance the intestinal microbiota, act as an antimicrobial compound, stimulate the immune system, induce anti-tumor effects, and induce anti-cholesterolemic effects [6]. Today, the vast majority of yogurt needs are met by commercially produced yogurts, and the goal of companies is

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to produce more solid, continuous sweet yogurt with a variety of additives. Commercial yogurts are produced thereby using standard starters.

During the fermentation of food stuffs, the main physical and chemical changes occur due to the growth and fermentative activity of the LAB that are used as starter cultures. LAB are also used as starter cultures for the fermentation of milk [22] and have been used for centuries as food preservatives [7,11]. Some LAB ferment lactose into lactic acid and thus help to acidify the final product. Others digest the proteins and lipids in milk producing several other substances that contribute to the formation of the unique flavor, aroma, appearance, and structure of a fermented product [18].

LAB comprise a genetically diverse group of bacteria that includes the following genera: Carnobacterium, Enterococcus, Lactobacillus, Lactococcus, Leuconostoc, Oenococcus, Pediococcus, Streptococcus, Tetragenococcus, Vagococcus and Weissella [15]. Those of industrial importance comprise Lactococcus, Enterococcus, Oenococcus, Pediococcus, Streptococcus, Leuconostoc and Lactobacillus species [17]. LAB are typically classified according to categories such as morphology, Gram staining, oxidase and catalase tests [7,13,20], growth at different temperatures and salt concentrations [26,27], carbohydrate fermentation [27,31], and cell wall composition. Knowledge of the biochemical properties of the starter cultures is necessary to determine the final characteristics of the fermented food product, such as acidity, aroma and flavor. To gain such knowledge, it is crucial to identify and characterize the bacteria in a starter culture.

Analysis of protein profiles of microorganisms carried out with the MALDI-TOF (Matrix-assisted laser desorption-ionization time-of-flight) mass spectrometry are compatible with the results of identification made by using molecular methods [25].Therefore, MALDI-TOF can be considered a robust and reliable method, as well as fast and inexpensive to identify Gram-positive bacteria such as lactic acid bacteria [25,30].

The purpose of this study was to isolate and characterize LAB from collected non-commercial yogurts in southern Anatolia and find out whether they differed from the LAB used for the production of commercial yogurts.

Materials and methods Traditional yogurt production and samples collection. A total of 208 non-commercial yogurt samples were collected from nine vil-

KARADUMAN ET AL.

lages in southeastern Anatolia, Turkey, from March to May, 2015 (The various locations are indicate with codes in the Tables.). Yogurts had been produced with traditionally methods by people from those villages. They used cow milk that was heated to 85 °C for 20 min and was then cooled rapidly to 46 °C. After this sort of pasteurization, one cup the inoculum obtained from a previous fermentation was added to approximate 1 liter of cooled milk. Every yogurt sample came from a different inoculum. After inoculation, milk was incubated at room temperature (or 5 °C higher) for one night or 6–7 h. Afterwards, the yogurt obtained was kept in a refrigerator at 4 °C. The pH values of the home-made yogurts were measured with a pH meter (Martini Instruments, Romania) and recorded.

Bacterial isolates. Agar plates of media M17 (Oxoid, UK) (50 ml of 10% lactose solution was added) and MRS (Oxoid, UK) were inoculated by dilution. The plates were incubated aerobically at 37 °C and 30 °C for 72 h.

The numbers of the resulting colonies were determined as CFU (colony forming units)/g. Phenotypic characterization (colony morphology, Gram staining and catalase assay) of LABs isolated from colonies was performed. Afterwards, they were identified by using MALDI-TOF mass spectroscopy.

Isolation and total bacterial counts. The samples were diluted before inoculating them on M17 agar (Oxoid, England) and MRS (de Man, Rogosa and Sharpe) agar (Oxoid, England). The M17 and MRS plates were incubated for 3 days in aerobic environments at 37 °C and 30 °C, respectively [10,12,14,21]. All of the suspect colonies in petri dishes were recorded as CFU/g [21].

Stock culture preparation. A sterile loop was used to sample colonies and to inoculate them into Eppendorf tubes containing 1 ml of M17 Broth (Oxoid, England) or MRS Broth (Oxoid, England). Bacterial samples inoculated into M17 were incubated for 48 h. Stocks were prepared from 500 µl of LAB cultured in MRS broth or M17 broth and 500 µl of sterile liquid glycerol that were mixed in a 1 ml Eppendorf tube and stored at –20 °C [5].

Identification of LAB. The phenotypic identification of colonies was conducted according to their culture properties (Gram staining), their cell morphology (cocci or rods) under the light microscope and their biochemical properties (catalase test) [4,5,24]. For these tests, stock cultures were grown on M17 agar and MRS agar as described in the previous paragraph.

The identification of LAB colonies isolated from non-commercial yogurts was carried out according to their protein and peptide registered information analysis using MALDI-TOF MS, which is a new technology for identifying LAB cultures. The advantages of this embodiment are that it has rapid and routine properties. For these tests, stock cultures were grown on M17 and MRS agar, and applied according to Susakol et al. [5]. The isolates were tested against various surface proteins. All suspicious isolates were selected for LAB identification with the MALDI-TOF MS test kit. Protein patterns of these isolates were analyzed by MALDI. Bruker MS (Becton-Dickinson USA) was used to identify the microorganisms. In the MALDI-TOF method, the Bruker device transforms the so-called matrix into a molecule of interest (DNA or proteins) by absorbing the matrix light after the ion beam is applied to a chemical surface that has been exposed to light for 24 h.

This method, which is used for extracting protein products from microorganisms, is easily recognizable by the result of comparing these profiles with a reference spectrum. In definition: 1 drop of physiological saline was sprayed onto the matrix, and fresh (24 h) samples were taken from the cultures. After

STARTER CULTURES IN ANATOLIA YOGURTS

mixing with the serum physiological solution in the matrix and drying in the air, identification was made in the MS device.

Characterization of LAB. The growth properties of LAB species that were isolated from the yogurt samples were determined at 4 °C, 25 °C and 45 °C [8] at pH 2, pH 5 and pH 7 [29] and at 3%, 6% and 10% NaCl (table salt) [20].

Results pH measuring. The pH values of the 208 yogurt samples analized ranged from 3.9 to 6.0. The average pH from same sources ranged between 4.35 and 4.97 (Table 1).

Isolation and total bacterial counts. The total bacterial counts on MRS agar medium were 0–2.3 × 1010 CFU/g, while the total bacterial counts on M17 agar medium were 3.7 × 108–3.1 × 1010 CFU/g following the isolation of LAB from the yogurt samples. Out of the 731 variety analyzed isolates, 503 were cocci (360 cocci were identified on M17 medium and 143 cocci were identified on MRS medium) while 228 of the analyzed LAB were rods (100 on M17 medium and 128 on MRS medium). Overall, 69% of the isolates were cocci and 31% were rods. All of the LAB were non-motile.

Bacterial isolates. Of the 503 cocci isolates, 86 (17%) belonged to the genus Enterococcus: 45 were Enterococcus hirae (9%), 36 were Enterococcus faecium (7%), 5 were Enterococcus durans (1%). Of the 228 rod isolates, 54 (24%) belonged to the genus Lactobacillus, the sepcies Lactobacillus casei or Lactobacillus paracasei. We attempted to classify the rest of cocci and rod isolates, but the data obtained were not reliable. The distribution of different LAB among the yogurts collected is shown in Table 2.

All of the LAB were Gram-positive. Gram (+) bacteria were selected by Gram staining. Additionally, LAB do not encode a catalase enzyme, and thus they cannot degrade hydrogen peroxide (H2O2) to water (H2O) and oxygen (O2) molecules. The results of catalase tests were negative (–) for the isolated LAB. The LAB identification results revealed that the LAB microbiota in the Turkish yogurts analyzed contained similarity 99,9%; 7% Enterococcus faecium, 9% E. hirae, 1% E. durans; contained similarity 34,0%; 24% Lactobacillus casei or L. paracasei (rod isolates are presented with either of them by 34,0% similarity). Spectra of four

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Table 1. The pH values of yogurts collected from villages in Turkey Villages

n

pH values Smallest

Biggest

Average

S1

30

4.00

5.90

4.40

S2

30

4.00

6.00

4.53

O

30

4.10

4.90

4.35

K

15

4.30

5.00

4.62

I

15

4.90

5.00

4.97

N1

15

4.40

4.80

4.63

N2

28

4.30

5.20

4.55

A

15

3.90

4.90

4.43

Y

30

4.30

4.70

4.43

(n = sample counter; S1, S2, O, K, I, N1, N2, A, Y = codes of nine locations, villages)

species obtained by MALDI-TOF MS are shown in Fig. 1.

The most broadly growing isolates were Enterococcus durans and E. faecium (the latter grew under all conditions except pH 2; 3%, 6% NaCl), Enterococcus hirae (it grew under all conditions except pH 2, 5; 3% NaCl). In contrast, the most sensitive isolates were Lactobacillus casei and L. paracasei (they former failed to grow at 4, 45 °C, at 10% NaCl and at pH 2; and the latter failed to grow at 4, 45 °C, at 6%,

Table 2. The distribution of different LAB among yogurts collected from villages in Turkey L. casei ?

L. paracasei ?

Villages

n

E. faecium

E. hirae

E. durans

S1

30

7

2

–

1

S2

30

18

3

–

1

O

30

11

8

–

5

K

15

–

20

–

11

I

15

–

7

5

3

N1

15

–

5

–

–

N2

28

–

–

–

18

A

15

–

–

–

9

Y

30

–

–

–

6

36 isolates

45 isolates

5 isolates

Total

86 isolates cocci

54 isolates 54 isolates rod

(n = sample counter; S1, S2, O, K, I, N1, N2, A, Y = codes of nine locations, villages)

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Fig. 1. Spectra of four species obtained by MALDI-TOF-MS. (A) Spectrum of Enterococcus hirae (99,9% similarity). (B) Spectrum of Enterococcus faecium (99,9% similarity). (C) Spectrum of Enterococcus durans (99,9% similarity). (D) Spectrum of Lactobacillus casei (34,0% similarity), or Lactobacillus paracasei (34,0% similarity).

10% NaCl and at pH 2). Almost all LAB could grow, either weakly or strongly, at 25 °C and at pH 7. The characteristics of the LAB species isolated are shown in Table 3.

Discussion Lactic acid bacteria (LAB) play an important role in food technology. They are the microorganisms that typically used for the production and long-term preservation of various fermented dairy products such as yogurt, buttermilk and cheese. LAB are considered generally recognized as safe (GRAS) microorganisms [16,23]. They are cocci or rodshaped bacteria that primarily produce lactic acid as the end product of carbohydrate fermentation [1,3,23]. The results of this study showed that the pH levels of then non-commercial yogurts from villages in Turkey that we analyzed were acidic (below 7) as a result of lactic acid fermentation; (their pH ranged from 3.9–6.0). As no standard inocula were applied in this study, the relatively high pH of some yogurts could be related to both yeasts and bacteria present in the yogurt total microbiota. In fact, 80 colonies of yeasts were

also found under the microscopy in 20 yogurt samples. With striking resemblance, yeasts were observed in the samples from S2 (30 colonies) and S1 (7 colonies) locations, which also had the highest pH values. These yeasts, however, have not been identified yet. The animal sources of the milk from which the yogurts were obtained varied; they included cow milk, sheep milk and goat milk. Erkaya and Sengul [9] have described pH levels of yogurts produced from different milk types ranged between 4.02 and 4.26 and they have reported that these pH values were related to the metabolic activities of the starter cultures and the fermentation time of the yogurt [9]. In addition, Omafuvbe and Enyioha [20] have found pH values ranging from 3.80 to 4.48 in yogurts obtained from villages in Nigeria [20]. They have also reported that these levels were lower than pH levels of commercial yogurts. In several countries, LAB are present in yogurt in numbers of at least 106 to 5 × 108 CFU/ml, depending on the type of yogurt [6]. According to the Turkish Food Codex, the total microorganism count in yogurt must exceed 106 CFU/g [19]. In this study on non-commercial yogurts, we found LAB counts of at least 3.7 × 108 CFU/g when the counts from M17 agar and MRS agar were considered together

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Int. Microbiol. Vol. 20, 2017

Table 3. The characteristics of the LAB species isolated from noncommercial yogurts from nine locations in Turkey.

LAB species

Characteristics Growth in medium with NaCl (%)

Growth at pH (acidity)

4, 25, 45

3, 6, 10

2, 5, 7

L. casei (R) or L. paracasei

–, +, –

+, –,–

–, +, +

E. durans (C)

w, +, w

–, –, +

–, +, +

E. hirae (C)

+, +, w

–, +, w

–, –, w

E. faecium (C)

w, +, w

–, –, +

–, w, w

Growth of temperature (0C)

Note: C: cocci, R: rod, +: positive growth, –: no growth, w: weak growth

(that is, adding the lowest number of CFU on MRS agar [zero growth] and the lowest number of CFU on M17 agar [3.7 × 108 CFU/g]). Therefore, the their LAB numbers were within the range of LAB abundancec in other yogurts in Turkey and in other countries. Enterococcus faecium, E. hirae, E. durans and Lactobacillus casei or L. paracasei were the most frequent members of the yogurt LAB microbiota.

These results suggested that non-commercial yogurts might develop in different ways and have a more wide profile of LAB species than commercial yogurts. Enterococcus faecium, E. hirae, E. durans (sensitive to low acid and low salinity) and Lactobacillus casei or L. paracasei (sensitive low acid and high salinity to low or high temperature) were the most interesting species among the isolated LAB species in terms of their characteristic properties. These LAB strains could be used for the fermentation process of yogurt production and are good candidate starter cultures.

MALDI has been successfully applied in recent times. It is a convenient and routine technique used to identify with 99% similarity of MALDI-TOF isolates. With only one colony, it is possible to identify it in just 10–15 min. MALDI is seen as a method that can be used in the food industry, in which rapid diagnostic procedures are needed. In addition, this method can be an inexpensive alternative to conventional methods. In fact, the identification fee for a single isolate is 1.2 US $. In our study, this method was strengthened by phenotypic identification and 2 different genus and 4 different species were identified. Rodríguez-Sánchez et al. [25] identified 3 L. casei [25] by same method such as 54 L. casei or Lactobacillus paracasei (similarity 34%), which we identified by MALDI-TOF MS in our study or 54 isolates Su-

29

sakul Palakawong et al. [28] defined two new bacterial strains of the Actinomyces genus, such as Gram-positive (+) anaerobic bacterial strains [28] that we identified with MALDI-TOF MS in our study. Streptococcus thermophilus and Lactobacillus delbrueckii ssp. bulgaricus, which are used as starter cultures in commercial yogurt production, were not isolated in this study. The reason why they not were found in our study is that both of them need microaerophilic or anaerobic conditions for isolation. Incubation on MRS at 30 ºC under aerobic conditions, as used in our experiment, might have hindered the growth of L. delbrueckii, especially when it had to compete with the mesophilic bacteria found.

Although lactose has to be present in the M17 medium, growth of S. thermophilus may be stunted under aerobic conditions. However, except for these species, other five species that we isolated had not been previously described to have the potential or ability to produce yogurt. These additional species might represent factors underliying the flavors and properties of non-commercial yogurts, different from commercial yogurts. Important results have been obtained for the provision of yogurts to Turkish consumers, who enjoy their local yogurt flavors. This study suggests different functions for different bacteria during the yogurt production, as we isolated bacterial species that have not often been isolated in other several studies. For instance, we isolated E. hirae, E. faecium, E. durans, Lactobacillus casei and L. paracasei strains from yogurts in which we could expect to find S. thermophilus and L. delbrueckii ssp. bulgaricus strains.

Other previous yogurt-related research in Turkey have identified only S. thermophilus and L. delbrueckii ssp. bulgaricus species.

Our study focused on the basic development of starter cultures for non-commercial yogurt production. The yogurt bacteria isolated in this study are used in several combinations that allow the development of a starter culture that is the basis for the unique taste of yogurt. To our knowledge, our study is the first to specifically analyze non-commercial yogurt microbiota in southeastern Anatolia, Turkey.

Acknowledgements. This study was supported by the Scientific Research Projects Governing Unit of Gaziantep University (FEF. 14.14). AK, MO and IHK designed the study and performed draft of working. BSK and NE did laboratory analysis. MO, IHK and SBO performed analysis evaluations.

Competing interests. None declared.

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RESEARCH ARTICLE International Microbiology 20(1): 31-41 (2017) doi:10.2436/20.1501.01.283. ISSN (print): 1139-6709. e-ISSN: 1618-1095

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Insights into the fecal microbiota of captive Iberian lynx (Lynx pardinus) José Alberto Núñez-Díaz,1 M. Carmen Balebona,1 Eva M. Alcaide,2 Irene Zorrilla,2 Miguel Ángel Moriñigo1* Universidad de Málaga, Andalucía Tech, Departmento de Microbiología, Campus de Teatinos, Málaga, Spain.

Division of Sustainability and Urban Environment Analysis and Diagnostic Center of Wildlife-C.A.D. Environment Agency and Water Andalusia, Málaga, Spain 1

2

Received 7 March 2017 · Accepted 30 March 2017

Summary. The Iberian lynx (Lynx pardinus) is an endangered species restricted to several areas of Spain and Portugal. Its low genetic diversity likely provokes immune depression and high susceptibility to infectious diseases. The intestinal microbiota is closely related to host health and nutrition. In order to contribute to the knowledge of the Iberian lynx intestinal microbiota, fecal microbiota of captive specimens from two breeding centers (“La Olivilla” and “El Acebuche”), located in Southern Spain, were studied by Denaturing Gradient Gel Electrophoresis (DGGE). Results grouped microbiota in two main clusters (I and III) which included DGGE patterns of 19 out of 36 specimens, cluster I being the most frequent in “La Olivilla” (50%) and cluster III in “El Acebuche” (55.55 %) specimens. Bacteroidetes, Firmicutes and Proteobacteria phyla were identified. Segregation of clusters I and III was attributed to different microorganism presence (Pseudomonas koreensis, Pseudomonas migulae, Carnobacterium sp., Arthrobacter, Robinsoniella peorensis and Ornithinibacillus sp.) and ability to use different carbon sources. Biolog EcoPlates® results indicate high functional diversity of fecal microbiota, it being higher in cluster III. The great impact of intestinal microbiota on host health supports the importance of its microbial composition understanding. This study is the first report of captive Iberian lynx fecal microbiota composition. [Int Microbiol 20(1): 31-41 (2017)] Keywords: Iberian lynx (Lynx pardinus) · fecal microbiota · biodiversity

Introduction The Iberian lynx (Lynx pardinus) is listed as endangered by the World Conservation Union (IUCN). At present, it is settled in Doñana-Aljarafe and Sierra Morena areas in Southern

*

Corresponding author: Miguel Ángel Moriñigo E-mail: morinigo@uma.es

Spain and it is being reintroduced in other areas of Spain and Portugal [47] (Iberlince project web). Studies related to phylogeny, histopathology and immunohistochemistry indicate low genetic diversity [1,10] and frequently depressed immune system [45], with high susceptibility to infectious diseases [34]. The precarious situation of the wild Iberian lynx led to the establishment of the Iberian Lynx Lynx pardinus Conservation Breeding Program [54], that follows a multidisciplinary approach encompassing management and applied re-

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search strategies in different areas. One of these areas is related to health and veterinary aspects, and the establishment of preventive disease protocols for the captive populations is one of the Program’s main lines of action [54].

Intestinal microbiota of mammals play a very important role mainly due to facilitating the absorption of ingested food by degradation of macromolecules [41] and protecting against potential pathogens [9]. In this way, research has convincingly demonstrated that the microbiota is crucial in order to prime and orchestrate innate and adaptive host immune responses [22]. This microbiota constitutes a complex ecosystem consisting of a big amount and diversity of bacterial species and the quantity and types of bacteria are closely linked to environmental variables. Understanding the gut microbial community structure is critical to identify and perform the potential fitnessrelated traits for the host [56]. In this context, knowledge of the captive Iberian lynx intestinal microbiota can be an useful tool in future preventive strategies against the disease. Several studies have characterized the gut microbiota of felines, such as cats [25], but in the case of the Iberian lynx, only one study that analyses distal gut microbial biodiversity of one specimen has been carried out [3]. For this reason, further studies with a higher number of animals are necessary to increase the understanding of the Iberian lynx intestinal microbiota.

The objective of this study was to assess the composition of the predominant microbial groups in the fecal microbiota of Iberian lynx specimens from two breeding centers where they are confined, “El Acebuche” and “La Olivilla” located in Doñana and Sierra Morena areas, respectively. These two breeding centers account for more than 50% of the captive Iberian lynx population.

Materials and methods Fecal samples of Iberian lynx. A total of 36 fecal samples of captive Iberian lynx specimens were kindly supplied by Centro de Análisis y Diagnóstico de la Fauna Silvestre de Andalucía (Málaga-Spain) within the context of the routine program of coprologic analysis of this species. Fecal samples were collected in November 2011 and within 3 h after defecation from 36 captive Iberian lynx specimens kept in the breeding centers of “El Acebuche” (10 males and 8 females) (15 ºC, average temperature) and “La Olivilla” (6 males and 12 females) (12 ºC, average temperature), located in Doñana and Sierra Morena areas (Andalucía, Southern Spain), respectively. Diet of Iberian lynx specimens consisted of rabbit obtained from the same supplier (Las Lomas, Cádiz, Spain) along the study period.

After collection, samples were immediately frozen at –20ºC until further analysis. Fecal samples came from apparently healthy specimens and none of them had a history of antibiotic use.

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Analysis of fecal microbiota. A portion of 50 mg of the center of feces was obtained in aseptic conditions. Total DNA was extracted from each sample as described by Martínez et al. [37]. DNA was quantified using a spectrophotometer (NanoDrop 1000, Thermo Scientific).

In order to compare DGGE patterns of the intestinal microbiota of Iberian lynx, the DNA was amplified using 16S rDNA bacterial domain-specific primers 309F (5′-ATCCCTACGGGAGGCWGCAG-3′) and 677R-CGGGG GGATMTCTACGCATTTCACCGCTAC-3′) [30]. PCR was carried out in a 50 µl reaction mixture that included 1 µl (10 µM) of each primer, 200 µM dNTPs, 5 µl 10X DreamTaq Green Buffer (with MgCl2 20 mM), 37 µl H2O Milli-Q, 5 µl of DNA template and 1 U DreamTaq Polymerase (Life-Technologies, Eugene, Oregon, USA). The standard cycling conditions were 94 ºC for 2 min, followed by 35 cycles at 94 ºC for 30 s, 56 ºC for 30 s and 68 ºC for 30 s and a final step at 68 °C for 5 min. Amplicons obtained (400 bp size) were electrophoresed on a 1% agarose gel (w/v) to check for product size and quantity. Later, amplicons were separated by denaturing gradient gel electrophoresis (DGGE) according Muyzer et al. [42] using a Dcode TM system (Bio-Rad). Electrophoresis was performed in an 8% polyacrylamide gel (37.5:1 acrylamide-bisacrylamide) using a 30 to 55% denaturing gradient (urea and formamide) for separation of PCR products. Electrophoresis was carried out for 5 hours at 150 V in 0.5 X TAE buffer (20 mM Tris acetate [pH 7.4], 10 mM sodium acetate, 0.5 mM Na2-EDTA) at 60 °C constant temperature. Gels were subsequently stained with AgNO3 [48]. The number and intensity of DGGE bands were calculated from the densitometric curves of the gel using the software FPQuest 4.0 (Applied Maths BVBA). A matrix of similarity between DGGE profiles was determined based on Pearson similarity coefficient. Clustering of DGGE patterns was achieved by construction of dendrograms using the unweighted pair group method with arithmetic averages (UPGMA). In order to determine the structural diversity of the microbial community corresponding to the DGGE banding pattern, three indices were calculated: (1) Shannon index (H′) was calculated according to the function H ′ = −Σ  pi * ln pi  , where pi = ni / N , ni is the peak surface of each band, and N is the sum of all the peak surfaces of all bands; (2) Simpson index (D) was calculated following the function D = Σpi2 , where pi = ni / N , ni is the peak surface of each band, and N is the sum of all the peak surfaces of all bands; and (3) Range-weighted richness (Rr) was calculated according to the function Rr = N 2 * Dg , where N represents the total number of bands in the pattern, and Dg the denaturing gradient comprised between the first and the last band of the pattern [38].

Predominant gel bands were retrieved for sequencing with sterile pipette tips, placed in 100 µl of double distilled Milli-Q water and incubated at 4 ºC overnight. DNA was amplified using universal primers, 309F and 677R without GC-clamp (5′-ATMTCTACGCATTTCACCGCTAC-3′). PCR was performed in a 50 µl reaction mixture containing 1 µl (10 µM) of each primer, 200 µM dNTPs, 5 µl 10X DreamTaq Green Buffer (with MgCl2 20 mM), 32 µl H2O Milli-Q, 10 µl of DNA template and 1U DreamTaq Polymerase (Life-Technologies, Eugene, Oregon, USA). The standard cycling conditions were 95 ºC for 2 min, followed by 45 cycles at 95 ºC for 30 s, 64 ºC for 30 s and 72 ºC for 60 s and a final step at 72 ºC for 5 min. The amplicons were used for sequence analysis (Macrogen Korea sequencing) and sequences were compared with those from the National Center for Biotechnology Information (NCBI) using the BLAST sequence algorithm [4].

(

)

Functional metabolic diversity of the fecal microbiota.

Functional diversity of fecal microbiota of Iberian lynx specimens included in clusters I and III was assessed by determining the community catabolic profiles (CLCPs) using Biolog EcoPlate® (Biolog, Hayward, CA, USA). Biolog EcoPlate® assays 31 carbon sources grouped by chemical class (carbohydrates, polymers, carboxylic acids, amino acids and amines) and incorporates

FECAL MICROBIOTA OF CAPTIVE IBERIAN LYNX

a control assay without a carbon source. All tests are carried out in triplicate.

Fecal samples (500 mg) of each specimen from clusters I and III were suspended in 50 ml PBS (pH 7.2) vortexed and allowed to rest for 15 min at 4ºC. Then, 150 µl supernatant aliquots were dispensed into each of the 96 wells of the Biolog Ecoplates plates and incubation was performed at 37 ºC in the dark. Activity measures were carried out at 590 nm with a microplate reader (Multiskan Ascent, Thermo Electron, Milford, MA, USA) every 24 h up to 144 h. To determine the level of functional diversity the following indices were calculated: (a) Shannon’s index as was applied to assess the substrate utilization pattern according to the following expression: H ′ = −Σ  pi * ln pi  , where pi = ni / N , ni is the is the value of OD590 of each well, and N is the sum of OD590 for all metabolic substrates. (b) Substrate richness (S) corresponding to the number of different substrates used.

Statistical analysis. Significance of differences in Shannon’s index (H’), Simpson’s index (D) and range-weighted richness (Rr) was determined after performing t-Student test with the software SPSS Statistics 17.0 (IBM).

Contribution of each bacterial species identified from sequencing of DGGE bands to similarity percentages (SIMPER) between clusters I and III was determined by using PAST software.

Results Clustering analysis was applied to fecal microbiota DGGE bands of Iberian lynx specimens, and the dendrogram obtained showed a high group similarity percentage. Cluster analysis based on Pearson coefficient of similarity between band patterns allowed the identification of clearly defined clusters (I to V) (Fig. 1). Distribution of the specimens included in these clusters did not respond to variables such as gender. Clusters I and III grouped 19 out of 36 specimens sampled, cluster I being the most frequent DGGE pattern detected in lynx from “La Olivilla” (50%). On the contrary, none DGGE pattern of this cluster was detected in specimens from “El Acebuche”. On the other hand, the DGGE patterns grouped in cluster III showed high values of similarity (about 76%) and included 55.55% of the specimens from “El Acebuche”, whilst only 5.55% DGGE patterns of animals from “La Olivilla” were included in this cluster.

The number of DGGE bands was high in all clusters and the ecological index values calculated from the bands and densitometric curves of the gel are summarized in Table 1.

Significant differences (p < 0.05) were not detected. Shannon diversity index (H’) calculated for all lynx specimen samples showed medium-high values (3.05–3.46), whereas low Simpson’s index values (0.05–0.09) were observed indicating low dominance of some microbial species (D). Range-weighted richness (Rr) showed high values (>30) indicating an habitable environment.

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The bacterial species identified based on the bands sequenced from DGGE gels of fecal samples grouped in clusters are summarized in Table 2. Phyla detected in all the samples included Actinobacteria (0.96% to 5.86%), Bacteroidetes (0.11% to 6.03%), Firmicutes (from 33.58% to 42.38%) and Proteobacteria, specifically γ-Proteobacteria, (ranging from 54.95% to 61.76%).

In order to break down the contribution of each bacterial group to the observed dissimilarity between clusters I and III fecal microbiota samples, similarity percentage analysis (SIMPER) was carried out. In this way, most important species responsible for the observed pattern of dissimilarity are summarized in Table 3. Almost 50% of the differences in bacterial communities detected in samples grouped in cluster I and III were attributed to Pseudomonas koreensis (13.9%), Pseudomonas migulae (8.9%), Carnobacterium sp (6.85%), Arthrobacter (5.6%), Robinsoniella peorensis (5.6%) and Ornithinibacillus sp. (5.3%).

Finally, metabolic analysis of fecal microbiota included in clusters I and III showed ability to use 24 and 30 out of 31 carbon sources assayed, respectively (Fig. 2). Catabolic profiles were determined and Shannon diversity values obtained indicate significantly lower (p < 0.05) number of substrates used by microbiota of the cluster I compared to fecal microbiota of cluster III after 24, 48 and 72 h incubation (Table 4).

Discussion Understanding the gut microbial community structure is critical to identify and establish the potential fitness-related traits for the host [56]. In this study, DGGE methodology was used to describe the microbial composition of feces of captive Iberian lynx. The limitations of DGGE in microbial analysis have been previously described [43], nevertheless substantial information about species composition can be obtained from very complex microbial communities such as the gut microbiota [31]. Changes in DGGE band abundance as reflected by bacterial diversity can indicate ecological shifts in the characteristics of bacterial communities [53]. It is commonly accepted that the intensity of the DGGE band is directly related to the density of the corresponding 16S rDNA. For this reason, Pearson similarity coefficient, that considers both position and band intensity, was used in this study to carry out the clustering analysis of the DGGE patterns. Although the same predominant DGGE bands were present in all DGGE pat-

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terns, clearly defined clusters were detected. Clusters I and III included profiles of 19 out of 36 specimens, these clusters displaying the most frequently DGGE patterns detected in

Fig. 1. Clustering analysis based on pair wise similarity index of DGGE patterns obtained from fecal samples of Iberian lynx specimens from “El Acebuche” and “La Olivilla” centers.

lynx specimens from “La Olivilla” and “El Acebuche”, respectively. Cluster III included 55.55% of the specimens from “El Acebuche”, percentage clearly higher than that (5.55%)

Table 1. Percentage of captive Iberian lynx specimens included in the clusters described in this study and Shannon diversity index (H′), Simpson dominance index (D) and range weighted richness (Rr) values obtained for the microbiota of the different clusters. Data of the ecological parameters are expressed as mean ± standard deviation. Significant differences (p < 0.05) were not detected.

CLUSTER I

II

III

IV

V

Specimens from “El Acebuche” (%)

0

5.55

55.55

27.70

11.11

Specimens from “La Olivilla” (%)

50

11

5.55

16.67

16.67

Shannon index (H’)

3.05 ± 0.11

3.13 ± 0.13

3.17 ± 0.07

3.46 ± 0.41

3.19 ± 0.09

Simpson index (D)

0.06 ± 0.02

0.05 ± 0.01

0.05 ± 0.01

0.09 ± 0.03

0.05 ± 0,00

67.18 ± 20.33

61.37 ± 21.58

89.39 ± 31.80

52.06 ± 34.45

83.48 ± 45.45

Ecological parameters

Range-weighted richness (Rr)

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Table 2. Nearest-match identification of 16S rDNA sequences corresponding to microbiota DNA recovered from the DGGE bands of each cluster.

Phylum / Band number

Closest relative

Similarity (%)

Genbank Number

I

II

III

IV

V

Arthrobacter sp. IARI-L-29

99

JF343188

5.86

2.11

2.16

0.96

4.10

Bacteroides xylanisolvens strain EBA22-11

97

JF298887

3.79

0.11

1.92

5.29

6.03

Actinobacteria 23 Bacteroidetes 17 Firmicutes 4

Bacillus psychrodurans strain CBG_LBI34

97

JF909578

2.55

3.24

6.70

7.90

4.46

10

Carnobacterium maltaromaticum

98

AB680942

1.76

3.78

3.94

4.20

2.59

18

Uncultured Carnobacterium sp. clone CTL-18

95

JQ798984

10.26

12.03

6.25

7.97

10.86

25

Clostridium hiranonis strain 45

100

JF693906

3.82

2.17

1.35

0.79

3.17

6

Kurthia zopfii

99

AB680230

1.15

4.32

4.41

2.86

1.75

5

Ornithinibacillus sp. XJSL10-7

98

GQ903476

1.29

1.18

5.98

3.61

2.01

15

Robinsoniella peoriensis strain PPC31

97

NR_041882

6.25

9.19

2.62

3.08

2.49

8

Sporosarcina ureae strain KNUC424

98

JQ071513

2.14

3.03

4.27

5.78

2.93

14

Uncultured bacterium clone TuCw28

90

DQ071463

4.36

3.44

3.08

2.66

3.52

26

Agarivorans sp. B29

97

AB542196

2.50

3.50

1.07

0.62

2.52

9

Escherichia coli strain G5B128

98

GU646167

1.52

3.36

5.76

5.24

2.11

21

Escherichia coli strain DP170

100

JF895181

2.39

2.98

2.04

1.38

5.68

22

Escherichia coli strain ADB-2

99

JX094849

4.09

3.37

1.16

1.69

2.86

13

Escherichia fergusonii strain G30

98

HQ259962

2.24

3.63

3.31

1.23

4.37

16

Escherichia fergusonii strain M5

99

HQ259946

2.60

2.65

4.19

0.85

2.21

7

Escherichia vulneris strain M3

100

HQ259947

1.07

2.56

4.44

3.38

3.14

1

Proteus mirabilis strain FFL2

99

JN222368

1.80

2.93

4.74

4.23

3.87

2

Pseudomonas brenneri strain KOPRI 25949

100

HQ825070

2.52

3.27

5.19

5.98

4.40

27

Pseudomonas koreensis strain J9B-42

99

HQ238778

10.74

2.49

0.94

0.26

3.09

19

Pseudomonas migulae strain A74

97

JN390959

11.81

5.30

6.36

11.50

6.27

12

Pseudomonas psychrophila strain Ibu-08

98

JQ782895

2.01

6.42

3.63

2.71

2.71

20

Pseudomonas tolaasii strain B44

98

EU169145

2.99

3.32

3.36

2.02

3.61

Proteobacteria

3

Shigella flexneri

99

AB639118

2.04

6.56

5.78

7.39

4.46

11

Shigella sonnei strain DY89

99

HQ591457

4.29

5.99

3.43

5.46

2.67

24

Uncultured bacterium clone Tutokochoana-138

96

JQ283151

2.13

3.43

1.94

1.01

1.70

detected in specimens from “La Olivilla”, for which the most frequently DGGE pattern detected corresponded to cluster I. On the contrary, cluster I did not group specimens from “El Acebuche”. The percentage of similarity showed by the DGGE patterns of cluster III was higher than that of profiles included in cluster I, showing higher homogeneity in microbiota composition of specimens grouped in cluster I. Ecosystem function and stability are influenced by species and functional group richness [7] biodiversity being essential in the protection of ecosystems against declines in their func-

tionality [58]. In this context, the number of DGGE bands was high in all clusters and ecological index values calculated from the band patterns did not show significant differences.

Shannon diversity index (H’) showed medium-high values corresponding to the typical profile of intestinal environment [14], with high evenness of the individuals among the bacterial species. Low Simpson’s index (D) values indicate low dominance of some species, whereas range-weighted richness (Rr) values obtained are among those classified as high according to the criteria proposed by Marzorati et al. [38]. These

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Table 3. SIMPER analysis indicating the contribution of each bacterial group to total dissimilarity among microbiota of captive Iberian lynx specimens included in clusters I and III. Bacterial groups contributing ≥ 5% to total dissimilarity are marked in bold Phylum/Band number

Closest relative

Dissimilarity (%)

Actinobacteria 23

Arthrobacter sp. IARI-L-29

5.59

17

Bacteroides xylanisolvens strain EBA22-11

2.99

Bacteroidetes Firmicutes 4

Bacillus psychrodurans strain CBG_LBI34

4.44

10

Carnobacterium maltaromaticum

2.24

18

Uncultured Carnobacterium sp. clone CTL-18

6.85

25

Clostridium hiranonis strain 45

3.73

6

Kurthia zopfii

3.65

5

Ornithinibacillus sp. XJSL10-7

5.33

15

Robinsoniella peoriensis strain PPC31

5.57

8

Sporosarcina ureae strain KNUC424

2.11

14

Uncultured bacterium clone TuCw28

2.40

26

Agarivorans sp. B29

2.21

9

Escherichia coli strain G5B128

4.76

21

Escherichia coli strain DP170

0.90

22

Escherichia coli strain ADB-2

4.32

13

Escherichia fergusonii strain G30

0.82

16

Escherichia fergusonii strain M5

1.37

7

Escherichia vulneris strain M3

3.81

1

Proteus mirabilis strain FFL2

3.15

Proteobacteria

2

Pseudomonas brenneri strain KOPRI 25949

2.69

27

Pseudomonas koreensis strain J9B-42

13.87

19

Pseudomonas migulae strain A74

8.88

12

Pseudomonas psychrophila strain Ibu-08

1.52

20

Pseudomonas tolaasii strain B44

0.16

3

Shigella flexneri

4.04

11

Shigella sonnei strain DY89

1.88

24

Uncultured bacterium clone Tutokochoana-138

0.65

range-weighted richness indicate increased genetic variability of fecal microbiota [13]. Based on these results the intestinal tract of Iberian lynx could be considered as a high habitable environment, hosting species phylogenetically different. This trait could protect this ecosystem against changes and declines in their functioning, because many species provide guarantees that some will maintain functioning even if others fail [58].

However, it is interesting to determine the predominant microbial species present in the feces and gut of Iberian lynx.

In this sense, Alcaide et al. [3] using high-throughput se-

quencing techniques reported the absolute dominance of phylogenetic lineages of Firmicutes, Bacteroidetes besides Fusobacteria, whereas members of Proteobacteria and Actinobacteria were also identified in relatively abundant quantities in the distal gut of Iberian lynx. In our study, sequencing of the predominant DGGE bands of all clusters showed that the prevailing phyla are Firmicutes, Proteobacteria, specifically γ-Proteobacteria class, Bacteroidetes and Actinobacteria, whereas Fusobacteria were not detected as predominant DGGE bands.

In the present work, predominant DGGE bands related to

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Fig. 2. Community metabolic diversity of fecal microbiota of Iberian lynx specimens included in clusters I and III based on the utilization of different carbon sources. Assayed substrates were grouped by chemical class (carbohydrates, carboxylic acids, polymers, amino acids and amines).

classes Clostridia and especially Bacilli were sequenced, similarly to Alcaide et al. [3] who reported these two classes as two of the predominant in the fecal microbiota of Iberian lynx. We also sequenced DGGE bands related to Lachnospiraceae family, specifically Robinsoniella genus. Similarly,

Alcaide et al. [3] also sequenced Lachnospiraceae as one of the largest clostridia families detected in fecal samples of Iberian lynx. However, other bacterial groups such as the family Ruminococcaceae were sequenced by Alcaide et al. [3] but not detected in our study. Differences between these two stud-

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Table 4. Shannon diversity values based on microbiota metabolic abilities of fecal suspensions of captive Iberian lynx specimens included in clusters I and III. Functional diversity was determined with Biolog EcoPlate® at different incubation periods. Data are expressed as mean ± standard deviation. Asterisk denote significant differences (p<0.05) between values of the same row.

Time of incubation (h)

Cluster I

Cluster III

t-Student value

p value

24 h

2,64 ± 0.07

2.97 ± 0.07*

2.22

0.0024

48 h

2.48 ± 0.09

3.20 ± 0.05*

2.22

< 0.0001

72 h

2.58 ± 0.09

3.25 ± 0.04*

2.22

< 0.0001

96 h

2,81 ± 0.08

3.31 ± 0.04*

2.22

< 0.0001

120 h

2.81 ± 0.08

3.39 ± 0.04*

2.22

< 0.0001

144 h

2.86 ± 0.08

3.38 ± 0.03*

2.22

< 0.0001

ies could be attributed to differential sensitivity of the technology used. In this way, DGGE band intensity is directly related to the density of the corresponding 16S rDNA, and it is commonly accepted that only those microbial groups representing ≥1% in terms of relative proportion are displayed in the DGGE patterns [18].

Our results showed the presence of Escherichia genus in the intestine of Iberian lynx, coinciding with those observed

by Gonçalves et al. [19], who demonstrated the presence of β–lactamase-producing E. coli strains in feces of Iberian lynx.

Microorganisms identified as Shigella sonnei and Shigella flexneri were also present in fecal samples analyzed, species described as disease agents of bacillary dysentery affecting humans and primates [12]. The results obtained in our study should be considered for future research related with potential inflammatory diseases affecting to Iberian lynx, as increased counts of mucosa-associated Enterobacteriaceae have been found in duodenal biopsies of cats with intestinal bowel disease (IBD) [23]. Similarly, dogs with IBD showed lower abundance of Bacteroidetes but higher abundance of Proteobacteria in comparison with healthy specimens [51].

In the present work, the presence of Bacteroidetes, in clusters II and III showed the lowest band intensity values. On the other hand, changes in Actinobacteria were mainly due to variations in Arthrobacter genus, which was more abundant in specimens grouped in clusters I and III. However, the level of Actinobacteria and Bacteroidetes in fecal and intestinal microbiota of cats has been reported that is affected by different conditions [15].

The main difference observed between the fecal microbiota of animals from the two breeding centers studied was the frequency of detection of DGGE patterns corresponding to

clusters I and III. In order to analyze the contribution of each bacterial group to the observed dissimilarity between clusters I and III, SIMPER analysis was carried out. The most important species responsible for the dissimilarity between clusters I and III were Pseudomonas koreensis, Pseudomonas migulae, Carnobacterium sp, Arthrobacter, Robinsoniella peorensis and Ornithinibacillus sp. Some P. koreensis strains have been described as spoilage bacteria from meat products [11], and have been isolated from eye lesions in fish [49], but others have also exhibited plant growth promoting traits [6] and ability to produce lipases and biosurfactant substances [5]. Pseudomonas migulae includes strains isolated from soil [33], water [55] and food [17]. Some strains have shown potential effects on plant growth [52] and ability to inhibit foodborne pathogen growth such as E. coli and Salmonella virulent strains [24,57].

On the other hand, Carnobacterium genus consists of species, mainly C. divergens and C. maltaromaticum, commonly found in the environment [28], foods [8] and intestinal tract of insects [50] and fish [2]. This microbial group is able to catabolize multiple carbohydrates [32] some species, such as C. maltaromaticum, showing chitinase activity [32] and being able to grow in meat at low temperature [26]. In addition, some Carnobacterium strains have the ability to produce bacteriocins [20] capable to inhibit pathogenic microorganisms [16] and to improve fish immune response [29]. These events have led to propose Carnobacterium strains as probiotics in aquaculture [44].

Ornithinibacillus species include aerobic, rod-shaped, motile and endospore-forming bacteria that have been isolated from several environments such as sludge compost [35], clams [57], Artic terrains [60], pasteurized milk [39] and hu-

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man blood [27]. On the other hand, the genus Arthrobacter includes a heterogeneous group of obligate aerobic species commonly found in soil, but also isolated from human clinical specimens [36].

The anaerobic and spore-forming genus Robinsoniella has been identified as the most abundant member of Firmicutes in wild mice [56], suggesting an important biological role [56].

In this context, changes in the level of this genus have been associated with colitic process in mice [59] and in nonalcoholic fatty liver disease patients [46]. The presence of enterotypes in several mammalian hosts or distinct clusters characterized by the abundance of signature bacterial genera has been proposed [21,40]. Wang et al. [56] described two clusters in the intestinal microbiome of wild mice with differences in bacterial taxons such as members of genus Robinsonella. It would be very suggestive to think that the results obtained in our study support the existence of enterotypes in the microbiome of Iberian lynx, although further studies are needed to confirm this findings.

Fecal microbiota of clusters I and III were able to use 24 and 30 out of 31 different carbon sources (carbohydrates, polymers, carboxylic acids, amino acids and amines), respectively. These results reflect high functional diversity of fecal microbiota, as they are able to metabolize most different substrates assayed in Biolog EcoPlates®. However, this high functional diversity was not equal along the incubation time.

Thus, the fecal microbiota of cluster III showed ability to use higher number of carbon substrates���������������������������� earlier than cluster I, especially amino acids and carboxylic acids, at 24, 48 and 72 h.

These differences could be related to the fact that Biolog EcoPlates® aerobic incubation carried out shows functional diversity of aerobes and facultative anaerobes only. In this way, the intensities of the DGGE bands corresponding to anaerobes such as Bacteroidetes and Firmicutes such as Bacteroides xylanisolvens, Clostridium hiraronis and Robinsoniella peorensis were higher in fecal microbiota from cluster I than in cluster III. This different microbiota composition could explain the delay in substrate metabolization observed. However, it is necessary consider that the environmental conditions of “El Acebuche” and “La Olivilla” are not identical. Thus, although time span between defecation and freezing of the feces was short, different environmental conditions in these areas could have some effect, which may be considered for future research.

This study is the first report comparing the composition of bacterial fecal microbiota of Iberian lynx specimens from two

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breeding centers of captive Iberian lynx where they are confined. Results obtained show that the microbiota present in the feces could indicate that the intestinal tract of Iberian lynx is an environment very habitable and with high biodiversity of microbial species. Changes in the gut microbiota are associated with far reaching consequences on host health and development. For this reason, the understanding of the gut microbiota may lead to targeted interventions for health promotion, disease prevention and management through tools such as diet and functional foods (with pre and probiotics), contributing for the future conservation of the Iberian lynx.

Acknowledgements. The authors thank the Consejería de Medio Ambiente del Gobierno de la Junta de Andalucía and the Iberian lynx breeding centers of “El Acebuche” and “La Olivilla”, for providing feces from the captive Iberian lynx specimens.

Competing interests: None declared.

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RESEARCH ARTICLE International Microbiology 20(1): 43-53 (2017) doi:10.2436/20.1501.01.284. ISSN (print): 1139-6709. e-ISSN: 1618-1095

www.im.microbios.org

Antimicrobial peptide from Bacillus subtilis CSB138: characterization, killing kinetics, and synergistic potency Sudip Regmi,1§ Yoon Seok Choi,1§ Yun Hee Choi,1 Young Kyun Kim,1 Seung Sik Cho,2 Jin Cheol Yoo,1* Joo-Won Suh3* Department of Pharmacy, College of Pharmacy, Chosun University, Gwangju, Korea. 2Department of Pharmacy, Mokpo National University, Muan, Jeonnam, Korea. 3Center for Nutraceutical and Pharmaceutical Materials, Myongji University, Myongji-ro, Cheoin-gu, Yongin, Gyeonggi-Do, Korea

1

Received 8 March 2017 · Accepted 30 March 2017 Summary. We studied the prospect of synergy between the antimicrobial peptide p138c and non-peptide antibiotics for increasing the potency and bacterial killing kinetics of these agents. The production of p138c was maximized in the late exponential growth phase of Bacillus subtilis CSB138. Purification of p138c resulted in a total of 4800 arbitrary units (AU) with 19.15-fold and 3.2% recovery. Peptide p138c was thermo-tolerant up to 50 °C and stable at pH 5.8 to 11. The biochemical nature of p138c was determined by a bioassay, similar to tricine-SDS-PAGE, indicating inhibition at 3 kDa. The amino acid sequence of p138c was Gly-Leu-Glu-Glu-Thr-Val-Tyr-Ile-Tyr-Gly-Ala-Asn-Met-X-Ser. Potency and killing kinetics against vancomycin-resistant Staphylococcus aureus improved considerably when p138c was synergized with oxacillin, ampicillin, and penicillin G. The minimal inhibitory concentration (MIC) of p138c showed a 4-, 8-, and 16-fold improvement when p138c was combined with oxacillin, ampicillin, and penicillin G, respectively. The fractional inhibitory concentration index for the combination of p138c and oxacillin, ampicillin, and penicillin G was 0.3125, 0.25, and 0.09, respectively. Synergy with non-peptide antibiotics resulted in enhanced killing kinetics of p138c. Hence, the synergy between antimicrobial peptide and non-peptide antibiotics may enhance the potency and bacterial killing kinetics, providing more potent and rapidly acting agents for therapeutic use. [Int Microbiol 20(1):43-53 (2017)] Keywords: Bacillus subtilis · antimicrobial peptides · killing kinetics

Introduction The wide use of antibiotics, in the past and recent decades, has caused the rapid emergence of antibiotic-resistant bacteria, indicating the need for the development of new antimicrobial Both authors contributed equally to the article.

§

*

Corresponding author: Jin Cheol Yoo E-mail: jcyu@chosun.ac.kr *Additional coresponding author: Jon-Won Suh E-mail: jwsuh@mju.ac.kr

agents. Antimicrobial peptides (AMPs), present in the host immune system, may be a new, anti-infective alternative to conventional antibiotics. Because of their broad-spectrum antimicrobial activity and exclusive membrane action mechanism, they may replace or accompany conventional antibiotics [26].

Therefore, AMPs can be used to develop future antibiotics. To date, approximately 1000 naturally occurring antibacterial peptides have been isolated. Despite this large number of iso-

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lated peptides, only a few have clinical applications because most of them display poor potency, specificity, and in vivo stability [5]. These drawbacks of AMPs need to be addressed in order to achieve effective development and application of new antimicrobial agents for clinical use. Hence, efforts have been placed into enhancing the killing kinetics, potency, and specificity of antibacterial peptides.

The activity and selectivity of AMPs are driven by their charge, amphipathicity, and hydrophobicity [4,25]. To address these factors, short AMP derivatives, with cell selective toxicity, have been developed to improve the activity and specificity based on the amino acid composition, charge, and hydrophobicity of natural peptides [10,13,15,21,27]. Toxicity has been reduced by minor sequence modifications [2,6,14]; however, such approaches are time consuming and costly.

The use of combination therapy is effective in overcoming antibiotic resistance. Employing β-lactamase inhibitors as codrugs with conventional antibiotics is well known [11,16].

The synergistic effect of antibiotic activity depends on the ability of the two molecules to exert a deleterious effect on the target microorganism higher than the sum of the effects of each drug alone. Synergy reduces the MIC of the combined drugs and helps prevent the development of resistance in microorganisms [16,24].

The widespread use of antibiotics plays a major role in the emergence of drug-resistant bacteria. Unhealthy practices in the pharmaceutical and manufacturing industries are also likely sources of the emergence of antibiotic-resistant strains [9].

Staphylococcus aureus and Enterococcus faecium are the major antibiotic-resistant pathogens found in hospitals. Methicillin-resistant Staphylococcus aureus (MRSA), detected for the first time in 1961 in Britain, is now common in hospitals. The first strain with complete resistance (>16 µg/ml) to vancomycin, termed vancomycin-resistant Staphylococcus aureus (VRSA), appeared in the United States in 2002. Vancomycinresistant Enterococcus (VRE) was reported in 1987. A report in 2004 showed that 9% of nosocomial bloodstream infections were caused by enterococci and 2% by the E. faecalis strain [23]. This rapid emergence of resistant bacteria highlights the need to find alternatives for conventional antibiotics.

In this study, we isolated, identified, and characterized a bacterial isolate from fermented food using gene sequencing.

Further, we studied the possible synergy between the antimicrobial peptide and non-peptide antibiotics. We have identified isolates with promising antimicrobial profiles and evaluated their biological potency. The combination of antimicro-

REGMI ET AL.

bial peptide and non-peptide antibiotics showed effective synergy, which we evaluated using fractional inhibitory concentration (FIC) indices. Our antimicrobial peptide was moderately potent and synergistic with commonly used antibiotics; it also showed robust potency and killing kinetics.

Materials and methods Materials. This The chemicals and solvents used in our study were of extra pure grade. Sepharose CL-6B and Sephadex G-50 were obtained from Pharmacia (Uppsala, Sweden). All other reagents were of the highest analytical grade available.

Bacterial strain and production of the antimicrobial peptide. More than 100 fermented food samples were used to isolate the bacterial strain. Isolation was performed according to the method described previously [16]. Briefly, 1g of the fermented food sample was mixed with 0.85% NaCl and incubated for 24 h at 37 °C. Following incubation, the sample was serially diluted up to 10−7 in Mueller-Hinton broth. The appropriate colonyforming units (CFU/ml) were estimated by streaking the dilutions on Mueller-Hinton agar media. The dilution with the required CFU/ml was considered working stock and maintained in 20% glycerol. The bacterial strain CSB138, with potent antimicrobial activity, was selected for further study.

The identification of the bacterial strain, based on morphological characteristics, was performed according to Bergey’s Manual of Systematic Bacteriology. Sequencing analysis of the 16S ribosomal RNA (rRNA) gene was conducted for further identification. Antimicrobial peptide production was performed using a bacterial culture grown in optimized medium for 28 h at 37 °C in a rotating incubator at 150 rpm.

Media optimization. Optimization of the culture media for strain CSB138 was conducted using various carbon and nitrogen sources, and metal ions. The carbon sources included mannitol, starch, fructose, sorbitol, sucrose, glucose, maltose, and lactose. The nitrogen sources included yeast and beef extract, malt, tryptone, peptone, oatmeal, and soymeal. The metal ions were Na2HPO4, NaH2PO4, MgSO4, ZnSO4, MgCl2, KH2PO4, FeSO4, NaCl, and CaCl2. The influence of carbon sources on the production of the antimicrobial compound was assessed using media supplemented with 1% yeast extract as a nitrogen source and various carbon sources. Fermentation was carried out in 250-ml Erlenmeyer flasks at 150 rpm and 37 °C. The influence of nitrogen sources on the production of the antimicrobial compound was assessed using a medium supplemented with 1% sorbitol as a carbon source and various nitrogen sources. The influence of metal ions on the production of the antimicrobial compound was determined using 1% sorbitol, 1% yeast extract, and various metal ions. The influence of sorbitol, yeast extract, and metal ions on the production of the antimicrobial compound by strain CSB138 was determined using different proportions of these supplemented compounds. The bacterial strain CSB138, cultured in fully optimized media containing 1.25% sorbitol and 1% yeast extract (S-YE), was optimal for the production of the antimicrobial compound. The antimicrobial compound, produced by strain CSB138, was designated as p138c.

Kinetics of bacterial growth. The S-YE fully optimized mediumwas inoculated with freshly grown Bacillus strain CSB138 and incubated at 37 °C. Samples (2 ml) were collected at the interval of 4 h and centrifuged

ANTIMICROBIAL PEPTIDE FROM B. SUBTILIS

at 10,000 ×g for 15 min. The obtained supernatant was assayed for antimicrobial activity. Optical density was recorded spectrophotometrically at 620 nm and pH was measured.

Purification of p138c. The seed culture, with the bacterial suspension turbidity equivalent to that of 0.5 McFarland standard solutions, was transferred into the main culture in a 2-liter baffled flask, containing 400 ml S-YE media, and incubated at 37 °C and 150 rpm for 28 h. Then, the fermented broth was centrifuged (Hanil Science Industrial, Supra 22K, Korea) at 10,000 × g at 6 °C for 15 min. The obtained cell-free supernatant was treated with diammonium sulfate at the saturation of 20–100%. The active precipitation fraction was recovered using centrifugation at 10,000 × g at 6 °C for 45 min, dialyzed with 10 mM/l Tris-HCl (pH 7.5) buffer, and subjected to ultrafiltration (Millipore). After ultrafiltration through the molecular weight cut-off membranes, the biologically active fraction was loaded onto a Sepharose CL-6B column (2.5 × 85 cm) (Pharmacia, Uppsala, Sweden) using 10 mM/l Tris-HCl (pH 7.5) buffer. The active fractions, obtained from Sepharose CL-6B gel permeation column chromatography, were pooled, concentrated, further purified with a Sephadex G-50 column (1.7 × 120 cm) (Pharmacia, Uppsala, Sweden) using the same buffering system, and subjected to purity analysis.

Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis and bioassay. The concentration of p138c was determined by the Bradford method [1] using bovine serum albumin (BSA) as the standard. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis (Tricine-SDS-PAGE) [17] was used to analyze the purity of the antimicrobial peptide. The electrophoresed gel was stained with Coomassie brilliant blue R-250 and destained with methanol/glacial acetic acid/distilled water (1:1:8, v/v/v). The inhibitory activity was assayed according to our previously described method [16]. Briefly, the electrophoresed gel was washed several times with 50 mM/liter Tris-HCl buffer (pH 7.5) containing 2.5% Triton X-100. Afterward, the processed gel was overlaid with 0.6% agar on MuellerHinton medium (Difco, USA) containing the indicator organisms (1.5 × 108 CFU/ml) and incubated at 37 °C for 12 h.

The N-terminal amino acid sequence. The N-terminal amino acid sequence of p138c was determined by Edman degradation using the automatic protein sequencing system model 492cLC (Applied Biosystems, Foster, CA, USA).

The effects of pH, temperature, proteases, and surfactant on p138c. The effect of temperature on the activity of p138c was evaluated by incubating this compound at the temperature of 0, 4, 10, 20, 37, 50, 60, 70, 80, and 121 °C for 24 h and assessing antimicrobial activity using the disc diffusion method. The effect of pH on the activity of p138c was tested by adjusting the pH 4.2 to 12.5 using 1 M HCl or 1 M NaOH, respectively, and assessing antimicrobial activity. In a separate procedure, 10 ml of cell-free supernatant were placed into a sterile Petri dish and exposed to UV light in a laminar airflow hood for 5 min; then, antimicrobial activity was examined using the disc diffusion method.

The effect of various proteases on the activity of p138c was examined using 50 µl of cell-free supernatant incubated for 150 min in the presence of 1 mg/ml or 0.1 mg/ml of trypsin, proteinase K, and catalase. Antimicrobial activity was determined using the disc diffusion method. In a separate experiment, Triton X-100, Tween 20, Tween 80, SDS, NaCl, and urea were added to p138c to yield the final concentrations of 1.0, 2.0, and 5.0 mM. Untreated p138c and the tested reagents, at their respective concentrations in distilled water, were used as controls. All the treated and untreated sam-

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ples were incubated at 37 °C for 6 h, after which the antimicrobial activity was examined using the disc diffusion method.

Antimicrobial inhibitory spectrum. Antimicrobial activity was determined by the disc diffusion method. A disc of filter paper (8 mm, Toyo Roshi Kaisha, Japan), saturated with 40 µl of p138c, was placed on the surface of a Mueller-Hinton agar plate, overlaid with indicator microorganisms, and incubated at 37 °C for 24 h. Following incubation, the clear zone of inhibition around the paper disc was measured. Antimicrobial activity, with respect to minimal inhibitory concentration (MIC), was determined according to our previous report [16]. Various microorganisms, including Bacillus sub­ tilis ATCC 6633, Enterococcus faecalis ATCC 29212, Mycobacterium smeg­ matis ATCC 9341, methicillin-resistant Staphylococcus aureus 5–3, methicillin-resistant S. aureus 4–5, vancomycin-resistant S. aureus, vancomycin-resistant Enterococci 4, vancomycin-resistant Enterococci 98, vancomycin-resistant Enterococci 89, S. aureus KCTC 1928, Micrococcus luteus ATCC 9341, Escherichia coli KCTC 1923, Salmonella typhimurium KCTC 1925, Pseudomonas aeruginosa KCTC 1637, extended-spectrum beta-lactamase V4 (E. coli), and Alcaligenes faecalis ATCC 1004 were used as test microorganisms. Vancomycin and bacitracin were used as reference antibiotics. After the inoculation of test organisms (1.5 × 108 CFU/mL), the plates were incubated at 37 °C overnight. Minimum bactericidal concentration (MBC) was determined using the broth dilution method. Antimicrobial activity was expressed in terms of arbitrary units (AU). An AU is defined as the highest dilution of the sample that produces a zone of inhibition. The reciprocal of the dilution was regarded as the titer of antimicrobial activity (AU/ml).

Assessment of antibiotic susceptibility. Susceptibility testing of the activity of p138c and various antibiotics (oxacillin [Sigma, USA], ampicillin [Sigma], penicillin G [USB, UK], ciprofloxacin [Sigma], bacitracin [USB] and vancomycin [Sigma]) against vancomycin-resistant Staphylococ­ cus aureus (VRSA), vancomycin-resistant Enterococci 4, vancomycin-resistant Enterococci 89, and vancomycin-resistant Enterococci 98 was conducted using the broth dilution method for the determination of MIC [22]. The bacterial culture (1.5 × 108 CFU/ml), grown overnight, was suspended in MuellerHinton broth; 100 µl of this suspension was aliquoted into a 96-well tissue culture plate (TCP). Various concentrations (0.156-80 µg/ml) of the antimicrobial peptide in 10 mM/l Tris-HCl (pH 7.5) buffer, and antibiotics in distilled water, were added to the suspension. The negative control was maintained without the addition of any antibiotics. The TCP was incubated at 37 °C in a shaking incubator at 150 rpm. After 12 h of incubation, optical density was measured at 620 nm. MIC is defined as the lowest concentration of any drug that inhibits the measurable growth of the indicator organism after a given incubation. Each experiment was performed in triplicate and repeated three times. Minimum bactericidal concentration (MBC) was measured by plating the content of the clear well, used in the MIC assessment, onto MuellerHinton agar plates. After the incubation, MBC was determined as the lowest concentration that inhibited bacterial growth (99.9%) on the surface of the agar.

Bacterial killing kinetics. The cultures (1.5 × 108 CFU/ml) of vancomycin-resistant Staphylococcus aureus (VRSA), vancomycin-resistant Enterococci 4 (VRE 4), vancomycin-resistant Enterococci 89 (VRE 89), and vancomycin-resistant Enterococci 98 (VRE 98), grown overnight, were diluted in Mueller-Hinton broth; antibiotics, along with the antimicrobial peptide p138c, were added to 100 µl of the diluted culture. The mixture was incubated at 37 °C and 150 rpm. The samples were collected at regular intervals, diluted, and plated on Mueller-Hinton agar plates. The plates were incubated at 37 °C for 18 h and the colonies were counted. For synergy assessment, MICs were determined by the addition of 0.5 MIC.

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Fig. 1. Phylogenetic tree, based on the complete 16S rRNA gene sequence, showing the relationships between the strain CSB138 and closely related taxa of the genus Bacillus. Reference sequences were retrieved from GenBank under the accession number indicated in parentheses after the strain name. Numbers of nodes are percentage bootstrap values based on 1000 replications; only values greater than 65 % are shown.

Bar: 0.005 substitutions per nucleotide position.

The checkerboard dilution assay for the assessment of synergy. The assessment of synergy was performed using the checkerboard method [7]. Separate antibiotics, and combinations of two antibiotics, were incubated with bacteria to observe the bactericidal effect. The MIC of combined antibiotics was calculated using the addition of 0.5 MIC of the antimicrobial peptide. The fractional inhibitory concentration (FIC) assay was used as the base for synergism. The fractional inhibitory concentration index (ΣFIC) was estimated according to our previous method [16]. Briefly, ΣFIC = FICA + FICB = (CA/MICA) + (CB/MICB), where MICA and MICB are the MICs of drugs A and B, and CA and CB are the concentrations of the combined drugs, respectively. The interaction was recorded, according to the guidelines provided by the European Committee on Antimicrobial Susceptibility Testing, as follows: FIC index ≤ 0.5 indicated synergy, FIC index > 0.5–1.0 indicated additive action, FIC index > 1.0 to ˂ 2.0 indicated indifference, and FIC index ≥ 2.0 indicated antagonism [3].

Results Identification of the bacterial strain. Strain CSB138 showed morphological resemblance to Bacillus. The gene sequencing analysis, obtained by gene sequencing 16S rRNA, indicated that the isolate showed the closest identity to Bacillus subtilis subsp. inaquosorum KCTC 13429 (accession no. AMXN01000021) with pairwise similarity of 99.93 %.

Based on similarities in morphology and gene sequence, our strain was classified as Bacillus subtilis CSB138. A phylogenetic tree, constructed from the 16S rRNA gene sequence, is shown in Fig.1.

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Table 1. Minimum inhibitory concentration of peptide p138c Antimicrobial spectrum of p138c MIC (µg/ml) p138c

Bacitracin

MBC (µg/ml) Vancomycin

p138c

Gram-negative bacteria Alcaligenes faecalis ATCC 1004

>80

>80

>80

>1280

Escherichia coli KCTC 1923

>80

>80

>80

>1280

Extended-spectrum beta-lactamase V4 (Escherichia coli)

>80

>80

>80

>1280

Pseudomonas aeruginosa KCTC 1637

>80

>80

>80

>1280

Salmonella typhimurium KCTC 1925

>80

>80

>80

>1280

Bacillus subtilis ATCC 6633

>80

20

0.312

640

Enterococcus faecalis ATCC 29212

2.5

5

2.5

160

Methicillin-resistant Staphylococcus aureus 4-5

0.625

2.5

0.625

640

Methicillin-resistant Staphylococcus aureus 5-3

0.625

2.5

1.25

640

Micrococcus luteus ATCC 9341

80

40

2.5

320

Mycobacterium smegmatis ATCC 9341

40

80

2.5

>1280

Staphylococcus aureus KCTC 1928

20

40

1.25

320

Vancomycin-resistant Staphylococcus aureus

20

80

>80

640

Vancomycin-resistant Enterococci 4

10

40

>80

160

Vancomycin-resistant Enterococci 89

10

>80

>80

640

Vancomycin-resistant Enterococci 98

10

>80

>80

640

Gram-positive bacteria

MIC, minimum inhibitory concentration; MBC, minimum bactericidal concentration

Culture media and production of the antimicrobial peptide. Microorganisms require specific biological culture media in order to grow under laboratory conditions.

Bacteria, grown in the laboratory culture media, are designed to meet all the requirements necessary for confluent growth.

However, the production of specific protein viz. antimicrobial peptide is based on optimized media. Among the carbon and nitrogen sources tested, sorbitol and yeast extract, respectively, were more effective. Metal ions did not significantly affect the production of p138c. Hence, the optimal medium that we used was supplemented with 1.25% sorbitol and 1% yeast extract (S-YE). The Bacillus isolate CSB138, producing p138c, was grown on S-YE medium at 37 °C for 28 h and 150 rpm.

p138c was detected starting at 4 h at pH 6.34, and maximal activity was recorded after 28 h at pH 7.1. The changes in the pH of the culture, from 6.25 to 8.38, were observed for up to 60 h of growth. We recorded increases in the cell density of the cultures at OD 620 nm, in the pH range of 6.34 to 8.36, with maximal density at pH 8.01 and maximal activity at 7.1. In this study, we evaluated the activity of p138c against various microorganisms. The antimicrobial activity of p138c was evaluated along with the standard reference antibiotics viz. vancomycin and bacitracin. The antimicrobial spectrum of p138c was 4 to > 8 times greater than that of bacitracin, and > 4 to > 8 times more effective than that of vancomycin, against vancomycinresistant Staphylococcus aureus and various VREs (Table 1).

Spectra of p138c. In the growth kinetics study, the maximal activity of p138c was observed after 28 h in the S-YE broth. The initial pH of the broth was 6.25. The activity of

Size exclusion chromatography and molecular weight determination of p138c. At the third step of purification, the biologically active sample was loaded onto a

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Fig. 2. Elution profile of peptide p138c from the Sepharose-CL-6B column (2.5 × 85 cm). The column was pre-equilibrated with 10 mM/l Tris/HCl buffer (pH 7.5).

Sepharose CL-6B and fractionated at the elution rate of 0.25 ml per min; the fraction eluted between 590 to 660 min showed maximal activity (Fig. 2). The biologically active fractions were collected and loaded onto Sephadex G-50 using the same buffering system. The active fractions were pooled together and desalted using a Sephadex G-25 column. The protein profile was determined using tricine SDS-PAGE, which showed a single protein band near 3 kDa. The inhibitory activity of p138c was examined in situ. Using an indicator microorganism, a clear zone of lysis was observed near 3 kDa. Purification resulted in a total of approximately 4800 AU with 19.15-

fold overall purification and recovery of 3.2% (Table 2).

Characterization. Residual activity was determined according to the retained biological activity. The activity of p138c was retained at 100% after heating at temperatures as high as 50 °C. Complete loss of activity occurred after heating at 121 °C and 105 kPa for 15 min. Peptide p138c tolerated a wide range of pH values (5.8 to 11). This ability to retain activity over a wide range of pH values may be the result of the reversible denaturation of the protein. The effects of temperature and pH on the activity of p138c are shown in Fig. 3. The

Table 2. Purification of peptide p138c Steps

Total protein (mg)

Total activity (AU)

Specific activity (AU/mg)

Purification fold

Recovery %

Crude

308.56

150000

486.12

1

100

Ammonium sulfate pallet

91.59

73600

803.60

1.65

49.1

Sepharose CL-6B

5.79

12000

2071.88

4.26

8

Sephadex G-50

0.52

4800

9310.50

19.15

3.2

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antimicrobial activity of p138c was completely inactivated after the cell-free supernatant was treated with proteinase K. The effect of trypsin was not detectable, indicating the proteinaceous nature of our antimicrobial peptide. Generally, antimicrobial peptides have proteinaceous characteristics. Our results show that H2O2 was not the factor responsible for the inhibition because no change in activity was recorded when p138c was treated with catalase. Peptide p138c was found to be sensitive to treatment with SDS, Tween 80, and NaCl; however, treatment with Triton X-100, Tween 20, and urea had no effect on the antimicrobial activity of p138c. UV treatment did not affect the antimicrobial activity, possibly because of the proteinaceous nature of p138c. The effects of enzymes, detergents, other reagents, and UV- light on the activity of p138c are shown in Table 3.

Amino acid sequence. The N-terminal sequence of the last 15 amino acids of p138c was determined to be Gly-LeuGlu-Glu-Thr-Val-Tyr-Ile-Tyr-Gly-Ala-Asn-Met-X-Ser. This sequence did not show significant homology with those of peptides of similar origin. No full query is covered during the search. Searching the National Centre for Biotechnology Information (NCBI) protein database using BLAST showed some similarity with other protein sequences from the Bacillus sp.

(Table 4).

49

Table 3. Effect of proteases, detergents, UV, and other reagents on the activity of p138c Proteases

Detergents

NaCl and urea

UV

Trypsin

ND

Proteinase K

+

Catalase

−

SDS

+

Triton X-100

−

Tween 20

−

Tween 80

+

NaCl

+

Urea

−

UV-light

−

ND: non-detectable; + inactivation by treatment; − resistant to treatment

monly used bacitracin and vancomycin. Peptide p138c showed higher antibacterial activity against vancomycin-resistant Staphy­lo­coccus aureus (MIC of 20 µg/ml, 4-fold greater than bacitracin and > 4-fold greater than vancomycin, and MBC of 640 µg/ml), vancomycin-resistant Enterococci 4 (MIC of 10 µg/ ml, 4-fold greater than bacitracin and > 8-fold greater than vancomycin, and MBC of 160 µg/ml), vancomycin-resistant Enterococci 89 (MIC of 10 µg/ml, > 8-fold greater than bacitracin and vancomycin, and MBC of 640 µg/ml), and vancomycin-resistant Enterococci 98 (MIC of 10 µg/ml, > 8-fold greater than bacitracin and vancomycin, and MBC of 640 µg/ml). The synergy between p138c and six conventional antibiotics was tested in vitro by the checkerboard dilution assay against VRSA and three vancomycin-resistant isolates of Enterococci (Table 5). Each combination of the drugs was assigned a ΣFIC. FIC indices

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Synergy between p138c and conventional antibiotics. The antimicrobial activity of p138c, determined in terms of MIC, was moderately active against gram-positive bacteria (MIC of 0.625–80 µg/ml). The antimicrobial spectrum of p138c possessed more potency than those of the com-

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Fig. 3. Effect of pH (A) and temperature (B) on the activity of peptide p138c.

50

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REGMI ET AL.

Table 4. Comparison of N-terminal amino acid sequence of peptide p138c with other related proteins from the Bacillus sp.

SN

Alignment

Identity

Peptide or protein

References

1

GLEETVYIYGANMXS

100 %

p138c [Bacillus subtilis]

Current study

2

EET+Y YGA EETIYFYGA

78 %

peptidase M16 [Bacillus subtilis]

NCBI Reference Sequence: WP_060399736.1

3

+ET YIY GA DETRYIYTGA

70 %

peptidase M20 [Bacillus subtilis]

NCBI Reference Sequence: WP_060399755.1

4

+ET YIY GA DETRYIYTGA

70 %

protein RocB [Bacillus subtilis]

NCBI Reference Sequence: WP_003235954.1

≤ 0.5 were regarded as synergistic. Peptide p138c showed synergy with the three β-lactam antibiotics (oxacillin, ampicillin, and penicillin G) used for inhibiting bacterial growth (VRSA). p138c, combined with oxacillin or ampicillin, or with penicillin G, substantially enhanced the potency of the combinatorial effect. The antibacterial activity of combined treatment with oxacillin and p138c against VRSA was 4-fold greater than that of p138c alone. Similarly, combining p138c with ampicillin produced antimicrobial activity that was 8-fold greater than that of p138c alone; the antimicrobial activity of p138c, combined with penicillin G, was 16-fold greater than that of p138c alone. These results show that synergizing p138c with β-lactam antibiotics is an effective strategy for improving antibacterial potency and cost effectiveness of antimicrobial peptides.

Effect of synergy on bactericidal activity. Drugs are categorized into bacteriostatic and bactericidal agents based on their effectiveness against infectious microorganisms. Bacteriostatic drugs inhibit the growth of bacteria, whereas bactericidal drugs kill them. We evaluated the minimum bactericidal concentration of the synergistic combination of p138c (MBC, 640 µg/ml) and oxacillin (MBC, 80 µg/ml), p138c (MBC, 640 µg/ml) and ampicillin (MBC, 20 µg/ml), and p138c (MBC, 640 µg/ml) and penicillin G (MBC, 40 µg/ml) against vancomycin-resistant Staphylococcus aureus. The combination of p138c (combined MBC, 160 µg/ml) and oxacillin (combined MBC, 5 µg/ml), p138c (combined MBC, 80 µg/ml) and ampicillin (combined MBC, 2.5 µg/ml), and p138c (combined MBC, 80 µg/ml) and penicillin G (combined MBC, 1.25 µg/ml) were bactericidal. The fractional inhibitory concentration (FIC) index for the combination of p138c and oxacillin was low (0.3125). Similarly, the FIC in-

dex for the combination of p138c and ampicillin was 0.25 and that of p138c and penicillin G was 0.09. FIC indices ˂ 0.5 indicate a strong synergy with respect to bactericidal activity.

Thus, oxacillin, ampicillin, or penicillin G potentiates bactericidal activity, demonstrating 4 to 8-fold improvement in the bactericidal action of p138c. Our results indicate that synergy with conventional antibiotics can transform a moderately bacteriostatic agent into a potent bactericidal drug.

The effect of synergy on bacterial killing kinetics. The bacterial load clearance in patients corresponds with how well antibiotics perform in killing the targeted bacterium. We tested the synergistic effect of p138c (MIC of 20 µg/ml) and various commonly used conventional antibiotics (oxacillin, MIC of 80 µg/ml; ampicillin, MIC of 20 µg/ml; and penicillin G, MIC of 40 µg/ml) against vancomycin-resistant Staphylococcus aureus to find a synergistic combination of the peptide and non-peptide antibiotics that affected the rate of bacterial killing kinetics. Peptide p138c alone was largely bacteriostatic against VRSA; in synergy with various tested antibiotics, p138c showed improved potency and more rapid killing kinetics (Fig. 4). Synergizing the peptide with commonly used antibiotics improved the potency of the antimicrobial peptide and the rate of bacterial killing kinetics.

Discussion Peptide p138c, produced by Bacillus subtilis CSB138, was isolated from fermented food. According to Selhub et al.

[18], the relationship between bioactive compounds in fermented food and fermentation-enriched chemicals affects the profile of the human intestinal microbiota, indicating that the

ANTIMICROBIAL PEPTIDE FROM B. SUBTILIS

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Table 5. Fractional inhibitory concentration (FIC) of peptide p138c Tested compounds

Oxacillin (OXA)

Ampicillin (AMP)

Penicillin G (PENG)

Ciprofloxacin (CIP)

Bacitracin (BAC)

Vancomycin (VAN)

Strains

MIC of p138c (Âľg/ml)

MIC in combination with p138c (Âľg/ml)

EFIC

Interpretation

VRSA

20

5

0.31

Synergy

VRE 4

10

5

0.63

VRE 89

10

10

<1.125

VRE 98

10

5

0.56

VRSA

20

2.5

0.25

Synergy

VRE 4

10

2.5

0.50

Synergy

VRE 89

10

5

0.63

VRE 98

10

5

0.75

VRSA

20

1.25

0.09

VRE 4

10

5

0.75

VRE 89

10

10

1.50

VRE 98

10

5

0.63

VRSA

20

5

2.25

VRE 4

10

5

1.50

VRE 89

10

5

4.50

VRE 98

10

2.5

1.25

VRSA

20

>80

>5

VRE 4

10

80

10

VRE 89

10

80

<9

VRE 98

10

>80

>9

VRSA

20

>80

>5

VRE 4

10

>80

>9

VRE 89

10

>80

>9

VRE 98

10

>80

>9

purified product from fermented food may alter the pre-consumption of dietary products. This ultimately amends the action by which fermentation-enriched chemicals act on the human intestinal microbiota. It is known that the antimicrobial peptides have functional activities including antimutagenic, anticancer, immunomodulatory, antiatherosclerotic, and anti-obesity effects. In our study, p138c inhibited a wide spectrum of bacteria and demonstrated antagonistic potential.

The lower potency, instability, and high production cost of antimicrobial peptides have caused major hurdles in their clinical development [5]. One of the solutions to these issues is synergizing antimicrobial peptides with conventional antibiotics, thereby lowering the dosage of each molecule in the synergistic combination.

Our study was designed to characterize p138c and study the prospect of synergy between the antimicrobial peptide and non-peptide antibiotics. The strain, isolated from fermented

Synergy

food, was identified via morphological and 16S rRNA gene sequencing analyses. p138c can be produced in inexpensive optimized culture media (S-YE). Generally, bacteriocin-like substances are produced in complex media [12]. The purification of p138c was conducted using sequential gel filtration, resulting in the eluted compound that was free of unwanted proteins and possessing robust antimicrobial activity. The purification of p138c, using precipitation in the presence of a low saturation of diammonium sulfate, and elution at void volume using gel permeation chromatography, indicated that p138c was secreted in the form of large aggregates. This result is consistent with those of other studies examining [8,16].

Peptide p138c was purified 19.15-fold with 3.2% recovery.

Bacterial growth kinetics and inhibitory activity indicated that antimicrobial activity occurred during the steady state of the growth curve, from early to late stationary phase, and declined in the very late stationary phase. Antimicrobial activity of the

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Fig. 4. Synergy improves killing kinetics of p138c. Killing kinetics of p138c, and of p138c combined with oxacillin (A), ampicillin (B), and penicillin G (C), with added 0.5 MIC of p138c, against vancomycin-resistant Staphylococcus aureus (VRSA).

test strain was detected in the mid-stationary phase, attaining maximum in the late exponential phase with significant decrease at 44 h of incubation. Our results suggest that sporulation had no effect on the production of the antimicrobial compound.

Peptide p138c was found to be stable at a temperature as high as 50 °C and a wide range of pH values (5.8–11). Our result is comparable to, and shows an even wider pH range than those in other studies [8,16,19]. Antimicrobial activity was retained over a wide range of pH values, which may be

due to the reversible denaturation of the protein. Peptide p138c was completely inactivated in presence of proteinase K, but this effect was not detected after treatment with trypsin, indicating the proteinaceous nature of p138c. Neither H2O2 nor UV light had any effect on the activity of p138c, again indicating the proteinaceous property of our antimicrobial peptide. The N-terminal amino acid sequence analysis of p138c was performed using automated Edman degradation.

Among the last 15 N-terminal amino acids in the sequence, one residue, which could not be identified, is shown as X. During the sequencing cycles of the analysis, the reaction was not blocked. The unknown residue in the amino-acid sequence may be the result of uncleaved amino acids.

Synergizing antimicrobial peptides and non-peptide antibiotics is an efficient approach for lowering the dosage of each molecule in combinatorial therapy. Our study showed an improvement in the MICs of p138c combined with oxacillin, ampicillin, and penicillin G. Synergy was indicated when FIC indices were ≤ 0.5. p138c, along with oxacillin, ampicillin, or penicillin G, caused dramatic decreases in MIC. The observed potentiation of antibacterial activity against VRSA was 4-fold greater when p138c was combined with oxacillin, 8-fold greater when p138c was combined with ampicillin, and 16fold greater when p138c was combined with penicillin G. The effectiveness of synergy can be evaluated by an effective utilization of a weak or moderately effective antimicrobial peptide, which, if used independently, would be considered a poor antibiotic. These combinations influence the antibiotic potency of the peptide-nonpeptide effect and decrease the dose of each agent in the drug combination, rendering synergy more economical. In our study, the synergy of the peptide with three β-lactam antibiotics (oxacillin, ampicillin, and penicillin G) suggested that the antimicrobial peptide might be synergistic with other antibiotics. Our results indicate that synergy could potentiate the bacteriostatic effect and improve bactericidal potency. In synergy with oxacillin, ampicillin, or penicillin G, the bactericidal potency increased 4 to 8-fold, demonstrating a potent bactericidal combination. According to Stratton, bactericidal drugs are preferable to bacteriostatic drugs because the risk for emergence of a resistant mutant is very low; the development of resistant mutants is prevented by killing the microorganism [20]. Hence, we can reduce the chances of antibiotic resistance by increasing the bactericidal potency of antimicrobial peptides. In addition to high potency, rapid killing kinetics are among the properties of antibio­ tics. In this study, the synergistic activity of oxacillin (MIC of

ANTIMICROBIAL PEPTIDE FROM B. SUBTILIS

80 µg/ml), ampicillin (MIC of 20 µg/ml), and penicillin G (MIC of 40 µg/ml), combined with p138c (MIC of 20 µg/ml), against VRSA showed a robust improvement in potency (Fig. 4).

As shown in this figure, p138c alone was largely bacteriostatic against VRSA, but in synergy with the three β-lactam anti­ biotics, p138c showed improved potency and killing kinetics.

In summary, our study investigated the potency and bacterial killing kinetics of the antimicrobial peptide p138c synergized with conventional antibiotics. The results of our study highlight the use of synergy to enhance the therapeutic potential of antimicrobial peptides. These strategies can be applied to a wide spectrum of antimicrobial peptides to produce a suitable agent for the treatment of MDR infections.

Acknowledgements. This work was supported by a grant from the National Research Foundation of Korea (NRF) funded by the Korean government (NRF-2015R1A2A1A15056120, NRF-2015R1D1A1A 01059483) and the Bio-industry Technology Development Program, Ministry of Agriculture, Food and Rural Affairs (115073-2).

Competing interests. None declared.

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