BioMed Research International
BIODESERT: Exploring and Exploiting
the Microbial Resource of Hot
and Cold Deserts
Guest Editors: Ameur Cherif, George Tsiamis, Stéphane Compant,
and Sara Borin
BIODESERT: Exploring and Exploiting
the Microbial Resource of Hot and Cold Deserts
BioMed Research International
BIODESERT: Exploring and Exploiting
the Microbial Resource of Hot and Cold Deserts
Guest Editors: Ameur Cherif, George Tsiamis,
Stéphane Compant, and Sara Borin
Copyright © 2015 Hindawi Publishing Corporation. All rights reserved.
This is a special issue published in “BioMed Research International.” All articles are open access articles distributed under the Creative
Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original
work is properly cited.
Contents
BIODESERT: Exploring and Exploiting the Microbial Resource of Hot and Cold Deserts, Ameur Cherif,
George Tsiamis, Stéphane Compant, and Sara Borin
Volume 2015, Article ID 289457, 2 pages
The Date Palm Tree Rhizosphere Is a Niche for Plant Growth Promoting Bacteria in the Oasis Ecosystem,
Raoudha Ferjani, Ramona Marasco, Eleonora Rolli, Hanene Cherif, Ameur Cherif, Maher Gtari,
Abdellatif Boudabous, Daniele Daffonchio, and Hadda-Imene Ouzari
Volume 2015, Article ID 153851, 10 pages
Pentachlorophenol Degradation by Janibacter sp., a New Actinobacterium Isolated from Saline
Sediment of Arid Land, Amel Khessairi, Imene Fhoula, Atef Jaouani, Yousra Turki, Ameur Cherif,
Abdellatif Boudabous, Abdennaceur Hassen, and Hadda Ouzari
Volume 2014, Article ID 296472, 9 pages
Biotechnological Applications Derived from Microorganisms of the Atacama Desert,
Armando Azua-Bustos and Carlos González-Silva
Volume 2014, Article ID 909312, 7 pages
Diversity and Enzymatic Profiling of Halotolerant Micromycetes from Sebkha El Melah, a Saharan Salt
Flat in Southern Tunisia, Atef Jaouani, Mohamed Neifar, Valeria Prigione, Amani Ayari, Imed Sbissi,
Sonia Ben Amor, Seifeddine Ben Tekaya, Giovanna Cristina Varese, Ameur Cherif, and Maher Gtari
Volume 2014, Article ID 439197, 11 pages
Geodermatophilus poikilotrophi sp. nov.: A Multitolerant Actinomycete Isolated from Dolomitic Marble,
Maria del Carmen Montero-Calasanz, Benjamin Hofner, Markus Göker, Manfred Rohde, Cathrin Spröer,
Karima Hezbri, Maher Gtari, Peter Schumann, and Hans-Peter Klenk
Volume 2014, Article ID 914767, 11 pages
Safe-Site Effects on Rhizosphere Bacterial Communities in a High-Altitude Alpine Environment,
Sonia Ciccazzo, Alfonso Esposito, Eleonora Rolli, Stefan Zerbe, Daniele Daffonchio, and Lorenzo Brusetti
Volume 2014, Article ID 480170, 9 pages
Contrasted Reactivity to Oxygen Tensions in Frankia sp. Strain CcI3 throughout Nitrogen Fixation and
Assimilation, Faten Ghodhbane-Gtari, Karima Hezbri, Amir Ktari, Imed Sbissi, Nicholas Beauchemin,
Maher Gtari, and Louis S. Tisa
Volume 2014, Article ID 568549, 8 pages
Screening for Genes Coding for Putative Antitumor Compounds, Antimicrobial and Enzymatic
Activities from Haloalkalitolerant and Haloalkaliphilic Bacteria Strains of Algerian Sahara Soils,
Okba Selama, Gregory C. A. Amos, Zahia Djenane, Chiara Borsetto, Rabah Forar Laidi, David Porter,
Farida Nateche, Elizabeth M. H. Wellington, and Hocine Hacène
Volume 2014, Article ID 317524, 11 pages
Absence of Cospeciation between the Uncultured Frankia Microsymbionts and the Disjunct Actinorhizal
Coriaria Species, Imen Nouioui, Faten Ghodhbane-Gtari, Maria P. Fernandez, Abdellatif Boudabous,
Philippe Normand, and Maher Gtari
Volume 2014, Article ID 924235, 9 pages
Hindawi Publishing Corporation
BioMed Research International
Volume 2015, Article ID 289457, 2 pages
http://dx.doi.org/10.1155/2015/289457
Editorial
BIODESERT: Exploring and Exploiting the Microbial Resource
of Hot and Cold Deserts
Ameur Cherif,1 George Tsiamis,2 Stéphane Compant,3 and Sara Borin4
1
University of Manouba, Biotechnology and Bio-Geo Resources Valorization (LR11-ES31), Higher Institute for Biotechnology,
BiotechPole Sidi Thabet, 2020 Ariana, Tunisia
2
Department of Environmental and Natural Resources Management, University of Patras, 2 Seferi Street, 30100 Agrinio, Greece
3
Bioresources Unit, Health & Environment Department, AIT Austrian Institute of Technology GmbH, 3430 Tulln, Austria
4
Department of Food, Environmental and Nutritional Sciences (DeFENS), University of Milan, Via Celoria 2, 20133 Milan, Italy
Correspondence should be addressed to Ameur Cherif; [email protected]
Received 15 February 2015; Accepted 15 February 2015
Copyright © 2015 Ameur Cherif et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Deserts are generally regarded as lifeless and inhospitable
ecosystems despite the general awareness of extremophilic
microorganisms. An amazing microbial diversity and a huge
biotechnological potential were unraveled during the last
decade using molecular approaches. Hot and cold deserts
were shown to host peculiar microbial assemblages able to
cope with hostile environment and/or to rapidly adapt to
changing conditions. This adaptation is inferred to particular community structure behavior and specific metabolic
capacities allowing cells to overcome water stress, fluctuating
temperature, and high salinity. Therefore, such microbes
could constitute a source of novel metabolites, biomolecules,
and enzymes potentially useful for environmental biotechnologies. With the global climate change, the aridification and
creeping desertification that constitute a worldwide serious
threat directly affecting agriculture and crop production, and
the growing food demands, desert microorganisms could
hold the key for green biotechnology and future applications
into soil bioreclamation and plant growth promotion for
vulnerable regions across the world.
This special issue was focused on the desert microbial
resource management (MRM) and how to explore and exploit
these resources from hot and cold deserts as well as from
arid areas. Aspects of this MRM concept are included in
this special issue and they are highlighting (a) the microbial
diversity and community structure behavior in desert environments and the identification of novel extremophiles, (b)
the influence of the biotic and abiotic factors on microbial
communities shape and dynamic and the functional networking including mechanism of adaptation and plantmicrobes interaction under extreme or changing conditions,
and (c) potential and case applications of desert microbes
and/or mixed cultures, such as in soil bioreclamation,
reverse-desertification, agriculture, and biomining.
This special issue contains one review and eight research
articles that address the three main aspects indicated above.
The paper of A. Jaouani et al., entitled “Diversity and Enzymatic Profiling of Halotolerant Micromycetes from Sebkha El
Melah, a Saharan Salt Flat in Southern Tunisia,” reported the
isolation of 21 alkali-halotolerant Ascomycetes assigned to the
6 genera Cladosporium, Alternaria, Aspergillus, Penicillium,
Ulocladium, and Engyodontium, basing on morphological
and molecular markers. Beside their salt and pH tolerance,
these saline-system fungi were shown to resist to oxidative
stress and low temperature and to produce extremozymes,
namely, cellulase, amylase, protease, lipase, and laccase, active
in high salt concentrations, which highlight their biotechnological potential. The paper authored by M. del C. MonteroCalasanz et al., “Geodermatophilus poikilotrophi sp. nov.: A
Multitolerant Actinomycete Isolated from Dolomitic Marble,”
described a new species within the genus Geodermatophilus.
Strain G18T , isolated from site near the Namib Desert,
is characterized by its resistance to heavy metals, metalloids, hydrogen peroxide, desiccation and ionizing, and UVradiations. Even though 16S rRNA sequence of strain G18T
showed 99% similarity with other Geodermatophilus species,
2
its taxonomic position and species definition was inferred
basing on polyphasic approach and its multitolerance towards
environmental stresses, justifying the original given epithet
“poikilotrophi.”
Four papers were dedicated to the ecological drivers
that shape microbial communities and functionalities. A
nice example of “cold desert” is presented by S. Ciccazzo et
al., “Safe-site Effects on Rhizosphere Bacterial Communities
in a High-Altitude Alpine Environment.” In this work, the
authors investigated rhizobacterial communities associated
with floristic consortia in different safe-sites located in
deglaciated terrain. Using DGGE and ARISA, they demonstrated a clear correlation between soil maturation and
bacterial diversity and a plant-specific effect leading to the
selection of specific rhizobacterial communities by the pioneer plants. Another model ecosystem was investigated by R.
Ferjani et al., in the paper “The Date Palm Tree Rhizosphere
Is a Niche for Plant Growth Promoting Bacteria in the Oasis
Ecosystem.” The work focused on the characterization of
the bacterial communities in the soil fractions associated
with the root system of date palms cultivated in seven
oases in Tunisia using culture-independent and dependent
approaches. It was shown that the date palm rhizosphere
bacterial communities were rather complex and correlate
with geoclimatic and macroecological factors along a northsouth aridity transect. However, with the wide diversity of
cultivable strains detected, interesting common features of
plant growth promoting (PGP) activity and abiotic stress
resistance were detected. The authors concluded that palm
root system and rhizosphere soil represent a reservoir of PGP
bacteria involved in the regulation of plant homeostasis. The
third example of plant-microbe interaction is the paper by
I. Nouioui et al., in which they demonstrated, as written in
the title, the “Absence of Cospeciation between the Uncultured Frankia Microsymbionts and the Disjunct Actinorhizal
Coriaria Species.” The investigation was achieved on five
Coriaria host species sampled from sites covering the full
geographical range of the genus (Morocco, France, New
Zealand, Pakistan, Japan, and Mexico). The reported findings argue that Frankia, the nitrogen-fixing actinobacteria
microsymbionts, have not evolved jointly with their host
plants and had probably dispersed globally as a protoFrankia, a free living nonsymbiotic ancestor. The authors
hypothesized also that cospeciation may have occurred but
subsequently lost after bacteria mixing and fitness selection in
the presence of indigenous symbionts. Frankia sp. was further
investigated in terms of nitrogen fixation under different
oxygen tensions. This work authored by F. Ghodhbane-Gtari
et al. was conducted on the actinorhizal plant Casuarina and
its compatible Frankia sp. strain CcI3. By studying the growth
of the strain, vesicle production, and several genes expression,
the authors confirmed the correlation between the biomass
and the vesicle production with elevated oxygen tension. It
was also shown that oxygen levels influenced nitrogenase
induction and that Frankia protects nitrogenase by the use of
multiple mechanisms including the vesicle-hopanoid barrier
and increased respiratory protection. Clearly, the microbial
assemblages selected by the plant roots in desert and arid soils
are shaped by the ecological biotic and abiotic drivers but with
BioMed Research International
the prerequisite of providing rhizosphere services and specific
functionalities.
Biotechnological potential and applications of desert
microbes have been reported in three papers. In one, A.
Khessairi et al. described a novel efficient pentachlorophenol(PCP-) degrading halotolerant actinobacterium, Janibacter
sp. FAS23. The strain was isolated from Sebkha El Naoual, a
saline ecosystem in southern Tunisia. Using HPLC analysis,
FAS23 was shown to be able to degrade high concentration
of PCP (up to 300 mg/l) and to tolerate salt fluctuation. PCP
degradation was further enhanced in the presence of glucose
and nonionic surfactant tween 80. The strain is considered as
a candidate for PCP bioremediation in polluted soils in arid
areas. In another paper authored by O. Selama et al., the isolation of haloalkalitolerant and haloalkaliphilic bacteria from
Algerian Sahara Desert soil was reported. Thirteen selected
isolates, mainly filamentous Actinobacteria, were screened
phenotypically for antibacterial, antifungal, and enzymatic
activities and by PCR for putative antitumor compounds
genes. The isolates were assigned to the genera Streptomyces,
Nocardiopsis, Pseudonocardia, Actinopolyspora, and Nocardia, with this latter constituting possibly a new branch in
the Actinomycetales order. Beside secreted extremozymes and
bioactives, several isolates showed antitumorigenic potential.
Another paper presented in this special issue is a review article nicely written by A. Azua-Bustos and C. González-Silva,
who focused on the Atacama Desert microbes and their current biotechnological applications. A large-scale application
in Chile is the copper bioleaching or biomining mediated by
indigenous halotolerant and acidophilic chemolithotrophic
bacteria like Acidithiobacillus ferrooxidans and Acidithiobacillus thiooxidans. Other potential applications in arsenic bioremediation and in biomedicine, including the discovery of
new antibiotics, antioxidant, antifungal, and immunosuppressive compounds, were cited. The authors reported also
application from eukaryotic microorganism as in the case
of the halophilic biflagellate unicellular green alga Dunaliella
that produce beta-carotene.
Definitely, desert environments represent a tremendous
reservoir where more efforts, relying not only on metagenomics but also on culturomics, should be dedicated to
unravel the hidden potential. The second decade for desert
biotechnology has just begun.
Acknowledgments
We thank the authors of the submitted papers for their
contribution. The preparation of this special issue would
not have been possible without the generous support and
dedication of experts who evaluated the papers submitted.
Ameur Cherif
George Tsiamis
Stéphane Compant
Sara Borin
Hindawi Publishing Corporation
BioMed Research International
Volume 2015, Article ID 153851, 10 pages
http://dx.doi.org/10.1155/2015/153851
Research Article
The Date Palm Tree Rhizosphere Is a Niche for Plant Growth
Promoting Bacteria in the Oasis Ecosystem
Raoudha Ferjani,1 Ramona Marasco,2 Eleonora Rolli,3 Hanene Cherif,1 Ameur Cherif,4
Maher Gtari,1 Abdellatif Boudabous,1 Daniele Daffonchio,2,3 and Hadda-Imene Ouzari1
1
LR03ES03 Laboratoire Microorganismes et Biomolécules Actives, Faculté des Sciences de Tunis, Université de Tunis El Manar,
Campus Universitaire, 2092 Tunis, Tunisia
2
Biological and Environmental Sciences and Engineering Division, King Abdullah University of Science and Technology,
Thuwal 23955-6900, Saudi Arabia
3
Department of Food, Environment, and Nutritional Sciences, University of Milan, Via Celoria 2, 20133 Milan, Italy
4
Université de La Manouba, Institut Supérieur de Biotechnologie de Sidi Thabet, LR11ES31 LR Biotechnologie & Valorisation
des Bio-Géo Ressources, BiotechPole Sidi Thabet, 2020 Ariana, Tunisia
Correspondence should be addressed to Hadda-Imene Ouzari; [email protected]
Received 16 May 2014; Accepted 6 October 2014
Academic Editor: Sara Borin
Copyright © 2015 Raoudha Ferjani et al. This is an open access article distributed under the Creative Commons Attribution
License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly
cited.
In arid ecosystems environmental factors such as geoclimatic conditions and agricultural practices are of major importance in
shaping the diversity and functionality of plant-associated bacterial communities. Assessing the influence of such factors is a key
to understand (i) the driving forces determining the shape of root-associated bacterial communities and (ii) the plant growth
promoting (PGP) services they provide. Desert oasis environment was chosen as model ecosystem where agriculture is possible
by the microclimate determined by the date palm cultivation. The bacterial communities in the soil fractions associated with the
root system of date palms cultivated in seven oases in Tunisia were assessed by culture-independent and dependent approaches.
According to 16S rRNA gene PCR-DGGE fingerprinting, the shapes of the date palm rhizosphere bacterial communities correlate
with geoclimatic features along a north-south aridity transect. Despite the fact that the date palm root bacterial community structure
was strongly influenced by macroecological factors, the potential rhizosphere services reflected in the PGP traits of isolates screened
in vitro were conserved among the different oases. Such services were exerted by the 83% of the screened isolates. The comparable
numbers and types of PGP traits indicate their importance in maintaining the plant functional homeostasis despite the different
environmental selection pressures.
1. Introduction
The southern regions of Tunisia are very arid and the date
palm (Phoenix dactylifera L.) is a key plant determining in
the oasis agroecosystem a microclimate that favours agriculture [1]. The palms protection provides many ecosystem
services, including ameliorating oasis temperature, changing
floodwater dynamics and facilitating wildlife, and making
agriculture possible under harsh environmental conditions
[2]. In the world, oases cover about 800 000 ha and support
the living of 10 million people. In Tunisia more than four millions of date palm trees are spread onto 32 000 ha of oasis in
the southern part of the country [3, 4]. As a result of the oases
overexploitation and strong anthropogenic pressures, these
ecosystems are becoming increasingly fragile. Furthermore,
despite the oasis potential to tolerate several abiotic stresses
typical of arid environment, the ongoing climate change
is enhancing the environmental pressure on the date palm
affecting growth and development, especially in the Middle
East [5].
Besides the well-known plant growth promoting properties typical of rhizospheres in temperate soils in nonarid
ecosystems, rhizosphere bacteria in arid soils contribute in
counteracting drought and salinity stresses, by providing
services such as, among others, physical protection of the
root from mechanical stress against the dry soil particles,
2
induction of plant physiological responses against water
losses [6], or productions of metabolites contributing to the
maintenance of the plant hormone and nutrient homeostasis,
[7]. In particular PGP (plant growth promoting) bacteria,
naturally associated with plants, have been shown to be
essential partners for improving plant tolerance to stressful
conditions [8]. The exploration of plants naturally adapted
to extreme condition may allow a reservoir of biodiversity
exploitable to understand the ecological service enclosed in
these ecosystems [8, 9]. In this context, ecological niche presented in the oasis ecosystem could provide a new model to
study and dissect the key factors driving the stability of this
ecosystem [10]. Little information is available about the
microbiological functionality of both oasis and date palm.
For instance the potential PGP services provided by the rootassociated bacteria appear to be invariant with respect to
geoclimatic factors despite provided by different bacterial
communities, according to observations across a north to
south aridity transect that included Tunisia [11].
Since plants contribute to shape soil microbial diversity
[12, 13], the aim of this work was to assess bacterial communities associated with the date palm rhizosphere soil, the
root surrounding soil and the bulk soil fractions in seven
Tunisian oases, in order to evaluate if along a north-south
transect (i) the assemblage of bacterial communities in the
palm root soil fractions was driven by the geoclimatic factors
and (ii) the ecological services were preserved in the soil
fractions of the root system. The structure of the bacterial
communities associated with the soil fractions of date palm in
the seven oases was dissected by 16S rRNA gene-based PCRDGGE (denaturing gradient gel electrophoresis) analysis. The
results were analysed in function of geoclimatic factor and
oasis origin, and compared with the diversity of the cultivable
bacteria and their PGP potential.
2. Materials and Methods
2.1. Site Description and Sampling. The sampling was carried
out from seven oases in different geographic locations in
Tunisia, along a latitude/longitude gradient, respectively from
32∘ to 34∘ N and from 7∘ to 9∘ E (Figure 1(a) and Supplementary Table 1 in the Supplementary Material available online at
http://dx.doi.org/10.1155/2015/153851). A traditional crop management was used in all the oases, including groundwaterbased flooding irrigation and fertilization with organic fertilizers. In each oasis, the roots of three date palm trees of
similar age, lying in the distance of less than 15 meters and
growing in the same soil were separately collected at 20–
30 cm depth in order to obtain the adhering rhizosphere soil
(R) tightly attached to roots. After removing the roots, the
root surrounding soil (S) was collected. Bulk soil samples (B)
not influenced by date palm root system were also sampled
as control. All soil samples were collected under sterile
conditions using sterile tools. Recovered samples were stored
at −20∘ C for molecular analysis or at 4∘ C for isolation.
2.2. Total DNA Extraction, PCR-DGGE, and Profile Analysis.
Total DNA from soil samples was extracted by commercial
kit FASTDNA SPIN KIT for soil (Qbiogene, Carlsbad, USA)
BioMed Research International
according to the manufacturer’s procedure. PCR amplification was performed in a final volume of 50 𝜇L using primers
907R and 357F, adding a GC-clamp to the forward primer
[14, 15]. The reaction mixture was prepared with 1X PCR
buffer, 2.5 mM MgCl2 , 0.12 mM deoxynucleoside triphosphate, 0.3 mM of each primer and 1U Taq DNA polymerase
and 10 ng of pooled DNA obtained from the three plant replicates were added as template. PCR products were resolved on
7% (w/v) polyacrylamide gel in 1X TAE pH 7.4 with a 40–60%
denaturing gradient. Gels were run at 90 V for 17 h at 60∘ C
in DCode apparatus (Bio-Rad, Italy). After electrophoresis,
gels were stained with ethidium bromide solution for 30 min,
washed with sterile distilled water, and photographed on a
UV transillumination table. The DGGE band profiles were
converted into numerical values using Image J (version 1.46)
and XLSTAT software.
2.3. Real Time PCR. Quantitative real time PCR (q-PCR)
was performed on a Chromo4 real time PCR machine (BioRad) to measure the presence and concentration of bacterial
16S rRNA gene associated with the rhizosphere fractions.
The reactions were performed with IQ SYBR Green Supermix (Bio-Rad), using primers targeting the 16S rRNA gene
(Bac357-F and Bac907-R) [16]. PCR SYBR green reactions
were prepared by using the “Brilliant SYBR Green QPCR
Master Mix” kit (Stratagene) in 96-well plates. The reaction
mix (25 mL) contained 1X Brilliant SYBR Green (2.5 mM
MgCl2 ), 0.12 mM of each primers, and approximately 100 ng
of extracted DNsA. The DNA obtained from the three plants
sampled in the same station was pooled and used as template
to carry out the real time assay in triplicate. At the end of
each real time PCR, a melting curve analysis was performed
for verifying the specificity of PCR products. To construct
standard curves, the 16S rRNA gene of Asaia sp. was amplified
by PCR and cloned using the pGEM T-easy Vector Cloning
Kit (Promega). q-PCR data relative to the 16S rRNA gene
concentration were log-transformed.
2.4. Isolation of Cultivable Bacteria. One gram of rhizosphere
soil (R) from each sample was suspended in 9 mL of sterile
physiological solution (9 g/L NaCl) and shaken for 15 min
at 200 rpm at room temperature. Suspensions were diluted
in tenfold series and plated in triplicate onto TSA (Tryptic
Soy Agar), YEM (Yest Extract Mannitol), and KB (King’B
agar) culture media. After three days at 30∘ C colonies were
randomly selected and spread on the original medium for
three times to avoid contamination risks. Pure strains were
frozen in 25% glycerol at −80∘ C. A total of 440 isolates were
collected. The isolates were named based on the station and
the medium from which they were isolated.
2.5. DNA Extraction, Dereplication, and Identification of Isolates. Genomic DNA was recovered from the isolates using a
boiling lysis. Bacterial cells were suspended in 50 𝜇L of sterile
TE (10 mM Tris/HCl, pH 8, 1 mM EDTA) and incubated
at 100∘ C for 8 min. After centrifugation (13000 g, 10 min),
the supernatant containing the released DNA was stored at
−20∘ C and used as template for PCR. Amplification of 16S–
23S internal transcribed spacer region (ITS) was performed
3
PCO2 (22.6% of total variation)
BioMed Research International
20
R
B
S
R
R R
0
S
B
B
−40
B
S
−20
0
20
40
PCO1 (62.2% of total variation)
Tozeur (BD-16)
Tamerza (BD-B)
Ain el Karma (BD-C)
Ksar Ghilane (BD-1)
Douz (BD-5)
El Faouar (BD-8)
Rjim Maatoug (BD-9)
(b)
(a)
10
7.5
5
Log10 (16S rRNA copies)
dbRDA2
(16.2% of fitted, 7.1% of total variation)
B
S
−20
Sampling station
S S
RR S
R
Tmin (∘ C)
Rainfall max (mm)
Long. E
0
Tmax (∘ C)
Lat. N
Rainfall min (mm)
−5
Alt (m)
−10
−10
−5
0
5
10
15
dbRDA1 (68.1% of fitted, 29.9% of total variation)
(c)
20
7.0
ab
a
a
a
a
6.5
a
6.0
b
5.5
5.0
BD-1
BD-5
BD-8
BD-9 BD-16 BD-B BD-C
(d)
Figure 1: Station location and analysis of bacterial community structure associated with soil fraction of date root system. (a) The location of
the studied oases is indicated on the map of Tunisia. (b) A principal coordinate analysis (PCO), deduced from the 16S rRNA gene-based PCRDGGE profiles, resumes the diversity of the bacterial communities associated with the date palm root soil fractions. 84.8% of total variation is
explained in the presented PCO. The soil fractions analysed are R: rhizosphere; S: root surrounding soil; and B: bulk soil. (c) Dist LM analysis
to evaluate which are the main geoclimatic variables influencing the structure of the bacterial communities associated with date palm root
soil fractions. Lat. N: latitude north; Long. E: longitude east; Alt.: altitude; 𝑇min : minimum temperature; 𝑇max : maximum temperature; Rainfall
min: minimum rainfall; Rainfall max: maximum rainfall. (d) Box-plot graph represents the quantification of 16S rRNA gene by qPCR. The
number of copies is expressed in Log10 . Statistical analysis (pairwise test) of bacterial assemblage across locations was indicated by the letter.
using the universal primers S-DBact-0008-a-S-20 (5󸀠 -CTA
CGG CTA CCT TGT TAC GA-3󸀠 ) and S-D-Bact-1495-a-S-20
(5󸀠 -AGA GTT TGA TCC TGG CTC AG-3󸀠 ) according to the
procedure described previously by Fhoula et al. [17]. Two 𝜇L
of the PCR products were checked by electrophoresis in 1.5%
agarose gel and stained with ethidium bromide. Gel images
were captured using Gel Doc 2000 system (Bio-Rad, Tunis,
Tunisia), and bacteria redundancy was reduced by evaluating
the different ITS profiles. One strain per each ITS haplotype
was used in the phylogenetic analysis and for further experiments. A total of 98 strains were characterized by 16S rRNA
gene sequencing using the primers S-D-Bact-1494-a-20 (5󸀠 GTC GTA ACA AGG TAG CCG TA-3󸀠 ) and L-D-Bact-0035a-15 (5󸀠 -CAA GGC ATC CAC CGT-3󸀠 ). PCR amplification
was carried out as described by Fhoula et al. [17]. The 16S
rRNA PCR amplicons were purified with Exonuclease-I and
Shrimp Alkaline Phosphatase (Exo-Sap, Fermentas, Life
Sciences) following the manufacturer’s standard protocol.
Sequencing of the purified amplicons was performed using
a Big Dye Terminator cycle sequencing kit V3.1 (Applied
Biosystems) and an Applied Biosystems 3130XL Capillary
DNA Sequencer machine. The obtained sequences, with an
average length of 750 bp, were compared with those available
at the National Centre for Biotechnology Information (NCBI)
database (http://www.ncbi.nlm.nih.gov) using the basic local
alignment search tool (BLAST) algorithm [18]. The 16S rDNA
sequences were submitted to the NCBI nucleotide database
under the accession number KJ956590 to KJ956687.
Phylogenetic analysis of the 16S rRNA gene sequences
was conducted with molecular evolutionary genetics analysis
(MEGA) software, version 6 [19]. Trees were constructed by
using neighbor-joining method [20].
4
2.6. Characterization of Plant Growth Promoting Activity and
Abiotic Stress Resistance. The 98 bacterial strains identified
were screened for production of indole acetic acid (IAA),
siderophores and ammonia, mineral phosphate solubilization, protease and cellulose activity, and tolerance to several abiotic stresses. Quantitative production of IAA was
determined as described by Ouzari et al. [21]. Briefly, after
incubation in minimal medium supplemented with glucose
(100 g/L) and L-tryptophane (10 𝜇g/mL), using Salkowski’s
reagent, the colour absorbance was read, after 20–25 min, at
535 nm. Concentration of IAA produced was measured by
comparison with a standard graph of IAA. Ability of bacteria
to solubilize inorganic phosphate was evaluated as described
by Nautiyal [22], by the observation of clear halo around the
bacterial colony grown in Pikovskaya medium. To demonstrate the production of siderophore, the tested strains were
spotted on nutrient agar plates. After incubation for 48–72 h
at 30∘ C, the grown strains were overlaid with CAS medium
supplemented with agarose (0.9% w/v). Positive test was
noted when colour modification around colonies from blue to
orange was observed [23]. Ammonia production was assayed
by inoculation of bacterial strains in 10 mL of peptone water
and using Nessler’s reagent (0.5 mL). Ammonia producing
strains were identified when brown to yellow colour was
developed [24]. Protease (casein degradation) and cellulase
activities were determined by spot inoculation of the strains
on Skimmed milk and CMC agar media, respectively. A clear
halo around the colonies indicates the ability of the strains to
produce the degrading enzymes [25].
Tolerance to osmotic stress was evaluated by adding to
tryptic soy broth (TSB) medium 30% of polyethylene glycol
(PEG 8000). Resistance to salt was assessed by adding 5, 10,
15, 20, 25, and 30% NaCl to the culture media and incubating
the plates at 30∘ C for 5 days. The ability to growth at 45, 50,
and 55∘ C was checked in TSA by incubation at the indicated
temperatures for 5 days. Tolerance to acid (3 and 4) and basic
(10 and 12) pH was assessed by adjusting the medium with
concentrated HCl (12 N) and NaOH (3 M), respectively.
2.7. Statistical Analysis. Significant differences in soil bacterial community structure were investigated by permutational
analysis of variance (PERMANOVA, [26]). Distance-based
multivariate analysis for a linear model (DistLM, [27]) was
used to determine which significant environmental variables
explain the observed similarity among the samples. The
Akaike information criterion (AIC) was used to select the
predictor variables [28]. The contribution of each environmental variable was assessed: firstly the “marginal test” is used
to assess the statistical significance and percentage contribution of each variable by itself and secondly the “sequential
test” was employed to explain the biotic similarity considering all the variable contributions. All the statistical tests
were performed by PRIMER v. 6.1 [29], PERMANOVA+for
PRIMER routines [30].
3. Results and Discussion
3.1. Environment Parameters Directly Influence Bacterial Communities Associated with Palm Rhizosphere. The diversity of
BioMed Research International
bacterial communities associated with the date palm root
system from each of the seven studied oasis was investigated
through the analysis of the diversity of the 16S rRNA gene in
the rhizosphere (R) and root surrounding soil (S) fractions.
Bulk soil (B) was also included as a comparative fraction not
directly influenced by the plant root. Separation among
palm bacterial communities located in north (BD-16, BDB, and BD-C) and south (BD-1, BD-5, BD-8, and BD9) oases was supported by a principal coordinate analysis
(PCO) (Figure 1(b)), suggesting that geoclimatic conditions
influence the bacterial community structure. Statistical analysis confirmed the grouping observed in the PCO analysis
with a significant difference between north and south oases
(PERMANOVA, df = 1.55; 𝐹 = 8.06; 𝑃 = 0.0017) but not a
significant separation mediated by the aquifer used to irrigate
the oases (PERMANOVA, df = 1.55; 𝐹 = 1.45; 𝑃 = 0.21).
Within the two macroregions, the north and south
groups of oases, we observed a significant difference among
oases (Supplementary Table 2) indicating the presence of
oasis-specific bacterial community supporting a concept of
“ecological island.” Pairwise analysis showed that such differences observed among the oases predominantly occurred
in the north regions (Supplementary Table 3), possibly
because the south region (closest to desert) presents harsher
conditions that select a more restricted type of bacteria. These
ecological islands represent specific cluster of biological
diversity that may contribute to the overall regional bacterial
community functionality and furthermore increase the level
of resilience to environmental change of the entire system
[30].
Along the transect, the soil fraction communities were
significantly different (PERMANOVA, df = 2.55; 𝐹 = 2.70;
𝑃 = 0.03). In particular the rhizosphere community, that
resides in the first millimeter of soil adhering to the root,
appeared completely different from the root surrounding soil
(PERMANOVA, 𝑡 = 2.04; 𝑃 = 0.017, p-pht) and bulk soil
(PERMANOVA, 𝑡 = 2.05; 𝑃 = 0.019, p-pht), suggesting the
influence of palm root exudates in shaping the bacterial
community. Generally, the rhizosphere is the transition zone
between the root surface and soil where the released exudates and the rhizodeposition favour microbial proliferation,
inducing changes in the structure and in the chemical-physical properties of the soil [31]. Indeed, the analysis of bacterial
abundance in the rhizosphere showed a numbers of 16S rRNA
copies ranging from 5.88 ± 0.78 to 6.63 ± 0.15 (Figure 1(d)).
Despite similar values observed in the rhizosphere community, a statistical difference among the stations was identified
(PERMANOVA, df = 6.20; 𝐹 = 2.93; 𝑃 = 0.041), mainly
influenced by environmental factor directly linked to location, such as altitude and temperature maximum (DistLM,
𝑃 = 0.03).
Despite the rhizosphere effect observed along the transect, in each oasis considered separately from the others, rhizobacterial community appeared directly connected to that
present in the root surrounding soil and the bulk soil, since
no statistically significant differences in the bacterial diversity
were observed among the different soil fractions within each
station (R, S e B: PERMANOVA, df = 12.55; 𝐹 = 1.62; 𝑃 =
0.057). The rhizosphere effect is particularly noticeable in
BioMed Research International
5
Table 1: Environmental factors associated to the structure of the date palm soil bacterial community. Relationships between bacterial
assemblages and climate features using nonparametric multivariate multiple regression analysis (DISTLM). (a) Marginal test considers each
single geographical variables and their contribution to explain the total variability. (b) Sequential test explaining the total variation with the
contribution of all the variables accounted together. Lat. N: latitude north; Long. E: longitude east; Alt.: altitude; 𝑇min : minimum temperature;
𝑇max : maximum temperature; Rainfall min: minimum rainfall; Rainfall max: maximum rainfall; 𝐹: statistic 𝐹; 𝑃: probability (in bold the
variables statistically significant; 𝑃 < 0.05); Prop.: proportion of total variation explained; Cumul.: cumulative variation explained by the
variables listed; Res df: residual degrees of freedom.
(a) Marginal test
Variable
Lat. N
Long. E
Alt (m)
𝑇min (∘ C)
𝑇max (∘ C)
Rainfall min (mm)
Rainfall max (mm)
𝐹
6.4211
4.8698
3.2116
3.8811
1.147
3.9401
1.8821
SS (trace)
1855.5
1444.3
980.15
1170.8
363.14
1187.3
588.06
𝑃
0.0028
0.0097
0.0376
0.0193
0.3034
0.0198
0.136
(b) Sequential test
Variable
(+) Lat. N
(+) Long. E
(+) Alt (m)
(+) 𝑇min (∘ C)
(+) 𝑇max (∘ C)
(+) Rainfall min (mm)
(+) Rainfall max (mm)
AIC
319.28
320.07
317.33
316.7
309.31
303.19
303.19
SS
1855.5
331.89
1240.6
644.65
2066
1528.3
< 0.01
nutrient-poor soils and under severe abiotic stresses, as previously observed for herbaceous and arboreal plants grown
in arid lands [7, 8, 32, 33]. In the oasis model the selection
mediated by “oasis ecosystem” appeared stronger than the
one exerted by the plant root system (Supplementary Table
2). Naturally, most of the desert microbial communities seem
to be structured solely by abiotic processes [34, 35]. However,
desert farming may strongly affect the sand/soil microbial
diversity reshaping the structure of the resident microbial
communities [8, 9, 36, 37]. During long-term desert farming
land management, such as that occurring in the studied oases,
the structure of rhizosphere bacterial community is strongly
influenced by the plant and the desert farming practices that
determine drastic shifts in the composition of the original
desert soil communities [7, 8].
Dist LM multivariate analysis was performed in order to
correlate the differences in the structure of bacterial communities in the different oases with environmental parameters.
The selection of soil microorganisms by the rhizosphere is a
complex process controlled by several factors, often not easily
correlated to the environmental settings [38]. Nevertheless,
Dist LM analysis showed that geoclimatic parameters contributed to drive the assemblage of the bacterial communities.
In particular, marginal test showed latitude, longitude, altitude, minimum temperature, and minimum rainfall as significant variables singularly involved in the selection of bacterial
assemblages (Table 1(a)). Sequential test confirmed latitude,
altitude, and temperature as variables involved in the bacterial community shaping (Table 1(b)). We can assume that
𝐹
6.4211
1.1517
4.5974
2.4558
9.1244
7.6468
0
𝑃
0.0022
0.2998
0.0122
0.084
0.0006
0.001
1
Cumul.
0.10627
0.12528
0.19633
0.23325
0.35158
0.43911
0.43911
Res. df
54
53
52
51
50
49
49
a concurrence of environmental factors, including a hot and
dry climate, may influence the differences among the bacterial communities of the soil fractions (R, S, and B) associated
with the root system of date palm cultivated in the oases in
the north and south macroregions examined (Figure 1(c)).
3.2. Cultivable Bacterial Communities Associated with Date
Palm Soil Fractions. The isolation of native bacterial species
associated with date palm root soil was performed using
nonspecific media, in order to select a wide range of genera
of possible plant growth promoters [39–41].
A total of 440 isolates were retrieved from the seven
analyzed stations. To manage such a large set of isolates, total
DNA was extracted from each isolate and 16S–23S rRNA
gene internal transcribed spacers (ITS) were amplified. ITS
characterization represents a useful molecular tool for the
discrimination of bacterial isolates up to the subspecies level
[42–45]. Within the whole bacterial collection, ITS-PCR
fingerprinting revealed 30 distinct haplotypes (H1-H30).
Haplotypes H4 and H20 were the most frequent and were
represented by 46 and 26 isolates, respectively. Representative isolates (from one to four strains for each haplotype,
summing up a total of 98 isolates) were subjected to species
identification using partial 16S rRNA gene sequencing (Supplementary Figure 1).
A wide diversity was detected into date palm rhizosphere bacterial community along the studied aridity transect in Tunisia. Significant differences were observed in the
structure of the bacterial communities in the rhizosphere of
6
3.3. Characterization of Rhizobacteria PGP Potential. The
plant microbiome is a key determinant of plant health and
productivity. Plant-associated microbes can help plants stimulate growth, promote biotic and abiotic stress resistance and
influence crop yield and quality by nutrient mobilization and
transport [6]. While the possibility to contribute to control
biotic stresses by plant-associated microorganisms is well
characterized, less is known for abiotic stress. However,
several promising examples of stress protecting bacteria are
already reported in the literature [7, 38, 53]. Recent works
demonstrated that drought-exposed plants cultivated under
desert farming are colonized by bacterial communities with
high PGP potential [7, 8]. Such a PGP potential can promote
increased tolerance to water shortage, mediated by the induction of a larger root system (up to 40%) that enhances water
uptake [7]. To assess if the oasis date palm PGP potential was
conserved in the rhizosphere soil, 98 isolates were evaluated
for a series of PGP traits. The majority (85%) of isolates
showed multiple PGP activities, which may promote plant
growth directly, indirectly, or synergistically. Only 15% of
the rhizobacteria showed one or no activity, while no strains
displayed all the screened PGP activities. The most common
PGP trait was auxin production (83%), followed by ammonia
synthesis (63%) and biofertilization activities such as solubilization of phosphates (48%) and siderophore production
100
90
80
Isolates (%)
70
60
50
40
30
20
10
0
BD-1
BD-5
BD-8
BD-9
Streptomyces
Arthrobacter
Labdella
Mycobacterium
Cellulomonas
Microbacterium
Staphylococcus
BD-16
Bacillus
Agrobacterium
Thalassospira
Rhizobium
Flavobacterium
Pantoea
Serratia
BD-B
BD-C
Providencia
Yersinia
Rahnella
Salinicola
Enterobacter
Pseudomonas
(a)
100
Isolates (%)
80
60
40
20
0
BD-1
BD-5
BD-8
BD-9
P Solubilization
Ammonia
IAA
BD-16
BD-B
BD-C
BD-B
BD-C
Siderophore
Protease
Cellulase
(b)
100
80
Isolates (%)
the analyzed oases, in particular for the differential distribution pattern of the major bacterial genera (Figure 2(a)).
According to the cluster analysis at the genus level performed
on the entire strain collection, the composition of the cultivable rhizobacterial communities associated with date palm
in the seven oases shared about the 65% similarity.
The phylogenetic identification of cultivable bacteria highlighted a predominance of gram-negative bacteria (66%), belonging to the Gammaproteobacteria (57%),
Alphaproteobacteria (7%), and Betaproteobacteria (1%) subclasses. The remaining isolates were affiliated to the Firmicutes (7%), Actinobacteria (26%), and Bacteroidetes (2%)
classes. Members of these taxa are frequently associated with
different plant species and types, confirming that soil is the
main reservoir of plant-associated bacteria [46]. The strains
were allocated into 20 different genera of variable occurrence
(Figure 2(a)), showing a high genetic diversity in the date
palm rhizosphere presumably influenced by the combined
effects of root exudates and agricultural management practices, particularly important under the arid pedoclimatic
conditions [47]. The rhizobacterial communities were dominated by Pseudomonas, as previously described in herbaceous
plants, arboreal and plant adapted to arid climates [11, 48–50].
Together with Pseudomonas, Pantoea and Microbacterium genera were observed in all stations followed by Bacillus
and Arthrobacter, which were reported in six out of seven stations. As well, Enterobacter, Salinicola, Rhizobium, and Staphylococcus were recorded among 5 stations, suggesting the
adaptation of these genera to the oasis environment. Except
Labedella, the genera found in association with date palm
rhizosphere have been previously recognized as being capable
of colonizing plant root systems in arid environment [11, 38,
48, 50–52].
BioMed Research International
60
40
20
0
BD-1
BD-5
PEG 30%
pH ≤ 4
pH ≥ 10
BD-8
BD-9
BD-16
∘
T ≥ 45 C
NaCl ≥ 10
(c)
Figure 2: Diversity and functionality of cultivable bacteria islated
from date palm rhizosphere. (a) Phylogenetic identification at the
genus level of culturable bacteria associated with date palm rhizosphere. (b) Percentage of date palm rhizosphere-associated bacteria
showing PGP activity. (c) Percentage of isolates displaying the
assayed abiotic stress tolerance in the bacterial collection of strains
associated with date palm cultivated in the seven oases analysed.
BioMed Research International
(44%). In our rhizobacterial collection, the IAA production
was equally distributed among the seven oases selected
along the aridity transect (Figure 2(b)), similarly to previous
observations in other arboreal plants cultivated along a
latitude transect [11], confirming that IAA synthesis is a
widespread PGP trait. The IAA production ranged from 2.5
to 85 𝜇g/mL with 49% of the strains producing an amount
ranging from 10 to 20 𝜇g/mL and the 38% showing higher
levels of IAA (more than 20 𝜇g/mL). As already described
in the literature, Pseudomonas, Bacillus, Pantoea, Staphylococcus, and Microbacterium were the most abundant taxa
implicated in IAA production [48, 54, 55]. The high frequency
of IAA producing strains suggests a role of PGP bacteria
in contributing to regulate the root surface extension and
consequently the potential of water and nutrient uptake [56].
Phosphorus, together with iron and nitrogen, is a key
nutrient for plant, particularly in oasis soil where the availability of nutrient sources of animal origin is scarce [57].
The ability of rhizobacteria to solubilize phosphate (48%)
through the production of organic acids or phytases can be
very important in arid ecosystems [58, 59]. Strains of Pantoea,
Enterobacter, Pseudomonas, Streptomyces, and Rhizobium
genera were the most efficient solubilizers, as previously
showed in other arid contexts such as Tunisian grapevine
[48] and different crops in Bolivia [60]. Several siderophoreproducing bacteria were observed in the rhizosphere (44%)
probably because this PGP trait confers competitive colonization ability in iron-limiting soils. Iron is made available
for the plant host and consequently exerts a biocontrol role
reducing iron-dependent spore germination of fungi [61].
The siderophore-producing bacteria belonged mainly to the
Pseudomonas genus (67%), followed by Bacillus (7%) and
Pantoea (7%). Predominance of siderophore release by Pseudomonas bacteria was already reported in the rhizosphere of
other plants [62, 63]. In addition to siderophore production,
cell wall degrading enzymes implicated in fungal inhibition
and the organic matter turnover [51] were investigated. The
49% and 15% of the examined isolates were able to produce
proteases and cellulases, respectively, with the most active
strains belonging to Serratia marcescens and Sinorhizobium
meliloti, respectively [64, 65]. Ammonia production can indirectly affect plant growth through nitrogen supply [66]. This
trait was represented in 64% of the isolates, confirming its
spread in the palm-bacteria association.
Further analyses were performed to evaluate the adaptability of isolates to abiotic stresses (Figure 2(c)). Drought
stress resistance was presented by 95% of the strains that
could grow in presence of increasing concentrations of PEG.
Most of the strains (98%) were able to grow at 45∘ C, while
only 39% at 50∘ C. The capacity to tolerate high temperature
drastically decreased (5%) at 55∘ C and only Bacillus and
Pseudomonas strains showed this ability [67, 68]. Moderate
halotolerance was presented by 75% of the isolates, while 50%
tolerated up to 15% NaCl, 20% actively grew in presence of
20% NaCl, and only the 6% were extremely halotolerant (25%
NaCl), indicating salinity as a major selective factor for the
bacterial microbiomes in the Tunisian date palm oases. The
formulation of halotolerant PGPR could be an interesting
alternative for agriculture productivity in the oasis [69].
7
The tested rhizobacteria could grow in a wide pH range.
Within the bacterial collection 96% and the 75% of the strains
were facultative alkalophiles able to grow in basic media (up
to pH = 12), while 34% of them could grow in acidic media
(pH = 4) and only 6% was facultative acidophiles growing
down to pH = 3.
4. Conclusion
Date palm represents the key plant species in desert oases
being essential in determining the oasis microclimate that can
allow agriculture. Palm exerts both physical and functional
services involved in the creation of ideal condition for desert
farming. Palm root system and rhizosphere soil showed a
complex diversity that enclosed a reservoir of PGP bacteria
involved in the regulation of plant homeostasis. Future
work is needed to perform experiment about the ability of
selected bacterial isolates in promoting plant growth under
greenhouse and field conditions. In this context, the selection
of autochthonous bacteria, together with the desert farming
practices, could have promising perspectives for sustainable
agriculture in oasis ecosystem.
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
Authors’ Contribution
Raoudha Ferjani, Ramona Marasco, and Eleonora Rolli
contributed equally to the work.
Acknowledgments
This work was supported by the project BIODESERT GA245746 “Biotechnology from desert microbial extremophiles
for supporting agriculture research potential in Tunisia and
Southern Europe” (European Union), Fondazione Project
BIOGESTECA n∘ 15083/RCC “Fondo per la promozione
di accordi istituzionali” (Regione Lombardia, Italy) through
a fellowship to RM. ER was supported by Università degli
Studi di Milano, DeFENS, European Social Fund (FSE), and
Regione Lombardia (contract “Dote Ricerca”). Thanks are
due to Marco Fusi for invaluable help in statistical analysis.
Research reported in this publication was supported by
the King Abdullah University of Science and Technology
(KAUST).
References
[1] C. T. Chao and R. R. Krueger, “The date palm (Phoenix dactylifera L.): overview of biology, uses, and cultivation,” HortScience,
vol. 42, no. 5, pp. 1077–1082, 2007.
[2] H. de Haas, Migration and Agricultural Transformations in the
Oases of Morocco and Tunisia, KNAG, Utrecht, The Netherlands, 2001.
8
[3] M. A. Elhoumaizi, M. Saaidi, A. Oihabi, and C. Cilas, “Phenotypic diversity of date-palm cultivars (Phoenix dactylifera L.)
from Morocco,” Genetic Resources and Crop Evolution, vol. 49,
no. 5, pp. 483–490, 2002.
[4] H. Hammadi, R. Mokhtar, E. Mokhtar, and F. Ali, “New
approach for the morphological identification of date palm
(Phoenix dactylifera L.) cultivars from Tunisia,” Pakistan Journal
of Botany, vol. 41, no. 6, pp. 2671–2681, 2009.
[5] Z. Sen, A. Al Alsheikh, A. M. Al-Dakheel et al., “Climate change
and Water Harvesting possibilities in arid regions,” International Journal of Global Warming, vol. 3, no. 4, pp. 355–371, 2011.
[6] C. M. Ryu, M. A. Farag, C. H. Hu et al., “Bacterial volatiles promote growth in Arabidopsis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 100, no. 8,
pp. 4927–4932, 2013.
[7] A. de Zelicourt, M. Al-Yousif, and H. Hirt, “Rhizosphere
microbes as essential partners for plant stress tolerance,” Molecular Plant, vol. 6, no. 2, pp. 242–245, 2013.
[8] R. Marasco, E. Rolli, B. Ettoumi et al., “A drought resistancepromoting microbiome is selected by root system under desert
farming,” PLoS ONE, vol. 7, no. 10, Article ID e48479, 2012.
[9] M. Köberl, H. Müller, E. M. Ramadan, and G. Berg, “Desert
farming benefits from microbial potential in arid soils and
promotes diversity and plant health,” PLoS ONE, vol. 6, no. 9,
Article ID e24452, 2011.
[10] K. Kumar, N. Amaresan, S. Bhagat, K. Madhuri, and R. C. Srivastava, “Isolation and characterization of rhizobacteria associated with coastal agricultural ecosystem of rhizosphere soils of
cultivated vegetable crops,” World Journal of Microbiology and
Biotechnology, vol. 27, no. 7, pp. 1625–1632, 2011.
[11] R. Marasco, E. Rolli, M. Fusi et al., “Plant growth promotion
potential is equally represented in diverse grapevine rootassociated bacterial communities from different biopedoclimatic environments,” BioMed Research International, vol. 2013,
Article ID 491091, 17 pages, 2013.
[12] A. A. Othman, W. M. Amer, M. Fayez, and N. A. Hegazi, “Rhizosheath of sinai desert plants is a potential repository for
associative diazotrophs,” Microbiological Research, vol. 159, no.
3, pp. 285–293, 2004.
[13] V. Saul-Tcherkas, A. Unc, and Y. Steinberger, “Soil microbial
diversity in the vicinity of desert shrubs,” Microbial Ecology, vol.
65, no. 3, pp. 689–699, 2013.
[14] G. Muyzer, E. C. de Waal, and A. G. Uitterlinden, “Profiling
of complex microbial populations by denaturing gradient gel
electrophoresis analysis of polymerase chain reaction-amplified
genes coding for 16S rRNA,” Applied and Environmental Microbiology, vol. 59, no. 3, pp. 695–700, 1993.
[15] V. C. Sheffield, D. R. Cox, L. S. Lerman, and R. M. Myers,
“Attachment of a 40-base-pair G + C-rich sequence (GC-clamp)
to genomic DNA fragments by the polymerase chain reaction
results in improved detection of single-base changes,” Proceedings of the National Academy of Sciences of the United States of
America, vol. 86, no. 1, pp. 232–236, 1989.
[16] G. Merlino, A. Rizzi, A. Schievano et al., “Microbial community structure and dynamics in two-stage vs single-stage thermophilic anaerobic digestion of mixed swine slurry and market
bio-waste,” Water Research, vol. 47, no. 6, pp. 1983–1995, 2013.
[17] I. Fhoula, A. Najjari, Y. Turki, S. Jaballah, A. Boudabous, and
H. Ouzari, “Diversity and antimicrobial properties of lactic acid
bacteria isolated from rhizosphere of olive trees and desert
truffles of tunisia,” BioMed Research International, vol. 2013,
Article ID 405708, 14 pages, 2013.
BioMed Research International
[18] S. F. Altschul, W. Gish, W. Miller, E. W. Myers, and D. J. Lipman,
“Basic local alignment search tool,” Journal of Molecular Biology,
vol. 215, no. 3, pp. 403–410, 1990.
[19] K. Tamura, D. Peterson, N. Peterson, G. Stecher, M. Nei, and
S. Kumar, “MEGA5: molecular evolutionary genetics analysis
using maximum likelihood, evolutionary distance, and maximum parsimony methods,” Molecular Biology and Evolution,
vol. 28, no. 10, pp. 2731–2739, 2011.
[20] N. Saitou and M. Nei, “The neighbor-joining method: a new
method for reconstructing phylogenetic trees,” Molecular Biology and Evolution, vol. 4, no. 4, pp. 406–425, 1987.
[21] H. Ouzari, A. Khsairi, N. Raddadi et al., “Diversity of auxinproducing bacteria associated to Pseudomonas savastanoiinduced olive knots,” Journal of Basic Microbiology, vol. 48, no.
5, pp. 370–377, 2008.
[22] C. S. Nautiyal, “An efficient microbiological growth medium
for screening phosphate solubilizing microorganisms,” FEMS
Microbiology Letters, vol. 170, no. 1, pp. 265–270, 1999.
[23] S. Pérez-Miranda, N. Cabirol, R. George-Téllez, L. S. ZamudioRivera, and F. J. Fernández, “O-CAS, a fast and universal
method for siderophore detection,” Journal of Microbiological
Methods, vol. 70, no. 1, pp. 127–131, 2007.
[24] J. C. Cappuccino and N. Sherman, Microbiology: A Laboratory
Manual, Benjamin Cummings, New York, NY, USA, 3rd edition, 1992.
[25] M. E. Cattelan, P. G. Hartel, and J. J. Fuhrmann, “Screening for
plant growth-promoting rhizobacteria to promote early soybean growth,” Soil Science Society of America Journal, vol. 63,
no. 6, pp. 1670–1680, 1999.
[26] M. J. Anderson, “A new method for non-parametric multivariate analysis of variance,” Austral Ecology, vol. 26, no. 1, pp. 32–
46, 2001.
[27] M. J. Anderson, DISTLM v.2: A FORTRAN Computer Program
to Calculate a Distance-Based Multivariate Analysis for a Linear
Model, Department of Statistics, University of Auckland, 2002.
[28] H. Bozdogan, “Model selection and Akaike’s Information Criterion (AIC): the general theory and its analytical extensions,”
Psychometrika. A Journal of Quantitative Psychology, vol. 52, no.
3, pp. 345–370, 1987.
[29] K. Clarke and R. Gorley, “PRIMER v6.” User Manual/Tutorial,
Plymouth Routine in Multivariate Ecological Research, Plymouth
Marine Laboratory, 2006.
[30] M. J. Anderson, R. N. Gorley, and K. R. Clarke, PERMANOVA+ for PRIMER: Guide to Software and Statistical Methods,
PRIMER-E, Plymouth, UK, 2008.
[31] W. M. Cook, K. T. Lane, B. L. Foster, and R. D. Holt, “Island
theory, matrix effects and species richness patterns in habitat
fragments,” Ecology Letters, vol. 5, no. 5, pp. 619–623, 2002.
[32] T. S. Walker, H. P. Bais, E. Grotewold, and J. M. Vivanco, “Root
exudation and rhizosphere biology,” Plant Physiology, vol. 132,
no. 1, pp. 44–51, 2003.
[33] R. Angel, M. I. M. Soares, E. D. Ungar, and O. Gillor, “Biogeography of soil archaea and bacteria along a steep precipitation
gradient,” ISME Journal, vol. 4, no. 4, pp. 553–563, 2010.
[34] A. Bachar, M. I. M. Soares, and O. Gillor, “The effect of resource
islands on abundance and diversity of bacteria in arid soils,”
Microbial Ecology, vol. 63, no. 3, pp. 694–700, 2012.
[35] S. C. Cary, I. R. McDonald, J. E. Barrett, and D. A. Cowan,
“On the rocks: The microbiology of Antarctic Dry Valley soils,”
Nature Reviews Microbiology, vol. 8, no. 2, pp. 129–138, 2010.
BioMed Research International
[36] T. Caruso, Y. Chan, D. C. Lacap, M. C. Y. Lau, C. P. McKay, and S.
B. Pointing, “Stochastic and deterministic processes interact in
the assembly of desert microbial communities on a global scale,”
ISME Journal, vol. 5, no. 9, pp. 1406–1413, 2011.
[37] M. Köberl, E. M. Ramadan, M. Adam et al., “Bacillus and Streptomyces were selected as broad-spectrum antagonists against
soilborne pathogens from arid areas in egypt,” FEMS Microbiology Letters, vol. 342, no. 2, pp. 168–178, 2013.
[38] E. Armada, A. Roldán, and R. Azcon, “Differential activity of
autochthonous bacteria in controlling drought stress in native
lavandula and salvia plants species under drought conditions in
natural arid soil,” Microbial Ecology, vol. 67, no. 2, pp. 410–420,
2014.
[39] A. Bachar, A. Al-Ashhab, M. I. M. Soares et al., “Soil microbial
abundance and diversity along a low precipitation gradient,”
Microbial Ecology, vol. 60, no. 2, pp. 453–461, 2010.
[40] H. Bertrand, R. Nalin, R. Bally, and J.-C. Cleyet-Marel, “Isolation and identification of the most efficient plant growthpromoting bacteria associated with canola (Brassica napus),”
Biology and Fertility of Soils, vol. 33, no. 2, pp. 152–156, 2001.
[41] M. A. Siddikee, P. S. Chauhan, R. Anandham, G.-H. Han, and
T. Sa, “Isolation, characterization, and use for plant growth promotion under salt stress, of ACC deaminase-producing halotolerant bacteria derived from coastal soil,” Journal of Microbiology
and Biotechnology, vol. 20, no. 11, pp. 1577–1584, 2010.
[42] F. Mapelli, R. Marasco, E. Rolli et al., “Potential for plant growth
promotion of rhizobacteria associated with Salicornia growing
in Tunisian hypersaline soils,” BioMed Research International,
vol. 2013, Article ID 248078, 13 pages, 2013.
[43] D. Daffonchio, A. De Biase, A. Rizzi, and C. Sorlini, “Interspecific, intraspecific and interoperonic variability in the 16S rRNA
gene of methanogens revealed by length and single-strand conformation polymorphism analysis,” FEMS Microbiology Letters,
vol. 164, no. 2, pp. 403–410, 1998.
[44] D. Daffonchio, S. Borin, G. Frova, P. L. Manachini, and C.
Sorlini, “PCR fingerprinting of whole genomes: the spacers
between the 16s and 23S rRNA genes and of intergenic tRNA
gene regions reveal a different intraspecific genomic variability
of Bacillus cereus and Bacillus licheniformis,” International Journal of Systematic Bacteriology, vol. 48, no. 1, pp. 107–116, 1998.
[45] D. Daffonchio, A. Cherif, and S. Borin, “Homoduplex and heteroduplex polymorphisms of the amplified ribosomal 16S-23S
internal transcribed spacers describe genetic relationships in
the “Bacillus cereus group”,” Applied and Environmental Microbiology, vol. 66, no. 12, pp. 5460–5468, 2000.
[46] V. Gürtler and V. A. Stanisich, “New approaches to typing
and identification of bacteria using the 16S-23S rDNA spacer
region,” Microbiology, vol. 142, no. 1, pp. 3–16, 1996.
[47] G. Berg and K. Smalla, “Plant species and soil type cooperatively
shape the structure and function of microbial communities in
the rhizosphere,” FEMS Microbiology Ecology, vol. 68, no. 1, pp.
1–13, 2009.
[48] F. Garcia-Pichel, S. L. Johnson, D. Youngkin, and J. Belnap,
“Small-scale vertical distribution of bacterial biomass and
diversity in biological soil crusts from arid lands in the Colorado
Plateau,” Microbial Ecology, vol. 46, no. 3, pp. 312–321, 2003.
[49] P. Joshi and A. B. Bhatt, “Diversity and function of plant growth
promoting Rhizobacteria associated with wheat Rhizosphere in
North Himalayan Region,” International Journal of Environmental Sciences, vol. 1, no. 6, p. 1135, 2011.
9
[50] D. Muleta, F. Assefa, K. Hjort, S. Roos, and U. Granhall, “Characterization of rhizobacteria isolated from wild Coffea arabica
L,” Engineering in Life Sciences, vol. 9, no. 2, pp. 100–108, 2009.
[51] G. Kumar, N. Kanaujia, and A. Bafana, “Functional and phylogenetic diversity of root-associated bacteria of Ajuga bracteosa
in Kangra valley,” Microbiological Research, vol. 167, no. 4, pp.
220–225, 2012.
[52] D. El Hidri, A. Guesmi, A. Najjari et al., “Cultivation-dependant
assessment, diversity, and ecology of haloalkaliphilic bacteria
in arid saline systems of southern Tunisia,” BioMed Research
International, vol. 2013, Article ID 648141, 15 pages, 2013.
[53] P. Alavi, M. R. Starcher, C. Zachow, H. Müller, and G. Berg,
“Root-microbe systems: the effect and mode of interaction
of stress protecting agent (SPA) Stenotrophomonas rhizophila
DSM14405T,” Frontiers in Plant Science, vol. 4, p. 141, 2013.
[54] O. Barazani and J. Friedman, “Is IAA the major root growth
factor secreted from plant-growth-mediating bacteria?” Journal
of Chemical Ecology, vol. 25, no. 10, pp. 2397–2406, 1999.
[55] P. Trivedi, T. Spann, and N. Wang, “Isolation and characterization of beneficial bacteria associated with citrus roots in
Florida,” Microbial Ecology, vol. 62, no. 2, pp. 324–336, 2011.
[56] R. Marasco, E. Rolli, G. Vigani et al., “Are drought-resistance
promoting bacteria cross-compatible with different plant models?” Plant Signaling and Behavior, vol. 8, no. 10, Article ID
e26741, 2013.
[57] D. P. Schachtman, R. J. Reid, and S. M. Ayling, “Phosphorus
uptake by plants: from soil to cell,” Plant Physiology, vol. 116, no.
2, pp. 447–453, 1998.
[58] P. Vyas, P. Rahi, and A. Gulati, “Stress tolerance and genetic
variability of phosphate-solubilizing fluorescent Pseudomonas
from the cold deserts of the trans-himalayas,” Microbial Ecology,
vol. 58, no. 2, pp. 425–434, 2009.
[59] K. W. Ndung’u-Magiroi, L. Herrmann, J. R. Okalebo, C. O. Othieno, P. Pypers, and D. Lesueur, “Occurrence and genetic diversity of phosphate-solubilizing bacteria in soils of differing
chemical characteristics in Kenya,” Annals of Microbiology, vol.
62, no. 3, pp. 897–904, 2012.
[60] M. Fürnkranz, H. Müller, and G. Berg, “Characterization of
plant growth promoting bacteria from crops in Bolivia,” Journal
of Plant Diseases and Protection, vol. 116, no. 4, pp. 149–155, 2009.
[61] C. M. Ribeiro and E. J. B. N. Cardoso, “Isolation, selection and
characterization of root-associated growth promoting bacteria in Brazil Pine (Araucaria angustifolia),” Microbiological
Research, vol. 167, no. 2, pp. 69–78, 2012.
[62] F. Tian, Y. Ding, H. Zhu, L. Yao, and B. Du, “Genetic diversity
of siderophore-producing Bacteria of tobacco rhizosphere,”
Brazilian Journal of Microbiology, vol. 40, no. 2, pp. 276–284,
2009.
[63] A. Sharma and B. N. Johri, “Growth promoting influence of
siderophore-producing Pseudomonas strains GRP3A and PRS9
in maize (Zea mays L.) under iron limiting conditions,” Microbiological Research, vol. 158, no. 3, pp. 243–248, 2003.
[64] D. J. M. Kumar, L. Lawrence, R. Rajan, S. Priyadarshini, S. Yachittybabu, and P. T. Kalaichelvan, “Characterization of lipase and
protease from Serratia marcescens DEPTK21 and its destaining
capability,” Asian Journal of Experimental Biological Sciences,
vol. 3, no. 3, pp. 621–628, 2012.
[65] P. Michaud, A. Belaich, B. Courtois, and J. Courtois, “Cloning,
sequencing and overexpression of a Sinorhizobium meliloti
M5N1CS carboxymethyl-cellulase gene,” Applied Microbiology
and Biotechnology, vol. 58, no. 6, pp. 767–771, 2002.
10
[66] P. A. Wani, M. S. Khan, and A. Zaidi, “Synergistic effects of the
inoculation with nitrogen-fixing and phosphate-solubilizing
rhizobacteria on the performance of field-grown chickpea,”
Journal of Plant Nutrition and Soil Science, vol. 170, no. 2, pp.
283–287, 2007.
[67] P. Scheldeman, L. Herman, S. Foster, and M. Heyndrickx,
“Bacillus sporothermodurans and other highly heat-resistant
spore formers in milk,” Journal of Applied Microbiology, vol. 101,
no. 3, pp. 542–555, 2006.
[68] S. S. Kanwar, M. Gupta, R. Gupta, R. K. Kaushal, and S. S.
Chimni, “Properties of hydrogel-entrapped lipase of thermophilic Pseudomonas aeruginosa BTS-2,” Indian Journal of Biotechnology, vol. 5, no. 3, pp. 292–297, 2006.
[69] I. L. D. Diatta, A. Kane, C. E. Agbangba et al., “Inoculation with
arbuscular mycorrhizal fungi improves seedlings growth of two
sahelian date palm cultivars (Phoenix dactylifera L., cv. Nakhla
hamra and cv. Tijib) under salinity stresses,” Advances in Bioscience and Biotechnology, vol. 5, no. 1, pp. 64–72, 2014.
BioMed Research International
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 296472, 9 pages
http://dx.doi.org/10.1155/2014/296472
Research Article
Pentachlorophenol Degradation by Janibacter sp., a New
Actinobacterium Isolated from Saline Sediment of Arid Land
Amel Khessairi,1,2 Imene Fhoula,1 Atef Jaouani,1 Yousra Turki,2 Ameur Cherif,3
Abdellatif Boudabous,1 Abdennaceur Hassen,2 and Hadda Ouzari1
1
Université Tunis El Manar, Faculté des Sciences de Tunis (FST), LR03ES03 Laboratoire de Microorganisme et
Biomolécules Actives, Campus Universitaire, 2092 Tunis, Tunisia
2
Laboratoire de Traitement et Recyclage des Eaux, Centre des Recherches et Technologie des Eaux (CERTE),
Technopôle Borj-Cédria, B.P. 273, 8020 Soliman, Tunisia
3
Université de Manouba, Institut Supérieur de Biotechnologie de Sidi Thabet, LR11ES31 Laboratoire de Biotechnologie et
Valorization des Bio-Geo Resources, Biotechpole de Sidi Thabet, 2020 Ariana, Tunisia
Correspondence should be addressed to Hadda Ouzari; [email protected]
Received 1 May 2014; Accepted 17 August 2014; Published 17 September 2014
Academic Editor: George Tsiamis
Copyright © 2014 Amel Khessairi et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Many pentachlorophenol- (PCP-) contaminated environments are characterized by low or elevated temperatures, acidic or alkaline
pH, and high salt concentrations. PCP-degrading microorganisms, adapted to grow and prosper in these environments, play an
important role in the biological treatment of polluted extreme habitats. A PCP-degrading bacterium was isolated and characterized
from arid and saline soil in southern Tunisia and was enriched in mineral salts medium supplemented with PCP as source of carbon
and energy. Based on 16S rRNA coding gene sequence analysis, the strain FAS23 was identified as Janibacter sp. As revealed by high
performance liquid chromatography (HPLC) analysis, FAS23 strain was found to be efficient for PCP removal in the presence of 1%
of glucose. The conditions of growth and PCP removal by FAS23 strain were found to be optimal in neutral pH and at a temperature
of 30∘ C. Moreover, this strain was found to be halotolerant at a range of 1–10% of NaCl and able to degrade PCP at a concentration
up to 300 mg/L, while the addition of nonionic surfactant (Tween 80) enhanced the PCP removal capacity.
1. Introduction
The polyorganochlorophenolic (POP) compounds have been
extensively used as wide spectrum biocides in industry
and agriculture [1]. The toxicity of these compounds tends
to increase according to their degree of chlorination [2].
Among chlorinated phenols, pentachlorophenol (PCP) has
been widely used as wood and leather preservative, owing
to its toxicity toward bacteria, mould, algae, and fungi [3].
However, PCP is also toxic to all forms of life since it is
an inhibitor of oxidative phosphorylation [4]. The extensive
exposure to PCP could cause cancer, acute pancreatitis,
immunodeficiency, and neurological disorders [5]. Consequently, this compound is listed among the priority pollutants
of the US Environmental Protection Agency [6]. Moreover, it
is recalcitrant to degradation because of its stable aromatic
ring and high chloride contents, thus persisting in the
environment [7]. Although contamination of soils and waters
with chemically synthesized PCP is a serious environmental
problem, their remediation may be possible using physical, chemical, and biological methods [8]. Bioremediation
represents a choice process, thanks to its low costs and
reduction of toxic residue generated in the environment. The
biodegradation of PCP has been studied in both aerobic and
anaerobic systems. Aerobic degradation of PCP especially
has been extensively studied and several bacterial isolates
were found to degrade and use PCP as a sole source of
carbon and energy. The most studied aerobic PCP-degrading
microorganisms included Mycobacterium chlorophenolicum
[9], Alcaligenes sp. [10], Rhodococcus chlorophenolicus [11],
Flavobacterium [12], Novosphingobium lentum [13] and Sphingomonas chlorophenolica [14], Bacillus [15], Pseudomonas
[16], and Acinetobacter [17], as well as some fungi species.
Saline and arid environments are found in a wide variety
2
of aquatic and terrestrial ecosystems. A low taxonomic
biodiversity is observed in all these saline environments [18],
most probably due to the high salt concentrations prevailing
in these environments. Moreover, the biodegradation process
is difficult to perform under saline conditions [19]. Besides
these metabolical and physiological features, halophilic and
halotolerant microorganisms are known to play important
roles in transforming and degrading waste and organic pollutants in saline and arid environment [20]. These microorganisms, particularly actinobacteria, are frequently isolated
from extreme environments such as Sabkha, Chott, and
Sahara which are known to have a great metabolic diversity
and biotechnological potential. The occurrence of actinobacteria in saline environment and their tolerance to high
salt concentrations were thus described [21]. However, few
actinobacteria genera, such as Arthrobacter [22] and Kocuria
[23], were reported for PCP-degradation process. The genus
Janibacter which is recognized by Martin et al. [24] belongs to
the family Intrasporangiaceae in the Actinomycetales order
and included five major species, J. limosus [24], J. terrae
[25], J. melonis [26], J. corallicola [27], and J. anophelis [28].
Interestingly, most of these species were reported for their
ability to degrade a large spectrum of aromatic and/or chlorinated compounds including polychlorinated biphenyls [29],
monochlorinated dibenzo-p-dioxin [30], dibenzofuran [31],
anthracene, phenanthrene [32], dibenzo-p-dioxin, carbazole,
diphenyl ether, fluorene [33], and polycyclic aromatic hydrocarbons [34]. However, no data reporting PCP degradation
by Janibacter members was described. PCP and other POP
compounds shared many physical properties, which limited
biodegradation processes, and one of these properties was
their lower solubility and therefore low bioavailability to the
degrading bacteria. Nevertheless, the use of surfactants such
as Tween 80 has the potential to increase the biodegradation
rates of hydrophobic organic compounds by increasing the
total aqueous solubility of these pesticides [35]. In this study,
we evaluated for the first time the PCP removal potential,
under different physicochemical conditions, by Janibacter sp.,
a halotolerant actinobacterium member isolated from arid
and saline land in southern Tunisia.
2. Materials and Methods
2.1. Chemicals and Solvents. PCP (MW 266.34 and >99%
purity) and acetonitrile (HPLC grade) were purchased from
Sigma Aldrich (USA). All other inorganic chemicals used to
prepare the different media are commercially available with
highest purity and are used without further purification.
2.2. Sample Collection and PCP-Degrading Bacterium Isolation. The sediment samples were collected in March 2011
from arid and saline ecosystems belonging to the site “Sebkha
El Naouel” with GPS coordinates: N 34∘ 26󸀠 951󸀠󸀠 E 09∘ 54󸀠 102󸀠󸀠
altitude 150 ft/46 m, in southern Tunisia. Bacterial isolation
was performed as described by Rösch et al. [36] with some
modifications: 10 g of soil sample was suspended in 100 mL of
phosphate-buffered salt solution (137 mM NaCl, 2.7 mM KCl,
10 mM Na2 HPO4 , and 2 mM KH2 PO4 ) and stirred vigorously
for 30 min. The soil suspension was diluted and 0.1 mL sample
BioMed Research International
was spread on the surface of yeast extract-mannitol medium
(YEM). YEM medium contained the following components
at the specified concentrations (in g/L): mannitol, 5; yeast
extract, 0.5; MgSO4 ⋅7H2 O, 0.2; NaCl, 0.1; K2 HPO4 , 0.5; Na
gluconate, 5; agar, 15; pH = 6, 8. After sterilization for 20 min
at 120∘ C, 1 mL of 16.6% CaCl2 solution was added to 1 liter
of YEM medium (1 : 1000). The plates were then incubated at
30∘ C for 7 days. Pure cultures of the isolates were obtained by
streaking a single colony on the same medium.
2.3. 16S rRNA Gene Amplification and Sequence Analysis.
For DNA extraction, the FAS23 strain was grown in tryptic
soy broth (TSB) containing (in g/L) casein peptone, 17; soya
peptone, 3; glucose, 2.5; sodium chloride, 5; dipotassium
hydrogen phosphate, 4. DNA extraction was performed
using CTAB/NaCl method as described by Wilson [37] and
modified by using lysozyme (1 mg/mL) for cell wall digestion.
The 16S rRNA gene was amplified using universal primers
SD-Bact-0008-a-S-20 (5󸀠 -AGA GTT TGA TCC TGG CTC
AG-󸀠 3) and S-D-Bact-1495-a-A-20 (5󸀠 -CTA CGG CTA CCT
TGT TAC GA-󸀠 3) [38]. PCR was performed in a final volume
of 25 𝜇L containing 1 𝜇L of the template DNA; 0.5 𝜇M of each
primer; 0.5 𝜇M of deoxynucleotide triphosphate (dNTP);
2.5 𝜇L 10X PCR buffer for Taq polymerase; MgCl2 1.5 mM;
1 UI of Taq polymerase. The amplification cycle was as
follows: denaturation step at 94∘ C for 3 min, followed by 35
cycles (45 sec at 94∘ C, 1 min at 55∘ C, and 2 min at 72∘ C) plus
one additional cycle at 72∘ C for 7 min as a final elongation
step. The 16S rDNA PCR amplicons were purified with
Exonuclease-I and Shrimp Alkaline Phosphatase (Exo-Sap,
Fermentas, Life Sciences) following the manufacturer’s standard protocol. Sequence analyses of the purified DNAs were
performed using a Big Dye Terminator cycle sequencing kit
V3.1 (Applied Biosystems) and an Applied Biosystems 3130XL
Capillary DNA Sequencer machine. Sequence similarities
were found by BLAST analysis [39] using the GenBank DNA
databases (http://www.ncbi.nlm.nih.gov/) and the Ribosomal Database Project (RDP). Phylogenetic analyses of the 16S
rRNA gene sequences were conducted with Molecular Evolutionary Genetics Analysis (MEGA) software, version 5 [40].
Trees were constructed by using neighbor-joining method
[41]. The sequence was deposited in GenBank database under
the accession number KC959984.
2.4. Degradation of PCP by Isolated Strain. The kinetics of
the PCP removal under different conditions were conducted
in 500 mL flasks, sealed with cotton stoppers, containing
100 mL of mineral salt medium (MSM) adjusted to pH
6.9, supplemented with 1% glucose and inoculated with
1% of 106 CFU/mL of the strain FAS23. The MSM contained the following components at the specified concentrations (in g/L): KH2 PO4 , 0.8; Na2 HP4 , 0.8; MgSO4 ⋅7H2 O,
0.2; CaCl2 ⋅2H2 O, 0.01; NH4 Cl, 0.5, plus 1 mL of trace
metal solution which includes (in mg/L) FeSO4 ⋅7H2 O, 5;
ZnSO4 ⋅7H2 O, 4; MnSO4 ⋅4H2 O, 0.2; NiCl⋅6H2 O, 0.1; H3 BO3 ,
0.15; CoCl2 ⋅6H2 O, 0.5; ZnCl2 0.25; and EDTA, 2.5. PCP
was added to the medium after autoclaving [19]. When
necessary, solid MSM plates were prepared by adding 15 g/L
bacteriological grade agar. The inoculum was prepared as
BioMed Research International
3
Janibacter melonis (JX865444)
Janibacter marinus (AY533561)
Janibacter terrae (KC469957)
52 Janibacter hoylei (FR749912)
Janibacter sanguinis (JX435047)
58
Janibacter corallicola (NR 041558)
Janibacter sp. FAS23
100
82 Janibacter sp. ATCC 33790 (GU933618)
Janibacter limosus (KC469951)
Terrabacter tumescens (NR 044984)
76
99 Terracoccus luteus (NR 026412)
Kocuria rhizophila (NR 026452)
Arthrobacter chlorophenolicus (EU102284)
40
0.01
Figure 1: The phylogenetic position of Janibacter sp. strain in relation to some members of actinobacteria (genus of Janibacter, Sphingomonas,
Terrabacter, Terracoccus, Kocuria, and Arthrobacter) based on 16S rRNA gene. Bootstrap values for a total of 1000 replicates are shown at the
nodes of the tree. The scale bar corresponds to 0.05 units of the number of base substitutions per site changes per nucleotide.
follows: overnight culture was centrifuged and the pellet was
rinsed twice with fresh MSM. PCP removal was monitored
during 144 h of incubation by varying different parameters:
(i) initial pH; (ii) initial PCP concentrations: 20, 50, 100,
200, and 300 mg/L corresponding to 0.075 mM, 0.19 mM,
0.37, 0.75 mM, and 1.14 mM, respectively; (iii) temperature
of incubation: 25, 30, and 37∘ C; (iv) NaCl concentrations:
10 g/L, 30 g/L, 60 g/L, and 100 g/L; (v) the addition of nonionic
surfactant Tween 80 (40 mg/L). Bacterial cell growth was
evaluated by measuring the optical density at 600 nm using
UV-VIS spectrophotometer (Spectro UVS-2700 Dual Beam
Labomed, Inc) every 24 h of the incubation. Three controls
were used: PCP-free MSM, uninoculated PCP containing
MSM, and PCP containing MSM inoculated with heated
inactivated cells. The cell suspension was centrifuged (5 min,
8000 rpm) and the supernatant was filtered through 0.22 𝜇m
filters [16]. Samples of 100 𝜇L were applied to C18 reverse
phase column (LiChrospher 100 RP-18 endcapped column,
250 mm × 4.6 mm i.d., and particle size of 5 𝜇m) at a flow
rate of 1 mL min−1 . The retained molecules were eluted over
35 min using the following gradient: 1% (v/v) phosphoric
acid in water for 4 min, followed by an increase to 100%
(v/v) acetonitrile within 21 min which was kept constant for
5 min and then decreased back to initial concentration and
kept constant for another 5 min. PCP was quantified using
external standards method. Percent removal was estimated
using the following formula: removal (%) = area − area/area
[42].
2.5. Statistical Analysis. Data were subjected to analysis of
variance using SPSS software (version 14.0) and the mean
differences were compared by Student-Newman-Keuls comparison test. A 𝑃 value of less than 0.05 was considered
statistically significant (test at 𝑃 < 0.05). Three replicates were
prepared for each treatment.
3. Results
3.1. Isolation, Identification of FAS23 Strain, and 16S rDNA
Sequence Based Phylogenetic Analysis. The bacterial strain
FAS23 was isolated from the saline and arid sediment.
The morphological aspect of FAS23 strain culture on the
isolation medium YEM showed opaque, pale, cream, and
convex colonies with glistening surface. Cells were Grampositive, rod-shaped, and positive for catalase and oxidase
tests. No growth under anaerobic conditions and no spore
formation were recorded. The optimal growth conditions of
FAS23 strain were pH of 7.0–8.5 and a temperature range
of 28–30∘ C. The strain was able to grow at a range of
salt concentrations from 1 to 100 g/L of NaCl. 16S rDNA
sequencing and phylogenetic analysis allowed the assignment
of FAS23 strain to Janibacter sp. (Figure 1).
3.2. The Optimum Growth Conditions of Janibacter sp. Strain.
The effect of physiological and biochemical variations (glucose supplement, temperature, pH, PCP concentration, and
presence of biosurfactant) on bacterial growth of Janibacter
sp. FAS23 and PCP removal was studied.
3.2.1. Effect of Glucose on the Growth of Janibacter sp. and PCP
Removal. The effect of glucose as cosubstrate on the growth
of Janibacter sp. strain and PCP removal was studied in MSM.
The result showed that the growth of the strain was possible
only after the addition of glucose (Figure 2(a)). As well, the
PCP was efficiently removed in the presence of glucose, and
71.84% of PCP was degraded within 24 hours and more than
90% after 72 h (Figure 2(b)). The obtained results indicated
the phenomenon of cometabolism in which microorganisms
do not obtain energy from the transformation reaction; they
rather require another substrate for growth [43].
3.2.2. Effect of pH and Temperature on the Growth of the
Strain and PCP Removal. The effect of pH variations (4.0,
6.9, and 9.0) on the growth and PCP removal was assessed
(Figure 3). At both pH 4.0 and 9.0, a low rate of growth and
PCP removal was observed after 24 and 48 h of incubation.
However, after 144 h of incubation, the rate of PCP removal
has reached values of 44.80% and 70.22% at pH 4.0 and pH
9.0, respectively. The optimum growth and PCP removal were
however observed at pH 6.9, as we noted a significant removal
of PCP of 71.84%, 84.47%, and 99.06% after 24, 48, and 144 h
of incubation, respectively. The strain FAS23 was able to grow
BioMed Research International
2.0
1.8
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.0
100
Residual PCP (%)
OD at 600 nm
4
80
60
40
20
0
0
24
48
72
96
Incubation period (h)
120
0
144
24
48
72
96
Incubation period (h)
120
144
PCP + Glucose
PCP
Glucose
PCP – Glucose
PCP
(b)
(a)
Figure 2: The growth (a) and the PCP removal (b) in the presence and in deficiency of the supplementary carbon source (glucose: 1%) by
Janibacter sp. FAS23. Error bars represent the standard deviation.
100
1.8
90
1.6
80
1.4
70
Residual PCP (%)
OD at 600 nm
2.0
1.2
1.0
0.8
0.6
60
50
40
30
0.4
20
0.2
10
0.0
0
0
24
48
72
96
Incubation periods (h)
pH 4-PCP
pH 4
pH 6.9-PCP
120
144
0
24
48
72
96
120
144
Incubation period (h)
pH 6.9
pH 9-PCP
pH 9
(a)
pH 4
pH 6.9
pH 9
(b)
Figure 3: (a) Growth of Janibacter sp. at different pH of culture medium: pH 4.0, pH 6.9, and pH 9.0 with 20 mg/L of PCP at 30∘ C. (b) Effect
of different pH on the PCP removal efficiency by Janibacter sp. Error bars represent the standard deviation.
in the temperature range of 25–37∘ C, with an optimum at
30∘ C. At 25 and 37∘ C, the growth of the bacterial strain, as
well as PCP removal, was affected (Figure 4). However, at
25∘ C, the strain showed a better growth and PCP removal
compared to temperature of 37∘ C. Likewise, the PCP removal
was optimal at 30∘ C reaching 71.84% and 99.06% after 24 h
and 144 h of incubation, respectively.
3.2.3. Effect of PCP Amount on the Growth and PCP Removal
by Janibacter. Variation of PCP amount in the medium
showed that the growth of the strain, as well as PCP removal,
decreased with the increase of PCP concentration (Figure 5).
At low concentrations (20 and 50 mg/L), the bacterial strain
was able to remove the majority of PCP after 72 h of
incubation time. Up to 100 mg/L, 50% of PCP could be
removed after 72 h of incubation. However, with higher
concentrations (200 and 300 mg/L), equivalent level of PCP
removal could be reached if the incubation time is extended.
3.2.4. Effect of Various NaCl Concentrations on the PCP
Removal. The strain was tested for its ability to remove PCP
(20 mg/L) at different NaCl concentrations (0%, 1%, 3%,
6%, and 10%). The best rate of growth and PCP removal
was recorded at 1% of NaCl (more than 92% after 144 h
of incubation). The growth and thus the capacity of PCP
removal were inhibited when the concentration of sodium
chloride was increased (Figure 6). At 3% NaCl, the PCP
removal was 72% after 144 h of incubation. When the NaCl
BioMed Research International
5
2.0
100
90
1.6
80
1.4
70
Residual PCP (%)
OD at 600 nm
1.8
1.2
1.0
0.8
0.6
60
50
40
30
0.4
20
0.2
10
0
0.0
0
24
48
72
96
Incubation period (h)
120
144
0
24
25∘ C
30∘ C
37∘ C
25∘ C-PCP
30∘ C-PCP
37∘ C-PCP
48
72
96
Incubation period (h)
120
144
25∘ C
30∘ C
37∘ C
(a)
(b)
Figure 4: (a) Growth of Janibacter sp. at different temperatures in presence of 20 mg/L of PCP and at pH 6.9. (b) Effect of temperature changes
on the PCP removal efficiency by Janibacter sp. Error bars represent the standard deviation.
1.8
100
1.6
90
1.4
80
Residual PCP (%)
OD at 600 nm
2.0
1.2
1.0
0.8
0.6
70
60
50
40
30
0.4
20
0.2
10
0
0.0
0
24
48
72
96
Incubation period (h)
120
144
100
200
300
0
20
50
0
24
48
72
96
Incubation period (h)
20
50
100
(a)
120
144
200
300
(b)
Figure 5: (a) Growth of Janibacter sp. in the presence of 0, 20, 50, 100, 200, and 300 mg/L of PCP. (b) Effect of different PCP concentrations
on the PCP removal by Janibacter sp. Error bars represent the standard deviation.
concentration was increased to 6% and 10%, PCP removal
falls to 46.53% and 17.62%, respectively.
of PCP (300 mg/mL) was improved by 30% after 72 h of
incubation, compared to the control (Figure 7(b)).
3.2.5. Effect of Nonionic Surfactant Tween 80 on the Biodegradation of PCP. In this study, the nonionic surfactant Tween
80 was found to enhance the growth and PCP-biodegradation
process (Figure 7). Interestingly, removal of high amount
4. Discussion
The strain FAS23 isolated from saline sediment collected
from Tunisian arid ecosystems was identified as an
BioMed Research International
2.0
1.8
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.0
Residual PCP (%)
OD at 600 nm
6
0
24
48
72
96
Incubation period (hours)
120
100
90
80
70
60
50
40
30
20
10
0
0
144
6%
10%
0%
1%
3%
24
48
72
96
120
Incubation period (hours)
144
6%
10%
0%
1%
3%
(a)
(b)
Figure 6: (a) Growth of Janibacter sp. in MS medium supplemented with different concentrations of NaCl: 0%, 1%, 3%, 6%, and 10% in the
presence of 20 mg/L of PCP at 30∘ C. (b) Effect of NaCl on the PCP removal efficiency by Janibacter sp. Error bars represent the standard
deviation.
2.0
100
1.8
90
80
1.4
Residual PCP (%)
OD at 600 nm
1.6
1.2
1.0
0.8
0.6
0.4
0.2
70
60
50
40
30
20
10
0
0.0
0
24
48
72
96
Incubation period (h)
20-T80
20
120
300-T80
300
(a)
144
0
24
48
72
96
120
144
Incubation period (h)
300-T80
300
20-T80
20
(b)
Figure 7: (a) Growth of Janibacter sp. in MS medium supplemented with nonionic surfactant Tween 80 (40 mg/L) containing 20 and 300 mg/L
of PCP. (b) Effect of nonionic surfactant Tween 80 on the PCP removal efficiency by Janibacter sp. Error bars represent the standard deviation.
actinobacterium belonging to the genus Janibacter sp., with
respect to morphological and biochemical tests and 16S
rRNA gene sequence. Despite their known high potential in
recalcitrant compounds biodegradation [29, 33], bacteria of
the genus Janibacter, described in this study, are reported
for the first time for their ability to degrade PCP. In
biodegradation process, glucose is commonly used as an
additional source of carbon and energy and is the most
metabolizable sugar which supported a maximum growth
[44]. In this context, our results are in agreement with those
of Singh et al. [45] and Singh et al. [15] who reported the
enhancement of bacterial growth and PCP-degradation
process using MSM supplemented with 1% of glucose [43].
This effect can be explained by the connection of the two
substrates metabolism. In fact, the NADH provided by
glucose metabolism may increase the biomass and thus
increase the total activity for PCP metabolizing [46].
In the present study, PCP removal is affected by pH
variation. As it was reported by Premalatha and Rajakumar
[43], Wolski et al. [47], Barbeau et al. [48], and Edgehill
[22] for Arthrobacter and different Pseudomonas species, the
neutral pH was found to be optimal for PCP degradation.
However, for other bacterial species, such as Sphingomonas
chlorophenolica, PCP degradation was more important at pH
9.2 [49].
Temperature is another important environmental factor
that can influence the rate of pollutants degradation [48].
The optimal temperature for the PCP removal was recorded
BioMed Research International
at 30∘ C, but lower temperatures (25∘ C) allowed significant
removal than the upper values. These results were in accordance with those of Wittmann et al. [9] and Crawford
and Mohn [50]. Overall, deviation in pH and temperature from the optimum results in alteration of microbial
growth and metabolism, as well as the pollutants properties
[51, 52].
The effect of different concentrations of PCP on growth of
the strain proved that the PCP removal was more efficient at
low concentrations (20 mg/L). This result was coherent with
data of Webb et al. [53] reporting that all strains tested were
able to degrade up to 90% of the PCP, when the concentration
was 10 mg/L. Moreover, as it was revealed by Karn et al.
[16], the ability of PCP removal of Janibacter sp. decreases
when PCP concentration was increased. Furthermore, we
found that the removal ability by Janibacter sp. has reached
40% after 144 h of incubation, when PCP concentration of
300 mg/L was used. These results are in accordance with
those of Chandra et al. [54] for Bacillus cereus strain and
may suggest that these bacteria may tolerate and remove high
concentrations of PCP if we increase the incubation time. On
the contrary, Kao et al. [55] reported that no PCP removal was
detected with PCP concentrations of 320 mg/L even after 20
days of incubation.
As for bioremediation, the strain should possess not only
the high removal efficiency for the target compounds but
also the strong abilities of adapting some conditions such
as pH, temperature, and salinity fluctuations. In this study,
it was shown that Janibacter sp. was able to remove PCP
even with salinity fluctuations (less than 10%). These results
were in accordance with those of Gayathri and Vasudevan
[56] suggesting that the reduction in phenolic components
removal efficiency above 10% NaCl may be due to increase
in salinity. These results indicated that Janibacter sp. strain
has an inherent flexibility to adapt to salinity fluctuations.
The use of surfactants has the potential to increase the
biodegradation rate of hydrophobic organic compounds in
contaminated environments. Nonionic surfactants are usually used in the bioavailability studies due to their relatively
low toxicity compared to ionic surfactants [57]. The enhanced
biodegradation in the micelles solution can be attributable
to the increased solubility, dissolution, and bioavailability
of compound to bacteria [58] and the surfactant enhanced
substrate transport through the microbial cell wall [59]. The
effects of the surfactants on PCP removal have been invariably attributed to the increased solubility and dissolution of
PCP or enhancement of mass transport in the presence of
surfactants. In this context, at high concentration of PCP
(300 mg/L), Tween 80 increases the removal rate of PCP when
the Tween 80 concentration is 40 mg/L. The enhancement of
PCP removal was slightly detected when the concentration
of PCP is 20 mg/L. These results can be confirmed with the
study of Cort et al. [60] when the biodegradation rate of
PCP was enhanced for the concentration of PCP at 140 and
220 mg/L but it was inhibited for the concentration of PCP
at 100 and 50 mg/L. Consequently, successful integration of
PCP and Tween 80 degradation was achieved by Janibacter
sp. strain.
7
5. Conclusion
In this study, a novel efficient PCP-degrading actinobacterium (Janibacter sp.) was isolated from saline soil of arid
land and investigated for its physiological characteristics.
Janibacter was able to remove high concentration of PCP and
to tolerate fluctuation of NaCl. This removal potential was
moreover accelerated by the addition of Tween 80. This study
suggested that strain Janibacter sp. could be widely used for
PCP bioremediation of polluted arid/extreme environments.
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
Acknowledgments
This work was financially supported by the NATO Project
SFP (ESP.MD.FFP981674) 0073 “Preventive and Remediation
Strategies for Continuous Elimination of Poly-Chlorinated
Phenols from Forest Soils and Groundwater.” It was in part
further supported by the European Union in the ambit of
the Project ULIXES (European Community’s Seventh Framework Programme, KBBE-2010-4 under Grant Agreement no.
266473).
References
[1] A. Vallecillo, P. A. Garcia-Encina, and M. Peña, “Anaerobic
biodegradability and toxicity of chlorophenols,” Water Science
and Technology, vol. 40, no. 8, pp. 161–168, 1999.
[2] S. Fetzner and F. Lingens, “Bacterial dehalogenases: biochemistry, genetics, and biotechnological applications,” Microbiological Reviews, vol. 58, no. 4, pp. 641–685, 1994.
[3] C. M. Kao, C. T. Chai, J. K. Liu, T. Y. Yeh, K. F. Chen, and S.
C. Chen, “Evaluation of natural and enhanced PCP biodegradation at a former pesticide manufacturing plant,” Water Research,
vol. 38, no. 3, pp. 663–672, 2004.
[4] D.-S. Shen, X.-W. Liu, and H.-J. Feng, “Effect of easily degradable substrate on anaerobic degradation of pentachlorophenol
in an upflow anaerobic sludge blanket (UASB) reactor,” Journal
of Hazardous Materials, vol. 119, no. 1–3, pp. 239–243, 2005.
[5] K. Sai, K. S. Kang, A. Hirose, R. Hasegawa, J. E. Trosko, and T.
Inoue, “Inhibition of apoptosis by pentachlorophenol in v-myctransfected rat liver epithelial cells: relation to down-regulation
of gap junctional intercellular communication,” Cancer Letters,
vol. 173, no. 2, pp. 163–174, 2001.
[6] EPA, “Final determination and indent to cancel and deny
applications for registrations of pesticide products containing pentachlorophenol (including but not limited to its salts
and esters) for non-wood uses,” US Environmental Protection
Agency. Federal Register, vol. 52, pp. 2282–2293, 1987.
[7] S. D. Copley, “Evolution of a metabolic pathway for degradation
of a toxic xenobiotic: the patchwork approach,” Trends in
Biochemical Sciences, vol. 25, no. 6, pp. 261–265, 2000.
[8] D. Errampalli, J. T. Trevors, H. Lee et al., “Bioremediation: a
perspective,” Journal of Soil Contamination, vol. 6, no. 3, pp.
207–218, 1997.
8
[9] C. Wittmann, A. P. Zeng, and W. D. Deckwer, “Physiological
characterization and cultivation strategies of the pentachlorophenol-degrading bacteria Sphingomonas chlorophenolica
RA2 and Mycobacterium chlorophenolicum PCP-1,” Journal
of Industrial Microbiology and Biotechnology, vol. 21, no. 6, pp.
315–321, 1998.
[10] I. S. Thakur, R. Sharma, S. Shukla, and A. H. Ahmed, “Enumeration and enrichment of pentachlorophenol degrading bacterial
of pulp and paper miller in the chemostat,” in Proceedings of
the AMI Conference, vol. 159, pp. 25–27, Jaipur, India, November
2000.
[11] J. H. A. Apajalahti, P. Karpanoja, and M. S. Salkinoja-Salonen,
“Rhodococcus chlorophenolicus sp. nov., a chlorophenol-mineralizing actinomycete,” International Journal of Systematic Bacteriology, vol. 36, no. 2, pp. 246–251, 1986.
[12] J. F. Gonzalez and W. S. Hu, “Effect of glutamate on the degradation of pentachlorophenol by Flavobacterium sp,” Applied
Microbiology and Biotechnology, vol. 35, no. 1, pp. 100–110, 1991.
[13] R. I. Dams, G. I. Paton, and K. Killham, “Rhizoremediation
of pentachlorophenol by Sphingobium chlorophenolicum ATCC
39723,” Chemosphere, vol. 68, no. 5, pp. 864–870, 2007.
[14] C.-F. Yang, C.-M. Lee, and C.-C. Wang, “Isolation and physiological characterization of the pentachlorophenol degrading
bacterium Sphingomonas chlorophenolica,” Chemosphere, vol.
62, no. 5, pp. 709–714, 2006.
[15] S. Singh, B. B. Singh, R. Chandra, D. K. Patel, and V. Rai,
“Synergistic biodegradation of pentachlorophenol by Bacillus
cereus (DQ002384), Serratia marcescens (AY927692) and Serratia marcescens (DQ002385),” World Journal of Microbiology and
Biotechnology, vol. 25, no. 10, pp. 1821–1828, 2009.
[16] S. K. Karn, S. K. Chakrabarty, and M. S. Reddy, “Pentachlorophenol degradation by Pseudomonas stutzeri CL7 in
the secondary sludge of pulp and paper mill,” Journal of
Environmental Sciences, vol. 22, no. 10, pp. 1608–1612, 2010.
[17] A. Sharma, I. S. Thakur, and P. Dureja, “Enrichment, isolation and characterization of pentachlorophenol degrading bacterium Acinetobacter sp. ISTPCP-3 from effluent discharge site,”
Biodegradation, vol. 20, no. 5, pp. 643–650, 2009.
[18] M. Kamekura, “Diversity of extremely halophilic bacteria,”
Extremophiles, vol. 2, no. 3, pp. 289–295, 1998.
[19] R. Margesin and F. Schinner, “Potential of halotolerant and
halophilic microorganisms for biotechnology,” Extremophiles,
vol. 5, no. 2, pp. 73–83, 2001.
[20] W. D. Grant, R. T. Gemmell, and T. J. McGenity, “Halophiles,”
in Extremophiles: Microbial Life in Extreme Environments, K.
Horikoshi and W. D. Grant, Eds., pp. 93–132, Wiley-Liss, New
York, NY, USA, 1998.
[21] C. K. Okoro, R. Brown, A. L. Jones et al., “Diversity of culturable
actinomycetes in hyper-arid soils of the Atacama Desert, Chile,”
Antonie van Leeuwenhoek, vol. 95, no. 2, pp. 121–133, 2009.
[22] R. U. Edgehill, “Pentachlorophenol removal from slightly acidic
mineral salts, commercial sand, and clay soil by recovered
Arthrobacter strain ATCC 33790,” Applied Microbiology and
Biotechnology, vol. 41, no. 1, pp. 142–148, 1994.
[23] S. K. Karn, S. K. Chakrabarti, and M. S. Reddy, “Degradation of
pentachlorophenol by Kocuria sp. CL2 isolated from secondary
sludge of pulp and paper mill,” Biodegradation, vol. 22, no. 1, pp.
63–69, 2011.
[24] K. Martin, P. Schumann, F. A. Rainey, B. Schuetze, and I. Groth,
“Janibacter limosus gen. nov., sp. nov., a new actinomycete
with meso- diaminopimelic acid in the cell wall,” International
BioMed Research International
[25]
[26]
[27]
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35]
[36]
[37]
[38]
[39]
Journal of Systematic Bacteriology, vol. 47, no. 2, pp. 529–534,
1997.
J.-H. Yoon, K.-C. Lee, S.-S. Kang, Y.-H. Kho, K.-H. Kang, and
Y.-H. Park, “Janibacter terrae sp. nov., a bacterium isolated from
soil around a wastewater treatment plant,” International Journal
of Systematic and Evolutionary Microbiology, vol. 50, no. 5, pp.
1821–1827, 2000.
J.-H. Yoon, H. B. Lee, S.-H. Yeo, and J.-E. Choi, “Janibacter melonis sp. nov., isolated from abnormally spoiled oriental melon
in Korea,” International Journal of Systematic and Evolutionary
Microbiology, vol. 54, no. 6, pp. 1975–1980, 2004.
A. Kageyama, Y. Takahashi, M. Yasumoto-Hirose, H. Kasai,
Y. Shizuri, and S. Omura, “Janibacter corallicola sp. nov., isolated from coral in Palau,” Journal of General and Applied
Microbiology, vol. 53, no. 3, pp. 185–189, 2007.
P. Kämpfer, O. Terenius, J. M. Lindh, and I. Faye, “Janibacter
anophelis sp. nov., isolated from the midgut of Anopheles
arabiensis,” International Journal of Systematic and Evolutionary
Microbiology, vol. 56, no. 2, pp. 389–392, 2006.
I. Sierra, J. L. Valera, M. L. Marina, and F. Laborda, “Study
of the biodegradation process of polychlorinated biphenyls in
liquid medium and soil by a new isolated aerobic bacterium
(Janibacter sp.),” Chemosphere, vol. 53, no. 6, pp. 609–618, 2003.
S. Iwai, A. Yamazoe, R. Takahashi, F. Kurisu, and O. Yagi,
“Degradation of monochlorinated dibeno -p-Dioxins by Janibacter sp. strain YA isolated from river sediment,” Current
Microbiology, vol. 51, no. 5, pp. 353–358, 2005.
S. Jin, T. Zhu, X. Xu, and Y. Xu, “Biodegradation of dibenzofuran
by Janibacter terrae strain XJ-1,” Current Microbiology, vol. 53,
no. 1, pp. 30–36, 2006.
A. Yamazoe, O. Yagi, and H. Oyaizu, “Degradation of polycyclic
aromatic hydrocarbons by a newly isolated dibenzofuranutilizing Janibacter sp. strain YY-1,” Applied Microbiology and
Biotechnology, vol. 65, no. 2, pp. 211–218, 2004.
A. Yamazoe, O. Yagi, and H. Oyaizu, “Biotransformation of fluorene, diphenyl ether, dibenzo-p-dioxin and carbazole by Janibacter sp,” Biotechnology Letters, vol. 26, no. 6, pp. 479–486,
2004.
G.-Y. Zhang, J.-Y. Ling, H.-B. Sun, J. Luo, Y.-Y. Fan, and Z.-J.
Cui, “Isolation and characterization of a newly isolated polycyclic aromatic hydrocarbons-degrading Janibacter anophelis
strain JY11,” Journal of Hazardous Materials, vol. 172, no. 2-3, pp.
580–586, 2009.
D. A. Edwards, R. G. Luthy, and Z. Liu, “Solubilization of polycyclic aromatic hydrocarbons in micellar nonionic surfactant
solutions,” Envirenmental Science and Technology, vol. 25, no. 1,
pp. 127–133, 1991.
C. Rösch, A. Mergel, and H. Bothe, “Biodiversity of denitrifying
and dinitrogen-fixing bacteria in an acid forest soil,” Applied and
Environmental Microbiology, vol. 68, no. 8, pp. 3818–3829, 2002.
K. Wilson, “Preparation of genomic DNA from bacteria,” in
Current Protocols in Molecular Biology, F. M. Ausubel, R. Brent,
R. E. Kingston et al., Eds., pp. 241–245, 1987.
D. Daffonchio, S. Borin, G. Frova, P. L. Manachini, and C.
Sorlini, “PCR fingerprinting of whole genomes: the spacers
between the 16s and 23S rRNA genes and of intergenic tRNA
gene regions reveal a different intraspecific genomic variability
of Bacillus cereus and Bacillus licheniformis,” International Journal of Systematic Bacteriology, vol. 48, no. 1, pp. 107–116, 1998.
S. F. Altschul, W. Gish, W. Miller, E. W. Myers, and D. J. Lipman,
“Basic local alignment search tool,” Journal of Molecular Biology,
vol. 215, no. 3, pp. 403–410, 1990.
BioMed Research International
[40] K. Tamura, D. Peterson, N. Peterson, G. Stecher, M. Nei, and
S. Kumar, “MEGA5: molecular evolutionary genetics analysis
using maximum likelihood, evolutionary distance, and maximum parsimony methods,” Molecular Biology and Evolution,
vol. 28, no. 10, pp. 2731–2739, 2011.
[41] N. Saitou and M. Nei, “The neighbor-joining method: a new
method for reconstructing phylogenetic trees.,” Molecular biology and evolution, vol. 4, no. 4, pp. 406–425, 1987.
[42] I. S. Thakur, “Structural and functional characterization of a stable, 4-chlorosalicylic-acid-degrading, bacterial community in a
chemostat,” World Journal of Microbiology and Biotechnology,
vol. 11, no. 6, pp. 643–645, 1995.
[43] A. Premalatha and G. S. Rajakumar, “Pentachlorophenol degradation by Pseudomonas aeruginosa,” World Journal of Microbiology and Biotechnology, vol. 10, no. 3, pp. 334–337, 1994.
[44] M. Tripathi and S. K. Garg, “Studies on selection of efficient bacterial strain simultaneously tolerant to hexavalent chromium
and pentachlorophenol isolated from treated tannery effluent,”
Research Journal of Microbiology, vol. 5, no. 8, pp. 707–716, 2010.
[45] S. Singh, R. Chandra, D. K. Patel, and V. Rai, “Isolation
and characterization of novel Serratia marcescens (AY927692)
for pentachlorophenol degradation from pulp and paper mill
waste,” World Journal of Microbiology and Biotechnology, vol. 23,
no. 12, pp. 1747–1754, 2007.
[46] M. Cai and L. Xun, “Organization and regulation of pentachlorophenol-degrading genes in Sphingobium chlorophenolicum
ATCC 39723,” Journal of Bacteriology, vol. 184, no. 17, pp. 4672–
4680, 2002.
[47] E. A. Wolski, S. E. Murialdo, and J. F. Gonzalez, “Effect of
pH and inoculum size on pentachlorophenol degradation by
Pseudomonas sp,” Water SA, vol. 32, no. 1, pp. 93–98, 2006.
[48] C. Barbeau, L. Deschênes, D. Karamanev, Y. Comeau, and R.
Samson, “Bioremediation of pentachlorophenol-contaminated
soil by bioaugmentation using activated soil,” Applied Microbiology and Biotechnology, vol. 48, no. 6, pp. 745–752, 1997.
[49] C.-F. Yang, C.-M. Lee, and C.-C. Wang, “Degradation of
chlorophenols using pentachlorophenol-degrading bacteria
Sphingomonas chlorophenolica in a batch reactor,” Current
Microbiology, vol. 51, no. 3, pp. 156–160, 2005.
[50] R. L. Crawford and W. W. Mohn, “Microbiological removal of
pentachlorophenol from soil using a Flavobacterium,” Enzyme
and Microbial Technology, vol. 7, no. 12, pp. 617–620, 1985.
[51] J. T. Trevors, “Effect of temperature on the degradation of
pentachlorophenol by Pseudomonas species,” Chemosphere, vol.
11, no. 4, pp. 471–475, 1982.
[52] M. A. Providenti, H. Lee, and J. T. Trevors, “Selected factors
limiting the microbial degradation of recalcitrant compounds,”
Journal of Industrial Microbiology, vol. 12, no. 6, pp. 379–395,
1993.
[53] M. D. Webb, G. Ewbank, J. Perkins, and A. J. McCarthy,
“Metabolism of pentachlorophenol by Saccharomonospora
viridis strains isolated from mushroom compost,” Soil Biology
and Biochemistry, vol. 33, no. 14, pp. 1903–1914, 2001.
[54] R. Chandra, A. Ghosh, R. K. Jain, and S. Singh, “Isolation and
characterization of two potential pentachlorophenol degrading
aerobic bacteria from pulp paper effluent sludge,” The Journal
of General and Applied Microbiology, vol. 52, no. 2, pp. 125–130,
2006.
[55] C. M. Kao, J. K. Liu, Y. L. Chen, C. T. Chai, and S. C. Chen,
“Factors affecting the biodegradation of PCP by Pseudomonas
mendocina NSYSU,” Journal of Hazardous Materials, vol. 124,
no. 1–3, pp. 68–73, 2005.
9
[56] K. V. Gayathri and N. Vasudevan, “Enrichment of phenol
degrading moderately halophilic bacterial consortium from
saline environment,” Journal of Bioremediation & Biodegradation, vol. 2011, article 104, 2010.
[57] D. H. Yeh, K. D. Pennell, and S. G. Pavlostathis, “Toxicity
and biodegradability screening of nonionic surfactants using
sediment-derived methanogenic consortia,” Water Science and
Technology, vol. 38, no. 7, pp. 55–62, 1998.
[58] F. Volkering, A. M. Breure, J. G. van Andel, and W. H. Rulkens, “Influence of nonionic surfactants on bioavailability and
biodegradation of polycyclic aromatic hydrocarbons,” Applied
and Environmental Microbiology, vol. 61, no. 5, pp. 1699–1705,
1995.
[59] W. H. Noordman, J. H. J. Wachter, G. J. de Boer, and D. B.
Janssen, “The enhancement by surfactants of hexadecane degradation by Pseudomonas aeruginosa varies with substrate availability,” Journal of Biotechnology, vol. 94, no. 2, pp. 195–212, 2002.
[60] T. L. Cort, M.-S. Song, and A. R. Bielefeldt, “Nonionic surfactant
effects on pentachlorophenol biodegradation,” Water Research,
vol. 36, no. 5, pp. 1253–1261, 2002.
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 909312, 7 pages
http://dx.doi.org/10.1155/2014/909312
Review Article
Biotechnological Applications Derived from Microorganisms of
the Atacama Desert
Armando Azua-Bustos1 and Carlos González-Silva2
1
2
Blue Marble Space Institute of Science, Seattle, WA 98109, USA
Centro de Investigación del Medio Ambiente (CENIMA), Universidad Arturo Prat, 1110939 Iquique, Chile
Correspondence should be addressed to Armando Azua-Bustos; [email protected]
Received 7 April 2014; Revised 29 June 2014; Accepted 7 July 2014; Published 23 July 2014
Academic Editor: Ameur Cherif
Copyright © 2014 A. Azua-Bustos and C. González-Silva. This is an open access article distributed under the Creative Commons
Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is
properly cited.
The Atacama Desert in Chile is well known for being the driest and oldest desert on Earth. For these same reasons, it is also
considered a good analog model of the planet Mars. Only a few decades ago, it was thought that this was a sterile place, but in
the past years fascinating adaptations have been reported in the members of the three domains of life: low water availability, high
UV radiation, high salinity, and other environmental stresses. However, the biotechnological applications derived from the basic
understanding and characterization of these species, with the notable exception of copper bioleaching, are still in its infancy, thus
offering an immense potential for future development.
1. Introduction
The Atacama Desert, located in northern Chile between
latitudes 17∘ and 27∘ south, has average annual rains of less
than 2 mm [1]. In comparison, other known deserts in the
world, like the Mojave Desert in North America [2] or the
Sahara Desert in Africa [3], have average annual rains of
116 mm and 100 mm, respectively. These extremely low rain
rates have determined the Atacama Desert to be classified as
a hyperarid desert [4] (a desert with an aridity index of less
than 0.05, as the evapotranspiration of water from its soils is
much higher than the inputs of rains). The Atacama Desert is
also unique as it is believed to be the oldest desert on Earth,
being arid for the last 150 million years and hyperarid for the
past 15 million years [5, 6].
Thus, the Atacama has been an extremely dry desert
for a very long time and only forty years ago it was
thought that nothing could live in its seemingly barren
landscapes (Figure 1(a)). However, during the past ten years,
culture dependent and independent methods have unveiled
a plethora of microorganisms (Bacteria, Archaea, and
Eukarya) that were able to adapt and evolve in very specific
and unexpected habitats of this desert [7]. Habitats as diverse
as the underside of quartz rocks [8], fumaroles at the Andes
Mountains [9], the inside of halite evaporites [10], and caves
of the Coastal Range [11, 12] showed that microbial life found
novel ways to adapt to the extreme conditions typical of
the Atacama: extremely low water availability, intense solar
radiation, and high salinity (for a more complete description
of Atacama’s microbial species, please see our recent review
on this subject [7]). However, up to date, very few works have
gone beyond the descriptive stage of establishing what types
of microorganisms may be found in specific microenvironments [13], thus explaining the incipient biotechnological
applications derived from knowledge that still being gained.
The study of the molecular strategies used by microbial
life in other extreme environments (high temperature, for
example) gave rise to many biotechnological applications that
are now of standard use [14]. In a similar way, the characterization of the molecular strategies evolved by microorganisms
of the Atacama to cope with its exceptional abiotic stresses
(desiccation in particular) should be multiple and unique,
and, thus, novel sources of metabolites and genes for the
biotechnological industry. In this review, the few reported
cases of the biotechnological use of Atacama Desert microorganisms to date are summarized.
2
BioMed Research International
(a)
(b)
(c)
Figure 1: Examples of habitats of the Atacama Desert from where biotechnological applications have been derived or used. (a) The central
valley, the hyperarid core of the Atacama Desert. (b) Heap bioleaching at Radomiro Tomic, an open pit copper mine owned by the Chilean
Copper Corporation (Codelco). Note the copper rich blue-green solution obtained from the heaps. (c) The Loa River, a typical arsenic rich
river of the Atacama Desert. Image credits: Panels A and C: Armando Azua-Bustos. Panel B: Armando Azua Aroz.
2. Applications Derived from Members of
the Bacteria Domain
Copper Bioleaching. Copper bioleaching or “biomining”
allowed the usage of insoluble copper sulphides and oxides
through hydrometallurgy, as opposed to the traditional
technology of pyrometallurgy. Compared to pyrometallurgy,
bioleaching has the advantage of being a simpler process,
requiring less energy and equipments (Figure 1(b)). In addition, bioleaching does not produce sulfur dioxide emissions,
an important factor for the Chilean mining towns which
were usually built alongside the extracting operations in
Chile (most of which are located in the Atacama Desert).
Bioleaching also offered a better treatment of low grade
(again the usual case in Atacama copper ores) or waste
ores and in many cases it is the only way to treat them.
Low-grade ores (0.6% and less) are abundant in Chile,
but their processing by pyrometallurgy in most cases is
not economical. Through bioleaching, copper was able to
be extracted from ore minerals like chalcopyrite (CuFeS2),
with the crucial contribution of chemolithotrophic microbial
species extremely tolerant to low pH, which use the reduced
sulphur as an energy source. The most known of these
microorganisms is Acidithiobacillus ferrooxidans [15], but
other species, like Leptospirillum ferrooxidans, Sulfobacillus
acidophilus, and Acidimicrobium ferrooxidans, are thought to
also participate in the bioleaching process [16, 17].
In Chile, the first mine that introduced bioleaching was
Sociedad Minera Pudahuel (a copper mine not located in
the Atacama) in the 1980s. Today, this process is extensively
used in the Chilean copper mining industry [18, 19], reaching
over 1.6 million tons of copper per year [19]. It has been
estimated that Chile’s copper actual reserves would increase
up to 50% if all copper sulphides could be economically
treated by bioleaching [19].
Acidithiobacillus ferrooxidans has been identified in different places of the Atacama Desert [20, 21]. Some of these
species have been found in sulfidic mine tailings dumps in
the marine shore at Chañaral Bay [21], located at the Coastal
Range of the Atacama. This is of particular interest as these
species were found to be halotolerant iron oxidizers, active
at NaCl concentrations up to 1 M in enrichment cultures.
High concentrations of chloride ions inhibit the growth of the
acidophilic microorganisms traditionally used in biomining
[22]. Thus, the finding of halotolerant bioleaching species
would allow the use of seawater for biomining operations in
the future, a very important advancement in a region, where
water availability has always been extremely low.
Bioleaching strains found in the Atacama Desert have
been recently patented, as is the case of Acidithiobacillus ferrooxidans strain Wenelen DSM 16786 [23] and Acidithiobacillus thiooxidans strain Licanantay DSM 17318 [24] (US Patent
numbers 7,601,530 and 7,700,343). Both strains showed
improved oxidizing activity when compared to standard
BioMed Research International
strains isolated elsewhere, like Acidithiobacillus ferrooxidans
ATCC 23270 and Acidithiobacillus thiooxidans ATCC 8085.
Strain Wenelen, an iron and sulfur oxidizing microorganism,
was particularly efficient in oxidizing chalcopyrite, while
strain Licanantay, a strict sulfur oxidizer, showed activity
in both primary and secondary sulfured minerals, such
as chalcopyrite, covellite, bornite, chalcocite, enargite, and
tennantite [24–26].
Recently, the comparative genomic analysis and metabolomic profiles of these two strains were obtained, which
turned helpful for determining basic aspects of its regulatory
pathways and functional networks, biofilm formation, energy
control, and detoxification responses [27, 28].
As for Archaea, although some species have been
reported in acid mine drainage in the Atacama [21], there are
yet no reports of strains specifically isolated for industrial use.
Biomedicine. Soils of the Atacama shelter a number
of bacterial species with promising characteristics for the
biomedical industry. One of the first descriptions of microorganisms that are known to produce such biomolecules was
published in 1966 by Cameron et al. [29]. Commissioned by
NASA, this group approached the Atacama as a way to obtain
basic information on terrestrial desert environments and its
microbiota in order to develop and test the instruments to be
taken to Mars ten years later by the Viking Mission. Among
others, they were among the first to report the presence
of Streptomyces species, Bacillus subtilis aterrimus, Bacillus
brevis, Bacillus cereus, and Micrococcus caseolyticus; however,
no details of biomolecules produced by these species were
later reported.
Almost forty years passed until a groundbreaking report
by McKay’s group in 2003 [30] showed that when the experiments performed by the Viking landers on the surface of Mars
were repeated with soils of the Yungay region of the Atacama
Desert, the same results were obtained essentially. This leads
to the recognition of the Atacama Desert as one of the driest
places on Earth, causing then a surge of reports focusing on
the characteristics of various microenvironmental conditions
in the Atacama and its related microbiology [7].
Later on, the interest in the potential biomedical use of
these recently reported species started, focusing on species
of the Actinobacteria, as these were previously known as
synthesizers of useful molecules [31]. Among this latter class,
species like Amycolatopsis, Lechevalieria, and Streptomyces
have been reported at various arid and hyperarid sites of the
Atacama [32].
Members of the Streptomycetes are a well-known source
of antibiotics [33], and Lechevalieria species are known to
have nonribosomal peptide synthase (NRPS) gene clusters
that synthesize antitumoral compounds [31]. Accordingly,
of the species found by Okoro’s group [32], all of the
Amycolatopsis and Lechevalieria and most of the Streptomyces
isolates tested positive for the presence of NRPS genes. This
same group determined later the metabolic profile of one
of these Streptomyces strains (strain C34), identifying three
new compounds from the macrolactone polyketides class
[34] and other compounds like deferoxamine E, hygromycin
A, and 5󸀠󸀠 -dihydrohygromycin. These compounds showed
a strong activity against the Gram-positive bacteria tested
3
(Staphylococcus aureus, Listeria monocytogenes, and Bacillus
subtilis), but weak activity against the tested Gram-negative
bacteria (E. coli and Vibrio parahaemolyticus).
In a parallel report, they also found that strain C34
synthetized four new antibiotics of the ansamycin-type
polyketides with antibacterial activity against both Staphylococcus aureus ATCC 25923 and Escherichia coli ATCC
25922 [35]. In particular, chaxamycin D4 showed a selective
antibacterial activity against S. aureus ATCC 25923. Another
of these strains, Streptomyces sp. C38, synthetized three new
macrolactone antibiotics (atacamycins A–C) which exhibited
moderate inhibitory activity against the enzyme phosphodiesterase (PDE-4B2) [36]. Inhibitors of PDE (the most
famous of this group being Viagra) can prolong or enhance
the effects of physiological responses mediated by cAMP
and cGMP by inhibition of their degradation by PDE and
are considered potential therapeutics for pulmonary arterial
hypertension, coronary heart disease, dementia, depression,
and schizophrenia [37]. In the case of atacamycin A, it also
showed anti proliferative activity against cell lines of colon
cancer (CXF DiFi), breast cancer (MAXF 401NL), uterus
cancer (UXF 1138L), and colon RKO cells [36].
Similar positive results were obtained by Leirós et al., 2014
[38] in which seven molecules synthetized by Streptomyces
sp. Lt 005, Atacama Streptomyces C1, and Streptomyces sp.
CBS 198.65 were tested against hydrogen peroxide stress in
primary cortical neurons as potentially new drugs for the
avoidance of neurodegenerative disorders such as Parkinson’s and Alzheimer’s diseases. The reported compounds
inhibited neuronal cytotoxicity and reduced reactive oxygen
species (ROS) release after 12 h of treatment. Among these
compounds, the quinone anhydroexfoliamycin and the red
pyrrole-type pigment undecylprodigiosin showed the best
protection against oxidative stress with mitochondrial function improvement, ROS production inhibition, and increase
of antioxidant enzymes like glutathione and catalase. In addition, both compounds showed a modest caspase-3 activity
induced by the apoptotic enhancer staurosporine.
In a different work, another group of secondary metabolites, called abenquines, were found to be synthetized by
Streptomyces sp. Strain DB634, isolated from the soils of
the Altiplano of the Atacama [39]. These abenquines (A–D)
showed modest inhibitory activity against Bacillus subtilis,
dermatophytic fungi, phosphodiesterase type 4b, and antifibroblast proliferation (NIH-3T3).
An interesting case to discuss in this section of a commercially successful, but controversial, example of a compound
produced from an Actinobacteria isolated from another
well-known Chilean environment is that of rapamycin (also
known as sirolimus), isolated by Brazilian researchers from
a strain of Streptomyces hygroscopicus endemic of Eastern
Island, or Rapa Nui [40]. Rapamycin was originally used
as an antibiotic, but later on it was discovered to show
potent immunosuppressive and antiproliferative properties
[41, 42] and even claimed to extend life span [43]. Sadly,
nothing of this development benefited the Chilean economy,
as agreements like the United Nations Rio Declaration on
Environment and Development were yet to be established.
4
Arsenic Bioremediation. Conventional arsenic removal in
drinking water such as reverse osmosis and nanofiltration are
effective and able to remove up to 95% of the initial arsenic
concentrations, but the operating costs of these plants are
high [44]. In addition, the oxidation of As (III) to As (V) is a
prerequisite for all conventional treatment processes, and as
this is an extremely slow reaction toxic and costly oxidants
such as chlorine, hydrogen peroxide, or ozone must be used
as catalysts [44, 45]. Thus, an attractive alternative solution
for arsenic removal is bioremediation, as a wide variety of
bacteria can use it as an electron donor for autotrophic
growth or as an electron acceptor for anaerobic respiration
[46–48].
In the case of the Atacama Desert, the first steps leading to
the biosequestration of arsenic by endemic microorganisms
are now being taken. This toxic metalloid is naturally found in
rivers of the Atacama Desert (Figure 1(b)) as arsenate As (V)
and the most toxic species arsenite As (III) [49–51]. Among
other negative biological effects, arsenate, being a chemical
analog of phosphate, inhibits oxidative phosphorylation and
arsenite binds to sulfhydryl groups of proteins [52]. It is
precisely in the sediments of one of these rivers, (Camarones
river near the coastal city of Arica) with arsenic concentration, in water (1100 𝜇g L−1 ) and sediments (550 𝜇g L−1 ) that
49 isolates were identified and distributed between the 𝛼Proteobacteria (5 isolates), 𝛽-Proteobacteria (13 isolates), and
𝛾-Proteobacteria (26 isolates) [44, 53]. Most of these species
belonged to the genera Alcaligenes, Burkholderia, Comamonas, Enterobacter, Erwinia, Moraxella, Pantoea, Serratia,
Sphingomonas, and Pseudomonas [53], of which Alcaligenes,
Burkholderia, Sphingomonas, Pantoea, Erwinia, and Serratia
were not previously reported in literature as arsenic tolerant.
Fittingly, eleven of the arsenic-tolerant isolates had the gene
ars that codes for the critical enzyme involved in this reaction,
arsenate reductase [53]. In a later work from this group,
it was found that one of the species isolated, Pseudomonas
arsenicoxydans strain VC-1, was able to tolerate up to 5 mM
of As (III), being also capable of oxidizing at high rates the
totality of the arsenite present in the medium, with lactate as a
carbon source [54]. Thus, the characterization of these species
in experimental bioreactors will certainly offer interesting
options for future water and soil bioremediation [55].
3. Applications Derived from Microbial
Members of the Eukarya Domain
Biomedicine. Carotenoids are lipid soluble tetraterpenoid pigments synthesized as hydrocarbons (carotene, e.g., lycopene,
𝛼-carotene, and 𝛽-carotene) or their oxygenated derivatives
(xanthophylls, e.g., lutein, 𝛼-cryptoxanthin, zeaxanthin, etc.)
by microorganisms and plants [56]. In these organisms,
they play multiple and critical roles in photosynthesis,
by maintaining the structure and function of photosynthetic complexes, contributing to light harvesting, quenching
chlorophyll triplet states, scavenging reactive oxygen species,
and dissipating excess energy [57, 58]. Up to date, more than
700 carotenoids have been described [59]. Yellow, orange,
and red carotenoids are used as pharmaceuticals, animal feed
BioMed Research International
additives, and colorants in cosmetics and foods. Interest in
dietary carotenoids has increased in the past years due to their
antioxidant and anti-inflammatory potential [60, 61], as they
are very efficient quenchers of singlet oxygen and scavengers
of other reactive oxygen species [62]. Carotenoids are also
important precursors of retinol (vitamin A) [62, 63].
Among other sources, species of the halophilic biflagellate unicellular green alga Dunaliella (Chlorophyta), like
Dunaliella salina, are industrially cultivated as a natural
source of beta-carotene around the world, including Chile
[64]. Under conditions of abiotic stress (high salinity, high
temperature, high light intensity, and nitrogen limitation) up
to 12% of the algal dry weight is 𝛽-carotene [65, 66] which
accumulates in oil globules in the interthylakoid spaces of
their chloroplast [65]. In addition, as a defense mechanism
against hypersalinity, D. salina synthetizes high amounts of
the compatible solute glycerol, another molecule of economic
value [67].
There are several species of the genus Dunaliella reported
in the Atacama Desert, mainly in hypersaline lagoons [68, 69]
and even growing aerophytically on cave walls [11]. In the
case of D. Salina Isolate Conc-007, isolated from the Salar de
Atacama, it was found to be capable of synthesizing 100 pg
of beta-carotene per cell, two to four times higher than
other species used in commercial beta-carotene production
[69, 70]. In turn, D. salina SA32007, also isolated from the
Salar de Atacama, synthesized triglycerides-enriched lipids
under nitrogen deficiency conditions, a potentially relevant
result for biodiesel production [71]. Important differences
in the carotenogenic capacity of the D. salina strains have
been shown to be dependent on the high genetic diversity
of member of this species [69]. This is highly relevant for
the case of the Chilean species, as it seems that they may
be better producers of 𝛽-carotene and other biomolecules in
comparison to other species of the world.
An additional factor to consider in this case is that
Dunaliella production facilities elsewhere (Australia, China,
and India) are located in areas where solar irradiance is
maximal, climate is warm, and hypersaline water is available
[57], which are precisely the characteristics of most areas of
the Atacama; thus, growth facilities may well be developed
in this desert using endemic strains. In addition, adaptive
laboratory evolution [72] and metabolic engineering may be
applied to the Atacama species in the future, as these methods
have recently been investigated and accomplished [72, 73].
4. Final Comments
The brevity of this review reflects how little has been advanced
to date in the biotechnological use of members of the
microbial world found in the Atacama Desert. This may be
understood. Although the Atacama is well known for its
extreme dryness, up to 2003, there was little interest in exploring and characterizing its potential microbial ecosystems, as it
was generally supposed to be sterile. Ten years later, microbial
life has been found in most if not all of its habitats, from
high thermal springs on the Andes Mountains to caves of the
Coastal Range, thus building a yet ongoing descriptive stage
BioMed Research International
of extant microbial ecosystems. Therefore, it is not surprising
that the technological stage of research is just beginning.
As previously mentioned, the Atacama Desert is unique
as it has been the most arid place on Earth for a very long
time, imposing the same selection pressure over the life forms
that arrived and then coevolved with it. Fittingly, all reports
to date have shown that these species are unique in the way
the capture, store and use water, tolerate solar radiation, high
saline conditions and low soil nutrients [7, 74]. Thus, with the
exception of copper bioleaching, now a multimillion dollar
industry, we believe there is an immense biotechnological
potential waiting to be discovered and developed in relation
to the tolerance to the aforementioned abiotic stresses.
Most groups now are still reporting various degrees of
tolerance to abiotic stresses in the frame of basic research
(extreme environments and astrobiology in particular), and
we foresee that during the next years the detailed understanding of the physiological and molecular mechanisms involved
in the many abiotic stress tolerances shown by these species
should increase to a great extent (see [13], e.g.). “Omics”
techniques, like genomics, proteomics, and metabolomics,
and high-throughput technologies will be key in elucidating
processes and mechanisms involved in these tolerances and
then in identifying and characterizing key molecules of
potential use.
In the case of bioleaching, we envision that new bacterial and archaeal strains will appear in the market. Tailormade combinations of mine-specific strains will probably be
isolated, characterized, and patented in order to maximize
the dissolution of copper from the particular complexity
of minerals characteristic of each place. In addition, other
properties of the mine may be taken into account in the
determination of the characteristics of the strain mixture, like
water quality, soil temperature, and so forth.
In the case of biomolecules of interest for the biomedical
industry, there are already a handful of groups characterizing potentially interesting biomolecules from bacterial
strains isolated from the hyperarid areas and hypersaline
lagoons of the Atacama Desert, biomolecules which have
just been identified, and their activities preliminary tested.
These isolates are few and are representatives of a very small
fraction of the habitats of the Atacama, so novel strains
and metabolites will certainly appear in the near future.
Microorganisms of the dry core of the Atacama will be of
particular interest, as we expect that these species, being
subjected to the most extreme conditions, should produce
a number of biomolecules involved in the competition for
scarce resources.
With the increasing pressure of finding new drugs able
to handle antibiotic-resistant pathogens [75], extreme environments are now being investigated in detail [76], and the
Atacama Desert, given its unique peculiarities, may be a
prime place to explore.
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
5
References
[1] C. P. McKay, E. I. Friedmann, B. Gómez-Silva, L. CáceresVillanueva, D. T. Andersen, and R. Landheim, “Temperature
and moisture conditions for life in the extreme arid region of
the atacama desert: four years of observations including the El
Niño of 1997-1998,” Astrobiology, vol. 3, no. 2, pp. 393–406, 2003.
[2] J. F. Reynolds, P. R. Kemp, K. Ogle, and R. J. Fernández,
“Modifying the ’pulse-reserve’ paradigm for deserts of North
America: precipitation pulses, soil water, and plant responses,”
Oecologia, vol. 141, no. 2, pp. 194–210, 2004.
[3] C. J. Tucker, H. E. Dregne, and W. W. Newcomb, “Expansion and
contraction of the Sahara desert from 1980 to 1990,” Science, vol.
253, no. 5017, pp. 299–300, 1991.
[4] United Nations Environment Programme (UNEP), World Atlas
of Desertification, 1992.
[5] A. J. Hartley, G. Chong, J. Houston, and A. E. Mather, “150
million years of climatic stability: evidence from the Atacama
Desert, northern Chile,” Journal of the Geological Society, vol.
162, no. 3, pp. 421–424, 2005.
[6] J. Houston and A. J. Hartley, “The central andean west-slope
rainshadow and its potential contribution to the origin of
hyper-aridity in the Atacama Desert,” International Journal of
Climatology, vol. 23, no. 12, pp. 1453–1464, 2003.
[7] A. Azua-Bustos, C. Urrejola, and R. Vicuña, “Life at the dry
edge: microorganisms of the Atacama Desert,” The FEBS Letters,
vol. 586, no. 18, pp. 2939–2945, 2012.
[8] A. Azúa-Bustos, C. González-Silva, R. A. Mancilla et al.,
“Hypolithic cyanobacteria supported mainly by fog in the
coastal range of the Atacama Desert,” Microbial Ecology, vol. 61,
no. 3, pp. 568–581, 2011.
[9] E. K. Costello, S. R. P. Halloy, S. C. Reed, P. Sowell, and S. K.
Schmidt, “Fumarole-supported islands of biodiversity within a
hyperarid, high-elevation landscape on socompa volcano, puna
de atacama, andes,” Applied and Environmental Microbiology,
vol. 75, no. 3, pp. 735–747, 2009.
[10] J. Wierzchos, C. Ascaso, and C. P. McKay, “Endolithic cyanobacteria in halite rocks from the hyperarid core of the Atacama
Desert,” Astrobiology, vol. 6, no. 3, pp. 415–422, 2006.
[11] A. Azúa-Bustos, C. González-Silva, L. Salas, R. E. Palma, and R.
Vicuña, “A novel subaerial Dunaliella species growing on cave
spiderwebs in the Atacama Desert,” Extremophiles, vol. 14, no.
5, pp. 443–452, 2010.
[12] A. Azúa-Bustos, C. González-Silva, R. A. Mancilla et al.,
“Ancient photosynthetic eukaryote biofilms in an Atacama
Desert coastal cave,” Microbial Ecology, vol. 58, no. 3, pp. 485–
496, 2009.
[13] A. Azua-Bustos, J. Zúñiga, C. Arenas-Fajardo, M. Orellana,
L. Salas, and R. Vicuña, “Gloeocapsopsis AAB1, an extremely
desiccation-tolerant cyanobacterium isolated from the Atacama Desert,” Extremophiles, vol. 18, no. 1, pp. 61–74, 2014.
[14] P. L. Bergquist, H. W. Morgan, and D. Saul, “Selected enzymes
from extreme thermophiles with applications in biotechnology,”
Current Biotechnology, vol. 3, no. 1, pp. 45–59, 2014.
[15] D. P. Kelly and A. P. Wood, “Reclassification of some species
of Thiobacillus to the newly designated genera Acidithiobacillus
gen. nov., Halothiobacillus gen. nov. and Thermithiobacillus
gen. nov.,” International Journal of Systematic and Evolutionary
Microbiology, vol. 50, no. 2, pp. 511–516, 2000.
[16] P. Norris, “Acidophile diversity in mineral sulfide oxidation,”
in Biomining, D. Rawlings and B. Johnson, Eds., pp. 199–216,
Springer, Berlin, Germany, 2007.
6
[17] A. Schippers, “Microorganisms involved in bioleaching and
nucleic acid-based molecular methods for their identification
and quantification,” in Microbial Processing of Metal Sulfides, E.
Donati and W. Sand, Eds., pp. 3–33, Springer, Dordrecht, The
Netherlands, 2007.
[18] E. M. Domic-Mihovilovic, “A review of the development and
current status of copper bioleaching operations in Chile: 25
years of successful commercial implementation,” in Biomining,
D. E. Rawlings and B. D. Johnson, Eds., pp. 81–95, Springer,
Berlin, Germany, 2007.
[19] J. C. Gentina and F. Acevedo, “Application of bioleaching to
copper mining in Chile,” Electronic Journal of Biotechnology, vol.
16, no. 3, pp. 1–16, 2013.
[20] K. P. Drees, J. W. Neilson, J. L. Betancourt et al., “Bacterial
community structure in the hyperarid core of the Atacama
Desert, Chile,” Applied and Environmental Microbiology, vol. 72,
no. 12, pp. 7902–7908, 2006.
[21] H. Korehi, M. Blöthe, M. A. Sitnikova, B. Dold, and A.
Schippers, “Metal mobilization by iron- and sulfur-oxidizing
bacteria in a multiple extreme mine tailings in the Atacama
Desert, Chile,” Environmental Science & Technology, vol. 47, no.
5, pp. 2189–2196, 2013.
[22] C. M. Zammit, S. Mangold, V. Rao Jonna et al., “Bioleaching in
brackish waters-effect of chloride ions on the acidophile population and proteomes of model species,” Applied Microbiology
and Biotechnology, vol. 93, no. 1, pp. 319–329, 2012.
[23] T. Sugio, A. Miura, P. A. Parada Valdecantos, and R. Badilla
Ohlbaum, “Bacteria strain Wenelen DSM 16786, use of said
bacteria for leaching of ores or concentrates containing metallic
sulfide mineral species and leaching processes based on the use
of said bacteria or other publications mixtures that contain said
bacteria,” United States Patent US 7,601,530 B2, 2009.
[24] A. Ohata, M. Manaba, and P. A. Parada Valdecantos, “Sulfuroxidizing bacteria and their use in bioleaching processes for
sulfured copper minerals,” United States Patent no. US 7,700,343
B2, 2009.
[25] A. Ohata, M. Manabe, and P. A. Parada, “Sulfur-oxidizing
bacteria and their use in bioleaching processes for sulfured
copper minerals,” United States Patent No. US 7,700,343, 2010.
[26] T. Sugio, A. Miura, P. A. Parada, and R. Badilla, “Bacteria
strain Wenelen DSM 16786, use of said bacteria for leaching of
ores or concentrates containing metallic sulfide mineral species
and leaching processes based on the use of said bacteria or
mixtures that contain said bacteria,” United States Patent No.
US 7,601,530, 2009.
[27] G. Levicán, J. A. Ugalde, N. Ehrenfeld, A. Maass, and P. Parada,
“Comparative genomic analysis of carbon and nitrogen assimilation mechanisms in three indigenous bioleaching bacteria:
predictions and validations,” BMC Genomics, vol. 9, article 581,
2008.
[28] P. Martı́nez, S. Gálvez, N. Ohtsuka et al., “Metabolomic study of
Chilean biomining bacteria Acidithiobacillus ferrooxidans strain
Wenelen and Acidithiobacillus thiooxidans strain Licanantay,”
Metabolomics, vol. 9, no. 1, pp. 247–257, 2013.
[29] R. E. Cameron, D. R. Gensel, and G. B. Blank, “Soil studies—
desert microflora XII, abundance of microflora in soil samples
from the Chile Atacama Desert,” in Space Programs Summary,
vol. 4, pp. 37–38, Jet Propulsion Laboratory, 1966.
[30] R. Navarro-González, F. A. Rainey, P. Molina et al., “Mars-like
soils in the Atacama Desert, Chile, and the dry limit of microbial
life,” Science, vol. 302, no. 5647, pp. 1018–1021, 2003.
BioMed Research International
[31] H. Onaka, “Biosynthesis of heterocyclic antibiotics in actinomycetes and an approach to synthesize the natural compounds,”
Actinomycetologica, vol. 20, no. 2, pp. 62–71, 2006.
[32] C. K. Okoro, R. Brown, A. L. Jones et al., “Diversity of culturable
actinomycetes in hyper-arid soils of the Atacama Desert, Chile,”
Antonie van Leeuwenhoek, International Journal of General and
Molecular Microbiology, vol. 95, no. 2, pp. 121–133, 2009.
[33] R. E. Procópio, I. R. Silva, M. K. Martins, J. L. Azevedo, and
J. M. Araújo, “Antibiotics produced by Streptomyces,” Brazilian
Journal of Infectious Diseases, vol. 6, no. 5, pp. 466–471, 2012.
[34] M. E. Rateb, W. E. Houssen, W. T. A. Harrison et al., “Diverse
metabolic profiles of a Streptomyces strain isolated from a hyperarid environment,” Journal of Natural Products, vol. 74, no. 9, pp.
1965–1971, 2011.
[35] M. E. Rateb, W. E. Houssen, M. Arnold et al., “Chaxamycins A—
D, bioactive ansamycins from a hyper-arid desert Streptomyces
sp.,” Journal of Natural Products, vol. 74, no. 6, pp. 1491–1499,
2011.
[36] J. Nachtigall, A. Kulik, S. Helaly et al., “Atacamycins A-C, 22membered antitumor macrolactones produced by Streptomyces
sp. C38,” Journal of Antibiotics, vol. 64, no. 12, pp. 775–780, 2011.
[37] Y. H. Jeon, Y. Heo, C. M. Kim et al., “Phosphodiesterase:
overview of protein structures, potential therapeutic applications and recent progress in drug development,” Cellular and
Molecular Life Sciences, vol. 62, no. 11, pp. 1198–1220, 2005.
[38] M. Leirós, E. Alonso, J. A. Sanchez et al., “Mitigation of
ROS insults by Streptomyces secondary metabolites in primary
cortical neurons,” ACS Chemical Neuroscience, vol. 15, no. 1, pp.
71–80, 2014.
[39] D. Schulz, P. Beese, B. Ohlendorf et al., “Abenquines A-D:
Aminoquinone derivatives produced by Streptomyces sp. strain
DB634,” Journal of Antibiotics, vol. 64, no. 12, pp. 763–768, 2011.
[40] C. Vézina, A. Kudelski, and S. N. Sehgal, “Rapamycin (AY
22,989), a new antifungal antibiotic. I. Taxonomy of the producing streptomycete and isolation of the active principle,” Journal
of Antibiotics, vol. 28, no. 10, pp. 721–726, 1975.
[41] B. K. Law, “Rapamycin: an anti-cancer immunosuppressant?”
Critical Reviews in Oncology/Hematology, vol. 56, no. 1, pp. 47–
60, 2005.
[42] F. Neff, D. Flores-Dominguez, D. P. Ryan et al., “Rapamycin
extends murine lifespan but has limited effects on aging,”
Journal of Clinical Investigation, vol. 123, no. 8, pp. 3272–3291,
2013.
[43] M. V. Blagosklonny, “Rapamycin extends life- and health span
because it slows aging,” Aging, vol. 5, no. 8, pp. 592–598, 2013.
[44] V. L. Campos, G. Escalante, J. Yañez, C. A. Zaror, and M.
A. Mondaca, “Isolation of arsenite-oxidizing bacteria from a
natural biofilm associated to volcanic rocks of Atacama Desert,
Chile,” Journal of Basic Microbiology, vol. 49, no. 1, pp. S93–S97,
2009.
[45] M. Kim, J. Nriagu, and S. Haack, “Carbonate ions and arsenic
dissolution by groundwater,” Environmental Science and Technology, vol. 34, no. 15, pp. 3094–3100, 2000.
[46] K. Hrynkiewicz and C. Baum, “Application of microorganisms
in bioremediation of environment from heavy metals,” in
Environmental Deterioration and Human Health, A. Malik,
E. Grohmann, and R. Akhtar, Eds., pp. 215–227, Springer,
Amsterdam, The Netherlands, 2014.
[47] S. Yamamura and S. Amachi, “Microbiology of inorganic
arsenic: from metabolism to bioremediation,” Journal of Bioscience and Bioengineering, vol. 118, no. 1, pp. 1–9, 2014.
BioMed Research International
[48] J. H. Huang, “Impact of microorganisms on arsenic biogeochemistry: a review,” Water, Air, & Soil Pollution, vol. 225, no.
2, article 1848, 2014.
[49] P. L. Smedley and D. G. Kinniburgh, “A review of the source,
behaviour and distribution of arsenic in natural waters,” Applied
Geochemistry, vol. 17, no. 5, pp. 517–568, 2002.
[50] D. K. Nordstrom, “Worldwide occurrences of arsenic in ground
water,” Science, vol. 296, no. 5576, pp. 2143–2145, 2002.
[51] D. D. Caceres, P. Pino, N. Montesinos, E. Atalah, H. Amigo,
and D. Loomis, “Exposure to inorganic arsenic in drinking
water and total urinary arsenic concentration in a Chilean
population,” Environmental Research, vol. 98, no. 2, pp. 151–159,
2005.
[52] S. Shen, X. F. Li, W. R. Cullen, M. Weinfeld, and X. C. Le,
“Arsenic binding to proteins,” Chemical Reviews, vol. 113, no. 10,
pp. 7769–7790, 2013.
[53] G. Escalante, V. L. Campos, C. Valenzuela, J. Yañez, C. Zaror,
and M. A. Mondaca, “Arsenic resistant bacteria isolated from
arsenic contaminated river in the Atacama Desert (Chile),”
Bulletin of Environmental Contamination and Toxicology, vol.
83, no. 5, pp. 657–661, 2009.
[54] V. L. Campos, C. Valenzuela, P. Yarza et al., “Pseudomonas
arsenicoxydans sp nov., an arsenite-oxidizing strain isolated
from the Atacama desert,” Systematic and Applied Microbiology,
vol. 33, no. 4, pp. 193–197, 2010.
[55] A. S. Butt and A. Rehman, “Isolation of arsenite-oxidizing
bacteria from industrial effluents and their potential use in
wastewater treatment,” World Journal of Microbiology and
Biotechnology, vol. 27, no. 10, pp. 2435–2441, 2011.
[56] A. Das, S. H. Yoon, S. H. Lee, J. Y. Kim, D. K. Oh, and S. W. Kim,
“An update on microbial carotenoid production: application of
recent metabolic engineering tools,” Applied Microbiology and
Biotechnology, vol. 77, no. 3, pp. 505–512, 2007.
[57] J. A. del Campo, M. Garcı́a-González, and M. G. Guerrero,
“Outdoor cultivation of microalgae for carotenoid production: current state and perspectives,” Applied Microbiology and
Biotechnology, vol. 74, no. 6, pp. 1163–1174, 2007.
[58] B. Demmig-Adams and W. W. Adams III, “Antioxidants in
photosynthesis and human nutrition,” Science, vol. 298, no.
5601, pp. 2149–2153, 2002.
[59] G. Britton, S. Liaaen-Jensen, and H. Pfander, Carotenoids
Handbook, Birkhäauser, Basel, Switzerland, 2004.
[60] A. Das, S. Yoon, S. Lee, J. Kim, D. Oh, and S. Kim, “An
update on microbial carotenoid production: application of
recent metabolic engineering tools,” Applied Microbiology and
Biotechnology, vol. 77, no. 3, pp. 505–512, 2007.
[61] M. M. Ciccone, F. Cortese, M. Gesualdo et al., “Dietary intake of
carotenoids and their antioxidant and anti-inflammatory effects
in cardiovascular care, mediators of inflammation,” Mediators of
Inflammation, vol. 2013, Article ID 782137, 11 pages, 2013.
[62] J. Fiedor J and K. Burda, “Potential role of carotenoids as
antioxidants in human health and disease,” Nutrients, vol. 6, no.
2, pp. 466–488, 2014.
[63] L. H. Reyes, J. M. Gomez, and K. C. Kao, “Improving
carotenoids production in yeast via adaptive laboratory evolution,” Metabolic Engineering, vol. 21, no. 1, pp. 26–33, 2014.
[64] A. Oren, “A hundred years of Dunaliella research: 1905–2005,”
Saline Systems, vol. 1, no. 2, 2005.
[65] W. Fu, G. Paglia, M. Magnúsdóttir et al., “Effects of abiotic
stressors on lutein production in the green microalga Dunaliella
salina,” Microbial Cell Factories, vol. 13, no. 3, pp. 1–9, 2014.
7
[66] A. Ben-Amotz, “Dunaliella 𝛽-carotene: from science to commerce,” in Enigmatic Microorganisms and Life in Extreme
Environments, J. Seckbach, Ed., pp. 399–410, Kluwer Academic
Publishers, Dordrecht, The Netherlands, 1999.
[67] A. Ben-Amotz and M. Avron, “The role of glycerol in the
osmotic regulation of the halophilic alga Dunaliella parva,”
Plant Physiology, vol. 51, no. 5, pp. 875–878, 1973.
[68] P. Araneda, C. Jimenez, and B. Goméz-Silva, “Microalgae from
Northern Chile. III. Growth and beta carotene content of three
isolates of Dunaliella salina from the Atacama Desert,” Revista
de Biologı́a Marina y Oceanografı́a, vol. 27, no. 2, pp. 157–162,
1992.
[69] P. I. Gómez and M. A. González, “Genetic variation among
seven strains of Dunaliella salina (Chlorophyta) with industrial
potential, based on RAPD banding patterns and on nuclear ITS
rDNA sequences,” Aquaculture, vol. 233, no. 1–4, pp. 149–162,
2004.
[70] A. S. Cifuentes, M. Gonzalez, O. Parra, M. Zu, and M. Zuñiga,
“Culture of strains of Dunaliella salina (Teodoresco 1905) in
different media under laboratory conditions,” Revista Chilena
de Historia Natural, vol. 69, no. 1, pp. 105–112, 1996.
[71] D. Arias-Forero, G. Hayashida, M. Aranda et al., “Protocol
for maximizing the triglycerides-enriched lipids production
from Dunaliella salina SA32007 biomass, isolated from the
Salar de Atacama (Northern Chile),” Advances in Bioscience and
Biotechnology, vol. 4, no. 8, pp. 830–839, 2013.
[72] W. Fu, O. Guðmundsson, G. Paglia et al., “Enhancement of
carotenoid biosynthesis in the green microalga Dunaliella salina
with light-emitting diodes and adaptive laboratory evolution,”
Applied Microbiology and Biotechnology, vol. 97, no. 6, pp. 2395–
2403, 2013.
[73] D. Geng, Y. Han, Y. Wang et al., “Construction of a system for
the stable expression of foreign genes in Dunaliella salina,” Acta
Botanica Sinica, vol. 46, no. 3, pp. 342–346, 2004.
[74] A. Azua-Bustos, C. González-Silva, C. Arenas-Fajardo, and R.
Vicuña, “Extreme environments as potential drivers of convergent evolution by exaptation: the Atacama Desert Coastal Range
case,” Frontiers in Microbiology, vol. 3, article 426, 2012.
[75] B. Cannon, “Microbiology: resistance fighters,” Nature, vol. 509,
no. 7498, pp. S6–S8, 2014.
[76] A. Tasiemski, S. Jung, C. Boidin-Wichlacz et al., “Characterization and function of the first antibiotic isolated from a vent
organism: the extremophile metazoan Alvinella pompejana,”
PLoS ONE, vol. 9, no. 4, Article ID e95737, 2014.
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 439197, 11 pages
http://dx.doi.org/10.1155/2014/439197
Research Article
Diversity and Enzymatic Profiling of Halotolerant Micromycetes
from Sebkha El Melah, a Saharan Salt Flat in Southern Tunisia
Atef Jaouani,1 Mohamed Neifar,1 Valeria Prigione,2 Amani Ayari,1
Imed Sbissi,1 Sonia Ben Amor,1 Seifeddine Ben Tekaya,1 Giovanna Cristina Varese,2
Ameur Cherif,3 and Maher Gtari1
1
Laboratoire Microorganismes et Biomolécules Actives, Faculté des Sciences de Tunis, Université Tunis El Manar, Campus Universitaire,
2092 Tunis, Tunisia
2
Dipartimento di Scienze della Vita e Biologia dei Sistemi, Università degli Studi di Torino, Viale Mattioli 25, 10125 Torino, Italy
3
Laboratoire Biotechnologie et Valorisation des Bio-Géo Ressources, Institut Supérieur de Biotechnologie de Sidi Thabet,
Université La Manouba, 2020 Sidi Thabet, Tunisia
Correspondence should be addressed to Atef Jaouani; [email protected]
Received 2 May 2014; Accepted 28 June 2014; Published 16 July 2014
Academic Editor: Sara Borin
Copyright © 2014 Atef Jaouani et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Twenty-one moderately halotolerant fungi have been isolated from sample ashes collected from Sebkha El Melah, a Saharan salt
flat located in southern Tunisia. Based on morphology and sequence inference from the internal transcribed spacer regions, 28S
rRNA gene and other specific genes such as 𝛽-tubulin, actin, calmodulin, and glyceraldehyde-3-phosphate dehydrogenase, the
isolates were found to be distributed over 15 taxa belonging to 6 genera of Ascomycetes: Cladosporium (𝑛 = 3), Alternaria (𝑛 = 4),
Aspergillus (𝑛 = 3), Penicillium (𝑛 = 5), Ulocladium (𝑛 = 2), and Engyodontium (𝑛 = 2). Their tolerance to different concentrations
of salt in solid and liquid media was examined. Excepting Cladosporium cladosporioides JA18, all isolates were considered as alkalihalotolerant since they were able to grow in media containing 10% of salt with an initial pH 10. All isolates were resistant to oxidative
stresses and low temperature whereas 5 strains belonging to Alternaria, Ulocladium, and Aspergillus genera were able to grow at 45∘ C.
The screening of fungal strains for sets of enzyme production, namely, cellulase (CMCase), amylase, protease, lipase, and laccase, in
presence of 10% NaCl, showed a variety of extracellular hydrolytic and oxidative profiles. Protease was the most abundant enzyme
produced whereas laccase producers were members of the genus Cladosporium.
1. Introduction
Sebkhas are salt flats occurring on arid coastline in North
Africa, Arabia, Baja California, and Shark Bay Australia [1].
They are considered among the most poikilotopic environments and characterized by extreme salt concentrations and
electromagnetic radiation exposure together with low water
and nutrient availabilities [2]. Regarded as detrimental to
“normal subsistence,” organisms copying such conditions to
survive and thrive are designed extremophiles [3]. Beside
halophytes plants and algae, the mostly diverse dwellers of
sebkhas being unveiled are members of bacterial, archaeal,
and fungal ranks [4–8]. Members of fungi kingdom recovered
from extreme environments such as sebkhas’ have shed light
on two promising viewpoints: first, as model for deciphering
stress adaptation mechanisms in eukaryotes [9] and secondary, as novel and largely unexplored materials for the
screening of novel bioactive natural products [10]. Over
the past decade, there is an increased awareness for new
hydrolytic enzymes useful under nonconventional conditions
[11].
Sebkha El Melah, a Saharan salt flat of southern Tunisia,
has an area of approximately 150 km2 and the level is slightly
below the sea. Fluvial basin excavation of Sebkha El Melah
appeared at the beginning of the Würmian Quaternary
period [12]. Around 40,000 BP the lagoon was highly desalinated by freshwater arrivals. At the upper Würm, seawater
withdrew and the basin evolves to a temporary lake or
Mediterranean
Sea
Tunisia
Libya
continental sebkha. More recently, around 8000 years BP,
the lagoon evolved into an evaporite basin. The sebkha
sediments are composed of several saliferous layers of rock
salt and gypsum (calcium sulfate) and/or polyhalite (sulfate
of potassium, calcium, and magnesium) [12]. Here we report
the isolation of moderately halotolerant fungi from Sebkha El
Melah. Strains have been identified based on morphological
and molecular markers and their resistance to salt, thermal,
alkaline, and oxidative stresses was assessed. Their ability to
produce different hydrolytic and oxidative enzymes under
salt stress was also evaluated.
BioMed Research International
Algeria
2
L3
L1
L2
Figure 1: Map of Sebkha El Melah (Google Earth). L1, L2, and L3
indicate locations of sampling.
2. Material and Methods
2.1. Sampling Site Description and Fungal Isolation. Three
locations from the Sebkha El Melah margins (L1:
33∘ 23󸀠 01.1󸀠󸀠 N 10∘ 54󸀠 56.8󸀠󸀠 E; L2: 33∘ 21󸀠 42.1󸀠󸀠 N 10∘ 55󸀠 05.5󸀠󸀠 E;
and L3: 33∘ 23󸀠 37.7󸀠󸀠 N 10∘ 53󸀠 40.2󸀠󸀠 E) were chosen for sampling
(Figure 1). From each location, a composite sample was
prepared aseptically from five subsamples (1–10 cm deep)
and collected from the arms and center of an X (each arm
was 1 m in length) [13]. One cm soil from the ground surface
was firstly removed to avoid contamination during sampling
procedure. Samples were then transported to the laboratory
in a cool box and stored at 4∘ C prior to processing.
Fungi were isolated on potato dextrose agar (PDA)
containing 10% of NaCl and 0.05% of chloramphenicol using
the soil plate method where few milligrams of sample were
directly spread on the agar medium. This method has a slight
edge over the dilution plate method since it allows higher
total number of isolates and limits invasion by species which
sporulate heavily [14].
2.2. Morphological and Molecular Identification. Isolated
fungi were identified conventionally according to their
macroscopic and microscopic features. After determination
of their genera [15–17], they were transferred to the media
recommended of selected genus monographs for species
identification.
DNA extraction was achieved as described by Liu et
al. [18]; the amplification of the internal transcribed spacer
regions (nuclear-encoded 18S rRNA-ITS1-5.8S rRNA-ITS228S rRNA) was performed using the couple of universal
primers ITS1 (5󸀠 -TCC GTA GGT GAA CCT GCG G-3󸀠 ) and
ITS4 (5󸀠 -TCC TCC GCT TAT TGA TAT GC-3󸀠 ) [19] and
the thermal cycler conditions according to Luo and Mitchell
[20]. PCR was carried out in 25 𝜇L volumes containing 2.5 𝜇L
of 1X PCR reaction buffer (100 mM Tris-HCl, 500 mM KCl,
pH 8.3), 1.5 𝜇L MgCl2 , 0.2 𝜇mol/L (each) primer, 0.2 𝜇mol/L
(each) dNTP, and 2.5 units of Taq polymerase (Dream Taq,
Fermentas) and 1 𝜇L of DNA template. Depending on the
fungus genus, different gene sequences were amplified. For
the Aspergillus flavus group, the calmodulin gene was amplified using the primers CL1 (5󸀠 -GARTWCAAGGAGGCCTTCTC-3󸀠 ) and CL2A (5󸀠 -TTTTGCATCATGAGTTGGAC3󸀠 ) according to Rodrigues et al. [21]; for the Cladosporium
genus, the actin gene was amplified using the primers ACT512F (5󸀠 -ATGTGCAAGGCCGGTTTCGC-3󸀠 ) and ACT783R (5󸀠 -TACGAGTCCTTCTGGCCCAT-3󸀠 ) according to
Bensch et al. [22]; for Alternaria genus, the glyceraldehyde3-phosphate dehydrogenase gene was amplified using the
primers GPD1 (5󸀠 -CAACGGCTTCGGTCGCATTG-3󸀠 ) and
GPD2 (5󸀠 -GCCAAGCAGTTGGTTGTGC-3󸀠 ) according to
Berbee et al. [23]; for Penicillium and Aspergillus genera,
the 𝛽-tubulin gene was amplified using the primers Bt2a
(5󸀠 -GGTAACCAAATCGGTGCTGCTTTC-3󸀠 ) and Bt2B
(5󸀠 -ACCCTCAGTGTAGTGACCCTTGGC-3󸀠 ) according to
Glass and Donaldson [24].
The PCR products were purified with QIAquick Wizard
PCR purification Kit (Promega) according to the manufacturer’s instructions, and the sequences were determined
by cycle sequencing using the Taq Dye Deoxy Terminator Cycle Sequencing kit (Applied Biosystems, HTDS,
Tunisia) and fragment separation in an ABI PrismTM 3130
DNA sequencer (Applied Biosystems, HTDS, Tunisia). The
sequences obtained were compared reference sequences in
the NCBI GenBank database using the BLASTN search
option [25].
2.3. Effect of pH, Salinity, Temperature, and Oxidative Stress.
PDA medium was used to study the effect of different stresses
on solid media. For oxidative stresses, H2 O2 or paraquat was
filter sterilized and added separately to melted PDA medium
previously autoclaved. Paraquat is a redox-cycling agent
widely used to generate reactive oxygen species and induce
oxidative stress in bacteria [26] and fungi [27]. For pH stress,
PDA medium was buffered with 100 mM Glycine-NaOH to
pH 10 before autoclaving. Salt stress in solid media was
studied in PDA medium containing different concentrations
of salts. The inoculated plates with 3 mm cylindrical mycelial
plugs were then incubated at 30∘ C for oxidative, salt, and
pH stresses and at 4∘ C and 45∘ C for thermal stresses, and
radial growth was measured daily. Results were expressed
as relative growth of fungal strains under different stresses
as follows: (Colony diameter under stress/colony diameter
without stress after 7 days incubation) × 100.
The effect of salinity in liquid medium was carried out in
Biolog system, a commercially redox based test (Biolog Inc.,
Hayward, CA). Malt extracts (2%) containing 0%, 5%, 10%,
15%, and 20% of salt were inoculated by a suspension of spores
BioMed Research International
and fragmented mycelium according to the supplier’s instructions in 96-well microtiter plates. After 15 days incubation
at 30∘ C, the numeric results were extracted using PM Data
Analysis 1.3 software. The fungal growth was assimilated to
the reduction of the redox indicator. The ability of the fungus
to grow in the presence of salt was expressed as the ratio of
kinetic curve surface under stress versus without stress.
2.4. Extracellular Enzymes Production Profiling. The capacity
of fungal isolates to produce extracellular enzymes, namely,
amylase, cellulase, protease, laccase and lipase, was assayed
in the presence of 10% of NaCl. Inoculation was made
by transferring 3 mm of cylindrical mycelial plugs on the
corresponding media. Amylase production was assayed on
PDA containing 1% soluble starch. Enzyme production is
shown by the presence of clear halo when iodine was
poured onto the plates. Cellulase production was tested
on PDA medium containing 1% of carboxymethylcellulose.
The presence of activity is reflected by a clear halo on red
background after flooding the plates with 0.2% Congo red
for 30 min. Protease production was revealed on skim milk
agar by the appearance of a clear zone corresponding to
casein hydrolysis/solubilization surrounding the microbial
colony. The laccase production was detected on PDA medium
containing 5 mM of 2,6 dimethoxyphenol (DMP). Oxidation
of the substrate is indicated by the appearance of brown color.
Lipase production was tested on PDA medium containing
10 mL/L of Tween 20 and 0.1 g/L of CaCl2 . Positive reaction
is accompanied by the presence of precipitates around the
fungal colony. The enzymes production was expressed as
activity ratio (PR) which corresponds to the activity diameter
(halo of enzymatic reaction) divided by the colony diameter
after 7 days incubation at 30∘ C.
2.5. Statistical Analysis. The data presented are the average of
the results of at least three replicates with a standard error of
less than 10%.
3. Results
3.1. Isolation and Identification of Halotolerant Fungi.
Twenty-one fungal isolates were obtained on halophilic
medium containing 10% NaCl and subjected to morphological and molecular identification. Seventeen strains were
identified at genus level based on 28S rRNA gene sequences,
while four were identified based on ITS regions. Final
assignment was based on combination of morphological
and 𝛽-tubulin, actin, calmodulin, and glyceraldehyde-3phosphate dehydrogenase genes sequencing (Table 1). The 21
strains have been identified as Cladosporium cladosporioides
(𝑛 = 2), Cladosporium halotolerans (𝑛 = 1), Cladosporium
sphaerospermum (𝑛 = 2), Alternaria tenuissima (𝑛 = 1),
Aspergillus flavus (𝑛 = 1), Aspergillus fumigatiaffinis (𝑛 = 1),
Aspergillus fumigatus (𝑛 = 1), Penicillium canescens (𝑛 = 1),
Penicillium chrysogenum (𝑛 = 3), Penicillium sp. (𝑛 = 1),
Alternaria alternata (𝑛 = 3), Ulocladium consortiale (𝑛 = 1),
Ulocladium sp. (𝑛 = 1), Engyodontium album (𝑛 = 1), and
Embellisia phragmospora (𝑛 = 1) species. All the strains have
3
been deposited at the Mycotheca Universitatis Taurinensis
(MUT) in the University of Turin.
3.2. Salt Tolerance of Fungal Isolates. Salt tolerance of the
fungal isolates was assessed on solid and liquid media for
NaCl content ranging from 5 to 20%. In solid media, salt
tolerance was estimated as relative growth represented by
the ratio of colony diameter under salt stress to that without
salt stress. As illustrated in Table 2, all the isolated strains
succeeded to grow in the presence of 10% of salt. While 19
isolates remain able to grow under 15% NaCl, only 7 isolates
tolerated 20% NaCl: Penicillium chrysogenum JA1 and JA22,
Cladosporium halotolerans JA8, Cladosporium sphaerospermum JA2, Cladosporium cladosporioides JA18, Aspergillus
flavus JA4, and Engyodontium album JA7.
When liquid cultures were used, fungal isolates seemed
to become more sensitive to salt stress. Indeed, none of the
strains was able to grow in the presence of 20% NaCl, whereas
only 8 strains and 19 strains tolerated 15% and 10% NaCl,
respectively (Table 2).
3.3. Alkaline, Temperature, and Oxidative Stress. Excepting
Cladosporium cladosporioides JA18, all tested strains were
able to grow at pH 10. All isolates were able to grow at 4∘ C
while only five strains Aspergillus fumigatus JA10, Aspergillus
fumigatiaffinis JA11, Alternaria alternata JA23, Ulocladium
consortiale JA12, and Ulocladium sp. JA17 showed a significant
growth at 45∘ C. All 21 strains tolerated oxidative stress
generated by 10 mM H2 O2 and 500 𝜇M paraquat (Table 3).
3.4. Enzymatic Profiling of Isolates. Among the 21 strains
tested, 13 strains displayed at least one of the five-screened
activities: protease, amylase, cellulase, lipase, and laccase, in
the presence of 10% NaCl (Table 4). Protease and amylase
were the most abundant activities shown by 9 and 6 strains,
respectively. Four strains belonging to Cladosporium and
Penicillium genera produced laccase while Cladosporium
sphaerospermum JA2, Aspergillus flavus JA4, and Engyodontium album JA7 were able to produce lipase. Cellulase activity
was detected only in Penicillium sp. JA15.
4. Discussion
With regard to bacteria that have been well explored in
southern desert region of Tunisia [28–31], data related to
fungi are scarce and are limited to truffle and mycorrhiza, so
far considered as real specialists of desert environments [32,
33]. To the best of our knowledge, this is the first report on the
isolation and characterization of fungi from Tunisian desert
and particularly from salt flat. A collection of 21 fungi isolates
have been established from samples ashes collected from
Sebkha El Melah. These alkalihalotolerant fungi have been
assigned to 15 taxa belonging to 6 genera of Ascomycetes.
Several studies showed that fungi belonging to Cladosporium,
Alternaria, and Ulocladium genera were clearly predominant
under desert and salty environments [34, 35]. These fungi
have in common thick-walled and strongly melanized spores
which are important for UV, radiation, and desiccation
Penicillium flavigenum JX997105 (100%)
P. confertum JX997081 (100%)
P. dipodomyis JX997080 (100%)
P. commune KC333882 (100%)
P. chrysogenum KC009827 (100%)
P. griseofulvum JQ781833 (100%)
Cladosporium sp. GU017498 (100%)
Hyalodendron sp. AM176721 (100%)
C. sphaerospermum AB572902 (99%)
C. cladosporioides EF568045 (99%)
ITS identification
Aspergillus
nd
Alternaria
nd
Cladosporium
Embellisia/Chalastospora
JA4
JA5
JA6
JA7
JA8
JA9
Embellisia phragmospora JN383493
(100%)
Cladosporium cladosporioides EF568045
(100%)
C. sphaerospermum AM176719 (100%)
C. halotolerans JX535318 (99%)
Engyodontium album HM214540 (100%)
Alternaria tenuissima similarity
Alternaria arborescens
Alternaria alternata (GPD)
Cladosporium halotolerans
(Actin)
Alternaria tenuissima
(GPD)
Penicillium canescens group
(𝛽-tubulin and calmodulin)
Penicillium desertorum JX997039 (100%)
P. chrysogenum KC009826 (99%)
Alternaria triticimaculans JN867470
(100%)
A. tenuissima JN867469 (100%)
A. mali JN867468 (100%)
A. alternata JQ690087 (100%)
Aspergillus flavus
(calmodulin)
Penicillium chrysogenum
(𝛽-tubulin)
Cladosporium sphaerospermum
(Actin)
Penicillium chrysogenum
(𝛽-tubulin)
Identification based on specific
primers
Aspergillus aureofulgens EF669617 (100%)
Penicillium chrysogenum Penicillium canescens HQ607858(99%)
Cladosporium
JA2
JA3
Penicillium
28S identification
JA1
Strain code
Table 1: Identification of fungal isolates.
Penicillium chrysogenum
Thom
28S KF417559
ITS KF417577
Final identification and
accession number in
NCBI
Cladosporium
sphaerospermum Penzig
nd
28S: KF417560
ITS: KF417578
Penicillium chrysogenum
Thom
Penicillium chrysogenum
28S: KF417561
ITS: KF417579
Aspergillus flavus Link
28S: KF417562
nd
ITS: KF417580
Penicillium canescens
Sopp
nd
ITS: KF417581
Alternaria tenuissima
(Nees)
Wiltshire
Alternaria alternata
28S: KF417563
ITS: KF417582
Engyodontium album
(Limber) de Hoog
Engyodontium album
ITS: KF417583
Cladosporium halotolerans
Zalar, de Hoog, and
Gunde-Cimerman
Cladosporium cladosporioides/halotolerans
28S: KF417564
ITS: KF417584
Embellisia phragmospora
(Emden) E.G.
Embellisia phragmospora
28S: KF417565
ITS: KF417585
Penicillium chrysogenum
Morphological identification
4
BioMed Research International
Alternaria alternata
nd
Alternaria tenuissima
Alternaria arborescens
Alternaria alternata
(GPD)
Cladosporium cladosporioides HQ380770
(100%)
Cladosporium
Alternaria
JA18
JA19
Alternaria sp. KC139473 (100%)
A. arborescens JQ781762 (100%)
A. alternata JN107734 (100%)
Cladosporium cladosporioides
nd
Ulocladium consortiale JQ585682 (100%)
U. chartarum JN942881 (99%)
Ulocladium
JA17
Ulocladium sp.
Penicillium glabrum
nd
Penicillium spinulosum KC167852 (100%)
P. glabrum KC009784 (100%)
nd
Cladosporium sphaerospermum
Penicillium
Cladosporium sphaerospermum
group
(Actin)
Cladosporium cladosporioides JX868638
(99%)
C. sphaerospermum HM999943 (99%)
Ulocladium tuberculatum/consortiale
JA15
Cladosporium
JA13
Ulocladium consortiale (GPD)
Ulocladium consortiale JQ585682 (100%)
Alternaria radicina HM204457 (99%)
nd
nd
Ulocladium
JA12
Aspergillus fumigatiaffinis
(𝛽-tubulin)
Aspergillus aff. fumigatus JN246066
(100%)
A. fumigatiaffinis KC253955 (99%)
Aspergillus fumigatiaffinis
Morphological identification
Cladosporium cladosporioides KC009539
Cladosporium/Davidiella (99%)
Davidiella tassiana GU248332 (98%)
nd
JA11
Aspergillus fumigatus
(𝛽-tubulin)
Identification based on specific
primers
Aspergillus lentulus JN943567 (99%)
A. aff. fumigatus JN246066 (99%)
A. fumigatiaffinis HF545316 (99%)
A. novofumigatus FR733874 (99%)
ITS identification
JA14
Aspergillus
28S identification
JA10
Strain code
Table 1: Continued.
Final identification and
accession number in
NCBI
Aspergillus fumigatus
Fresenius
28S: KF417566
ITS: KF417586
Aspergillus fumigatiaffinis
Hong, Frisvad, and
Samson
ITS: KF417587
Ulocladium consortiale
(Thümen) E.G. Simmons
28S: KF417567
ITS: KF417588
Cladosporium
sphaerospermum Penzig
28S: KF417568
ITS: KF417589
Cladosporium
cladosporioides
(Fresenius) G.A. de Vries
28S: KF417569
ITS: KF417590
Penicillium sp.
28S: KF417570
ITS: KF417591
Ulocladium sp.
28S: KF417572
ITS: KF417593
Cladosporium
cladosporioides
(Fresenius) G.A. de Vries
28S: KF417573
ITS: KF417594
Alternaria alternata
Keissler
28S: KF417574
ITS: KF417595
BioMed Research International
5
Alternaria
Penicillium
nd
JA22
JA23
28S identification
JA20
Strain code
Penicillium chrysogenum KC341721 (99%)
P. dipodomyicola JX232278 (99%)
P. rubens JX003126 (99%)
P. commune JN676122 (99%)
Alternaria alternata JQ809324 (100%)
A. quercus KC329620 (100%)
A. tenuissima KC329619 (100%)
A. atrans KC329618 (100%)
Alternaria brassicae JX290150 (100%)
A. porri HQ821479 (100%)
ITS identification
Penicillium chrysogenum
Alternaria alternata
Alternaria tenuissima
Alternaria arborescens
Alternaria alternata
(GPD)
Alternaria alternata
Alternaria tenuissima
Alternaria arborescens
Alternaria alternata
(GPD)
Penicillium chrysogenum
(𝛽-tubulin)
Morphological identification
Identification based on specific
primers
Table 1: Continued.
Alternaria alternata
Keissler
ITS: KF417598
Final identification and
accession number in
NCBI
Alternaria alternata
Keissler
28S: KF417575
ITS: KF417596
Penicillium chrysogenum
Thom
28S: KF417576
ITS: KF417597
6
BioMed Research International
BioMed Research International
7
Table 2: Effect of salt concentration on fungal growth in solid and liquid media.
Strain code
JA1
JA3
JA22
AJ5
JA15
JA8
JA2
JA13
JA14
JA18
JA4
JA10
JA11
JA19
JA20
JA23
JA6
JA9
JA12
JA17
JA7
Solid media (1)
Liquid media (2)
5% NaCl 10% NaCl 15% NaCl 20% NaCl 5% NaCl 10% NaCl 15% NaCl 20% NaCl
Penicillium chrysogenum
74
72
60
18
83
54
10
0
Penicillium chrysogenum
100
72
37
0
96
46
11
0
Penicillium chrysogenum
100
82
41
25
90
46
0
0
Penicillium canescens
70
30
20
0
79
44
0
0
Penicillium sp.
83
70
34
0
53
18
0
0
Cladosporium halotolerans
80
68
32
18
30
0
0
0
Cladosporium sphaerospermum
76
64
34
22
47
19
11
0
Cladosporium sphaerospermum
100
49
25
0
81
79
12
0
Cladosporium cladosporioides
40
30
10
0
0
0
0
0
Cladosporium cladosporioides
58
40
24
8
63
61
4
0
Aspergillus flavus
90
80
48
26
56
16
0
0
Aspergillus fumigatus
100
76
35
0
100
57
11
0
Aspergillus fumigatiaffinis
100
46
25
0
52
30
12
0
Alternaria alternata
52
38
24
0
57
9
0
0
Alternaria alternata
60
40
0
0
37
24
0
0
Alternaria alternata
100
68
20
0
65
12
0
0
Alternaria tenuissima
100
60
22
0
80
55
17
0
Embellisia phragmospora
94
50
10
0
78
26
0
0
Ulocladium consortiale
72
28
0
0
32
10
0
0
Ulocladium sp.
100
70
28
0
67
20
0
0
Engyodontium album
56
36
14
10
43
10
0
0
Strain
(1) Relative growth on solid media after 7 days incubation = (⌀ colony under salt stress/⌀ colony without salt stress) × 100. (2) Relative growth in liquid media
after 7 days incubation = (kinetic curve surface under salt stress/kinetic curve surface without salt stress) × 100.
Table 3: Effect of alkaline, thermal, and oxidative stresses on fungal growth.
Strain code
JA1
JA3
JA22
JA5
JA15
JA8
JA2
JA13
JA14
JA18
JA4
JA10
JA11
JA19
JA20
JA23
JA6
JA9
JA12
JA17
JA7
Strain
Penicillium chrysogenum
Penicillium chrysogenum
Penicillium chrysogenum
Penicillium canescens
Penicillium sp.
Cladosporium halotolerans
Cladosporium sphaerospermum
Cladosporium sphaerospermum
Cladosporium cladosporioides
Cladosporium cladosporioides
Aspergillus flavus
Aspergillus fumigatus
Aspergillus fumigatiaffinis
Alternaria alternata
Alternaria alternata
Alternaria alternata
Alternaria tenuissima
Embellisia phragmospora
Ulocladium consortiale
Ulocladium sp.
Engyodontium album
Alkaline stress (1)
pH 10
43
42
47
26
43
34
21
21
34
—
46
89
94
49
58
100
57
58
44
93
34
Thermal stress (2)
4∘ C
45∘ C
39
—
50
—
45
—
28
—
100
—
26
—
24
—
43
—
38
—
41
—
22
—
41
61
26
100
35
—
48
—
83
40
30
—
67
—
37
36
28
100
18
—
Oxidative stress (3)
H2 O2 [10 mM]
Paraquat [500 𝜇M]
66
74
84
71
68
53
59
63
100
100
44
40
52
48
55
44
20
31
18
16
47
39
100
100
100
100
69
89
100
100
57
52
81
100
100
100
56
100
81
100
66
53
Relative growth of fungal strains under different stresses after 7 days incubation was expressed as follows: (1) (⌀ colony at pH 10/⌀ colony at pH 5) × 100; (2)
(⌀ colony at 45∘ C or 4∘ C/⌀ colony at 30∘ C) × 100; (3) (⌀ colony with H2 O2 or paraquat/⌀ colony without stress) × 100. —: not significant growth.
8
BioMed Research International
Table 4: Enzymes activities of fungal isolates in the presence of 10% NaCl.
Strain code
JA1
JA3
JA22
AJ5
JA15
JA8
JA2
JA13
JA14
JA18
JA4
JA10
JA11
JA19
JA20
JA23
JA6
JA9
JA12
JA17
JA7
Strain
Penicillium chrysogenum
Penicillium chrysogenum
Penicillium chrysogenum
Penicillium canescens
Penicillium sp.
Cladosporium halotolerans
Cladosporium sphaerospermum
Cladosporium sphaerospermum
Cladosporium cladosporioides
Cladosporium cladosporioides
Aspergillus flavus
Aspergillus fumigatus
Aspergillus fumigatiaffinis
Alternaria alternata
Alternaria alternata
Alternaria alternata
Alternaria tenuissima
Embellisia phragmospora
Ulocladium consortiale
Ulocladium sp.
Engyodontium album
Protease
++
++
+
−
−
+
+++
−
+
−
+
−
−
−
−
−
+
−
−
−
+
Amylase
+
−
+
−
−
−
−
+
+
−
−
−
−
−
+
−
−
−
−
−
+
Cellulase
−
−
−
−
+
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
Lipase
−
−
−
−
−
−
+
−
−
−
+
−
−
−
−
−
−
−
−
−
++
Laccase
−
−
−
+
−
+
+
+
−
−
−
−
−
−
−
−
−
−
−
−
−
AR: activity ratio = (⌀ activity/⌀ colony). −: no activity; +: AR < 1; ++: 1 < AR < 2; +++: 2 < AR < 3.
tolerance [10]. On the other hand, Molitoris et al. [36]
reported that other halotolerant and halophilic fungi such
as Aspergillus and Cladosporium spp. are predominant in
saline desert soil of Dead Sea. Many Aspergillus species have
been also reported to constitute dominant fungi in desert of
Saudi Arabia and Libya [37, 38], and halotolerant species,
including Aspergillus spp., Penicillium spp., and Cladosporium
sphaerospermum, have been consistently isolated from hypersaline environments around the globe [39]. In this study,
contrary to many reports on hypersaline environments, no
species belonging to the genera Eurotium, Thrimmatostroma,
Emericella, and Phaeotheca [9] have been obtained, probably
because of the initial alkaline pH of the Sebkha El Melah
salt lake. Actually, the effect of pH on the fungal diversity is
controversial. Misra [40] observed that fungal diversity varies
with the pH while other investigators found no significant
effect of pH values of water and soil habitats on fungal
occurrence [41]. It is more likely that the number of the
isolated fungi is directly correlated to the organic matter
content of water, mud, and soil samples [42].
Beside the identification of the recovered fungal isolates
from Sebkha El Melah, the second goal of the current study
was the detection of some of their physiological and biochemical features. This allows understanding ecological adaptation
to extreme environment and predicts some biotechnological
usage. The 21 strains have been screened for tolerance to
extreme NaCl concentrations, basic pH, temperature, and
oxidative stress and for the production of important enzymatic activities in presence of 10% NaCl.
Excepting Cladosporium cladosporioides JA18, all isolates
obtained in this study can be considered as moderately
haloalkaliphilic fungi as deduced from their ability to grow
at pH 10 and 10% of NaCl. However, the isolates were able
to grow when salt was not added to their growing media.
Excepting some Wallemia ichthyophaga the most strictly
halophilic fungus [43], all other fungal strains known to
date are able to grow without salt, a fact confirmed in
our study. However, gradual decrease in fungal growth was
observed with the increasing of salt concentration in the
culture medium. Nineteen strains remain able to grow under
15% of NaCl, whereas 7 strains were able to tolerate 20% of
NaCl. This result was confirmed by salt tolerance assay in
liquid media as estimated by Biolog system. It is noteworthy
that fungi were more sensitive to salt stress in liquid media
than in solid media. This could be explained by the alteration
of the osmotic gradient, forcing the fungi to expend more
energy in the osmoregulatory processes, resulting in slower
growth [44]. Moreover, at higher salt concentration death
occurs.
Regarding the stress of pH, the capacity of the majority of
isolates to growth at pH 10 implies firstly that some habitats in
the salt lake may have a varying pH and secondly that fungi
can tolerate a wide pH range. Prima facie, the overall results
in solid and liquid media showed that Penicillium chrysogenum JA1 and JA3, Cladosporium sphaerospermum JA2 and
JA13, Cladosporium cladosporioides JA18, Aspergillus fumigatus JA10, Aspergillus fumigatiaffinis JA11, and Alternaria
tenuissima JA6 are the most alkalihalotolerant isolates in this
study.
The tolerance of the strains to extreme 45∘ C was tested
and results indicated that Aspergillus fumigatus JA10, Alternaria alternata JA23, Ulocladium sp. JA17, and Aspergillus
fumigatiaffinis JA11 were able to grow. Of particular interest,
the latter two strains retained 100% of the growth rate
BioMed Research International
and biomass production as estimated by colony diameter.
Moreover, their ability to grow at low temperature may allow
them to better adapt to the big temperature fluctuation in
desert environments. Additionally, exposure to substrates
generating oxidative stress such as H2 O2 at 10 mM and
paraquat at 500 𝜇M did not alter significantly the growth of
almost tested strains demonstrating their ability to tolerate
oxidative stress. These findings may explain their presence in
desert regions that are considered amongst the most stressful
environments on Earth because of the high UV radiation,
desiccation, increased salinity, low nutrient availability, seasonal and daily temperature variation, and solar irradiation
[6, 10].
It has been postulated that microorganisms sharing a
rich and particular extracellular enzymatic activities are
common in harsh conditions characterizing their ecological
habitat including high level of aridity, temperature, ionic
strength, and particularly the low nutrient availability [31].
This implies the need by microorganisms for an effective
utilization of each possible available organic compound [45].
Moreover, fungal isolates from hot desert were revealed to
play an important role in seeds germination by breaking
dormancy and increasing water uptake [46]. In the present
study, the capacity of fungal isolates to produce extracellular
enzymes was assayed in the presence of 10% of NaCl.
Enzymes tested were the following: amylase for degradation
of starch, abundant carbohydrate polymer in many plant
tissues; protease for degradation of plant and animal proteins;
cellulase which hydrolyses the cellulose, the main component
of wood, ubiquitous substrate for fungi; and finally the laccase
involved in plant material delignification and in the synthesis
of the melanin and related compounds to protect fungi
against radiation. Thirteen strains displayed high productions
at least for one of the five-screened activities while no
clear correlation of enzyme production profile with fungal
systematic groups was noted. The abundance of protease
activity is in line with previous data on fungal isolates
from extreme environments showing high caseinase activities
with little effect of salinity and temperature on enzyme
production [36]. The relative limited number of isolates
displaying cellulase, amylase, lipase, and laccase activities
suggests that high concentration of salt may have an adverse
effect on enzyme production and/or activity. Their energy was
probably oriented to avoid salt stress due to 10% NaCl rather
than the production of bioactive extrolites [47]. However,
not detecting the enzyme is not absolute confirmation of
an isolate inability to produce it. It could also mean that
the enzyme was not released from the mycelium or that
the medium is inadequate for its detection [48]. Laccase
production in the presence of 10% of salt by the Cladosporium
group may be of biotechnological interest, for example, in
mycoremediation of high salty environments contaminated
by organic pollutants.
In conclusion, fungal community described in this study
was similar to those reported in inhospitable habitats char
acterized by limitation of nutrients, moisture deficit, and
9
exposure to high solar radiation. Further studies are needed
in order to elucidate their biogeochemical roles in such an
extreme environment and to exploit their promising potential
to produce new biomolecules such as enzymes and protective
agents against oxidative stress.
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
Acknowledgments
The authors acknowledge the financial support from the
European Union in the ambit of the Project BIODESERT
(EU FP7-CSA-SA REGPOT-2008-2, Grant agreement no.
245746) and the Tunisian Ministry of Higher Education and
Scientific Research in the ambit of the laboratory Project LR
MBA20. Atef Jaouani wants to thank the Tunisian Society for
Microbial Ecology (ATEM) for supporting publication fees of
this work.
References
[1] J. K. Warren, “Sabkhas, saline mudflats and pans,” in Evaporites:
Sediments, Resources and Hydrocarbons, pp. 139–220, Springer,
Berlin, Germany, 2006.
[2] A. A. Gorbushina and W. E. Krumbein, “Rock dwelling fungal
communities: diversity of life styles and colony structure,” in
Journey to Diverse Microbial Worlds, J. Seckbach, Ed., pp. 317–
334, Kluwer Academic Publishers, Amsterdam, The Netherlands, 2000.
[3] L. J. Rothschild and R. L. Mancinelli, “Life in extreme environments,” Nature, vol. 409, no. 6823, pp. 1092–1101, 2001.
[4] S. B. Humayoun, N. Bano, and J. T. Hollibaugh, “Depth distribution of microbial diversity in mono lake, a meromictic soda lake
in California,” Applied and Environmental Microbiology, vol. 69,
no. 2, pp. 1030–1042, 2003.
[5] C. Demergasso, E. O. Casamayor, G. Chong, P. Galleguillos,
L. Escudero, and C. Pedrós-Alió, “Distribution of prokaryotic
genetic diversity in athalassohaline lakes of the Atacama Desert,
Northern Chile,” FEMS Microbiology Ecology, vol. 48, no. 1, pp.
57–69, 2004.
[6] N. Gunde-Cimerman, A. Oren, and A. Plemenitaš, “Adaptation
to life in high salt concentrations,” in Archaea, Bacteria, and
Eukarya, p. 576, Springer, Dordrecht, The Netherlands, 2005.
[7] L. Maturrano, F. Santos, R. Rosselló-Mora, and J. Antón,
“Microbial diversity in Maras salterns, a hypersaline environment in the Peruvian Andes,” Applied and Environmental
Microbiology, vol. 72, no. 6, pp. 3887–3895, 2006.
[8] Q. Wu, G. Zwart, M. Schauer, M. Kamst-van Agterveld, and
M. Hahn, “Bacterioplankton community composition along
a salinity gradient of sixteen high-mountain lakes located
on the Tibetan Plateau, China,” Applied and Environmental
Microbiology, vol. 72, no. 8, pp. 5478–5485, 2006.
[9] C. Gostinčar, M. Grube, S. De Hoog, P. Zalar, and N. GundeCimerman, “Extremotolerance in fungi: evolution on the edge,”
FEMS Microbiology Ecology, vol. 71, no. 1, pp. 2–11, 2010.
10
[10] K. Sterflinger, D. Tesei, and K. Zakharova, “Fungi in hot and
cold deserts with particular reference to microcolonial fungi,”
Fungal Ecology, vol. 5, no. 4, pp. 453–462, 2012.
[11] M. J. Liszka, M. E. Clark, E. Schneider, and D. S. Clark,
“Nature versus nurture: developing enzymes that function
under extreme conditions,” Annual Review of Chemical and
Biomolecular Engineering, vol. 3, pp. 77–102, 2012.
[12] J. P. Perthuisot, La Sebkha El Melah de Zarzis: genèse et évolution
d’un bassin salin paralique; Les gisements néolithiques des abords
de la Sebkha el Melah, vol. 9 of de Travaux du Laboratoire de
géologie/École normale supérieure, École Normale Supérieure,
1975.
[13] I. cLellan, A. Varela, M. Blahgen et al., “Harmonisation of
physical and chemical methods for soil management in Cork
Oak forests—lessons from collaborative investigations,” African
Journal of Environmental Science and Technology, vol. 7, no. 6,
pp. 386–401, 2013.
[14] J. H. Warcup, “The soil-plate method for isolation of fungi from
soil,” Nature, vol. 166, no. 4211, pp. 117–118, 1950.
[15] K. H. Domsch, W. Gams, and T. H. Anderson, Compendium of
Soil Fungi, Academic Press, London, UK, 1980.
[16] J. A. von Arx, The Genera of Fungi Sporulating in Pure Culture,
J. Cramer, Vaduz, Germany, 1981.
[17] E. Kiffer and M. Morelet, Les deutéromycètes: Classification et
clés d'identification gènérique, INRA Editions, Paris, France,
1997.
[18] D. Liu, S. Coloe, R. Baird, and J. Pedersen, “Rapid minipreparation of fungal DNA for PCR,” Journal of Clinical Microbiology, vol. 38, no. 1, article 471, 2000.
[19] T. J. White, T. Bruns, S. Lee, and J. Talor, “Amplification
and direct sequencing of fungal ribosomal RNA genes for
phylogenetics,” in PCR Protocols: A Guide to Methods and
Applications, M. A. Innes, D. H. Gelfand, J. J. Sninsky, and T. J.
White, Eds., pp. 315–322, Academic Press, New York, NY, USA,
1990.
[20] G. Luo and T. G. Mitchell, “Rapid identification of pathogenic
fungi directly from cultures by using multiplex PCR,” Journal of
Clinical Microbiology, vol. 40, no. 8, pp. 2860–2865, 2002.
[21] P. Rodrigues, C. Santos, A. Venâncio, and N. Lima, “Species
identification of Aspergillus section Flavi isolates from Portuguese almonds using phenotypic, including MALDI-TOF
ICMS, and molecular approaches,” Journal of Applied Microbiology, vol. 111, no. 4, pp. 877–892, 2011.
[22] K. Bensch, U. Braun, J. Z. Groenewald, and P. W. Crous, “The
genus cladosporium,” Studies in Mycology, vol. 72, pp. 1–401,
2012.
[23] M. L. Berbee, M. Pirseyedi, and S. Hubbard, “Cochliobolus phylogenetics and the origin of known, highly virulent
pathogens, inferred from ITS and glyceraldehyde-3-phosphate
dehydrogenase gene sequences,” Mycologia, vol. 91, no. 6, pp.
964–977, 1999.
[24] N. L. Glass and G. C. Donaldson, “Development of primer
sets designed for use with the PCR to amplify conserved genes
from filamentous Ascomycetes,” Applied and Environmental
Microbiology, vol. 61, no. 4, pp. 1323–1330, 1995.
[25] S. F. Altschul, W. Gish, W. Miller, E. W. Myers, and D. J. Lipman,
“Basic local alignment search tool,” Journal of Molecular Biology,
vol. 215, no. 3, pp. 403–410, 1990.
[26] H. M. Hassan and I. Fridovich, “Paraquat and Escherichia coli.
Mechanism of production of extracellular superoxide radical,”
Journal of Biological Chemistry, vol. 254, no. 21, pp. 10846–10852,
1979.
BioMed Research International
[27] M. B. Angelova, S. B. Pashova, B. K. Spasova, S. V. Vassilev,
and L. S. Slokoska, “Oxidative stress response of filamentous
fungi induced by hydrogen peroxide and paraquat,” Mycological
Research, vol. 109, no. 2, pp. 150–158, 2005.
[28] A. de Groot, V. Chapon, P. Servant et al., “Deinococcus deserti
sp. nov., a gamma-radiation-tolerant bacterium isolated from
the Sahara Desert,” International Journal of Systematic and
Evolutionary Microbiology, vol. 55, no. 6, pp. 2441–2446, 2005.
[29] A. Chanal, V. Chapon, K. Benzerara et al., “The desert of
Tataouine: an extreme environment that hosts a wide diversity
of microorganisms and radiotolerant bacteria,” Environmental
Microbiology, vol. 8, no. 3, pp. 514–525, 2006.
[30] M. Gtari, D. Daffonchio, and A. Boudabous, “Assessment of
the genetic diversity of Frankia microsymbionts of Elaeagnus
angustifolia L. plants growing in a Tunisian date-palm oasis by
analysis of PCR amplified nifD-K intergenic spacer,” Canadian
Journal of Microbiology, vol. 53, no. 3, pp. 440–445, 2007.
[31] I. Essoussi, F. Ghodhbane-Gtari, H. Amairi et al., “Esterase as
an enzymatic signature of Geodermatophilaceae adaptability
to Sahara desert stones and monuments,” Journal of Applied
Microbiology, vol. 108, no. 5, pp. 1723–1732, 2010.
[32] I. Sbissi, M. Neffati, A. Boudabous, C. Murat, and M. Gtari,
“Phylogenetic affiliation of the desert truffles Picoa juniperi and
Picoa lefebvrei,” Antonie van Leeuwenhoek, vol. 98, no. 4, pp.
429–436, 2010.
[33] I. Sbissi, F. Ghodhbane-Gtari, M. Neffati, H. Ouzari, A. Boudabous, and M. Gtari, “Diversity of the desert truffle Terfezia
boudieri chatin in southern Tunisia,” Canadian Journal of
Microbiology, vol. 57, no. 7, pp. 599–605, 2011.
[34] C. A. Conley, G. Ishkhanova, C. P. McKay, and K. Cullings, “A
preliminary survey of non-lichenized fungi cultured from the
hyperarid Atacama Desert of Chile,” Astrobiology, vol. 6, no. 4,
pp. 521–526, 2006.
[35] E. V. Smolyanyuk and E. N. Bilanenko, “Communities of
halotolerant micromycetes from the areas of natural salinity,”
Microbiology, vol. 80, no. 6, pp. 877–883, 2011.
[36] H. P. Molitoris, A. S. Buchalo, I. Kurchenko et al., “Physiological
diversity of the fIrst fIlamentous fungi isolated from the hypersaline Dead Sea,” in Aquatic Mycology Across the Millennium, K.
D. Hyde, W. H. Ho, and S. B. Pointing, Eds., vol. 5, pp. 55–70,
Fungal Diversity Press, 2000.
[37] S. I. I. Abdel-Hafez, “Survey of the mycoflora of desert soils in
Saudi Arabia,” Mycopathologia, vol. 80, no. 1, pp. 3–8, 1982.
[38] A. H. M. El-Said and A. Saleem, “Ecological and physiological
studies on soil fungi at Western Region, Libya,” Mycobiology,
vol. 36, pp. 1–9, 2008.
[39] P. Zalar, G. S. de Hoog, H.-J. Schroers, P. W. Crous, J. Z. Groenewald, and N. Gunde-Cimerman, “Phylogeny and ecology of
the ubiquitous saprobe Cladosporium sphaerospermum, with
descriptions of seven new species from hypersaline environments,” Studies in Mycology, vol. 58, pp. 157–183, 2007.
[40] J. K. Misra, “Occurrence, distribution and seasonality of aquatic
fungi as affected by chemical factors in six alkaline ponds of
India,” Hydrobiologia, vol. 97, no. 2, pp. 185–191, 1982.
[41] F. T. El-Hissy, S. A. El-Zayat, A. M. Khallil, and M. S. Massoud,
“Aquatic fungi from the submerged mud of Aswan High Dam
Lake,” Microbiological Research, vol. 152, no. 1, pp. 27–32, 1997.
[42] A. M. Khallil, F. T. El-Hissy, and E. H. Ali, “Seasonal fluctuations
of aquatic fungi recovered from Egyptian soil (Delta Region),”
Journal of Basic Microbiology, vol. 35, no. 2, pp. 93–102, 1995.
BioMed Research International
[43] P. B. Matheny, J. A. Gossmann, P. Zalar, T. K. A. Kumar,
and D. S. Hibbett, “Resolving the phylogenetic position of the
Wallemiomycetes: an enigmatic major lineage of Basidiomycota,” Canadian Journal of Botany, vol. 84, no. 12, pp. 1794–1805,
2006.
[44] A. Blomberg and L. Adler, “Tolerence of fungi to NaCl,” in Stress
Tolerance of Fungi, D. Jennings, Ed., pp. 209–231, 1993.
[45] A. Oren, “Bioenergetic aspects of halophilism,” Microbiology
and Molecular Biology Reviews, vol. 63, no. 2, pp. 334–348, 1999.
[46] P. Delgado-Sánchez, M. A. Ortega-Amaro, J. F. JiménezBremont, and J. Flores, “Are fungi important for breaking seed
dormancy in desert species? Experimental evidence in Opuntia
streptacantha (Cactaceae),” Plant Biology, vol. 13, no. 1, pp. 154–
159, 2011.
[47] J. C. Frisvad, “Halotolerant and halophilic fungi and their
extrolite production,” in Adaptation to Life at High Salt Concentrations in Archaea , Bacteria and Eukarya, N. GundeCimerman, A. Oren, and A. Plemenitaš, Eds., pp. 426–440,
Springer, Amsterdam, The Netherlands, 2005.
[48] A. Abdel-Raheem and C. A. Shearer, “Extracellular enzyme
production by freshwater ascomycetes,” Fungal Diversity, vol. 11,
pp. 1–19, 2002.
11
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 914767, 11 pages
http://dx.doi.org/10.1155/2014/914767
Research Article
Geodermatophilus poikilotrophi sp. nov.: A Multitolerant
Actinomycete Isolated from Dolomitic Marble
Maria del Carmen Montero-Calasanz,1,2 Benjamin Hofner,3
Markus Göker,1 Manfred Rohde,4 Cathrin Spröer,1 Karima Hezbri,5
Maher Gtari,5 Peter Schumann,1 and Hans-Peter Klenk1
1
Leibniz Institute DSMZ-German Collection of Microorganisms and Cell Cultures, Inhoffenstraße 7B, 38124 Braunschweig, Germany
Instituto de Investigacióon y Formacióon Agraria y Pesquera (IFAPA), Centro Las Torres-Tomejil,
Carretera Sevilla-Cazalla de la Sierra, Km 12.2, 41200 Alcalá del Rı́o, Sevilla, Spain
3
Institut für Medizininformatik, Biometrie und Epidemiologie, Friedrich-Alexander-Universität Erlangen-Nürnberg,
Waldstraße 6, 91054 Erlangen, Germany
4
Helmholtz Centre for Infection Research (HZI), Inhoffenstraße 7, 38124 Braunschweig, Germany
5
Laboratoire Microorganismes et Biomolécules Actives, Université de Tunis Elmanar (FST) et Université de Carthage (INSAT),
2092 Tunis, Tunisia
2
Correspondence should be addressed to Maria del Carmen Montero-Calasanz; [email protected]
and Hans-Peter Klenk; [email protected]
Received 1 April 2014; Revised 3 June 2014; Accepted 9 June 2014; Published 9 July 2014
Academic Editor: Sara Borin
Copyright © 2014 Maria del Carmen Montero-Calasanz et al. This is an open access article distributed under the Creative
Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the
original work is properly cited.
A novel Gram-reaction-positive, aerobic actinobacterium, tolerant to mitomycin C, heavy metals, metalloids, hydrogen peroxide,
desiccation, and ionizing- and UV-radiation, designated G18T , was isolated from dolomitic marble collected from outcrops in
Samara (Namibia). The growth range was 15–35∘ C, at pH 5.5–9.5 and in presence of 1% NaCl, forming greenish-black coloured
colonies on GYM Streptomyces agar. Chemotaxonomic and molecular characteristics of the isolate matched those described
for other representatives of the genus Geodermatophilus. The peptidoglycan contained meso-diaminopimelic acid as diagnostic
diaminoacid. The main phospholipids were phosphatidylethanolamine, phosphatidylcholine, phosphatidylinositol, and small
amount of diphosphatidylglycerol. MK-9(H4 ) was the dominant menaquinone and galactose was detected as diagnostic sugar. The
major cellular fatty acids were branched-chain saturated acids iso-C16:0 and iso-C15:0 and the unsaturated C17:1 𝜔8c and C16:1 𝜔7c.
The 16S rRNA gene showed 97.4–99.1% sequence identity with the other representatives of genus Geodermatophilus. Based on
phenotypic results and 16S rRNA gene sequence analysis, strain G18T is proposed to represent a novel species, Geodermatophilus
poikilotrophi. Type strain is G18T (= DSM 44209T = CCUG 63018T ). The INSDC accession number is HF970583. The novel R
software package lethal was used to compute the lethal doses with confidence intervals resulting from tolerance experiments.
1. Introduction
The family cursive was originally proposed by Normand
et al. [1], but a formal description of the family name was
only published a decade later [2]. At the time of writing,
the family comprises the genera Blastococcus, Modestobacter,
and Geodermatophilus (as the type genus). Geodermatophilus
was proposed by Luedemann [3] and was included in the
Approved Lists of Bacterial Names [4]. This genus was poorly
studied for a long time due to difficulties in culturing isolates
[5], in spite of the fact that its members are frequently
isolated from arid soils [5] and occasionally from arid and
semiarid rock substrates such as rock vanish and marble
[6, 7], where a variety of environmental changing factors
influence their settlement, growth, and development [8].
Some of them were also isolated from rhizosphere soil [9, 10].
To enable the survival in such extreme ecological niches,
where bacterial cells are suppressed to reactive oxygen species
2
(ROS) generating-stresses, those should exhibit a very broad
range of tolerance to multiple and fluctuating environmental
stresses, such as solar radiation, desiccation and rehydration,
temperature fluctuations, salts, and metals [8, 11], and a
probable ionizing-radiation (IR) resistance. The origin of
this last capability cannot be explained as adaptation to
environment, suggesting an “incidental” result of tolerance
to desiccation, whose DNA damage pattern is similar to that
generated by ionizing radiation in Deinococcus species [12].
Furthermore, tolerance to hydrogen peroxide and mitomycin
C as indicators of the presence of an efficient microbial
oxidative stress repair and double-strand break repair system,
characteristics also attributed to radiation resistance, have
been widely studied [13, 14]. Multiple-stress tolerance of the
type strain Geodermatophilus obscurus was already described
by Gtari et al. [11], suggesting a correlation between tolerance
profiles to desiccation, mitomycin C, hydrogen peroxide, and
ionizing- and UV-radiation. Previous works of Rainey et al.
[15] and Giongo et al. [16] already revealed the prevalence of
IR resistant Geodermatophilus isolates from arid soil sample
at comparatively the same radiation levels as observed for
Deinococcus species and the predominance of species belongs
to the family Geodermatophilaceae detected from intercontinental dust, illustrating, therefore, to resist radiation and
desiccation stresses during travel in the high atmosphere.
Fourteen named species have been classified in the
genus Geodermatophilus (ordered by the dates of effective
publication of the names): G. obscurus [3], G. ruber [9],
G. nigrescens [17], G. arenarius [18], G. siccatus [19, 20], G.
saharensis [20, 21], G. tzadiensis [22, 23], G. telluris [24], G.
soli and G. terrae [10], G. africanus [5, 23], G. normandii
[25], G. taihuensis [26], and G. amargosae [27, 28]. Until now,
only the genome of the type strain of the type species, G.
obscurus G-20T , has been sequenced [29]. Moreover, three
subspecies have been identified and named, but their names
were not validly published yet: “G. obscurus subsp. utahensis,”
“G. obscurus subsp. dictyosporus” [3], and “G. obscurus subsp.
everesti” [30, 31]. This study describes the taxonomic position
of a novel species into the genus Geodermatophilus based
on a polyphasic approach and its tolerance to different
environmental stresses.
2. Materials and Methods
2.1. Isolation. During screening for microorganisms from
dolomitic marble outcrops in an agriculture area at 1150 masl
in Samara, near to Namib desert (Namibia), a greenish-black
strain designated as G18T was isolated (in 1993) and purified
as described by Eppard et al. [7].
2.2. Morphological and Biochemical Characterization. Cultural characteristics were tested on GYM Streptomyces
medium (DSMZ medium 65), TSB agar (DSMZ medium
535), GPHF medium (DSMZ medium 553), R2A medium
(DSMZ medium 830), GEO medium (DSMZ medium 714),
PYGV medium (DSMZ medium 621), and Luedemann
medium (DSMZ medium 877) for 15 days at 28∘ C. To
determine its morphological characteristics, strain G18T was
BioMed Research International
cultivated on GYM Streptomyces medium at 28∘ C. Colony
features were observed at 4 and 15 days under a binocular
microscope according to Pelczar Jr. [32]. Exponentially growing bacterial cultures were observed with an optical microscope (Zeiss AxioScope A1) with a 100-fold magnification
and phase-contrast illumination. Micrographs of bacterial
cells grown on GYM Streptomyces broth after 7 days were
taken with a field-emission scanning electron microscope
(FE-SEM Merlin, Zeiss, Germany). Gram reaction was performed using the KOH test described by Gregersen [33]. Cell
motility was observed on modified ISP2 [34] swarming agar
(0.3%, w/v) at pH 7.2 supplemented with (l−1 ) 4.0 g dextrin,
4.0 g yeast extract, and 10.0 g malt extract. Oxidase activity
was analysed using filter-paper disks (Sartorius grade 388)
impregnated with 1% solution of N,N,N 󸀠 ,N 󸀠 -tetramethylp-phenylenediamine (Sigma-Aldrich); a positive test was
defined by the development of a blue-purple colour after
applying biomass to the filter paper. Catalase activity was
determined based on formation of bubbles following the
addition of 1 drop of 3% H2 O2 . Growth rates were determined
on plates of GYM Streptomyces medium for temperatures
from 10 to 50∘ C at 5∘ C increments and for pH values from
4.0 to 12.5 (in increments of 0.5 pH units) on modified
ISP2 medium by adding NaOH or HCl, respectively, since
the use of a buffer system inhibited growth of the strains.
The utilization of carbon compounds and acid production
were tested at 28∘ C using API 20 NE strips (bioMérieux)
and GEN III Microplates in an Omnilog device (BIOLOG
Inc., Hayward, CA, USA) in comparison with the reference
strains G. africanus DSM 45422T , G. amargosae DSM 46136T ,
G. arenarius DSM 45418T , G. nigrescens DSM 45408T , G.
normandii DSM 45417T , G. obscurus DSM 43160T , G. ruber
DSM 45317T , G. saharensis DSM 45423T , G. siccatus DSM
45419T , G. soli DSM 45843T , G. taihuensis DSM 45962T , G.
telluris DSM 45421T , G. terrae DSM 45844T , and G. tzadiensis
DSM 45416𝑇 in parallel assays. The GEN III Microplates were
inoculated with cells suspended in a viscous inoculating fluid
(IF C) provided by the manufacturer at a cell density of 70%
transmittance (T) for G. amargosae DSM 46136T , at 75–79%
T for G. africanus DSM 45422T , at 90% T for G. arenarius
DSM 45418T and G. taihuensis DSM 45962T , and at 80–83% T
for all other reference strains. Respiration rates (and growth)
were measured yielding a total running time of 5 or 10 days,
depending on the strain, in phenotype microarray mode.
Each strain was studied in two independent repetitions. Data
were exported and analysed using the opm package for R
[35, 36] v.1.0.6. Reactions with a distinct behaviour between
the two repetitions were regarded as ambiguous. Clustering
analyses of the phenotypic microarrays were constructed
using the pvclust package for R v.1.2.2. [37]. Enzymatic
activities were tested using API ZYM galleries according to the instructions of the manufacturer (bioMérieux).
Chemotaxonomic procedures. Whole-cell sugars were prepared according to Lechevalier and Lechevalier [38], followed
by thin layer chromatography (TLC) analysis [39]. Polar
lipids were extracted, separated by two-dimensional TLC,
and identified according to procedures outlined by Minnikin
et al. [40] with modifications proposed by Kroppenstedt and
BioMed Research International
Goodfellow [41]. Additionally, choline-containing lipids were
detected by spraying with Dragendorff ’s reagent (Merck)
[42]. Menaquinones (MK) were extracted from freeze-dried
cell material using methanol as described by Collins et al. [43]
and analysed by high-performance liquid chromatography
(HPLC) [44]. The extraction and analysis of cellular fatty
acids were carried out in two independent repetitions from
biomass grown on GYM agar plates held at 28∘ C for 4
days and harvested always from the same sector (the last
quadrant streak). Analysis was conducted using the microbial
identification system (MIDI) Sherlock Version 4.5 (method
TSBA40, ACTIN6 database) as described by Sasser [45]. The
annotation of the fatty acids in the ACTIN6 peak naming
table is consistent with IUPAC nomenclature (i.e., double
bond positions identified with reference to the carboxyl
group of the fatty acid), but for consistency with other
publications this has been altered to numbering from the
aliphatic end of the molecule (i.e., 16 : 1 CIS 9 become 16 : 1
𝜔7c, etc.). The composition of peptidoglycan hydrolysates (6
N HCl, 100∘ C for 16 h) was examined by TLC as described by
Schleifer and Kandler [46]. All chemotaxonomical analyses
were conducted under standardized conditions with strain
G18T and cultures of the same set of reference strains as listed
above for morphological and biochemical characterisations.
2.3. Genetic and Phylogenetic Analysis. G + C content of
chromosomal DNA of strain G18T was determined by HPLC
according to Mesbah et al. [47]. Genomic DNA extraction,
PCR-mediated amplification of the 16S rRNA gene, and
purification of the PCR product were carried out as described
by Rainey et al. [48]. Phylogenetic analysis was based on an
alignment of 16S rRNA gene sequences from type strains of
all species with effectively published names in the Geodermatophilaceae inferred as described by Montero-Calasanz et
al. [5]. Pairwise similarities were calculated as recommended
by Meier-Kolthoff et al. [49]. For DNA-DNA hybridization
tests, cells were disrupted by using a Constant Systems TS
0.75 KW (IUL Instruments, Germany). DNA in the crude
lysate was purified by chromatography on hydroxyapatite as
described by Cashion et al. [50]. DNA-DNA hybridization
was carried out as described by De Ley et al. [51] under
consideration of the modifications described by Huss et al.
[52] using a model Cary 100 Bio UV/VIS-spectrophotometer
equipped with a Peltier-thermostatted 6 × 6 multicell changer
and a temperature controller with in situ temperature probe
(Varian).
2.4. Tolerance Experiments. The tolerance of strain G18T
and G. obscurus G-20T (DSM 43160), as a positive control
[11], to ionizing- and UV-radiation, mitomycin C, hydrogen peroxide, desiccation, and heavy metals/metalloids, was
assayed using nonsporulating cultures obtained by growth
in TYB medium [53] at 28∘ C for 5 days, washed twice with
0.9% NaCl, homogenized, and subsequently resuspended in
saline solution. Ionizing-radiation experiments were carried
out according to a protocol outlined by Gtari et al. [11].
To test the resistance to UV-radiation, 0.5 mL aliquots of
culture suspensions was spread onto GYM Streptomyces agar
3
plates in duplicate in two independent experiments and then
exposed to a dose of 5–10 J⋅s−1 m−2 in a laminar flow hood
equipped with crossbeam 254 nm UV sources in both side
walls (Safe 2020, Thermo Scientific) for 1, 10, 30, 60, 120, 240,
and 600 min. After 2 weeks at 28∘ C, the survival fractions
were calculated based on the c.f.u. mL−1 . The UV shadow
zone was avoided. The tolerance to DNA damaging agent
mitomycin C was tested in two independent experiments
by incubation of cell suspension at room temperature with
the antibiotic at a final concentration of 5 𝜇g⋅mL−1 . After
1, 5, 10, 20, 40, 60, and 120 min, samples were centrifuged
at 3500 rpm for 4 min, washed twice in 0.9% NaCl, and,
subsequently, serially diluted. Aliquots were spread on GYM
Streptomyces agar in duplicate. After incubation, the survival
fractions were calculated based on the c.f.u. mL−1 . To test
the resistance to hydrogen peroxide, equal volumes of cell
suspensions and 0.5% hydrogen peroxide were incubated at
room temperature in two independent experiments. After 1,
5, 10, 20, 40, 60, and 120 min, samples were handled as was
previously described in mitomycin experiments to calculate
the survival fractions. For desiccation tolerance, 25 𝜇L of cell
suspension were transferred to individual wells of microtiter
plates in triplicate. Unsealed microtiter plates were placed in
a desiccator (23.5% relative humidity) containing silica gel
rubin (Fluka) at room temperature. After 20, 40, 60, 80, and
100 days, 250 𝜇L of sterile water was added to individual wells
to rehydrate the desiccated cells and then incubated at room
temperature for 1 hour and plated on GYM Streptomyces
agar. The determination of survival fractions was conducted
as described above. The sensitivity of strain G18T to heavy
metals and metalloids was determined by a growth inhibition
plate assay as described by Richards et al. [54]. AgNO3 ,
CuCl2 , CoCl2 , NiCl2 , K2 CrO4 , Pb(NO3 )2 , and Na2 HAsO4
were added to GYM Streptomyces medium at 0.1, 0.3, 0.5, 1.0,
2.0, 4.0, 8.0, 10.0, 30.0, and 50.0 mM. Growth was evaluated
after 1 month at 28∘ C, determining minimum inhibitory
concentration (MIC).
2.5. Statistical Analysis of Tolerance Experiments. To evaluate
the tolerance of strain G18T and G. obscurus DSM 43160T
with respect to the various physiological challenges, the
median lethal dose (LD50) and the lethal dose 10 (LD10)
values were computed for both strains. As the number of
bacteria initially used in each experiment cannot directly
be obtained and consequently, death rates or survival rates
cannot be directly computed; standard models based on
logistic regression models to obtain LD values are thus not
available. A negative binomial model for count data [55]
was used to estimate of number of survivors dependent on
dose, strain, and experiment. Penalized splines [56], one for
each strain, were used to allow the dose to have a nonlinear
influence on survival fractions. The estimation process was
stabilised by using of a square root transformation on dose.
LD50 and LD10 values were subsequently estimated from the
model and 95% confidence intervals were obtained using a
parametric bootstrap approach [57, Chapter 5.4]. Details on
model fitting and the estimation of the confidence intervals
as well as code to derive LD values from survival count
4
Figure 1: Scanning electron micrograph of strain G18T grown on
GYM Streptomyces medium for 7 days at 28∘ C.
data with one or two strains can be found in the supplementary material (see Figure S4 in Supplementary Material
available online at http://dx.doi.org/10.1155/2014/914767). All
computations were done with R [58] using the R software
packages mgcv [57] and lethal [59].
3. Results and Discussion
3.1. Morphological and Biochemical Characteristics. Cells of
strain G18T were pleiomorphic and Gram-reaction-positive.
Individual cells and aggregates were observed, confirming
reports by Ishiguro and Wolfe [53] of synchronous morphogenesis on unspecific media and previous observations on
other representatives of the genus Geodermatophilus [27].
In line with the original description by Luedemann [3],
circular or elliptical motile zoospores and septated filaments
from zoospore germination were observed (Figure 1). Young
colonies were light-red in colour and turned greenish-black at
maturity. Similar colours conversions were already observed
by Nie et al. [17] and Montero-Calasanz et al. [18, 19, 21,
22, 25] for type strains of other representatives of the genus,
such as G. nigrescens, G. arenarius, G. siccatus, G. saharensis,
G. tzadiensis, and G. normandii, when cultivated under the
same growth conditions (Table 1). Colonies were convex,
nearly circular and opaque with a moist surface and an entire
margin. Strain G18T grew well on GYM Streptomyces and
GEO media but did not grow on TSA, R2A, GPHF, PYGV,
and Luedemann media. It grew best at 25–30∘ C but did not
grow below 15∘ C or above 35∘ C. Growth was observed in
presence of 1% NaCl and between pH 5.5–9.5 (optimal range
pH 7.0–9.5). Results from phenotype microarray analysis
are shown as a heatmap in the supplementary material
(Figure S1) in comparison to the reference type strains of the
genus Geodermatophilus. A summary of selected differential
phenotypic characteristics is presented in Table 1. In the
phenotypic clustering significant support (>95%) is obtained
for G. poikilotrophi DSM 44209T , G. nigrescens DSM 45408T
and G. normandii DSM 45417T being most similar to each
other regarding the characters present in GEN III Microplates
(Suppl. Figure S2).
3.2. Chemotaxonomic Characteristics. Analysis of cell wall
components revealed the presence of DL-diaminopimelic
BioMed Research International
acid (cell wall type III), which is consistent with other species
of the genus Geodermatophilus [27, 38]. Strain G18T displayed
primarily menaquinone MK-9(H4 ) (82.5%), in agreement
with values reported for the family Geodermatophilaceae [2],
but also MK-9(H0 ) (8.8%) and MK-9(H2 ) (4.8%). Major
fatty acids were iso-C16:0 (24.5 ± 0.2%), iso-C15:0 (16.6 ±
1.3%), C17:1 𝜔8c (13.9 ± 0.1%) and C16:1 𝜔7c (8.3 ± 0.1%),
complemented by iso-C16:1 H (5.6 ± 0.9%), anteiso-C15:0
(4.1 ± 0.4%), anteiso-C17:0 (4.4 ± 0.2%), C18:1 𝜔9c (3.6 ±
0.1%) and C16:0 (2.4 ± 0.9%). The phospholipids pattern
consisted of phosphatidylethanolamine (PE), phosphatidylcholine (PC), phosphatidylinositol (PI), and small amount
of diphosphatidylglycerol (DPG) in accordance with profiles obtained for representatives of other Geodermatophilus
species investigated in this study (Table 1). Phosphatidylglycerol was not detectable (see Supplementary Figure S3).
This fact was already predictable based on phospholipids
profiles displayed for other representatives of the genus such
as G. arenarius, G. siccatus, G. tzadiensis, G. normandii, or G.
amargosae, whose phosphatidylglycerol amounts were nearly
imperceptible. Whole-cell sugar analysis revealed galactose
as the diagnostic sugar [38] but also glucose and ribose.
Genomic G + C content was 74.4 mol%.
3.3. Molecular Analysis. The almost complete (1514 bp) 16S
rRNA gene sequence of strain G18T was determined. The 16S
rRNA sequence showed the highest degree of similarity with
the type strains of G. siccatus (99.1%), G. africanus (99.0%),
G. amargosae (98.5%), G. normandii (98.4%), G. obscurus
(98.3%), G. tzadiensis (98.2%), G. nigrescens (98.1%), G. ruber
(98.0%), and G. arenarius (98.0%). All listed closely related
type strains were placed within the same phylogenetic group
by both, maximum likelihood and maximum-parsimony
estimations (Figure 2). The 16S rRNA gene sequences analysis
thus strongly supports the assignment of strain G18T to the
genus Geodermatophilus. However, similarities in 16S rRNA
gene sequence between G18T and some closely related type
strains indicated the need to prove the genomic distinctness
of the type strain representing the novel species by DNADNA hybridization. Strain G18T displayed a DNA-DNA
relatedness of 35.3 ± 1.0% with the type strain of G. siccatus
and 28.1 ± 2.1% with G. africanus. DNA-DNA hybridizations
of strain G18T with the type strains of G. amargosae, G.
normandii, G. obscurus, G. tzadiensis, G. nigrescens, G. ruber,
and G. arenarius were not conducted, according to MeierKolthoff et al. [49] that statistically confirmed that the
threshold value previously established at 97% 16S rRNA gene
sequence similarity was too conservative in microbial species
discrimination and determined a Actinobacteria-specific 16S
rRNA threshold at 99.0% with a maximun probability of error
of 1.00% to get DNA-DNA hybridization values above the
70% threshold recommended by Wayne et al. [60] to confirm
the species status of novel strains.
3.4. Tolerance. Gamma-radiation survival of strain G18T
(Figure 3(a)) showed not significantly different inactivation
kinetic as for G. obscurus DSM 43160T , which is considered
i-C15:0 ,
i-C16:0 ,
C17:1 𝜔8c
Major fatty acidsb
4
5
+
+
+
+
+/−
−
+/−
+/−
−
−
−
−
+
+
−
+
+/−
+/−
+
Moist
+
−
−
+
Moist
+
−
+
+
−
−
−
−
+
+
−
−
−
−
−
MK9(H4 ),
MK-9(H4 ),
MKMKMK-9(H4 )
MK-9(H2 ),
8(H4 ),
9(H4 )
MK-8(H4 )
MK-9
(H0 )
PE, PC,
DPG, PC, PE, DPG, PE, PC, DPG, PE,
DPG, PI,
PI, PG
PI, 2PL, PG PC, PI, PG
PG
i-C15:0 ,
ai-C15:0 ,
i-C15:0 ,
i-C15:0 ,
i-C15:0 ,
i-C16:0 ,
i-C16:0 ,
i-C16:0
i-C16:0
C17:1 𝜔8c
ai-C17:0 ,
C17:1 𝜔8c
+
+
−
−
+
−
−
−
−
+/−
−
+
−
−
+
+
+
−
−
Moist
Light-red, Light-red,
Light-red, red
black
brown
3
6
7
MK-9(H4 ),
MK-8(H4 )
MK-9(H4 ),
MK-8(H4 ),
MK-9(H0 )
i-C15:0 ,
i-C16:0 ,
C17:1 𝜔8c
MK-9(H4 ),
MK-9(H0 )
+
+/−
+
+
+
+
+
−
+
−
+
−
+
+
−
+
+
+
+
8
Light-red,
greenishblack
Moist
MK-9(H4 )
+
+
+
+
−
+
+
+
+
−
+
−
+
+
+
+
−
−
+
Dry
Black
9
i-C15:0 ,
i-C16:0 ,
i-H-C16:1
i-C15:0 ,
i-C16:0
i-C15:0 ,
i-C16:0
DPG, PC, PI, DPG, PC, PE, DPG, PC, PE,
PE, PG
PI, PG
PI, APL, PG
+
+
+
+
+
+
+
+
−
−
+
+
+
+
−
−
−
+
−
−
+
+
+
+
+
+
−
+
+
+
PE, PC, PI,
DPG, PG
+
−
+/−
+
Moist
Light-red,
black
−
−
−
−
Moist
Light-red,
black
i-C15:0 ,
i-C16:0 ,
C17:0
DPG, PME,
PE, PI, 3PL†
MK-9(H4 ),
MK-9(H0 ),
MK-9(H2 )
+/−
−
+
+
+
+/−
+
−
+
+
−
−
+
+
+
+
+
+
+
Moist
Light red
10
i-C15:0 ,
i-C16:0 ,
C18:1 𝜔9c
DPG, PME,
PE, PI, 5PL†
MK-9(H4 ),
MK-9(H0 )
+
−
+
+
−
+/−
+
−
+
+
−
−
+
+
+
+
−
−
+/−
Moist
Light red
11
MK9(H4 )
+
−
+
+
+
−
−
−
+
−
−
−
+
+
−
+
−
+/−
+/−
MK-9
(H4 ), MK9(H0 )#
+
+
−
−
+
+/−
+
+
+
+/−
−
−
+
−
+
+
+/−
+
+
13
14
Light-red,
greenish- Coral pink
black
Moist
Moist
MK9(H4 )
+/−
+/−
+/−
−
−
+/−
+/−
−
−
+
−
+/−
−
−
−
+
−
+
−
Dry
Black
15
i-C16:0
i-C15:0 ,
i-C16:0
i-C15:0 ,
i-C16:0 ,
C17:1 𝜔8c
i-C15:0 ,
i-C16:0
DPG, PC, DPG, PC, DPG, PE, DPG, PC,
PE, PI, PG PE, PI, PG PI, PIM# PE, PI, PG
MK9(H4 )
+
+
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
Dry
Black
12
+, positive reaction; −, negative reaction; +/−, ambiguous; MK, menaquinones; DPG, diphosphatidylglycerol; PE, phosphatidylethanolamine; PME, phosphatidyl-N-methylethanolamine; PE-OH, hydroxyphosphatidylethanolamine; PG, phosphatidylglycerol; PC, phosphatidylcholine; PI, phosphatidylinositol; PIM, phosphatidylinositol mannoside; PL, unknown phospholipid; APL, unknown amino-phospholipid; i-,
iso-branched; ai-, anteiso-branched.
a
Only components making up ≥ 5% peak area ratio are shown; b only components making up ≥ 10% peak area ratio are shown; ∗ the components are listed in decreasing order of quantity.
†
Data taken from Jin et al. [10]. # Data taken from Qu et al. [26].
PE, PC, PI,
DPG
+
+/−
+
−
−
+
+
+
+
−
−
−
+
+
+
+
−
+
+
−
+
+
−
+
−
+
−
+
+
+
Phospholipids∗
+
+/−
−
+/−
−
+
−
−
MK-9(H4 )
Dry
Black
2
1
Light-red,
greenishblack
Moist
Predominant
menaquinone(s)a
Colony surface on GYM
Utilization of
Turanose
Stachyose
D-Melibiose
D-Salicin
NaCl range (w/v)
1%
4%
D-Mannose
L-Rhamnose
Inosine
D-Sorbitol
D-Mannitol
D-Arabitol
Glycerol
L-Alanine
L-Arginine
L-Histidine
Pectin
D-Gluconic acid
Quinic acid
Colony colour on GYM
Characteristics
Table 1: Differential phenotypic characteristics of strain G18T and the type strains of other Geodermatophilus species. Strains: 1, G. poikilotrophi sp. nov. G18T ; 2, G. obscurus DSM 43160T ; 3,
G. ruber DSM 45317T ; 4, G. nigrescens DSM 45408T ; 5, G. arenarius DSM 45418T ; 6, G. siccatus DSM 45419T ; 7, G. saharensis DSM 45423T ; 8, G. tzadiensis DSM 45416T ; 9, G. telluris DSM
45421T ; 10, G. soli DSM 45843T ; 11, G. terrae DSM 45844T ; 12, G. africanus DSM 45422T ; 13, G. normandii DSM 45417T ; 14, G. taihuensis DSM 45962T ; 15, G. amargosae DSM 46136T . All
physiological data are from this study.
BioMed Research International
5
6
BioMed Research International
Geodermatophilus arenarius DSM 45418T (HE654547)
84/67
Geodermatophilus telluris DSM 45421T (HE815469)
Geodermatophilus tzadiensis DSM 45416T (HE654545)
88/75
79/83
Geodermatophilus saharensis DSM 45423T (HE654551)
Geodermatophilus amargosae DSM 46136T (HF679056)
Geodermatophilus normandii DSM 45417T (HE654546)
82/66
Geodermatophilus nigrescens DSM 45408T (JN656711)
Geodermatophilus africanus DSM 45422T (HE654550)
81/77
—/64
68/66
79/73
Geodermatophilus poikilotrophi G18T (HF970583)
Geodermatophilus obscurus DSM 43160T (X92356)
Geodermatophilus siccatus DSM 45419T (HE654548)
Geodermatophilus ruber DSM 45317T (EU438905)
69/85
Geodermatophilus soli DSM 45843T (JN033772)
Geodermatophilus taihuensis DSM 45962T (JX294478)
100/100
Geodermatophilus terrae DSM 45844T (JN033773)
Modestobacter roseus DSM 45764T (JQ819258)
73/79
Modestobacter marinus DSM 45201T (EU18 1225)
100/100
Modestobacter versicolor DSM 16678T (AJ871304)
Modestobacter multiseptatus DSM 44406T (Y18646)
T
Blastococcus jejuensis DSM 19597 (DQ200983)
—/64
Blastococcus aggregatus DSM 4725T (L40614)
Blastococcus saxobsidens DSM 44509T (FN600641)
Blastococcus endophyticus DSM 45413T (GQ494034)
0.008
Figure 2: Maximum likelihood phylogenetic tree inferred from 16S rRNA gene sequences, showing the phylogenetic position of strain G18T
relative to the type strains within the family cursive. The branches are scaled in terms of the expected number of substitutions per site (see
size bar). Support values from maximum-likelihood (left) and maximum-parsimony (right) bootstrapping are shown above the branches if
equal to or larger than 60%.
as highly resistant, according to data reported by Gtari
et al. [11]. Strain G18T strains exhibited a shoulder of resistance similar to D. radiodurans R1 to approximately 5 KGy
[61], but comparatively lower than the observed one by G.
obscurus DSM 43160T . Nevertheless, LD10 of both, G18T
and G. obscurus DSM 43160T , was around 9 KGy, a dose
comparatively higher than the displayed one for the high
radiation resistant strain D. radiodurans R1 [61], although
other authors reported a LD10 around 10 KGy by using the
same strain [62]. UV-radiation survival curves revealed a
similar progressive loss of viability in both strains during the
first 10 min of exposure until levels below 50%. However, the
differences between the two resistant phenotypes increased
along the curve, observing a significant variation on viability
at 10% survival (Figure 3(b)). According to radiated doses,
strain G18T and G. obscurus DSM 43160T were capable
to support the lethal effects of 6300–12600 J⋅s−1 m2 and
63600–31800 J⋅s−1 m2 , respectively, sustaining a survival rate
higher than 10%. Battista [63] and Shukla et al. [62] reported
LD10 values of 700–1000 J⋅s−1 m2 for the highly resistant
D. radiodurans R1. The tolerance to UV-radiation in the
genus Geodermatophilus was already observed, in addition
to G. obscurus DSM 43160T , in G. tzadiensis DSM 45416T
by Montero-Calasanz et al. [22]. Cultures of strain G18T
tolerated an exposure to mitomycin of nearly 120 min showing a viability rate of 10%, a value significantly higher than
the one observed for the positive control (LD10 = 71 min)
(Figure 3(c)). Tolerance of strain G18T (LD10 = 7 min)
in comparison with the positive control G. obscurus DSM
43160T (LD10 = 8 min) to 0.5% hydrogen peroxide along the
curves did not show any significant differences (Figure 3(d)).
Based on desiccation survival curves given in Figure 3(e),
both strains initially exhibited a similar resistance (LD50).
At the first sample point (20 days), strain G18T showed a
survival of less than 10%, a value comparatively different to
the results observed by G. obscurus DSM 43160T , whose LD10
is reached after 38 days. However, it is worth mentioning
that after 110 days a remaining bacterial population of strain
G18T was still observed. Strain G18T demonstrated thus a
high tolerance to ROS-generating stresses gamma- and UVradiation, mitomycin C, hydrogen peroxide, and desiccation comparable to the positive control G. obscurus DSM
43160T and, in general terms, to DNA damaging-resistant
D. radiodurans R1. This correlative tolerance between ROSgenerating stresses was already widely described [11, 62] and
support the hypothesis of efficient and common cellular
BioMed Research International
7
50000
c.f.u (mL−1 )
c.f.u (mL−1 )
10000
5000
20000
10000
5000
2000
1000
500
1000
500
100
50
10
2
0
6
8
4
Radiation dose (KGy)
0
10
200
400
Time (min)
600
Lethal dose(s)
Lethal dose(s)
LD50, strain: 43160T
LD10, strain: 43160T
LD50, strain: G18T
LD10, strain: G18T
LD50, strain: 43160T
LD50, strain: G18T
LD10, strain: 43160T
LD10, strain: G18T
95% confidence intervals of lethal doses
95% confidence intervals of lethal doses
Strain: G18T
Strain: 43160T - G18T
−5
0
5
LD10
LD20
Strain: G18T
LD30
LD40
Strain: 43160
LD10
LD20
LD30
LD40
LD50
LD60
Strain: 43160T
−10
T
T
Strain: 43160 - G18
T
LD50
LD60
0
10
50
100
150
200
Time (min)
Radiation dose (KGy)
(a)
(b)
50000
360000
20000
250000
c.f.u (mL−1 )
c.f.u (mL−1 )
490000
10000
5000
2000
160000
90000
40000
1000
10000
500
0
0
20
40
60
80
Time (min)
0
120
100
LD50, strain: 43160T
LD10, strain: 43160T
LD10, strain: G18T
LD10
LD20
Strain: G18T
LD30
LD40
LD50
LD60
T
50
100
120
0
50
LD50, strain: G18T
LD10, strain: G18T
95% confidence intervals of lethal doses
T
Strain: 43160 - G18
60
80
Time (min)
LD50, strain: 43160T
LD10, strain: 43160T
LD50, strain: G18T
95% confidence intervals of lethal doses
T
40
Lethal dose(s)
Lethal dose(s)
Strain: 43160
20
T
LD10
LD20
Strain: G18T
LD30
LD40
Strain: 43160
T
Strain: 43160 - G18
100
T
LD50
LD60
2
0
2
Time (min)
4
Time (min)
(c)
(d)
Figure 3: Continued.
6
8
10
8
BioMed Research International
c.f.u (mL−1 )
1e + 06
1e + 05
1e + 04
1e + 03
1e + 02
0
20
40
60
Time (days)
Lethal dose(s)
LD50, strain: 43160T
LD10, strain: 43160T
80
100
LD50, strain: G18T
LD10, strain: G18T
95% confidence intervals of lethal doses
Strain: 43160T
LD10
LD20
T
LD30
LD40
Strain: 43160T - G18T
LD50
LD60
Strain: G18
0
10
20
30
40
50
Time (days)
(e)
Figure 3: Estimation of survival following exposure to gamma-radiation (a), UV-radiation (b), mitomycin C (c), hydrogen peroxide (d), and
desiccation (e) for strain G18T and G. obscurus DSM 43160T as positive control. The mean c.f.u.mL−1 per strain is given together with the
LD50 and LD10 values in the upper panel of each figure; 𝑦-axis is on a logarithmic scale ((a)–(c), (e)), or on a square root scale (d). The lower
panel depicts LD10 and LD50 values per strain and the differences between strains together with confidence intervals. Confidence intervals
that do not contain zero (dashed vertical line) indicate significant differences to zero; in case of strain differences this indicates significant
differences between strains.
DNA repair mechanisms. Strain G18T showed the highest
tolerance to AsO4 3− (MIC = 8.0 mM) followed by Pb2+
(MIC = 4.0 mM), CrO4 2− (MIC = 4.0 mM) and Ag1+ (MIC
= 1.0 mM). Whereas the growth of G. obscurus DSM 43160T
was mainly inhibited by concentrations below 1.0 mM, except
AsO4 3− whose sensitivity was 10 times higher (MIC =
80.0 mM) than the one observed for strain G18T (Table 2). It
has been widely described that the heavy metals/metalloids
exposure also produces ROS generation [64]. In this study, a
correspondence with other ROS-generating stresses was not
observed, in agreement with data reported by Gtari et al.
[11] for G. obscurus DSM 43160T , but also for Modestobacter
multiseptatus BC501 and Blastococcus saxobsidens DD2, suggesting the presence of alternative mechanisms to counteract
the heavy metals/metalloids stress, such as transport outside
the cells [65], adsorption on exocellular structures such as
melanin [66], or enzymatic reduction to less toxic forms [67,
68]. Although it is noteworthy that toxicity levels of lead and
copper in G. obscurus DSM 43160T by comparison with the
results displayed by Gtari et al. [11] were much different from
each other. These divergences in the levels of tolerance might
be due to the differences in the media compositions [69]. In
addition, it was confirmed that neither phosphate buffer nor
carbon source concentration present in GYM Streptomyces
medium caused an overestimated metals tolerance of strains,
justified by the different tolerance range found in both strains
and its mostly correlation with the results described by Gtari
et al. [11].
Apart from the phylogenetic analysis based on 16S rRNA
gene sequences, several phenotypic features support the
distinctiveness of strain G18T from representatives of all
other Geodermatophilus species (Table 1). Based on the
phenotypic and genotypic data presented, we propose that
strain G18T represents a novel species within the genus Geodermatophilus, with the name Geodermatophilus poikilotrophi
sp. nov.
Description of Geodermatophilus poikilotrophi sp. nov.. Geodermatophilus poikilotrophi (poi.kil.o.troph’i N. L. fem. gen. n.
poikilotrophi referring to a bacterium that can tolerate diverse
environmental stresses).
Colonies are greenish-black-coloured, circular, and convex with a moist surface. Cells are Gram-reaction-positive,
catalase positive, and oxidase negative. No diffusible pigments are produced on any of the tested media. Utilizes dextrin, D-maltose, D-trehalose, D-cellobiose, sucrose,
stachyose, D-glucose, D-mannose, D-fructose, D-galactose,
L-rhamnose, D-sorbitol, D-mannnitol, myo-inositol, glycerol, L-arginine, pectin, D-gluconic acid, quinic acid, methyl
BioMed Research International
9
Table 2: Minimum inhibitory concentration of seven heavy metals and metalloids for strain G18T and G. obscurus DSM 43160T .
Strain
G18T
DSM 43160T
AgNO3
1.0
0.3
CuCl2
0.1
0.1
CoCl2
0.3
0.3
pyruvate, D-lactic acid methyl ester, 𝛼-ketoglutaric acid,
D-malic acid, bromosuccinic acid, potassium tellurite, Υamino-N-butyric acid, acetoacetic acid, propionic acid,
acetic acid, as sole carbon source for energy and growth,
but not turanose, D-raffinose, D-melibiose, 𝛽-methyl-Dglucoside, D-salicin, N-acetyl-D-glucosamine, N-acetyl-Dgalactosamine, N-acetylneuraminic acid, 3-O-methyl-Dglucose, D-fucose, inosine, sodium lactate, D- and Lserine, D-arabitol, D-glucose-6-phosphate, D-aspartic acid,
glycyl-L-proline, L-alanine, L-glutamic acid, L-histidine, Lpyroglutamic acid, L-galactonic acid-𝛾-lactone, glucuronamide, mucic acid, D-saccharic acid, p-hydroxyphenylacetic
acid, citric acid, 𝛾-amino-n-butyric acid, and butyric acid.
Acid is produced from L-arginine and Υ-amino-N-butyric
acid and can be used as a sole nitrogen source but
not N-acetyl-D-glucosamine, N-acetyl-D-galactosamine, Nacetyl-neuraminic acid, D- and L-serine, D-aspartic acid,
glycyl-L-proline, L-alanine, L-histidine, L-glutamic acid,
L-histidine, L-pyroglutamic acid, glucuronamide, and 𝛾amino-n-butyric acid. Positive for aesculin degradation.
Negative for reduction of nitrate, denitrification, indole
production and gelatin degradation. Tests for alkaline
phosphatase, esterase lipase (C8), esterase (C4), leucine
arylamidase and 𝛼-glucosidase are positive, but those
for urease, 𝛽-glucosidase, acid phosphatase, valine arylamidase, Naphthol-AS-BI-phosphohydrolase, lipase (C14),
cystine arylamidase, trypsin, 𝛼-chymotrypsin, 𝛼- and 𝛽galactosidase, 𝛽-glucuronidase, N-acetyl-𝛽-glucosamidase,
𝛼-mannosidase and 𝛼-fucosidase are negative. Cell growth
ranges from 15 to 35∘ C and from pH 5.5 to 9.5. It is tolerant to gamma- and UV-radiation, mitomycin C, hydrogen
peroxide, desiccation and heavy metals/metalloids AsO4 3− ,
Pb2+ , CrO4 2− and Ag1+ . The peptidoglycan in the cell
wall contains meso-diaminopimelic acid as diamino acid,
with galactose as the diagnostic sugar. The predominant
menaquinone is MK-9(H4 ). The main polar lipids are phosphatidylethanolamine, phosphatidylcholine, phosphatidylinositol, and small amount of diphosphatidylglycerol. Cellular
fatty acids consist mainly of iso-C16:0 , iso-C15:0 , C17:1 𝜔8c, and
C16:1 𝜔7c. The type strain has a genomic DNA G + C content of
74.4 mol %. The INSDC accession number for the 16S rRNA
gene sequences of the type strain G18T (= DSM 44209T =
CCUG 63018T ) is HF970583.
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
MIC (mM) of
NiCl2
K2 CrO4
0.5
4.0
0.3
1.0
Pb(NO3 )2
4.0
1.0
NaHAsO4
8.0
80.0
Acknowledgments
The authors would like to acknowledge the help of Bettina
Sträubler and Birgit Grün for DNA-DNA hybridization analyses, Gabi Pötter for assistance with chemotaxonomy, Brian
J. Tindall (all at DSMZ, Germany) for his guidance in the
chemotaxonomic analyses, and Haitham Sghaier (CNSTN,
Tunisia) for providing access to the gamma irradiation
facility. Maria del Carmen Montero-Calasanz is the recipient
of a postdoctoral contract from the European Social Fund
Operational Programme (2007–2013) for Andalusia and also
recipient of a DSMZ postdoctoral fellowship (2013–2015).
References
[1] P. Normand, S. Orso, B. Cournoyer et al., “Molecular phylogeny
of the genus Frankia and related genera and emendation of
the family Frankiaceae,” International Journal of Systematic
Bacteriology, vol. 46, no. 1, pp. 1–9, 1996.
[2] P. Normand, “Geodermatophilaceae fam. nov., a formal description,” International Journal of Systematic and Evolutionary
Microbiology, vol. 56, part 10, pp. 2277–2278, 2006.
[3] G. M. Luedemann, “Geodermatophilus, a new genus of the
Dermatophilaceae (Actinomycetales),” Journal of Bacteriology,
vol. 96, no. 5, pp. 1848–1858, 1968.
[4] V. B. D. Skerman, V. McGowan, and P. H. A. Sneath, “Approved
lists of bacterial names,” International Journal of Systematic
Bacteriology, vol. 30, no. 1, pp. 225–420, 1980.
[5] M. D. Montero-Calasanz, M. Göker, G. Pötter et al., “Geodermatophilus africanus sp. nov., a halotolerant actinomycete
isolated from Saharan desert sand,” Antonie van Leeuwenhoek,
vol. 104, no. 2, pp. 207–216, 2013.
[6] C. Urzı̀, L. Brusetti, P. Salamone, C. Sorlini, E. Stackebrandt, and
D. Daffonchio, “Biodiversity of Geodermatophilaceae isolated
from altered stones and monuments in the Mediterranean
basin,” Environmental Microbiology, vol. 3, no. 7, pp. 471–479,
2001.
[7] M. Eppard, W. E. Krumbein, C. Koch, E. Rhiel, J. T. Staley, and
E. Stackebrandt, “Morphological, physiological, and molecular
characterization of actinomycetes isolated from dry soil, rocks,
and monument surfaces,” Archives of Microbiology, vol. 166, no.
1, pp. 12–22, 1996.
[8] A. A. Gorbushina, “Life on the rocks,” Environmental Microbiology, vol. 9, no. 7, pp. 1613–1631, 2007.
[9] Y. Q. Zhang, J. Chen, H. Y. Liu, Y. Q. Zhang, W. J. Li, and L.-Y. Yu,
“Geodermatophilus ruber sp. nov., isolated from rhizosphere
soil of a medicinal plant,” International Journal of Systematic and
Evolutionary Microbiology, vol. 61, part 1, pp. 190–193, 2011.
[10] L. Jin, H.-G. Lee, H.-S. Kim, C.-Y. Ahn, and H.-M. Oh,
“Geodermatophilus soli sp. nov. and Geodermatophilus terrae sp.
nov., two actinobacteria isolated from grass soil,” International
10
[11]
[12]
[13]
[14]
[15]
[16]
[17]
[18]
[19]
[20]
[21]
[22]
[23]
[24]
[25]
BioMed Research International
Journal of Systematic and Evolutionary Microbiology, vol. 63,
part 7, pp. 2625–2629, 2013.
M. Gtari, I. Essoussi, R. Maaoui et al., “Contrasted resistance of
stone-dwelling Geodermatophilaceae species to stresses known
to give rise to reactive oxygen species,” FEMS Microbiology
Ecology, vol. 80, no. 3, pp. 566–577, 2012.
V. Mattimore and J. R. Battista, “Radioresistance of Deinococcus
radiodurans: functions necessary to survive ionizing radiation
are also necessary to survive prolonged desiccation,” Journal of
Bacteriology, vol. 178, no. 3, pp. 633–637, 1996.
K. W. Minton, “DNA repair in the extremely radioresistant
bacterium Deinococcus radiodurans,” Molecular Microbiology,
vol. 13, no. 1, pp. 9–15, 1994.
K. W. Minton, “Repair of ionizing-radiation damage in the radiation resistant bacterium Deinococcus radiodurans,” Mutation
Research: DNA Repair, vol. 363, no. 1, pp. 1–7, 1996.
F. A. Rainey, K. Ray, M. Ferreira et al., “Extensive diversity of
ionizing-radiation-resistant bacteria recovered from Sonoran
Desert soil and description of nine new species of the genus
Deinococcus obtained from a single soil sample,” Applied Environmmental Microbiology, vol. 71, no. 9, pp. 5225–5235, 2005.
A. Giongo, J. Favet, A. Lapanje et al., “Microbial hitchhikers on
intercontinental dust: high-throughput sequencing to catalogue
microbes in small sand samples,” Aerobiologia, vol. 29, no. 1, pp.
71–84, 2013.
G. X. Nie, H. Ming, S. Li et al., “Geodermatophilus nigrescens sp.
nov., isolated from a dry-hot valley,” Antonie van Leeuwenhoek,
vol. 101, no. 4, pp. 811–817, 2012.
M. C. Montero-Calasanz, M. Göker, G. Pötter et al., “Geodermatophilus arenarius sp. nov., a xerophilic actinomycete isolated
from Saharan desert sand in Chad,” Extremophiles, vol. 16, no.
6, pp. 903–909, 2012.
M. del Carmen Montero-Calasanz, M. Göker, M. Rohde et al.,
“Geodermatophilus siccatus sp. nov., isolated from arid sand of
the Saharan desert in Chad,” Antonie van Leeuwenhoek, vol. 103,
no. 3, pp. 449–456, 2013.
J. Euzéby, “List of new names and new combinations previously
effectively, but not validly, published,” International Journal of
Systematic and Evolutionary Microbiology, vol. 63, part 5, pp.
1577–1580, 2013.
M. C. Montero-Calasanz, M. Göker, G. Pötter et al., “Geodermatophilus saharensis sp. nov., isolated from sand of the Saharan
desert in Chad,” Archives of Microbiology, vol. 195, no. 3, pp. 153–
159, 2013.
M. D. C. Montero-Calasanz, M. Göker, W. J. Broughton et al.,
“Geodermatophilus tzadiensis sp. nov., a UV radiation-resistant
bacterium isolated from sand of the Saharan desert,” Systematic
and Applied Microbiology, vol. 36, no. 3, pp. 177–182, 2013.
A. Oren and G. M. Garrity, “List of new names and new
combinations previously effectively, but not validly, published,”
International Journal of Systematic and Evolutionary Microbiology, vol. 63, pp. 3931–3934.
M. D. C. Montero-Calasanz, M. Göker, G. Pötter et al., “Geodermatophilus telluris sp. nov., an actinomycete isolated from
Saharan desert sand,” International Journal of Systematic and
Evolutionary Microbiology, vol. 63, part 6, pp. 2254–2259, 2013.
M. C. Montero-Calasanz, M. Göker, G. Pötter et al., “Geodermatophilus normandii sp. nov., isolated from Saharan desert
sand,” International Journal of Systematic and Evolutionary
Microbiology, vol. 63, part 9, pp. 3437–3443, 2013.
[26] J.-H. Qu, M. Hui, J.-Y. Qu et al., “Geodermatophilus taihuensis
sp. nov., isolated from t he interfacial sediment of a eutrophic
lake,” International Journal of Systematic and Evolutionary
Microbiology, vol. 63, pp. 4108–4112, 2013.
[27] M. C. Montero-Calasanz, M. Göker, M. Rohde et al., “Description of Geodermatophilus amargosae sp. nov., to accommodate
the not validly named Geodermatophilus obscurus subsp. amargosae (Luedemann,1968),” Current Microbiology, vol. 68, no. 3,
pp. 365–371, 2014.
[28] A. Oren and G. M. Garrity, “Validation List no. 158. List of
new names and new combinations previously effectively, but
not validly, published,” International Journal of Systematic and
Evolutionary Microbiology, vol. 64, part 5, pp. 1455–1458, 2014.
[29] N. Ivanova, J. Sikorski, M. Jando et al., “Complete genome
sequence of Geodermatophilus obscurus type strain (g-20 T),”
Standards in Genomic Sciences, vol. 2, no. 2, pp. 158–167, 2010.
[30] E. E. Ishiguro and D. W. Fletcher, “Characterization of Geodermatophilus strains isolated from high altitude Mount Everest
soils,” Mikrobiologika, vol. 12, pp. 99–108, 1975.
[31] P. Normand and D. R. Benson, “Genus I. Geodermatophilus
Luedemann 1968. 1994,” in Bergey's Manual of Systematic
Bacteriology: The Actinobacteria, vol. 5, part 1, pp. 528–530,
Springer, New York, NY, USA, 2nd edition, 2012.
[32] M. J. Pelczar Jr., Manual of Microbiological Methods, McGrawHill, New York, NY, USA, 1957.
[33] T. Gregersen, “Rapid method for distinction of gram negative
from gram positive bacteria,” European Journal of Applied
Microbiology, vol. 5, no. 2, pp. 123–127, 1978.
[34] E. B. Shirling and D. Gottlieb, “Methods for characterization
of Streptomyces species,” International Journal of Systematic
Bacteriology, vol. 16, pp. 313–340, 1966.
[35] L. A. I. Vaas, J. Sikorski, V. Michael, M. Göker, and H.-P. Klenk,
“Visualization and curve-parameter estimation strategies for
efficient exploration of phenotype microarray kinetics,” PLoS
ONE, vol. 7, no. 4, Article ID e34846, 2012.
[36] L. A. I. Vaas, J. Sikorski, B. Hofner et al., “Opm: an R package
for analysing OmniLog (R) phenotype microarray data,” Bioinformatics, vol. 29, no. 14, pp. 1823–1824, 2013.
[37] R. Suzuki and H. Shimodaira, “pvclust: hierarchical clustering with p-values via multiscale bootstrap resampling,” R
package version 01.2-2, http://cran.r-project.org/web/packages/
pvclust/index.html.
[38] M. P. Lechevalier and H. A. Lechevalier, “Chemical composition
as a criterion in the classification of aerobic actinomycetes,”
International Journal of Systematic Bectoriology, vol. 20, pp. 435–
443, 1970.
[39] J. L. Staneck and G. D. Roberts, “Simplified approach to identification of aerobic actinomycetes by thin layer chromatography,”
Journal of Applied Microbiology, vol. 28, no. 2, pp. 226–231, 1974.
[40] D. E. Minnikin, A. G. O’Donnell, M. Goodfellow et al., “An
integrated procedure for the extraction of bacterial isoprenoid
quinones and polar lipids,” Journal of Microbiological Methods,
vol. 2, no. 5, pp. 233–241, 1984.
[41] R. M. Kroppenstedt and M. Goodfellow, “The family Thermomonosporaceae: Actinocorallia, Actinomadura, Spirillispora
and Thermomonospora,” in The Prokaryotes, vol. 3 of Archaea
and Bacteria: Firmicutes, Actinomycetes, pp. 682–724, Springer,
New York, NY, USA, 3rd edition, 2006.
[42] B. J. Tindall, “A comparative study of the lipid composition of
Halobacterium saccharovorum from various sources,” Systematic and Applied Microbiology, vol. 13, no. 2, pp. 128–130, 1990.
BioMed Research International
[43] M. D. Collins, T. Pirouz, M. Goodfellow, and D. E. Minnikin, “Distribution of menaquinones in actinomycetes and
corynebacteria,” Journal of General Microbiology, vol. 100, no.
2, pp. 221–230, 1977.
[44] R. M. Kroppenstedt, “Separation of bacterial menaquinones
by HPLC using reverse phase (RP18) and a silver loaded ion
exchanger,” Journal of Liquid Chromatography, vol. 5, no. 12, pp.
2359–2387, 1982.
[45] M. Sasser, “Identification of bacteria by gas chromatography of
cellular fatty acids,” USFCC Newsl, vol. 20, p. 16, 1990.
[46] K. H. Schleifer and O. Kandler, “Peptidoglycan types of bacterial
cell walls and their taxonomic implications,” Bacteriological
Reviews, vol. 36, no. 4, pp. 407–477, 1972.
[47] M. Mesbah, U. Premachandran, and W. B. Whitman, “Precise
measurement of the G+C content of deoxyribonucleic acid
by high-performance liquid chromatography,” International
Journal of Systematic Bacteriology, vol. 39, no. 2, pp. 159–167,
1989.
[48] F. A. Rainey, N. Ward-Rainey, R. M. Kroppenstedt, and E.
Stackebrandt, “The genus Nocardiopsis represents a phylogenetically coherent taxon and a distinct actinomycete lineage:
proposal of Nocardiopsaceae fam. nov.,” International Journal of
Systematic Bacteriology, vol. 46, no. 4, pp. 1088–1092, 1996.
[49] J. P. Meier-Kolthoff, M. Göker, C. Spröer, and H. Klenk,
“When should a DDH experiment be mandatory in microbial
taxonomy?” Archives of Microbiology, vol. 195, no. 6, pp. 413–418,
2013.
[50] P. Cashion, M. A. Holder Franklin, J. McCully, and M. Franklin,
“A rapid method for the base ratio determination of bacterial
DNA,” Analytical Biochemistry, vol. 81, no. 2, pp. 461–466, 1977.
[51] J. de Ley, H. Cattoir, and A. Reynaerts, “The quantitative
measurement of DNA hybridization from renaturation rates,”
European Journal of Biochemistry, vol. 12, no. 1, pp. 133–142, 1970.
[52] V. A. R. Huss, H. Festl, and K. H. Schleifer, “Studies on the
spectrophotometric determination of DNA hybridization from
renaturation rates,” Systematic and Applied Microbiology, vol. 4,
no. 2, pp. 184–192, 1983.
[53] E. E. Ishiguro and R. S. Wolfe, “Induction of morphogenesis
in Geodermatophilus by inorganic cations and by organic
nitrogenous cations,” Journal of Bacteriology, vol. 117, no. 1, pp.
189–195, 1974.
[54] J. W. Richards, G. D. Krumholz, M. S. Chval, and L. S. Tisa,
“Heavy metal resistance patterns of Frankia strains,” Applied
and Environmental Microbiology, vol. 68, no. 2, pp. 923–927,
2002.
[55] P. McCullagh and J. A. Nelder, Generalized Linear Models,
Chapman & Hall, London, UK, 2nd edition, 1989.
[56] P. H. C. Eilers and B. D. Marx, “Flexible smoothing with 𝐵splines and penalties,” Statistical Science, vol. 11, no. 2, pp. 89–
121, 1996.
[57] S. N. Wood, Generalized Additive Models: An Introduction with
R, CRC press, 2006.
[58] R Core Team, A Language and Environment for Statistical
Computing, R Foundation for Statistical Computing, Vienna,
Austria, 2014, http://www.R-project.org/.
[59] B. Hofner, “lethal: Compute lethal doses (LD) with confidence
intervals,” R package, http://r-forge.r-project.org/projects/lethal/.
[60] L. G. Wayne, D. J. Brenner, R. R. Colwell et al., “Report of the
Ad Hoc committee on reconciliation of approaches to bacterial
systematics,” International Journal of Systematic Bacteriology,
vol. 37, no. 4, pp. 463–464, 1987.
11
[61] J. R. Battista, A. M. Earl, and M. J. Park, “Why is Deinococcus
radiodurans so resistant to ionizing radiation?” Trends in
Microbiology, vol. 7, no. 9, pp. 362–365, 1999.
[62] M. Shukla, R. Chaturvedi, D. Tamhane et al., “Multiplestress tolerance of ionizing radiation-resistant bacterial isolates
obtained from various habitats: correlation between stresses,”
Current Microbiology, vol. 54, no. 2, pp. 142–148, 2007.
[63] J. R. Battista, “Against all odds: the survival strategies of
Deinococcus radiodurans,” Annual Review of Microbiology, vol.
51, pp. 203–224, 1997.
[64] S. J. Stohs and D. Bagchi, “Review article oxidative mechanisms
in the toxicity of metals ions,” Free Radical Biology and Medicine,
vol. 18, no. 2, pp. 321–336, 1995.
[65] D. H. Nies, “The cobalt, zinc, and cadmium efflux system
CzcABC from Alcaligenes eutrophus functions as a cationproton antiporter in Escherichia coli,” Journal of Bacteriology,
vol. 177, no. 10, pp. 2707–2712, 1995.
[66] R. V. Fogarty and J. M. Tobin, “Fungal melanins and their
interactions with metals,” Enzyme and Microbial Technology,
vol. 19, no. 4, pp. 311–317, 1996.
[67] A. M. Spain, A. D. Peacock, J. D. Istok et al., “Identification
and isolation of a Castellaniella species important during
biostimulation of an acidic nitrate- and uranium-contaminated
aquifer,” Applied and Environmental Microbiology, vol. 73, no.
15, pp. 4892–4904, 2007.
[68] A. C. Poopal and R. S. Laxman, “Studies on biological reduction
of chromate by Streptomyces griseus,” Journal of Hazardous
Materials, vol. 169, no. 1–3, pp. 539–545, 2009.
[69] I. V. N. Rathnayake, M. Megharaj, G. S. R. Krishnamurti, N.
S. Bolan, and R. Naidu, “Heavy metal toxicity to bacteria: are
the existing growth media accurate enough to determine heavy
metal toxicity?” Chemosphere, vol. 90, no. 3, pp. 1195–1200, 2013.
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 480170, 9 pages
http://dx.doi.org/10.1155/2014/480170
Research Article
Safe-Site Effects on Rhizosphere Bacterial Communities in
a High-Altitude Alpine Environment
Sonia Ciccazzo,1 Alfonso Esposito,2 Eleonora Rolli,1 Stefan Zerbe,2
Daniele Daffonchio,1 and Lorenzo Brusetti2
1
2
Department of Food, Environmental and Nutritional Sciences (DeFENS), University of Milan, Via Celoria 2, 20133 Milan, Italy
Faculty of Science and Technology, Free University of Bozen-Bolzano, Piazza Università 5, 39100 Bolzano, Italy
Correspondence should be addressed to Lorenzo Brusetti; [email protected]
Received 2 April 2014; Accepted 14 May 2014; Published 4 June 2014
Academic Editor: George Tsiamis
Copyright © 2014 Sonia Ciccazzo et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
The rhizosphere effect on bacterial communities associated with three floristic communities (RW, FI, and M sites) which differed
for the developmental stages was studied in a high-altitude alpine ecosystem. RW site was an early developmental stage, FI was
an intermediate stage, M was a later more matured stage. The N and C contents in the soils confirmed a different developmental
stage with a kind of gradient from the unvegetated bare soil (BS) site through RW, FI up to M site. The floristic communities were
composed of 21 pioneer plants belonging to 14 species. Automated ribosomal intergenic spacer analysis showed different bacterial
genetic structures per each floristic consortium which differed also from the BS site. When plants of the same species occurred
within the same site, almost all their bacterial communities clustered together exhibiting a plant species effect. Unifrac significance
value (𝑃 < 0.05) on 16S rRNA gene diversity revealed significant differences (𝑃 < 0.05) between BS site and the vegetated sites
with a weak similarity to the RW site. The intermediate plant colonization stage FI did not differ significantly from the RW and
the M vegetated sites. These results pointed out the effect of different floristic communities rhizospheres on their soil bacterial
communities.
1. Introduction
A glacier foreland after glacier retreat can be considered
a cold desert, being composed of habitats characterized
by severe climatic regimes and barren substrate with low
total carbon and nitrogen contents [1]. Rock cracks, concave
surfaces, and little depressions could ensure protection from
wind, cold, and other harsh environmental conditions [2, 3]
helping the accumulation of nutrients and the growth of
pioneer plants. Safe-sites are defined as little areas, often
surrounded by big stones, filled up of stone debris or mineral
mud [4]. Here, opportunistic pioneer plants could settle
down and form first floristic consortia, significantly affected
by the geochemistry of the lytic material. Indeed, physical
and biogeochemical weathering processes provide soils of
soluble nutrients and when the plant colonization on parent
materials occurs, the development of glacier foreland into
fertile soils is enhanced by rhizodeposition, root exudation,
decaying biomass, and root mass development. Safe-sites
can be severely affected by geological dynamics, such as
sudden mudslides, alluvial fan sliding, and scree movement,
that take back the habitat to an earlier pioneer condition.
Consequently, safe-sites cannot reach the climax but only a
stable stage of middle maturity [5].
Furthermore, pioneer plants could select rhizosphere
microbial communities able to promote plant growth thanks
to the interactions in nutrient cycling and carbon sequestration [6]. Nevertheless, in a natural ecosystem it is difficult to
assess the effect of vegetation on the rhizosphere bacterial
communities, especially in high mountain environments
characterized by variable environmental parameters (successional stage, pH, rainfall, moisture, mineral composition,
sampling season, and slope) within a size-limited area typical
of early and transitional successional stages. The impact
of single plants on microbial communities in an alpine
glacier forefield has previously been studied to highlight
the relationship between the rhizosphere bacterial communities of pioneer plants or of the related bare soil and the
2
chronosequence [7–10]. In an early chronosequential stage,
the rhizosphere microbial community of Poa alpina L. was
strongly influenced by the environmental conditions but, in
transition and mature stage, plants could select a specific
microbial community [9]. Along a similar chronosequence,
the pioneer plant Leucanthemopsis alpina (L.) Heywood
showed an opposing rhizosphere effect with a specific microbial community in the early successional stage only [7]. The
study of the spatial extent of Lc. alpina on the microbial
community and on the physical-chemical parameters in an
early successional stage (5, 10 years) did not exhibit significant
trends, supporting the conclusion of Tscherko et al. [9].
However, in a safe-site, the pioneer vegetation interrelated
in floristic consortia often exhibited ground stems and root
tangle with large nets. In this case, a safe-site could be equaled
to a transitional or even a mature grassland for root tangle and
plant community structure. The floristic community effect in
such a habitat was observed in natural as well as in artificial
experimental sites [11]. Osanai et al. [12] demonstrated that
cooccurring plant species from native grassland selected their
microbial communities. The effect was generally smaller than
for species that generally do not cooccur naturally, such as
those from agricultural crop systems [13], improved grassland
systems, or fertilized grassland fields [14, 15]. Nunan et al.
[16] found a weak influence of plant community or no
effect of plant species on the structure and diversity of the
root-colonizing bacterial community when comparing five
cooccurring grass species from an upland grazed grassland in
Scotland. Moreover, topography and other uncharacterized
environmental factors seemed to be main drivers of the
bacterial community composition.
On the other hand, studies about the effect of plant cover
on microbial community in cold environments regarded
different ecological niches and pointed out the higher significance of environmental parameters than the influence
of the floristic consortia. In Antarctic environments along
a latitudinal gradient, bacterial diversity of dense vegetation
from different locations was comparable whereas bacterial
diversity of “fell-field” vegetation decreased with increasing
latitude [17, 18]. In permafrost meadow, steppe, or desert
steppe, soil characteristics were driving factors of the microbial diversity [19]. In high elevation arid grassland, a strong
plant effect was demonstrated for the perennial bunchgrasses
Stipa, Hilaria, and for the invading annual grass Bromus [20].
Consequently, the aim of this work was to investigate if,
in different safe-sites on a deglaciated terrain of the same
chronosequential age, floristic consortia could select specific
rhizobacterial communities.
2. Materials and Methods
2.1. Study Site and Soil Samples. The study site is located in
the upstream subcatchment of Saldur river (46∘ 46󸀠 30󸀠󸀠 N; 10∘
41󸀠 46󸀠󸀠 E; 2,400 a.s.l.) in the high Matsch valley (South Tyrol,
Italy) with a drainage area of 11 km2 . The main geological
processes are periglacial and the streamflow is characterized
by the glacier dynamics. During 1970–2000, the valley had
an average rainfall of about 550 mm per year. In 2011, the
mean temperatures of the plant growing season were 7.3∘ C
BioMed Research International
in July, 10.3∘ C in August, and 8∘ C in September and the mean
precipitations were 2.7, 2.5, and 3.6 mm per day, respectively.
The dominant rock types are schist and gneiss [21] and the
most common soil types are acidic leptosols, regosols, and
umbrisols (mean pH = 4.3) derived from carbonate-free
bedrocks. The study site, a foreland of about 3.3 Km2 left after
a quick glacier retreat in the last 160 years [22], was located
above the tree line (2,100 m a.s.l). The analysis of the historical
maps of the third Austro-Hungarian topographic survey
(the so-called “Franzisco-Josephinische Landesaufnahme”)
dated 1850 and the aerial photographs of 1945 and of 2006
orthophotos were helpful to reconstruct the different stages
of glacier retreat. Thus, comparing these photos, our sampling
site was ice-free since 1850.
Rhizosphere and soil sampling were carried out in 2011
May, at the beginning of the plant growing season. Three safesites (RW, FI, and M sites) characterized by loosely organized
assemblages of different plant species and a bare soil (BS site)
were sampled. The sites were less than 20 × 20 cm. RW site,
below an iron rich rock-face, was colonized by Diphasiastrum
alpinum and Gnaphalium supinum L.; FI site, a floristic island
between big rocks, was colonized by Cladonia sp., Festuca
halleri All., Polytrichum sp., Racomitrium sp., Sedum alpestre
Vill., and Senecio carniolicus (Willd.) Braun-Blanq.; M site,
a safe-site surrounded by big rocks and characterized by a
flatter area, was colonized by Cetraria islandica (L.) Ach.,
Leucanthemopsis alpina (L.) Heywood, Potentilla aurea L.,
Rhododendron ferrugineum, Sibbaldia procumbens L., and
Silene acaulis (L.) Jacq. These sampling sites were carefully
chosen in order to share similar conditions in terms of
altitude, features, and geology.
The rhizosphere samples of all the single plant individuals
within a floristic community were collected. Each individual
plant was carefully pulled out the soil, without damaging its
single root system. After pulling out each plant and avoiding
roots, 4 g of rhizosphere soil strictly adhering to the roots
was collected with a pair of sterile tweezers. Three replicates
of bulk soil were collected as a control. Moreover, from
each safe-site, 50 g of root-free soil was collected and put
into plastic bags for soil chemical analysis. All the samples
were immediately transported in refrigerated boxes to the
laboratory as soon as the logistic constraints permitted and
they were stored at −80∘ C until analysis.
2.2. Soil Chemical Analysis. Soil samples for chemical analysis were oven-dried at 105∘ C until constant weight and
then acid was digested (HNO3 concentrated 65% and H2 O2
30%) in a milestone high performance microwave oven (MLS
Mega, Gemini BV Laboratory, Apeldoorn, The Netherlands).
To determine the total organic carbon content, soil samples
were also acidified with HCl (6 M) to eliminate all carbonates.
Metals and total phosphorous were determined by inductively coupled plasma-optical emission spectroscopy (ICPOES, Spectro Ciros CCD, Spectro GmbH, Kleve, Germany).
Nitrogen and C were quantified with an elemental analyzer
(Flash 2000, Thermo Scientific). The pHH2 O was measured
using an Accumet AP85 pH (Fisher Scientific Ltd., Pittsburgh, PA, USA). To test the level of significance of the
observed chemical differences among sites, a Kruskal-Wallis
BioMed Research International
3
test was done by using Mann-Whitney pairwise comparison
post hoc test and Bonferroni correction in Past software [23].
Table 1: Percentage of total nitrogen and carbon content and C/N
ratio in the four safe-sites.
2.3. Molecular Analysis of the Bacterial Communities. Total
DNA of the rhizosphere and soil samples was extracted
using Ultraclean Soil DNA Extraction kit (MO-BIO, Arcore,
Italy). Microbial analyses were carried out using denaturing
gradient gel electrophoresis (DGGE) [24] to describe the
rhizobacterial taxa diversity and automated ribosomal intergenic spacer analysis (ARISA) [25] to describe the structure
of the rhizobacterial communities.
For DGGE analysis, primers GC357f and 907r were used
as described [26]. DGGE was run in a BioRad DCode
universal mutation detection system (Bio-Rad, Milan, Italy).
Polyacrylamide gels were done according to Muyzer et al.
[24]. Gels were stained for 30 min in 1x TAE buffer containing SYBR Safe-DNA gel stain (Invitrogen, Milan, Italy).
Visualization and digital image recording were performed
with UVTec (Cambridge, UK). All the visible DGGE bands
were excised and reamplified [24]. Sequencing was performed by STAB-Vida Inc. (Caparica, Portugal). Identification of 16S rRNA genes was done by comparison with
EMBL/Genebank/DDBJ database and RDP database using
BLASTN and Classifier, respectively. All sequences were
submitted to the Ribosomal Database Project (RDP) web
server [27] to assign taxonomy. Sequences were submitted
to the Genbank/EMBL/DDBJ databanks under the accession
numbers HG763876-HG764130.
ARISA fingerprint was performed as described by Cardinale et al. [25] with the ITSF/ITSREub primer set. Denatured
ARISA fragments were run by STAB-Vida Inc. The data
were analyzed with Peak Scanner software v1.0 (Applied
Biosystems, Monza, Italy) and a threshold of 40 fluorescent
units was used, corresponding to two times the highest peak
detected during the negative control run. Output matrix was
obtained as in Rees et al. [28].
Safe-site
2.4. Statistical Analysis of ARISA and DGGE. ARISA matrix
was normalized with the formula (x/∑x)∗1000, where “x”
is the fragment height in units of fluorescence, and then
transformed on a logarithmic scale for multivariate analysis.
Log-transformation was used to stabilize the sample variance,
to reduce the interaction effect, and to normalize the distribution of data. Moreover, log-transformation can combine
the information of a binary matrix with those of a nontransformed data matrix, hence preserving the relative abundance
information and down-weighting dominant groups.
In order to assess changes in rhizobacterial community
structure between floristic consortia, nonmetric multidimensional scaling (NMDS) was applied with Bray-Curtis algorithm. NMDS does not need the assumption of linear associations among variables being described as the most efficient
ordination method for microbial ecology [29]. Bray-Curtis
is not influenced by recurrent absent values into the matrix,
a characteristic very common in ARISA matrices [30].
ANOSIM (based on Bray-Curtis similarity) was performed
to test significant differences in the profile composition of
the different sites. ANOSIM is a nonparametric statistical
test, based on permutation, which uses rank similarity matrix
BS
RW
FI
M
Nitrogen %
Carbon %
C/N
Average St. dev. Average St. dev. Average St. dev.
0.05
0.01
0.62
0.16
11.5
0.61
0.27
0.11
3.48
1.47
12.7
0.94
0.72
0.35
10.4
6.03
14.2
1.46
0.98
0.85
19.3
18.3
17.5
3.79
of an ordination plot to calculate an 𝑅 test statistic on the
null hypothesis 𝐻0 that there are no differences among
groups. When 𝑅 is near to 0, 𝐻0 is true, while when 𝑅 is
reaching 1, 𝐻0 can be rejected and there is a discrimination
between groups. When ANOSIM statistics approaches 1,
the similarities within groups are larger than the similarity
between groups. We rejected 𝐻0 when significance 𝑃 value
was <0.05. To test the level of significance between/within
plant species ARISA clusters, a Kruskal-Wallis test was done
as above.
The Nexus format of the phylogenetic tree of the DGGE
identified bands performed by MEGA5 was submitted to
the UniFrac web server to test differences among sites
based in the UniFrac metric with 100 permutations and the
Bonferroni correction factor [31]. A principal coordinates
analysis (PCoA) on the DGGE sequence distance matrix
for each pair of safe-sites was calculated through UniFrac
metric. On the basis of the DGGE sequences, similar safesites tended to cluster together. In order to allow a broader
view of those similarities, the first three principal components
were considered.
3. Results
3.1. Soil Chemical Analysis. Soil resulted to be a sandy silt soil
with an average texture of 72.3 ± 5.0% of sand, 21.0 ± 4.1%
of silt, 6.6 ± 1.3% of clay, and 4.6 ± 1.3% of humus; pH was
4.5 ± 0.3%. Average chemical composition of sampled soils
was total P 0.7 ± 0.1 mg/kg d.m., total K 7.4 ± 1.0 mg/kg
d.m., total Ca 3.4 ± 0.6 mg/kg d.m., total Mg 13.4 ± 1.7 mg/kg
d.m., total Fe 45.4 ± 6.9 mg/kg d.m., and total Al 29.4 ±
5.6 mg/kg d.m. No calcium carbonate was detected. Since
those safe-sites were located in proximity of each other,
their soil chemical composition did not differ substantially
between sites (Kruskal-Wallis test 𝑃 < 0.05; data not shown).
No nitrate was detected, while all the nitrogen found was
represented by ammonia only. Nitrogen increased along an
ideal gradient from bare soil (0.05% dry weight) to the most
vegetated M site (0.98% dry weight) and also total organic
carbon grew up from BS site (0.62% dry weight) to M site
(19.3% dry weight; Table 1). The trend was confirmed by the
C/N ratio which tended to increase constantly among sites
of more complex vegetative patterns. Bonferroni-corrected
Kruskal-Wallis nonparametric analysis of variance showed
significant differences among sites for both total nitrogen,
organic carbon content and C/N ratio, except for C and N
content between RW versus FI and RW versus M (𝑃 values
4
BioMed Research International
Table 2: Level of significance (𝑃 values) of the differences in C,
N, and C/N content among sites by Bonferroni-corrected KruskalWallis test.
BS versus RW
BS versus FI
BS versus M
RW versus FI
RW versus M
FI versus M
C
0.023
0.028
0.019
0.175
0.772
0.004
N
0.023
0.028
0.019
0.197
0.954
0.004
C/N
0.023
0.043
0.032
0.012
0.023
0.045
shown in Table 2) explained by a higher standard deviation
of C and N content in M sites.
3.2. Genetic Structure of Bacterial Communities in Alpine
Bulk Soils and Plant Colonized Safe-Sites. Due to the high
sensitivity of the automated capillary electrophoresis, ARISA
fingerprints of both rhizosphere and bare soil bacterial communities provided complex profiles with peaks ranging from
151 bp to 1437 bp and the 16S-23S rRNA internal transcribed
spacer region (ITS) richness varied from 43 to 168 peaks.
The electropherograms, characterized by distinct peaks number and intensity, revealed a shift in bacterial community
structure across the different safe-sites plant communities.
On the NMDS plot (stress value = 0.18), samples from rootfree soil (BS), safe-site of early developmental stage (RW),
intermediate stage (FI), and from the most mature one (M)
showed four separate clusters based on microbial community
structure (Figure 1). According to axis 1, RW site and BS site
are separated from M and FI sites. According to axis 2, BS and
M sites are separated by RW and FI sites. The unvegetated
BS site clustered in a specific group, differentiated by the
plant rhizospheres, is clustering closer to the rhizosphere
bacterial communities of RW site than to those of FI and
M sites. The NMDS separation is partially explained by N
and C content, as shown by those variable vectors, which
influenced more the M site than the other safe-sites. ANOSIM
analysis confirmed a highly significant difference among the
four microbial community structures (𝑅 = 0.81; 𝑃 = 0.0001)
and the performed test showed significant differences in the
pairwise comparisons of the sites with 𝑅 values approaching 1
in most of the cases (Table 3). Where replicated individuals of
the same plant species within each safe-site were found, it was
possible to denote a plant species effect. This is recognizable
within RW safe-site, where individuals from D. alpinum and
G. supinum formed two clusters significantly different along
the first axis of NMDS (𝑃 = 0.032 at the Kruskal-Wallis
test). In FI and M sites the tendency of individuals of the
same species to cluster together seems to disappear, except for
R. ferrugineum, maybe due to the higher number of species
interconnected in the safe-site.
3.3. Diversity of the Bacterial Communities Associated with
Alpine Bulk Soils and Pioneer Plants in Safe-Sites. DGGE
was performed to investigate the different microenvironments of the three safe-sites and bulk soil in terms of their
dominant bacterial population composition. A total of 255
Table 3: 𝑃 and 𝑅 values of ANOSIM based on Bray-Curtis similarity
of the four safe-sites as grouped after ARISA-NMDS plot analysis.
P/R value
BS
RW
FI
M
BS
0.0124
0.0077
0.0092
RW
0.9630
0.0005
0.0009
FI
0.9758
0.9390
M
0.7937
0.7434
0.7055
0.0004
sequences of more than 300 bp were obtained from all sample
profiles. RDP facilitated the determination of putative taxonomic affiliation of the recovered sequences. Major bacterial
taxa included Acidobacteria Gp3 and Gp1, Sphingobacteria,
Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria, and Actinobacteria (Figure 2). A noteworthy amount of
uncultured bacteria was found. Shifts in bacterial communities were visible. Members of the Acidobacteria order were
present in all the sites samples. They generally represented
the most abundant taxon, although a decrease of their relative
abundance is visible with percentage from BS site (57%) to
M site (33%). Proteobacteria were not found in BS site, while
they were scarcely present in RW and FI site rhizospheres
(4%, 8%, resp.). In M site Proteobacteria became more abundant than Acidobacteria (35%). In particular, the increasing
abundance of Proteobacteria was due to Alphaproteobacteria,
being more represented than Gammaproteobacteria and
Betaproteobacteria. A considerable amount of unclassified
Proteobacteria was also evident in M site. Sphingobacteria
were recovered with low percentages in RW, FI, and M sites
rhizospheres whereas members of Actinobacteria taxa were
even less abundant being present in FI and M sites rhizospheres only. We did not find Sphingobacteria or Actinobacteria taxa associated with BS samples. According to RDP
classification, unclassified Acidobacteria or Proteobacteria, as
well as other uncharacterized bacteria, were quite common
within all sites. For example, RW site was almost completely
colonized by unclassified Acidobacteria and unknown Bacteria, except few sequences affiliated to uncultured Burkholderia or to a Chitinophagaceae bacterium. Similarly, FI safe-site
was mostly colonized by unclassified Acidobacteria, although
more frequent sequences belonging to Bradyrhizobiaceae,
Chitinophagaceae, and other rarer taxa such as Flavisolibacter
sp. or Granulicella sp. were found. Finally, M site, the most
differentiated safe-site, counted the presence of unknown
Bradyrhizobiaceae, Bradyrhizobium sp., and uncultured Rhizobiales, as well as Chitinophagaceae, Streptacidiphilus sp.,
Thermomonosporaceae, and Xanthomonadaceae.
Despite bias associated with sampling, DNA extraction,
PCR amplification, and DGGE run, the pattern of differences in bacterial communities composition between the
unvegetated soils of the BS site and the rhizospheres of
the three safe-sites was supported by the pairwise UniFrac
distance ordinations. Comparing each pair of environments
using the Bonferroni correction, the UniFrac permutation
test significance (𝑃 values < 0.05) showed that the BS site
samples were significantly different from FI and M sites
rhizospheres, but not from RW site rhizospheres. Moreover,
the FI site rhizosphere did not differ significantly from M
BioMed Research International
5
0.30
M P aur 40
0.24
0.18
M S aca 37
M R fer 16
BS 13
M S pro 35 %C
%N
M L alp 39 0.12
M R fer 34
M C isl 36
0.06
BS 11
BS 12
−0.2
−0.3
FI S car 64
FI Pol sp 66
FI F hal 65
FI Clad sp 69
FI Pol sp 68
−0.06
FI Rac sp 62
FI S car 67 −0.12
FI S alp 61
−0.18
0.1
RW D alp 21
0.2
0.3
0.4
RW D alp 23
RW G sup 19
RW D alp 22
RW G sup 20
RW G sup 18
Figure 1: NMDS plot of the three safe-sites and the bare soil site according to UniFrac distance matrix. BS site was a root-free safe-site,
RW site was an early developmental floristic stage, FI site was an intermediate stage, and M site was a later stage. Plant sample names are
the following: C isl—Cetraria islandica (L.) Ach.; Clad sp—Cladonia sp.; D alp—Diphasiastrum alpinum; F hal—Festuca halleri All.; Gsup—Gnaphalium supinum L.; L alp—Leucanthemopsis alpina (L.) Heywood; Pol sp—Polytrichum sp.; P aur—Potentilla aurea L.; Rac sp—
Racomitrium sp.; R fer—Rhododendron ferrugineum; S alp—Sedum alpestre Vill.; S car—Senecio carniolicus (Willd.) Braun-Blanq.; S pro—
Sibbaldia procumbens L.; S aca—Silene acaulis (L.) Jacq.
100
and RW sites rhizospheres, while the M site rhizospheres
exhibited significant differences with the RW site. A PCoA
analysis of the UniFrac distance matrix was calculated to
assess the overall sequence population similarity among safesites (Figure 3). The first axis of PCoA analysis, explaining
45.6% of the total variance, showed a shift of the BS site
from RW, FI, and M sites. The FI and M communities were
located very close together in the same quadrant suggesting
a similar bacterial community composition influenced by
variables related to PC1. On the other hand, PC2 (32.6% of
the variation) explained the differences between RW site and
the other three sites. Finally, the third component (21.8% of
the variance) differentiated FI from M and from BS and RW
sites.
90
80
70
(%)
60
50
40
30
20
10
0
4. Discussion
BS
RW
Unc. bacteria
Actinobacteria
Unc. Proteobacteria
Betaproteobacteria
Gammaproteobacteria
FI
M
Alphaproteobacteria
Sphingobacteria
Acidobacteria Gp1
Acidobacteria Gp3
Figure 2: Percentage abundance of each taxonomic group for each
individual rhizobacterial communities of the three safe-sites (RW,
FI, and M) and the bare soil site (BS) after 16S rRNA gene DGGEPCR analysis and band sequencing.
Safe-sites are defined as environments immediately nearby a
pool of seeds, where their germination, growth, and establishment are favorable [4]. In this respect, their availability,
accessibility, and geomorphological diversity in high mountain represent important characteristics of this environment,
since they represent a microsite where a list of ecological
hazards (snow, wind, frost, and irradiation) are less severe
than in open terrains and where plant propagules can resist,
grow, and reproduce. In Matsch valley, belonging to south
Tyrolean Alps, additional ecological hazards are represented
6
BioMed Research International
0.4
0.4
0.3
0.3
M
0.2
0.2
0.0
−0.1
−0.1
−0.2
−0.3
−0.3
−0.4
−0.4
−0.4
−0.2
FI
0.0
−0.2
RW
0.0
BS
0.1
FI
PC2 (32.6%)
PC2 (32.6%)
0.1 BS
−0.5
−0.6
M
RW
0.2
PC1 (45.6%)
(a)
−0.5
−0.3
−0.2
−0.1
0.0
0.1
0.2
0.3
0.4
PC3 (21.8%)
(b)
Figure 3: Principal coordinates analysis of the UniFrac distance matrix calculated to assess the overall sequence population similarity among
safe-sites. Percentage of variance of the single principal coordinates axis is indicated.
by hot and dry summers, instability of the soil substrate,
and excessive animal grazing [32]. Within each safe-site,
more than one plant species can grow from seeds, specialized
vegetative propagules, or plant fragments [33]. In such kind
of environments, pioneer plants tend to grow in very complex
coenosis, where roots are strictly intermixed and interrelated.
A great diversity of root exudates from all these plants is
released in rhizosphere, increasing the carbon amount of
the safe-site. Due to the characteristics of safe-sites, usually
well isolated among each other by rocks, sand, or mud,
an analysis to understand the occurrence of a vegetation
effect on rhizobacterial communities cannot be done with
traditional squared-plots, where more safe-sites are sampled
smoothing possible differences between them. Hence, we
decided to study three kinds of safe-sites at different stages
of morphological development, by sampling each single
rhizosphere from all the growing plant individuals.
The vegetation complexity of the three safe-sites (RW,
FI, and M) raised from a simple colonization of two species
(RW site) to the colonization of lichens, mosses, and few
herbaceous plant species (FI site) till M site, where five
herbaceous species and one woody species (R. ferrugineum)
were found. We discovered a distinct clustering of bacterial
communities according to RW, FI, and M vegetation types
that are significantly diverse from the unvegetated soil (BS
site). We also found that a gradient in terms of C and N
enrichments from BS site to the most developed M site was
an important determinant of microbial community profiles.
UniFrac analysis showed site-shifts in bacterial diversity
which suggest a specialized physiology adapted to the peculiar site environmental conditions. Moreover, the differences
among safe-sites, according to C and N gradients, support the
occurrence of a plant cover effect on the rhizosphere bacterial
community within those safe-sites.
Previous investigations of the rhizosphere effect were
conducted on few single pioneer plants or in grassland plots.
Almost all the researches on the rhizosphere effect associated
with a single plant species were achieved on crop or other
plants either in artificial microcosms such as pots or on
agricultural soils such as orchards and crop monocultures.
Most of these researches demonstrated that peculiar root
exudation and rhizodeposition of different plant species
could select the structural and functional diversity of the
associated rhizosphere bacterial communities [34–36]. On
the other hand, a consistent number of studies have showed
that several environmental parameters, that is, soil type,
soil characteristics, growth stage, management practices,
and growing season may influence the composition of the
microbial communities in the rhizosphere [37–44]. Past
studies about a natural alpine ecosystem investigated single
plant species along successional chronosequences and found
inconsistent effects of pioneer plants on rhizosphere microbial communities. For example, while the rhizobacterial
community of Lc. alpina was different from the interspace
community in an early successional chronosequential stage,
in a later stage it became similar to the interspace community.
In this case, it seemed that the influence of Lc. alpina
depended on soil age and that nutrient availability could
influence the bacterial community structure [7]. In another
study case, Lc. alpina individuals in the early successional
stage (5, 10 years) of a glacier forefield showed no selective
effect on the microbial community, since a similar bacterial
community structure was apparent up to 40 cm of distance
to the plant [8]. Another single pioneer plant, P. alpina,
BioMed Research International
did not exhibit a selective role on its rhizosphere bacterial
community in the pioneer stage of a chronosequence, maybe
due to the harsh environmental conditions of the plot where
it was growing [9]. However, by investigating a more mature
soil, the same plant species could select a specific microbial
community but related to soil properties and carbon supply.
On the other hand, safe-sites are more complex than
single pioneer plant individuals in a cold environment,
but they show less complexity than a homogeneous plot
carefully designed in mountain grasslands. Real safe-sites
are much less homogeneous, being shaped by the history of
the microarea where they are such as dynamical differences
in climate, in geophysical features, or in biota colonization
which determine complicated patterns and often unique rates
of soil development [1]. In our case, due to the quick glacier
melting in the last 80 years, the 160-year soil represents the
only transitional step of the glacier moraine between earliest
stages (<10 years) and mature soil (>500 years). As shown
by aerial photos, orthophotos, and a topographic survey, one
of the glacier tongues of the Weisskugel glacier has been
retreating with a discontinuous movement. Consequently,
there was no constant gradient of soil age but distinct block
stages where soil age is invariable. In this sense, the 160-yearold stage is more stable than an earlier successional soil and
it can host a larger number of plant species. Nevertheless it
was possible to distinguish hundreds of safe-sites of which
the three chosen were the most represented. Within the stable
block stage of 160 years old, the measured differences in
rhizobacterial composition and soil parameters supported
the hypothesis that the plant community composition of each
floristic consortium exhibited an effect on the rhizobacterial
communities widely documented in studies done in quite
different ecosystems. For example, Nunan et al. [16] demonstrated a more important influence of the plant community
composition than of the individual plant species on the
root colonizing bacterial community in an upland grazed
grassland, whereas Osanai et al. [12] showed a significant
impact of the plant species on the soil bacterial community
composition. Similar results were obtained comparing the
rhizosphere bacterial communities of three plant species of
an arid grassland [20].
The rhizosphere bacterial communities of RW site, characterized by only two different plant species, clustered more
closely with the BS site than with the vegetated ones showing
a simpler bacterial community, as confirmed by the UniFrac
analysis which detected no significant difference between the
two sites. Although FI and M sites had a similarity of about
56%, inside the FI site were found rhizobacterial communities
of mosses and lichens which did not cluster strictly with
the plant ones. The presence of lichens and mosses in the
same site could explain why the bacterial community of
the FI site represented an intermediate stage between the
RW site and the M site. The M site, colonized by individuals of six plant species, could be considered a later stage
where floristic consortia selected a more complex bacterial
community which significantly differs from the one of BS
and RW sites. The UniFrac analysis showed that the BS
communities were distinct from ones of the FI and M sites
and were weakly similar to the ones of RW site. Moreover,
7
the intermediate plant colonization stage, FI site, did not
differ significantly from the RW and the M vegetated sites.
Previous studies [9, 45, 46] showed that the development
of the soil microbial community in alpine glaciers was
determined by the accumulation of soil TOC and total
nitrogen. The increasing content of C and N in the floristic
consortia corresponded with increased floristic developmental stage. Soil nutrients and C influenced the bacterial
community composition along a chronosequence [7], while
in the Mendenhall glacier chronosequence [47] they were
not correlated with the rhizobacterial communities. These
different conclusions seem to strongly depend on the adopted
experimental design. Cultural-independent techniques based
on phospholipid fatty acid (PLFA) determination [9, 10], to
point out the different concentration of bacterial/fungal fatty
acids and to compare the Gram-positives/Gram-negatives
ratio, or molecular methods like restriction fragment length
polymorphism (RFLP) and DGGE analyses [7, 8] could not
have enough resolution to detect little changes in the bacterial
community genetic structure due to faint environmental
variables [48]. The ARISA analysis we used, targeting the
intergenic 16S-23S rRNA gene highly variable ITS region,
showed more sensitivity and enabled the detection up to
subspecies level, increasing the chance of the analysis to
detect very little effects on complex bacterial communities
[49].
5. Conclusions
Despite the harsh environmental condition of the natural
alpine ecosystem and the tight complex root system of
the safe-site, our results support the capability of different
pioneer plant consortia to select specific rhizobacterial communities with an increase of bacterial diversity according to
the increase of soil maturation. Moreover, when plants of
the same species occurred in the same site, the associated
rhizobacterial communities clustered more strictly together
according to their genetic structures, confirming the high
similarity of the rhizobacterial communities within individuals of the same pioneer plant species.
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
Acknowledgments
This research was financed by the Dr. Erich-Ritter and the
Dr. Herzog-Sellenberg Foundation within the Stifterverband
für die Deutsche Wissenschaft, Project “EMERGE: retreating
glaciers and emerging ecosystems in the Southern Alps”
(CUP n. I41J11000490007). Partial funds came from the Free
University of Bozen/Bolzano internal funds TN5026 “Effects
of climate change on high-altitude ecosystems” (CUP n.
I41J10000960005). The authors would like to thank Elisa
Varolo for plant species identification.
8
References
[1] J. A. Matthews, The Ecology of Recently-Deglaciated Terrain: A
Geoecological Approach to Glacier Forelands and Primary Succession, Cambridge University Press, Cambridge, UK, 1992.
[2] A. Jumpponen, H. Väre, K. G. Mattson, R. Ohtonen, and J. M.
Trappe, “Characterization of “safe sites” for pioneers in primary
succession on recently deglaciated terrain,” Journal of Ecology,
vol. 87, no. 1, pp. 98–105, 1999.
[3] K. Mizuno, “Succession processes of alpine vegetation in
response to glacial fluctuations of tyndall glacier, Mt. Kenya,
Kenya,” Arctic and Alpine Research, vol. 30, no. 4, pp. 340–348,
1998.
[4] J. L. Harper, J. N. Clatworthy, I. H. McNaughton, and G. S. Sagar,
“The evolution of closely related species living in the same area,”
Evolution, vol. 15, no. 2, pp. 209–227, 1961.
[5] S. Pignatti, Ecologia del Paesaggio, U.T.E.T, Milan, Italy, 1994.
[6] R. Hayat, S. Ali, U. Amara, R. Khalid, and I. Ahmed, “Soil
beneficial bacteria and their role in plant growth promotion: a
review,” Annals of Microbiology, vol. 60, no. 4, pp. 579–598, 2010.
[7] I. P. Edwards, H. Bürgmann, C. Miniaci, and J. Zeyer, “Variation
in microbial community composition and culturability in the
rhizosphere of Leucanthemopsis alpina (L.) Heywood and
adjacent bare soil along an alpine chronosequence,” Microbial
Ecology, vol. 52, no. 4, pp. 679–692, 2006.
[8] C. Miniaci, M. Bunge, L. Duc, I. Edwards, H. Bürgmann, and J.
Zeyer, “Effects of pioneering plants on microbial structures and
functions in a glacier forefield,” Biology and Fertility of Soils, vol.
44, no. 2, pp. 289–297, 2007.
[9] D. Tscherko, U. Hammesfahr, M.-C. Marx, and E. Kandeler,
“Shifts in rhizosphere microbial communities and enzyme
activity of Poa alpina across an alpine chronosequence,” Soil
Biology and Biochemistry, vol. 36, no. 10, pp. 1685–1698, 2004.
[10] D. Tscherko, U. Hammesfahr, G. Zeltner, E. Kandeler, and R.
Böcker, “Plant succession and rhizosphere microbial communities in a recently deglaciated alpine terrain,” Basic and Applied
Ecology, vol. 6, no. 4, pp. 367–383, 2005.
[11] F. E. Z. Haichar, C. Marol, O. Berge et al., “Plant host habitat and
root exudates shape soil bacterial community structure,” ISME
Journal, vol. 2, no. 12, pp. 1221–1230, 2008.
[12] Y. Osanai, D. S. Bougoure, H. L. Hayden, and M. J. Hovenden,
“Co-occurring grass species differ in their associated microbial
community composition in a temperate native grassland,” Plant
and Soil, vol. 368, no. 1-2, pp. 419–431, 2013.
[13] R. Costa, M. Götz, N. Mrotzek, J. Lottmann, G. Berg, and
K. Smalla, “Effects of site and plant species on rhizosphere
community structure as revealed by molecular analysis of
microbial guilds,” FEMS Microbiology Ecology, vol. 56, no. 2, pp.
236–249, 2006.
[14] E. Benizri and B. Amiaud, “Relationship between plants and soil
microbial communities in fertilized grasslands,” Soil Biology and
Biochemistry, vol. 37, no. 11, pp. 2055–2064, 2005.
[15] A. K. Patra, L. Abbadie, A. Clays-Josserand et al., “Effects of
management regime and plant species on the enzyme activity
and genetic structure of N-fixing, denitrifying and nitrifying
bacterial communities in grassland soils,” Environmental Microbiology, vol. 8, no. 6, pp. 1005–1016, 2006.
[16] N. Nunan, T. J. Daniell, B. K. Singh, A. Papert, J. W. McNicol,
and J. I. Prosser, “Links between plant and rhizoplane bacterial
communities in grassland soils, characterized using molecular
techniques,” Applied and Environmental Microbiology, vol. 71,
no. 11, pp. 6784–6792, 2005.
BioMed Research International
[17] E. Yergeau, S. Bokhorst, A. H. L. Huiskes, H. T. S. Boschker, R.
Aerts, and G. A. Kowalchuk, “Size and structure of bacterial,
fungal and nematode communities along an Antarctic environmental gradient,” FEMS Microbiology Ecology, vol. 59, no. 2, pp.
436–451, 2007.
[18] E. Yergeau, K. K. Newsham, D. A. Pearce, and G. A. Kowalchuk,
“Patterns of bacterial diversity across a range of Antarctic
terrestrial habitats,” Environmental Microbiology, vol. 9, no. 11,
pp. 2670–2682, 2007.
[19] X. F. Zhang, L. Zhao, S. J. Xu Jr., Y. Z. Liu, H. Y. Liu, and G. D.
Cheng, “Soil moisture effect on bacterial and fungal community
in Beilu River (Tibetan Plateau) permafrost soils with different
vegetation types,” Journal of Applied Microbiology, vol. 114, no.
4, pp. 1054–1065, 2013.
[20] C. R. Kuske, L. O. Ticknor, M. E. Miller et al., “Comparison
of soil bacterial communities in rhizospheres of three plant
species and the interspaces in an arid grassland,” Applied and
Environmental Microbiology, vol. 68, no. 4, pp. 1854–1863, 2002.
[21] G. Habler, M. Thöni, and B. Grasemann, “Cretaceous metamorphism in the Austroalpine Matsch Unit (Eastern Alps): the
interrelation between deformation and chemical equilibration
processes,” Mineralogy and Petrology, vol. 97, no. 3-4, pp. 149–
171, 2009.
[22] C. Knoll and H. Kerschner, “A glacier inventory for South
Tyrol, Italy, based on airborne laser-scanner data,” Annals of
Glaciology, vol. 50, no. 53, pp. 46–52, 2009.
[23] Ø. Hammer, D. A. T. Harper, and P. D. Ryan, “Past: paleontological statistics software package for education and data analysis,”
Palaeontologia Electronica, vol. 4, no. 1, pp. 1–9, 2001.
[24] G. Muyzer, E. C. de Waal, and A. G. Uitterlinden, “Profiling
of complex microbial populations by denaturing gradient gel
electrophoresis analysis of polymerase chain reaction-amplified
genes coding for 16S rRNA,” Applied and Environmental Microbiology, vol. 59, no. 3, pp. 695–700, 1993.
[25] M. Cardinale, L. Brusetti, P. Quatrini et al., “Comparison of
different primer sets for use in automated ribosomal intergenic
spacer analysis of complex bacterial communities,” Applied and
Environmental Microbiology, vol. 70, no. 10, pp. 6147–6156, 2004.
[26] A. M. Sass, H. Sass, M. J. L. Coolen, H. Cypionka, and J.
Overmann, “Microbial communities in the chemocline of a
hypersaline deep-sea basin (Urania basin, Mediterranean Sea),”
Applied and Environmental Microbiology, vol. 67, no. 12, pp.
5392–5402, 2001.
[27] J. R. Cole, Q. Wang, E. Cardenas et al., “The Ribosomal Database
Project: improved alignments and new tools for rRNA analysis,”
Nucleic Acids Research, vol. 37, no. 1, pp. D141–D145, 2009.
[28] G. N. Rees, D. S. Baldwin, G. O. Watson, S. Perryman, and D.
L. Nielsen, “Ordination and significance testing of microbial
community composition derived from terminal restriction
fragment length polymorphisms: application of multivariate
statistics,” Antonie van Leeuwenhoek, vol. 86, no. 4, pp. 339–347,
2004.
[29] K. R. Clarke, “Non-parametric multivariate analyses of changes
in community structure,” Australian Journal of Ecology, vol. 18,
no. 1, pp. 117–143, 1993.
[30] K. R. Clarke and R. M. Warwick, Change in Marine Communities: An Approach to Statistical Analysis and Interpretation,
PRIMER-E, Plymouth, UK, 2nd edition, 2001.
[31] M. Hamady, C. Lozupone, and R. Knight, “Fast UniFrac:
facilitating high-throughput phylogenetic analyses of microbial
communities including analysis of pyrosequencing and PhyloChip data,” ISME Journal, vol. 4, no. 1, pp. 17–27, 2010.
BioMed Research International
[32] K. M. Urbanska, Safe Sites—Interface of Plant Population Ecology and Restoration Ecology. Restoration Ecology and Sustainable
Development, Cambridge University Press, Cambridge, UK,
1997.
[33] R. Chou, C. Vardy, and R. L. Jefferies, “Establishment from
leaves and other plant fragments produced by the foraging
activites of geese,” Functional Ecology, vol. 6, no. 3, pp. 297–301,
1992.
[34] H. P. Bais, S.-W. Park, T. L. Weir, R. M. Callaway, and J. M.
Vivanco, “How plants communicate using the underground
information superhighway,” Trends in Plant Science, vol. 9, no.
1, pp. 26–32, 2004.
[35] J. Farrar, M. Hawes, D. Jones, and S. Lindow, “How roots control
the flux of carbon to the rhizosphere,” Ecology, vol. 84, no. 4, pp.
827–837, 2003.
[36] A. M. Hirsch, W. D. Bauer, D. M. Bird, J. Cullimore, B. Tyler,
and J. I. Yoder, “Molecular signals and receptors: controlling
rhizosphere interactions between plants and other organisms,”
Ecology, vol. 84, no. 4, pp. 858–868, 2003.
[37] C. M. Hansel, S. Fendorf, P. M. Jardine, and C. A. Francis,
“Changes in bacterial and archaeal community structure and
functional diversity along a geochemically variable soil profile,”
Applied and Environmental Microbiology, vol. 74, no. 5, pp.
1620–1633, 2008.
[38] N. M. Kennedy, D. E. Gleeson, J. Connolly, and N. J. W. Clipson,
“Seasonal and management influences on bacterial community
structure in an upland grassland soil,” FEMS Microbiology
Ecology, vol. 53, no. 3, pp. 329–337, 2005.
[39] G. A. Kowalchuk, D. S. Buma, W. de Boer, P. G. L. Klinkhamer,
and J. A. van Veen, “Effects of above-ground plant species composition and diversity on the diversity of soil-borne microorganisms,” Antonie van Leeuwenhoek, vol. 81, no. 1–4, pp. 509–
520, 2002.
[40] C. L. Lauber, M. S. Strickland, M. A. Bradford, and N. Fierer,
“The influence of soil properties on the structure of bacterial
and fungal communities across land-use types,” Soil Biology and
Biochemistry, vol. 40, no. 9, pp. 2407–2415, 2008.
[41] P. Marschner, C.-H. Yang, R. Lieberei, and D. E. Crowley, “Soil
and plant specific effects on bacterial community composition
in the rhizosphere,” Soil Biology and Biochemistry, vol. 33, no. 11,
pp. 1437–1445, 2001.
[42] M. E. Schutter, J. M. Sandeno, and R. P. Dick, “Seasonal, soil
type, and alternative management influences on microbial communities of vegetable cropping systems,” Biology and Fertility of
Soils, vol. 34, no. 6, pp. 397–410, 2001.
[43] N. Weinert, Y. Piceno, G.-C. Ding et al., “PhyloChip hybridization uncovered an enormous bacterial diversity in the rhizosphere of different potato cultivars: many common and few
cultivar-dependent taxa,” FEMS Microbiology Ecology, vol. 75,
no. 3, pp. 497–506, 2011.
[44] T. Wu, D. O. Chellemi, J. H. Graham, K. J. Martin, and E. N.
Rosskopf, “Comparison of soil bacterial communities under
diverse agricultural land management and crop production
practices,” Microbial Ecology, vol. 55, no. 2, pp. 293–310, 2008.
[45] D. A. Lipson, S. K. Schmidt, and R. K. Monson, “Links between
microbial population dynamics and nitrogen availability in an
alpine ecosystem,” Ecology, vol. 80, no. 5, pp. 1623–1631, 1999.
[46] R. Ohtonen, H. Fritze, T. Pennanen, A. Jumpponen, and
J. Trappe, “Ecosystem properties and microbial community
changes in primary succession on a glacier forefront,” Oecologia,
vol. 119, no. 2, pp. 239–246, 1999.
9
[47] J. E. Knelman, T. M. Legg, S. P. O’Neill et al., “Bacterial
community structure and function change in association with
colonizer plants during early primary succession in a glacier
forefield,” Soil Biology and Biochemistry, vol. 46, pp. 172–180,
2012.
[48] R. Danovaro, G. M. Luna, A. Dell’Anno, and B. Pietrangeli,
“Comparison of two fingerprinting techniques, terminal
restriction fragment length polymorphism and automated
ribosomal intergenic spacer analysis, for determination of
bacterial diversity in aquatic environments,” Applied and
Environmental Microbiology, vol. 72, no. 9, pp. 5982–5989,
2006.
[49] A. Okubo and S.-I. Sugiyama, “Comparison of molecular fingerprinting methods for analysis of soil microbial community
structure,” Ecological Research, vol. 24, no. 6, pp. 1399–1405,
2009.
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 568549, 8 pages
http://dx.doi.org/10.1155/2014/568549
Research Article
Contrasted Reactivity to Oxygen Tensions in Frankia sp.
Strain CcI3 throughout Nitrogen Fixation and Assimilation
Faten Ghodhbane-Gtari,1,2 Karima Hezbri,1 Amir Ktari,1 Imed Sbissi,1
Nicholas Beauchemin,2 Maher Gtari,1,2 and Louis S. Tisa2
1
Laboratoire Microorganismes et Biomolécules Actives, Université Tunis El Manar (FST) and Université Carthage (INSAT),
Campus Universitaire, 2092 Tunis, Tunisia
2
Department of Molecular, Cellular & Biomedical Sciences, University of New Hampshire, 46 College Road, Durham,
NH 03824-2617, USA
Correspondence should be addressed to Louis S. Tisa; [email protected]
Received 18 April 2014; Revised 28 April 2014; Accepted 15 May 2014; Published 28 May 2014
Academic Editor: Ameur Cherif
Copyright © 2014 Faten Ghodhbane-Gtari et al. This is an open access article distributed under the Creative Commons Attribution
License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Reconciling the irreconcilable is a primary struggle in aerobic nitrogen-fixing bacteria. Although nitrogenase is oxygen and reactive
oxygen species-labile, oxygen tension is required to sustain respiration. In the nitrogen-fixing Frankia, various strategies have been
developed through evolution to control the respiration and nitrogen-fixation balance. Here, we assessed the effect of different
oxygen tensions on Frankia sp. strain CcI3 growth, vesicle production, and gene expression under different oxygen tensions.
Both biomass and vesicle production were correlated with elevated oxygen levels under both nitrogen-replete and nitrogendeficient conditions. The mRNA levels for the nitrogenase structural genes (nif HDK) were high under hypoxic and hyperoxic
conditions compared to oxic conditions. The mRNA level for the hopanoid biosynthesis genes (sqhC and hpnC) was also elevated
under hyperoxic conditions suggesting an increase in the vesicle envelope. Under nitrogen-deficient conditions, the hup2 mRNA
levels increased with hyperoxic environment, while hup1 mRNA levels remained relatively constant. Taken together, these results
indicate that Frankia protects nitrogenase by the use of multiple mechanisms including the vesicle-hopanoid barrier and increased
respiratory protection.
1. Introduction
The genus Frankia is comprised of nitrogen-fixing actinobacteria that are able to establish a mutualistic symbiosis with
a variety of dicotyledonous host plants that results in the
establishment of a root nodule structure [1–6]. The bacteria
nourish their host plant with combined nitrogen and the
plants provide in return carbon and energy. This symbiosis
allows actinorhizal host plants to colonize nutrient-poor
soils. Besides its life style within the host plant, these bacteria
are members of soil community although less information
is known about this life style [7]. Under arid tropic and
subtropic conditions of North Africa, actinorhizal plants are
essentially represented by fast growing and highly tolerant
trees from the family Casuarinaceae [8].
Under atmospheric oxygen conditions, Frankia actively
fixes dinitrogen to ammonium within the root nodules of the
host plants and aerobically in culture [9–15]. The oxygenlabile nitrogenase enzyme is localized within specialized
thick-walled structures, termed vesicles that are formed in
planta and in vitro [2, 16–18]. Their shape is strain dependent and host-plant-influenced. Vesicles act as specialized
structures for the nitrogen fixation process and are formed
terminally on short side branches of hyphae that have a
septum near their base. The mature vesicle is surrounded
by an envelope that extends down the stalk of the vesicle
past the basal septum, which separates the vesicle from the
hypha. The envelope surrounding the vesicle is composed
of multilaminated lipid layers containing primarily bacteriohopanetetrol and its derivatives [19–22]. It is believed that this
lipid envelope acts as an oxygen diffusion barrier to protect
the nitrogenase enzyme from oxygen inactivation [19].
Unlike other actinorhizal plants, Frankia found within
the root nodules of Casuarina and Allocasuarina plants are
2
devoid of symbiotic vesicle structures [23, 24]. A positive
correlation was observed between the differentiation of intracellular hyphae and the lignifications of the host-infected cell
walls [23]. In several actinorhizal nodules, a low oxygen tension was shown to be consistent with the high concentrations
of hemoglobin [2]. Frankia are known to produce truncated
hemoglobins [25–27]. Besides hemoglobins, Frankia possess
hydrogenases that may act as oxygen-scavenging enzymes
[28]. Sequencing of several Frankia genomes [29–34] has
provided insight on the physiology and opened up new
genomics tools for these microbes. These databases have
been used in transcriptomics [35–37] and proteomics studies
[38–40] on these bacteria. The aim of the present study
was to investigate the expression levels for several selected
genes involved under different oxygen concentration for the
Casuarina compatible Frankia sp. strain CcI3. These genes
were involved in the following functions: nitrogen fixation
and assimilation, hopanoid biosynthesis, hydrogen uptake,
and oxidative stress.
2. Materials and Methods
2.1. Culture Conditions and Experimental Design. Frankia sp.
strain CcI3 [41] was grown and maintained at 28∘ C in basal
MP growth medium with 5.0 mM propionate and 5.0 mM
NH4 Cl as carbon and nitrogen sources, respectively, as
described previously [42].
In all experimental procedures, Frankia cells were grown
for 7 days in 250 mL cylindrical bottles with a working
MP medium volume of 50 mL with and without NH4 Cl for
nitrogen-deficient and nitrogen-replete conditions, respectively. Three sets of oxygen tensions were considered: oxic
(atmospheric condition), hypoxic (reduced partial pressure
of oxygen), and hyperoxic (elevated oxygen levels). Hypoxic
conditions were generated by placing the cultures in Brewer’s
jar that contained reduced partial pressures of oxygen by the
use of gas packets (BBL GasPak BBL CampyPak System).
For this system, water interacts with catalyst in the packet
generating a reduced partial pressure of oxygen within the
chamber. Hyperoxic conditions were generated by continuously air-sparging the cultures via an aquarium pump.
2.2. Growth Assessment and Vesicle Count. For dry weight
determinations, cell cultures were collected on tarred membrane filters (type HA, 0.45 um pore size; Millipore Corp.).
The filters were placed in a Petri dish over desiccant and
dried at 90∘ C to constant weight [43]. In parallel, protein
content was measured. Briefly, cell samples were solubilized
by heating for 15 min at 90∘ C in 1.0 N NaOH and total proteins
were measured using BCA method [44].
Vesicle numbers were determined as previously described
[45, 46]. Briefly, cells were sonicated for 30 s with a Braun
model 350 sonifier under power setting of 3 using microtip
probe. This treatment disrupted the mycelia and released
vesicles. The numbers of vesicles were counted by using
a Petroff-Hausser counting chamber with a phase-contrast
microscope at magnification of 400x.
BioMed Research International
2.3. Determination of Ammonia. Ammonium concentration
was determined in cell-free media using modified protocol of
Berthelot’s reagent [47].
2.4. RNA Extraction, RT-PCRs, and Q-PCR. For these experiments, all solutions and materials were DEPC-treated to
prevent RNA degradation. RNA extractions were performed
by the Triton X100 method as previously described [48]. RNA
samples were treated with DNase I (New England Biolabs)
according to the manufacturer’s recommendations. RNA
samples were quantified with a Nanodrop 2000c spectrophotometer (Thermo Scientific) and stored at −80∘ C until use.
The cDNA synthesis was performed using hexamer primers,
400 ng RNA and SuperScript III reverse transcriptase (Invitrogen) according to the manufacturer’s recommendations.
The cDNA was quantified by a Nanodrop 2000c spectrophotometer, diluted to 10 ng/𝜇L working stocks in DNAse-free,
RNAse-free H2 O, and stored at −20∘ C until use.
Frankia gene expression analyses were performed by
qRT-PCR using specific primers (Table 1) and SYBR Green
PCR Master Mix (Applied Biosystems) as described previously [49]. Briefly, each 25 𝜇L reaction contained 50 ng template cDNA, 300 nM of the forward and reverse primer mix,
and SYBR Green PCR Master Mix. Parameters for the Agilent
MP3000 were as follows: (1) 95∘ C for 15 min, (2) 40 cycles of
95∘ C for 15 s and 60∘ C for 30 s, and (3) thermal disassociation
cycle of 95∘ C for 60 s, 55∘ C for 30 s, and incremental increases
in temperature to 95∘ C for 30 s. Reactions were performed
in triplicates and the comparative threshold-cycle method
was used to quantify gene expression. The results were
standardized with rpsA expression levels. Relative expression
(fold changes) was determined by the Pfaffl method [50] with
the control as the calibrator. Two biological replicates of the
triplicate samples were averaged.
3. Results
3.1. Growth and Vesicle Production under Different Oxygen
Pressures. Figure 1 shows the effect of oxygen on the growth
yield of Frankia sp. strain CcI3. Under nitrogen-replete
conditions (NH4 ), the biomass of cells grown under hyperoxic conditions was greater than both cultures grown under
oxic and hypoxic conditions. Under nitrogen-deficient (N2 )
conditions, the biomass correlated with the oxygen level with
the hyperoxic conditions generating the greatest biomass.
Furthermore, vesicle production under nitrogen-deficient
(N2 ) conditions positively correlated with oxygen tension.
Cells under hyperoxic (air-sparged) conditions produced 2.6and 5.4-fold more vesicles (6.50 ± 0.41 × 106 /mg) than oxic
(2.45 ± 0.29 × 106 /mg) and hypoxic (1.20 ± 0.36 × 106 /mg)
conditions, respectively. Analysis of ammonia metabolism by
Frankia CcI3 indicates that it was correlated with oxygen
tension. With nitrogen-replete conditions, hyperoxic conditions resulted in the highest ammonia consumption, followed
by oxic condition and lastly hypoxic condition (Figure 1(c)).
Under nitrogen-deficient conditions the level of ammonium ions increased under lower oxygen tension. This level
decreased with corresponding increases in oxygen tension.
BioMed Research International
3
Table 1: Primers used in this study.
Locus tag
Gene
Gene identity
francci3 4488
nif H
Nitrogenase reductase iron-sulfur protein
francci3 4487
nif D
Nitrogenase molybdenum-iron protein alpha chain
francci3 4486
nif K
Nitrogenase molybdenum-iron protein beta chain
francci3 4496
hup1
Nickel-dependant hydrogenase, large subunit
francci3 1076
hup2
Uptake hydrogenase, large subunit
francci3 1149
hboO
Truncated hemoglobin
francci3 2581
hboN
Truncated hemoglobin
francci3 0823
sqhC
Squalene hopene cyclase
francci3 0819
hpnC
Squalene synthase
francci3 2949
katA
Catalase
francci3 2817
sodA
Superoxide dismutase
francci3 3012
gltD
Glutamate synthase, small subunit
francci3 3013
gltB
Glutamate synthase, large subunit
francci3 3142
glnA
Glutamine synthetase, type I
francci3 3143
glnA
Glutamine synthetase, type II
francci3 4059
glnA
Glutamine synthetase, catalytic region
francci3 1057
rpsA
30S ribosomal protein S1
Sequence
5󸀠 -CGACAACGACATGAAGACC-3󸀠
5󸀠 -CTTGCCGATGATGCTCTC-3󸀠
5󸀠 -AAGGACATCGTCAACATCAGCCAC-3󸀠
5󸀠 -AACTGCATCGCGGCGAAGTTATTC-3󸀠
5󸀠 -TGACGACGACTCCGGAAACAAACA-3󸀠
5󸀠 -TGTGGTAGACCTCGTCCTTGAACA-3󸀠
5󸀠 -AACAAATCTGCGACGTCACGGTCA-3󸀠
5󸀠 -ACTCTCGATCCATTCACCGCAGTA-3󸀠
5󸀠 -TGGAAGGTCAACTGGCTGGAGAA-3󸀠
5󸀠 -ATGTCTAGGCAGTACCGGAGGAAGAA-3󸀠
5󸀠 -GGGACGCCTGGCTGAAGA-3󸀠
5󸀠 -CCAGAGCTGCCTGTCGAGATC-3󸀠
5󸀠 -CACCCCTCTTTGCCAACCG-3󸀠
5󸀠 -GGTGGTTTCCGTCGGGAC-3󸀠
5󸀠 -TGCAATGGCTGCTGGACAA-3󸀠
5󸀠 -TGCCGTAGACGTGGTTGAT-3󸀠
5󸀠 -AACTTCCCGGTCTCGCCGTT-3󸀠
5󸀠 -AACGCGTTGAAGTGGAAACGAACC-3󸀠
5󸀠 -ACATGCCGGTGTTCTTCATTCAGG-3󸀠
5󸀠 -ACATCATCATGTGGCATCGACTCGG-3󸀠
5󸀠 -GTGCCAATGACACCCTTGAGAAGA-3󸀠
5󸀠 -AGTGGAGAATATGCCCGGAAAGGT-3󸀠
5󸀠 -TGCATGCGACGAACAACTTCCC-3󸀠
5󸀠 -ATGATGCTGACCTCGATCTGCTTG-3󸀠
5󸀠 -CGTGCTGAAGGTGATGTCCAAGAT-3󸀠
5󸀠 -AAATAGGCGTCGATCAGTTCCTGG-3󸀠
5󸀠 -ATGACCCGATCACCAAGGAACAGT-3󸀠
5󸀠 -GGGTTGTAGTCATAACGGACATCG-3󸀠
5󸀠 -AACTTCTCCACCAGGCAGACGAT-3󸀠
5󸀠 -AGAACTTGTTCCACGGAGCTGTCT-3󸀠
5󸀠 -TACAACATCGACTACGCGCTTTCC-3󸀠
5󸀠 -ATACCGGAACACGATCTCGAACTG-3󸀠
5󸀠 -CGAAGTCCGTTCCGAGTTC-3󸀠
5󸀠 -CGCCGAAGTTGACGATGG-3󸀠
Locus tag and gene designation were determined from the Integrated Microbial Genomes System (IMG) at the Joint Genome Institute (https://img.jgi.doe.gov/)
[51].
3.2. Expression of Nitrogen Fixation and Assimilation Genes
under Different Oxygen Pressures. The effect of oxygen on the
expression of several genes involved in nitrogen fixation and
assimilation was measured by detecting changes in mRNA
levels via qRT-PCR (Figure 2). For nitrogen-deficient conditions, the level of structural nitrogenase genes (nif HDK)
mRNA increased >10-fold under hyperoxic and hypoxic
conditions compared to oxic condition (Figure 2(a)). Under
nitrogen-replete conditions, the expression levels for these
genes were very low and there was no change with different
oxygen tensions.
The Frankia genome contains two glutamate synthase
genes (gltB and gltD) encoding the large and small subunits of
the enzyme. These two glutamate synthase genes were studied
for their expression levels under three oxygen tensions. The
mRNA levels of the gltB gene were reduced except under
hyperoxic and nitrogen-replete conditions (Figure 2(b)). The
gltD mRNA levels increased slightly (1.3–2.5-fold) under the
different nitrogen and oxygen conditions. There were four
glutamine synthetase orthologs found within the Frankia sp.
strain CcI3 genome. We were able to follow the expression
of three of these glnA genes (Figure 2(c)). The level of
francci3 3143 mRNA was controlled by nitrogen. Under all
oxygen conditions, francci3 3143 mRNA levels increased 10–
15-fold under nitrogen-deficient (N2 ) conditions. Both high
and low oxygen tensions increased the level of francci3 3143
mRNA. The level of francci3 3142 mRNA was decreased
under nitrogen-deficient (N2 ) conditions and showed 7-fold
increase under hyperoxic under nitrogen-replete conditions.
The levels of francci3 4059 mRNA remained constant except
under hyperoxic conditions, in which levels increased 15fold. Under hyperoxic conditions, the levels of francci3 4059
mRNA were controlled by nitrogen status and increased
approximately 2-3-fold from nitrogen-replete (NH4 ) conditions.
3.3. Expression of Genes Known to Protect Nitrogenase from
Oxygen and Reactive Oxygen Species. The biosynthesis of
4
BioMed Research International
0.16
0.3
0.14
Protein (mg/mL)
Dry weight (mg/mL)
0.25
0.2
0.15
0.1
0.05
0.12
0.1
0.08
0.06
0.04
0.02
0
LN2
LNH4
NN2
NNH4
HN2
0
HNH4
LN2
LNH4
(a)
NN2
NNH4
HN2
HNH4
(b)
Ammonium ions (mg/L)
70
60
50
40
30
20
10
0
LN2
LNH4
NN2
NNH4
HN2
HNH4
(c)
Figure 1: Biomass yields of Frankia sp. strain CcI3 grown under nitrogen fixation (N2 ) and nitrogen-replete (NH4 ) at hypoxic (L), oxic (N),
and hyperoxic (H) conditions as estimation by (a) dry weight and (b) total protein and determination of (c) ammonium ion concentrations.
hopanoids has been correlated with vesicle development
[19]. The effect of oxygen tension on the expression of the
squalene synthase (hpnC) and squalene/phytoene cyclase
(sqhC) genes was examined (Figure 2(d)). Under nitrogenreplete conditions (NH4 ), the level of mRNA for sqhC showed
a 2-fold increase for hyperoxic conditions. A smaller increase
was observed for hpnC mRNA levels. In general, sqhC and
hpnC were expressed constitutively with comparable mRNA
levels for hypoxic and oxic levels. Under nitrogen-deficient
(N2 ) conditions, the mRNA levels of both genes (sqhC and
hpnC) increased 2- and 1.5-fold, respectively.
The Frankia CcI3 genome contains two hydrogenase
operons [30, 52, 53]. We tested the effects of oxygen tension and nitrogen status of their gene expression levels
(Figure 2(e)). Under nitrogen-replete (NH4 ) conditions, the
level of mRNA for hup2 increased proportionally with the
level of oxygen present, while the level of mRNA for hup1
only increased under hyperoxic conditions. The expression of
hup2 was influenced by the nitrogen status of the cells and by
the oxygen levels. Under both conditions, hup2 mRNA levels
increased, but hup1 expression remained constant.
The effect of oxygen tension and nitrogen status was
investigated on the expression of two truncated hemoglobins
(hboO and hboN). The level of mRNA of hboO and hboN
increased under hyperoxic condition for both nitrogen conditions (Figure 2(f)). Under nitrogen-replete (NH4 ) conditions, mRNA levels for hboO increased proportionally to
the oxygen tension levels. Under hypoxic nitrogen-deficient
conditions, mRNA levels for hboN increased about 1.5-fold.
The effects of oxygen tension and nitrogen status on the
expression levels of two oxygen defense enzymes, catalase
(katA) and superoxide dismutase (sodA), were also tested
(Figure 2(g)). Under hyperoxic conditions, the mRNA levels
of katA increased 6.5- and 8-fold under nitrogen-deficient
(N2 ) and nitrogen-replete (NH4 ) conditions, respectively. The
expression of the sodA gene appeared to be constitutive under
all oxygen tensions and both nitrogen statuses.
4. Discussion
Without a doubt, the vesicle is the most characteristic
morphogenetic structure produced by Frankia [1]. Vesicles
are functionally analogous to cyanobacterial heterocysts
providing unique specialized cells that allow nitrogen fixation
under aerobic condition [54, 55]. In this study, the growth
of Frankia strain CcI3 was evaluated under three oxygen
tensions. The results indicate that growth increased with
elevated oxygen tensions (Figure 1) confirming the aerobic
nature of the microbe. Although the dry weight measurement
increased, the total protein values were reduced under hyperoxic nitrogen-deficient (N2 ) conditions. This result would
imply that the cells were producing other metabolic products
under this condition and a similar level of protein compared
to hypoxic nitrogen-deficient (N2 ) condition. Thus, this result
BioMed Research International
5
3
35
2.5
30
250
200
25
2
150
20
1.5
100
15
1
50
0.5
0
0
LN2 LNH4 NN2 NNH4 HN2 HNH4
nifH
nifD
nifK
10
5
LN2 LNH4 NN2 NNH4 HN2 HNH4
gltD
gltB
LN2 LNH4 NN2 NNH4 HN2 HNH4
francci3 3142
francci3 3143
(a)
(b)
6
30
5
25
4
20
3
15
2
10
1
5
0
0
0
LN2 LNH4 NN2 NNH4 HN2 HNH4
(c)
5
4.5
4
3.5
3
2.5
2
1.5
1
0.5
0
LN2 LNH4 NN2 NNH4 HN2 HNH4
(d)
LN2 LNH4 NN2 NNH4 HN2 HNH4
HboO
HboN
hup2
hup1
sqhC
hpnC
francci3 4059
(e)
(f)
9
8
7
6
5
4
3
2
1
0
LN2 LNH4 NN2 NNH4 HN2 HNH4
SodA
KatA
(g)
Figure 2: Relative gene expression (fold change) in response to hyperoxic and hypoxic conditions. Frankia cultures were grown under
nitrogen-replete (NH4 ) or nitrogen-deficient (N2 ) conditions. These cultures were exposed to oxic (N), hyperoxic (H), and hypoxic (L)
conditions as described in Section 2. Experimental gene expression was normalized to the rpsA housekeeping gene and compared to the
calibrator (NH4 oxic conditions). The following genes were analyzed: (a) nif HDK (b) gltB and gltD, (c) glnA genes, (d) hpnC and sqhC, (e)
hup1 and hup2, (f) hboN and hboO, and (g) sodA and katA.
suggests that part of the respiration was uncoupled providing
some oxygen protection. Frankia contains two respiratory
systems and a cyanide-insensitive system was proposed to
help protect nitrogenase from oxygen inactivation [46]. With
other aerobic nitrogen-fixing bacteria, increased respiratory
rates in response to elevated oxygen tensions help maintain
low levels of intracellular oxygen protecting nitrogenase from
inactivation [56, 57]. Under nitrogen-deficient (N2 ) conditions, vesicles were produced and correlated with oxygen
tensions. The numbers of vesicles produced per mg dry
weight increased with elevated oxygen levels. These results
confirm those obtained previously [58, 59].
In our study, we investigated the effects of oxygen on
gene expression for a variety of functional genes involved in
nitrogen fixation, nitrogen assimilation, and protection from
oxygen and other reactive oxygen species [60]. The levels
of expression for the structural nitrogenase genes (nif HDK)
indicate a concordant profile with clear induction under
6
nitrogen-deficient (N2 ) conditions. Transcriptome studies
on Frankia sp. strain CcI3 under nitrogen-deficient and
nitrogen-replete conditions also show an increase in nif HDK
gene expression [35, 36]. The levels of nif HDK mRNA
showed an increase under hypoxic and hyperoxic conditions
indicating that nitrogenase induction was influenced by
oxygen levels.
The hopanoid envelope has been postulated to be
involved in the protection of nitrogenase from oxygen
inactivation [19]. We found that mRNA levels of squalene
synthase (hpnC) and squalene-hopene cyclase (sqhC) genes
increased in response to oxygen tension under nitrogendeficient conditions, but remained constant under nitrogenreplete conditions (Figure 2(d)). The results correlate with
the increase in vesicle envelope observed under high oxygen
levels [61]. Nalin et al. [62] found only a slightly higher
hopanoid content under nitrogen-deficient conditions suggesting remobilization rather than nascent biosynthesis. Furthermore, the Frankia sp. strain CcI3 transcriptome profiles
under nitrogen-deficient and nitrogen-replete conditions did
not show any significant differences in hopanoid biosynthetic
genes [35, 36]. However, these studies were performed under
one oxygen tension while our study has investigated three
different oxygen tensions.
Analysis of the nitrogen assimilation genes (gltB, gltD,
and glnA) is a bit more complex. The Frankia CcI3 genome
contained several homologues of glnA. The mRNA level of
francci3 3143 correlated the best with nitrogen regulation,
being increased under nitrogen-deficient conditions. Transcriptome studies have shown that francci3 3143 expression
increased significantly under nitrogen-fixing conditions [35,
36], while all of the other homologues remained consistent.
This result would suggest that this gene encoded primary
nitrogen scavenging enzyme. The levels of expression were
also influenced by elevated oxygen tensions during increased
nitrogenase activity. The expression levels of the gltB and gltD
appear to be less influenced by oxygen tension. These effects
seemed in agreement with the ammonia metabolism results
that showed an increase in consumption under hyperoxic
conditions.
Our results on hemoglobin gene expression correlate with
previous results [48] that showed no increase in hboN and
hboO expression in response to nitrogen status increased
under low oxygen tension. However, our results conflict in
response to oxygen. We found that both hboN and hboO
mRNA levels increased under hyperoxic conditions. The use
of the more sensitive qRT-PCR in our study compared to RTPCR is the best explanation for these differences.
Frankia possesses two uptake hydrogenase systems [52,
53]. One of them has been correlated with symbiotic growth
and the other to free-living conditions [53]. Our results show
that hup2 gene expression was influenced by nitrogen status
suggesting that it was associated with vesicle production,
while hup1 gene expression was relatively constant. The
levels of hup2 mRNA increased proportionally with oxygen
tensions suggesting potential oxygen protection mechanism.
Anoxic conditions have no effect on hydrogenase gene
expression by Frankia CcI3 but increased by 30% for Frankia
BioMed Research International
alni ACN14a [60]. We did not test anoxic conditions in our
study.
Increased oxygen tension can lead to elevated oxidative
stress conditions. We investigated the influence of oxygen
tensions on reactive oxidative stress genes. While sodA
expression levels were constitutive, katA gene expression
increased under hyperoxic conditions. In general, our results
confirm those of Steele and Stowers [63], which examined
enzymatic activity levels. They reported an increase in catalase activity in cultures derepressed for nitrogen fixation
compared to ammonium-grown cultures.
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
Acknowledgments
Louis S. Tisa was supported in part by Agriculture and Food
Research Initiative Grant 2010-65108-20581 from the USDA
National Institute of Food and Agriculture, Hatch Grant
NH530, and the College of Life Sciences and Agriculture
at the University of New Hampshire, Durham, NH, USA.
This is scientific contribution number 2556 from the NH
Agricultural Experimental Station. Maher Gtari and Faten
Ghodhbane-Gtari were supported in part by a Visiting
Scientist and Postdoctoral Scientist Program administered by
the NH AES at the University of New Hampshire.
References
[1] D. R. Benson and W. B. Silvester, “Biology of Frankia strains,
actinomycete symbionts of actinorhizal plants,” Microbiological
Reviews, vol. 57, no. 2, pp. 293–319, 1993.
[2] K. Huss-Danell, “Actinorhizal symbioses and their N2 fixation,”
New Phytologist, vol. 136, no. 3, pp. 375–405, 1997.
[3] L. G. Wall, “The actinorhizal symbiosis,” Journal of Plant Growth
Regulation, vol. 19, no. 2, pp. 167–182, 2000.
[4] C. Santi, D. Bogusz, and C. Franche, “Biological nitrogen fixation in non-legume plants,” Annals of Botany, vol. 111, no. 5, pp.
743–767, 2013.
[5] J. Schwencke and M. Carú, “Advances in actinorhizal symbiosis:
host plant-Frankia interactions, biology, and applications in
arid land reclamation. A review,” Arid Land Research and
Management, vol. 15, no. 4, pp. 285–327, 2001.
[6] A. Sellstedt and K. H. Richau, “Aspects of nitrogen-fixing actinobacteria, in particular free-living and symbiotic Frankia,”
FEMS Microbiology Letters, vol. 342, no. 2, pp. 179–186, 2013.
[7] E. E. Chaia, L. G. Wall, and K. Huss-Danell, “Life in soil by the
actinorhizal root nodule endophyte Frankia. A review,” Symbiosis, vol. 51, no. 3, pp. 201–226, 2010.
[8] M. Gtari and J. O. Dawson, “An overview of actinorhizal plants
in Africa,” Functional Plant Biology, vol. 38, no. 8-9, pp. 653–661,
2011.
[9] J. D. Tjepkema, W. Ormerod, and J. G. Torrey, “Vesicle formation and acetylene reduction activity in Frankia sp. CPI1
cultured in defined nutrient media,” Nature, vol. 287, no. 5783,
pp. 633–635, 1980.
BioMed Research International
[10] J. D. Tjepkema, W. Ormerod, and J. G. Torrey, “Factors affecting
vesicle formation and acetylene reduction (nitrogenase activity)
in Frankia sp. CpI1,” Canadian Journal of Microbiology, vol. 27,
no. 8, pp. 815–823, 1981.
[11] D. Gauthier, H. G. Diem, and Y. Dommergues, “In vitro nitrogen fixation by 2 actinomycete strains isolated from Casuarina
nodules,” Applied and Environmental Microbiology, vol. 41, pp.
306–308, 1981.
[12] M. A. Murry, Z. Zhongze, and J. G. Torrey, “Effect of O2 on
vesicle formation, acetylene reduction, and O2 -uptake kinetics
in Frankia sp. HFPCc13 isolated from Casuarina cunninghamiana,” Canadian Journal of Microbiology, vol. 31, no. 9, pp. 804–
809, 1985.
[13] Z. Zhongze and J. G. Torrey, “Biological and cultural characteristics of the effective Frankia strain HFPCcl3 (Actinomycetales)
from Casuarina cunninghamiana (Casuarinaceae),” Annals of
Botany, vol. 56, no. 3, pp. 367–378, 1985.
[14] M. S. Fontaine, S. A. Lancelle, and J. G. Torrey, “Initiation and
ontogeny of vesicles in cultured Frankia sp. strain HFPArI3,”
Journal of Bacteriology, vol. 160, no. 3, pp. 921–927, 1984.
[15] A. J. P. Burggraaf and W. A. Shipton, “Studies on the growth of
Frankia isolates in relation to infectivity and nitrogen fixation
(acetylene reduction),” Canadian Journal of Botany, vol. 61, no.
11, pp. 2774–2782, 1983.
[16] T. M. Meesters, “Localization of nitrogenase in vesicles
of Frankia sp. Cc1.17 by immunogoldlabelling on ultrathin
cryosections,” Archives of Microbiology, vol. 146, no. 4, pp. 327–
331, 1987.
[17] T. M. Meesters, S. T. Van Genesen, and A. D. L. Akkermans,
“Growth, acetylene reduction activity and localization of nitrogenase in relation to vesicle formation in Frankia strains Cc1.17
and Cp1.2,” Archives of Microbiology, vol. 143, no. 2, pp. 137–142,
1985.
[18] T. M. Meesters, W. M. Vanvliet, and A. D. L. Akkermans,
“Nitrogenase is restricted to the vesicles in Frankia strain
EAN1pec,” Physiologia Plantarum, vol. 70, no. 2, pp. 267–271,
1987.
[19] A. M. Berry, O. T. Harriott, R. A. Moreau, S. F. Osman, D. R.
Benson, and A. D. Jones, “Hopanoid lipids compose the Frankia
vesicle envelope, presumptive barrier of oxygen diffusion to
nitrogenase,” Proceedings of the National Academy of Sciences of
the United States of America, vol. 90, no. 13, pp. 6091–6094, 1993.
[20] A. M. Berry, R. A. Moreaua, and A. D. Jones, “Bacteriohopanetetrol: abundant lipid in Frankia cells and in nitrogenfixing nodule tissue,” Plant Physiology, vol. 95, no. 1, pp. 111–115,
1991.
[21] O. T. Harriott, L. Khairallah, and D. R. Benson, “Isolation and
structure of the lipid envelopes from the nitrogen-fixing vesicles
of Frankia sp. strain CpI1,” Journal of Bacteriology, vol. 173, no.
6, pp. 2061–2067, 1991.
[22] H. C. Lamont, W. B. Silvester, and J. G. Torrey, “Nile red fluorescence demonstrates lipid in the envelope of vesicles from N2 fixing cultures of Frankia,” Canadian Journal of Microbiology,
vol. 34, no. 5, pp. 656–660, 1988.
[23] R. H. Berg and L. McDowell, “Endophyte differentiation in
Casuarina actinorhizae,” Protoplasma, vol. 136, no. 2-3, pp. 104–
117, 1987.
[24] R. H. Berg and L. Mcdowell, “Cytochemistry of the wall of
infected-cells in Casuarina actinorhizae,” Canadian Journal of
Botany, vol. 66, no. 10, pp. 2038–2047, 1988.
[25] V. Coats, C. R. Schwintzer, and J. D. Tjepkema, “Truncated
hemoglobins in Frankia CcI3: effects of nitrogen source, oxygen
7
[26]
[27]
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35]
[36]
[37]
[38]
[39]
[40]
concentration, and nitric oxide,” Canadian Journal of Microbiology, vol. 55, no. 7, pp. 867–873, 2009.
J. D. Tjepkema, “Hemoglobins in the nitrogen-fixing root
nodules of actinorhizal plants,” Canadian Journal of Botany, vol.
61, no. 11, pp. 2924–2929, 1983.
J. D. Tjepkema, R. E. Cashon, J. Beckwith, and C. R. Schwintzer,
“Hemoglobin in Frankia, a nitrogen-fixing actinomycete,”
Applied and Environmental Microbiology, vol. 68, no. 5, pp.
2629–2631, 2002.
A. Sellstedt, P. Reddell, and P. Rosbrook, “The occurrence
of hemoglobin and hydrogenase in nodules of 12 CasuarinaFrankia symbiotic associations,” Physiologia Plantarum, vol. 82,
no. 3, pp. 458–464, 1991.
F. Ghodbhane-Gtari, N. Beauchemin, D. Bruce et al., “Draft
genome sequence of Frankia sp. strain CN3, an atypical,
non-infective (Nod-) ineffective (Fix-) isolate from Coriaria
nepalensis,” Genome Announcements, vol. 1, no. 2, Article ID
e0008513, 2013.
P. Normand, P. Lapierre, L. S. Tisa et al., “Genome characteristics of facultatively symbiotic Frankia sp. strains reflect host
range and host plant biogeography,” Genome Research, vol. 17,
no. 1, pp. 7–15, 2007.
A. Sen, N. Beauchemin, D. Bruce et al., “Draft genome sequence
of Frankia sp. strain QA3, a nitrogen-fixing actinobacterium
isolated from the root nodule of Alnus nitida,” Genome
Announcements, vol. 1, no. 2, Article ID e0010313, 2013.
S. R. Mansour, R. Oshone, S. G. Hurst, K. Morris, W. K. Thomas,
and L. S. Tisa, “Draft genome sequence of Frankia sp. strain
CcI6, a salt-tolerant nitrogen-fixing actinobacterium isolated
from the root nodule of Casuarina cunninghamiana,” Genome
Announcements, vol. 2, no. 1, Article ID e01205-13, 2014.
I. Nouioui, N. Beauchemin, M. N. Cantor et al., “Draft
genome sequence of Frankia sp. strain BMG5.12, a nitrogenfixing actinobacterium isolated from Tunisian soils,” Genome
Announcements, vol. 1, no. 4, Article ID e00468-13, 2013.
L. G. Wall, N. Beauchemin, M. N. Cantor et al., “Draft genome
sequence of Frankia sp. strain BCU110501, a nitrogen-fixing
actinobacterium isolated from nodules of Discaria trinevis,”
Genome Announcements, vol. 1, no. 4, Article ID e00503-13,
2013.
D. M. Bickhart and D. R. Benson, “Transcriptomes of Frankia
sp. strain CcI3 in growth transitions,” BMC Microbiology, vol. 11,
article 192, 2011.
H.-I. Lee, A. J. Donati, D. Hahn, L. S. Tisa, and W.-S.
Chang, “Alteration of the exopolysaccharide production and
the transcriptional profile of free-living Frankia strain CcI3
under nitrogen-fixing conditions,” Applied Microbiology and
Biotechnology, vol. 97, no. 24, pp. 10499–10509, 2013.
N. Alloisio, C. Queiroux, P. Fournier et al., “The Frankia
alni symbiotic transcriptome,” Molecular Plant-Microbe Interactions, vol. 23, no. 5, pp. 593–607, 2010.
N. Alloisio, S. Félix, J. Maréchal et al., “Frankia alni proteome
under nitrogen-fixing and nitrogen-replete conditions,” Physiologia Plantarum, vol. 130, no. 3, pp. 440–453, 2007.
J. E. Mastronunzio and D. R. Benson, “Wild nodules can be
broken: proteomics of Frankia in field-collected root nodules,”
Symbiosis, vol. 50, no. 1-2, pp. 13–26, 2010.
J. E. Mastronunzio, Y. Huang, and D. R. Benson, “Diminished
exoproteome of Frankia spp. in culture and symbiosis,” Applied
and Environmental Microbiology, vol. 75, no. 21, pp. 6721–6728,
2009.
8
[41] Z. Zhang, M. F. Lopez, and J. G. Torrey, “A comparison of
cultural characteristics and infectivity of Frankia isolates from
root nodules of Casuarina species,” Plant and Soil, vol. 78, no.
1-2, pp. 79–90, 1984.
[42] N. J. Beauchemin, T. Furnholm, J. Lavenus et al., “Casuarina
root exudates alter the physiology, surface properties, and plant
infectivity of Frankia sp. strain CcI3,” Applied and Environmental Microbiology, vol. 78, no. 2, pp. 575–580, 2012.
[43] L. Tisa, M. Mcbride, and J. C. Ensign, “Studies of growth
and morphology of Frankiastrains EAN1pec , EuI1c , CpI1, and
ACN1AG ,” Canadian Journal of Botany, vol. 61, no. 11, pp. 2768–
2773, 1983.
[44] P. K. Smith, R. I. Krohn, G. T. Hermanson et al., “Measurement
of protein using bicinchoninic acid,” Analytical Biochemistry,
vol. 150, no. 1, pp. 76–85, 1985.
[45] L. S. Tisa and J. C. Ensign, “Comparative physiology of nitrogenase activity and vesicle development for Frankia strains CpI1,
ACN1AG , EAN1pec and EUN1f ,” Archives of Microbiology, vol. 147,
no. 4, pp. 383–388, 1987.
[46] L. S. Tisa and J. C. Ensign, “The calcium requirement for
functional vesicle development and nitrogen fixation by Frankia
strains EAN1pec and CpI1,” Archives of Microbiology, vol. 149, no.
1, pp. 24–29, 1987.
[47] E. D. Rhine, G. K. Sims, R. L. Mulvaney, and E. J. Pratt,
“Improving the Berthelot reaction for determining ammonium
in soil extracts and water,” Soil Science Society of America
Journal, vol. 62, no. 2, pp. 473–480, 1998.
[48] J. Niemann and L. S. Tisa, “Nitric oxide and oxygen regulate
truncated hemoglobin gene expression in Frankia strain CcI3,”
Journal of Bacteriology, vol. 190, no. 23, pp. 7864–7867, 2008.
[49] F. Perrine-Walker, P. Doumas, M. Lucas et al., “Auxin carriers
localization drives auxin accumulation in plant cells infected
by Frankia in Casuarina glauca actinorhizal nodules,” Plant
Physiology, vol. 154, no. 3, pp. 1372–1380, 2010.
[50] M. W. Pfaffl, “A new mathematical model for relative quantification in real-time RT-PCR,” Nucleic Acids Research, vol. 29, no.
9, article e45, 2001.
[51] V. M. Markowitz, F. Korzeniewski, K. Palaniappan et al., “The
integrated microbial genomes (IMG) system,” Nucleic Acids
Research, vol. 34, pp. D344–D348, 2006.
[52] P. Normand, C. Queiroux, L. S. Tisa et al., “Exploring the
genomes of Frankia,” Physiologia Plantarum, vol. 130, no. 3, pp.
331–343, 2007.
[53] M. Leul, P. Normand, and A. Sellstedt, “The organization,
regulation and phylogeny of uptake hydrogenase genes in
Frankia,” Physiologia Plantarum, vol. 130, no. 3, pp. 464–470,
2007.
[54] N. A. Noridge and D. R. Benson, “Isolation and nitrogenfixing activity of Frankia sp. strain CpI1 vesicles,” Journal of
Bacteriology, vol. 166, no. 1, pp. 301–305, 1986.
[55] L. S. Tisa and J. C. Ensign, “Isolation and nitrogenase activity of
vesicles from Frankia sp. strain EAN1pec ,” Journal of Bacteriology, vol. 169, no. 11, pp. 5054–5059, 1987.
[56] A. Hochman and R. H. Burris, “Effect of oxygen on acetylene
reduction by photosynthetic bacteria,” Journal of Bacteriology,
vol. 147, no. 2, pp. 492–499, 1981.
[57] H. Dalton and J. R. Postgate, “Effect of oxygen on growth of
Azotobacter chroococcum in batch and continuous cultures,”
Journal of General Microbiology, vol. 54, no. 3, pp. 463–473, 1968.
[58] M. A. Murry, Z. Zhongze, and J. G. Torrey, “Effect of O2 on vesicle formation, acetylene reduction, and O2 -uptake kinetics in
BioMed Research International
[59]
[60]
[61]
[62]
[63]
Frankia sp. HFPCcI3 isolated from Casuarina cunninghamiana,”
Canadian Journal of Microbiology, vol. 31, no. 9, pp. 804–809,
1985.
Z. Zhongze, M. A. Murry, and J. G. Torrey, “Culture conditions influencing growth and nitrogen fixation in Frankia sp.
HFPCcI3 isolated from Casuarina,” Plant and Soil, vol. 91, no. 1,
pp. 3–15, 1986.
K. H. Richau, R. L. Kudahettige, P. Pujic, N. P. Kudahettige,
and A. Sellstedt, “Structural and gene expression analyses of
uptake hydrogenases and other proteins involved in nitrogenase
protection in Frankia,” Journal of Biosciences, vol. 38, no. 4, pp.
703–712, 2013.
R. Parsons, W. B. Silvester, S. Harris, W. T. Gruijters, and
S. Bullivant, “Frankia vesicles provide inducible and absolute
oxygen protection for nitrogenase,” Plant Physiology, vol. 83, no.
4, pp. 728–731, 1987.
R. Nalin, S. R. Putra, A.-M. Domenach, M. Rohmer, F. Gourbiere, and A. M. Berry, “High hopanoid/total lipids ratio in
Frankia mycelia is not related to the nitrogen status,” Microbiology, vol. 146, no. 11, pp. 3013–3019, 2000.
D. B. Steele and M. D. Stowers, “Superoxide dismutase and
catalase in Frankia,” Canadian Journal of Microbiology, vol. 32,
no. 5, pp. 409–413, 1986.
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 317524, 11 pages
http://dx.doi.org/10.1155/2014/317524
Research Article
Screening for Genes Coding for Putative Antitumor
Compounds, Antimicrobial and Enzymatic Activities from
Haloalkalitolerant and Haloalkaliphilic Bacteria Strains of
Algerian Sahara Soils
Okba Selama,1 Gregory C. A. Amos,2 Zahia Djenane,1 Chiara Borsetto,2 Rabah Forar Laidi,3
David Porter,2 Farida Nateche,1 Elizabeth M. H. Wellington,2 and Hocine Hacène1
1
Microbiology Group, Laboratory of Cellular and Molecular Biology, Faculty of Biological Sciences, USTHB, BP 32, EL ALIA,
Bab Ezzouar, Algiers, Algeria
2
School of Life Sciences, University of Warwick, Coventry CV4 7AL, UK
3
Department de Biologie, Ecole Normale Superieure (ENS), Vieux Kouba, Alger, Algeria
Correspondence should be addressed to Hocine Hacène; h [email protected]
Received 26 February 2014; Revised 13 April 2014; Accepted 6 May 2014; Published 27 May 2014
Academic Editor: Ameur Cherif
Copyright © 2014 Okba Selama et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Extreme environments may often contain unusual bacterial groups whose physiology is distinct from those of normal
environments. To satisfy the need for new bioactive pharmaceuticals compounds and enzymes, we report here the isolation of
novel bacteria from an extreme environment. Thirteen selected haloalkalitolerant and haloalkaliphilic bacteria were isolated from
Algerian Sahara Desert soils. These isolates were screened for the presence of genes coding for putative antitumor compounds using
PCR based methods. Enzymatic, antibacterial, and antifungal activities were determined by using cultural dependant methods.
Several of these isolates are typical of desert and alkaline saline soils, but, in addition, we report for the first time the presence of a
potential new member of the genus Nocardia with particular activity against the yeast Saccharomyces cerevisiae. In addition to their
haloalkali character, the presence of genes coding for putative antitumor compounds, combined with the antimicrobial activity
against a broad range of indicator strains and their enzymatic potential, makes them suitable for biotechnology applications.
1. Introduction
There is an increasingly urgent need for new active biomolecules and enzymes for use in industry and therapy [1].
However, the rate of discovery of new useful compounds has
been in decline [2, 3] and because of this there is an interest in
investigating previously unexplored ecological niches [4, 5],
particularly extreme environments. These environments have
provided a useful source of novel biologically active compounds in recent years [1, 6, 7].
Extreme environments are distributed worldwide. These
ecosystems were thought to be lifeless as insurmountable
extreme physical and chemical barriers to life exhibit. With
the advancement of our knowledge, we now see them as yet
another niche harbouring “extremophiles” [8]; major categories of extremophiles include halophiles, thermophiles, acidophiles, alkaliphiles, and haloalkaliphiles [6, 9].
The haloalkaliphiles bacteria have attracted a great deal of
attention from researchers in this last decade [9]. In 1982, the
term haloalkaliphile was used for the first time to describe
bacteria that are both halophilic and alkaliphilic [10]. This
group of bacteria is able to grow optimally or very well at pH
values at or above 10 along with high salinity (up to 25% (w/v)
NaCl) [11].
To encounter such harsh conditions, haloalkaliphilic
microorganisms have found various physiological strategies
to sustain their cell structure and function [12, 13]. These
bacteria have widely been identified and studied from the
2
2. Material and Methods
2.1. Sampling and Strains Isolation. Samples from different
soils (7 sites) of Algeria’s Sahara Desert were collected on
March 2010 (100–300 g per site in sterile bags) (Figure 1).
Most samples were saline and alkaline soils, with an electrical
conductivity between 1.4 and 20.2 mS/cm (at 20∘ C) and pH
range of 7.5–9; the temperature varies from 22∘ C north to
44∘ C south of the Sahara. One gram from each sample was
suspended in 9 ml sterile water (of 0.9%, 10%, and 20% NaCl
w/v) and serial dilutions to 10−4 . For each dilution and for
each concentration, soil particles were allowed to sediment;
then 0.1 mL of the liquid phase was spread onto the surface
of each of the modified International Streptomyces Project 2
(ISP2) [27] media agar supplemented with NaCl with respect
to the various concentrations of salt used for dilutions (0.9%,
10%, and 20% NaCl w/v) and adjusted to either pH= 7 or
pH = 10 by adding 5 M NaOH before autoclaving and spread
onto nutrient agar plates. The plates were maintained at
constant humidity incubated at either 30∘ C or 50∘ C for 15
days. Colonies were picked out and repeatedly restreaked
until purity was confirmed. All bacterial culture isolates were
stored at 4∘ C in the same medium used for isolation.
2.2. Physiological Growth Parameters. Physiological growth
parameters for the thirteen selected strains were determined
by agar plate method on modified ISP2 medium depending
on the modified parameter. Salinity tolerance was examined
for 0, 1, 5, 10, 15, 20, and 25% NaCl w/v. The pH growth range
was investigated between pH 5 and 12 at intervals of 1 pH
unit. The temperatures tested were 4, 10, 15, 20, 25, 30, 37,
40, 42, 45, 55, and 60∘ C. Incubation time was one week for
Actinobacteria and two days for non-Actinobacteria.
Spain
Mediterranean Sea
Atlantic
Ocean
Biskra
Djelfa
Morocco
Tunisia
Bechar
Ouargla
Algerian Sahara
Adrar
Mauritania
hypersaline environments, soda lakes, solar saltern, salt
brines, carbonate springs, and Dead Sea [14]. Their survival
obviously indicated the widespread distribution of such
organisms in natural saline environments [12, 15].
The interest in haloalkaliphilic microorganisms is due not
only to the necessity for understanding the mechanisms of
adaptation to multiple stresses and detecting their diversity,
but also to their possible application in biotechnology [9].
The present work involved the isolation and characterization of new haloalkalitolerant and haloalkaliphilic bacteria able to produce extremozymes and elaborate natural
bioactive compounds effective against pathogenic bacteria
and fungi as well. The screening for genes coding for putative
antitumor compounds by PCR with three sets of primers
was also performed. We have been interested in soils of
Algerian Sahara Desert, which is one of the biggest deserts
and encompasses one of the most extreme environments
worldwide (Sabkha and Chott). However, it is also considered
to be one of the less explored parts. Our team has been
interested in these magnificent ecosystems for many years
and the few studies that have been published have shown
great active biomolecules [16–18], biodiversity of interesting
new taxa [19–24], and enzymes [25, 26].
BioMed Research International
In Salah
Tamanrasset
N
300 (km)
Libya
Mali
Niger
Figure 1: Location of the sampled sites from Algerian Sahara Desert.
Djelfa: 35∘ 16󸀠 47.5󸀠󸀠 N 3∘ 43󸀠 25.4󸀠󸀠 E; Biskra: 34∘ 11󸀠 01.1󸀠󸀠 N 6∘ 07󸀠 21.8󸀠󸀠 E;
Ouargla: 33∘ 29󸀠 28.85 N 5∘ 59󸀠 10.52󸀠󸀠 E; Tamanrasset 23∘ 00󸀠 30.01󸀠󸀠 N
5∘ 13󸀠 33.32󸀠󸀠 E; In Salah: 27∘ 11󸀠 31.60󸀠󸀠 N 2∘ 27󸀠 12.52󸀠󸀠 E; Adrar:
27∘ 44󸀠 48.14󸀠󸀠 N 0∘ 16󸀠 10.21󸀠󸀠 W; Bechar: 30∘ 51󸀠 25.71󸀠󸀠 N 1∘ 59󸀠 58.56󸀠󸀠
W.
2.3. Molecular Study
2.3.1. DNA Extraction. Total genomic DNA from the different selected bacteria for this study was isolated and purified
using Qiagen Blood and Tissue DNA extraction kit (Qiagen,
UK). DNA was eluted in Tris-HCL and its quantity and quality were tested using NanoDrop 2000 (Thermo Scientific).
DNA was stored at −20∘ C until use.
2.3.2. Molecular Identification. The amplification of 16S
rRNA gene for the selected strains was performed using
the universal bacterial primer pairs pA/pH designed by
Edwards et al. [28] (Table 1). PCR reactions were performed
in final reaction volume of 50 𝜇L containing 1 𝜇L (10–100 ng)
of DNA template, 25 𝜇L master mix (Promega, Madison,
WI, USA), 1 𝜇L (10 𝜇M) of each primer, and 1 𝜇L of BSA
(10 mg/mL) (Promega, Madison, WI, USA). PCR products
were analyzed on 1.5% (w/v) agarose (Sigma, UK) in 40 mM
Tris-acetate with 1 mM EDTA (TAE) buffer at pH of 8.0
stained with ethidium bromide at 0.5 𝜇g mL−1 . Bands of
the corresponding size were cut out and purified with
gel extraction kit (Qiagen; Venlo, Netherlands) as per the
manufacturer’s instructions.
The nucleotide sequences for the 16S rRNA gene of the
different strains were carried out by GATC Biotech (UK).
The isolates were identified using the EzTaxon-e server
(http://eztaxon-e.ezbiocloud.net/) on the basis of 16S rRNA
sequence data [32].
BioMed Research International
3
Table 1: Primers used in this study.
Primers
Gene
Molecules
pA: AGAGTTTGATCCTGGCTCAG
pH: AAGGAGGTGATCCAGCCGCA
1,5 Kb
16S RNA
///////////
Glu1: CSGGSGSSGCSGGSTTCATSGG
dNDP-Glucose-4,6Glu2: GGGWRCTGGYRSGGSCCGTAGTTG
dehydratases
546 bp
//////
Oxytryptophan
dimerization genes
StaDVF: GTSATGMTSCAGTACCTSTACGC
(StaD/RebD/VioB)
StaDVR: YTCVAGCTGRTAGYCSGGRTG.
(indolotryptoline
570 bp
biosynthetic gene
cluster)
BE-54017,
(tryptophan
dimmers)
AuF3: GAACTGGCCSCGSRTBTT
AuR4: CCNGTGTGSARSKTCATSA
600–700 bp
Iadomycin
cyclase gene of
Streptomyces
venezuelae ISP5230
The Molecular Evolutionary Genetics Analysis (MEGA)
software, version 4.0.2, was used to assist the phylogenetic
analyses and the phylogenetic tree construction [33]. Similar
16S rRNA gene sequences for the studies of the strains were
obtained by using Eztaxon [32]. Multiple alignments of data
were performed by CLUSTAL W [34]. Evolutionary distances
were calculated by using maximum composite likelihood
method and are in the units of the number of base substitutions per site [35]. Phylogenetic tree was reconstructed
with the neighbour-joining algorithm [36]. Topology of the
resultant tree was evaluated by bootstrap analyses of the
neighbour-joining dataset, based on 1000 resamplings [37].
The sequences reported in this study have been submitted
to NCBI GenBank and the accession numbers are listed in
appendices.
2.4. Screening
2.4.1. Primers and Molecular Screening. From the thirteen
selected strains, six were subjected to molecular screening for
genes coding for putative antitumor compounds using three
primer sets (Table 1). These strains were chosen on the basis
of the presence of nonribosomal peptide synthetases/ polyketide synthases (NRPS/PKS) genes within their genomes
(data not published). The first set designed by Decker et al.
[29] amplified dNDP-glucose dehydratase genes. The second
set was that of Chang and Brady [30] used to screen for
biosynthesis of the antitumor substance BE-54017. The final
set was used from the study of Ouyang et al. [31] targeting
the jadomycin cyclase gene which intervenes in angucycline
production.
Angucycline
cyclases
Marine
sponge
Reference PCR programs
PCR cycles were as follows: 1 cycle at
95∘ C for 10 min; 35 cycles at 94∘ C for
[28]
1 min, 55∘ C for 1 min, and 72∘ C for
2 min; one final cycle at 72∘ C for
10 min.
PCR conditions used were 95∘ C for
4 min; 30 cycles of 95∘ C for 30 s, 65∘ C
[29]
for 30 s, and 68∘ C for 1.30 min; and a
final extension cycle at 68∘ C for 5 min.
PCR protocol: 1 cycle of 95∘ C for
5 min; 7 cycles of 95∘ C for 30 sec, 65∘ C
for 30 sec with 1∘ C decrement per cycle
[30]
to 59∘ C, and 72∘ C for 40 sec; 30 cycles
of 95∘ C for 30 sec, 58∘ C for 30 sec, and
72∘ C for 40 sec; 1 cycle of 72∘ C for
7 min; hold at 4∘ C
Optimized PCR conditions were as
follows: (1) denaturation at 94∘ C for
5 min, (2) 30 amplification cycles with
[31]
denaturation (45 s, 94∘ C), annealing
(60 s, 60∘ C), and extension (60 s,
72∘ C), and (3) a final extension at 72∘ C
for 8 min.
The PCR mixture included 1-2 𝜇L of genomic DNA,
15 𝜇L master mix (Sigma,UK), 1 𝜇L each of forward and
reverse primers (10 𝜇M each) (Sigma, UK), 1 𝜇L of BSA
(10 mg/mL) (Promega, Madison, WI, USA), and 6 𝜇L sterile
distilled water in a final volume of 25 𝜇L. PCR was performed
with Mastercycler pro (Eppendorf). Agarose gels (1% w/v)
were photographed after staining with ethidium bromide at
0.5 𝜇g mL−1 with a minivisionary imaging system. Sizes of the
fragments were estimated using the Fermentas 1 kb Plus DNA
ladder (Fermentas, UK).
2.4.2. Antimicrobial Activities Test. Antimicrobial activity
was determined by the agar cylinder diffusion method.
A 6 mm diameter cylinder was taken from solid cultures
and put on preseeded nutrient agar plate of the targeted
microorganisms mentioned below. Up to five cylinders of
different bacteria per plate were tested. Inhibition zones
were expressed as diameter and measured after incubation
at 37∘ C for 24 h for bacteria and at 28∘ C for 48–72 h for the
filamentous fungus and yeasts [38].
Reference strains used in this study were as follows.
Sa: Staphylococcus aureus ATCC 25923, Ml: Micrococcus
luteus ATCC 9341, Ec: Escherichia coli ATCC 25922, Pa:
Pseudomonas aeruginosa ATCC 27853, Ca: Candida albicans
(clinical isolate, Algerian Central Hospital of Army of Algeria), Foa: Fusarium oxysporum f. sp. albedinis a filamentous
phytopathogenic fungi for date palm (Algerian National
Institute for Plant Protection), and Sc: Saccharomyces cerevisiae.
2.4.3. Enzymatic Screening. Enzymatic activities “amylolytic,
proteolytic (caseinase), and lipolytic” were screened using
4
zone clearance assays. The enzymatic substrate was incorporated to the media, and the strains were restreaked by spots
[39]. The tests were conducted with respect to physiological
growth parameters of each strain.
3. Results
3.1. Strains Isolation and Selection. Isolation plates developed
various types of colonies. Sixty to hundred colonies were
found per plate in the first dilution for almost all soils, two to
ten colonies were observed in the third dilution, and almost
nothing in the fourth dilution plates. We have also seen that
for the same dilution the number of colonies decreases when
the concentration of NaCl increases. One to five colonies
which looked less represented were selected from each plate
with respect to the haloalkaliphilic character. A total of thirtynine isolates were distinguished. Amongst these thirty-nine
isolates (17 were filamentous, 17 bacilli form, and 5 were cocci
form), thirteen strains—eleven with particular morphology
(filamentous, which may indicate Actinobacteria that are best
known for the production of active biomolecules), one bacilli
form, and one cocci form—were the subject of our study.
The macroscopic and microscopic aspects of three of the
thirteen strains are represented in Figure 2. The molecular
identification by EzTaxon-e, physiological growth parameters, and enzymatic screening are described in Table 2.
The alphabetical strains code used in our study refers to the
geographical area origin of isolation; the numerical strains
code part is a simple sequential order to differentiate strains.
3.2. Physiological Growth Study. All strains could tolerate up
to 5% NaCl. Strains Reg1, Ker5, and HHS1 were able to tolerate
up to 10%, whereas Bisk4 could tolerate up to 15%. Tag5
growth started at 1% and M5A started growing at 10%; these
two strains could grow up to 20% NaCl. Reg1, Ker5, and HHS1
are considered as halotolerant. M5A and Tag5 are considered
to be halophilic [40].
All strains except A60 had a versatile range of growth
pH (5–12) indicating alkaliphilic growth; A60 (5–9) was only
alkalitolerant.
Beside the alkalitolerant character of strain A60, it presented a thermophilic profile (45–60∘ C). With the exception
of strain Bisk4, which may be considered as thermotolerant
bacteria since it grows up to 55∘ C, the other selected bacteria
are considered to be mesophiles.
3.3. Identification. Most isolated strains belonged to the
genus Streptomyces (AT1, ASB, GB1, Ig6, and GB3). The five
Actinobacteria, other than Streptomyces, were identified as
follows: Reg1 and Ker5 as two different Nocardiopsis sp., HHS1
as Pseudonocardia sp., M5A as Actinopolyspora sp., and Bisk2
as Nocardia sp. Bisk2 looks like a new member as it branches
out 100% of the time from its nearest relative Nocardia
jejuensis determined by EzTaxon-e with 95% similarity for
the 750 recovered bases. One filamentous strain A60 was
identified as Thermoactinomyces sp. The bacilli Bisk4 is part of
the Bacillus mojavensis complex and the cocci Tag5 belonged
to the genus Marinococcus (Table 2; Figure 3).
BioMed Research International
3.4. Screening for Biotechnological Potential
3.4.1. Screening for Genes Coding for Putative Antitumor
Compounds. Glu1/Glu2 primer set had 4/6 positives. High
intensity band was registered for the strain Ig6. The primers
targeted two different regions for the strain Bisk2. Multiple
bands were recovered from the strain GB1 while no one
range 500–700 pb. PCR using this primer was negative for
the strain A60 (Figure 4(a)). The StaDVF/StaDVR primer
set was positive in one strain (Figure 4(b)). The PCR with
AuF3/AuF4 primer set was negative for all tested strains
(Figure 4(c)).
3.4.2. Antimicrobial Activity. The antimicrobial activity of the
thirteen selected strains differed between strains (Table 2;
Figure 5). Among these, eight showed at least an antimicrobial activity against one of the targeted microorganisms.
A highly broad spectrum antimicrobial activity inhibition
was seen by the strain Streptomyces sp. (GB3). The strain
Bacillus sp. (Bisk4) had gram positive antibacterial activity
and antifungal activity against the filamentous fungi. The
strain Actinopolyspora sp. (M5A) inhibited the growth of
Micrococcus luteus. The isolate Nocardia sp. (Bisk2) showed a
unique and selective activity against the yeast Saccharomyces
cerevisiae (Figure 5(c)). However, none of the thirteen strains
demonstrate specific and unique activity against the gram
negative bacteria.
3.4.3. Enzymatic Activity. Strains from Bacillus and Streptomyces were more enzymatically active and possess at least two
of the screened enzymes. The strain Thermoactinomyces sp.
(A60) was able to degrade casein and lipids. Strains Bisk2,
TAG5, and HHS1 seemed to have none of these screened
enzymes (Figure 6).
4. Discussion
In this study we looked at extreme environment of the
Algerian Sahara Desert as a source for novel strains possessing interesting bioactive properties. In total, we isolated
a collection of thirty-nine haloalkalitolerant and haloalkaliphilic isolates, thirteen of which were selected and
screened for genes coding for putative antitumor compounds, as well as screening for antimicrobial and enzymatic activities. All strains were identified using 16S rRNA
gene sequencing. This study represents novelty in looking
at the relatively understudied areas of Sabkha and Chott
and has yielded at least thirteen strains which potentially have antitumorgenic, antimicrobial, and enzymatic
properties.
Although often extreme and hostile ecosystems diversity
and abundance of bacteria can be low ranging from 10 to
104 UFC/g of soil where the physicochemical parameters are
controlling factors [19], the strains retrieved and identified
in our study, in particular, of Actinobacteria strains, which
belong to various taxa, indicate a great diversity. Diversity in
environments such as the one in this study has previously
been investigated such as in Tunisia [9], China [41], and
5–12
5–12
5–12
5–9
5–12
5–12
5–12
5–12
5–12
5–12
5–12
5–12
0–5
10–20
0–12
0–5
0–10
0–10
1–20
0–10
0–5
0–5
0–7
0–7
Bisk2
M5A
HHS1
A60
AT1
Reg1
Tag5
Ker5
IG6
ASB
GB1
GB3
20–37
20–37
20–42
20–40
20–42
10–42
25–42
20–42
45–60
25–30
30–40
4–42
20–55
+: positive activity, −: negative activity, and N: not tested.
5–12
0–15
Growth parameters
Salinity interval
pH
Temperature
(% g/L)
interval interval (∘ C)
Bisk4
Strains
Enzymatic activity
Antimicrobial activity
−
−
N
N
−
−
N
−
N
−
−
N
N
N
N
N
−
N
N
N
N
+
N
N
N
N
−
−
N
N
N
+
+
−
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
−
+
+
−
−
−
−
−
−
−
−
+
+
−
−
−
−
+
−
+
+
−
+
−
−
+
−
+
+
−
−
−
+
−
+
−
−
N
+
+
+
+
+
+
+
−
−
−
+
−
−
+
−
+
+
+
−
−
−
−
−
−
−
−
+
−
−
−
−
−
−
−
−
−
+
−
−
−
−
−
+
+
−
−
−
−
−
−
−
−
−
−
+
+
+
−
+
−
−
−
−
−
−
−
−
−
+
+
−
+
−
−
−
−
−
−
−
+
−
Glu StaDV AuF Proteolytic Amylolytic Lipolytic Sa Ml Pa Ec Foa Ca Sc
Antitumor genes
Bacillus tequilensis 10b(T)
(Bacillus mojavensis group)
Nocardia jejuensis N3-2(T)
Actinopolyspora dayingensis TRM
4064(T)
Pseudonocardia ammonioxydans
H9(T)
Thermoactinomyces vulgaris
KCTC 9076(T)
Streptomyces mutabilis NBRC
12800(T)
Nocardiopsis dassonvillei subsp.
albirubida DSM 40465(T)
Marinococcus halophilus DSM
20408(T)
Nocardiopsis dassonvillei subsp.
albirubida DSM 40465(T)
Streptomyces sparsus YIM
90018(T)
Streptomyces pilosus NBRC
12807(T) (Streptomyces pilosus
group)
Streptomyces celluloflavus NBRC
13780(T)
Streptomyces cyaneofuscatus JCM
4364(T)
Most related species
Table 2: Physiologic characterization, antitumoral genes, enzymatic activity, antimicrobial activity, and most related species of the thirteen selected strains of this study.
BioMed Research International
5
6
BioMed Research International
Gx400
Gx200
(a)
Gx300
Gx200
(b)
Gx200
Gx400
(c)
Figure 2: Macroscopic morphology (left) on ISP2 and microscopic filamentous morphology (right) of three strains of this study. (a) Strain
IG6: spiral chain of spores on aerial mycelium. (b) Strain Bisk2: nocardioform mycelium. (c) Strain M5A: long straight chains of spores on
aerial mycelium.
previously in the Algerian Sahara soils [19, 42], which
has revealed that members of these extreme ecosystems
are mainly halotolerant or halophilic organisms. Many of
the isolated taxa in this study have previously been found
in this environment, particularly of the Actinopolyspora,
Nocardiopsis, and Marinococcus [9, 41–43]. Despite this
their community structure differs both quantitatively and
qualitatively for each different ecosystem. This would be
due not only to the adaptation to environmental obstacles
but also to the geolocalisation [43], the difference of the
study protocol (method, media) [41], and the sampling sites
[42].
Genome sequencing followed by bioinformatics analysis
for some of the already sequenced microorganisms such
as Actinobacteria and Bacillus has revealed the presence of
several gene clusters per genome that can produce different
molecules [44]. Among the validly described halotolerant
and halophilic bacteria, particularly Actinobacteria, only few
numbers have been subjected to analysis of their bioactive
compounds [45]. In addition, many compounds are usually
produced in very low amounts (or not at all) under typical
laboratory conditions [46]. PCR based methods for specific
enzymes activating specific molecules are excellent screening
tools for these strains; they would not only indicate the
BioMed Research International
7
Strain GB3 (JQ690542)
68
72 Streptomyces cyaneofuscatus; JCM 4364 (AY999770)
54
Strain GB1 (JQ690543)
41 Streptomyces cellulofavus; NBRC 13780 (AB184476)
70
Strain Ig6 (JQ690545)
Streptomyces sparsus; YIM 90018 (AJ849545)
Strain ASB (JQ690544)
Streptomyces mutabilis; NBRC 12800 (AB184156)
95
Strain AT1 (JQ690546)
71
100
Streptomyces pilosus; NBRC 12807 (AB184161)
Strain Ker5 (JQ690548)
100
Strain Reg1 (JQ690549)
Nocardiopsis dassonvillei subsp. albirubida; DSM 40465 (X97882)
Strain M5A (KJ409655)
100
52
Actinopolyspora dayingensis; TRM 4064 (KC461229)
100 Strain HHS1 (JQ690547)
78
Pseudonocardia ammonioxydans; H9 (AY500143)
Strain Bisk2 (JQ690551)
57
98 Nocardia jejuensis; N3-2 (AY964666)
Strain Bisk4 (JQ690553)
100
Bacillus tequilensis; 10b (HQ223107)
Strain A60 (JQ690550)
99
Termoactinomyces vulgaris; KCTC 9076 (AF138739)
Strain Tag5 (JQ690552)
100 Marinococcus halophilus; DSM 20408 (X90835)
Escherichia coli strain KCTC 2441 (EU014689)
98
99
57
0.02
700
500
700
A60
M5A
Bisk4
GB1
Ig6
Bisk2
W
M
W
A60
M5A
Bisk4
Ig6
GB1
Bisk2
M
W
A60
M5A
Bisk4
Ig6
GB1
Bisk2
M
Figure 3: Molecular phylogeny of thirteen selected bacteria and the most related type strains species using partial 16S rRNA sequences.
The evolutionary distances were computed using the maximum composite likelihood method and are in the units of the number of base
substitutions per site. Tree topology was constructed using MEGA 4.0. Bootstrap values (𝑛 = 1000 replicates) were indicated at the nodes.
Escherichia coli KCTC2441 sequence was added as an out group for this tree.
700
500
500
(a)
(b)
(c)
Figure 4: Agarose gel electrophoresis of PCR products from genomic DNA of six strains of the present study with selective fragments
amplification range 500–700 bp using primers: (a) Glu1/Glu2, (b) StaDVF/StaDVR, and (c) AuF3/AuF4. M: 1 kb Plus DNA ladder; W: water
control.
8
BioMed Research International
ASB
GB3
Bisk4
Bisk2
Bisk4
GB3
Foa
Bisk2
(a)
(b)
(c)
Figure 5: Antimicrobial activity of some strains among the selected strains: (a) antibacterial activity against Staphylococcus aureus, (b)
antifungal activity against Fusarium oxysporum f. sp. albedinis, and (c) antifungal activity of the strain Nocardia sp. (Bisk2) against the yeast
Saccharomyces cerevisiae.
ASB
AT1
A60
Ig6
ASB
Ker5
GB3
Reg1
(a)
(b)
(c)
Figure 6: Enzymatic activities of some strains among the selected strains. (a) Proteases (caseinase), (b) lipases, and (c) amylases.
presence of probable genes clusters but also help in biochemical characterisation of the molecules. These methods
would help in reducing the number of strains that need to
be screened by cultural methods. The PCR based methods
not only are limited to genomic DNA but also can be applied
for the screening of eDNA that lead to the discovery of new
active biomolecules [30]. Screening for potential production
of a particular type of biomolecules such as antibiotics and
antitumorales, without going through the tedious biochemistry process, is more efficient when the typing protocol
is targeting the biosynthesis gene cluster rather than the
taxonomic marker genes (e.g., 16S rRNA gene) which often
give misleading results [47, 48].
In our study, we have been interested in molecular
screening of bioactive genes coding for putative antitumor compounds. The degenerate primers Glu1/Glu2 for
the conserved N-terminal sequence of dNDP-glucose 4,6dehydratase genes have been extensively used to screen
out for clusters of active biomolecules with antitumoral
activity such as novobiocin [49], enediyne [50], elloramycin
[51], sibiromycin [52], ravidomycin, and chrysomycin [53].
The primer set has also been reported in other screening
studies for talosins A and B cluster, an antifungal [54], for
caprazamycin biosynthesis, an antimycobacterial [55], and
more recently we have used this set to screen for amicetin
biosynthesis gene cluster, an antibacterial and antiviral agent
[56]. The second primer set was designed by Chang and
Brady [30] who screened a previously archived soil eDNA
cosmid library by PCR using degenerate primers designed
to recognize conserved regions in known oxytryptophan
dimerization genes (StaD/RebD/VioB etc). The oxytryptophan dimerization enzymes were chosen as probes because
this enzyme family is used in the biosynthesis of structurally
diverse tryptophan dimmers, which have shown an antitumoral activity. Both indolocarbazole biosynthetic gene clusters (e.g., staurosporine, rebeccamycin, K-252a, and AT2433)
and violacein biosynthetic gene clusters contain homologous
enzymes that carry out the oxidation (StaO/RebO/VioA) and
subsequent dimerization (StaD/RebD/VioB) of tryptophan.
One among the six screened strains was positive for the set
of the primers, strain M5A. This would signify that the strain
M5A could produce tryptophan dimmers compound(s).
BioMed Research International
The sequencing result followed by blast for the PCR products of M5A using StaDVF/StaDVR primers set (GenBank:
KJ560370) has shown 76% homology to the uncultured bacterium clone AR1455 rebeccamycin-like tryptophan dimer
gene cluster (GenBank: KF551872) that was studied by Chang
and Brady [30], while, for the strains Streptomyces sp. Ig6, it
has shown a mixed PCR product; we think this is probably
due to the presence of multiple variable copies of this gene in
this strain.
The different patterns of activity against the targeted
microorganisms observed in this study may indicate a variety of the produced active biomolecules. The antimicrobial
activity of Bisk2, most closely related to Nocardia jejuensis
[57], has never been reported to our knowledge. This result
encourages us to consider Bisk2 as probably a new member or
at least a new strain of Nocardia. Genome sequencing, DNADNA hybridising, and molecular chemotaxonomy would
give more knowledge about its taxonomic position among the
Nocardia species.
The Sahara Desert is subject to large fluctuations in
parameters such as temperature, pH, or salinity. It is populated by communities of organisms with intrinsic genomic
heterogeneity for adaptation. The mechanisms of cell adaptation engage several enzymatic processes that may be a source
of enzymes that show a higher level of stability and activity
over a wider range of conditions. The screened enzymes
found in this study (proteases, amylases, and lipases) would
be economically valuable since they were screened from such
environments and are likely to exhibit rare properties; these
extremozymes are of great value to biotechnology industries
[7, 58, 59].
5. Conclusion
Exploration of biodiversity and biotechnological potential
of desert microorganisms has gone several steps forward
in recent years. The Sahara Desert is one of the biggest
worldwide. It spreads upon several countries of Africa.
These countries are among the countries worldwide to have
the smallest registration rates of biodiversity in biological
databases [60].
In addition to the insights on the biodiversity of Algerian
Sahara Desert, to our knowledge, this is the first time to use
the molecular screening of these genes coding for putative
antitumor compounds to analyse Algerian strains. In this
study, we have highlighted the interesting presence of diverse
haloalkalitolerant and haloalkaliphilic strains with potential
antitumorigenic, bioactive, and other interesting enzymes.
Future work will concentrate on more cloning and sequencing for whole clusters, chemical characteristics, identification
by application of mass spectrum, and other enzymatic and
biochemical techniques that would be more suitable for better
determination of the nature of the elaborated compounds
produced by the strains identified in this study particularly of Nocardia sp. Bisk2, Actinopolyspora sp. M5A, and
Streptomyces sp. Ig6.
9
Appendix
GenBank accession numbers for 16S RNA gene sequences of
13 strains of this study are GB3 (JQ690542), GB1 (JQ690543),
ASB (JQ690544), Ig6 (JQ690545), AT1 (JQ690546), HHS1
(JQ690547), Ker5 (JQ690548), Reg1 (JQ690549), A60
(JQ690550), Bisk2 (JQ690551), Tag5 (JQ690552), Bisk4
(JQ690553), and M5A (KJ409655).
Conflict of Interests
The authors declare that there is no conflict of interests
regarding the publication of this paper.
Acknowledgments
The authors acknowledge Warwick University staff, in particular Dr. Calvo-Bado A. L., Dr. Khalifa A., and Dr. Witcomb L. The authors also acknowledge Professor Naim M.
from HCA, Dr. Antri K. USTHB for providing the targeted
microorganisms, and Mr. Bouhzila F. from environmental
Biotechnology, Polytechnical School, Algiers, for physicalchemical soils parameters determination. The authors give
special thanks to Mr. Mohammed A., Mr. Slama G., and Mr.
Natèche M. for the help. The authors would also like to thank
the anonymous reviewers for the analysis and the enrichment
of this paper. In the end, the authors would like to thank
the Algerian Ministry of Higher Education and Scientific
Research and the University of Warwick for supporting this
work. CB has received funding from the European Union’s
Seventh Framework Programme for research, technological
development, and demonstration under Grant no. 289285.
References
[1] M. Podar and A. L. Reysenbach, “New opportunities revealed
by biotechnological explorations of extremophiles,” Current
Opinion in Biotechnology, vol. 17, no. 3, pp. 250–255, 2006.
[2] S. Subramaniam, “Productivity and attrition: key challenges for
biotech and pharma,” Drug Discovery Today, vol. 8, no. 12, pp.
513–515, 2003.
[3] K. S. Lam, “New aspects of natural products in drug discovery,”
Trends in Microbiology, vol. 15, no. 6, pp. 279–289, 2007.
[4] N. Perić-Concha and P. F. Long, “Mining the microbial
metabolome: a new frontier for natural product lead discovery,”
Drug Discovery Today, vol. 8, no. 23, pp. 1078–1084, 2003.
[5] J. Spı́žek, J. Novotná, T. Řezanka, and A. L. Demain, “Do we
need new antibiotics? The search for new targets and new compounds,” Journal of Industrial Microbiology and Biotechnology,
vol. 37, no. 12, pp. 1241–1248, 2010.
[6] F. A. Rainey and A. Oren, “Extremophile microorganisms and
the methods to handle them,” in Extremophiles Methods in
Microbiology, F. A. Rainey and A. Oren, Eds., vol. 35, pp. 1–25,
Academic Press, London, UK, 2006.
[7] Y. Ma, E. A. Galinski, W. D. Grant, A. Oren, and A. Ventosa,
“Halophiles 2010: life in saline environments,” Applied and
Environmental Microbiology, vol. 76, no. 21, pp. 6971–6981, 2010.
[8] L. J. Rothschild and R. L. Mancinelli, “Life in extreme environments,” Nature, vol. 409, no. 6823, pp. 1092–1101, 2001.
10
[9] D. El Hidri, A. Guesmi, A. Najjari et al., “Cultivation-dependant
assessment, diversity, and ecology of haloalkaliphilic bacteria
in arid saline systems of southern Tunisia,” BioMed Research
International, vol. 2013, Article ID 648141, 15 pages, 2013.
[10] G. S. H. Soliman and H. G. Trueper, “Halobacterium pharaonis
sp.nov., a new, extremely haloalkaliphilic archaebacterium with
low magnesium requirement,” Zentralblatt für Bakteriologie
Mikrobiologie und Hygiene: I. Abt. Originale C, vol. 3, no. 2, pp.
318–329, 1982.
[11] K. Horikoshi, Alkaliphiles, Wiley Online Library, 2008.
[12] K. Horikoshi, “Alkaliphiles: some applications of their products for biotechnology,” Microbiology and Molecular Biology
Reviews, vol. 63, no. 4, pp. 735–750, 1999.
[13] H. Takami, Y. Takaki, and I. Uchiyama, “Genome sequence
of Oceanobacillus iheyensis isolated from the Iheya Ridge and
its unexpected adaptive capabilities to extreme environments,”
Nucleic Acids Research, vol. 30, no. 18, pp. 3927–3935, 2002.
[14] S. P. Singh, “Extreme environments and extremophiles,” in EBook: Environmental Microbiology, pp. 1–35, National Science
Digital Library (CSIR), New Delhi, India, 2006.
[15] A. A. Joshi, P. P. Kanekar, A. S. Kelkar et al., “Cultivable bacterial
diversity of alkaline Lonar lake, India,” Microbial Ecology, vol.
55, no. 2, pp. 163–172, 2008.
[16] H. Hacène, N. Sabaou, N. Bounaga, and G. Lefebvre, “Screening
for non-polyenic antifungal antibiotics produced by rare Actinomycetales,” Microbios, vol. 79, no. 319, pp. 81–85, 1994.
[17] H. Hacene, K. Kebir, D. S. Othmane, and G. Lefebvre, “HM17, a
new polyene antifungal antibiotic produced by a new strain of
Spirillospora,” Journal of Applied Bacteriology, vol. 77, no. 5, pp.
484–489, 1994.
[18] H. Hacène, F. Daoudi-Hamdad, T. Bhatnagar, J. C. Baratti, and
G. Lefebvre, “H107, a new aminoglycoside anti-Pseudomonas
antibiotic produced by a new strain of Spirillospora,” Microbios,
vol. 102, no. 402, pp. 69–77, 2000.
[19] H. Hacěne, F. Rafa, N. Chebhouni et al., “Biodiversity of
prokaryotic microflora in El Golea Salt lake, Algerian Sahara,”
Journal of Arid Environments, vol. 58, no. 3, pp. 273–284, 2004.
[20] S. Boutaiba, H. Hacene, K. A. Bidle, and J. A. Maupin-Furlow,
“Microbial diversity of the hypersaline Sidi Ameur and Himalatt
Salt Lakes of the Algerian Sahara,” Journal of Arid Environments,
vol. 75, no. 10, pp. 909–916, 2011.
[21] A. Bouanane-Darenfed, M. L. Fardeau, P. Grégoire et al.,
“Caldicoprobacter algeriensis sp. nov. a new thermophilic anaerobic, xylanolytic bacterium isolated from an Algerian hot
spring,” Current Microbiology, vol. 62, no. 3, pp. 826–832, 2011.
[22] A. N. Addou, P. Schumann, C. Spröer, H. Hacene, J. L. Cayol,
and M. L. Fardeau, “Melghirimyces algeriensis gen. nov., sp. nov.,
a member of the family Thermoactinomycetaceae, isolated from
a salt lake,” International Journal of Systematic and Evolutionary
Microbiology, vol. 62, part 7, pp. 1491–1498, 2012.
[23] A. N. Addou, P. Schumann, C. Spröer et al., “Melghirimyces
thermohalophilus sp. nov., a thermoactinomycete isolated from
an Algerian salt lake,” International Journal of Systematic and
Evolutionary Microbiology, vol. 63, no. part 5, pp. 1717–1722,
2013.
[24] M. Amziane, F. Metiaz, A. Darenfed-Bouanane et al., “Virgibacillus natechei sp. nov., a moderately halophilic bacterium
isolated from sediment of a saline lake in southwest of Algeria,”
Current Microbiology, vol. 66, no. 5, pp. 462–466, 2013.
[25] T. Bhatnagar, S. Boutaiba, H. Hacene et al., “Lipolytic activity
from Halobacteria: screening and hydrolase production,” FEMS
Microbiology Letters, vol. 248, no. 2, pp. 133–140, 2005.
BioMed Research International
[26] S. Boutaiba, T. Bhatnagar, H. Hacene, D. A. Mitchell, and J. C.
Baratti, “Preliminary characterisation of a lipolytic activity from
an extremely halophilic archaeon, Natronococcus sp.,” Journal
of Molecular Catalysis B: Enzymatic, vol. 41, no. 1-2, pp. 21–26,
2006.
[27] E. B. Shirling and D. Gottlieb, “Methods for characterization
of Streptomyces species,” International Journal of Systematic
Bacteriology, vol. 16, pp. 313–340, 1966.
[28] U. Edwards, T. Rogall, H. Blocker, M. Emde, and E. C. Bottger,
“Isolation and direct complete nucleotide determination of
entire genes. Characterization of a gene coding for 16S ribosomal RNA,” Nucleic Acids Research, vol. 17, no. 19, pp. 7843–7853,
1989.
[29] H. Decker, S. Gaisser, S. Pelzer et al., “A general approach for
cloning and characterizing dNDP-glucose dehydratase genes
from actinomycetes,” FEMS Microbiology Letters, vol. 141, pp.
195–201, 1996.
[30] F. Y. Chang and S. F. Brady, “Cloning and characterization of
an environmental DNA-derived gene cluster that encodes the
biosynthesis of the antitumor substance BE-54017,” Journal of
the American Chemical Society, vol. 133, no. 26, pp. 9996–9999,
2011.
[31] Y. Ouyang, H. Wu, L. Xie et al., “A method to type the potential
angucycline producers in actinomycetes isolated from marine
sponges,” International Journal of Sciences: Basic and Applied
Research, vol. 99, no. 4, pp. 807–815, 2011.
[32] O. S. Kim, Y. J. Cho, K. Lee et al., “Introducing EzTaxon-e: a
prokaryotic 16S rRNA Gene sequence database with phylotypes
that represent uncultured species,” International Journal of
Systematic and Evolutionary Microbiology, vol. 62, part 3, pp.
716–721, 2012.
[33] K. Tamura, J. Dudley, M. Nei, and S. Kumar, “MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version
4.0,” Molecular Biology and Evolution, vol. 24, no. 8, pp. 1596–
1599, 2007.
[34] J. D. Thompson, D. G. Higgins, and T. J. Gibson, “CLUSTAL
W: improving the sensitivity of progressive multiple sequence
alignment through sequence weighting, position-specific gap
penalties and weight matrix choice,” Nucleic Acids Research, vol.
22, no. 22, pp. 4673–4680, 1994.
[35] K. Tamura, M. Nei, and S. Kumar, “Prospects for inferring
very large phylogenies by using the neighbor-joining method,”
Proceedings of the National Academy of Sciences of the United
States of America, vol. 101, no. 30, pp. 11030–11035, 2004.
[36] N. Saitou and M. Nei, “The neighbor-joining method: a new
method for reconstructing phylogenetic trees,” Molecular Biology and Evolution, vol. 4, no. 4, pp. 406–425, 1987.
[37] J. Felsenstein, “Confidence limits on phylogenies: an approach
using the bootstrap,” Evolution, vol. 39, pp. 783–791, 1985.
[38] R. Forar-Laidi, A. Abderrahmane, and A. A. Hocine-Norya,
“Identification and antibiosis of a novel actinomycete strain
RAF-11 isolated from Iraqi soil,” International Journal of Sciences: Basic and Applied Research, vol. 12, no. 1, pp. 141–159, 2013.
[39] E. Phillips and P. Nash, “Culture media,” in Manual of Clinical
Microbiology, E. H. Lennette, A. Balows, W. J. Hausler Jr.,
and H. J. Shadomy, Eds., pp. 1064–1065, American Society for
Microbiology, Washington, DC, USA, 4th edition, 1985.
[40] A. Ventosa, E. Mellado, C. Sanchez-Porro, and M. C. Marquez,
“Halophilic and halotolerant micro-organisms from soils,” in
Microbiology of Extreme Soils, Soil Biology, P. Dion and C. S.
Nautiyal, Eds., vol. 13, pp. 87–115, Springer, Berlin, Germany,
2008.
BioMed Research International
[41] J. Wu, T. Guan, H. Jiang et al., “Diversity of actinobacterial
community in saline sediments from Yunnan and Xinjiang,
China,” Extremophiles, vol. 13, no. 4, pp. 623–632, 2009.
[42] A. Meklat, N. Sabaou, A. Zitouni, F. Mathieu, and A. Lebrihi,
“Isolation, taxonomy, and antagonistic properties of halophilic
actinomycetes in saharan soils of algeria,” Applied and Environmental Microbiology, vol. 77, no. 18, pp. 6710–6714, 2011.
[43] E. Pagaling, H. Wang, M. Venables et al., “Microbial biogeography of six salt lakes in Inner Mongolia, China, and a salt lake
in Argentina,” Applied and Environmental Microbiology, vol. 75,
no. 18, pp. 5750–5760, 2009.
[44] “Genomes OnLine Database,” http://genomesonline.org/cgibin/GOLD/index.cgi.
[45] J. Hamedi, F. Mohammadipanah, and A. Ventosa, “Systematic
and biotechnological aspects of halophilic and halotolerant
actinomycetes,” Extremophiles, vol. 17, pp. 1–13, 2013.
[46] A. K. Chaudhary, D. Dhakal, and J. K. Sohng, “An insight into
the “-omics” based engineering of streptomycetes for secondary
metabolite overproduction,” BioMed Research International,
vol. 2013, Article ID 968518, 15 pages, 2013.
[47] M. Metsä-Ketelä, L. Halo, E. Munukka, J. Hakala, P. Mäntsäla,
and K. Ylihonko, “Molecular evolution of aromatic polyketides
and comparative sequence analysis of polyketide ketosynthase
and 16S ribosomal DNA genes from various Streptomyces
species,” Applied and Environmental Microbiology, vol. 68, no.
9, pp. 4472–4479, 2002.
[48] C. P. Ridley, Y. L. Ho, and C. Khosla, “Evolution of polyketide
synthases in bacteria,” Proceedings of the National Academy of
Sciences of the United States of America, vol. 105, no. 12, pp. 4595–
4600, 2008.
[49] M. Steffensky, A. Mühlenweg, Z. X. Wang, S. M. Li, and
L. Heide, “Identification of the novobiocin biosynthetic gene
cluster of Streptomyces spheroides NCIB 11891,” Antimicrobial
Agents and Chemotherapy, vol. 44, no. 5, pp. 1214–1222, 2000.
[50] W. Liu and B. Shen, “Genes for production of the enediyne
antitumor antibiotic C-1027 in Streptomyces globisporus are
clustered with the cagA gene that encodes the C-1027 apoprotein,” Antimicrobial Agents and Chemotherapy, vol. 44, no. 2, pp.
382–392, 2000.
[51] A. Ramos, F. Lombó, A. F. Braña, J. Rohr, C. Méndez, and J.
A. Salas, “Biosynthesis of elloramycin in Streptomyces olivaceus
requires glycosylation by enzymes encoded outside the aglycon
cluster,” Microbiology, vol. 154, no. 3, pp. 781–788, 2008.
[52] W. Li, A. Khullar, S. Chou, A. Sacramo, and B. Gerratana,
“Biosynthesis of sibiromycin, a potent antitumor antibiotic,”
Applied and Environmental Microbiology, vol. 75, no. 9, pp.
2869–2878, 2009.
[53] M. K. Kharel, S. E. Nybo, M. D. Shepherd, and J. Rohr, “Cloning
and characterization of the ravidomycin and chrysomycin
biosynthetic gene clusters,” ChemBioChem, vol. 11, no. 4, pp.
523–532, 2010.
[54] T. M. Yoon, J. W. Kim, J. G. Kim, W. G. Kim, and J. W. Suh,
“Talosins A and B: new isoflavonol glycosides with potent
antifungal activity from Kitasatospora kifunensis MJM341 I.
Taxonomy, fermentation, isolation, and biological activities,”
Journal of Antibiotics, vol. 59, no. 10, pp. 633–639, 2006.
[55] L. Kaysser, E. Wemakor, S. Siebenberg et al., “Formation
and attachment of the deoxysugar moiety and assembly of
the gene cluster for caprazamycin biosynthesis,” Applied and
Environmental Microbiology, vol. 76, no. 12, pp. 4008–4018,
2010.
11
[56] G. Zhang, H. Zhang, S. Li et al., “Characterization of the
amicetin biosynthesis gene cluster from Streptomyces vinaceusdrappus NRRL 2363 implicates two alternative strategies for
amide bond formation,” Applied and Environmental Microbiology, vol. 78, no. 7, pp. 2393–2401, 2012.
[57] S. D. Lee, “Nocardia jejuensis sp. nov., a novel actinomycete
isolated from a natural cave on Jeju Island, Republic of Korea,”
International Journal of Systematic and Evolutionary Microbiology, vol. 56, no. 3, pp. 559–562, 2006.
[58] R. Margesin and F. Schinner, “Potential of halotolerant and
halophilic microorganisms for biotechnology,” Extremophiles,
vol. 5, no. 2, pp. 73–83, 2001.
[59] A. Oren, “Industrial and environmental applications of
halophilic microorganisms,” Environmental Technology, vol. 31,
no. 8-9, pp. 825–834, 2010.
[60] O. Selama, P. James, F. Nateche, E. M. H. Wellington, and H.
Hacène, “The world bacterial biogeography and biodiversity
through databases: a case study of NCBI Nucleotide Database
and GBIF Database,” BioMed Research International, vol. 2013,
Article ID 240175, 11 pages, 2013.
Hindawi Publishing Corporation
BioMed Research International
Volume 2014, Article ID 924235, 9 pages
http://dx.doi.org/10.1155/2014/924235
Research Article
Absence of Cospeciation between the Uncultured Frankia
Microsymbionts and the Disjunct Actinorhizal Coriaria Species
Imen Nouioui,1 Faten Ghodhbane-Gtari,1 Maria P. Fernandez,2
Abdellatif Boudabous,1 Philippe Normand,2 and Maher Gtari1,3
1
Laboratoire Microorganismes et Biomolécules Actives, Université de Tunis El Manar (FST) et Université Carthage (INSAT),
2092 Tunis, Tunisia
2
Ecologie Microbienne, Centre National de la Recherche Scientifique UMR 5557, Université Lyon I, 69622 Villeurbanne Cedex, France
3
Laboratoire Microorganismes et Biomolécules Actives, Faculté des Sciences de Tunis, Campus Universitaire, 2092 Tunis, Tunisia
Correspondence should be addressed to Maher Gtari; [email protected]
Received 4 March 2014; Revised 25 March 2014; Accepted 27 March 2014; Published 22 April 2014
Academic Editor: Ameur Cherif
Copyright © 2014 Imen Nouioui et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Coriaria is an actinorhizal plant that forms root nodules in symbiosis with nitrogen-fixing actinobacteria of the genus Frankia. This
symbiotic association has drawn interest because of the disjunct geographical distribution of Coriaria in four separate areas of the
world and in the context of evolutionary relationships between host plants and their uncultured microsymbionts. The evolution
of Frankia-Coriaria symbioses was examined from a phylogenetic viewpoint using multiple genetic markers in both bacteria and
host-plant partners. Total DNA extracted from root nodules collected from five species: C. myrtifolia, C. arborea, C. nepalensis, C.
japonica, and C. microphylla, growing in the Mediterranean area (Morocco and France), New Zealand, Pakistan, Japan, and Mexico,
respectively, was used to amplify glnA gene (glutamine synthetase), dnaA gene (chromosome replication initiator), and the nif
DK IGS (intergenic spacer between nifD and nifK genes) in Frankia and the matK gene (chloroplast-encoded maturase K) and
the intergenic transcribed spacers (18S rRNA-ITS1-5.8S rRNA-ITS2-28S rRNA) in Coriaria species. Phylogenetic reconstruction
indicated that the radiations of Frankia strains and Coriaria species are not congruent. The lack of cospeciation between the two
symbiotic partners may be explained by host shift at high taxonomic rank together with wind dispersal and/or survival in nonhost
rhizosphere.
1. Introduction
The genus Frankia comprises nitrogen-fixing actinobacteria that are able to induce perennial root nodules on
woody dicotyledonous plants called actinorhizals [1]. The
actinorhizal plant families belong to three dicotyledonous
orders: Fagales (Betulaceae,Casuarinaceae, and Myricaceae),
Rosales (Elaeagnaceae, Rhamnaceae, and Rosaceae), and
Cucurbitales (Coriariaceae and Datiscaceae) [2]. Analysis of
the molecular phylogeny of members of Frankia genus consistently identifies four main clusters regardless of the typing
locus used [3]. Three symbiotic Frankia clusters containing
strains able to establish effective nodules and fulfill Koch’s
postulates and one atypical with strains unable to establish
effective nodulation on their host plants have been defined
among Frankia genera. Cluster 1 includes Frankia strains in
association with Betulaceae, Myricaceae, and Casuarinaceae.
Cluster 2 contains Frankia nodulating species from the
Coriariaceae, Datiscaceae, and Rosaceae families as well as
Ceanothus of the Rhamnaceae. Frankia strains in cluster 3
form effective root nodules on plants from members of the
Myricaceae, Rhamnaceae, Elaeagnaceae, and Gymnostoma of
the Casuarinaceae.
Symbiotic Frankia strains have been only isolated from
Fagales (Frankia cluster 1) and the families Elaeagnaceae
and Rhamnaceae (Frankia cluster 3) of the Rosales, while
Frankia of cluster 2 have still not yet been isolated in culture
despite repeated attempts [2]. The position in the Frankia
phylogenetic tree of cluster 2 relative to the other clusters has
varied depending on the marker used. It was proposed at the
base using glnA and 16S rRNA genes [4, 5], derived with ITS
16S–23S rRNA genes [6] and concatenated gyrB, nif H and
2
glnII genes [7] and should be clarified by the upcoming whole
genome phylogeny. Nevertheless, a position at the base of all
symbiotic lineages has been retained in the latest treatment of
Bergey’s manual [8].
Cross-inoculation studies using crushed nodules suggest
that cluster 2 strains form a separate and unique host
specificity group [9–11], even though provenances from the
full geographical range have not yet been tested. Despite
the high taxonomic diversity of host plants belonging to
the cross-inoculation group of cluster 2 and its disjunct
range, uncultured Frankia in root nodules of several host
plants have so far shown a low level of diversity regardless
of the typing locus used [6, 7, 11–16], suggesting a recent
emergence, a strong and recent evolutionary bottleneck, or
a nonrepresentative sampling. The time of emergence of all
Frankia lineages is poorly documented as no convincing
fossil remains. An equivalence between 16S rRNA sequences
distance and time of emergence has been proposed by
Ochman and Wilson [17] where 1% is equivalent to 50 million
years, and since 4% divergence exists between Frankia cluster
2 and the other clusters, one would conclude that Frankia
emerged 200 million years ago [5], which would mean that
there is missing diversity either due to a recent evolutionary
bottleneck or due to a lack of sampling [16]. A possibility thus
exists that the missing variability in cluster 2 strains is due to
the fact that sampling has so far been limited essentially to
North American and Mediterranean areas.
Evidence for cospeciation has been found so far only in
the case of Casuarina species growing in Australia and their
Frankia [18] that are in their immense majority resistant to
growth in pure culture. Among actinorhizal plants of the
Cucurbitales subclade, the family Coriariaceae, with only one
genus, Coriaria, contains about 17 species [19] that occur in
four disjunct areas of the world: the Mediterranean, Southeast
Asia, Central and South America, and the Pacific islands of
New Zealand and Papua New Guinea [20–24]. Yokoyama et
al. [19] considered that the Eurasian species are basal and have
emerged some 60 million years ago. This date is in agreement
with the 65 million years proposed by Bell et al. [25] based on
multiple genes (rbcL, 18S rDNA, atpB) phylogeny, while the
same authors propose an emergence of the Casuarinaceae at
about 30 million years.
The present study was aimed at testing the hypothesis
of cospeciation between uncultured Frankia microsymbionts
and their Coriaria host species sampled from sites covering
the full geographical range of the genus: Coriaria myrtifolia
(Morocco and France), C. nepalensis (Pakistan), C. arborea
(New Zealand), C. japonica (Japan), and C. microphylla
(Mexico).
2. Materials and Methods
2.1. DNA Extraction, PCR Amplification, and Sequencing.
Root nodules from naturally occurring Coriaria species
(Table 1) were kindly provided by Dr. Marı́a Valdés (Escuela
Nacional de Ciencias Biológicas, México, DF, México), Dr.
Sajjad Mirza (National Institute for Biotechnology Genetic
Engineering, Faisalabad, Pakistan), Dr. Warwick Silvester
(University of Waikato, Waikato, New Zealand), Dr. Kawther
BioMed Research International
Benbrahim (University of Fes, Fes, Morocco), Dr. Takashi
Yamanaka (Forest and Forestry Products Research Institute,
Ibaraki, Japan), and Dr. Jean-Claude Cleyet-Marel (INRAIRD, Montpellier, France). Individual lobes were selected,
surface-sterilized in 30% (vol/vol) H2 O2 , and rinsed several
times with distilled sterile water. The DNA extraction from
single nodule lobes was performed as previously described
by Rouvier et al. [26]. Nodule lobes were crushed with sterile
plastic mortars and pestles in 300 𝜇L of extraction buffer
(100 mM Tris (pH 8), 20 mM EDTA, 1.4 M NaCl, 2% (wt/vol)
CTAB (cetyltrimethyl ammonium bromide), and 1% (wt/vol)
PVPP (polyvinyl polypyrrolidone)). The homogenates were
incubated at 65∘ C for 60min, extracted with chloroformisoamyl alcohol (24 : 1, vol/vol) and the resulting DNA was
ethanol-precipitated and resolubilized. The extracted DNA
was used for PCR amplification of both bacterial and plant
DNA regions using the primers listed in Table 2. The amplicons were then cycle-sequenced in both directions using
an ABI cycle sequencing kit (Applied Biosystem 3130). The
nucleotide sequences obtained in this study were deposited in
the NCBI nucleotide sequence database under the accession
numbers given in Table 1.
2.2. Phylogenetic Analysis. Frankia strain CcI3 and Casuarina
equisetifolia were used as outgroups in this study because
they are physiologically distinct from the group studied yet
phylogenetically close. The data sets were completed with
homologous sequences present in the databases (Table 1).
Alignments of Frankia glnA, dnaA, and IGS nif D-K and Coriaria matK and 18S rRNA-ITS1-5.8S rRNA-ITS2-28S rRNA
were generated with ClustalW [27], manually edited with
MEGA 5.0 [28]. Bacterial and plant sequences were separately
concatenated and then used to examine maximum-likelihood
cladogram evolutionary relationships of each symbiotic partner using 1000 bootstraps by following the GTR + G base
substitution model. The distance between the sequences was
calculated using Kimura’s two-parameter model [29]. Phylogenetic trees were constructed using the Neighbor-Joining
method [30] with 1000 bootstraps [31] as implemented in
MEGA 5.0. In parallel, a Bayesian inference was realized
with MrBayes [32] using the GTR + G model and 1,000,000
generations.
A statistical test for the presence of congruence between
Coriaria and Frankia phylogenies was evaluated through
global distance-based fitting in ParaFit program [33] as
implemented in CopyCat [34] and tests of random association were performed with 9999 permutations globally across
both phylogenies for each association.
An additional statistical test for correlation between
geographical distances (obtained using http://www.daftlogic
.com/projects-google-maps-distance-calculator.htm) and
phylogenetic distances was made using Pearson’s r correlation
implemented in the R software [35].
3. Results
To avoid taxonomic ambiguities, DNAs from both Coriaria
hosts and Frankia microsymbionts were characterized on the
same root nodule tissues. The method of DNA isolation from
C. nepalensis
C. japonica
C. myrtifolia
Species
∘
󸀠
󸀠󸀠
∘
󸀠
󸀠󸀠
∘
󸀠
󸀠󸀠
∘
󸀠
󸀠󸀠
∘
Murree, +33 54 15 N 73 23 25 E/33.9042 N
73.3903∘ E/2291.2 m
Pakistan
Tosa district, +33∘ 45󸀠 39.18󸀠󸀠 , +133∘ 27󸀠 42, 89󸀠󸀠 /10 m
Japan
Montpellier, 43∘ 36󸀠 51.48󸀠󸀠 N/3∘ 52󸀠 23.97󸀠󸀠 E/41 m
Nyons, 44 21 46.50 N/5 08 21.82 E/259 m
France
Bab Berred, Chefchaouen:
35∘ 00󸀠 979N/04∘ 58󸀠 092󸀠󸀠 E/1290 m
Oued El Koub, Ouezzane:
35∘ 01󸀠 879N/05∘ 20󸀠 565E/140 m
Morocco
Locality coordinates/altitude (asl)
CnP1
CnP2
CnP3
CnP4
CjJA
CjJB
CjJC
CjJD
CjJE
CmF1
CmF2
CmF3
CmF4
CmF5
CmNy1
CmNy2
CmNy3
CmNy4
CmNy5
KC796590 KC796599
CmM1a
CmM1b
CmM1c
CmM2a
CmM2b
AB016456
KC796605
AB016459
AF280103
KC796597 KC796607
AF280101
KC796594
AF280102
KC796593 KC796602
KC796598 KC796603
KC796591 KC796600
KC796592 KC796601
KC796550
KC796551
KC796552
KC796553
KC796554
KC796555
KC796556
KC796557
KC796558
KC796503
KC796504
KC796505
KC796506
KC796507
KC796576
KC796577
KC796578
KC796579
KC796580
KC796586 KC796559
KC796587 KC796560
KC796588 KC796561
KC796589 KC796562
KC796590 KC796563
KC796591 KC796564
KC796592 KC796565
KC796593
—
KC796594 KC796566
KC796595 KC796567
KC796578
KC796579
KC796580
—
KC796581
KC796582
KC796583
KC796584
KC796585
KC796544 KC796508 KC796584
KC796545 KC796509 KC796585
KC796546 KC796510 KC796586
KC796536
KC796537
KC796538
KC796539
KC796540
KC796526
KC796527
KC796528
KC796529
KC796530
KC796531
KC796532
KC796533
KC796534
KC796535
KC796517
KC796518
KC796519
KC796520
KC796521
KC796522
KC796523
KC796524
KC796525
Yang et al., unpublished
This study
This study
This study
This study
This study
This study
This study
This study
Yang et al., unpublished
(Yokoyama et al., 2000 [19])
This study
This study
This study
This study
This study
Yang et al., unpublished
(Yokoyama et al., 2000 [19])
This study
This study
This study
This study
This study
This study
This study
This study
This study
This study
This study
This study
This study
This study
Plant sequence accession number Bacterial sequence accession number
References
ITS1-ITS2
matK
glnA
dnaA
IGS nif D-K
CmMs1
CmMs2
CmMs3
CmMs4
Nodule
labels
Table 1: List of Coriaria root nodules and sequences used in this study.
BioMed Research International
3
Casuarina
equisetifolia
Datisca
glomerata
C. papuana
C. sarmentosa
C. ruscifolia
C. terminalis
C. intermedia
C. microphylla
C. arborea
Species
Morelos, 99∘ 30󸀠 , 19∘ 30󸀠 /2400 m
Mexico
Hapuku river, North Canterbury, South island:
−42∘ 23󸀠 42.24󸀠󸀠 , +173∘ 41󸀠 18.07󸀠󸀠 /64 m
New Zealand
Locality coordinates/altitude (asl)
EF635457
EF635475
AF277293
AB16454
KC796595 KC796604
AY864057
AY968449
AB015462
AF485250
CP000249 CP000249
CP002801 CP002801
KC796547 KC796514
KC796548 KC796515
KC796549 KC796516
KC796542 KC796511
KC796543 KC796512
KC796544 KC796513
CP000249
CP002801
KC796587
KC796588
KC796589
KC796581
KC796582
KC796583
This study
This study
This study
Yang et al., unpublished
(Yokoyama et al., 2000 [19])
Yang et al., unpublished
(Yokoyama et al., 2000 [19])
Yang et al., unpublished
Yang et al., unpublished
Yang et al., unpublished
Yang et al., unpublished
(Yokoyama et al., 2000 [19])
Yang et al., unpublished
(Yokoyama et al., 2000 [19])
(Yokoyama et al., 2000 [19])
(Persson et al., 2011 [50])
Zhang et al., unpublished
Forrest and Hollingsworth
unpublished
(Normand et al., 2007 [51])
Sogo et al., unpublished
Herbert et al., unpublished
This study
This study
This study
(Yokoyama et al., 2000 [19])
Rotherham et al., unpublished
Rotherham et al., unpublished
Yang et al., unpublished
Plant sequence accession number Bacterial sequence accession number
References
ITS1-ITS2
matK
glnA
dnaA
IGS nif D-K
Table 1: Continued.
CmicMx1 KC796596 KC796606
CmicMx2
CmicMx3
AY091813
AB016458
AF280100
AB016455
AY091817
AY091815
AY091814
AF280104
AB016462
AY091816
AB016464
AB016461
CaNZ1
CaNZ2
CaNZ3
Nodule
labels
4
BioMed Research International
BioMed Research International
5
Table 2: Primers used for PCR amplification and DNA sequencing.
Gene primers
glnA
DB41
DB44
dnaA
F7154 dnaAF
F7155 dnaAR
IGS nif D-K
F9372 nifD1 5
F9374 nifK1 5
F9373 nifD2 5
F9375 nifK2 5
18S-ITS1-5.8S-ITS2-28S
ITS1
ITS4
F9030-CJ-ITSF
F9031-CJ-ITSR
matK
F9249-matKF
F9250-matkR
Sequence (5󸀠 -3󸀠 )
Amplicons approximate size (bp)
References
TTCTTCATCCACGACCCG
GGCTTCGGCATGAAGGT
500
(Clawson et al., 2004 [4])
GAGGARTTCACCAACGACTTCAT
CRGAAGTGCTGGCCGATCTT
700
Bautista et al. unpublished
GTCATGCTCGCCGTCGGNG
GTTCTTCTCCCGGTAyTCCCA
700
This study
ACCGGCTACGAGTTCGCNCA
TGCGAGCCGTGCACCAGNG
700
This study
TCCGTAGGTGAACCTGCGG
TCCTCCGCTTATTGATATGC
700
(White et al., 1990 [52])
AGCCGGACCCGCGACGAGTTT
CGACGTTGCGTGACGACGCCCA
400
This study
ACATTTAAATTATGTGTCAG
TGCATATACGCACAAATC
700
This study
root nodules used in this study yielded PCR-amplifiable DNA
for both bacterial and plant PCR target sequences in all cases.
However, in several instances it was easier to amplify Frankia
than Coriaria DNA, which may have been mostly due to the
specificity of the primer sets used. Thus, in this study, new
primers were designed (Table 2).
For the bacterial microsymbionts, the average uncorrected p-distances (proportion of differences between sequences) were computed for each region and were found to be
relatively small for dnaA (𝑝 = 0.0378), intermediate for glnA
(𝑝 = 0.0625), and high for IGS nif D-K region (𝑝 = 0.0833).
Blast analyses of the individual genes permitted assigning
them all to Frankia cluster 2. Nearly 3000 nucleotides were
obtained by concatenating sequences of the three DNA
regions.
Sequences variation for Coriaria species was small based
on matK gene (𝑝 = 0.0205) compared to ITS1-ITS2 sequences
(𝑝 = 0.0423). By concatenating matK and ITS1- ITS2 region,
a composite sequence of 1500 nt was used for phylogenetic
inference.
All studied sequences were analyzed independently to test
for incongruence between the data sets for each symbiotic
partner. Similar topologies have been generally observed
between phylogenetic trees inferred from glnA, dnaA, and
IGS nif D-K sequences for Frankia and from matK and ITS
sequences for Coriaria regardless of the used phylogenetic
methods (not shown).
The topologies of the trees obtained for the two symbiotic
partners were not congruent (Figure 1). Moreover, global
distance-based ParaFit analysis recovered mostly random
associations between Frankia and Coriaria host plant species
(𝑝 = 0.33) and rejected cospeciation hypothesis. On the
microbial side, the New Zealand microsymbionts were at
the root (Group A); then three groups emerged, group
B comprising the Pakistani, Mexican, and Mediterranean
symbionts from France, group C comprising microsymbionts from Morocco, and then group D comprising French
and Japanese microsymbionts as well as the Dg1 reference
sequence obtained initially from a Pakistani soil. On the
host plant side, group 1 at the root comprises New Zealand
and South American sequences, while group 2 comprises the
Japanese, Mediterranean, and Pakistani sequences.
On the other hand, no significant correlations were found
for Frankia symbionts (𝑟2 = 0.772; Fgeneticdist = (geogdist ×
5.830E−06 ) + 2.541E−02 ) nor for the Coriaria host plants (𝑟2
= 0.883; Fgeneticdist = (geogdist × 2.023E−06 ) + 6.460E−03 )
(data not shown).
4. Discussion
Cospeciation has been postulated to have occurred in
some Frankia actinorhizal host plants, in particular in the
Casuarina-Frankia cluster 1b [18] but not in Alnus-infective
and Elaeagnus-infective Frankia strains where many isolates
able to fulfill Koch’s postulates have been obtained. To
test if cospeciation was general or an exception, it was
decided to study uncultured Frankia microsymbionts and
representative Coriaria hosts, a lineage where no Frankia
isolate exists and where geographic discontinuities may have
limited dispersion. DNA sequences were obtained from root
nodules collected from New Zealand (C. arborea), Pakistan
(C. nepalensis), Japan (C. japonica), Mexico (C. microphylla),
and France and Morocco (C. myrtifolia) and multiple molecular markers were analyzed for phylogenetic inference.
6
BioMed Research International
99
99
68
Group C
80
CmF2
CmNy5
F.CmMs4
F.CmM1a
F.CmM1c
F.CmM1b
F.CmNy4
F.CmMs3
75
81
Group B
91
99
99
97
Group A
100
F.CmM2b
Oceania
Asia
100
C. nepalensis
90
75
54
C. terminalis
C. intermedia
C. japonica
CjJA
CmicMx1
C. microphylla
F.CnP1
C. papuana
F.CnP2
CaNZ1
F.CmMx1
C. arborea
F.CaNZ1
76
CnP1
F.CmNy5
F.CmMx2
86
C. myrtifolia
C. myrtifolia
F.CmNy1
F.CaNZ2
Frankia
CmMs1
F.CmF2
F.CmMs1
89
CmM2a
F.CmF1
F.CmMs2
92
CmM1a
Group 2
Group D
100
F.CjJA
Dg1
96
70
90
87
90
51
C. ruscifolia
73
C. sarmentosa
C. lurida
Group 1
F.CjJB
69
83
70
Coriaria
Europe/N. Africa
America
Figure 1: Phylogenetic trees of the Frankia microsymbionts (left) and the Coriaria host plants (right). The Frankia tree was constructed using
the glnA, dnaA, and the nif D-K intergenic spacer, while the Coriaria tree was done using the matK and the 18S rRNA-ITS1-5.8S rRNA-ITS228S rRNA with ML method using strain CcI3 and Casuarina as outgroups respectively for Frankia and hot plant phylogenetic trees. The
numbers at branches indicate bootstrap results above 50%. Lines are drawn between the microsymbionts and their hosts. The color code
indicates the place of origin of the leave or of the set when homogenous. The groups numbers 1 and 2 on the right are according to Yokoyama
et al. [19].
Paleontological data based on macrofossils and pollen
fossils have brought several authors [36–40] to conclude that
the Coriariaceae had a Laurasian origin (North America
and Eurasia). There have been a few dissenting opinions, in
particular those of Croizat [41] and Schuster [42] who considered that Coriaria originated in Gondwana and migrated
to the Northern Hemisphere. However, such paleontological
studies are not very convincing, as it is recognizably hard
to ascribe fossils to a given family and even more so to a
given genus. Thus, several authors have been surprised by
the results of molecular phylogeny positioning Coriariaceae
close to the Datiscaceae. Molecular approaches would thus
give support to a Gondwanan origin.
Yokoyama et al. [19] proposed that Coriaria species had
emerged 59–63 million years ago, which is coherent with
the date of 70 million years proposed by Bell et al. [25],
considerably older than that proposed (30 million years) by
the same authors for the Casuarinaceae.
Topology and clustering of Coriaria phylogeny obtained
in the current study are similar to those obtained by
Yokoyama et al. [19], while the position at the base of the
host plant species from New Zealand, C. arborea, and the
South American C. ruscifolia and C. microphylla species was
contrary to that of Yokoyama et al. [19] who found the
Eurasian species at the base using rbcL (a large subunit of
ribulose 1,5-bisphosphate carboxylase/oxygenase) and matK
(maturase K) genes. The present study suggests that the
Coriaria ancestor may have emerged between Asia and NZ
and then dispersed worldwide and that the Asian lineage
may have given rise relatively recently to the Mediterranean
species, while the NZ lineage gave rise to the North American
species (Figure 2).
Previous studies had concluded that Frankia cluster 2 had
a low genetic diversity [6, 7, 16] but these studies had been
focused on only part of the full diversity of the symbiotic
Coriaria-Frankia, essentially in North America and Mediterranean. In this work we aimed to expand the scope of the
study to the worldwide diversity and phylogeny of microsymbionts of Coriaria species. Four microbial subgroups were
identified that did not fit to the geographic range of the host
plants, while two host plant subgroups were identified. The
position of subgroup A containing microsymbionts of New
Zealand C. arborea at the base of Frankia cluster 2 is in
agreement with previous study [16]. In view of previously
BioMed Research International
7
CmNy1-2-3-4-5
CmF1-2-3-4-5
Coriaria myrtifolia
CmM1a-b-c
CmM2a-b
CmMs1-2-3-4-5
C. microphylla
CmicMx1-2-3
C. nepalensis
CnP1-2-3
C. terminalis
C. japonica
CjJA-B-C-D-E
C. intermedia
C. papuana
Coriaria sp.
C. sarmentosa
C. ruscifolia
Coriaria agustissima
C. arborea CaNZ1-2-3
C. kingiana
C. lurida
C. plumosa
C. pottsiana
C. pteroides
C. sarmentosa
Figure 2: Distribution of Coriaria species. Root nodules have been sampled from C. myrtifolia, C. arborea, C. nepalensis, C. japonica, and C.
microphylla growing in Mediterranean areas (Morocco and France), New Zealand, Pakistan, Japan, and Mexico, respectively. Short arrows
indicate sampling sites for this study while long arrows indicate possible routes of dispersal as discussed.
reported data, members of cluster 2 Frankia studied here
were found to have relatively higher sequences variation (pdistance = 0.0625) than those reported by Vanden Heuvel et
al. [16] (𝑝 = 0.00454) based on the same 460 nt of the glnA
gene.
Molecular clock dating suggests that Frankia genus has
emerged much earlier, 125 Myr bp before the appearance of
angiosperm fossils in the Cretaceous period and the extant
actinorhizal plants [4]. Normand et al. [5] using the 4%
divergence in the 16S rRNA between cluster 2 and other
Frankia lineages as equivalent to 50 MY/1% distance [17]
concluded that the genus Frankia had emerged long before
the extant dicotyledonous lineages. These authors proposed
Frankia cluster 2 as the proto-Frankia as nonsymbiotic
ancestor of 62–130 Myr bp [43] and 100–200 Myr bp [5]. Since
the distance in the 16S rRNA gene between cluster 1a (Frankia
alni) and cluster 1b is less than 1%, the date of emergence of the
Casuarina-infective lineage has been proposed to be less than
50 million years [5]. Thus the Casuarina/Frankia 1b lineage is
considerably younger than the Coriaria/Frankia lineage and
would have had less time to migrate out of its cradle and
mingle with other hosts in its new territories and lose the
cospeciation signal.
Symbiotic partnership often tends to become obligatory,
as in the case of Casuarina host plants, where Frankia is only
present in soils close to the host plant [44], which means that
the bacterium loses autonomy and becomes dependent on its
host. Speciation of the host could then lead to synchronous
speciation of its microsymbiont unless dispersal through
long-distance carriers such as winds or migratory birds
occurred or if there is survival of Frankia cluster 2 in the
rhizosphere of nonhosts as was recently demonstrated for
Alnus glutinosa in Tunisia [45]. The numerous transitions
seen in the Frankia phylogenetic tree from one continent to
another would reinforce the idea.
Yokoyama et al. [19] concluded from their study of the
Coriaria species phylogeny that the Eurasian species had
diverged earlier and are more diverse than other groups, but
that nevertheless the origin of the genus could have been in
North America, whence the South America and the Pacific
species could have originated. Our study brings us to suggest
a third possibility, Oceania, which could also be the origin
of this actinorhizal symbiosis, which can be concluded from
phylogenetic inferences positioning both bacterial and host
plant partners as at the base to Frankia-Coriaria symbiosis.
Another element that would support this hypothesis is the
large number of extant species there; according to Yokoyama
et al. [19] New Zealand would be home to 8 of the 17 existing
species. A similar argument has often been made to establish
Sub-Saharan Africa as the cradle of humankind [46] or
Mexico for maize [47].
Comparison of both the plant and the microbe phylogenetic topologies did not show any evidence for cospeciation
of Frankia microsymbiontsand their Coriaria host species.
The results obtained in this study suggest that Frankia
microsymbionts hosted currently by Coriaria species had
probably dispersed globally as a proto-Frankia, a free living
and nonsymbiotic ancestor. In parallel, the proto-Coriaria
then diversified into the extant Coriaria species that appear to
have been retreating given their scattered distribution, a trend
8
possibly reinforced recently due to man uprooting because
of the toxicity of the fruits for mammals [48, 49]. It can
thus be hypothesized that Coriaria appeared in the Pacific
Islands more than 70 million years ago and presumably was
symbiotic from the start, before dispersing over all continents
as they drifted apart. The Coriaria species diversified in
their different biotopes, as they saw the appearance of other
plants hosting the same microsymbiont of Frankia cluster 2
such as Datiscaceae, Rosaceae, Ceanothus, or even nonhost
species such as Alnus glutinosa that was recently found to
host Frankia cluster 2 in its rhizosphere [45]. Members of
these alternative host plant species cooccur sympatrically
with Coriaria such as Ceanothus and Purshia species in
Mexico and Datisca cannabina in Pakistan. These Frankia
cluster 2 host plant species have more extended geographic
distribution and overlap in some instances Coriaria’s disjunct
area and as a result can compensate Frankia microsymbionts
remoteness, which would thus obscure the cospeciation
signal. Cospeciation may also occur but subsequently is lost
after bacterial mixing and fitness selection in the presence of
“indigenous” and “dispersal” symbionts.
Conflict of Interests
BioMed Research International
[5]
[6]
[7]
[8]
[9]
[10]
The authors declare that there is no conflict of interests
regarding the publication of this paper.
[11]
Acknowledgments
This work is supported by CMCU (Comité Mixte TunisoFrançais pour la Coopération Inter-Universitaire No.
10/G0903). The authors are grateful to Dr. Marı́a Valdés
(Escuela Nacional de Ciencias Biológicas, México, México),
Dr. Sajjad Mirza (National Institute for Biotechnology
Genetic Engineering, Faisalabad, Pakistan), Dr. Warwick
Silvester (University of Waikato, Waikato, New Zealand),
Dr. Kawther Benbrahim and Dr. A. Ennabili (University of
Fes, Fes, Morocco), Mr. Spick (Montpellier Botanical garden,
France), Dr. J. C. Cleyet-Marel (Montpellier INRA, France),
Mr. D. Moukouanga (IRD Montpellier, France), and Dr.
Takashi Yamanaka (Forest and Forestry Products Research
Institute, Ibaraki, Japan) for providing Coriaria nodules.
[12]
[13]
[14]
References
[1] M. P. Lechevalier, “Taxonomy of the genus Frankia (Actinomycetales),” International Journal of Systematic Bacteriology, vol. 44,
no. 1, pp. 1–8, 1994.
[15]
[2] D. R. Benson, B. D. Vanden Heuvel, and D. Potter, “Actinorhizal
symbioses: diversity and biogeography,” in Plant Microbiology,
M. Gillings, Ed., pp. 97–127, BIOS Scientific Publishers, Oxford,
UK, 2004.
[16]
[3] M. Gtari, L. S. Tisa, and P. Normand, “Diversity of Frankia
strains, actinobacteria symbionts of actinorhizal plants,” in
Symbiotic Endophytes, Soil Biology, R. Aroca, Ed., vol. 37,
Chapter 7, pp. 123–148, Springer, Berlin, Germany, 2013.
[4] M. L. Clawson, A. Bourret, and D. R. Benson, “Assessing the
phylogeny of Frankia-actinorhizal plant nitrogen-fixing root
[17]
nodule symbioses with Frankia 16S rRNA and glutamine synthetase gene sequences,” Molecular Phylogenetics and Evolution,
vol. 31, no. 1, pp. 131–138, 2004.
P. Normand, S. Orso, B. Cournoyer et al., “Molecular phylogeny
of the genus Frankia and related genera and emendation of
the family Frankiaceae,” International Journal of Systematic
Bacteriology, vol. 46, no. 1, pp. 1–9, 1996.
F. Ghodhbane-Gtari, I. Nouioui, M. Chair, A. Boudabous, and
M. Gtari, “16S-23S rRNA intergenic spacer region variability in
the genus Frankia,” Microbial Ecology, vol. 60, no. 3, pp. 487–
495, 2010.
I. Nouioui, F. Ghodhbane-Gtari, N. J. Beauchemin, L. S. Tisa,
and M. Gtari, “Phylogeny of members of the Frankia genus
based on gyrB, nifH and glnII sequences,” Antonie van Leeuwenhoek, vol. 100, no. 4, pp. 579–587, 2011.
P. Normand and D. R. Benson, “Genus I Frankia Brunchorst
1886, 174AL,” in Bergey’s Manual of Systematic Bacteriology, The
Actinobacteria, M. Goodfellow, P. Kämpfer, H.-J. Busse et al.,
Eds., vol. 5 of Bergey’s Manual Trust, pp. 512–520, Springer, New
York, NY, USA, 2012.
J. G. Torrey, “Cross-inoculation groups within Frankia and
host-endosymbiont associations,” in The Biology of Frankia and
Actinorhizal Plants, C. R. Schwintzer and J. D. Tjepkema, Eds.,
pp. 83–106, Academic Press, San Diego, Calif, USA, 1990.
S. J. Kohls, J. Thimmapuram, C. A. Buschena, M. W. Paschke,
and J. O. Dawson, “Nodulation patterns of actinorhizal plants
in the family Rosaceae,” Plant and Soil, vol. 162, no. 2, pp. 229–
239, 1994.
M. S. Mirza, K. Pawlowski, F. Y. Hafeez, A. H. Chaudhary,
and A. D. L. Akkermans, “Ultrastructure of the endophyte and
localization of nifH transcripts in root nodules of Coriaria
nepalensis Wall. by in situ hybridization,” New Phytologist, vol.
126, no. 1, pp. 131–136, 1994.
G. Nick, E. Paget, P. Simonet, A. Moiroud, and P. Normand,
“The nodular endophytes of Coriaria spp. form a distinct
lineage within the genus Frankia,” Molecular Ecology, vol. 1, no.
3, pp. 175–181, 1992.
M. Bosco, S. Jamann, C. Chapelon, and S. P. Normand, “Frankia
microsymbiont in Dryas drummondii nodules is closely related
to the microsymbiont of Coriaria and genetically distinct from
other characterized Frankia strains,” in Nitrogen Fixation with
Non-Legumes, N. A. Hegazi, M. Fayez, and M. Monib, Eds., pp.
173–183, The American University in Cairo Press, 1994.
D. R. Benson, D. W. Stephens, M. L. Clawson, and W. B.
Silvester, “Amplification of 16S rRNA genes from Frankia strains
in root nodules of Ceanothus griseus, Coriaria arborea, Coriaria
plumosa, Discaria toumatou, and Purshia tridentata,” Applied
and Environmental Microbiology, vol. 62, no. 8, pp. 2904–2909,
1996.
D. R. Benson and M. L. Clawson, “Evolution of the actinorhizal
plant symbioses,” in Prokaryotic Nitrogen Fixation: A Model
System for Analysis of Biological Process, E. W. Triplett, Ed., pp.
207–224, Horizon Scientific Press, Wymondham, UK, 2000.
B. D. Vanden Heuvel, D. R. Benson, E. Bortiri, and D. Potter,
“Low genetic diversity among Frankia spp. strains nodulating
sympatric populations of actinorhizal species of Rosaceae,
Ceanothus (Rhamnaceae) and Datisca glomerata (Datiscaceae)
west of the Sierra Nevada (California),” Canadian Journal of
Microbiology, vol. 50, no. 12, pp. 989–1000, 2004.
H. Ochman and A. C. Wilson, “Evolution in bacteria: evidence
for a universal substitution rate in cellular genomes,” Journal of
Molecular Evolution, vol. 26, no. 1-2, pp. 74–86, 1987.
BioMed Research International
[18] P. Simonet, E. Navarro, C. Rouvier et al., “Co-evolution
between Frankia populations and host plants in the family
Casuarinaceae and consequent patterns of global dispersal,”
Environmental Microbiology, vol. 1, no. 6, pp. 525–533, 1999.
[19] J. Yokoyama, M. Suzuki, K. Iwatsuki, and M. Hasebe, “Molecular phylogeny of Coriaria, with special emphasis on the disjunct
distribution,” Molecular Phylogenetics and Evolution, vol. 14, no.
1, pp. 11–19, 2000.
[20] R. D. O. Good, “The geography of the genus Coriaria,” New
Phytologist, vol. 29, pp. 170–198, 1930.
[21] H. H. Allan, “Coriariaceae,” in Flora of New Zealand, L. B.
Moore, Ed., pp. 300–305, Government printer, Wellington, New
Zealand, 1961.
[22] R. Melville, “Continental drift, mesozoic continents and the
migrations of the angiosperms,” Nature, vol. 211, no. 5045, pp.
116–120, 1966.
[23] L. E. Skog, “The genus Coriaria Coriariaceae in the Western
Hemisphere,” Rhodora, vol. 74, pp. 242–253, 1972.
[24] R. Melville, “Vicarious plant distributions and paleogeography
of the Pacific region,” in Vicariance Biogeography, G. Nelson and
E. D. Rosen, Eds., pp. 413–435, Columbia University Press, New
York, NY, USA, 1981.
[25] C. D. Bell, D. E. Soltis, and P. S. Soltis, “The age and diversification of the angiosperms re-revisited,” American Journal of
Botany, vol. 97, no. 8, pp. 1296–1303, 2010.
[26] C. Rouvier, J. Schwenke, Y. Prin et al., “Biologie et diversité
génétique des souches de Frankia associées aux Casuarinacées,”
Acta Botanica Gallica, vol. 143, pp. 567–580, 1996.
[27] J. D. Thompson, D. G. Higgins, and T. J. Gibson, “CLUSTAL
W: improving the sensitivity of progressive multiple sequence
alignment through sequence weighting, position-specific gap
penalties and weight matrix choice,” Nucleic Acids Research, vol.
22, no. 22, pp. 4673–4680, 1994.
[28] K. Tamura, D. Peterson, N. Peterson, G. Stecher, M. Nei, and
S. Kumar, “MEGA5: molecular evolutionary genetics analysis
using maximum likelihood, evolutionary distance, and maximum parsimony methods,” Molecular Biology and Evolution,
vol. 28, no. 10, pp. 2731–2739, 2011.
[29] M. Kimura, “A simple method for estimating evolutionary rates
of base substitutions through comparative studies of nucleotide
sequences,” Journal of Molecular Evolution, vol. 16, no. 2, pp. 111–
120, 1980.
[30] N. Saitou and M. Nei, “The neighbor-joining method: a new
method for reconstructing phylogenetic trees,” Molecular Biology and Evolution, vol. 4, no. 4, pp. 406–425, 1987.
[31] J. Felsenstein, “Confidence limits on phylogenies: an approach
using the bootstrap,” Evolution, vol. 39, pp. 783–791, 1985.
[32] F. Ronquist and J. P. Huelsenbeck, “MrBayes 3: bayesian phylogenetic inference under mixed models,” Bioinformatics, vol. 19,
no. 12, pp. 1572–1574, 2003.
[33] P. Legendre, Y. Desdevises, and E. Bazin, “A statistical test for
host-parasite coevolution,” Systematic Biology, vol. 51, no. 2, pp.
217–234, 2002.
[34] J. P. Meier-Kolthoff, A. F. Auch, D. H. Huson, and M. Göker,
“CopyCat: cophylogenetic analysis tool,” Bioinformatics, vol. 23,
no. 7, pp. 898–900, 2007.
[35] R. Ihaka and R. Gentleman, “R: a language for data analysis and
graphics,” Journal of Computational and Graphical Statistics, vol.
5, no. 3, pp. 299–314, 1996.
[36] A. N. Gladkova, “Fragments of the history of the Myricaceae
family,” Pollen and Spore, vol. 4, p. 345, 1962.
9
[37] J. A. Doyle, “Cretaceous angiosperm pollen of the Atlantic
coastal plain and its evolutionnary significance,” Journal of the
Arnold Arboretum, vol. 50, pp. 1–35, 1969.
[38] P. H. Raven and D. I. Axelrod, “Angiosperm biogeography and
past continental movements,” Annals of the Missouri Botanical
Garden, vol. 61, pp. 539–673, 1974.
[39] J. Muller, “Fossil pollen records of extant angiosperms,” The
Botanical Review, vol. 47, no. 1, pp. 1–142, 1981.
[40] O. Eriksson and B. Bremer, “Pollination systems, dispersal
modes, life forms, and diversification rates in angiosperm
families,” Evolution, vol. 46, pp. 258–266, 1992.
[41] L. Croizat, Manual of Phytogeography: An Account of Plant
Dispersal Throughout the World, Junk, The Hague, 1952.
[42] R. M. Schuster, “Plate tectonics and its bearing on the geographical origin and dispersal of angiosperms,” in Origin and Early
Evolution of Angiosperms, C. B. Beck, Ed., pp. 48–138, Columbia
University Press, New York, 1976.
[43] S.-C. Jeong, A. Liston, and D. D. Myrold, “Molecular phylogeny
of the genus Ceanothus (Rhamnaceae) using rbcL and ndhF
sequences,” Theoretical and Applied Genetics, vol. 94, no. 6-7, pp.
852–857, 1997.
[44] J. F. Zimpfer, C. A. Smyth, and J. O. Dawson, “The capacity of
Jamaican mine spoils, agricultural and forest soils to nodulate
Myrica cerifera, Leucaena leucocephala and Casuarina cunninghamiana,” Physiologia Plantarum, vol. 99, no. 4, pp. 664–672,
1997.
[45] I. Nouioui, I. Sbissi, F. Ghodhbane-Gtari, K. Benbrahim, P.
Normand, and M. Gtari, “First report on the occurrence of the
uncultivated cluster 2 Frankia microsymbionts in soil outside
the native actinorhizal host range area,” Journal of Biosciences,
vol. 38, pp. 695–698, 2013.
[46] S. Ramachandran, O. Deshpande, C. C. Roseman, N. A.
Rosenberg, M. W. Feldman, and L. L. Cavalli-Sforza, “Support
from the relationship of genetic and geographic in human
populations for a serial founder effect originating in Africa,”
Proceedings of the National Academy of Sciences of the United
States of America, vol. 102, no. 44, pp. 15942–15947, 2005.
[47] B. Prasanna, “Diversity in global maize germplasm: characterization and utilization,” Journal of Biosciences, vol. 37, pp. 843–
855, 2012.
[48] L. De Haro, P. Pommier, L. Tichadou, M. Hayek-Lanthois, and J.
Arditti, “Poisoning by Coriaria myrtifolia Linnaeus: a new case
report and review of the literature,” Toxicon, vol. 46, no. 6, pp.
600–603, 2005.
[49] S. F. Belcher and T. R. Morton, “Tutu toxicity: three case
reports of Coriaria arborea ingestion, review of literature and
recommendations for management,” The New Zealand Medical
Journal, vol. 126, pp. 103–109, 2013.
[50] T. Persson, D. R. Benson, P. Normand et al., “Genome
sequence of “Candidatus Frankia datiscae” Dg1, the uncultured
microsymbiont from nitrogen-fixing root nodules of the dicot
Datisca glomerata,” Journal of Bacteriology, vol. 193, no. 24, pp.
7017–7018, 2011.
[51] P. Normand, P. Lapierre, L. S. Tisa et al., “Genome characteristics of facultatively symbiotic Frankia sp. strains reflect host
range and host plant biogeography,” Genome Research, vol. 17,
no. 1, pp. 7–15, 2007.
[52] T. J. White, T. Bruns, S. Lee, and J. Taylor, “Amplification
and direct sequencing of fungal ribosomal RNA genes for
phylogenetics,” in PCR Protocols: A Guide to Methods and
Applications, M. A. Innis, D. H. Gelfand, J. J. Sninsky, and T.
J. White, Eds., pp. 315–322, Academic Press, San Diego, Calif,
USA, 1990.