University of Groningen Normal operating range of the ... · and activity of the rhizosphere...
Transcript of University of Groningen Normal operating range of the ... · and activity of the rhizosphere...
University of Groningen
Normal operating range of the microbial community under potatoInceoğlu, Özgul
IMPORTANT NOTE: You are advised to consult the publisher's version (publisher's PDF) if you wish to cite fromit. Please check the document version below.
Document VersionPublisher's PDF, also known as Version of record
Publication date:2011
Link to publication in University of Groningen/UMCG research database
Citation for published version (APA):Inceolu, Ö. (2011). Normal operating range of the microbial community under potato. Groningen: s.n.
CopyrightOther than for strictly personal use, it is not permitted to download or to forward/distribute the text or part of it without the consent of theauthor(s) and/or copyright holder(s), unless the work is under an open content license (like Creative Commons).
Take-down policyIf you believe that this document breaches copyright please contact us providing details, and we will remove access to the work immediatelyand investigate your claim.
Downloaded from the University of Groningen/UMCG research database (Pure): http://www.rug.nl/research/portal. For technical reasons thenumber of authors shown on this cover page is limited to 10 maximum.
Download date: 10-02-2020
Chapter 8
General Discussion
134
Impact of plants (including GM plants) on the soil microbiota
It has been suggested that each soil harbors its own characteristic microflora (28, 63).
Furthermore, it has been shown that the community structure and diversity of the soil
microbiota are important determinants of the stability of key functions of agro-
ecosystems, even though the relationship between the functioning of agro-ecosystems
and soil microbial diversity is not well defined yet (51). Soil type, nutrient status, pH,
moisture and plant factors (e.g. plant species and age) can affect the abundance,
community composition and activities of soil microorganisms (31). In fact, plants are the
most important contributors to the formation of organic matter in soil, as a result of their
release of nutrients, mucilages and complex polymers. Thus, "hot spots" for microbial
growth and activity form in zones around plant roots, which are collectively called the
rhizosphere. However, the local interactions between plant roots and the soil microbiota
are still poorly understood (11), in spite of the fact that the composition and quantity of
particular carbonaceous substrates in root exudates are important. These may vary
depending on plant species, rhizosphere microsite location and plant growth stage (91).
Due to changes in the exudation patterns over the time of plant growth, the composition
and activity of the rhizosphere microflora can also be altered as a function of time (36,
91). The rhizosphere microbiota is further influenced by plant species type as well as by
the genetic make-up of a plant species (15, 53, 68, 74, 94).
Depending on the objective of the modification, genetically modified (GM) plants
can have great potential to advance agriculture. On the other hand, there are concerns
about the potential ecological effects of GM plants intended for routine cropping. For
instance, GM plants might exert undesirable effects on soil organisms, in comparison to
the parent plant. Such effects might be related to changes in root exudation patterns,
plant residues and even to changed agricultural management practices that go with the
genetic modification (49). In previous studies, transgenic potatoes carrying traits for
antibacterial activities or altered starch composition have been shown to affect the
rhizosphere microbial community (57, 71, 72). However, the effects were found to be of a
magnitude that was comparable with other factors such as soil type and plant genotype
(57, 71).
GM plants have been shown to exert variable effects on the soil ecosystem, which
are often relatively minor compared with the impacts of differences in cultivars or those
associated with biotic and abiotic factors. Thus, determinations of the natural variation in
a cropping system, in which effects of cultivar differences are also considered, provide
valuable baseline reference data (47). The effects of GM plants should ideally be
considered in comparison to the baseline, establishing the so-called normal operating
conditions of functioning (the normal operating range or NOR) of the living soil under
cultivation.
135
This study
The main idea behind the work executed in the context of this thesis was to (1) determine
the variation among soil microbiological parameters as a result of cropping of a series of
different potato cultivars, and (2) weigh the putative effects of GM potato plants to those
of the selected cultivars. This way, as a first outcome, an assessment of the NOR of potato
cropping would become tangible. In addition, the question whether the putative effects of
the GM plant remained within or reached outside the NOR would receive an answer. We
placed an emphasis on the assessment of the overall bacterial community composition as
well as on that of individual microbial populations as these are affected by plants (42).
For the experiments, six different potato cultivars (Aveka [A], Aventra [Av], Karnico
[K], Modena [M; modified from Karnico for low amylose content of the tubers] (16),
Premiere [P] and Désirée [D]) were used. Cultivars A, Av, K and M produced tubers with
high starch contents and had a low and/or medium growth rate, whereas cultivars P and D
yielded tubers with relatively low starch contents and had high growth rates. The different
cultivars had different parental cultivars in the first generation, so that their overall
pedigree was complex. For instance, cultivar A was related to D in the fifth generation and
to K in the third generation (90).
Which bacterial community DNA extraction method gives the most representative results?
For the current study, we tried to establish a soil DNA extraction method with minimal
bias with regard to the yield of DNA pertaining to the key bacterial communities studied.
The soil DNA extraction method has been suggested to be the major determinant of the
apparent bacterial diversity and community structure in a soil sample (4, 5). Thus, at the
beginning of each study, the best extraction method for the purpose of the study should
be determined. Next, standardized methods must be used for all subsequent elements of
the study. The suitability of four selected soil DNA extraction methods to obtain
representative DNA from three agricultural soils of divergent texture, organic matter (OM)
content and pH was thus assessed. We confirmed that the soil DNA extraction method
was indeed a major determinant of the apparent bacterial diversity and community
structure. Strikingly, different new or already known bacterial groups were found in the
same soil as a result of something as simple as the application of a different soil DNA
extraction method. Specifically, the so-called Powersoil (P) method was shown to directly
extract large amounts of DNA of sufficient purity from the three soils, enabling direct PCR-
based molecular analyses. Across methods and samples, the superior yields produced by
method P over the enzymatic lysis based C (CIAT) method confirmed previous data on soil
DNA extraction biases (46, 57, 59). Method P was thus suggested to offer a good choice
for the purpose of the current study in two soils, as it quite consistently yielded high
amounts of DNA and the resulting PCR-DGGE patterns revealed high apparent bacterial
diversities.
136
Is PCR-DGGE sensitive enough to show the effect of environmental factors on the soil bacterial community?
During the work, PCR-DGGE was used as a standard tool to assess the soil microbial
community structures across soils and potato cultivars. In previous studies, PCR-DGGE had
been shown to represent a reliable tool for the assessment of differences or changes in
microbial community structures, especially when those occur in the most abundant
organisms (34, 60). On the other hand, there are limitations of the approach, as it detects
primarily the most abundant ribotypes (62). Moreover, the possible existence of multiple
gene copies in a single organism could present problems of interpretation. As our research
questions addressed the relative changes in the community structures of dominant
organisms, these caveats were considered to not strongly influence the overall results and
interpretations obtained in the study. Indeed, assays like PCR-DGGE have previously
proven to be sensitive enough to reveal changes in a soil community as a result of a
perturbation (13, 61, 87). In the current study, PCR-DGGE also provided a useful indicator
of changes arising from different environmental factors, i.e. the presence of a rhizosphere,
the plant growth stage and time in the season. Indeed, our aim was to find indicators that
are sensitive to such environmental impacts, allowing us to monitor the cropping system
in respect of influences from the plant (cultivar) and the environment.
What are the major drivers of the bacterial communities associated with different potato cultivars?
As indicated earlier, soil bacterial communities are affected by factors like soil
characteristics, environmental conditions and crop management practices like rotation
and crop residue removal (30, 54). This complexity of factors is almost prohibitive to a
detailed understanding of the impact of each single factor. In some cases, plant species
type may have a greater influence on the bacterial community composition of a soil than
soil type (31, 95). On the other hand, the effect of soil type on the bacterial community
may be superior to the effects from plant species type in other cases (24, 86). In Chapters
3-5 and 7, we assessed the dynamics in the abundance, diversity and community structure
of selected soil bacterial communities as a function of plant cultivar type, plant growth
stage and soil type, using six potato cultivars in two soils of different texture.
Accordingly, in Chapter 3 the bacterial community changes were observed over
time in the bulk soil of two fields (B-low organic, V-high organic). Such community changes
in bulk soil, especially between the pre-planting and young plant stages, have previously
been suggested to occur (44). Interestingly, in our study more drastic changes were
observed in the V soil after one year. We cogitated that this change might have been
caused by the heavy water accumulation in this field the year after harvesting, which was
related to a water-impermeable layer that was present at about 80 cm depth in this soil
(Chapter 3). Thus, in this particular case, the changes in the bacterial community
structures that were observed might be explained by differences in water contents and
oxygen limitations that had occurred during the water logging.
137
Besides, both the B and V fields were under crop rotation. The B soil was under
short rotation, including barley and potato. In order to better understand the effect of
seasonal variation on the bacterial diversity in the B soil, clone libraries of bulk soils from
six different time points (spanning three years) were compared. The time points were
April 2008- before potato planting, May 2008 – young potato plant, June 2008 – flowering,
September 2008 - senescence, December 2008 – after removal of plants, July 2009 –
barley flowering, May 2010 - young potato plant, June 2010 – flowering and September
2010 – senescence (Chapter 7). The six libraries varied in the estimates of bacterial
richness based on the CHAO1 richness estimator. Specifically, the richness in the bulk soil -
December 2008 sample revealed the highest value (86), whereas bulk soil collected in
June 2008 had the lowest richness estimation (49). Overall, the libraries had not sampled
the extant diversity to completion. Furthermore, the data showed that the soil bacterial
community make-up was changing over the time as the concomitant PCR-DGGE analyses
revealed the occurrence of shifted patterns. Since each band in such pattern might
represent a bacterial species, the appearance of different species was time-dependent.
What implications these differences and changes have for soil functioning is as yet
unknown. In soil, a range of species may contribute, in equivalent or different ways, to a
particular soil function. This phenomenon has been denoted “functional redundancy”.
Thus, one species may substitute for another one that eventually got lost due to a local
perturbation. Although for many soil functions there is no clear evidence of any
relationship between soil functioning and bacterial diversity, there may be impacts of the
shifts. For instance, in our work phyla like the Proteobacteria showed trends in becoming
more, or less, dominant over time (for example increasing from the flowering to the
senescence stages in the rhizosphere). These changes might be important in functional or
stability terms, but an assessment of this possible linkage to function or resilience has to
await further work. It is logical to posit here that the observed shifts resulted from the
combined effects of the presence of a plant, the plant type and the reigning
environmental conditions such as temperature and moisture.
Moreover, soil type was found to be a key determinant of the bacterial community
composition in the rhizospheres of the potato plants that were grown across soils, as
evidenced by PCR-DGGE and phenotype microarray analysis (Chapters 3 and 5). For
instance, when analyzing the effect of the different cultivars on the bacterial abundance
and community structure, significant differences were observed between the
communities in both fields at all growth stages. Expectedly, increases were noted in the
bacterial abundances in the rhizospheres of all potato cultivars in comparison to those in
the bulk soil. In the V soil, however, the bacterial abundance was not strongly affected by
the presence of a potato plant, and it remained roughly constant over the growth season.
It is possible that, due to the higher nutritional status (OM content and level of other
nutrients), the bacterial abundance in the V soil was affected to a much lower extent by
any root exudates that eventually became available (Chapter 3).
In Chapter 5, the microbial communities from selected samples were analyzed in
phenotype microarrays for their capacities to use different carbon, sulfur and phosphorus
sources. PCR-DGGE analysis was then applied to fingerprint those communities that
allowed high consumption of carbon and sulfur sources. Clearly, soil type affected the
structure of the communities that were active on the selected sulfur and carbon sources.
138
In previous studies, it was also shown that soil type is a key factor that determines the
bacterial community composition in the rhizospheres of plants grown across soils (10, 29).
In the current study, an overall main finding was that soil type exerted the most profound
influence on the analyzed functional bacterial communities in both bulk and rhizosphere
settings. This clearly revealed that the soil bacterial communities differed between the
two soils analyzed and that the communities in the respective rhizospheres were also
different, most likely as a result of the selection of different bacterial consortia by the
same plants growing in different soils.
Indeed, plants are thought to selectively attract particular soil microorganisms to
their rhizospheres, under which avid consumers of root-excreted compounds may be
dominant (13, 64). The abundance and make-up of the rhizosphere bacterial communities
are also known to be governed by the often complex interactions of soil type, plant
species (genotypes) and plant growth (48, 51, 55, 56, 89). In our observations, the
rhizosphere bacterial and betaproteobacterial communities were indeed significantly
different from their respective bulk soil counterparts (Chapter 3). Besides, the plant
growth stages also strongly influenced the community make-ups. Root exudation patterns
may change over time, inciting changes in the microbial community composition in the
rhizosphere (19). In Chapter 5, we found that the bacterial communities from the B and V
bulk soils with plants in the senescence stage either consumed different C sources than
those from the corresponding rhizospheres or the utilization rates were lower than those
in the latter. However, in this study, potential rather than actual utilization of each
substrate was measured, which may only remotely reflect what is going on in situ.
Furthermore, the PCR-DGGE bacterial community analyses that were performed on
selected substrates of the phenotype microarray revealed the bulk soil communities to
cluster apart from the corresponding rhizosphere ones. Some of the compounds that were
analyzed are possibly important as key determinants of plant-bacterium interactions and
these were therefore selected for closer analysis (77). For instance, L-malic acid is known
to be secreted by the roots of varied plants and apparently provides an effective signaling
molecule that modulates the establishment of beneficial rhizobacteria in the plant
rhizosphere, suggesting it has a regulatory role (77). Besides, α-ketobutyric acid, D-alanine
and L-ornithine have also been found to be specific to the rhizosphere, in this case of
white potato (26). In our study, L-malic acid, next to three other compounds i.e., L-aspartic
acid, γ-amino butyric acid and glycyl-L-proline, might indeed be a specific modulator of the
potato root-associated bacterial communities, since organisms with clear utilization
capacities were apparently selected by the rhizosphere.
Also, the amount and types of compounds in the root exudates might show
genotype-specific variations. Different potato cultivars with different rates of growth and
root development are likely to release organic compounds to different extents, and the
bacterial populations in the rhizospheres of the two cultivar groups with different tuber
starch contents likely consisted of species that utilize different carbon sources. A plant
genotype-specific selective effect of plant root type on the root-associated rhizobacterial
community structures was also observed in previous studies (24, 25, 56, 71). In order to
better understand the effects of the differences in plant physiology (related to tuber
starch content and root development) on the rhizospheric bacterial diversity, clone
libraries from the rhizospheres of two cultivars growing in the B soil during flowering stage
139
were compared to those of the respective bulk soil (Chapter 3). Surprisingly, the three
libraries varied remarkably in the estimated diversity of the Betaproteobacteria.
Specifically, the rhizosphere of the cultivar with high-starch-content tubers (Aveka)
showed the highest betaproteobacterial diversity, whereas the communities associated
with the low-starch-tuber cultivar (Premiere) were the least diverse. We currently ignore
why the two cultivars apparently selected communities with such divergent diversities and
can only speculate on how niches might be shaped by both.
Besides, to allow a more high-throughput analysis of the soil bacterial communities,
we used, as a second approach, soil DNA-based pyrosequencing to assess the bacterial
community dynamics (Chapter 4). Around 350,000 sequences were obtained (5,700 to
38,000 per sample). The cultivars analyzed were grouped based on their starch contents
(A, Av, K- high starch tuber, P and D- low starch tuber). The bacterial community make-up
changed in conjunction with the tuber starch content of the host plants. In some cases,
even the rhizosphere effect was different based on plant tuber starch content and,
presumably, plant physiology. In this study, the communities at the GM plant (M) were
compared against the collective data. Interestingly, no significant differentiating effect on
the bacterial community was found for cultivar M. To be able to weigh the effect of
cultivar M against the NOR defined by the communities in the rhizosphere of all cultivars
and bulk soil over three growth stages, the borders of the NOR were established by the
maxima and minima, i.e. the upper (75%) and the lower (25%) percentiles of the relative
abundances of phyla and/or classes. This was accomplished using the average of five
cultivars, GM plant and bulk soil, including all sampling times (Chapter 4). From this
assessment, the values obtained for the GM plant fitted in the overall NOR (in relation to
the rhizosphere or the bulk soil). Strikingly, here, the fluctuations of Betaproteobacteria
and of different orders and genera within this class, observed at the GM plant, slightly
exceeded the NOR for the collective rhizospheres (Fig. 1). Here, orders or genera that are
important for soil functioning were chosen for the analyses. Unfortunately, the data
revealed many unclassified bacteria and, in addition, important genera such Polaromonas
and Acidovorax were not detectable. Given the foregoing, the order Burkholderiales and
the genera Burkholderia, Variovorax, Achromobacter and Nitrosospira were chosen to
serve as potential key indicators (Chapter 4). The data from the clone library analysis
(Chapter 6) and pyrosequencing (Chapter 4) were thus examined to establish the NOR
with respect to these taxa. Since the data obtained from the clone library were (five- to
seven-fold) lower than those of the pyrosequencing (indicating a possible cloning bias),
data from clone library for Betaproteobacteria and Burkholderiales were corrected with
factors 7 and 5 respectively, to allow comparison with the pyrosequencing data. The
resulting NOR for the various taxonomic levels is depicted in figure 1. Considerable
variation in the fractions over the total of the different taxonomic levels was observed in
the potato cropping system over time.
From the results obtained, we distill that the Betaproteobacteria and subgroups
therein might serve as indicator classes for the establishment of a NOR (baseline) of key
bacterial species in soil under potato. The main argument supporting this thesis is that
many members of the betaproteobacterial group are important mediators in steps of
the cycling of nitrogen, sulfur and carbon through the soil ecosystem (43, 81). Secondly,
they were found to be sensitive to shifts induced by potato cultivar type. Not only in the
140
work done under Chapter 4, but also in that of Chapter 6, it was shown that the
community make-up of the Betaproteobacteria is strongly responsive to the presence of
plants, which relates to cultivar type and plant growth stage. The study on potential
function (Chapter 5) also showed a clear rhizosphere and cultivar effect on the
betaproteobacterial communities. However, we could not discern a particular effect of the
GM plant. Thus, cultivar M, as compared to the other cultivars (including K), did not exert
any specific effect on the utilization patterns of the carbon, phosphorus and sulfur sources
by its associated microflora. The responses of the soil/rhizosphere microbial communities
to these nutrient sources were investigated individually. However, one needs to realize
that root exudates normally contain diverse compounds in fluctuating concentrations in
an integrated environment. Hence, due to the resulting complexity, pinpointing the effects
on the real in situ microbial populations at the potato root surface is inherently difficult.
What about the unclassified bacteria?
Pyrosequencing offered us a chance to explore the bacterial community and their
interactions at a more thorough level. However, a large fraction of the soil bacterial
communities studied was found to abide in unknown territory, much as what was found in
the clone libraries. The DNA obtained from the bacterial community in the bulk soil
collected during the young plant and flowering stages was actually dominated by
sequences of as-yet-unclassified bacteria (15-35%). Hence, this part of the soil community
remains a challenging reservoir of novel bacterial diversity. Subsamples, consisting of 100
sequences each, were taken from the as-yet-unclassified sequences of three bulk and
three rhizosphere soil samples. Per subsample (soil or rhizosphere), phylogenetic trees
were built and the clustering was analyzed. In all cases, most (>95%) of the sequences fell
in 7-10 branches, in which individual reads often showed deep branching. “Flat” branches
containing more than 5 sequences were never observed using the 97% cut-off level,
indicating that none of the tested sequences showed overall dominance (i.e. roughly >
1.3% of the total). High percentages of the as-yet-unclassified bacteria have also been
found in other relevant studies (2, 89). There is a strong need to identify such as-yet-
unclassified bacteria and ideally organisms should be cultured in order to start to
understand their ecology. This pleas strongly for the position that the new and highly
powerful culture-independent techniques should go hand-in-hand with advanced culture-
dependent ones.
141
0
0,05
0,1
0,15
0,2
0,25
0,3
0,35
0 1 2 3 4 5 6
relative abundance
of V
ariovorax [%
]
bulk rhizosphere GM
0
1
2
3
4
5
6
0 5 10 15 20 25 30
relative abundance of Betaproteobacteria [%]
bulk-pyroseq
rhizosphere-pyroseq
GM-pyroseq
clone library
0
1
2
3
4
5
0 5 10 15 20 25 30
relative abundance of Burkholderiales [%]
bulk-pyroseq
rhizosphere-pyroseq
GM-pyroseq
clone library
A
B
C
0
0,05
0,1
0,15
0,2
0,25
0,3
0,35
0 1 2 3 4 5 6
relative abundance
of V
ariovorax [%
]
bulk rhizosphere GM
0
1
2
3
4
5
6
0 5 10 15 20 25 30
relative abundance of Betaproteobacteria [%]
bulk-pyroseq
rhizosphere-pyroseq
GM-pyroseq
clone library
0
1
2
3
4
5
0 5 10 15 20 25 30
relative abundance of Burkholderiales [%]
bulk-pyroseq
rhizosphere-pyroseq
GM-pyroseq
clone library
A
B
C
142
Figure 1 - Normal
operating range
(NOR) of the bulk and
rhizosphere soil
against the
genetically modified
plant (M) across the
season 2008 for (A)
Betaproteobacteria
(B) Burkholderiales
(C) Variovorax. (D)
Nitrosospira (E)
Burkholderia (F)
Achromobacter. 1-
before planting (April
2008) 2- young plant
potato (May 2008), 3-
flowering (June
2008), 6-senescence
(September 2008), 9-
after removal of
plants (December
2008), 15 – barley
during flowering of
potato plants (June
2009), 28 – flowering
(July 2010), 30 –
senescence
(September 2010).
Rhizosphere indicates
the average of 5
cultivars. Blue lines
indicate the NOR for
bulk soil. The pink
area represents the
NOR for rhizosphere
and the green area
represents the range
for the GM plant
(Chapter 4). Data in
(A) and (B) from
clone libraries were
harmonized with
number 7 and 5
respectively.
0,01
0,02
0,03
0,04
0,05
0,06
0,07
0 1 2 3 4 5 6
relative abundance of Nitrosospira [%
]
bulk rhizosphere GM
0,00
0,05
0,10
0,15
0,20
0,25
0,30
0 1 2 3 4 5 6
Relative abundance of Burkholderia [%
]
bulk rhizosphere GM
0,00
0,02
0,04
0,06
0,08
0,10
0 1 2 3 4 5 6
Relative abundance of Achromobacter [%]
bulk rhizosphere GM
D
F
E
0,01
0,02
0,03
0,04
0,05
0,06
0,07
0 1 2 3 4 5 6
relative abundance of Nitrosospira [%
]
bulk rhizosphere GM
0,00
0,05
0,10
0,15
0,20
0,25
0,30
0 1 2 3 4 5 6
Relative abundance of Burkholderia [%
]
bulk rhizosphere GM
0,00
0,02
0,04
0,06
0,08
0,10
0 1 2 3 4 5 6
Relative abundance of Achromobacter [%]
bulk rhizosphere GM
D
F
E
143
Can microbial diversity be related to ecological behavior?
The large pyrosequencing-based data set also gave us the opportunity to assess the
ecological behavior, in terms of the dynamics of relative abundance, of particular pre-
specified bacterial phyla and classes. Here, I would like to posit that the general
prevalence of many members of, e.g., the Acidobacteria in bulk soils (as opposed to
rhizosphere soils) may relate to their generally oligotrophic (K strategist) lifestyle (22).
Acidobacterium has appeared as a highly recalcitrant phylum for a long time, in terms of
understanding behavior, due to the difficulty of isolation of its member strains. The
phylum consists of a wide range of species, with presumably diverse ecological behavior
(65). Contradictory information has been provided on the occurrence of different
members of the Acidobacteria in the plant-soil environment (38, 40, 65). In contrast, many
of the Betaproteobacteria (22) as well as Pseudomonas species (88) are known as typical
copiotrophs (r strategists) which tend to be strongly favored in soil under nutrient-rich
conditions. It is possible that each plant growth stage is characterized by a specific but
different root exudation pattern which drives different bacterial communities. During the
young, flowering and senescence stages of plant growth, the rhizosphere may offer
varying niches that are characterized by different levels and compositions of root
exudates. Besides, in the bulk soil, the early (young) growth stage was preferred by
Acidobacteria (oligotroph) whereas the late (senescence) stage was abundant with
copiotrophs (e.g. Beta-, Alpha-proteobacteria and Pseudomonas). An explanation might be
that, in the senescence stage, the bulk soil experienced more pronounced nutrient
supplies, in this case from senescent (“leaky”) plant roots than at earlier stages. Thus, the
root tissue may have started to enrich the surrounding soil environment with nutrients. In
our observations, Pseudomonas was found in similar amounts across all potato cultivars
(so independently of cultivar type), whereas the prevalence of the class
Betaproteobacteria was likely cultivar-dependent. The latter might also provide an
argument in favor of the use of Betaproteobacteria as a group of indicator organisms.
In a recent opinion paper by Philippot et al. (69), it was claimed that many high
taxonomic levels of bacteria show 'ecological coherence' (22, 70). It was argued that the
ecological coherence within a taxon is greater than across taxa. Ecological coherence here
is the fact that members of a given taxon share general life strategies or traits that
distinguish them from members of other taxa (69). The authors assumed that during the
course of evolution, adaptations to different ecological niches have shaped and
differentiated bacterial lineages, and then the divergences are reflected not only at the
strain or species level but also or mainly at higher taxonomic ranks. There are studies that
support the idea of ecological coherence of bacterial groups at taxonomic ranks higher
than the species (8, 14, 45, 88). Other studies report either site- or time-specific
community structures at high taxonomic levels (9, 23, 32), as well as differential responses
of bacterial lineages to changing environmental factors (9, 37). On the other hand,
Philippot et al suggested to keep in mind the imperfection in recognizing ecological
cohesiveness of high bacterial taxa; they advocated a focus on unifying principles (69).
One might compare this to the dietary preferences of people based on regional
conditions. For instance, if you check different regions of Turkey, the climate and land
144
conditions define the dietary habits of people. In the southeast, people have meat- and
grain-based diets, whereas the diet is more fish-based in the north. In the Netherlands,
diets are completely different and based mainly on potato. However, there are also
exceptions, like in a country where they use hot spices. There might be people who do not
like the spices, the other way around for countries which do not know any spices other
than salt. These examples lead us to general dietary predictions, based on region. In soil,
one might cogitate the existence of such local diet-determined communities: strains of the
same taxon might show similar behaviour as determined by site. Thus, once the ecology of
single species can be described, one might start to discern unified ecological traits at
higher ranks.
How does the level of aromatic sulfonates affect the bacterial community in soil?
In Chapter 6, the effect of different concentrations of linear alkyl benzenesulfonate (LAS)
and sulfate in soil on which the potato cultivars Aveka (high starch tuber) and Premiere
(low starch tuber) were growing, was studied in a microcosm experiment. The idea was
that (1) soils may become deprived of inorganic sulfur turning the transformation of other
(organic) sulfur sources indispensable for microbes and plants, (2) aromatic sulfonates in
soil are potentially important sulfur sources for plant nutrition and (3) many organic sulfur
sources are biodegraded by microorganisms in the soil. Therefore, the degradation of LAS
by particular microbial desulfonators in soil was predicted, however their ecology was
unknown.
In previous work, changes in the community structures of Alphaproteobacteria and
Actinobacteria in bulk soil as a result of the addition of LAS (78) have been noted.
Moreover, short-term effects of sulfate on the diversity of Betaproteobacteria in the
Agrostis stolonifera rhizosphere have also been shown (83). However, other studies have
shown no effects of added LAS on the diversity or community structures of bacteria in
soils (7, 78, 92). In the current study, for the first time, an effect of LAS on the soil
betaproteobacterial community make-up was revealed.
We also found that there are desulfonators active under sulfate-containing
conditions in soil. The desulfonating enzymes produced by such organisms are often not
constitutive, but only become expressed under low-sulfate conditions as part of a broader
sulfate starvation response (39). On the other hand, desulfonation of aromatic sulfonates
has been found to occur in natural soil (81). Also, a previous study suggested that some
desulfonating bacteria, evidenced via detection of Cupriavidus sp. like asfA sequences, can
also take advantage of elevated sulfate levels (83). In our work, we did find rapid
desulfonation in the sulfate-containing soil. The sulfate level in this soil had initially been
underestimated (performed using water extraction). How can we explain the
simultaneous occurrence of desulfonation and sulfate? Possibly, the sulfate may have
been heterogeneously distributed. Hence, it might have been less available to local
desulfonators, which perceived LAS as their prime sulfur (and/or carbon) source.
Together, these factors might have resulted in the activation of desulfonators in sulfate-
containing soil.
145
Overall, our results suggest that particular bacterial phylotypes that are involved in
desulfonation and/or carbon mineralization quickly utilized LAS in the soil. The
composition of this desulfonating guild is utterly unknown, but Variovorax paradoxus may
have played a role – and thus become selected – at high LAS levels in the potato
rhizosphere, as was clearly indicated from the clone library analysis for LAS-50-treated
rhizosphere in the flowering stage.
Concluding remarks
Since the ecosystems on Earth differ in their buffering capacity against perturbation as a
function of biotic and abiotic factors, natural variations within a system inciting
community changes are to be considered as determinants of the NOR of that system (41).
This argument holds for both natural and man-made systems, such as agricultural ones. In
the latter, effects of all (agronomic) measures that are normal in the system need to be
included in the NOR. We here argued this NOR should place a strong focus on the soil
microbiota, which underpins soil functioning. Afterwards, the NOR can be used to assess
the effect of a GM plant on the soil microbial community. In our study, we used the GM
cultivar M, which has been derived from the parent cultivar K. Cultivar K was thus
modified to prevent the formation of amylase in the tubers. We cogitated that the
modification might have altered the root exudation pattern of cultivar M. To allow
assessing the putative effects of the modification, its effects are ideally weighed against a
NOR that was built on the variation among the other (unmodified) cultivars used in this
study. A baseline was thus built up with selected potato cultivars that differed in the
starch content of their tubers. Overall, the facet tuber starch content turned out to be an
important factor impacting on the community structure of the potato associated
microbiota. Overall, most assessments performed showed the absence of a significant
effect of the modification that gave origin to cultivar M. In fact, the collective data
obtained with all cultivars established the NOR of potato cropping in the used soils. The
putative effects of future genetic modifications in potato should be weighed against the
NOR provided by this suite of cultivars.
However, the assessed NOR may not be of use for another crop or even for same
crop in different soils, since the environmental factors may be too varied. For instance,
one study assessed the community structure of the microbiota in agricultural soil under
four systems and found Actinobacteria, Bacteriodetes and Firmicutes to be the most
dominant phyla (2). In our study, Actinobacteria and Alphaproteobacteria comprised the
most dominant phylum and class, respectively, whereas the prevalences of members of
the Bacteriodetes and Firmicutes were found to be relatively low. This appears consistent
with the contention that each soil may have its own characteristic community make-up
(28). Moreover, another study from Acosta-Martinez (1) showed that the effects of
grazing on pasture by cattle were significant for the Actinobacteria, Proteobacteria and
Chlorofexi, indicating that the particular use of a soil can affect particular bacterial groups
in a soil. Besides, soil pH was shown to explain a significant portion of the variability
associated with changes in the community structures that were observed (63), indicating
the key role exerted by pH in shaping the community make-up of soil.
146
Growing GM or conventional plants in the field has raised concerns about the
potential risk of effects on non-target organisms. The possibility of long-term effects from
GM plants obviously cannot be excluded and must be examined on a case-by-case basis.
There are currently a range of studies on the effects of differently-modified plants on the
soil microbial community (3, 6, 17, 19-21, 27, 33, 48, 50, 57, 66, 67, 72, 73, 79, 82, 85, 93).
In certain cases, the degree of difference between the modified and parent cultivar (or
any other cultivar of the same plant species) was not defined (35, 58). In regulatory terms,
a GM plant is ideally characterized as “substantially equivalent” (with the exception of the
modification) to its parent, and concerns are whether there is any reason to suppose it
poses an unacceptable risk to soil health, including the soil microbiota and its functioning
(47). One key bacterial group involved in soil functioning is represented by the
Betaproteobacteria, as members of this group are important mediators in key steps of
the cycling of nitrogen, sulfur and carbon through soil (43, 81). For instance, the
Nitrosomonadaceae form an important cluster of betaproteobacterial ammonia oxidizers,
whereas Burkholderia types play important roles in soil transformations, the
mycorrhization of plants as well as in symbiotic nitrogen fixation (12, 75). Other members
of the Betaproteobacteria promote plant growth by virtue of their synthesis of
phytohormones and vitamins (18). For instance, B. phytofirmans typically produces
aminocyclopropane-1-carboxylate (ACC) deaminase, which assists in the lowering of the
level of the stress hormone ethylene at plants. Accordingly, the growth of plant roots may
increase when B. phytofirmans is present, by virtue of the reduction of this root
elongation inhibitor (84). Furthermore, other Burkholderia species are important
producers of antibiotics that antagonize bacterial and fungal phytopathogens (52).
Recently, the betaproteobacterial genera Variovorax and Polaromonas have been found
to be capable of desulfonating aromatic sulfonates in the wheat rhizosphere (80).
Encouragingly, our study showed that several members of the Betaproteobacteria are
sensitive to effects from different potato cultivars as well as soil and time, allowing their
potential use as bioindicators that define the NOR (baseline) for potato cropping.
Moreover, in our study, high-throughput pyrosequencing on the basis of directly
extracted soil DNA was used. In spite of its limitations, e.g. read length (76), it gave us an
angle at the entire microbial community, and showed the clear effect of cultivar type per
different level of taxa. This also enabled to see the ecological coherence of different
taxonomic levels.
Plant-soil systems are highly heterogeneous and dynamic, and here we supplied
data concerning the NOR of soil microbial community properties in response to potato
plant-induced variables. The effects of the GM plant were minor compared with the
variation seen across all cultivars or those associated with season. Concerning potato
cropping, we advocate the use of particular Betaproteobacteria as sensitive indicators for
the assessment of the quality of the potato-cropped soils (47). Thus, members of the soil
Betaproteobacteria can be monitored to assess perturbations, resulting in a framework
that may monitor changes and provide a framework for risk assessment. Furthermore, in
the current study, the asfA gene of species belonging to the Betaproteobacteria was also
found to be “sensitive” to soil type, the presence of a plant and plant growth stage.
However, it needs further study to assess the validity of the use of asfA as an indicator. In
this respect, in case of detection of a change, the question is how we should handle it
147
(42). This will require a strengthening of the theoretical link between the soil microbial
community structure and its functions, including the aspect of resilience/resistance of the
system (42).
References
1. Acosta-Martínez, V., S. E. Dowd, Y. Sun, D. Wester, and V. Allen. 2010. Pyrosequencing analysis
for characterization of soil bacterial populations as affected by an integrated livestock-cotton
production system. Appl. Soil Ecol. 45:13-25.
2. Acosta-Martinez, V., K. S, S. E. Dowd, Y. Sun, and V. Allen. 2008. Tag encoded pyrosequencing
analysis of bacterial diversity in a single soil type as affacted by management and land use. Soil Biol.
Biochem. 40:2762-2770.
3. Andreote, F., R. Mendes, F. Dini-Andreote, R. P.B., C.A. Labate, A.A. Pizzirani-Kleiner, J.D. van Elsas, J.L. Azevedo, and W.L. Araújo. 2008. Transgenic tobacco revealing altered bacterial diversity
in the rhizosphere during early plant development. Antonie Van Leeuwenhoek. 93:415-424.
4. Ashelford, K. 2005. At least 1 in 20 16s rRNA sequence records currently held in public
repositories in estimated to contain substantial anomalies. Appl. Environ. Microbiol. 71:7724-7736.
5. Bachoon, D., E. Otero, and R. Hodson. 2001. Effects of humic substances on fluorometric DNA
quantification and DNA hybridization. J. Microbiol. Methods. 47:73-82.
6. Becker, R., U. Behrendt, B. Hommel, S. Kropf, and A. Ulrich. 2008. Effects of transgenic fructan-
producing potatoes on the community structure of rhizosphere and phyllosphere bacteria. FEMS
Microbiol. Ecol. 66:411-425. doi: 10.1111/j.1574-6941.2008.00562.x.
7. Brandt, K. K., N. O. G. Jørgensen, T. H. Nielsen, and A. Winding. 2004. Microbial community-level
toxicity testing of linear alkylbenzene sulfonates in aquatic microcosms. FEMS Microbiol.Ecol.
49:229-241.
8. Bruce C., T., O. Nick, M. Niall, B. Mark J., W. Andrew S., and W. Andrew. 2009. Vegetation affects
the relative abundances of dominant soil bacterial taxa and soil respiration rates in an upland
grassland soil. Microbial Ecology. 59:335-343.
9. Buckley, D. H., and T. M. Schmidt. 2001. The structure of microbial communities in soil and the
lasting impact of cultivation. Microbial Ecology. 52:11-21.
10. Buyer, J. S., D. P. Roberts, and E. Russek-Cohen. 2002. Soil and plant effects on microbial
community structure. Can. J. Microbiol. 48:955-964.
11. Carney, K., and P. Matson. 2005. Plant communities, soil microorganisms and soil carbon
cycling: does altering the world below ground matter to ecosysyem functioning? Ecosystems. 9:928-
940.
12. Chen, W., L. Moulin, C. Bontemps, P. Vandamme, G. Bena, and C. Boivin-Masson. 2003.
Legume symbiotic nitrogen fixation by beta-proteobacterial is widespread in nature. J. Bacterio.
185:7266-72.
13. Costa, R., M. Götz, N. Mrotzek, K. Lottmann, G. Berg, and K. Smalla. 2006. Effects of site and
plant species on rhizosphere community structure as revealed by molecular analysis of microbial
guild. FEMS Microbiol.Ecol. 56:236-249.
14. Cruz-Martínez, K., K. B. Suttle, E. L. Brodie, M. E. Power, G. L. Andersen, and J. F. Banfield. 2009. Despite strong seasonal responses, soil microbial consortia are more resilient to long-term
changes in rainfall than overlying grassland. ISME J. 3:738-744.
15. Dalmastri, C., L. Chiarini, C. Cantale, A. Bevivino, and S. Tabacchioni. 1999. Soil type and maize
cultivar affect the genetic diversity of maize root-associated Bulkoderia cepacia populations.
Microbial Ecology. 38:273-284.
148
16. de Vetten, N., A. Wolters, K. Raemakers, I. van der Meer, R. ter Stege, E. Heeres, P. Heeres, and R. Visser. 2003. A transformation method for obtaining marker-free plants of a cross-pollinating and
vegatatively propagated crop. Nature Biotech. 21:439-442.
17. Di Giovanni, G. D., L. S. Watrud, R. J. Seidler, and F. Widmer. 1999. Comparison of Parental and
Transgenic Alfalfa Rhizosphere Bacterial Communities Using Biolog GN Metabolic Fingerprinting and
Enterobacterial Repetitive Intergenic Consensus Sequence-PCR (ERIC-PCR). Microbial Ecology.
37:129-139.
18. Duineveld, B., A. Rosado, J. D. van Elsas, and J. A. van Veen. 1998. Analysis of dynamics of
bacterial communities in the rhizosphere of chrysanthemum via DGGE and substrate utilization
patterns. Appl. Environ. Microbiol. 64:4950-4957.
19. Dunfield, K., and J. Germida. 2003. Seasonal changes in the rhizosphere microbial communities
associated with field-grown genetically modified canola. Appl. Environ. Microbiol. 69:7310-7318.
20. Dunfield, K., and J. Germida. 2004. impact of genetically modified crops on soil- and plant-
associated microbial communities. J Environ Qual. 33:806-815.
21. Fang, M., J. Kremer, P. Motavalli, and G. Davis. 2005. Bacterial diversity in rhizosphere of
nontransgenic and transgenic corn. Appl. Environ. Microbiol. 71:4132-4136.
22. Fierer, N., M. A. Bradford, and R. B. Jackson. 2007. Toward an ecological classification of soil
bacteria. Ecology. 88:1354-1364.
23. Fulthorpe, R. R., L. F. W. Roesch, A. Riva, and E. W. Triplett. 2008. Distantly sampled soils carry
few species in common. ISME J. 2:901-910.
24. Garbeva, P., J. Postma, J. A. van Veen, and J. D. van Elsas. 2006. Effect of above ground plant
species on soil microbial community structure and its impact on supply of Rhizoctonia solani AG3.
Environ. Microbiol. 8:233-246.
25. Garbeva, P., J. A. van Veen, and J. D. van Elsas. 2004. Microbial diversity in soil: selection of
microbial population by plant and soil type and implications for disease suppressiveness. Ann. Rev.
Phytopat. 42:243.
26. Garland, J. L. 1996. Patterns of potential C source utilization by rhizosphere communities. Soil
Biol. Biochem. 28:223-230.
27. Gebhard, F., and S. K. 1999. Monitoring filed releases of genetically modified sugar beets for
persistence of transgenic plant DNA and horizontal gene transfer. FEMS Microbiol.Ecol. 28:261-272.
28. Gelsomino, A., A. Keijzer-Wolters, G. Cacco, and J. D. van Elsas. 1999. Assessment of bacterial
community structure in soil by PCR and DGGE. J. Microbiol. Methods. 38:1-15.
29. Girvan, M. S., J. Bullimore, J. N. Pretty, A. M. Osborn, and A. S. Ball. 2003. Soil Type Is the
Primary Determinant of the Composition of the Total and Active Bacterial Communities in Arable
Soils. Appl. Environ. Microbiol. 69:1800-1809. doi: 10.1128/aem.69.3.1800-1809.2003.
30. Govaerts, B., M. Mezzalama, Y. Unno, K. Sayre, M. Luna-Guido, K. Vanherck, L. Dendooven, and J. Deckers. 2007. Influence of tillage, residue management, and crop rotation on soil microbial
biomass and catabolic diversity. Appl. Soil Ecol. 37:18-30.
31. Grayston, S. J., S. Wang, C. D. Campbell, and A. C. Edwards. 1998. Selective influence of plant
species on microbial diversity in the rhizosphere. Soil Biol. Biochem. 30:369-378.
32. Hackl, E., S. Zechmeister-Boltenstern, L. Bodrossy, and A. Sessitsch. 2004. Comparison of
Diversities and Compositions of Bacterial Populations Inhabiting Natural Forest Soils. Appl. Environ.
Microbiol. 70:5057-5065. doi: 10.1128/AEM.70.9.5057-5065.2004.
33. Hannula, S. E., W. de Boer, and J. A. van Veen. 2010. In situ dynamics of soil fungal communities
under different genotypes of potato, including a genetically modified cultivar. Soil Biol. Biochem.
42:2211-2223. doi: DOI: 10.1016/j.soilbio.2010.08.020.
34. Heuer, H., G. Wieland, J. Schönfeld, A. Schönwälder, N. Gomes, and K. Smalla. 2001. Bacterial
community profiling using DGGE and TGGE analysis, p. 177-190. In Anonymous Environmental
Molecular Microbiology: Protocols and Applications.
149
35. Hilbeck, A., M. Meier, J Römbke, J. S, H Teichmann, and T. B. 2011. Environmental risk
assessment of genetically modified plants - concepts and controversies. Environ. Scie. Eur. 23:1.
36. Hu, Z., Z. Yang, C. Xu, S. Haneklaus, Z. Cao, and E. Schnug. 2002. Effect of crop growth on the
distribution and mineralization of soil sulfur in the rhizosphere. J Plant Nutr. Soil Sci. 165:249-254.
37. Jangid, K., M. A. Williams, A. J. Franzluebbers, J. S. Sanderlin, J. H. Reeves, M. B. Jenkins, D. M. Endale, D. C. Coleman, and W. B. Whitman. 2008. Relative impacts of land-use, management
intensity and fertilization upon soil microbial community structure in agricultural systems. Soil Biol.
Biochem. 40:2843-2853. doi: DOI: 10.1016/j.soilbio.2008.07.030.
38. Jones, R., M. Robenson, C. Lauber, M. Hamady, R. Knight, and N. Fierer. 2009. A comprehensive
survey of soil acidobacterial diversity using pyrosequencing and clone libraries. ISME J. 3:442-453.
39. Kertesz, M. 1999. Riding the sulfur cycle - metabolism of sulfonates and sulfate esters in Gram-
negative bacteria. FEMS Microbiol Rev. 24:135-175.
40. Kielak, A., A. S. Pijl, J. A. van Veen, and G. A. Kowalchuk. 2008. Phylogenetic diversity of
Acidobacteria in a former agricultural soil. ISME J. 3:378-382.
41. Kowalchuk, G., M. Bruinsma, and J. van Veen. 2003. Assessing responses of soil microorganisms
to GM plants. Trends Ecol. Evol. 18:403-410.
42. Kowalchuk, G., W. de Boer, and J. van Veen. 2003. Knowledge gaps with respect to the effect of
genetically modified crops on the functioning of soil ecosystems. COGEM Research. .
43. Kowalchuk, G., J. Stephen, W. de Boer, J. Prosser, T. Embley, and J. Woldendorp. 1997. Analysis
of ammonia-oxidizing bacterial of the ß subdivision of the class Proteobacteria in coastal sand dunes
by DGGE and sequencing of PCR-amplified 16S ribosomal DNA fragements. Appl. Environ. Microbiol.
63:1489-1497.
44. Larkin, R. 2003. Characterization of soil microbial communities under different potato cropping
systems by microbial population dynamics, substrate utilization and fatty acid profiles. Soil Biol.
Biochem. 35:1451-1466.
45. Lauber, C. L., M. Hamady, R. Knight, and N. Fierer. 2009. Pyrosequencing-based assessment of
soil pH as a predictor of soil bacterial community structure at the continental scale. Appl. Environ.
Microbiol. 75:5111-5120. doi: 10.1128/aem.00335-09.
46. Leff, L., J. R. Dana, J. V. McArthur, and L. J. Shimkets. 1995. Comparison of Methods of DNA
Extraction from Stream Sediments. Appl. Environ. Microbiol. 61:1141-1143.
47. Lilley, A. K., M. J. Bailey, Cartwright CL, S. Turner, and P. Hirsch. 2006. Life in earth: the impact
of GM plants on soil ecology? Trends in Biotechol. 24:9-14.
48. Liu, W. 2010. Do genetically modified plants impact arbuscular mycorrhizal fungi? Ecotoxicology.
19:229-238. doi: 10.1007/s10646-009-0423-1.
49. Liu, Z., T. DeSantis, G. ANdersen, and R. Knight. 2008. Accurate taxonomy assignments from 16S
rRNA sequences produced by highly parallel pyrosequencers. Nuc. Acids Res. 36:120.
50. Lottmann, J., H. Heuer, K. Smalla, and G. Berg. 1999. Influence of transgenic T4-lysozyme-
producing potato plants on potentially beneficial plant-associated bacteria. FEMS Microbiol.Ecol.
29:365-377.
51. Lupwayi, N., W. Rice, and G. Clayton. 1998. Soil microbial diversity and community structure
under wheat as influenced by tillage and crop rotation. Soil Biol. Biochem. 30:1733-1741.
52. MacDonald, K., and D. Speert. 2006. Interaction of Burkholderia species with phagocytic
systemIn T. Coenye and P. Vandamme (eds.), Burkholderia molecular microbiology and genomics.
Horizon Bioscience, UK.
53. Manter, D., J. Delgado, D. Holm, and R. Stong. 2010. Pyrosequencing reveals a highly diverse
and cultivar-specific bacterial endophyte community in potato roots. Microbial Ecology. 60:157-166.
54. Mclaughlin, A., and P. Mineau. 1995. The impact of agricultural practices on biodiversity. Agricul
Ecosys Environ. 55:201-212.
150
55. Micallef, S., S. Channer, S. Michael P, and C. Adán. 2009. Plant age and genotype impact the
progression of bacterial community succession in the Arabidopsis rhizosphere. Plant Signal Behav.
4:777-780.
56. Micallef, S., M. Shiaris, and A. Colon-Carmona. 2009. Influence of Arabidopsis thaliana
accessions on rhizobacterial communites and natural variation in root exudates. J Exp Botany.
60:1729-1742.
57. Milling, A., K. Smalla, F. Maidl, M. Schloter, and J. Munch. 2004. Effects of transgenic potatoes
with an altered starch composition on the diversity of soil and rhizosphere bacteria and fungi. Plant
and Soil. 266:23-39.
58. Millstone, E., E. Brunner, and S. Mayer. 1999. Beyond 'substantial equivalence'. Nature.
401:525-526. doi: 10.1038/44006.
59. More, M., J. Herrick, M. Silva, W. Ghiorse, and E. Madsen. 1994. Quantitative cell lysis of
indigenous microorganisms and rapid extraction of microbial DNA from sediment. Appl. Environ.
Microbiol. 60:1572-1580.
60. Muyzer, G., T. Brinkhoff, U. Nübel, C. Santegoeds, H. Schäfer, and C. Waver. 1998. DGGE in
microbial ecology, p. 1-27. In G. Kowalchuk, F. de Bruijn, I. Head, A. Akkermand, and J. van Elsas
(eds.), Molecular Microbial Ecology Manualvol. 3.4.4. Kluwer Ac. Pub., Netherlands.
61. Muyzer, G., E. de Waal, and A. Uitterlinden. 1993. Profiling of complex microbial populations by
DGGE analysis of PCR-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol. 59:695-700.
62. Muyzer, G., and K. Smalla. 1998. Application of DGGE and TGGE in microbial ecology. Antonie
Van Leeuwenhoek. 73:127-141.
63. Nacke, H., Thürmer A, Wollherr A, Will C, and Hodac L. 2011. Pyrosequencing-Based
Assessment of Bacterial Community Structure Along Different Management Types in German Forest
and Grassland Soils. PLoS ONE. 6:e17000.
64. Nannipieri, P., J. Ascher, M. T. Ceccherini, L. Landi, G. Pietramellara, G. Renella, and F. Valori. 2008. Effects of Root Exudates in Microbial Diversity and Activity in Rhizosphere Soils, p. 339-365. In
C. S. Nautiyal and P. Dion (eds.), Molecular Mechanisms of Plant and Microbe Coexistencevol. 15.
Springer, Berlin Heidelberg.
65. Nunes da Rocha, U., J. D. van Elsas, and L. S. van Overbeek. 2010. Real-time PCR detection of
Holophagae (Acidobacteria) and Verrucomicrobia subdivision 1 groups in bulk and leek (Allium
porrum) rhizosphere soils. J. Microbiol. Metods. 83:141-148.
66. Oger, P., P. Annik, and D. Yves. 1997. Genetically engineered plants producing opines alter their
biological environment. Nat. Biotech. 15:369-372.
67. Oliveira, A. P., M. E. Pampulha, and J. P. Bennett. 2008. A two-year field study with transgenic
Bacillus thuringiensis maize: effects on soil microorganisms. Sci. Total Environ. 405:351-357. doi:
10.1016/j.scitotenv.2008.05.046.
68. Petra, M., C. David, and Ching Hong Yang. 2004. Development of specific rhizosphere bacterial
communities in relation to plant species, nutrition and soil type. Plant and Soil. 261:199-208.
69. Philippot, L., S. G. E. Andersson, T. J. Battin, J. I. Prosser, J. P. Schimel, W. B. Whitman, and S. Hallin. 2010. The ecological coherence of high bacterial taxonomic ranks. Nat.Rev.Microbiol. 8:523-
529.
70. Philippot, L., D. Bru, N. P. A. Saby, J. Čuhel, D. Arrouays, M. Šimek, and S. Hallin. 2009. Spatial
patterns of bacterial taxa in nature reflect ecological traits of deep branches of the 16S rRNA
bacterial tree. Environ. Microbiol. 11:3096-3104.
71. Rasche, F., V. Hödl, C. Poll, E. Kandeler, M. Gerzabek, J. D. van Elsas, and A. Sessitsch. 2006.
Rhizosphere bacteria affected by transgenic potatoes with antibacterial activities compared with the
effect of soil. wild type potatoes, vegetation stage and pathogen exposure. FEMS Microbiol.Ecol.
56:219-235.
72. Rasche, F., E. Marco-Noales, H. Velvis, L. S. van Overbeek, M. Lopez, J. D. van Elsas, and A. Sesstisch. 2006. Structural characteristics and plant-beneficial effects of colonizing the shoots of
151
field grown conventional and genetically modified T4-lysozyme producing potatoes. Plant and Soil.
289:123-140.
73. Rasche, F., H. Velvis, C. Zachow, G. Berg, J. D. van Elsas, and A. Sessitsch. 2006. Impact of
transgenic potatoes expressing antibacterial agents on bacterial endophtes is comparable with the
effect of plant genotype, soil type and pathogen infection. J. Appl. Ecol. 43:555-566.
74. Rengel, Z., R. Gutteridge, P. Hirsch, and D. Hornby. 1996. Plant growth, micronutrient
fertilization and take-all infection influence bacterial populations in the rhizosplere of wheat. Plant
and Soil. 183:269-277.
75. Reuus, V., P. Estrada-De los Santos, S. Tenorio-Salgado, V. Baldani, M. Schmid, J. B. J. Balandreau, A. Hartmann, and J. Caballero-Melloado. 2004. Burkholderia tropica sp. nov., a novel
nitrogen-fixing, plant associated bacterium. J. Sys. Evol. Microb. 54:2155-2162.
76. Ronaghi, M. 2001. Pyrosequencing sheds light on DNA sequencing. Genome Research. 11:3-11.
77. Rudrappa, T., K. J. Czymmek, P. W. Pare, and H. P. Bais. 2008. Root-Secreted Malic Acid Recruits
Beneficial Soil Bacteria. Plant Physiol. 148:1547-1556. doi: 10.1104/pp.108.127613.
78. Sanchez-Peinado, M., L. Gonzelaz-Lopez, M. Martinez-Toledo, C. Pozo, and B. Rodelas. 2010.
Influence of LAS on the structure of Alphaproteobacteria, Actinobacteria and Acidobacteria
communities in soil microcosm. Environ Sci Pollut Res. 17:779-790.
79. Sarkar, B., A. K. Patra, T. J. Purakayastha, and M. Megharaj. 2009. Assessment of biological and
biochemical indicators in soil under transgenic Bt and non-Bt cotton crop in a sub-tropical
environment. Environ. Monit. Assess. 156:595-604. doi: 10.1007/s10661-008-0508-y.
80. Schmalenberger, A., S. Hodge, A. Bryant, M. Hawkesford, M. Singh, and M. Kertesz. 2008. The
role of Variovorax and other Comamonadaceae in sulfur transformations by microbial wheat
rhizosphere communities exposed to different fertilization regimes. Environ. Microbiol. 10:1486-
1500.
81. Schmalenberger, A., and M. Kertesz. 2007. Desulfurization of aromatic sulfonates by
rhizosphere bacteria: high diversity of the asfA gene. Environ. Microbiol. 9:535-545.
82. Schmalenberger, A., and C. C. Tebbe. 2002. Bacterial community composition in the rhizosphere
of a transgenic, herbicide-resistant maize (Zea mays) and comparison to its non-transgenic cultivar
Bosphore. FEMS Microbiol.Ecol. 40:29-37.
83. Schmalenberger, A., A. Telford, and M. A. Kertesz. 2010. Sulfate treatment affects desulfonating
bacterial community structures in Agrostis rhizospheres as revealed by functional gene analysis
based on asfA. Eur.J.Soil Biol. 46:248-254.
84. Sessitsch, A., T. Coenye, A. V. Sturz, P. Vandamme, E. A. Barka, J. F. Salles, J. D. van Elsas, D. Faure, B. Reiter, B. R. Glick, G. Wang-Pruski, and J. Nowak. 2005. Burkholderia phytofirmans sp.
nov., a novel plant-associated bacterium with plant-beneficial properties. Int. J. Syst. Evol. Microbiol.
55:1187-1192.
85. Shen, R., H. Cai, and W. Gong. 2006. Transgenic Bt cotton has no apparent effect on enzymatic
activities or functional diversity of microbial communities in rhizosphere soil. Plant and Soil.
285:149-159.
86. Singh, B., S. Munro, J. Potts, and P. Millard. 2007. Influence of grass species and soil type on
rhizosphere microbial community structure in grassland soils. Appl. Soil Ecol. 36:147-155.
87. Smalla, K., G. Wieland, A. Buchner, A. Zock, J. Parzy, and S. Kaiser. 2001. Bulk and rhizosphere
soil bacterial communities studied by denaturing gradient gel electrophoresis: plant-dependent
enrichment and seasonal shifts revealed. Appl. Environ. Microbiol. 87:4742-51.
88. Smit, E., P. Leeflang, S. Gommans, J. van den Broek, S. van Mil, and K. Wernars. 2001. Diversity
and seasonal fluctuations of the dominant members of the bacterial soil community in a wheat field
as determined by cultivation and molecular methods. Appl. Environ. Microbiol. 67:2284-2291. doi:
10.1128/aem.67.5.2284-2291.2001.
152
89. Uroz, S., M. Buée, C. Murat, P. Frey-Klett, and F. Martin. 2010. Pyrosequencing reveals a
contrasted bacterial diversity between oak rhizosphere and surrounding soil. Environmental
Microbiology Reports. 2:281-288. doi: 10.1111/j.1758-2229.2009.00117.x.
90. van Berloo, R., R. Hutten, H. van Eck, and R. Visser. 2007. An online potato pedigree database
resource. Potato Research. 50:45-57.
91. van Overbeek, L., and J. D. van Elsas. 2008. Effects of plant genotype and growth stage on the
structure of bacterial communities associated with potato (Solanum tuberosum L.). FEMS
Microbiol.Ecol. 64:283-296.
92. Vinther, F., G. Mortensen, and L. Elsgaard. 2003. Effects of linear alkylbenzene sulfonates on
functional diversity of microbial communities. Environ Tox Chem. 22:35-39.
93. Weinert, N., R. Meincke, C. Gottwald, H. Heuer, N. C. Gomes, M. Schloter, G. Berg, and K. Smalla. 2009. Rhizosphere communities of genetically modified zeaxanthin-accumulating potato
plants and their parent cultivar differ less than those of different potato cultivars. Appl. Environ.
Microbiol. 75:3859-3865. doi: 10.1128/AEM.00414-09.
94. Weinert, N., Y. Piceno, G. C. Ding, R. Meincke, H. Heuer, G. Berg, M. Schloter, G. Andersen, and K. Smalla. 2011. PhyloChip hybridization uncovered an enormous bacterial diversity in the
rhizosphere of different potato cultivars: many common and few cultivar-dependent taxa. FEMS
Microbiol. Ecol. 75:497-506. doi: 10.1111/j.1574-6941.2010.01025.x; 10.1111/j.1574-
6941.2010.01025.x.
95. Wieland, G., R. Neumann, and H. Backhaus. 2001. Variation of microbial communities in soil
rhizosphere and rhizoplane in response to crop species, soil type, and crop development. Appl.
Environ. Microbiol. 67:58349-5854.