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Deciphering the microbial
composition of biodynamic
preparations and their
effects on the apple
rhizosphere microbiome
Expedito Olimi
1
*, Samuel Bickel
1
, Wisnu Adi Wicaksono
1
,
Peter Kusstatscher
1
, Robert Matzer
4
, Tomislav Cernava
1
and Gabriele Berg
1,2,3
*
1
Institute of Environmental Biotechnology, Graz University of Technology, Graz, Austria,
2
Leibniz Institute for Agricultural Engineering and Bioeconomy (ATB), Potsdam, Germany,
3
Institute for Biochemistry and Biology, University of Potsdam, Potsdam, Germany,
4
Independent consultant for soil management (BODENmanagement.net), Graz, Austria
Soil microbial communities are crucial for plant growth and are already
depleted by anthropogenic activities. The application of microbial transplants
provides a strategy to restore beneficial soil traits, but less is known about the
microbiota of traditional inoculants used in biodynamic agriculture. In this
study, we used amplicon sequencing and quantitative PCR to decipher
microbial communities of composts, biodynamic manures, and plant
preparations from Austria and France. In addition, we investigated the effect
of extracts derived from biodynamic manure and compost on the rhizosphere
microbiome of apple trees. Microbiota abundance, composition, and diversity
of biodynamic manures, plant preparations, and composts were distinct.
Microbial abundances ranged between 10
10
-10
11
(bacterial 16S rRNA genes)
and 10
9
-10
11
(fungal ITS genes). The bacterial diversity was significantly higher
in biodynamic manures compared to compost without discernible differences
in abundance. Fungal diversity was not significantly different while abundance
was increased in biodynamic manures. The microbial communities of
biodynamic manures and plant preparations were specificforeach
production site, but all contain potentially plant-beneficial bacterial genera.
When applied in apple orchards, biodynamic preparations (extracts) had the
non-significant effect of reducing bacterial and fungal abundance in apple
rhizosphere (4 months post-application), while increasing fungal and lowering
bacterial Shannon diversity. One to four months after inoculation, individual
Frontiers in Soil Science frontiersin.org01
OPEN ACCESS
EDITED BY
David Johnston-Monje,
University of Valle, Colombia
REVIEWED BY
Oluwadara Pelumi Omotayo,
North-West University, South Africa
Marketa Sagova-Mareckova,
Crop Research Institute (CRI), Czechia
*CORRESPONDENCE
Expedito Olimi
expedito.olimi@tugraz.at
SPECIALTY SECTION
This article was submitted to
Soil Biology, Ecosystems and
Biodiversity,
a section of the journal
Frontiers in Soil Science
RECEIVED 16 August 2022
ACCEPTED 31 October 2022
PUBLISHED 16 November 2022
CITATION
Olimi E, Bickel S, Wicaksono WA,
Kusstatscher P, Matzer R, Cernava T
and Berg G (2022) Deciphering the
microbial composition of biodynamic
preparations and their effects on the
apple rhizosphere microbiome.
Front. Soil Sci. 2:1020869.
doi: 10.3389/fsoil.2022.1020869
COPYRIGHT
© 2022 Olimi, Bickel, Wicaksono,
Kusstatscher, Matzer, Cernava and Berg.
This is an open-access article
distributed under the terms of the
Creative Commons Attribution License
(CC BY). The use, distribution or
reproduction in other forums is
permitted, provided the original
author(s) and the copyright owner(s)
are credited and that the original
publication in this journal is cited, in
accordance with accepted academic
practice. No use, distribution or
reproduction is permitted which does
not comply with these terms.
TYPE Original Research
PUBLISHED 16 November 2022
DOI 10.3389/fsoil.2022.1020869
taxa indicated differential abundance. We observed the reduction of the
pathogenic fungus Alternaria, and the enrichment of potentially beneficial
bacterial genera such as Pseudomonas. Our study paves way for the
science-based adaptation of empirically developed biodynamic formulations
under different farming practices to restore the vitality of agricultural soils.
KEYWORDS
biodynamic farming, compost microbiome, biodynamic manures, biodynamic
preparations, rhizosphere microbiome, 16S rRNA/ITS amplicon sequencing, organic
soil amendments
Introduction
The soil microbiome plays an essential role in crop
production, but is a strongly influenced component in the
Anthropocene (1,2). Therefore, it is critical to identify threats
that cause microbiome disturbances, as well as to find
sustainable restoration strategies for affected microbiomes (2).
Microbial diversity is crucial for soil and plant health (3) and for
human health as well, which can be measured in taxonomic and
functional diversity as well as by total gene count within the
microbiota (4). However, enriching microbial diversity in the
soil is a challenge, which can be managed directly by applying (i)
microbiome transplants, (ii) microbes with beneficial properties,
(iii) microbiota-active metabolites, or indirectly by changing
environmental conditions in a way that microbiomes also shift
their structure and function from dysbiosis into a healthy state
(1,5). The use of compost in field fertilization, a classical
microbiome transplant, can be traced back to the advent of
human civilization, marked by the agricultural revolution (6).
Numerous benefits for enhancing soil ecosystem functions, soil
quality, and plant health, are attributed to composting in arable
and natural ecosystems (7–9). However, many extensive farming
systems from the past have been replaced by what is nowadays
known as intense, conventional agriculture. It is characterized by
high-yield cultivars, which need high inputs of synthetic
fertilizer, and pesticides and commonly result in reduced soil
vitality, emerging and multi-resistant soil-borne pathogens and
an overall yield decline, which prompts a re-evaluation of
agricultural practices (10–12).
An ongoing evaluation of current agricultural practices has
shifted attention towards alternative and environmentally
sustainable systems, including biodynamic farming to restore
the “lost soil microbiome”(13–15). Furthermore, the globally
increasing demand for foods and beverages produced under
organic management practices (16–18) motivates a deeper
investigation of microbial constituents that are present in the
utilized organic amendments. Biodynamic (BD) farming was
first conceptualized by the Austrian philosopher Rudolf Steiner
(1861-1925) (19) and is considered the first systematic form of
ecological or organic farming (15,19,20). Although organic and
BD farming are related, the latter considers specific practices
which are intended to influence the biological metaphysical
aspects including natural rhythms concerning the sun and
moon, weather, and seasons of the farm (10,15,21). A special
feature of BD agriculture is the use of specific BD manure and
fermented plant preparations (20). These products are applied as
solid manure or liquid extracts in form of field sprays (i.e., soil or
foliar application) to enhance soil quality and stimulate plant
health (22–24). Recent studies highlighted the role of BD
preparations, especially in enhancing soil diversity (15,18,25),
and have also found application in viticulture (17), especially in
enhancing winegrape quality (26). In addition, long-term field
trials indicated differences in the soil microbiome between BD,
integrated, and organic farming systems (27,28) even under
desert farming conditions (29).
BD farming is rooted in anthroposophy, which views
humans as the primary link between the cycles of the earth
and the cosmos, with humans bridging a gap between the
spiritual and material worlds (17). Special natural biodynamic
preparations (dubbed “Steiner preparations”)wereformulated
and utilized in this system to replace synthetic products (17).
The procedure to obtain BD formulations involves fermenting
cow manure in cow horns to create the so called biodynamic
extracts (compost teas), which are then applied according to
Steiner (30). While the reason for fermentation in cow horns
may still be unknown, preparations from this method have
been proven to have special qualities including soil fertility
improvement and enhancing plant physiological responses to
light radiation (25). Moreover, (26) found a substantial impact
of BD preparations in the enhancement of wine quality,
winegrape canopy and chemistry, despite contradicting
results regarding effects on soil quality. Yet, the microbiome
of BD formulations (manures and plant preparations), the
differences between BD manures and non-biodynamic
composts as well as their impact on the rhizosphere
microbiota remain unexplored.
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Therefore, we analysed bacterial and fungal communities of
BD formulations (manures and plant preparations) and
composts (standard-, horse-, and apple composts). Extracts
from the BD manures and composts are commonly applied as
field sprays; thus, we also investigated the microbiome of liquid
extracts and their precursor materials. The two analysed extracts
were applied in differently managed (organic and integrated)
apple orchards and samples were taken at bloom (one month),
and post-harvest (four months) to investigate the temporal
variability in the rhizosphere microbiome. The study was
based on the hypotheses that: (i) BD manures and composts
have a different microbiome; (ii) the microbiome of BD manures
is affected by the amendment with biodynamic plant
preparations; (iii) the microbiome of BD manures differs
between regions and the year of production; (iv) the
microbiomes of extracts obtained from BD manure and
compost are different from the precursor materials, and (v) the
microbiome of the apple rhizosphere is affected by the
application of extracts.
Material and methods
Description of the biodynamic and
compost formulations
We sampled four BD manure products and six biodynamic
plant preparations for amplicon sequence analysis. BD horn
manures were obtained from Demeter (Vienna, Austria) and
BioDynamie Services sarl (Chateau, France), while all plant
preparations were obtained from France. In the BD farming
community, the plant preparations are referred to as P-502, P-
503, P-504, P-505, P-506, and P-507. In the production process,
BD manure (500) is prepared from cow manure that is
fermented in cow horns and buried in the soil at a depth of
50 cm for six months (i.e., during autumn and winter). After the
horns are recovered from the soil, the retrieved horn manure
(500) can be amended with six different plant preparations (P-
502 to P-507) and referred to as 500P.
The BD formulations used in this study originated from two
different regions of production: France (FR-500/FR-500P) and
Austria (AT-500/AT-500P). Apart from differences in the region
of origin, we included BD products from different production
years (Table 1). According to the producers, BD preparations
were stored in ceramic pots and placed in a double-walled
wooden box covered with a wooden lid. The boxes were kept
in cellars exclusively dedicated for storage of preparations at
constant environmental conditions. We analysed BD manure
samples without preparations (AT/FR-500) produced in the
years 2019, 2020, and 2021 and BD manures with preparations
(AT/FR-500P) from the years 2012, 2016, and 2020 (Table 1).
The BD manure FR-500P produced in 2016 was amended with
four plant preparations coded F, T, S, and V, according to the
producer in France.
For comparison between biodynamic manures and compost,
three composts (standard-, apple, and horse compost) were
included in the analysis (Table 1). Briefly, the different
composts were constituted as follows: (i) Standard compost
(70% shrub cuttings, 5% soil, and 25% organic waste); (ii)
Horse compost (60% shrub cuttings, 5% soil, and 35% horse
manure with straw); and (iii) Apple compost (75% shrub
cuttings, 5% soil, and 20% pressed organic apple fruits).
Field treatments and experimental design
As a common practice in BD farming, sprays for field
application are extracted from BD formulations. Similarly,
compost can be applied in liquid form as “compost tea”;itis
prepared by mixing liquid extracts from compost with molasses
and stone dust. We applied two extracts from the BD manure
(AT-500P) and compost (standard compost) in two apple
orchards, which were maintained following organic and
integrated management practices. The organic and integrated
orchards were located at 47.1289°N, 15.7807°E and 47.2125°N,
15.8569°E, respectively. Apple trees were grown for
approximately ten years in each orchard. Treatments were
performed at the beginning of spring (May 2021). The BD
preparations were applied by sprinkling the extracts in the
vicinity of the plant roots, using the “Demeter”recommended
application rate of 1:10 (i.e. liquid extract: water) liters per acre.
For each orchard, a randomized plot design of 4 plots per
treatment and 8-10 trees per plot was chosen. To account for
spatial variations, plots were randomly distributed across six
neighbouring rows. Rhizosphere samples were randomly taken
from each plot using an auger of 5 cm diameter. Ten soil cores
from each plot were pooled to comprise a biological replicate.
Rhizosphere (soil associated with roots) sampling was
performed one- and four- months after treatment and
corresponded with the spring and autumn seasons. Samples
were kept cooled and transported within six hours to the
laboratory at the Institute of Environmental Biotechnology
(Graz University of Technology, Graz, Austria).
Sample processing and deoxyribonucleic
acid extraction
For compost, BD manures, and BD plant preparations, four
grams of sample were stored at -70°C until DNA was extracted.
Before DNA extraction from liquid extracts (“compost tea”), the
extracts from compost and BD manure were centrifuged at
16000g for 16 minutes, and the microbial pellet was frozen at
-70°C until DNA extraction. For orchard samples, approx. four
Olimi et al. 10.3389/fsoil.2022.1020869
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grams of rhizosphere soil were stored at -70°C. For all samples,
total microbial genomic DNA was extracted using the E.Z.N.A.®
Soil DNA Kit (Omega Bio-tek, Inc.; Norcross-Georgia, United
States) following the manufacturer’s instructions. DNA was
quality checked using a Nanodrop 2000 (Thermo Scientific,
Wilmington, DE, USA) and stored at -20°C before performing
PCR and amplicon sequencing.
Real-time quantitative PCR of total
bacterial and fungal abundance
The primer pairs Unibac-II-515f/Unibac-II-806r for bacteria
as previously used by (29) and ITS1f/ITS2r for fungi (31) were
used for real-time qPCR quantifications to determine the copy
numbers of 16S rRNA and ITS genes in the different samples.
Reactions were performed in a total volume of 10 mLina
reaction mix composed of 5 mL of KAPA SYBR Green (Bio-Rad,
Hercules, CA, U.S.A.), 0.5 mL of each primer (10 µM), 3 mLof
PCR grade water and 1 mL template DNA (samples were diluted
1:10 in PCR grade water). Amplifications were performed in
triplicates for each sample using a Rotor-GeneTM 6000 series
thermal cycler (Corbett Research, Sydney, Australia) with the
following program settings: initial denaturation (95°C,5 min)
followed by 35 cycles of denaturation (95°C,10 s); annealing
(54°C, 15 s); extension (72°C, 10 s); then melt down from 72 to
96°C. Serial dilutions of standards containing defined copy
numbers were generated according to (29)andusedto
calculate gene copy numbers in different samples. As standards,
known concentrations of 16S rRNA gene fragments from a Bacillus
sp., and the ITS region from a Penicillium sp. were used.
Amplicon library preparation
The extracted DNA was further used for amplicon library
preparation based on the hypervariable V4 region of the
bacterial 16S rRNA gene and the ITS1 regions of fungal DNA.
TABLE 1 Overview of the studied composts (i.e., apple-, horse-, and standard compost) and biodynamic manures (with/without plant
preparations).
Sample ID Formulation* Year Sample type Region
#
Description
C1 Compost tea
A
2021 Extract Field spray extract (from C12)
C2 Compost tea 2021 Extract Extract from C12, but same as C1
C3 Compost tea 2021 Extract Extract from C13
C4 FR-500P 2012 Solid FR 500P- Horn manure preparation -2012
C5 FR-500 2019 Solid FR 500-Horn manure-2019
C6 FR-500P 2016 Solid FR 500P- Horn manure preparation-2016(F)
C7 FR-500P 2016 Solid FR 500P- Horn manure preparation-2016(T)
C8 FR-500P 2016 Solid FR 500P- Horn manure preparation-2016(S)
C9 FR-500P 2016 Solid FR 500P- Horn manure preparation-2016(V)
C10 FR-500P 2020 Solid FR 500- Horn manure-2020
C11 FR-500P 2020 Solid FR 500P- Horn manure preparation-2020
C12 Standard compost Solid AT
C13 Apple compost Solid AT
C14 Horse compost Solid AT
D1 AT-500 2020 Solid AT 500- Horn manure
D2 AT-500 2021 Solid AT 500- Horn manure
D3 AT-500P 2020 Solid AT 500P- Horn manure Preparation
D4 AT-500P
A
2021 Extract AT Field spray extract (from D3)
D5 AT-500P 2021 Extract AT Field spray extract (from D3)
P1 P-502 Solid FR Achillea millefolium (Achillee; P-502)
P2 P-503 Solid FR Matricaria rectutita (Camomille; P-503)
P3 P-504 Solid FR Urtica dioica (Ortie (P-504)
P4 P-505 Solid FR Quercus robur (Ecorce DeChene; P-505)
P5 P-506 Solid FR Taraxacum dens-leonis (Pissenlit; P-506)
P6 P-507 Liquid FR Valeriana officinalis (Valeriane; P-507)
#
Acronyms FR, France, AT, Austria indicate regions of manure and preparation production.
*The superscript A indicates extracts which were used in field treatments.
BD plant preparations (P502-507) and their respective plant species from which they are derived were also included.
Olimi et al. 10.3389/fsoil.2022.1020869
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We employed one-step PCR using the primer pair 515F (5’-
GTGCCAGCMGCCGCGGTAA-3’), and 806R (5’-
GGACTACHVGGGTWTCTAAT-3’) for the bacterial library
preparation (32). Both forward and reverse primers contained
sample-specific barcodes to facilitate multiplexed sequencing.
For each PCR, 1 µL of extracted DNA was used as a template in a
30 µL reaction. Peptide nucleic acid (PNA) PCR clamps were
used to block the amplification of plant plastid and
mitochondrial 16S rRNA genes during the PCR amplification
of the bacterial community (33,34). The reaction mixture
contained 6 µL (5xTaq &GO, PCR pre-mix, MP Biomedicals),
0.6 µL (10 µM 515F/806R) primers, 0.45 µL (50 µM mPNA and
pPNA), and 20.9 µL of PCR grade water. All reactions were
performed in triplicates in a thermocycler (Bio-metra GmbH,
Jena, Germany). The PCR program included an initial
denaturation (96°C, 5 min), followed by 30 cycles (94°C for 60
s, 78°C PNA step for 5 s, 54°C for 1 min, 74°C for 60 s), followed
by 74°C for 10 min and then a cool down to 10°C.
For the fungal community, we used the primer pair ITS1f
(5’-CTTGGTCATTTAGAGGAAGTAA-3’)andITS2r(5’-
GCTGCGTTCTTCATCGATGC-3’)(31,35) for library
preparation. All amplifications were performed in triplicates.
One µL of extracted DNA was used in each 30 µL reaction. The
reaction mixture contained 6 µL (5xTaq &GO, PCR pre-mix,
MP Biomedicals), 0.6 µL (10 µM ITS1F/ITS2R) primers, 21.8 µL
of PCR grade water. The PCR program included the first step of
initial denaturation (96°C, 5 min), followed by 30 cycles
(denaturation: 96°C for 60 s, annealing: 58°C for 60 s, and
extension: 74°C for 60 s), followed by 74°C for 10 min (final
extension) and a cool-down step to 10°C. Successful PCR
amplifications at the correct amplicon size were confirmed by
gel electrophoresis. PCR amplicons were purified using Wizard
SV Gel and PCR Clean-Up System (Promega, Madison, WI)
following the manufacturer’s instructions. Purified PCR
amplicons were quantified using a Nanodrop 2000 (Thermo
Scientific, Wilmington, DE, USA) and pooled in equimolar
concentrations. Paired-end Illumina MiSeq 2x 250 sequencing
of the amplicon library was performed at Eurofin(Berlin,
Germany). All raw reads obtained from the sequencing
company were deposited at the European Nucleotide Archive
(ENA) under study accession number PRJEB53400.
Bioinformatic pipeline
Paired-end reads from the sequencing facility were quality-
checked and demultiplexed using Cutadapt (36). Demultiplexed
reads were analysed using QIIME2 version 2021.11.0 (37).
Primer sequences were removed, and the DADA2 algorithm in
theQIIME2environmentwasemployedtoqualityfilter,
denoise, and remove chimeric sequences (38); thus generating
amplicon sequence variants (ASVs), and a feature table of
microbial counts. Taxonomic assignment of denoised reads
was performed using the VSEARCH (39) tool in QIIME2 by
comparing the reads against reference databases: SILVA132 for
16S rRNA (40,41) and UNITE v7 (42) for ITS sequence reads.
Statistical analysis
All data were analysed with R statistical software (version
4.0.3) using RStudio (Version 1.1.423) (43), supplemented by the
web-based microbiome analyst tools (44). The obtained
microbial ASV tables and taxonomic assignment were
processed using phyloseq (45) and vegan 2.5.7 (46) packages.
For alpha and beta diversity analyses, the statistics were
performed on datasets rarefied to minimum sampling depths of
1000 and 700 reads per sample for 16S and ITS analyses,
respectively. The sample-specific rarefaction curves for
bacterial and fungal communities were visualized (Figure S1).
One sample of a biodynamic plant preparation (P-507) was
excluded from the analysis due to insufficient replicates (one
sample was retained).
Microbial alpha diversity was calculated using the Shannon
diversity index (H’). The non-parametric Kruskal-Wallis test
(47), followed by Dunn’s test (corrected for multiple
comparisons using the Bonferroni method) was performed
on Shannon diversity index and abundance data to test for
differences between the samples (e.g., BD manures vs.
composts, extracts vs. precursor materials); and treatment
comparisons (extract from standard compost and AT-500P)
post-inoculation in the different orchards. Also, the effect of
treatment on the rhizosphere microbiome at different
sampling times was tested. Permutational analysis of
variance (PERMANOVA, 999 permutations) based on Bray-
Curtis dissimilarity was used to test for differences in
community structure between sample groups. For significant
factors (and combinations), pairwise comparisons were
performed using the ‘pairwise.adonis2’function in vegan
(46) with p-values adjusted using the Bonferroni method.
The differences in microbial community structure between
the sample groups were visualized using dendrograms
(generated by complete clustering of Bray-Curtis distances),
and non-metric multidimensional scaling (NMDS).
Microbial taxonomic composition based on the percentage
relative abundance of top20 taxa at class, order, and family levels
was represented using stacked bar plots. Linear discriminant
analysis Effect Size (LEfSE) (48) implemented in microbiome
analyst (44) was used to identify the taxa underlying the observed
microbiome differences between the composts and biodynamic
preparations, as well as revealing treatment effects in the different
orchards and sampling times. The common microbiome (defined
by a prevalence of 50%, and detection threshold of 0.001% relative
abundance) was used to obtain unique and shared ASVs associated
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with the different samples. The list of ASVs was visualized using a
Venn diagram generated by the systemPipeR package (49)in
Rstudio.
We used the SourceTracker R script (50)toestimatethe
proportion of microbiota in the apple rhizosphere that potentially
originated from the field sprays extracted from compost and BD
manures after soil treatment. SourceTracker estimates the
proportion of microbiota that originates from a set of source and
sink environments as previously described (50). In our case, the
extracts from BD manure (AT-500P) and standard compost which
were used for orchard treatment were considered as the source
environment and the rhizosphere soil as the sink environment. By
considering control soil as the microbial source, we also tracked
microbiota in the treated rhizosphere soil which was not of extract
origin, but likely originated from the soil.
Results
The biodynamic manures and composts
were distinct in microbiome structure
and composition
Amplicon sequencing of bacterial and fungal communities
yielded 2,213,856 and 4,534,963 high-quality reads, assigned to
9,387 bacterial and 2,742 fungal ASVs, respectively. Generally, we
observed significant differences in microbiome structures between
BD manures (AT-500/P or FR-500/P) and composts (i.e. apple-,
horse-, standard composts), especially for the bacterial community
(Figure 1). Moreover, a separation based on the country of origin of
BD manures was observed (Figures 1A,B). Significant differences
(P=0.008) in Shannon diversity between compost and BD manures
were observed for bacterial communities (Table S3A), while no
significant differences (P=0.19) were observed for the fungal
community (Table S4A). Microbial abundance was significantly
different for bacterial (P=0.04; Table S1A) and fungal communities
(P=0.009; Table S2A).
The bacterial abundance (16S rRNA gene fragments per
gram soil) was higher in BD manure than in compost and ranged
as follows and ranged as follows: (BD manure: 2.52×10
11
to
4.17×10
11
vs compost: 8.89.16×10
10
to 1.46×10
11
). Meanwhile,
the fungal community (ITS region copies per gram soil) were:
(BD manure: 1.34×10
9
to 1.18×10
11
vs compost: 8.89×10
10
to
1.46×10
11
). However, a significant difference (P<0.05) was
observed only for the fungal abundance between BD manure
(AT-500) and horse compost (Figure S2B), while no significant
differences (P>0.05) were observed for the bacterial community
(Figure S2A).
The bacterial Shannon index was higher in BD manures than
in composts and ranged as follows: BD manure (5.28-5.63)
B
CD
EF
A
FIGURE 1
Microbial community structure and composition of BD manures and composts. Coloured circles represent the different BD manures (500/P)
and non-biodynamic (NBD) composts. Figures (A, B) are dendrogram representation of the bacterial and fungal community obtained by
complete clustering of the Bray Curtis distance matrix. (C, D) are stacked bar plots showing the bacterial, and fungal taxonomic composition in
the different BD and compost samples; while (E, F) represent shared and unique ASVs between BD manures and composts for bacterial and
fungal communities, respectively. The common microbiome was defined as ASVs with prevalence >50% and relative abundance >0.001%. AT
and FR indicate countries Austria and France, respectively.
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versus compost (1.81-3.54). Significantly higher bacterial
diversity (P=0.02) in BD manures (AT-500 and FR-500)
compared with horse compost was observed (Figure S2C). In
contrast, the fungal Shannon diversity ranged between (2.63-
3.13) for BD manures as compared to compost (2.41-2.60), and
significant differences (P>0.05) in Shannon diversity were
observed for the fungal community (Figure S2D).
PERMANOVA between BD manures and composts
revealed significant differences in community compositions
between these organic formulations for bacteria (R
2
= 0.85,
P=0.001) and fungi (R
2
= 0.85, P=0.001). In addition, BD
manures amended with plant preparations (AT/FR-500P)
could be distinguished from those without preparations (AT/
FR-500), especially for the bacterial community (Figures 1A,B).
Comparing higher-level taxonomy of BD manures and
composts revealed differences in microbial composition between
samples. Dominant phyla included Chloroflexi (average percentage
abundance: 35.0%), Proteobacteria (29.5%), and Firmicutes (8.9%).
The phylum Chloroflexi was dominant in composts (standard-,
apple-, and horse), with abundance (69.8%, 72.4%, and 87.7%,
respectively). Meanwhile, Proteobacteria,Firmicutes,and
Actinobacteria were highest in BD manures with relative
abundances of 43.7%, 14.8%, and 9.5%, respectively. For the
fungal community, the phyla Ascomycota (59.7%), Basidiomycota
(5.7%), and Mortierellomycota (5.1%) were predominant. We
observed differences among compost microbiomes, especially for
the bacterial community. The bacterial family Anaerolineaceae was
present only in composts; however, with a dominance of uncultured
bacteria (Figure 1C). Meanwhile, BD manures were seen to contain
members of families Pseudomonadaceae,Peptostreptococcaceae,
Microbacteriaceae,Lentimicrobiaceae,Lachnospiraceae,
Devosiaceae,andBurkholderiaceae (Figure 1C). The fungal
families including unidentified Microascales, unidentified
Hypocreales,andCladosporiaceae were observed in composts
(Figure 1D). Trichosporonaceae,Microascaceae,Lasiosphaeriaceae,
Mortierellaceae,andChaetomiaceae were observed in BD manures,
while the fungal family Cordycipitacea was only found in the apple
compost (Figure 1D).
Moreover, bacterial genera including Bacillus,Devosia, and
Pseudomonas were mainly associated with BD manures (AT-
500/P and FR-500/P). The fungal genera Lecanicillium, and
Arthrobotrys were mainly present in composts, while
Trichosporon, Scutellinia, and Sporobolomyces, were associated
with BD manures.
Analysis of the common microbiome analysis revealed that
109, and 82 bacterial ASVs were unique to BD manures and
composts, respectively, while only two ASVs were shared
(Figure 1E). Some bacterial genera in the common
microbiome of BD manure were Paenibacillus,Mesorhizobium,
Mycobacterium,Cellulomonas, and Clostridium. On the other
hand, 13 and 54 unique fungal ASVs were detected in BD
manures and compost, while five ASVs were shared between
both microbiomes (Figure 1F). The shared fungal ASVs included
unidentified Capnodiales and unidentified Microascacea.
Biodynamic manures varied in structure
and composition between the different
countries of production, but shared a
common microbiome
We compared BD manures that differed in production year
and region of production (Austria or France). Differences
(P<0.05) in fungal diversity were observed between FR-500P
[2012] and AT-500 [2021] as shown (Figure S3D), while the
fungal abundance was substantially higher in AT-500 [2020/
2021] as compared to FR-500P [2016: S, T, and V] (Figure S3B).
No significant differences (P>0.05) in bacterial abundance and
diversity were observed among the biodynamic manures, for the
two regions and different years of production (Figures S3A,C).
In contrast to microbial diversity and abundance, we observed
substantial differences in microbial composition between BD
manures based on the country of production as follows: bacterial
community (BD manure: R
2
= 0.78, P=0.001, country: R
2
=0.24,
P=0.001), and fungal community (BD manure: R
2
= 0.78, P=0.001,
country: R
2
= 0.26, P=0.001). These differences were also shown by
NMDS in which clear separation of samples according to the region
and BD manure type was observed (Figure S4).
The dominant bacterial classes were Gammaproteobacteria
(19.3 average percentage relative abundance), Clostridia (14.5%)
and Alphaproteobacteria (12.1%). Predominant fungal classes
included Sordariomycetes (47.9%), Tremellomycetes (6.7%),
Pezizomycetes (5.9%) and Dothideomycetes (5.2%). Differences
between regions were visible, especially for the bacterial
community. For instance, the order Actinomarinales was only
found in BD manures produced in France (Figure S3C).
Meanwhile, Sphingobacteriales were observed exclusively in
Austrian BD manure samples (Figure S4C). The fungal
community between the two regions was separated by the
presence of orders Capnodiales,Ascosphaerales,and
Pleosporales were associated with the BD manures produced in
France. Independent of the region of production, BD manures
were found to contain bacterial families such as Bacillaceae,
Burkholderiaceae,Clostridiaceae,Flavobacteriaceae,
and Pseudomonadaceae.
The assessment of the common microbiome of BD manures
from the two countries revealed that 30 bacterial and 13 fungal
ASVs were shared. Most bacterial ASVs were unique (120 and
123 for Austria and France, respectively). In contrast, a total of
16 and 11 fungal ASVs were unique for Austria and France
(Figures S4E,F). Some of the shared bacterial ASVs included
Devosia, Azotobacter, Acidibacter, Clostridium, Romboutsia,
Cellulomonas, Sorangium, Solirubrobacter,andTuricibacter.
The shared fungal genera were Trichosporon,Mortierella,
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unidentified Microascaceae, unidentified Lasiosphaeriaceae and
unidentified Sordariales.
Common microbiome of plant
preparations and preparation-amended
BD manures
To establish whether BD plant preparations altered the
microbiome of BD manures (FR/AT-500P), the community
structure and diversity of BD preparations (manure and plant
preparations) were analysed. Significant differences in
microbiome composition were seen between amended BD
manures and the different plant preparations (bacterial: R
2
=
0.83, P=0.001; fungal community: R
2
= 0.94, P=0.001. NMDS
visualization showed a clear separation between BD manures
and plant preparations. Microbiome separation between plant
preparations was observed for bacterial and fungal communities,
respectively (Figures S5A,C).
There was a high microbiota load both in BD plant
preparations and manures (bacterial: 8.12 × 10
10
to 2.65 ×
10
11
), and fungal (9.70 × 10
9
to 1.34 × 10
10
). A significant
difference (P<0.05) was observed for the fungal abundance
between FR-500 and P-505 (Figure S6B), and no differences
(P>0.05) were observed for the bacterial community (Figure
S6A). On the other hand, the Shannon diversity index of BD
plant preparations and amended manure (i.e. FR-500P) ranged
between 4.38-5.28 and 2.20-3.18, both for bacterial and fungal
communities, respectively. Significant differences (Kruskal-
Wallis, P<0.05) in bacterial diversity were observed between
BD manure (FR-500P) and plant preparation (P-505), as well as
between P-504 with P-505 and P-506 (Figure S5C). The fungal
diversity was substantially higher (P<0.05) in preparations (P-
505 and P-506) as compared to P-503 (Figure S6D).
The bacterial order Bacillales was present both in BD
manures and plant preparations, except in preparation P-505,
and was highest in P-503 and P-504 with average relative
abundances of 11.8% and 36.0%, respectively. The bacterial
orders Rhizobiales,Pseudomonadales,andFlavobacteriales
were observed in both plant preparations and BD manures
(Figure 5B). Meanwhile, Bacteroidales was especially present in
AT-500/P and plant preparations (P-503 and P-505). For the
fungal community, BD manures were distinguished from plant
preparations by orders Capnodiales,Hypocreales,
Ascosphaerales,Eurotiales and Unidentified fungi. The fungal
orders Microascales,andPezizales were associated with BD
manures and plant preparations (Figure S4D).
A comparison between preparation amended and non-
amended manures revealed that 120 and 155 bacterial ASVs
were shared between (AT-500 and AT-500P), as well as (FR-500
and FR-500P), respectively (Figures S7B,D). Similarly, for the
fungal community 25 and 23 ASVs were shared (Figures S8B,
D). There was a common microbiome shared between BD
manures and plant preparations, both for bacterial (Figures
S7A,C) and fungal communities (Figures S8A,C),
respectively.Moreuniquethansharedmembersofthe
bacterial (Figures S7A,C) and fungal (Figures S8A,C) ASVs
were observed for plant preparations and BD manures. The
shared part of the common microbiome between BD manures
and plant preparations was composed of bacterial genera
Romboutsia,Azotobacter,Pedomicrobium,andBacillus.
Meanwhile, shared fungal genera were Fusarium,
Arthrographis, Unidentified Microascaceae, Unidentified
Chaetomiaceae, and Unidentified Nectriaceae.
LEfSe analysis identified the bacterial and fungal genera
explaining the differences between BD manures (FR-500P) and
plant preparations (Figures 2A,B, respectively). The BD plant
preparations (P-502, P-503, P-504, P-505, and P-506) were
differentiated by the bacterial genera Luteibacter,
Oceanobacillus,Virgibacillus,Mycobacterium,and
Staphylococcus (Figure 2A). Lentimicrobium,Romboutsia, and
Devosia were associated with BD manures (AT/FR-500P).
However, the genus Bacillus was shared between BD manures
(AT/FR-500P) and plant preparations. Meanwhile, fungal
genera Scutellinia,Trichosporon,andSporobolomyces were
mainly associated with BD manures, while Fusarium,
Ascosphaera,Pseudallescheria,Arthrographis,Chaetomium,
Cyberlindnera,andScedosporium were associated with BD
plant preparations (Figure 2B).
Extracts and precursor materials of BD
manure and standard compost contained
a common microbiome
The microbial diversity and abundance comparison between
extracts and precursor materials of BD manure (AT-500P) and
standard compost revealed that extracts generally contained a
higher microbial diversity than precursor materials, except for
AT-500P (Figures S9C,D); and vice versa for microbial
abundance (Figures S9A,B). Significantly higher (t-test: P<0.05)
microbial abundance was observed in the precursor of BD manure
(AT-500P) in comparison to the respective extract, both for
bacterial and fungal communities (Figures S9A,B). No significant
differences (P>0.05) in microbial abundance were observed between
the precursor and extract of standard compost (Figures S9A,B).
The bacterial diversity in the standard compost extract was
significantly lower (t-test, P=0.004) than in the extract derived
from the compost materials (Figure S9C). Contrarily, the fungal
diversity in extract derived from standard compost and BD manure
(AT-500P) was significantly higher (t-test, P<0.05) in comparison
to precursor material (Figure S9D).
The microbial community structure between extract and
precursor materials of BD manure (AT-500P) and standard
compost was distinct. This was confirmed by PERMANOVA
analysis in which we observed significant differences between the
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extracts and their derivative materials (i.e., bacteria: R
2
=0.65,
P=0.001 and fungi: R
2
= 0.72, P=0.001). In addition, pairwise
community comparison revealed significant differences (P<0.05)
in community structure between precursor materials (standard
compost and AT-500P) and their respective extracts, both for
bacterial and fungal communities (Tables S7 and S8, respectively).
The extracts and precursor materials of standard compost were
distinct in microbial composition. BD manure and their respective
extract displayed similar microbiome compositions (Figures 3A,B).
The bacterial classes Anaerolineae (61.6%), and Chloroflexia (8.2%)
predominated in standard compost, while Gammaproteobacteria
(52.3%) and Bacilli (34.0%) were mainly present in the extract of
standard compost (Figure 3A).TheBDmanure(AT-500P)and
extract were mainly associated with the bacterial classes
Gammaproteobacteria (precursor: 30.8% and extract: 20.6%),
Bacteroidia (18.0% and 11.3%), Alphaproteobacteria (11.8% and
B
A
FIGURE 2
Dot plot representation of differentially abundant bacterial (A) and fungal (B) genera between BD manures and BD plant preparations. Plant
preparations are shown as P-502, P-503, P-504, P-505, and P-506; while biodynamic manures were represented as AT -500/P and FR-500/P.
Acronym P- represent preparation, while AT and FR represent the countries Austria and France where the BD manures were produced. The
differentially abundant genera with LDA scores between 5 and 7 and were based on FDR adjusted P-value. The coloured bar represents the
relative abundances of the differentially abundant genera, with the lowest indicated as blue and highest as red.
Olimi et al. 10.3389/fsoil.2022.1020869
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18.1%), Clostridia (11.8% and 11.5%)), and Deltaproteobacteria
(5.2% and 8.2%). For the fungal community, extracts of standard
compost and BD manure as compared to the precursor materials
contained a higher abundance of Dothideomycetes (10.0% versus
1.2%, and 2.6% versus 0.1%, respectively), Eurotiomycetes (7.7%
versus 1.2%, and 4.1% versus 0.3%), as well as Leotiomycetes (2.0%
versus 0.7%, and 3.1% versus 0.5%) (Figure 3B).
Interestingly, BD manure (AT-500P) and the associated extract
shared 228 and 110 ASVs of the common microbiome, both for
bacterial and fungal communities, respectively (Figures 3C,E).
Conversely, standard compost and extract were found to share
196 and 111 ASVs (Figures 3D,F). Differential abundance analysis
revealed the enrichment of bacterial genera like Bacillus,
Pseudomonas, Paenibacillus,Flavobacterium,andAzotobacter in
the extract of standard compost as compared to precursor materials
(Figure S10A). Meanwhile, in extract (AT-500P) as compared to
precursor material, an enrichment of bacterial genera including
Roseobacter and Polynucleobacter was observed, while
Lentimicrobium, Devosia,andSimplicispira were enriched in AT-
500P (precursors) Figure S10B.
For the fungal community, genus-level differences between
extracts and precursor materials of standard compost and BD
manure (AT-500P) were marked by the enrichment of fungal
genera Fusarium,Trichosporon and Aspergillus in the extract as
compared to standard compost and AT-500P precursors
(Figures S10C,D). In addition, Penicillium,andScutellinia
comprised the extracts of standard compost (Figure S10C),
while Arthrographis was enriched in AT-500P extract
(Figure S10D).
Extracts have a time-dependent effect
on the apple rhizosphere microbiome
under organic and integrated
management systems
The microbial structure and composition, as well as diversity
and abundance analyses, were performed with rhizosphere
samples from two apple orchards managed using organic and
integrated management systems. Samples were obtained at two
B
C
D
E
F
A
FIGURE 3
Comparison of microbial structure composition between AT-500P and standard compost as well as their respective extract. Panels (A, B) are
stacked barplot representations of the bacterial and fungal community composition in precursors and extracts of compost and BD manure.
Panels (C, D) are Venn diagrams representing the common bacterial ASVs shared between extract and precursor materials of compost and BD
manure, while (E, F) represent the same for the fungal community. The acronyms [E], [C], and [M] represent extract, compost, and BD manure,
respectively.
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sampling times (spring and autumn) from both orchards. There
were no significant differences (P>0.05) in microbial abundance
between treatments and controls, both for organic and
integrated orchards at one- and four-months post-inoculation
(Figure S11). Generally, after four months, bacterial diversity
was lower in treatments in comparison to controls. In contrast,
the fungal diversity was higher in treatments as compared to
controls (Figure S12). At four months post-inoculation, the
bacterial diversity was significantly lower (P<0.05) in standard
compost (extract treatment) compared to the control in the
organic orchard, as well as between AT-500P and the control for
the integrated orchard (Figure S12B). Furthermore, the fungal
diversity was significantly higher in the AT-500P treatment
compared to the control in the integrated orchard four-month
post-inoculation (Figure S12D).
For beta diversity, significant microbial structure differences
between orchard types and sampling time were observed: orchard
type (bacterial: R
2
= 0.14, P=0.001; fungal: R
2
= 0.30, P=0.001) and
sampling time (bacterial: R
2
= 0.06, P=0.001; fungal: R
2
=0.06,
P=0.004). An overall effect of treatment on the microbiome was
observed in the organic orchard, both after one month
(PERMANOVA; bacterial: R
2
= 0.32, P=0.02; fungal: R
2
=0.26,
P=0.001) and four months (bacterial: R
2
= 0.27, P=0.002; fungal: R
2
= 0.23, P=0.05) as shown (Tables S7,S8 for the bacterial and fungal
community, respectively). However, in the integrated orchard, a
significant effect of treatment on bacterial community structure was
observed four months after treatment (R
2
= 0.32, P=0.002), while no
significant effect (P>0.05) was seen for the fungal community
(Tables S7,S8). Pairwise significant differences (P<0.05) between
treatment and control were evidenced for the fungal community at
one-month post-inoculation (Table S8), and for the bacterial
community in both orchards at four months post-inoculation
(Table S7). The microbial community clustering by NMDS was
shown for different treatments and orchards, at one month
(Figures S13A,C), and four months (Figures S14A,C)post
inoculation, both for bacterial and fungal community, respectively.
The apple rhizosphere of the two orchards was dominated
by the bacterial phyla Proteobacteria (Organic: Integrated;
26.1%: 25.8%), Firmicutes (30.0%: 3.4%), Actinobacteria
(18.8%: 26.9%), Chloroflexi (13.1%: 15.5%), and Acidobacteria
(17.2%; 10.4%). On the other hand, the fungal community was
predominated by Ascomycota (70.1%: 68.1%), Mortierellomycota
(21.8%: 21.8%), and Basidiomycota (6.5%: 8.3%). For both
orchards, the bacterial composition was similar at one-month
post-inoculation, but a higher average relative abundance of less
abundant taxa categorized into (“others”) was seen in
rhizosphere soil treated with extract of BD manure (AT-500P)
and standard compost as compared to the control (Figure S13B).
For the fungal community, we observed that families such as
Microascaceae, and Chaetomiaceae were higher in treatments as
compared to the control (Figure S13D).
At four months post-inoculation, the family-level taxonomic
composition was similar, but differences were observed in the
less abundant taxa (“others”), whose abundance was high in
treatments as compared to the control (Figures S14B,D). In
contrast, a high abundance of the family Chaetomiaceae was
observed in treatments as compared to controls, both for organic
and integrated orchards (Figure S14D).
All microbial genera which were responsible for significant
pairwise differences in community composition between
treatments and controls were further explored. At one-month
post-inoculation, enrichment in the control of the fungal genera
Alternaria,Thelonectria, and Solicoccozyma was observed in the
organic orchard (Figure 4A). Meanwhile, the genus Chaetomium
was highly abundant in the rhizosphere which was treated with
the extract from BD manure (AT-500P). At four months post-
inoculation, bacterial genera including Bacillus,Bradyrhizobium,
Pseudolabrys, and unidentified Burkholderiacea were enriched in
the controls, compared to the treatments, while Chthniobacter
and Candidatus Xiphinemebacter were enriched in the
rhizosphere after treatments in the organic orchard
(Figure 4B). For the integrated orchard, the bacterial genera
Pseudomonas,Mycobacterium, and Candidatus Udeobacter were
enriched in the rhizosphere that had been treated with AT-500P
(extract), while Candidatus Xiphinemebacter, and uncultured
Pirellulaceae were enriched in rhizosphere treated with the
extract from standard compost (Figure 4B).
Application of extracts from standard
compost and BD introduces new
microbes into the apple rhizosphere,
especially in the integrated
apple orchard
SourceTracker was employed to reveal the portion of the
rhizosphere microbiome, which was potentially introduced by
the treatments that included extracts from standard compost
and AT-500P for organic and integrated orchards (Figure 5)at
two sampling times. We observed time-dependent differences in
the proportion of the rhizosphere microbiome that was
introduced by extracts. A higher proportion of the microbiota
at one month post-inoculation, than at four months post-
inoculation could be traced back to the treatments, especially
for the integrated orchard.
We observed that treatments introduced a higher proportion
of bacteria in integrated than in the organic orchard, especially at
one-month post-inoculation, and the average percentage
proportions ranged as follows: organic (1%-2%) and integrated
management (1%-20%). However, a high proportion of bacteria
(10% to 60%), and fungi (0%- 20%) from the extracts (i.e.
labelled as “unknown”) could not be traced in the rhizosphere
(Figure 5). For the fungal community, both in the organic and
integrated orchard, the extract from standard compost as
compared to AT-500P was found to potentially introduce
more fungi into the rhizosphere: organic (AT-500P: 0%-2% vs.
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standard compost: 0%-3%), and integrated management (0%-
1% vs. 0%-2%). Considering control soil as the microbial source,
we observed that a high proportion of the microbiome (bacteria:
40% to 90%, and fungi: 80% to 95%) was shared with the treated
rhizosphere soil.
Discussion
Biodynamic manures, composts, and plant preparations carried
distinct microbiomes. We observed a higher microbial diversity in
BD manures than in BD plant preparations and composts (apple-,
horse-, and standard composts). While the impact of type and
quantity of organic material on compost microbiomes was
previously reported (51), our study reveals for the first time
specific microbiome differences between BD manures and
composts. These variations could be attributed to various factors
such as the type of starting material and the composting technology
used (52). Moreover, the role of physical and chemical parameters
such as oxygen, temperature, moisture content, pH, and nutrient
availability shape the compost microbiome (53).
SpecifictoBDfarming,thefermentation of animal waste and
the subsequent amendment with plant preparations form a critical
part of BD products (20). This study revealed that plant
preparations and amended BD manures contain a distinct
microbiome. In addition, the microbiome varied among the
different plant preparations. Such variation among preparations
might be attributed to plant materials from which they are derived.
Moreover, the effect of the plant genotype on the microbiome has
been reviewed elsewhere (54,55). Our study provides a link between
the plant microbiome and the microbiome of plant-derived organic
amendments. We propose that the specific procedures used in
making BD manures might have contributed significantly to the
uniqueness of BD products. This is apparent from the clear
differences between composts and their BD counterparts
(Figures 1A,Bfor bacterial and fungal communities, respectively).
However, a common microbiome comprised of Paenibacillus,
Cellulomonas,andClostridium was shared between all BD
manures and composts. These taxa play essential roles as
proficient degraders of plant biomass (56). The presence of
members under the family Clostridiaceae could be attributed to
the use of cow manure in BD preparations. Clostridium is associated
B
C
A
FIGURE 4
Dot plot representation of differentially abundant genera in treatments which showed significant pairwise differences to the control for the
different orchards and sampling times. (A) shows differentially abundant fungal genera at one-month post inoculation in organic orchard, while
(B, C) represent significant differentially abundant bacterial genera between treatment and control in organic and integrated orchard at four-
months post inoculation, respectively. The significant (P<0.05) differentially abundant genera were obtained by LEfSe, and the genera with high
LDA scores (3 to 6) were shown. The [E] in the legend represents extract, and the coloured scale bar represents relative abundances, with the
lowest indicated as blue, and highest as red.
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with severe bacillary hemoglobinuria in cattle and sheep, and it was
suggested that its incidence can be reduced by the annual
vaccination of herds (57). Clostridia in traditional BD
preparations could fulfil similar functions. Additionally, Nitrogen
fixation by Clostridia (58,59), underpins the potential effect of
organic amendments in promoting beneficial soil microbial traits.
Biodynamic manures amended with preparations contained a
high abundance of Actinobacteria, a phylum known for its active
role in the composting process (60,61). Evidence linking
Actinobacteria in a consortium of Firmicutes,Proteobacteria,and
Bacteroidetes to the breakdown of lignocellulose has recently been
revealed (56). Potential plant-beneficial bacterial families like
Pseudomonadaceae, Burkholderiaceae,andBacillaceae (62)were
observed to dominate the BD formulations and plant preparations.
This suggests a potential contribution of the microbiome associated
with BD manure and plant preparations in farming systems.
Interestingly, a common microbiome of the bacterial genus
Azotobacter, and the fungal genera Fusarium and Arthrographis
was observed between BD manures and plant preparations and
further emphasizes the potential soil enrichment with beneficial
bacteria. The genus Fusarium mainly contains plant pathogenic
species (63), and was observed to be mainly associated with BD
plant preparations. However, non-pathogenic and beneficial
Fusarium species are well known (64) supporting the idea that
intraspecific microbial diversity can supress pathogenicity (3).
The fungal family Ascosphaeracea was observed in some of the
BD plant preparations and has previously been found associated
with the microbiome of bees (65). We suggest that this could
have originated from the flowers of the plants used as starting
materials in the production of BD plant preparations.
Independent of the production year (storage effect), the
microbiome of BD manures was distinct between the regions of
production (Austria and France). We suggest that various factors
including plant and animal raw materials, as well as product
handling in the two regions, have contributed significantly to the
observed microbiome differences. Our study extends this insight
to explain the likely cause of the region-specific difference
observed in BD manures. Furthermore, the microbiome
consistency across years suggests that BD manures can be
reproduced in a production line. The specific procedures to
B
CD
A
FIGURE 5
Tracking the microbiome transfers from standard compost and BD manure (AT-500P) extracts (i.e. microbiome source) into the treated
rhizosphere soil under different management systems and sampling times. Panel (A) and (B) shows the proportion of microbiota tracked from
treatment into rhizosphere in the integrated orchard for bacterial and fungal community, respectively, while (C) and (D) represent the same for
the organic orchard. The proportion of the microbiome in the treated rhizosphere which was shared with the untreated soil (i.e. control soil) is
shown both for organic and integrated orchards. The unknown part of the microbiome includes the percentage proportion of microbiota in
treatments which could not be traced in the rhizosphere. Coloured stacked bars represent proportions of microbiome in treated rhizosphere
(i.e. AT-500P and standard compost extracts), shared microbiome between control and treated rhizosphere soil, as well as the unknown
microbiome (i.e. the unassigned part of treatment microbiome). The x-axis represents time in months post inoculation. The y-axis shows the
proportion of the community originating from the different sources.
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make BD preparations have been practised for decades by the BD
farming communities and were also standardized (10,15,20),
which provides an explanation for the reproducibility.
The use of liquid extracts from BD manures as fieldspraysisa
common practice in BD farming (15). While the biochemical
characteristics of such extracts were already deciphered (66,67),
our study serves as a primer to further understand the microbiome
of BD extracts. The microbiomes of extracts and precursor
materials of composts and BD manures were distinct but shared
a common microbiome. We likely attribute the shared microbiome
to the material from which the extracts are derived; also the
procedure that materials are subjected during the fermentation.
Moreover, the common microbiome of compost, mainly composed
of Firmicutes and Actinobacteria was recently revealed (68).
Bacterial genera including Pseudomonas,Paenibacillus,
Azotobacter,andunidentified Sphingomonadaceae were associated
with standard compost extract Microbially enhanced compost
extracts were previously found to be essential in improving plant
growth and nutrition (69). Thus, their microbiome composition
could potentially explain the effects attributed to extract application
on plant health.
Several fungal genera such as Fusarium,Aspergillus Penicillium,
Mortierella,andLecanicillium were observed in extracts of the
different compost and BD manures. Interestingly, Lecanicillium has
been found to have entomopathogenic properties, which led to its
use as a biocontrol agent against aphids (70), as well as
phytoparasitic nematodes and fungi (71,72). The fungal genus
Penicillium is diverse and widely distributed in the soil environment
(73–75). It can exert positive effects through the production of
secondary products (76), as well as play critical roles in plant growth
and defence (77–79). High microbial diversity was found in
extracts, as compared to the precursor standard compost and BD
manure. This could be attributed to additives such as molasses and
stone dust which were used to make the extracts. In addition, water
as a solvent used in the extract production process may have
contributed to its microbiome. Recent studies often found high
microbial diversity in water microbiomes (80–82).
Overall, a positive impact of BD manure and compost
amendments on soil fungal diversity was found, especially at
four months post-inoculation, both for organic and integrated
orchards. Enrichment effects of BD formulations on the soil
microbiome were reported in a forgoing study (27), and further
confirmed by the current study. Previous studies indicated that
the use of BD manures and associated extracts in viticulture
affects soil biodiversity and ecosystem functions (13,18,83,84),
and plays important roles in improving soil quality (23); as well
as their potential role in biodynamic viticulture including the
enhancement of wine quality (17,25,26). In the present study,
treatments increased Acidobacteria, both in organic and
integrated orchards. Acidobacteria is a rather underrepresented
phylum, yet ubiquitous in the soil environment (85,86), and
involved in various processes in biogeochemical cycles,
decomposition of biopolymers, exopolysaccharide secretion,
and plant growth promotion (85,87,88). The fungal genus
Alternaria was found to be decreased in treatments as compared
to the untreated rhizosphere at one-month post-inoculation in
the organic orchard. Thus, we suggest that one potential role of
BD treatments is the reduction of plant pathogenic fungi, which
also includes Alternaria. This genus is ubiquitous with
saprophytic, endophytic, and pathogenic species which are
commonly found associated with various plants, including
fruits (89–91). Moreover, some Alternaria species were linked
with allergies in humans (92,93). Interestingly, strains of
Fusarium and Alternaria have recently been found to be
strongly associated with seeds of numerous crop plants, where
they are vertically transmitted to the root and rhizosphere (94).
On the other hand, the bacteria genera Pseudomonas and
Mycobacterium werefoundtobeenrichedintheapple
rhizosphere of the integrated orchard which had been treated
with extracts from BD manure. Thus, in the context of BD
farming, our findings provide insights into the potential role of
BD products especially in increasing the abundance of beneficial
bacterial genera. Moreover, the determination of various soil
quality indicators suggests larger response in the BD system,
than in conventional management systems (95,96).
Our attempt to track the microbiome from extracts to the
rhizosphere revealed that a specific proportion of the extract
microbiome was transferable. In contrast to a previous study by
(27), we observed that a higher proportion of the bacterial and
fungal community was transferred to the rhizosphere in the
integrated management orchard when compared the organic
orchard. This implies that the use of BD extracts differently
affects local soil microbiomes. This might be due to competition
with indigenous microorganisms that are present in these soils.
However, microbial signatures of organic amendment origin can
survive in the soil microbial biomass (97,98). Interestingly, even
with the low applied rates for BD preparations (i.e. 1L:10L;
extract: water) as shown by Koepf etal. (99) and the international
“Demeter”guidelines for biodynamic farming (https://demeter.
net/), we observed a significant effect of treatment on the
microbial community structure and diversity, particularly an
increase in fungal diversity during autumn for the integrated
orchard. Thus, we suggest that an increase in the frequency of
treatment application could more effectively modulate the local
soil microbiome. Moreover, the application of small quantities
(i.e. ≥5%) of suppressive soil was found to induce
suppressiveness to disease conducive soils (100,101). The low
proportion of microbes transferred to the apple rhizosphere of
the organic as compared to the integrated management orchard
could be associated with high competition from the resident
Olimi et al. 10.3389/fsoil.2022.1020869
Frontiers in Soil Science frontiersin.org14
microbiota. Limitations for the establishment of bioinoculant
strains in soil environments were previously described (102,
103), and may include competition, as well as limitation in
colonization due to different soil physico-chemical parameters.
Conclusion
The findings of this study provide first insights into the
microbiome of biodynamic manures and their applied liquid
extracts. Overall, our findings provide knowledge on approaches
to enhance microbial diversity in agricultural soils, especially in
intensively managed fields. Moreover, the in-depth assessment
of biodynamic products in orchards subjected to different field
management systems provides valuable information related to
their contribution to plant rhizosphere microbiomes.
Interestingly, the low standardization of compost amendments
criticized in the past (104),
can be an advantage in future agriculture, which needs more
diversification at all levels. We detected potentially beneficial
microbes originating from biodynamic treatments that may be
established in different farming systems upon their introduction;
although, consistent application will be needed to provide
expected benefits in the long term.
Data availability statement
The datasets presented in this study can be found in online
repositories. The names of the repository/repositories and accession
number(s) can be found in the article/Supplementary Material.
Author contributions
GB, PK, EO, and RM designed the study. EO, PK, and RM
performed the experiment and took the samples. EO performed
library preparation and other laboratory work. EO, PK and WW
analysed the data and interpreted it together with TC, SB, and
GB. EO, SB, and PK wrote the manuscript. All authors read,
corrected, and approved the final version.
Funding
We gratefully acknowledge funding by EXCALIBUR
research project with funding from the European Union’s
Horizon 2020 Research and Innovation Program under grant
agreement No. 817946 (to GB).
Acknowledgments
We sincerely thank Robert Matzer (Gleisdorf, Steirmark-
Austria) and Josef Singer (Gleisdorf Steirmark-Austria) for
allocating us experimental orchards to apply our treatments,
perform sampling, as well as in maintaining the experimental
plots. The cooperation and hospitality from their families is also
highly appreciated.
Conflict of interest
Author RM was employed by the company
BODENmanagement.net.
The remaining authors declare that the research was conducted
in the absence of any commercial or financial relationships
that could be construed as a potential conflict of interest.
Publisher’s note
All claims expressed in this article are solely those of the
authors and do not necessarily represent those of their affiliated
organizations, or those of the publisher, the editors and the
reviewers. Any product that may be evaluated in this article, or
claim that may be made by its manufacturer, is not guaranteed
or endorsed by the publisher.
Supplementary material
The Supplementary Material for this article can be found
online at: https://www.frontiersin.org/articles/10.3389/
fsoil.2022.1020869/full#supplementary-material
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