# The short-term response of soil microbial communities to digestate application depends on the characteristics of the digestate and soil type

F. Vautrin, P. Piveteau, M. Cannavacciuolo, P. Barré, C. Chauvin, C. Villenave, D. Cluzeau, K. Hoeffner, P. Mulliez, V. Jean-Baptiste, et al.

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## To cite this version:

F. Vautrin, P. Piveteau, M. Cannavacciuolo, P. Barré, C. Chauvin, et al. The short-term response of soil microbial communities to digestate application depends on the characteristics of the digestate and soil type. Applied Soil Ecology, 2024, 193, pp.art. 105105. 10.1016/j.apsoil.2023.105105. hal-04266661

HAL Id: hal-04266661

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Submitted on 12 Jan 2025

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1 The short-term response of soil microbial communities to digestate application depends

2 on the characteristics of the digestate and soil type

3 **Authors:**

4 F. Vautrin¹, P. Piveteau², M. Cannavacciuolo³, P. Barré⁴, C. Chauvin⁵, C. Villenave⁵, D.

5 Cluzeau⁶, K. Hoeffner⁶, P. Mulliez⁷, V. Jean-Baptiste⁸, G. Vrignaud⁹, J. Tripied¹, S.

6 Dequiedt¹, P.A. Maron¹, L. Ranjard¹, S. Sadet-Bourgeteau¹

7 **Affiliations:**

8 ¹ Agroécologie, INRAE, Institut Agro, Univ. Bourgogne, Univ. Bourgogne Franche-Comté,

9 F-21000 Dijon, France

10 ² INRAE, UR OPAALE, Rennes, France

11 ³ USC 1432 LEVA, Ecole Supérieure d'Agricultures (ESA), INRAE, Angers, France

12 ⁴ Laboratoire de Géologie, Ecole normale supérieure, CNRS, PSL Univ., IPSL, Paris, France

13 ⁵ ELISOL Environnement, Congénies, France

14 ⁶ University of Rennes, CNRS, ECOBIO [(Ecosystèmes, biodiversité, évolution)] - UMR

15 6553, Rennes, France

16 ⁷ Chambre régionale d'agriculture des Pays de la Loire

17 ⁸ GRDF, Paris, France

18 ⁹ ACE méthanisation, Thouars, France

19 **E-mail adresses:**

20 F. Vautrin: florianvautrin@gmail.com; P. Piveteau: pascal.piveteau@inrae.fr; M.

21 Cannavacciuolo: m.cannavacciuolo@groupe-esa.com; P. Barré: barre@geologie.ens.fr; C.
22 Chauvin: camille.chauvin@elisol-environnement.fr; C. Villenave : cecile.villenave@elisol-environnement.fr; D. Cluzeau: daniel.cluzeau@univ-rennes1.fr; K. Hoeffner: kevin.hoeffner@univ-rennes1.fr; P. Mulliez: pierre.mulliez@pl.chambagri.fr; V. Jean-Baptiste: vincent.jean-baptiste@grdf.fr; J. Tripied: julie.tripied@inrae.fr; S. Dequiedt: samuel.dequiedt@inrae.fr; P.A. Maron: pierre-alain.maron@inrae.fr; G. Vrignaud: gregory.vrignaud@gmail.com; L. Ranjard: lionel.ranjard@inrae.fr; S. Sadet-Bourgeteau: sophie.bourgeteau-sadet@agrosupdijon.fr

29 **Keywords:** Digestate; soil types; high-throughput DNA sequencing; microcosm experiment;
30 microbial community

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32 **Corresponding author:** Sophie Sadet-Bourgeteau, sophie.bourgeteau-sadet@agrosupdijon.fr, Agroécologie, INRAE, Institut Agro, Univ. Bourgogne, Univ. Bourgogne Franche-Comté, F-21000 Dijon, France

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36 **Abstract**

37 Anaerobic digestion of organic waste is a key process to produce renewable energy and meet 38 the growing demand for sustainable energy. The residues of anaerobic digestion – called 39 digestates – can be used as soil amendments to improve crop yields. However, the effect of 40 digestates on the soil biota, especially on microorganisms, needs to be better documented 41 before a large scale use of digestates in agriculture. In addition, how the quality and 42 composition of the digestate may affect soil microbial communities has not been properly 43 addressed yet. We designed a microcosm experiment under controlled experimental 44 conditions to compare effects (42 days) of four digestates produced from varying intakes
(cattle manure and/or energy crop and/or food residues and/or slurry) on soil microbial communities; a control microcosm made of undigested cattle manure was also used. Each digestate was applied on three contrasting soils representing contrasted pedo-climatic conditions (especially soil type and climate). These three soils presented different prokaryotic and fungal communities structures. The effect of digestate inputs on the soil microbial biomass and diversity was assessed using molecular DNA-based tools (quantification of extracted soil DNA and high-throughput sequencing, respectively) in comparison to the untreated cattle manure control condition. Our results show that 42 days after digestate application, significant differences of soil microbial communities were observed according to the digestate characteristics; these differences were soil-dependent. Thus, in the silty clay loam soil, no effect of digestates was observed on soil microbial biomass or diversity (P > 0.05), as compared to the undigested cattle manure. In the two other soil types (loam and sandy loam), soil microbial biomass decreased (around -40%, P<0.001) when digestates having a low total organic carbon content (from 0.61 to 3.3 g.100g⁻¹) were applied. None of the digestates affected the soil prokaryotic diversity whatever the soil type (P > 0.05). Digestate application resulted in higher fungal diversity (around +35%; P<0.001) in soils with low C/N ratio (9.14 in average). The microbial community structure of coarse-textured soil appeared more impacted by organic inputs than fine-textured soils. To conclude, our results show that different soil types, harboring distinct microbial community structures, responded differently to different digestates application. This response was also digestate-dependent.

# 1. Introduction

Biogas production has been rapidly growing in Europe for more than a decade (Appel et al., 2016), in line with Europe's target to meet 32% of renewable energy production by 2030 (European Parliament, 2018). This increase may be further amplified by the current
69 geopolitical context raising questions about Europe's gas self-sufficiency. It is also an
70 alternative way to manage farmyard manure and/or industrial organic waste while producing
71 green energy (Panuccio et al., 2021). In addition to biogas production, anaerobic digestion
72 produces digestate – a by-product used as an organic fertilizer (Pivato et al., 2016). Thus,
73 biogas production contributes to promote circular bioeconomy by closing the nutrient loop, it
74 contributes also to the mitigation of greenhouse gas emissions from farm effluents and
75 increases carbon sequestration in soils (Nayal et al., 2016; Tambone et al., 2017).

76 According to the literature, the organic matter of digestates may improve the soil
77 structure, its infiltration rate and its water-holding capacity (Barłóg et al., 2020; Makádi et al.,
78 2012; Möller and Müller, 2012), and thus increase productivity while lowering fertilization
79 costs. However, thorough investigations of possible side effects on the soil biota – especially
80 on microorganisms that play important roles in the soil ecosystem – are required to properly
81 assess the sustainability of large-scale land application of digestates in agricultural fields.

82 It is now well established that organic amendments improve soil microbial communities
83 by increasing the soil microbial biomass (Chaudhry et al., 2012; Francioli et al., 2016; Ros et
84 al., 2006) and inducing shifts of prokaryotic and fungal community structures (Chaudhry et
85 al., 2012; Francioli et al., 2016; Ros et al., 2006; Sadet-Bourgeteau et al., 2018). The
86 response of microbial communities to organic waste inputs depends on the soil type and on
87 the nature of the organic materials (Sadet-Bourgeteau et al., 2019). In addition, it is important
88 to consider the time lag between organic material application and their impact on the soil.
89 Transient changes in the bacterial functional and genetic structures occur within the first three
90 months following organic amendment. Then, a period of resilience is observed, eventually
91 leading to similar communities in both the amended and control plots (Calbrix et al., 2007).
92 This strong transient alteration of soil microbial functioning may lead to modifications of the
93 carbon and nitrogen cycles (Sadet-Bourgeteau et al., 2019). This could explain why the period
within days directly following land application of organic amendments is considered of the highest importance for environmental hazards. However, the impact of digestates on the soil microbial communities has not been extensively studied on the short term, i.e. up to eight weeks after application. In addition, to our knowledge, no studies assessed this effect on soil microbial communities from different soil types.

Reports on the impacts of digestates on the soil microbiota are scarce, and those on the short-term effects of digestates on the soil microbial communities are rather contradictory, as reviewed by Karimi *et al.* (2022). Some studies suggest that biogas digestates increase the soil microbial biomass (Caracciolo et al., 2015; Cattin et al., 2021; García-Sánchez et al., 2015; Muscolo et al., 2017; Panuccio et al., 2021) and prokaryotic diversity (Caracciolo et al., 2015; García-Sánchez et al., 2015) from 21 to 180 days after inputs compared to unfertilized soil. Other studies show no significant effect of digestate inputs on the soil microbial biomass or prokaryotic diversity from 1 to 10 weeks after application (Akari and Uchida, 2021; Barduca et al., 2021; Podmirseg et al., 2019). To our knowledge, the short-term impact of digestates on the soil fungal community has never been assessed. Interestingly, a short-term effect of digestates on the microbial community structure is consistently observed whatever the experimental conditions (Caracciolo et al., 2015; Gómez-Brandón et al., 2016; Hupfauf et al., 2016; Johansen et al., 2013; Sapp et al., 2015), leading to growth of *Actinobacteria* and *Acidobacteria*, a decrease of the *Bacteroidetes* (Sapp *et al.*, 2015) or a shift in the ammonia-oxidizing bacteria (AOB) community (Gómez-Brandón *et al.*, 2016). Differences in experimental setups and conditions could explain conflicting results observed for some microbial indicators: the soil type, the digestate nature and dose, the delay between digestate application and soil sampling, the presence and type of vegetation varied across studies. All these factors make it difficult to reach generic conclusions about the impact of digestates on
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the soil microbial quality. However, as previously mentioned, studies combining different
soil types and contrasting digestates are lacking (Karimi et al., 2022).

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Therefore, thorough investigations of possible side effects on the soil biota – especially
on microorganisms that play important roles in the soil ecosystem – are required to properly
assess the sustainability of large-scale land application of digestates in agricultural fields.

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Based on previous studies, we hypothesized that the responses of microbial communities to digestate depend on the soil type and on the nature of digestate. Therefore, the objective of the present study was to compare, under controlled experimental conditions, the short-term impact of different types of digestates on soil microbial communities, and to assess if these impacts depended on the type of soil. This laboratory microcosm experiment was designed with four types of digestate and three soils, with a range of physicochemical and climatic characteristics, selected to represent different microbial communities. Soils were sampled from fields that had never received any digestate amendment. The four digestates mainly differed in their physicochemical composition, especially their C/N ratio and nutrient content.

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Digestates were applied to the microcosms at an agronomic dose of 25 t.ha⁻¹. The responses of prokaryotic and fungal communities (microbial biomass, 16S and 18S ribosomal gene inventories) were assessed 42 days after digestate application. The novelty of this work is to address concomitantly the factors “type of digestate” and “type of soil” within the same experimental setup to assess how these factors affect the responses of prokaryotic and fungal communities upon digestate application to soil. High throughput sequencing respectively targeting 16S and 18S ribosomal genes was used to characterize the vast microbial diversity of soil and assess microbial communities’ composition changes after amendment.

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## 2. Materials and methods

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2.1. *Soils, biogas digestates and control manure used in this study*
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Three soils were selected according to their pedoclimatic diversity. One silty clay loam soil was collected from the continental Burgundy region (BOU, 47°38′86.88N; 5°06′29.01E). One loam soil originated from the oceanic Pays de la Loire region (PDL, 47°72′86.90N; 0°49′71.88W). The third soil, of sandy loam texture, came from the Mediterranean Provence-Alpes-Côte d'Azur region (PACA, 43°17′80.55N; 6°20′88.88E). These three soils, thereafter called BOU, PDL and PACA did not have any record of digestate land application; BOU and PDL were under annual cropping systems, while PACA was a vineyard soil.

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The top layer (0-20 cm) of these soils was sampled in February 2021. Each soil sample was homogenized and sieved to 4 mm to remove aboveground plant debris, roots and stones. An aliquot was air-dried for physicochemical analysis. Particle size distribution, pH, soil total organic carbon (TOC), soil total nitrogen (Total N), soil C/N ratio, Olsen P, Cation Exchange Capacity (CEC) and exchangeable cations (calcium, magnesium, sodium and potassium) were determined by the Soil Analysis Laboratory of INRAE (Arras, France, https://www6.hautsdefrance.inrae.fr/las). The water-holding capacity (WHC; gravimetric humidity) was estimated in the laboratory. The initial soil physicochemical parameters are summarized in Table 1. The remaining soil was used for microcosm setup.

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Whole digestates were collected from four commercial anaerobic digestors fed with contrasting input organic materials: i) **FYM**: farmyard manure – 75% cattle manure, 10% cattle slurry, 10% energy crop and 5% milk-derived matter, ii) **CMF**: cattle manure + fat – 45% duck slurry, 45% biowaste products, 10% fatty waste from agroindustry, iii) **MFW**: manure + food waste – 52% manure, 17% vegetables, 19% waste from agroindustry, 10% slurry and 2% blood and iv) **SMS**: silage maize and sorghum – 90% energy crop and 10% maize silage. FYM underwent a sanitation stage at 70 °C for 1h. Undigested cattle manure (**CM**) from a dairy farm was chosen as control to assess conventional agronomic practices. All fresh organic amendments were sent to Aurea Co. (France, https://aurea.eu/) to determine
moisture, total nitrogen (Total N), pH, and exchangeable cations (phosphorus, potassium, calcium, magnesium and sodium). The carbon fractions (total organic carbon (TOC), soluble organic carbon (SOC), hemicellulosis, cellulosis and lignin-cutin) were assessed by the Regional Laboratory of Analyses and Research (France, https://aisne.com/a-votre-service/laboratoire-departemental). The physicochemical characteristics of the organic amendments are summarized in **Table 2**.

## 2.2. Microcosm setup and incubation

The experimental design consisted of a factorial arrangement with three soils (BOU, PACA and PDL), five amendments (four different types of digestates (CMF, FYM, MFW, SMS), one untreated organic amendment (CM) and one unfertilized control (sterile water). Four replicate microcosms were prepared per condition. In total, 72 microcosms (3 soils × 6 treatments × 4 replicates) were prepared. Unfertilized control microcosms were added in order to take into account the natural evolution of the microbial communities during incubation of soil microcosms, as reported previously (Sadet-Bourgeteau et al., 2019; Tardy et al., 2014). Microbial community results after 42 days of incubation of these unfertilized control were subtracted to those of the treated samples. Thus, for each soil type, results were expressed as percent differences between the test condition (CMF, FYM, MFW, SMS or CM) and unfertilized control.

For each condition, four sterile flasks (250 mL) were filled with 60 g (dry weight) of soil. After two weeks of pre-incubation at 15 °C, 0.682 g of organic waste products were thoroughly mixed with a soil sample to reach the agronomic dose of 25 t.ha⁻¹ according to agricultural practices in French cropping systems. Sterile water was added to these microcosms to adjust the water content to 70 % soil water-holding capacity (WHC). The flasks were incubated in the dark at 15 °C for 42 days to simulate spring weather conditions.
Every week, they were aerated under a biological safety cabinet, and the water content was adjusted (70% of the soil WHC) when necessary to correct for any water loss through evaporation. Germinating plants were removed when required. After incubation 42 days incubation, samples were harvested, lyophilized and stored at -40 °C until use for molecular analyses based on soil DNA extraction. Aliquots of each of the four replicates were pooled, and soil chemical analyses (except particle size distribution) were performed by the Soil Analysis Laboratory at the INRAE of Arras (France) as described for the initial stage of the experiment.

## 2.3. DNA Extraction, purification and molecular microbial biomass determination

Microbial DNA was extracted from 1 g (dry weight) of soil according to a standardized procedure of the GenoSol platform (INRAE, Dijon, France, https://www2.dijon.inrae.fr/plateforme_genosol/) (Terrat et al., 2012). Since a highly positive linear relationship has been shown between soil DNA recovery and C-biomass measurement (Dequiedt et al., 2011; Marstorp et al., 2000), DNA concentrations of crude extracts were determined by electrophoresis in 1% agarose gel stained with ethidium bromide using a calf thymus DNA standard curve, and used as estimates of microbial biomass (Dequiedt et al., 2011). After quantification, residual impurities were separated from nucleic acids by centrifuging 100 µL of crude DNA through two types of minicolumns (Macherey-Nagel, Hoerdt, France) according to the manufacturer's instructions. Purified DNA concentrations were finally measured on a 5-µL load using the Quantifluor staining kit (Promega, Lyon, France) according to the manufacturer's protocol.

## 2.4. High-throughput sequencing of 16S and 18S rRNA gene sequences

For prokaryotic diversity (16S), the V3-V4 region of the bacterial chromosome was targeted using primers F479 (5'-CAG CMG CYG CNG TAA NAC-3') and R888 (5'-CCG
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YCA ATT CMT TTR AGT-3') on a 440-bp fragment (Tardy et al., 2014). For each sample, 5 ng of DNA were used in a final PCR volume of 25 µL, and amplified under the following conditions: 94 °C for 2 min, 35 cycles of 30 s at 94 °C, 30 s at 52 °C and 1 min at 72 °C, followed by 7 min at 72 °C (Tardy *et al.*, 2014).

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As demonstrated by George *et al.*, 2019, soil fungal richness follows the same trends across land use, irrespective of primer set (18S rRNA or ITS (internal transcribed spacer)). Thus, as proceeded in previous studies (Chemidlin Prévost-Bouré *et al.*, 2011, Sadet-Bourgeteau *et al.*, 2019) we used a 350-bp 18S rRNA fragment to target fungi with primers FR1 (5'-ANC CAT TCA ATC GGT ANT-3') and FF390 (5'-CGA TAA CGA ACG AGA CCT-3') (Chemidlin Prévost-Bouré et al., 2011). For each sample, 5 ng of DNA were used in a final PCR volume of 25 µL and amplified under the following conditions: 94 °C for 3 min, 35 cycles of 30 s at 94 °C, 1 min at 52 °C and 1 min at 72 °C, followed by 5 min at 72 °C (Sadet-Bourgeteau *et al.*, 2019).

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All PCR products were purified using the ProNex® Size-Selective DNA Purification System (Promega, Lyon, France) and quantified with the Quantifluor staining kit (Promega, Lyon, France) according to the manufacturer's instructions.

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Series of 10-bp multiplex identifiers were added to the 5' end of the primers to identify each sample and prevent PCR biases. For bacteria and archaea, 7.5 ng of DNA were used and amplified with seven cycles under the same conditions as those of the first PCR. For fungi, 5 ng of DNA were used under optimized PCR conditions with a denaturation step at 94 °C for 1 min and seven cycles.

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The PCR products were purified with the MinElute gel extraction kit (Qiagen NV) and quantified with the Quantifluor staining kit (Promega, Lyon, France) according to the manufacturer's instructions. All samples were pooled in equal amounts and cleaned with the

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solid-phase reverse immobilization method (SPRI) using the ProNex® kit (Promega, Lyon, France). The pool was finally sequenced with a MiSeq Illumina instrument (Illumina Inc, San Diego, CA) operating with V3 chemistry and producing 2x300 bp paired-reads by GenoScreen (https://www.genoscreen.fr/fr/).

## 2.5. Bioinformatic analysis of the 16S and 18S rRNA gene sequences

All paired-end Illumina MiSeq reads were processed using the BIOCOM-PIPE, as described by Djemiel et al. (2020). The parameters retained for each step can be found in **Supplementary Table S1**. The resulting high-quality reads were used for (i) taxonomy-independent analysis, determining a biodiversity richness index, and (ii) taxonomy-based analysis using similarity approaches against dedicated reference databases from SILVA v132 (Quast et al., 2012). The raw datasets are available in the EBI database system under project accession PRJEB53389.

## 2.6. Statistical analysis

All datasets were analyzed with R software (R version 4.1.3) (Verzani, 2004). As previously mentioned, in order to take into account the natural evolution of the soil microbial community throughout the incubation (molecular microbial biomass and microbial richness), the results at the end of the experiment (42 days) from the soils that only received sterile water (un-fertilized control) were subtracted from those of the treated samples. Then, the results were expressed as percent differences. Normality was tested for each parameter using Shapiro-Wilk's test followed by Bartlett's test of the homogeneity of variances. Then, an analysis of variance (ANOVA) and a *post-hoc* Tukey's honestly significant difference (HSD) test ($P < 0.05$) were performed to test the effect of soil type, organic materials type and soil type × organic material type on physicochemical properties and microbial community
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indicators (microbial biomass, prokaryotic and fungal richness). The effects with p-values < 0.05 were considered statistically significant.

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In order to highlight the organic amendment or soil physicochemical parameters involved in changes in the microbial community indicators, a redundancy analysis (RDA) was performed, based on the following parameters: soil granulometry, total nitrogen (Total N), total organic carbon (TOC), C/N ratio, pH, Olsen P, together with the following organic amendment variables: total nitrogen (Total N), total organic carbon (TOC), SOC, C/N ratio, pH, Olsen P. These variables were used to evaluate whether the soil and organic inputs physico-chemical characteristics discriminated the different treatments. To determine the abiotic effects on the microbial community indicators, the *rda* and *ordistep* functions (*vegan* package) were used in R software (R version 4.1.3) (Verzani, 2004).

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The prokaryotic and fungal community structures were also compared using a taxonomic abundance matrix at the phylum level. Non-metric multidimensional scaling (nMDS) was performed on raw data (no subtraction of the unfertilized control) to graphically depict the distribution patterns of the microbial community structures across treatments, constructed with the 1% most abundant phyla. Differences between treatments were assessed by non-parametric PERMANOVA tests (P < 0.05) run with the *adonis* function in the *vegan* package. Explanatory phyla and physicochemical parameters were fitted on the graphs (Total N, TOC, SOC, C/N ratio, pH and Olsen P for the organic amendments; Total N, TOC, C/N ratio, pH, Olsen P, clay and silt for the soils).

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# 3. Results

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## 3.1. Soil physicochemical characteristics

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Before organic material application, the three soils exhibited contrasting textures, with a greater clay content in BOU (386 g clay.kg⁻¹ dry soil) and a greater sand content in PACA

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(666 g sand.kg⁻¹ dry soil), whereas PDL was considered as a loamy soil (392 g silt kg⁻¹ dry soil) (Table 1). Forty-two days after organic material application, the Principal Component Analysis (PCA) of the soil physico-chemical characteristics highlighted that soil samples were not discriminated according to organic treatments, but remained mainly discriminated by their type (Fig. 1). Thus, at the end of the experiment, BOU was characterized by a higher pH (7.9), higher TOC (20.57 g.kg⁻¹ dry soil) and nitrogen (1.79 g.kg⁻¹ dry soil) contents (Fig. 1). PACA was characterized by lower TOC (5.99 g.kg⁻¹ dry soil) and nitrogen (0.63 g.kg⁻¹ dry soil) contents (Fig. 1). PDL was characterized by a lower pH (6.8).

## 3.2. Effects of digestate inputs on soil microbial community indicators

Forty-two days after organic material application, an effect of soil, digestate and soil×digestate interaction was observed for all measured soil microbial indicators (molecular microbial biomass, prokaryotic and fungal richness; **Fig. 2, 3 and 4**; P < 0.001). More precisely, in BOU, no significant digestate-related difference was observed (P > 0.05; **Fig. 2, 3 and 4**). All digestates (CMF, FYM, MFW and SMS) induced the same effect on the soil microbial biomass and the microbial diversity indexes (prokaryotic and fungal richness) as cattle manure did.

In PACA, 42 days after application, most of digestates (CMF, FYM and SMS) inputs induced lower microbial biomass compared to CM (-48%, -53% and -37%, respectively; P < 0.001; **Fig. 2**). No significant difference in soil microbial biomass was observed between MFW- and CM-amended PACA (P > 0.05; **Fig. 2**). In PACA soil, digestate application induced no differences of prokaryotic richness in comparison to the CM control, excepted for MFW-treated PACA, which exhibited 20% lower prokaryotic diversity compared to CM treatment (P < 0.001, **Fig. 3**). Regarding PACA soil fungal richness, no significant differences were
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observed between digestates (CMF, FYM, MFW and SMS)- and CM-treatments (P > 0.05; **Fig. 4**).

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In PDL, 42 days after application, significant lower soil microbial biomass was observed when CMF and SMS were applied compared to CM (-44% and -48%, respectively; P < 0.001; **Fig. 2**). Regarding soil prokaryotic diversity, no difference was noted between CM- and digestates-amended PDL (P > 0.05; **Fig. 3**). On the opposite, all digestates induced 21% higher fungal richness in PDL soil compared with cattle manure input (P < 0.001; **Fig. 4**).

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The redundancy analysis (RDA) approach allowed us to decipher the determinants of these variations of soil microbial parameters. Whatever the soil considered, the total organic carbon content of the inputs (29% of the variance due to the TOC) and the silt content (30% of the variance) of soil explained the significant variations of soil molecular microbial biomass (**Supplementary Table S2**). Concerning the prokaryotic diversity, none of the physicochemical characteristics tested could explain observed variance. Fungal diversity variation was explained by the soil C/N ratio (67% of the variance) (**Supplementary Table S2**).

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### 3.3. Soil microbial community structure

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The nMDS ordination highlighted distinct prokaryotic and fungal community structures among the three unfertilized soils at the end of the 42-day incubation (**Supplementary Fig. S1 A and B**). Consequently, the microbial structures of each soil, 42 days after organic amendment application, were analyzed separately. In BOU, the prokaryotic and fungal community structures differed among treatments at the end of the 42-day incubation (**Fig. 5A and Fig. 6A**). The prokaryotic community structure of CMF-, FYM- and MFW-amended BOU seemed to differ from those of CM- and SMS-amended BOU (**Fig. 5A**). However, PERMANOVA analysis highlighted that prokaryotic community structure of BOU soil

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amended with digestates or undigested cattle manure was similar 42 days after organic
materials inputs (P > 0.05, **Supplementary Table S3**). The fungal community structure of
CMF- and MFW-amended BOU differed from those of CM-amended BOU (Fig. 6A).

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PERMANOVA analysis confirmed these observations highlighting significant differences
between fungal community structures from BOU soil amended with CM and those amended
with the following digestates: CMF, FYM and MFW (P < 0.05, **Supplementary Table S4**). A

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greater abundance of *Ascomycota* was found in CM-amended BOU. BOU soil amended with
digestates such MFW and CMF recorded a greater abundance of *Mucoromycota*,
*Cryptomycota* and *Basidiomycota* (Fig. 6A). FYM-amended BOU soil was supported by a

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greater abundance of *Chytridiomycota* (Fig. 6A).

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The prokaryotic community structure of CM-amended PACA significantly differed from
FYM and CMF treatments (Fig. 5B, PERMANOVA, **Supplementary Table S3**, P < 0.05).

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To note that PERMANOVA analysis highlighted that the prokaryotic structure from PACA
amended with CMF differed from those amended with SMS (Supplementary Table S3, P <
0.05). These differences were supported by a greater abundance of *Bacteroidetes*, *Chloroflexi*
and *Spirochaetes* and a lower abundance of *Firmicutes*, *Actinobacteria* and *Cloacimonetes* in
CM-amended PACA than in CMF-, FYM-, MFW- and SMS-amended PACA (Fig. 5B).

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Regarding the fungal community structure, SMS-amended PACA differed from all other
amended PACA, CM included (Fig. 6B, PERMANOVA, **Supplementary Table S4**, P <
0.05). PACA amended with SMS recorded a greater abundance of *Basidiomycota* (Fig. 6B).

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The prokaryotic community structure of CM-amended PDL seemed to differ from all
digestates-amended PDL (Fig. 5C), but this was not confirmed by PERMANOVA analysis,
(Supplementary Table S3, P > 0.05). Regarding the fungal community structure, CM-
amended PDL seemed to differ from FYM-, MFW- and CMF-amended PDL (Fig. 6C). This
was not confirmed by PERMANOVA analysis, (Supplementary Table S4, P > 0.05),

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<figure><img src="image_3.png" /></figure>

however, CMF-amended PDL differed from MFW- and SMS-amended PDL (PERMANOVA, **Supplementary Table S4**, P < 0.01). As observed in BOU soil, PDL soil amended with CMF recorded a greater abundance of *Mucoromycota* and *Cryptomycota* (**Fig. 6C**).

<figure><img src="image_4.png" /></figure>

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## 4. Discussion

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<figure><img src="image_25.png" /></figure>

Widespread development of anaerobic digestion is expected in the coming years to meet the rising demand for green energy. As a result, agronomic use of digestates may be generalized. However, impacts of digestates on soil biodiversity still need to be evaluated to prevent potential environmental side effects associated to the use of these inputs across a wide range of biogeographic regions and soil contexts. To our knowledge, the comparison of different digestates in different soil types has never been addressed simultaneously under the same controlled experimental conditions. Contrasting digestates were selected and raw cattle manure was added as control treatment to illustrate the most common organic materials used as crop amendments. A dose of digestate of 25 t.ha⁻¹ was applied to mimic common agronomic practices in French fields (Monard *et al.*, 2020). Application of the same agronomic dose of digestates with contrasting composition resulted in different doses of elements brought, especially nitrogen. Interestingly, the total nitrogen and C/N ratio of CMF differed from those of the other digestates: the nitrogen content was two times higher in CMF (the largest amount) than in SMS, and TOC was seven times lower in CMF than in FYM. These differences in carbon and nitrogen amounts induced a high variability of the C/N ratio across treatments, probably leading to different fates of nitrogen in the amended soils. A C/N ratio > 15 (e.g. in CM) leads to nitrogen immobilization in the soil, whereas a low C/N ratio leads to NOx emissions, so that a value 5 ≤ C/N ratio ≤ 6 was proposed to be optimal
381 (Abubaker et al., 2015; Cheng et al., 2020). The differences of C/N ratios of the applied
382 organic materials led us to expect differences among the microcosms.

383 Differences in molecular microbial biomass were observed depending on the soil type
384 and the digestate. Microbial biomass remained unchanged in the silty clay loam soil (BOU)
385 regardless of the treatment, but decreased in the sandy loam (PACA) and loam (PDL) soils
386 down to 48% under CMF and SMS treatment compared to the CM treatment. The effects of
387 digestate on soil microbial biomass reported in the literature are often conflicting, mainly due
388 to great variations between the experimental systems. Thus, several authors observed an
389 increase of soil microbial biomass (assessed by fumigation extraction or soil DNA
390 measurement) after digestate application (Cattin et al., 2021; García-Sánchez et al., 2015;
391 Muscolo et al., 2017; Panuccio et al., 2021), whereas others reported no change (Barduca et
392 al., 2021; Podmirseg et al., 2019). In the present study, redundancy analysis highlighted silt
393 content as a major driver of the observed differences according to the soil type. This suggests
394 that the soil texture can influence the response of microbial biomass to digestate application.
395 This hypothesis is supported by Panuccio et al. (2021), who found greater changes of
396 microbial biomass in sandy soil than in silty soil both amended with digestate. Fine-textured
397 soil is indeed known to be a favourable habitat for microbes because of a better supply of
398 organic resources and protection from desiccation, gas diffusion, toxic exogenous compounds
399 and predation by protozoa (Dequiedt et al., 2011; Ranjard et al., 2003).

400 Our work also highlighted total organic carbon of digestates as a second major driver
401 of soil microbial biomass variations. Thus, the lower microbial biomass quantified in some
402 digestate-amended soils than in CM-amended soils can be explained by the low total organic
403 carbon content of the digestates, hence a lower amount of organic carbon available for
404 microbial growth.
405

As observed for the soil microbial biomass, 42 days after organic material application, microbial diversity variations depended on the characteristics of the digestate applied and the soil type. This was consistent with previous results collected from different soil microbial communities amended with other types of organic waste products (Sadet-Bourgeteau et al., 2019). A increase of prokaryotic richness could be observed but a significant decrease was observed in MFW-amended sandy loam soil (PACA) (20% less than in CM-amended soil). These results are not in accordance with the literature available where an increase of soil prokaryotic diversity (Shannon) was reported after digestate application (Caracciolo et al., 2015; García-Sánchez et al., 2015; Hupfauf et al., 2016). These conflicting results could be due to the sensitivity of the method used (fingerprint techniques, PLFA... versus 16S metabarcoding), on the difference of metric used (Shannon vs Richness), and, more largely, on the differences between experimental designs (soil type, digestate...).

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In the present work, the prokaryotic community structures of the three unfertilised soils (control microcosms) were significantly different (Supplementary Fig. S1). Whatever the treatment and soil type, at the end of the experiment the prokaryotic community structures at the phylum level were dominated by *Proteobacteria*, *Bacteroidetes*, *Thaumarchaeota* and *Actinobacteria*, major taxa found in French soils (Karimi *et al.*, 2018), but changes of microbial community structure were detected depending on the soil type and the quality of the amendment applied (Fig. 4 and 5). In comparison with manure amendment, differences in prokaryotic community structures were significant only in sandy loam soil (PACA) amended with FYM and CMF digestates. Similar trends were recorded in the loam (PDL) and silty clay loam (BOU) soils, but differences were not significant. As already mentioned for the soil microbial biomass, the coarse texture also influenced the prokaryotic community structure more than the fine texture did. This is consistent with other studies highlighting an effect of digestate on prokaryotic community structure limited to coarse-textured soils (Gómez-
Brandón et al., 2016; Hupfauf et al., 2016; Johansen et al., 2013; Sapp et al., 2015). This was confirmed by Abubaker *et al.* (2013) who observed greater variability of community structure in sandy soil than in clay soil. To summarize, our results highlighted that the observed effects on distinct prokaryotic community structures were digestate- and soil-dependent. However, in the three studied soils, the CM and CMF treatments consistently led to the most different soil prokaryotic community structures, with *Spirochaetes* more abundant in all CM-amended soils and *Actinobacteria* in all CMF-amended soils. This can be explained by the abundance of *Spirochaetes* in manure because *Spirochaetes* are known free-living, anaerobic symbionts in the stomachs of cows and other ruminants (Gupta et al., 2013). Thus, they may have come from non-sanitized livestock manure. CMF was the digestate with the largest amount of nitrogen and $NH_4^+$. High amounts of ammonium ions lower the soil pH and modify the microbial community structure (Chen et al., 2020). Furthermore, *Actinobacteria* are stimulated in acidic soils (Karimi et al., 2018). In CMF, these conditions probably favoured the abundance of *Actinobacteria*. This is also in accordance with a previous study highlighting that the proportion of *Actinobacteria* was significantly higher in inorganically fertilized (NPK) soils (Francioli *et al*., 2016).

Increases of fungal richness were captured but only in loam soil (PDL) amended with CMF and FYM (21% more than in CM-amended soil). To our knowledge, this is the first report of the effect of digestates on soil fungal diversity. In the present study, the redundancy analysis did not explain prokaryotic richness variation, but a 67% negative effect of the soil C/N ratio of digestates on fungal richness was captured. This is in accordance with previous works reporting that the soil fungal richness was negatively correlated with the soil C/N ratio (George et al., 2019; Yang et al., 2017).

The fungal community structure was influenced by the input type, but the soil type balanced this effect. To our knowledge, the impact of digestate application on soil fungal
community structure had never been assessed before. In the present study, the sandy loam soil (PACA) exacerbated the effect of organic inputs on soil fungal community structure compared to the silty clay loam soil (BOU). In the three soils, the communities were dominated by *Ascomycota*, *Mucoromycota* and *Basidiomycota* whatever the condition, as frequently observed in French soils (Maron *et al.*, 2018). Contrary to what was observed with the soil prokaryotic communities, different fungal phyla could be stimulated by the same organic material depending on the type of soil in which it was applied. This suggests that different soil types, harbouring distinct fungal community structures, responded differently to digestates application.

The present work monitored under controlled conditions enabled us to address the impact of the characteristics of digestates on the response of soil microbial communities. Moreover, high throughput sequencing was appropriate to characterize the vast microbial diversity of soil but it provides limited information on soil microbial functioning. In order to better characterize the environmental impacts of digestates, further research is needed under field conditions. Future field studies should be implemented with repeated applications of digestates over several years, in order to address holistically the sustainability of digestate application. A special focus on microbial communities involved in biogeochemical cycles would be of interest in combination with measurements of GHG emissions.

473

## 5. Conclusions

The forthcoming increase of the amounts of anaerobic digestates produced requires a proper assessment of the environmental safety of land application of digestates, notably for soil health and the telluric microorganisms that play key roles in biogeochemical cycles. This study provided new insights regarding the short-term effects of contrasting anaerobic
digestates on soil microorganisms. Our results highlighted that the response of soil microbial communities to the application of digestate depended firstly on the characteristics of the digestate and secondly that these effects of digestate on soil microbial community depended on the soil type. Indeed, coarse-textured soil was most affected that the finest ones.

Importantly, when applied on coarse-textured soil, digestates characterized by a low C/N ratio (high levels of ammonium and low levels of organic carbon) induced the greatest impact on soil microbial communities (lower microbial biomass and a shift of microbial structure and composition) compared to untreated manure amendment. This pattern suggests that digestates with low C/N ratios could be recommended for fertilisation of heavy soils with high clay and C content while digestates with higher C/N ratios seem more appropriate for lighter, sandier soils, containing less organic carbon.

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## 6. Acknowledgments

The authors thank the GenoSol platform for technical support https://doi.org/10.15454/L7QN45 in molecular biology. The authors are grateful to the OPAALE (Optimisation des Procédés en Agriculture, Agroalimentaire et Environnement) and LEVA (Légumineuses, Ecophysiologie Végétale, Agroécologie) technical staff for their technical support for soil and biogas digestate harvesting and microcosm experiment setup, and Aurélien Cottin for his help with the bioinformatic tools. Calculations were performed using HPC resources from DNUM CCUB (Centre de Calcul de l'Université de Bourgogne). They also thank Annie Buchwalter for correction and improving English language in the manuscript.

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## 501 Funding
502

This work was supported by grants GRAINE from the ADEME, the CASDAR Innov et Part

503

Metha-BioSol and GrDF.

504

**Declaration of competing interest**

505

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

506

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<figure><img src="image_26.png" /><figcaption>**Figure 1.** Principal Component analysis (PCA) performed on the soil physicochemical characteristics 42 days after organic materials application (CM; CMF; FYM; MFW; SMS) or sterile water (Unfertilized control) for the three soils (BOU; PACA and PDL).</figcaption></figure>

Abbreviations: BOU, Bourgogne; PACA, Provence-Alpes-Côte d'Azur; PDL, Pays de la Loire; CM, cattle manure; CMF, cattle manure + fat; FYM, farmyard manure; MFW, manure + food waste; SMS, silage maize and sorghum; Total N, total nitrogen; TOC, total organic carbon; C/N, carbon/nitrogen ratio; pH, potential of hydrogen; P₂O₅, phosphorus pentoxide; K₂O, potassium oxide; MgO, magnesium oxide; Na₂O, sodium oxide.
<figure><img src="image_27.png" /><figcaption>**Figure 2.** Effect of different organic materials (digestate and cattle manure) on soil microbial biomass from three soils, 42 days after application.</figcaption></figure>

In order to take into account the natural evolution of soil microbial biomass throughout the incubation, results at the end of the experiment (42 days) from un-fertilized control were subtracted from those of the treated samples; results were expressed as percent of differences.

Crossed effect soil x digestate, significant at the 0.05 probability level.

Abbreviations: BOU, Bourgogne; PACA, Provence-Alpes-Côte d'Azur; PDL, Pays de la Loire; CM, cattle manure; CMF, cattle manure + fat; FYM, farmyard manure; MFW, manure + food waste; SMS, silage maize and sorghum.
<figure><img src="image_28.png" /><figcaption>**Figure 3.** Effect of different organic materials (digestate and cattle manure) on the prokaryotic richness from three soils, 42 days after application.</figcaption></figure>

In order to take into account the natural evolution of soil prokaryotic richness throughout the incubation, results at the end of the experiment (42 days) from un-fertilized control were subtracted from those of the treated samples; results were expressed as percent of differences.

Crossed effect soil x digestate, significant at the 0.05 probability level.

Abbreviations: BOU, Bourgogne; PACA, Provence-Alpes-Côte d'Azur; PDL, Pays de la Loire; CM, cattle manure; CMF, cattle manure + fat; FYM, farmyard manure; MFW, manure + food waste; SMS, silage maize and sorghum.
<figure><img src="image_29.png" /><figcaption>**Figure 4.** Effect of different organic materials (digestate and cattle manure) on the fungal richness from three soils, 42 days after application.</figcaption></figure>

In order to take into account this natural evolution of soil fungal richness throughout the incubation, results at the end of the experiment (42 days) from un-fertilized control were subtracted from those of the treated samples; results were expressed as percent of differences.

Crossed effect soil x digestate, significant at the 0.05 probability level.

Abbreviations: BOU, Bourgogne; PACA, Provence-Alpes-Côte d'Azur; PDL, Pays de la Loire; CM, cattle manure; CMF, cattle manure + fat; FYM, farmyard manure; MFW, manure + food waste; SMS, silage maize and sorghum.
<figure><img src="image_30.png" /></figure>

<figure><img src="image_31.png" /></figure>

<figure><img src="image_32.png" /></figure>
**Figure 5.** Non-metric multi-dimensional scaling (nMDS) ordination plot derived from the taxonomy table of the prokaryotic phyla (abundance > 1%) from three soils ((A) BOU, (B) PACA, and (C) PDL), 42 days post treatment.

Abbreviations: CM, cattle manure; CMF, cattle manure + fat; FYM, farmyard manure; MFW, manure + food waste; SMS, silage maize and sorghum. The most explanatory phyla and soil chemical parameters are fitted on the plot. TOC, total organic carbon; C/N, carbon/nitrogen ratio; pH, potential of hydrogen.

*Colors are not mandatory for this figure in print.*
<figure><img src="image_33.png" /></figure>

<figure><img src="image_34.png" /></figure>

<figure><img src="image_35.png" /></figure>
**Figure 6.** Non-metric multi-dimensional scaling (nMDS) ordination plot derived from the taxonomy table of the fungal *phyla* (abundance > 1%) from three soils ((A) BOU, (B) PACA, and (C) PDL), 42 days post treatment.

Abbreviations: CM, cattle manure; CMF, cattle manure + fat; FYM, farmyard manure; MFW, manure + food waste; SMS, silage maize and sorghum. The most explanatory *phyla* and soil chemical parameters are fitted on the plot. TOC, total organic carbon; C/N, carbon/nitrogen ratio; pH, potential of hydrogen.

*Colors are not mandatory for this figure in print.*
<table><caption>**Table 1.** Initial physicochemical characteristics of the three soils.</caption><thead><tr><th></th><th>BOU</th><th>PACA</th><th>PDL</th></tr></thead><tbody><tr><td>Water-holding capacity (%)</td><td>33</td><td>23</td><td>26</td></tr><tr><td>Clay (g.kg<sup>-1</sup> DM)</td><td>386</td><td>102</td><td>115</td></tr><tr><td>Silt (g.kg<sup>-1</sup> DM)</td><td>530</td><td>232</td><td>392</td></tr><tr><td>Sand (g.kg<sup>-1</sup> DM)</td><td>80</td><td>666</td><td>493</td></tr><tr><td>pH *</td><td>7.96 ± 0.013 a</td><td>6.68 ± 0.017 b</td><td>6.63 ± 0.005 c</td></tr><tr><td>CEC (cmol<sup>+</sup>.kg<sup>-1</sup> DM) *</td><td>25.44 ± 0.192 a</td><td>7.10 ± 0.477 b</td><td>6.94 ± 0.527 b</td></tr><tr><td>TOC (g.kg<sup>-1</sup> DM) *</td><td>19.81 ± 0.360 a</td><td>7.68 ± 0.748 c</td><td>9.65 ± 0.239 b</td></tr><tr><td>Total N (g.kg<sup>-1</sup> DM) *</td><td>1.74 ± 0.011 a</td><td>0.77 ± 0.043 c</td><td>0.99 ± 0.019 b</td></tr><tr><td>NH<sub>4</sub><sup>+</sup> (mg.kg<sup>-1</sup> DM) †</td><td>1.01 ± 0.118 a</td><td>1.15 ± 0.220 a</td><td>1.54 ± 0.564 a</td></tr><tr><td>NO<sub>3</sub><sup>-</sup> (mg.kg<sup>-1</sup> DM) *</td><td>5.58 ± 0.193 b</td><td>9.53 ± 2.408 a</td><td>2.90 ± 0.359 b</td></tr><tr><td>C/N ratio *</td><td>11.4 ± 0.236 a</td><td>10.0 ± 0.503 b</td><td>9.7 ± 0.087 b</td></tr><tr><td>P<sub>2</sub>O<sub>5</sub> (g.kg<sup>-1</sup> DM) *</td><td>0.070 ± 0.001 a</td><td>0.014 ± 0.002 c</td><td>0.040 ± 0.001 b</td></tr><tr><td>K<sub>2</sub>O (g.kg<sup>-1</sup> DM) *</td><td>0.17 ± 0.002 b</td><td>0.22 ± 0.478 a</td><td>0.18 ± 0.008 b</td></tr><tr><td>CaO (g.kg<sup>-1</sup> DM) *</td><td>6.86 ± 0.032 a</td><td>1.41 ± 0.053 c</td><td>1.73 ± 0.025 b</td></tr><tr><td>MgO (g.kg<sup>-1</sup> DM) *</td><td>0.12 ± 0.002 b</td><td>0.29 ± 0.018 a</td><td>0.09 ± 0.002 c</td></tr><tr><td>Na<sub>2</sub>O (g.kg<sup>-1</sup> DM) *</td><td>0.011 ± 0.001 b</td><td>0.021 ± 0.002 a</td><td>0.010 ± 0.001 b</td></tr></tbody></table>

*Significant at the 0.001 probability level; †Values with the same lower-case letters in a row are not significantly different at P < 0.05.

Abbreviations: BOU, Bourgogne; PACA, Provence-Alpes-Côte d'Azur; PDL, Pays de la Loire; DM, dry matter; CEC, cation exchange capacity; TOC, total organic carbon; NH<sub>4</sub><sup>+</sup>, ammonium nitrogen; NO<sub>3</sub><sup>-</sup>, nitric nitrogen; TOC, total organic carbon; C/N, carbon/nitrogen ratio; pH, potential of hydrogen; P<sub>2</sub>O<sub>5</sub>, phosphorus pentoxide; K<sub>2</sub>O, potassium oxide; CaO, calcium oxide; MgO, magnesium oxide, Na<sub>2</sub>O, sodium oxide.
<table><caption>**Table 2.** Elements brought with the digestates and the manure, expressed as the amount loaded in each microcosm.</caption><thead><tr><th rowspan="2">Organic amendments</th><th>Total N</th><th>Organic N</th><th>NH<sub>4</sub><sup>+</sup></th><th>NO<sub>3</sub><sup>-</sup></th><th>TOC</th><th>C/N</th><th>pH</th><th>P<sub>2</sub>O<sub>5</sub></th><th>K<sub>2</sub>O</th><th>CaO</th><th>MgO</th><th>Na<sub>2</sub>O</th><th>SOC</th><th>Hemicellulosis</th><th>Cellulosis</th><th>Lignin cutin</th></tr><tr><th>mg</th><th></th><th></th><th></th><th>µg</th><th>mg</th><th>-</th><th>-</th><th>mg</th><th></th><th></th><th></th><th></th><th></th><th></th><th></th><th></th></tr></thead><tbody><tr><td>FYM</td><td>3.75</td><td>2.39</td><td>1.33</td><td>2.75</td><td>26.60</td><td>6.30</td><td>8.80</td><td>1.84</td><td>5.59</td><td>2.46</td><td>0.89</td><td>1.02</td><td>10.29</td><td>4.57</td><td>6.54</td><td>5.16</td></tr><tr><td>MFW</td><td>4.23</td><td>2.30</td><td>1.92</td><td>2.70</td><td>14.32</td><td>2.80</td><td>8.30</td><td>1.09</td><td>4.84</td><td>2.11</td><td>0.34</td><td>0.82</td><td>9.62</td><td>1.76</td><td>1.27</td><td>1.65</td></tr><tr><td>SMS</td><td>2.52</td><td>1.70</td><td>0.81</td><td>2.73</td><td>13.30</td><td>4.50</td><td>8.00</td><td>0.82</td><td>3.27</td><td>1.02</td><td>0.34</td><td>0.14</td><td>8.07</td><td>2.25</td><td>1.25</td><td>1.73</td></tr><tr><td>CMF</td><td>4.64</td><td>1.43</td><td>3.11</td><td>2.70</td><td>3.82</td><td>0.80</td><td>8.40</td><td>0.34</td><td>1.43</td><td>0.20</td><td>0.07</td><td>1.09</td><td>3.81</td><td>0.00</td><td>0.00</td><td>0.01</td></tr><tr><td>CM</td><td>2.66</td><td>2.26</td><td>0.38</td><td>2.76</td><td>42.97</td><td>18.30</td><td>8.90</td><td>1.16</td><td>3.27</td><td>1.57</td><td>0.68</td><td>0.27</td><td>9.88</td><td>11.51</td><td>14.22</td><td>7.35</td></tr></tbody></table>

Weights are expressed as the amount loaded in each microcosm.

Abbreviations: NH<sub>4</sub><sup>+</sup>, ammonium nitrogen; NO<sub>3</sub><sup>-</sup>, nitric nitrogen; TOC, total organic carbon; C/N, carbon/nitrogen ratio; pH, potential of hydrogen; P<sub>2</sub>O<sub>5</sub>, phosphorus pentoxide; K<sub>2</sub>O, potassium oxide; CaO, calcium oxide; MgO, magnesium oxide; Na<sub>2</sub>O, sodium oxide; SOC, soluble organic carbon; FYM, farmyard manure; MFW, manure + food waste; SMS, silage maize and sorghum; CMF, cattle manure + fat; CM, cattle manure.
<figure><img src="image_36.png" /></figure>

Different soils with distinct microbial community structures, responded differently to different digestates application. Response also digestate-dependent