Next Article in Journal
From Lab to Field: Biofertilizers in the 21st Century
Previous Article in Journal
Optimized Fertilization Shifted Soil Microbial Properties and Improved Vegetable Growth in Facility Soils with Obstacles
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Horticulturae
Volume 9
Issue 12
10.3390/horticulturae9121304
horticulturae-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Composts Obtained by Mixing Hop Leaves with Wheat Straw or Farmyard Manure Improved Soil Properties and Increased Microbial Communities

by
Sandra Afonso
1,2,
Ermelinda L. Pereira
3,4,
Margarida Arrobas
3,4,
M. Ângelo Rodrigues
3,4 and
Altino Choupina
3,4,*
1
CFE-Centre for Functional Ecology—Science for the People and the Planet, Department of Life Sciences, University of Coimbra, 3000-456 Coimbra, Portugal
2
Associate Laboratory TERRA, University of Coimbra, 3000-456 Coimbra, Portugal
3
Centro de Investigação de Montanha (CIMO), Instituto Politécnico de Bragança, Campus de Santa Apolónia, 5300-253 Bragança, Portugal
4
Laboratório Associado para a Sustentabilidade e Tecnologia em Regiões de Montanha (SusTEC), Instituto Politécnico de Bragança, Campus de Santa Apolónia, 5300-253 Bragança, Portugal
*
Author to whom correspondence should be addressed.
Horticulturae 2023, 9(12), 1304; https://doi.org/10.3390/horticulturae9121304
Submission received: 10 November 2023 / Revised: 29 November 2023 / Accepted: 30 November 2023 / Published: 5 December 2023
Browse Figures
Review Reports Versions Notes

Abstract

:
Hop (Humulus lupulus L.) leaves are rich in nutrients, particularly nitrogen (N). After harvest, they can be recycled through composting for use as a soil amendment. In this study, we report the effect of composts obtained from mixtures of hop leaves with other organic materials (wheat straw, farmyard manure, and ash from hop stems) at different ratios on soil properties and microbial diversity. Data on total N, total organic carbon (TOC), microbial N (Mic-N), microbial C (Mic-C), soil basal respiration (SBR), metabolic quotient (qCO2), Mic-C/TOC ratio, acid phosphatase activity (APA), microbial density, and species identification were assessed after each one of the two growing seasons of potted lettuce (Lactuca sativa L.). The diversity of microbial species was evaluated using Simpson and Shannon diversity indexes, and the interactions between soil properties and the microbial community were explored. Higher microbial activity was found among the soils amended with leaves plus straw (HS), which exhibited higher levels of TOC, APA, Mic-N, and total N in the first growing cycle and higher levels of Mic-C, Mic-C/TOC, SBR, TOC, and Mic-N in the second growing cycle. Fungi identified belong to the Ascomycota and Zygomycota phyla, while bacteria belong to the Actinobacteria, Bacillota, Bacteroidetes, Firmicutes, and Proteobacteria phyla. Differences in the prevalent microbial genera were observed between compost treatments and growing cycles. Correlation analysis revealed significant relationship between soil bacteria and fungi abundance and higher levels of N and C in the soils, indicating the relevance of specific microbial genera, such Acrostalagmus, Doratomyces, Talaromyces, and Aspergillus fungi, as well as Gordonia and Bacillus bacteria. Overall, the results indicate that hop leaves-based compost, particularly with a higher proportion of leaves and straw, influenced the composition of the soil microbial community, ultimately enhancing soil N availability for plant development.

Graphical Abstract

1. Introduction

The reconversion and valorisation of agricultural waste present significant challenges at the present time. In the case of hops, a crop that is destined mostly for the brewing industry [1,2], a considerable quantity of organic residues is produced every year all over the world. The main world producers are Germany and the United States, though commercial production is spread over many countries [3,4]. Recently, the craft brewing movement expanded the interest in hop production to new emerging areas of production such as Italy and Brazil [5,6,7]. The increasing interest in hop production also increased the interest in valuing the waste resulting from hop harvest [8,9] and from the brewing industry [10,11].
Composting is a process that allows the management of agricultural waste in a sustainable way, decreasing the negative environmental impact. The organic waste is transformed into composts that can be used as soil amendments [12,13]. The benefits of the application of these organic materials on soil quality result from the increase in soil available nutrients, soil organic matter, and beneficial microorganisms [14,15].
Soil biochemical and biological parameters such as microbial biomass C and N, soil respiration, enzyme activity, and mineralizable N are widely used to assess the effects of composts on soil properties. These parameters have the sensitivity to reflect slight modifications that soil can undergo after the application of an organic substrate [16,17,18]. Organic amendments are usually rich in organic C and N, which promote microbial activity and growth [19,20,21]. Microbial biomass represents the living component of soil organic matter, while microbial respiration reflects the basic turnover rates, and both respond rapidly to changes in soil management [16,21]. The decomposition of complex organic compounds through microorganisms requires the activity of enzymes, which will be reflected in the mineralization rates [18,22]. Enzyme phosphatases, for instance, are involved in the process of mineralizing of phosphate, increasing the availability of phosphorus (P), an important nutrient for soil microbes’ growth, which consequently influences C decomposition and cycling [23,24]. The mineralization of N is also carried out through microorganisms, and the rate at which this occurs depends on the content of C and N in organic residues [24,25].
The application of organic substrates to the soil not only influences microbial activity and growth but can also lead to changes in microbial diversity and in the dominance of functional groups. Soil properties such as nutrient content, pH, total N, organic C, and C/N ratio are major factors affecting soil microbial community composition [26,27,28,29,30]. These changes will in turn be reflected in the quality of the soil [31,32,33]. Pospíšilová et al. [34] showed that the application of organic amendments resulted in chemical and structural changes in the humic substances of soil organic matter.
Hop waste resulting from hop harvest is mostly leaves and stems since the flowers (cones) are the most valuable commercial part of the plant. The hop plants are able to grow nearly 7 m high, producing a large quantity of biomass. In the hop fields of the north-eastern of Portugal, Afonso et al. [35] measured an average dry matter yield of 14.4 t ha−1. In addition, this considerable amount of biomass is very rich in nutrients, particularly N [35]. Therefore, the composts obtained from hop leaves may have adequate levels of nutrients and organic C to improve soil quality. For the present study, we hypothesised that hop leaves-based composts may improve soil properties and enhance soil microbial biomass and its diversity. Thus, the main purpose of the study was to evaluate the effect of composts obtained through suitable mixtures of hop leaves with other local organic resources (wheat straw, farmyard manure, and ash from hop stems) on soil properties and microbial composition from a pot experiment with lettuce carried out during two growing seasons. The relationships between soil properties and the microbial community among the different compost treatments were also explored. The results of the agronomic behaviour of lettuce were already reported in a previous paper [8].

2. Materials and Methods

2.1. Experimental Trials

The composts were obtained after 9 months of composting of hop leaves mixed with other raw materials (wheat straw, farmyard manure, and ash from hop stems) at different ratios (Table 1). The composting process included seven domestic composters that were prepared with mixtures of hop leaves and cow manure and hop leaves and wheat straw at different ratios. Additionally, one of the compost mixtures included hop stem ash. The hop leaves used as raw materials originated from the hop harvesting process and resulted from the separation of cones. The ash also originated from the hop harvest, resulting from the burning of hop stems, a common practice among hop farmers. The cow manure and wheat straw were obtained from local farmers and employed without further processing. The raw materials were introduced into the composters in intercalated layers at predetermined rates to achieve different C/N ratios (Table 1). The process was regularly monitored, including the physical turning of piles, temperature recording, and watering when necessary to ensure adequate moisture. The final composts obtained were used in a pot experiment with lettuce, cv. Wonder of Summer, carried out outdoors under a tile roof during two growing cycles (June–August 2018; April–July 2019). The experiment included nine fertilised treatments, an untreated control (C), and four replicates per treatment. Each pot (~3 L) was filled with 3 kg of dry and sieved soil (2 mm), sampled from the 0–0.20 m layer of a fallow plot. The initial soil had a sandy clay loam texture (soil separates, 23.9% clay, 21.8% silt, and 54.3% sand), a pH (H2O) of 6.51, organic C (Walkley–Black method) of 5.2 g kg−1, total N (Kjeldahl) of 1.28 g kg−1 and extractable P and K (Egner–Riehm method) of 47.8 mg P2O5 kg−1 and 53.3 mg K2O kg−1. The composts were previously mixed with the soil at rates of 20 t ha−1 (dry weight basis) (D1) and 40 t ha−1 (D2), considering that the dry mass of the <2 mm soil fraction of the arable layer (0.2 m) is 2240 t ha−1. The final compost samples were collected in triplicate and subjected to analysis to determine total organic C (Walkley–Black method) and N (Kjeldahl method) for the calculation of the C/N ratio. All the composts were applied at the single (D1) rate, and the composts with the highest and lowest C/N ratios were also applied at the double rate (D2) (Table 1). The pots were watered regularly to maintain optimal moisture levels for plant growth. The amount of water applied varied over time and between pots, since evapotranspiration and the need for watering depend on the development stage of the plants. In the second cycle, the soils were maintained within each pot and were the same soils used in the first cycle, with no additional compost added.

2.2. Soil Analysis

Soil samples were collected in each pot after the lettuce harvest in the two growing seasons and were split into two subsamples. One subsample was frozen in fresh and used for microbial analysis. The other subsample was oven-dried at 40 °C (Memmert UFE 800-Germany ) and used for the determination of total organic C (TOC), total N, and acid phosphatase activity (APA). TOC was determined by dry combustion (Thermolyne 6000 furnace- Germany) and total N by the Kjeldahl method (FOSS KjeltecTM 8400). APA was determined by measuring p-nitrophenol released from 1 g of soil after incubation with 0.2 mL toluene, 4 mL MVB (Modified Universal Buffer) adjusted to pH 6.5, and 1-mL p-nitrophenylphosphate, at 37 °C for 1 h [36] (T80 + UV/VIS spectrometer, PG Instruments- Leicestershire, UK).
Microbial analyses were performed in three replicates for each treatment. Soil microbial biomass C (Mic-C) and N (Mic-N) were determined using the chloroform fumigation–extraction method [37]. The soil samples, adjusted to 60% water holding capacity (WHC), were weighed (20 g) in bottles, fumigated directly with 1 mL of ethanol-free chloroform, and incubated in the dark for 24 h at 25 °C. The extraction from fumigate and non-fumigated samples was performed with 0.5 M K2SO4 (1:4, soil/solution). The extracts were frozen until further analysis. The Mic-C and Mic-N were calculated using a KEC (non-extractable biomass N fraction) factor of 0.33 [38] and a KEN (extractable biomass N fraction) factor of 0.54 [39], respectively. All results are expressed on a dry weight (105 °C) basis. Soil basal respiration (SBR) was measured, according to Pereira et al. [40], in a 7-day laboratory incubation. Briefly, a moist sample (25 g) adjusted to 60% WHC was placed in a 1.5 L glass jar with a plastic vial containing 20 mL of NaOH (1M) solution. After incubation at 25 °C for 1, 3, and 7 days, BaCl2 (1.5 M) solution was added to a plastic vial and then titrated with 2M HCl until the colour changed from pinkie to transparent. Four jars without soil served as controls. SBR was calculated as the accumulative CO2-C released from the soil during 7 days’ laboratory incubation. The metabolic quotient (qCO2) was calculated as the ratio of respiration (mg C-CO2 mg−1 d−1) to Mic-C. The reagents currently in use in the above-mentioned analyses are PanReac p.a.
The densities of cultivable bacteria and fungi were estimated in soil samples by using a standard dilution-plating procedure on agar media. The bacteria were quantified on plate count agar (PCA, Liofilchem, Roseto degli Abruzzi Italy) supplemented with cycloheximide (100 mg L−1; Sigma-Aldrich, Milan, Italy) to suppress fungal growth and incubated at 30 °C for 3 days. Fungi were quantified using rose bengal agar medium (RBA, Liofilchem, Italy), and the plates were incubated at 25 °C for 5 days. Actinomycetes were quantified using DifcoTM Actinomycete isolation agar medium, supplemented with cycloheximide (100 μg mL−1) and nalidixic acid (10 μg mL−1), and incubated at 30 °C for 4 to 10 days. After incubation, the colonies were counted as colony-forming units (cfu). Results were expressed as log cfu g−1 of dry soil. Selected colonies were then inoculated onto yeast malt extract agar (MEA, Liofilchem, Italy) in the case of fungi and plate count agar (PCA, Liofilchem, Italy) in the case of bacteria, for purification, and stored in 30% glycerol at −18 °C.

2.3. DNA Extraction, PCR Amplification, and Sequences Analysis

Total genomic DNA from bacteria was extracted using the REDExtract-N-AmpTM Blood PCR Kit (Sigma-Aldrich), following the manufacturer’s instructions. Total genomic DNA from fungi was extracted according to Martins et al. [41]. The extractions were performed on the soil samples collected after the harvest of lettuce from the first and second growing cycles.
PCR amplification of the 16S rDNA gene was performed using the primer pairs 338F (5′-ACTCCTACGGGAGGCAGCA-3′) and 518R (5′-ATTACCGCGGCTGCTGG-3′) for the V3 region [42], and the primer pairs 338F and 797R (5′-GCGTGGACTACCAGGGTATCTAATCC-3′) for the V3-V4 regions of the gene [43]. PCR amplification of the 18S rDNA for the eukaryota ITS1 region was performed using the primer pairs ITS5F (5′-GGAAGTAAAAGTCGTAACAAGG-3′) and ITS1R (5′-GCTGCGTTCTTCATCGATGC-3′) [26]. The PCR mixture was prepared in a 50 μL volume containing 100 ng of DNA extract, 0.4 μM of each primer, 0.2 mM of each dNTP, 2 U of Taq polymerase, 5X PCR buffer (Promega), 3 mM of MgCl2, 400 μg of bovine serum albumin, and adjusted to the final volume with sterile deionized water. The PCR amplification of 16S rDNA was performed as follows: initial denaturation step of 3 min at 98 °C, followed by 30 cycles (95 °C for 30 s, 57 °C for 45 s, and 72 °C for 50 s), with a final elongation step of 8 min at 72 °C, for primers 338F-518R; initial denaturation step of 2 min at 95 °C, followed by 30 cycles (95 °C for 30 s, 60 °C for 30 s, and 72 °C for 50 s), with a final elongation step of 5 min at 72 °C, for primers 338F-797R. The 18S rDNA PCR programme was carried out with an initial denaturation step at 98 °C for 2 min, followed by 30 cycles (98 °C for 15 s, 55 °C for 30 s, and 72 °C for 30 s), and a final elongation step at 72 °C for 5 min. Amplified PCR products were checked by gel electrophoresis on a 1% agarose gel. PCR amplifications were first extracted from a 0.8% w/v agarose gel and then purified with the Geneclean II Kit (MP Biomedicals). The DNA was quantified by a nano-spectrophotometer (mySPEC Twin 732-2535, VWR, Darmstadt, Germany). The DNA was sequenced with the Sanger method using an automatic sequencer, ABI Prism 377TM (Salamanca University, Salamanca, Spain), that performed the electrophoretic separation and detection of the fluorescence-labelled DNA fragments. Four different fluorescence colours identify the four didexoxynucleotides incorporated in the extension reaction (ATCG).

2.4. Data Analysis

Data were subject to analysis of variance to identify significant differences between treatments, according to the experimental design, with a one-way ANOVA using the SPSS v. 25.0 program. When significant differences were found among experimental treatments, the means were separated by the Tukey–Kramer HSD test (α = 0.05). The analysis of DNA sequences was carried out through the BioEdit programme, and the sequence identity was determined by a BLAST and FASTA nucleotide search from the NCBI and EMBL databases, respectively. It was chosen the closest match sequence, considering the identification percent, score bits, and expected value. The diversity indexes, Simpson and Shannon, were determined using PAST (paleontological statistics software package) version 2 [44], and the data were subjected to analysis of variance according to the procedure described above.

3. Results

3.1. Soil Chemical and Microbiological Evaluation

After the first lettuce cycle, it was found that the composts prepared with a higher proportion of leaves increased the total N and TOC in the soil, particularly the HS2:1D2 treatment, which corresponds to a double rate of leaves (Table 2). The mixture of leaves plus straw tended to increase Mic-N compared to the mixture of leaves plus manure, particularly when a high proportion of leaves (HS2:1D1) was used, which was also reflected in the treatment applied at a double rate (HS2:1D2). Mic-C was particularly high in the treatment in which an equivalent amount of leaves and straw (HS1:1D1) was used and was much lower in all the other treatments. SBR did not vary significantly between treatments. The metabolic coefficient showed very high values in the control treatment and very low values in the HS1:1D1 treatment. The opposite was observed with the Mic-C/TOC ratio. The highest average value of APA was observed in the HS2:1D2 treatment, which was significantly higher than those of the other treatments, except for that of the HS1:2D2 treatment. After the second plant cycle, all values related to the microbial activity of the soil maintained, in some way, the trends between treatments observed in the previous cycle, but the differences faded in a general way (Table 2). SBR was the variable that appeared out of the standard. In the first cycle, there were no significant differences between treatments, while in the second cycle, treatments with a double rate of compost (HS1:2D2 and HS2:1D2) showed significantly higher values than the other treatments.

3.2. Microbial Community Composition and Species Identification

3.2.1. Effect of Compost Treatments on Soil Microbial Composition

The numbers of culturable heterotrophic bacteria differed significantly between compost treatments in the soils collected after the first growing cycle, while significant differences were not found for the culturable actinomycetes population and fungi. The average soil fungi population was higher in the treatments with leaves plus straw applied at the double rate (HS1:2D2 and HS2:1D2) and in the treatment with the same proportion of leaves plus straw (HS1:1D1). The lowest average value was found in the control treatment. The average soil bacteria population was higher in the treatment with a higher leaves proportion plus straw and a single rate of application (HS2:1D1). The average soil actinomycetes population did not show significant differences between treatments. The average population of fungi and bacteria in the soils collected after the second growing cycle was higher in the treatment with a double rate of leaves and a double rate of application (HS2:1D2). The average soil actinomycetes population was higher in the treatment with leaves plus manure with a higher proportion of manure (HM1:5D1). The lower average levels for the fungi population were found in the treatments with leaves plus manure and the control treatment. The lower average levels of bacteria soil population were registered in the control and HM treatments, which were also significantly lower than the HS2:1D2 treatment. The differences between treatments were not significant for the actinomycetes soil population. The proportion of actinomycetes in relation to total bacteria was the lowest in the treatments with leaves plus straw applied at the double rate of application (HS1:2D2 and HS2:1D2) in the soils collected after the first cycle. In the soils collected after the second cycle, the proportion of actinomycetes in relation to total bacteria was significantly lower in the HS2:1D2 treatment in comparison with the HM1:1D1 and control treatments (Table 3).

3.2.2. Correlation Analysis between Microbial Population and Soil Properties

The relation between microbial population and soil properties was analysed through Spearman’s correlation coefficient. Positive and significant correlations were identified for the bacteria and actinomycetes populations in the soils collected after the first growing cycle (2018). The abundance of bacteria and actinomycetes was positive and significantly correlated with the levels of total N (Table 4). Correlation results were more expressive in relation to the soils collected after the second growing cycle (2019). The abundance of fungi and bacteria was positively and significantly correlated with total N, TOC, Mic-N, and Mic-C. Significant correlations were not found for the actinomycetes population.

3.2.3. Microbial Species Identification

A total of 10 fungi colonies and 23 bacteria colonies were identified from the isolates obtained from the soil samples collected after the harvest of lettuce in the first and second growing cycles. Between the fungi colonies isolated, the species identified were found to belong to two phyla, namely Ascomycota and Zygomycota. Between the bacteria colonies isolated, the species identified were found to belong to five different phyla, namely Actinobacteria, Bacillota, Bacteroidetes, Firmicutes, and Proteobacteria (Figure 1).
In the soil samples collected after the first growing cycle, among the fungi communities, Ascomycota was the predominant phyla in all treatments. The genus Aspergillus and Mortierella, respectively, from the Ascomycota and Zygomycota phylum’s, were the most abundant in the control and HM treatments. The dominance among genera varied depending on the treatment. In this regard, Aspergillus was prevalent in the control treatment, exhibiting the highest average values among treatments (38.46%), while Mortierella had the highest rate of abundance in the HM1:3D1 treatment (26.03%). The predominant genera in the treatments with leaves plus straw (HS) were Acrostalagmus, Aspergillus, Doratomyces, and Mortierella. Among these groups, the higher average values were associated with the treatments applied at a double rate, particularly for Doratomyces (45.62%) and Acrostalagmus (31.86%), respectively, in the treatments with a higher proportion of straw (HS1:2D2) and a higher proportion of leaves (HS2:1D2). Among bacterial communities found in the soil samples collected after the first growing cycle, Actinobacteria and Firmicutes were the predominant phyla in all treatments (Figure 1). For most treatments, both phyla shared the dominance with variations, though Firmicutes clearly dominated in HS treatments applied at a double rate. Between these phyla, the main genera were Arthrobacter and Bacillus, respectively, from Actinobacteria and Firmicutes. The higher average values found for Arthrobacter were recorded in the control treatment (58.87%) and the lower in the treatment with a higher leaves proportion plus straw at a double rate of application (HS2:1D2; 25.32%). Conversely, the higher average value found for Bacillus was recorded in the HS2:1D2 treatment (62.25%) and the lower in the control treatment (33.59%).
In soil samples collected after the second growing cycle, among the fungi communities, Ascomycota continued to be the predominant phylum in all treatments, but no clear dominance was found between the different genera of this phylum. The higher average values were found for the genus Talaromyces (41.18%) and Mortierella (30.41%), respectively, in the treatment with a higher proportion of straw (HS1:2D1) and in the treatment with a higher proportion of leaves (HS2:1D1), both at a single rate of application. Between bacteria communities, Firmicutes continued to be the dominant phyla on the soil samples collected after the second growing cycle in all treatments. The genus Bacillus belonging to the Firmicutes phylum was the dominant genus in all treatments, and markedly in the treatments with leaves plus straw (HS). The higher average value for Bacillus (88.56%) was recorded in the treatment with a higher proportion of leaves plus straw at a double rate of application (HS2:1D2), and the lower value was recorded in the control treatment (45.53%), highlighting the trend found in the first growing cycle.
Between the fungi species identified, six species belong to the Ascomycota phylum (Acrostalagmus luteoalbus, Aspergillus flavus, Aspergillus fumigatus, Cephalotrichum asperulum, Penicillium restrictum, and Talaromyces pinophilus), and two species belong to the Zygomycota phylum (Mortierella alpina and Rhizopus arrhizus). The majority of the bacterial species identified belong to the Actinobacteria phylum (six species) and Firmicutes phylum (nine species). Streptomyces and Bacillus were the predominant species identified, respectively, in the Actinobacteria and Firmicutes phylums. The other species identified were Neobacillus endophyticus (Bacillota phylum), Chryseobacterium sp. (Bacteroidetes phylum), Pseudoxanthomonas sp., and Variovorax guangxiensis (Proteobacteria phylum).

3.2.4. Correlation Analysis between Microbial Taxonomic Distribution and Soil Nitrogen and Organic Carbon Content

The abundance of fungi and bacteria genus, soil total N, and TOC were analysed by Spearmen’s correlation coefficients to determine their relationships (Table 5). Positive and significant correlations were identified for some fungi and bacteria taxa in the soils collected after the first growing cycle (2018). The abundance of Acrostalagmus and Doratomyces Ascomycota fungi was significantly and positively correlated with soil TOC (r = 0.544 and 0.547, respectively; p < 0.01). Significant and positive correlations were also found between the abundance of Acrostalagmus and total N (r = 0.390; p < 0.05). Between bacteria communities, the abundance of Gordonia Actinobacteria was also significantly and positively correlated with soil TOC (r = 0.627; p < 0.01). Positive and significant correlations were also found for different fungi and bacteria taxa in relation to the soils collected after the second growing cycle (2019). The abundance of Aspergillus and Talaromyces fungi and of Streptomyces and Bacillus bacteria was shown to be positively and significantly correlated with both soil indicators, total N and TOC, at p < 0.05 and p < 0.01.

3.2.5. Diversity Indices of Soil Fungi and Bacteria Communities

The diversity of fungi and bacteria among the compost treatments was estimated by Simpson and Shannon indexes, which indicated, respectively, the evenness and richness of the communities. As shown in Figure 2, significant differences were not found among fungi and bacterial communities in the soils collected after the first growing cycle and after the second growing cycle. However, a more consistent trend was found in the treatments with leaves plus straw at the double rate of application (HS1:2D2 and HS2:1D2). Overall, these treatments presented the highest and lowest average index values and in the opposite direction when comparing fungi and bacteria results. For instance, in the soils collected after the first growing cycle, the higher Simpson’s index average registered between bacteria communities (0.783), indicating lower species evenness, was found in the treatment with the higher proportion of straw and at the double rate of application (HS1:2D2). Conversely, the lower Simpson’s index average registered between fungi communities (0.697), indicating higher species evenness, was found in the same treatment. Similarly, in the soils collected after the second growing cycle, the higher and lower average values of both indexes were found in the treatment with a higher proportion of leaves plus straw and at the double rate of application (HS2:1D2). The average values recorded in that treatment indicated lower evenness (Simpson index, 0.814) and higher richness (Shannon index, 1.82) of species between the fungal communities, whereas higher evenness and lower species richness were observed among the bacterial communities (Simpson index, 0.473; Shannon index, 1.0.6).

4. Discussion

The effect of composts on soil properties differed between treatments and more clearly between growing cycles. In the soils collected after the first growing cycle, the lower levels of TOC and Mic-N in the treatments with leaves plus manure (HM) indicate a higher stabilisation of OM due to the high maturity of HM compost. On the contrary, some of the treatments with leaves plus straw (HS), in particular with high straw proportions, presented high levels of Mic-C and Mic-C/TOC, which suggests a high availability of C for microbial biomass production [45]. The microbial biomass C is a soil quality indicator, and the increase in soils due to organic manure application has been reported in other studies [46,47,48]. The treatment leaves plus straw with a higher leaves proportion at the double rate of application (HS2:1D2) differentiated from all due to the high levels of TOC, APA, Mic-N, and total N, suggesting an intense activity of microorganisms in the degradation of OM. Thus, the higher availability of organic C likely stimulates phosphatase enzyme activity (APA) due to P demand by microorganisms [18]. At the same time, the degradation of OM by microorganisms resulted in high mineralization and immobilisation of N by microbe biomass, increasing N availability (total N and Mic-N) for plant uptake.
In the soils collected after the second growing cycle, the treatments with the lower and higher straw proportions at the double rate of application, respectively, HS2:1D2 and HS1:2D2, diverge from all due to the high levels of microbiologic activity (Mic-C, Mic-C/TOC, SBR, TOC, Mic-N) and higher availability of total N. As previously discussed, the higher levels of OM, particularly easily degradable OM (since in the second year the composts in soil were more degraded), increased microorganisms’ activity, as reported in other studies [46,49]. On the opposite, the high levels of qCO2 in the control treatment in both growing cycles, along with the lower levels of Mic-C/TOC, indicate a high maintenance energy requirement and a low efficiency in the use of energy for microbial growth [45]. Probably this was due to the lower levels of OM and thus lower organic C availability, resulting in less microbial N and C biomass. In the comparison between the treatments with ash (HSA1:1:0.04D1) and without (HS1:1D1), the addition of ash resulted in lower C availability for microbial growth (lower levels of Mic-C and Mic-C/TOC) and higher availability of N (higher levels of Mic-N and total N), suggesting an enhanced effect on the mineralization rate. It is known that under laboratory conditions, the growth of ammonia-oxidising microorganisms is inhibited at pH values less than 6.5 [50]. The buffering effect of ash may have increased nitrifying microorganisms’ growth, thus increasing the nitrification rate.
Concerning the effect of composts on soil microbial composition, in the soils collected after the first growing cycle, no significant differences between treatments were observed for soil fungi and actinomycetes. However, for soil bacteria, the treatments with leaves plus straw (HS2:1D1) exhibited the highest average (7.44 log cfu g−1), while the treatments with leaves plus manure (HM1:3D1) showed the lowest (7.00 log cfu g−1), and the differences were statistically significant. When straw-based composts were incorporated into the soil, they were less mature than manure-based composts, thereby promoting the continuation of the decomposition process [8]. The high prevalence of bacteria in the soil may be associated with the decomposition of leaves and straw from immature composts, as bacteria tend to dominate when large amounts of organic carbohydrates are available [51].
The effect of composts on soil microbial composition was more pronounced in the second growing cycle for bacteria and fungi, revealing significant differences between treatments. However, no significant differences were observed for soil actinomycetes. The HS treatments exhibited a higher abundance of soil fungi and bacteria compared to the HM and control treatments, with the highest average values recorded in the HS2:1D2 treatments, respectively, of 5.00 log cfu g−1 and 7.32 log cfu g−1. Overall, the results indicate that HS treatments, especially those with higher proportions of leaves, fostered a greater abundance of fungi and bacteria. This outcome could be attributed to the higher concentration of nutrients, particularly N in leaves, along with a higher concentration of C in straw, which likely promoted the growth of bacteria and fungi [52,53]. The results of the correlation analysis between microbial populations and soil properties support this hypothesis, as they revealed a significant relationship between the abundance of soil bacteria and soil fungi with higher levels of N and C in the soils. This result aligns with the findings of Afonso et al. [8], suggesting a period of N immobilization in the first growing cycle followed by a period of net mineralization in the second growing cycle. The activity of bacteria and fungi is closely associated with the rates of soil N transformation [53]. The abundance of C and N in the soil strongly influences the growth of bacteria and fungi, driving the response of heterotrophic nitrification [52,53].
The fungi colonies isolated from the soil samples collected after the harvest of lettuce from the first and second growing cycles were identified as belonging to the Ascomycota and Zygomycota phyla. The bacteria colonies included Actinobacteria, Bacillota, Bacteroidetes, Firmicutes, and Proteobacteria phyla. In the soil samples collected after the first growing cycle, the dominant fungus phylum was Ascomycota. Ascomycota are prominent decomposers among fungi and have been identified as major phyla in soils treated with sewage sludge compost [54]. Ma et al. [55] also recognised Ascomycota members as dominant during various stages of fungal succession while decomposing straw residues in arable soil. Voříšková and Baldrian [56] studied fungal communities during litter decomposition and found Ascomycota to be the most abundant during the early stages of this process. These findings align with our results. The genera Acrostalagmus, Aspergillus, and Doratomyces from the Ascomycota phylum, along with Mortierella from the Zygomycota phylum, were the most prevalent in the HS treatments. Within this group, Doratomyces, followed by Acrostalagmus, showed higher abundance in the treatments with leaves plus straw applied at a double rate (HS1:2D2; HS2:1D2). On the other hand, the genus Aspergillus (Ascomycota) was most abundant in the control treatment, while Mortierella (Zygomycota) was most abundant in the HM treatments. Acrostalagmus, Aspergillus, and Mortierella are commonly found in soils [24]. In accordance with our findings, Acrostalagmus and Doratomyces genera were identified during the composting of organic material [57]. Aspergillus is known for its role in phosphate and potassium solubilization in the soil [24,58]. Some Aspergillus species, such as Aspergillus niger, are commonly used in bioleaching for heavy metal removal [59], and can enhance the mineralization of lignocellulose [60,61]. Mortierella, on the other hand, was found to be associated with systems exclusively receiving organic fertilisers such as farmyard manure and slurry [28]. This genus is known to promote plant growth and exhibit high cellulose and hemicellulose degradation activity [62]. Hence, it aligns with the observation that these genera were abundant in the soil samples, particularly in the treatments of leaves plus straw, with Mortierella also being the most abundant in the treatments of leaves plus manure. The correlation analysis revealed that the abundance of Acrostalagmus and Doratomyces fungi correlated with high levels of organic C, and for Acrostalagmus, it was also related to high levels of N, likely indicating their role in straw degradation and N cycling. However, further information is needed to fully understand the specific roles of these genera in the decomposition process.
Regarding bacteria, in the soil samples collected after the first growing cycle, Actinobacteria and Firmicutes were the most abundant phyla across all the treatments. This is consistent with other studies that reported these phyla as abundant in various types of organic waste compost [54,63,64]. Among the different genera, Arthrobacter (Actinobacteria) and Bacillus (Firmicutes) were the most prevalent in all the treatments. These genera are commonly found in soil and are known to colonise the rhizoplane, promoting plant growth [24]. The highest rate of Arthrobacter was found in the control treatment (58.87%), and the lowest was found in the treatment of leaves plus straw, with a higher proportion of leaves at a double rate (HS2:1D2: 25.32%). Conversely, Bacillus abundance was highest in the treatments of leaves plus straw applied at a double rate (HS2:1D2: 62.5% > HS1:2D2: 59.3%) and lowest in the control treatment (33.59%). Arthrobacter bacteria are typically abundant in mature compost [57], which may explain their highest presence in the control treatment, where OM was more stabilized. Regarding Bacillus species, they possess the ability for N fixation [65], P and K solubilisation [58], and the capability to degrade cellulose and starch [66]. Hence, it is reasonable that they were found at higher rates in the treatments of leaves plus straw at a double rate. The correlation analysis indicated that the abundance of the genus Gordonia, which belongs to the phylum Actinobacteria, was related to high levels of organic C. Although this genus had low rates in all treatments, the highest levels were found in the treatments of leaves plus straw at a double rate, particularly with higher levels of straw (HS1:2D2: 0.76%; HS2:1D2: 0.54%). Gordonia species are known to contribute to soil quality improvement and plant growth promotion. They produce several enzymes and are capable of degrading lignin [67], which may explain their higher presence in treatments with higher proportions of straw and leaves.
In the soil samples collected after the second growing cycle, Ascomycota remained the predominant fungus phylum across all treatments. Similarly, Tan et al. [61] found Ascomycota to be the dominant phylum throughout all stages of wheat straw composting. Although there was no clear dominance between fungi genera, higher rates were found for the Talaromyces (41.18%) and Mortierella (30.41%) genera in the treatments of leaves plus straw, with higher proportions of straw (HS1:2D1) and leaves (HS2:D1), respectively. Our findings align with those of Xu et al. [68], who demonstrated that Talaromyces is a crucial fungus in promoting the degradation of lignin and humification during lignocellulosic waste composting. Mortierella, along with Talaromyces, likely played a relevant role in the decomposition of these subtracts, maybe related to the degradation of cellulose or hemicellulose [62]. The correlation analysis revealed a connection between the abundance of Talaromyces and Aspergillus with high levels of organic C and N in the soil. Talaromyces species, including T. pinophilus identified in this study, are frequently employed in phytoremediation and can enhance nutrient availability in soil with organic amendments [69]. Microorganisms can utilise lignin as a source of C, contributing to an increase in organic C availability in the soil [70]. This process seems to be influenced by interactions between lignin decomposition products (microbial biomass and necromass) and soil minerals [71]. As previously stated, Aspergillus can mineralize metals and lignocellulose [59,61]. Therefore, both genera most likely played a role in lignin breakdown and contributed to the increase in soil organic C and N.
Regarding bacteria, in the soil samples collected after the second growing cycle, Firmicutes emerged as the most abundant phylum across all treatments, with Bacillus being the dominant genus, particularly in the treatments with leaves plus straw. These findings may be related to the role of Firmicutes, especially Bacillus, in the degradation of cellulose rich materials such as straw [66,72] as discussed earlier. Moreover, Firmicutes tend to dominate during the late stage of cellulose degradation, operating at high ammonia levels [72]. This observation can explain the positive correlation found between Bacillus, the dominant genus within Firmicutes, and the concentration of organic C and N in the soil. Positive relationships between soil C and N were also observed for Streptomyces, a genus belonging to the Actinobacteria phylum. Actinobacteria, in general, and particularly Streptomyces, are known for their ability to decompose plant litter and more recalcitrant compounds like lignin. They are typically present during the final stages of decomposition [24,66].
The diversity indexes, Simpson and Shannon, which, respectively, estimated the evenness and richness of the fungi and bacteria communities, did not exhibit significant differences for soil microbe communities in either the first or second growing cycle. Conversely, other studies have reported changes in soil microbial communities after compost amendments, leading to improved microbial diversity [32,54]. However, a consistent trend was observed in the treatments of leaves plus straw, at the double rate of application. In the first growing cycle, the treatment with a higher proportion of straw exhibited high fungi species evenness and low bacterial species evenness. The distribution of fungi was less equitable, with a clear dominance of Doratomyces fungi, likely related to straw decomposition at early stages. Among bacteria, dominance was shared between Arthrobacter (Actinobacteria) and Bacillus (Firmicutes). In the second growing cycle, the treatments with a higher proportion of leaves plus straw showed lower evenness and higher richness of fungi, indicating a more balanced distribution among the taxa present. For bacteria, higher evenness and lower richness were observed, likely due to the clear dominance of Bacillus species and their importance in the decomposition of cellulose straw at late stages [66,72]. Furthermore, previous research with lettuce pots and field experiments in soil fertilised with different types of organic compost demonstrates that composts can improve soil chemical, physical, and microbiological properties, ultimately promoting plant growth [73,74,75]. Likewise, the results of our short-term pot experiment also revealed that hop leaves-based composts have the potential to be used in crop fields to enhance soil fertility, soil microbiology, and plant growth while contributing to improved circular management of crop residue. However, further research with long-term field experiments is required to substantiate the preliminary findings of the current study.

5. Conclusions

The present study explored the effect of hop leaves-based compost on soil properties and microbial communities during two growing cycles of lettuce. Compost with leaves plus straw increased microbial abundance compared to leaves plus manure composts and control treatments. This was likely due to the increased N availability from leaves, C from straw, and the presence of degradable OM from immature compost. The treatment with leaves plus straw, especially during the first growing cycle, exhibited intense microbial activity and stimulated phosphatase enzyme activity, leading to substantial mineralisation and immobilisation of N by microbial biomass, ultimately enhancing N availability for plant uptake. The findings indicate that the higher availability of C and N in the soil played a crucial role in driving the growth and activity of bacteria and fungi, as well as influencing the dominance between microbial genera.
The changes in fungi and bacteria genera in response to compost treatments were more pronounced in the treatments with leaves plus straw, particularly during the first growing cycle when the decomposition process was more intense. The relationship found between soil properties and microbial communities revealed that the activity of bacteria and fungi was closely associated with the abundance of N and C in the soil, likely related to straw decomposition and N cycling at different stages of the growing cycles. The microbial genera found in these associations included Acrostalagmus, Doratomyces, Talaromyces, and Aspergillus fungi, as well as Gordonia, Bacillus, and Streptomyces bacteria. While significant effects on microbial diversity due to compost treatments were not observed, the dominance of certain genera, such as Doratomyces fungi in the first growing cycle and Bacillus bacteria in the second growing cycle, suggested their relevance in early-stage straw decomposition and decomposition of cellulose straw at later stages, respectively. However, further research is needed to uncover the specific functions and interactions of these microorganisms in the context of OM transformation.
Overall, our findings contribute to the understanding of compost-induced changes in soil microbial communities and underscore the importance of organic compost to promote soil health and crop productivity, suggesting the potential applicability of hop-leaves based compost in agricultural fields.

Author Contributions

Conceived and designed the experiments: S.A., E.L.P., M.A., M.Â.R. and A.C. Performed the experiments: S.A. Analysed the data: S.A. Wrote the paper: S.A., E.L.P., M.A., M.Â.R. and A.C. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Foundation for Science and Technology (FCT, Portugal) through national funds FCT/MCTES (PIDDAC) to CIMO (UIDB/00690/2020 and UIDP/00690/2020) and SusTEC (LA/P/0007/2020) and through the scholarship grant of Sandra Afonso (number BD/116593/2016) under the Programme PT2020 (UID/AGR/00690/2019).

Data Availability Statement

Data are contained within the article.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Almaguer, C.; Schönberger, C.; Gastl, M.; Arendt, E.K.; Becker, T. Humulus lupulus—A story that begs to be told. A review. J. Inst. Brew. 2014, 120, 289–314. [Google Scholar] [CrossRef]
  2. Teghtmeyer, S. Hops. J. Agric. Food Inf. 2018, 19, 9–20. [Google Scholar] [CrossRef]
  3. Small, E. Hop (Humulus lupulus)—A bitter crop with sweet prospects. Biodiversity 2016, 17, 115–127. [Google Scholar] [CrossRef]
  4. FAOSAT. Production: Crops. Available online: https://www.fao.org/faostat/en/#data/QCL/visualize (accessed on 20 August 2022).
  5. Rossini, F.; Loreti, P.; Provenzano, M.E.; De Santis, D.; Ruggeri, R. Agronomic performance and beer quality assessment of twenty hop cultivars grown in Central Italy. Ital. J. Agron. 2016, 11, 180–187. [Google Scholar] [CrossRef]
  6. Mozzon, M.; Foligni, R.; Mannozzi, C. Brewing Quality of Hop Varieties Cultivated in Central Italy Based on Multivolatile Fingerprinting and Bitter Acid Content. Foods 2020, 9, 541. [Google Scholar] [CrossRef] [PubMed]
  7. da Rosa Almeida, A.; Maciel, M.V.d.O.B.; Cardoso Gasparini Gandolpho, B.; Machado, M.H.; Teixeira, G.L.; Bertoldi, F.C.; Noronha, C.M.; Vitali, L.; Block, J.M.; Barreto, P.L.M. Brazilian Grown Cascade Hop (Humulus lupulus L.): LC-ESI-MS-MS and GC-MS Analysis of Chemical Composition and Antioxidant Activity of Extracts and Essential Oils. J. Am. Soc. Brew. Chem. 2020, 79, 156–166. [Google Scholar] [CrossRef]
  8. Afonso, S.; Arrobas, M.; Pereira, E.L.; Rodrigues, M.Â. Recycling nutrient-rich hop leaves by composting with wheat straw and farmyard manure in suitable mixtures. J. Environ. Manag. 2021, 284, 112105. [Google Scholar] [CrossRef] [PubMed]
  9. Kanai, N.; Nishimura, K.; Umetani, S.; Saito, Y.; Saito, H.; Oyama, T.; Kawamura, I. Upcycling of Waste Hop Stems into Cellulose Nanofibers: Isolation and Structural Characterization. ACS Agric. Sci. Technol. 2021, 1, 347–354. [Google Scholar] [CrossRef]
  10. Kopeć, M.; Mierzwa-Hersztek, M.; Gondek, K.; Wolny-Koładka, K.; Zdaniewicz, M.; Jarosz, R. Biological activity of composts obtained from hop waste generated during the brewing. Biomass Convers. Biorefinery 2020, 12, 1271–1279. [Google Scholar] [CrossRef]
  11. Hrnčič, M.K.; Španinger, E.; Košir, I.J.; Knez, Ž.; Bren, U. Hop compounds: Extraction techniques, chemical analyses, antioxidative, antimicrobial, and anticarcinogenic effects. Nutrients 2019, 11, 257. [Google Scholar] [CrossRef]
  12. Pergola, M.; Persiani, A.; Palese, A.M.; Di Meo, V.; Pastore, V.; D’Adamo, C.; Celano, G. Composting: The way for a sustainable agriculture. Appl. Soil Ecol. 2018, 123, 744–750. [Google Scholar] [CrossRef]
  13. Lal, R. Soils and sustainable agriculture. A review. Agron. Sustain. Dev. 2008, 28, 57–64. [Google Scholar] [CrossRef]
  14. Wong, J.W.C.; Wang, X.; Selvam, A. Improving compost quality by controlling nitrogen loss during composting. In Current Developments in Biotechnology and Bioengineering; Wong, J.W.C., Tyagi, R.D., Pandey, A., Eds.; Elsevier: Amsterdam, The Netherlands, 2017; pp. 59–82. [Google Scholar]
  15. Singh, J.S.; Pandey, V.C.; Singh, D.P. Efficient soil microorganisms: A new dimension for sustainable agriculture and environmental development. Agric. Ecosyst. Environ. 2011, 140, 339–353. [Google Scholar] [CrossRef]
  16. Gil-Sotres, F.; Trasar-Cepeda, C.; Leirós, M.C.; Seoane, S. Different approaches to evaluating soil quality using biochemical properties. Soil Biol. Biochem. 2005, 37, 877–887. [Google Scholar] [CrossRef]
  17. Zagal, E.; Muñoz, C.; Quiroz, M.; Córdova, C. Sensitivity of early indicators for evaluating quality changes in soil organic matter. Geoderma 2009, 151, 191–198. [Google Scholar] [CrossRef]
  18. Yang, L.; Zhang, Y.; Li, F. Soil enzyme activities and soil fertility dynamics. In Advances in Citrus Nutrition; Spring: Berlin/Heidelberg, Germany, 2012; pp. 143–156. [Google Scholar]
  19. Raviv, M. Production of high- quality composts for horticultural purposes: A mini-review. HortTechnology 2005, 15, 52–57. [Google Scholar] [CrossRef]
  20. Gent, D.H.; Barbour, J.D.; Dreves, A.J.; James, D.G.; Parker, R.; Walsh, D.B.; O’Neal, S. Field Guide for Integrated Pest Management in Hops; Oregon State University: Corvallis, OR, USA; University of Idaho: Moscow, ID, USA; USDA-ARS: Washington, DC, USA; Washington State University: Pullman, WA, USA, 2010. [Google Scholar]
  21. Knapp, B.A.; Ros, M.; Insam, H. Do Composts Affect the Soil Microbial Community? In Microbes at Work: From Wastes to Resources; Insam, H., Franke-Whittle, I., Goberna, M., Eds.; Springer: Berlin/Heidelberg, Germany, 2010; pp. 271–291. [Google Scholar]
  22. Caldwell, B.A. Enzyme activities as a component of soil biodiversity: A review. Pedobiologia 2005, 49, 637–644. [Google Scholar] [CrossRef]
  23. Hartman, W.H.; Richardson, C.J. Differential Nutrient Limitation of Soil Microbial Biomass and Metabolic Quotients (qCO2): Is There a Biological Stoichiometry of Soil Microbes? PLoS ONE 2013, 8, e57127. [Google Scholar] [CrossRef]
  24. Osman, K.T. Soils: Principles, Properties and Management, 1st ed.; Springer: Dordrecht, The Netherlands, 2013. [Google Scholar]
  25. Maier, R.M. Biogeochemical Cycling. In Environmental Microbiology, 3rd ed.; Pepper, I.L., Gerba, C.P., Gentry, T.J., Eds.; Academic Press: San Diego, CA, USA, 2015; pp. 339–373. [Google Scholar]
  26. Ma, J.; Nergui, S.; Han, Z.; Huang, G.; Li, H.; Zhang, R.; Zhu, L.; Liao, J. The Variation of the Soil Bacterial and Fungal Community Is Linked to Land Use Types in Northeast China. Sustainability 2019, 11, 3286. [Google Scholar] [CrossRef]
  27. Lauber, C.L.; Strickland, M.S.; Bradford, M.A.; Fierer, N. The influence of soil properties on the structure of bacterial and fungal communities across land-use types. Soil Biol. Biochem. 2008, 40, 2407–2415. [Google Scholar] [CrossRef]
  28. Hartmann, M.; Frey, B.; Mayer, J.; Mäder, P.; Widmer, F. Distinct soil microbial diversity under long-term organic and conventional farming. ISME J. 2015, 9, 1177–1194. [Google Scholar] [CrossRef] [PubMed]
  29. Manoharan, L.; Kushwaha, S.K.; Ahrén, D.; Hedlund, K. Agricultural land use determines functional genetic diversity of soil microbial communities. Soil Biol. Biochem. 2017, 115, 423–432. [Google Scholar] [CrossRef]
  30. Navrátilová, D.; Tláskalová, P.; Kohout, P.; Dřevojan, P.; Fajmon, K.; Chytrý, M.; Baldrian, P. Diversity of fungi and bacteria in species-rich grasslands increases with plant diversity in shoots but not in roots and soil. FEMS Microbiol. Ecol. 2019, 95, fiy208. [Google Scholar] [CrossRef] [PubMed]
  31. Zhong, W.; Gu, T.; Wang, W.; Zhang, B.; Lin, X.; Huang, Q.; Shen, W. The effects of mineral fertilizer and organic manure on soil microbial community and diversity. Plant Soil 2010, 326, 511–522. [Google Scholar] [CrossRef]
  32. Cozzolino, V.; Di Meo, V.; Monda, H.; Spaccini, R.; Piccolo, A. The molecular characteristics of compost affect plant growth, arbuscular mycorrhizal fungi, and soil microbial community composition. Biol. Fertil. Soils 2016, 52, 15–29. [Google Scholar] [CrossRef]
  33. Ding, G.-C.; Bai, M.; Han, H.; Li, H.; Ding, X.; Yang, H.; Xu, T.; Li, J. Microbial taxonomic, nitrogen cycling and phosphorus recycling community composition during long-term organic greenhouse farming. FEMS Microbiol. Ecol. 2019, 95, fiz042. [Google Scholar] [CrossRef] [PubMed]
  34. Pospíšilová, L.; Horáková, E.; Fišera, M.; Jerzykiewicz, M.; Menšík, L. Effect of selected organic materials on soil humic acids chemical properties. Environ. Res. 2020, 187, 109663. [Google Scholar] [CrossRef]
  35. Afonso, S.; Arrobas, M.; Rodrigues, M.Â. Soil and plant analyses to diagnose hop fields irregular growth. J. Soil Sci. Plant Nutr. 2020, 20, 1999–2013. [Google Scholar] [CrossRef]
  36. Eivazi, F.; Tabatabai, M. Phosphatases in soils. Soil Biol. Biochem. 1977, 9, 167–172. [Google Scholar] [CrossRef]
  37. Witt, C.; Gaunt, J.L.; Galicia, C.C.; Ottow, J.C.G.; Neue, H.U. A rapid chloroform-fumigation extraction method for measuring soil microbial biomass carbon and nitrogen in flooded rice soils. Biol. Fertil. Soils 2000, 30, 510–519. [Google Scholar] [CrossRef]
  38. Vance, E.D.; Brookes, P.C.; Jenkinson, D.S. An extraction method for measuring soil microbial biomass C. Soil Biol. Biochem. 1987, 19, 703–707. [Google Scholar] [CrossRef]
  39. Brookes, P.; Kragt, J.; Powlson, D.; Jenkinson, D. Chloroform fumigation and the release of soil nitrogen: The effects of fumigation time and temperature. Soil Biol. Biochem. 1985, 17, 831–835. [Google Scholar] [CrossRef]
  40. Pereira, E.; Santos, S.; Arrobas, M.; Patrício, M.D.S. Microbial biomass and N mineralization in mixed plantations of broadleaves and nitrogen-fixing species. For. Syst. 2011, 20, 516–524. [Google Scholar] [CrossRef]
  41. Martins, I.; Martins, F.; Belo, H.; Vaz, M.; Carvalho, M.; Cravador, A.; Choupina, A. Cloning, characterization and in vitro and in planta expression of a glucanase inhibitor protein (GIP) of Phytophthora cinnamomi. Mol. Biol. Rep. 2014, 41, 2453–2462. [Google Scholar] [CrossRef] [PubMed]
  42. Wu, X.; Wu, F.; Zhou, X.; Fu, X.; Tao, Y.; Xu, W.; Pan, K.; Liu, S. Effects of Intercropping with Potato Onion on the Growth of Tomato and Rhizosphere Alkaline Phosphatase Genes Diversity. Front Plant Sci 2016, 7, 846. [Google Scholar] [CrossRef] [PubMed]
  43. Sune, D.; Rydberg, H.; Augustinsson, Å.N.; Serrander, L.; Jungeström, M.B. Optimization of 16S rRNA gene analysis for use in the diagnostic clinical microbiology service. J. Microbiol. Methods 2020, 170, 105854. [Google Scholar] [CrossRef]
  44. Hammer, O.; Harper, D.; Ryan, P. PAST: Paleontological Statistics Software Package for Education and Data Analysis. Palaeontol. Electron. 2001, 4, 1–9. [Google Scholar]
  45. Anderson, T.-H.; Domsch, K.H. Soil microbial biomass: The eco-physiological approach. Soil Biol. Biochem. 2010, 42, 2039–2043. [Google Scholar] [CrossRef]
  46. Hu, J.L.; Lin, X.G.; Wang, J.H.; Dai, J.; Chen, R.R.; Zhang, J.B.; Wong, M.H. Microbial functional diversity, metabolic quotient, and invertase activity of a sandy loam soil as affected by long-term application of organic amendment and mineral fertilizer. J. Soils Sediments 2011, 11, 271–280. [Google Scholar] [CrossRef]
  47. Jannoura, R.; Joergensen, R.G.; Bruns, C. Organic fertilizer effects on growth, crop yield, and soil microbial biomass indices in sole and intercropped peas and oats under organic farming conditions. Eur. J. Agron. 2014, 52, 259–270. [Google Scholar] [CrossRef]
  48. Tu, C.; Ristaino, J.B.; Hu, S. Soil microbial biomass and activity in organic tomato farming systems: Effects of organic inputs and straw mulching. Soil Biol. Biochem. 2006, 38, 247–255. [Google Scholar] [CrossRef]
  49. Dinesh, R.; Srinivasan, V.; Hamza, S.; Manjusha, A.; Kumar, P.S. Short-term effects of nutrient management regimes on biochemical and microbial properties in soils under rainfed ginger (Zingiber officinale Rosc.). Geoderma 2012, 173–174, 192–198. [Google Scholar] [CrossRef]
  50. Prosser, J.I. Nitrogen in soils: Nitrification. In Encyclopedia of Soils in the Environment; Hillel, D., Ed.; Elsevier: Oxford, UK, 2005; pp. 31–39. [Google Scholar]
  51. Nemet, F.; Perić, K.; Lončarić, Z. Microbiological activities in the composting process—A review. J. Agric. Environ. Sci. 2021, 8, 41–53. [Google Scholar] [CrossRef]
  52. Zhang, Y.; Wang, J.; Dai, S.; Zhao, J.; Huang, X.; Sun, Y.; Chen, J.; Cai, Z.; Zhang, J. The effect of C:N ratio on heterotrophic nitrification in acidic soils. Soil Biol. Biochem. 2019, 137, 107562. [Google Scholar] [CrossRef]
  53. Isobe, K.; Koba, K.; Otsuka, S.; Senoo, K. Nitrification and nitrifying microbial communities in forest soils. J. For. Res. 2011, 16, 351–362. [Google Scholar] [CrossRef]
  54. Ai, Y.-J.; Li, F.-P.; Gu, H.-H.; Chi, X.-J.; Yuan, X.-T.; Han, D.-Y. Combined effects of green manure returning and addition of sewage sludge compost on plant growth and microorganism communities in gold tailings. Environ. Sci. Pollut. Res. 2020, 27, 31686–31698. [Google Scholar] [CrossRef] [PubMed]
  55. Ma, A.; Zhuang, X.; Wu, J.; Cui, M.; Lv, D.; Liu, C.; Zhuang, G. Ascomycota Members Dominate Fungal Communities during Straw Residue Decomposition in Arable Soil. PLoS ONE 2013, 8, e66146. [Google Scholar] [CrossRef]
  56. Voříšková, J.; Baldrian, P. Fungal community on decomposing leaf litter undergoes rapid successional changes. ISME J. 2013, 7, 477–486. [Google Scholar] [CrossRef]
  57. Ryckeboer, J.; Mergaert, J.; Vaes, K.; Klammer, S.; De Clercq, D.; Coosemans, J.; Insam, H.; Swings, J. A survey of bacteria and fungi occurring during composting and self-heating processes. Ann. Microbiol. 2003, 53, 349–410. [Google Scholar]
  58. Bhardwaj, D.; Ansari, M.W.; Sahoo, R.K.; Tuteja, N. Biofertilizers function as key player in sustainable agriculture by improving soil fertility, plant tolerance and crop productivity. Microb. Cell Factories 2014, 13, 66. [Google Scholar] [CrossRef]
  59. Xu, Y.; Feng, Y.-Y. Feasibility of Sewage Sludge Leached by Aspergillus Niger in Land Utilization. Pol. J. Environ. Stud. 2016, 25, 405–412. [Google Scholar] [CrossRef] [PubMed]
  60. Duan, Y.; Awasthi, S.K.; Chen, H.; Liu, T.; Zhang, Z.; Zhang, L.; Awasthi, M.K.; Taherzadeh, M.J. Evaluating the impact of bamboo biochar on the fungal community succession during chicken manure composting. Bioresour. Technol. 2019, 272, 308–314. [Google Scholar] [CrossRef] [PubMed]
  61. Tan, H.; Yu, Y.; Zhu, Y.; Liu, T.; Miao, R.; Hu, R.; Peng, W.; Chen, J. Impacts of size reduction and alkaline-soaking pretreatments on microbial community and organic matter decomposition during wheat straw composting. Bioresour. Technol. 2022, 360, 127549. [Google Scholar] [CrossRef] [PubMed]
  62. Ozimek, E.; Hanaka, A. Mortierella Species as the Plant Growth-Promoting Fungi Present in the Agricultural Soils. Agriculture 2021, 11, 7. [Google Scholar] [CrossRef]
  63. Awasthi, M.K.; Li, J.; Kumar, S.; Awasthi, S.K.; Wang, Q.; Chen, H.; Wang, M.; Ren, X.; Zhang, Z. Effects of biochar amendment on bacterial and fungal diversity for co-composting of gelatin industry sludge mixed with organic fraction of municipal solid waste. Bioresour. Technol. 2017, 246, 214–223. [Google Scholar] [CrossRef] [PubMed]
  64. Partanen, P.; Hultman, J.; Paulin, L.; Auvinen, P.; Romantschuk, M. Bacterial diversity at different stages of the composting process. BMC Microbiol. 2010, 10, 94. [Google Scholar] [CrossRef] [PubMed]
  65. Goswami, D.; Thakker, J.N.; Dhandhukia, P.C. Portraying mechanics of plant growth promoting rhizobacteria (PGPR): A review. Cogent Food Agric. 2016, 2, 1127500. [Google Scholar] [CrossRef]
  66. Krishna, M.P.; Mohan, M. Litter decomposition in forest ecosystems: A review. Energy Ecol. Environ. 2017, 2, 236–249. [Google Scholar] [CrossRef]
  67. Sowani, H.; Kulkarni, M.; Zinjarde, S. An insight into the ecology, diversity and adaptations of Gordonia species. Crit. Rev. Microbiol. 2018, 44, 393–413. [Google Scholar] [CrossRef]
  68. Xu, Z.; Ma, L.; Zhao, B.; Li, Y.; Chen, Y.; Deng, Y.; Wang, Y. Humification process enhancement through relative abundance promotion of Talaromyces and Coprinopsis by inoculated Phanerochaete chrysosporium during the secondary fermentation of composting. Environ. Sci. Pollut. Res. 2023, 30, 9060–9065. [Google Scholar] [CrossRef]
  69. El-Shahir, A.A.; El-Tayeh, N.A.; Ali, O.M.; Abdel Latef, A.A.H.; Loutfy, N. The Effect of endophytic Talaromyces pinophilus on growth, absorption and accumulation of heavy metals of Triticum aestivum grown on sandy soil amended by sewage sludge. Plants 2021, 10, 2659. [Google Scholar] [CrossRef] [PubMed]
  70. Silva, J.P.; Ticona, A.R.P.; Hamann, P.R.V.; Quirino, B.F.; Noronha, E.F. Deconstruction of Lignin: From Enzymes to Microorganisms. Molecules 2021, 26, 2299. [Google Scholar] [CrossRef] [PubMed]
  71. Huang, W.; Hammel, K.E.; Hao, J.; Thompson, A.; Timokhin, V.I.; Hall, S.J. Enrichment of Lignin-Derived Carbon in Mineral-Associated Soil Organic Matter. Environ. Sci. Technol. 2019, 53, 7522–7531. [Google Scholar] [CrossRef] [PubMed]
  72. Sun, L.; Liu, T.; Müller, B.; Schnürer, A. The microbial community structure in industrial biogas plants influences the degradation rate of straw and cellulose in batch tests. Biotechnol. Biofuels 2016, 9, 128. [Google Scholar] [CrossRef]
  73. Hernández, T.; Chocano, C.; Moreno, J.-L.; García, C. Use of compost as an alternative to conventional inorganic fertilizers in intensive lettuce (Lactuca sativa L.) crops—Effects on soil and plant. Soil Tillage Res. 2016, 160, 14–22. [Google Scholar] [CrossRef]
  74. Trupiano, D.; Cocozza, C.; Baronti, S.; Amendola, C.; Vaccari, F.P.; Lustrato, G.; Di Lonardo, S.; Fantasma, F.; Tognetti, R.; Scippa, G.S. The effects of biochar and its combination with compost on lettuce (Lactuca sativa L.) growth, soil properties, and soil microbial activity and abundance. Int. J. Agron. 2017, 2017, 3158207. [Google Scholar] [CrossRef]
  75. Solaiman, Z.M.; Yang, H.; Archdeacon, D.; Tippett, O.; Tibi, M.; Whiteley, A.S. Humus-rich compost increases lettuce growth, nutrient uptake, mycorrhizal colonisation, and soil fertility. Pedosphere 2019, 29, 170–179. [Google Scholar] [CrossRef]
Horticulturae 09 01304 g001
Figure 1. Fungi (a) and bacteria (b) community composition at the genus and phylum level in the soils samples collected in August 2018 and July 2019 after the harvest of lettuce of the first and second growing cycle, respectively (HM, hop leaves + cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS+ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1).
Figure 1. Fungi (a) and bacteria (b) community composition at the genus and phylum level in the soils samples collected in August 2018 and July 2019 after the harvest of lettuce of the first and second growing cycle, respectively (HM, hop leaves + cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS+ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1).
Horticulturae 09 01304 g001
Horticulturae 09 01304 g002
Figure 2. Simpson (a) and Shannon (b) diversity indexes of fungi and bacteria species identified in the soils samples collected in August 2018 and July 2019 after the harvest of lettuce of the first and second growing cycle, respectively (HM, hop leaves + cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS + ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1). Means followed by the same letter are not statistically different by Tukey HSD test (α = 0.05).
Figure 2. Simpson (a) and Shannon (b) diversity indexes of fungi and bacteria species identified in the soils samples collected in August 2018 and July 2019 after the harvest of lettuce of the first and second growing cycle, respectively (HM, hop leaves + cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS + ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1). Means followed by the same letter are not statistically different by Tukey HSD test (α = 0.05).
Horticulturae 09 01304 g002
Table 1. Compost mixtures, respective mixture ratios, carbon/nitrogen (C/N) ratio, soil application rate, and treatment abbreviation.
Table 1. Compost mixtures, respective mixture ratios, carbon/nitrogen (C/N) ratio, soil application rate, and treatment abbreviation.
Compost MixturesRatioC/NSoil Application RateAbbreviation
Hop leaves + cow manure (HM)1:511.5820 t ha−1 dry weight (D1)HM1:5D1
Hop leaves + cow manure (HM)1:312.0320 t ha−1 dry weight (D1)HM1:3D1
Hop leaves + cow manure (HM)1:110.5320 t ha−1 dry weight (D1)HM1:1D1
Hop leaves + wheat straw (HS)1:227.7520 t ha−1 dry weight (D1)HS1:2D1
Hop leaves + wheat straw (HS)1:121.9520 t ha−1 dry weight (D1)HS1:1D1
Hop leaves + wheat straw + hop stems ash (HSA)1:1:0.0425.2920 t ha−1 dry weight (D1)HSA1:1:0.04D1
Hop leaves + wheat straw (HS)1:0.515.7920 t ha−1 dry weight (D1)HS1:0.5D1
Hop leaves + wheat straw (HS)1:227.7540 t ha−1 dry weight (D2)HS1:2D2
Hop leaves + wheat straw (HS)1:0.515.7940 t ha−1 dry weight (D2)HS1:0.5D2
Table 2. Total nitrogen (N), total organic carbon (TOC), microbial nitrogen (Mic-N), microbial carbon (Mic-C), soil basal respiration (SBR; cumulative respiration during a 7-day incubation), metabolic quotient (qCO2), Mic-C/TOC ratio and acid phosphatase activity (APA) from soil samples collected in August 2018 after the harvest of lettuce of the first and second growing cycle (HM, hop leaves+cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS + ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1).
Table 2. Total nitrogen (N), total organic carbon (TOC), microbial nitrogen (Mic-N), microbial carbon (Mic-C), soil basal respiration (SBR; cumulative respiration during a 7-day incubation), metabolic quotient (qCO2), Mic-C/TOC ratio and acid phosphatase activity (APA) from soil samples collected in August 2018 after the harvest of lettuce of the first and second growing cycle (HM, hop leaves+cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS + ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1).
Total NTOCMic-NMic-CSBRqCO2Mic-C/TOCAPA
Treatment-------- g kg−1 -------------------- mg kg−1 -------------mg CO2-C g–1mg g−1µg PNP g−1
2018
Control0.94 c22.0 e15.5 c55.1 b84.8 a92.3 a2.5 b195.6 bc
HM1:5D11.50 b21.9 e32.6 bc257.8 b127.6 a23.1 ab11.9 b193.6 bc
HM1:3D11.32 bc23.8 de36.7 abc98.6 b86.8 a39.5 ab4.1 b193.2 bc
HM1:1D11.77 b21.0 e27.4 bc262.5 b91.2 a14.5 b12.9 ab199.5 bc
HS1:2D11.30 bc26.6 cd43.9 abc321.7 b95.1 a22.2 ab12.1 ab188.4 c
HS1:1D11.47 b30.4 b40.1 abc776.2 a92.7 a5.1 b26.5 a198.6 bc
HS2:1D11.68 b30.7 b67.3 ab425.6 ab93.6 a12.7 b13.8 ab207.6 bc
HSA1:1:0.04D11.60 b29.7 bc51.8 abc210.5 b97.4 a21.0 ab7.2 b217.7 bc
HS1:2D21.56 b32.3 ab49.0 abc280.6 b107.5 a16.1 b8.7 b243.6 ab
HS2:1D22.34 a35.4 a74.8 a178.8 b108.0 a29.4 ab5.0 b276.1 a
Prob. > F<0.0001<0.00010.00170.00040.27130.03090.00070.0125
Standard error0.060.923.8041.593.655.851.436.35
2019
Control1.31 f23.0 d24.4 d78.3 c56.8 b117.1 a3.4 bc173.8 b
HM1:5D11.52 de27.6 bc49.4 bcd113.4 c55.7 b72.3 a4.1 bc189.9 b
HM1:3D11.59 cde26.9 c27.3 cde72.4 c50.3 b114.0 a2.7 c183.1 b
HM1:1D11.55 e27.7 bc31.4 cde123.9 c54.3 b65.7 a4.5 bd195.6 ab
HS1:2D11.52 e28.2 bc45.4 bcde127.1 c59.9 b68.6 a4.5 bc199.6 ab
HS1:1D11.64 bcd28.3 bc37.8 bcde135.3 c51.2 b55.7 a4.8 bc192.4 b
HS2:1D11.69 bc28. bc54.3 abc120.3 c56.0 b67.6 a4.3 bc186.8 b
HSA1:1:0.04D11.48 e27.3 c44.4 bcde137.1 bc56.0 b66.4 a5.0 bc189.0 b
HS1:2D21.73 b30.1 ab60.0 ab262.0 a121.1 a67.3 a8.7 a244.8 a
HS2:1D22.08 a31.9 a73.3 a204.6 ab150.1 a106.6 a6.4 ab186.9 b
Prob. > F<0.0001<0.0001<0.0001<0.0001<0.00010.0534<0.00010.0084
Standard error0.030.382.699.4819.765.540.2904.16
Means followed by the same letter are not statistically different by Tukey HSD test (α = 0.05).
Table 3. Effect of different compost treatments on microbial populations of the soils samples collected in August 2018 and July 2019 after the harvest of lettuce of the first and second growing cycle, respectively (HM, hop leaves + cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS+ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1).
Table 3. Effect of different compost treatments on microbial populations of the soils samples collected in August 2018 and July 2019 after the harvest of lettuce of the first and second growing cycle, respectively (HM, hop leaves + cow manure at the ratios 1:5, 1:3, and 1:1; HS, hop leaves + wheat straw at the ratios 1:2, 1:1, and 2:1; and HSA, HS+ash from hop stems at the ratios 1:1:0.04; D1 and D2, 20 and 40 kg dw ha−1).
TreatmentFungiBacteriaActinomycetesAct./Bacteria
------------------------ [log (cfu g−1)] ------------------------(%)
2018Control4.52 a7.04 ab6.78 a56.06 a
HM1:5D15.01 a7.39 ab7.13 a56.20 a
HM1:3D15.08 a7.00 b6.61 a43.08 a
HM1:1D15.00 a7.23 ab6.92 a52.78 a
HS1:2D14.93 a7.14 ab6.74 a45.80 a
HS1:1D15.11 a7.12 ab6.85 a54.95 a
HS2:1D15.03 a7.44 a7.11 a46.85 a
HSA1:1:0.04D14.90 a7.25 ab7.02 a61.60 a
HS1:2D25.11 a7.21 ab6.69 a30.45 a
HS2:1D25.11 a7.23 ab6.72 a30.89 a
Prob. > F0.07330.03530.07010.3741
Standard error0.250.180.2417.57
2019Control4.44 bcde6.63 c6.31 a48.89 a
HM1:5D14.24 de6.88 bc6.49 a42.39 ab
HM1:3D14.05 e6.77 bc6.29 a33.77 ab
HM1:1D14.32 cde6.86 bc6.47 a50.45 a
HS1:2D14.52 abcde6.99 abc5.97 a13.86 ab
HS1:1D14.83 abc7.06 abc6.39 a22.96 ab
HS2:1D14.88 ab7.15 ab6.26 a14.48 ab
HSA1:1:0.04D14.62 abcd7.05 abc6.18 a13.65 ab
HS1:2D24.75 abcd7.14 ab6.24 a13.01 ab
HS2:1D25.00 a7.32 a6.25 a8.96 b
Prob. > F>0.00010.00100.65450.0029
Standard error0.330.230.2919.15
Means followed by the same letter are not statistically different by Tukey HSD test (α = 0.05).
Table 4. Spearman’s correlation coefficients between microbial communities and soil properties— total nitrogen (N), total organic carbon (TOC), microbial nitrogen (Mic-N), microbial carbon (Mic-C), soil basal respiration (SBR; average daily respiration), metabolic quotient (qCO2), and acid phosphatase activity (APA)—of the soil samples collected in August 2018 and July 2019, after the harvest of lettuce of the first and second growing cycle, respectively.
Table 4. Spearman’s correlation coefficients between microbial communities and soil properties— total nitrogen (N), total organic carbon (TOC), microbial nitrogen (Mic-N), microbial carbon (Mic-C), soil basal respiration (SBR; average daily respiration), metabolic quotient (qCO2), and acid phosphatase activity (APA)—of the soil samples collected in August 2018 and July 2019, after the harvest of lettuce of the first and second growing cycle, respectively.
Total NTOCMic-NMic-CSBRqCO2APA
---------- g kg−1 --------------------- mg kg−1 ------------mg CO2-C g–1µg PNP g−1
2018
Fungi0.3440.2640.1180.3460.058−0.3450.174
Bacteria0.524 **0.1610.3170.3170.261−0.2370.246
Actinomycetes0.394 *−0.0900.2250.3030.183−0.2070.106
2019
Fungi0.494 **0.509 **0.463 *0.555 **0.161−0.2020.105
Bacteria0.534 **0.486 **0.588 **0.574 **0.154−0.3140.084
Actinomycetes0.159−0.064−0.158−0.153−0.0560.086−0.113
Correlation are significant at the <0.05 level * and 0.01 level **.
Table 5. Spearman’s correlation coefficients between fungi and bacteria Genus taxa level and soil total nitrogen (N) and total organic carbon (TOC) of the soil samples collected in August 2018 and July 2019, after the harvest of lettuce of the first and second growing cycle, respectively.
Table 5. Spearman’s correlation coefficients between fungi and bacteria Genus taxa level and soil total nitrogen (N) and total organic carbon (TOC) of the soil samples collected in August 2018 and July 2019, after the harvest of lettuce of the first and second growing cycle, respectively.
PhylumGenus20182019
Total NTOCTotal NTOC
---------- g kg−1 ------------------ g kg−1 --------
Fungi
AscomycotaAcrostalagmus0.390 *0.544 **0.0910.263
AscomycotaAspergillus−0.146−0.3610.420 *0.460 *
AscomycotaDoratomyces0.2350.547 **0.2260.197
AscomycotaPenicillium−0.118−0.323n.d.n.d.
AscomycotaTalaromyces0.1100.2060.413 *0.432 *
ZygomycotaMortierella0.167−0.0670.1950.222
ZygomycotaRhizopus0.126−0.1750.2490.184
n.d.0.222−0.1550.3180.266
Bacteria
ActinobacteriaArthrobacter0.332−0.150−0.091−0.308
ActinobacteriaGordonia0.3190.627 **0.0340.117
ActinobacteriaMicrobacterium−0.147−0.0850.0930.185
ActinobacteriaStreptomyces0.3040.2430.460 *0.478 **
BacteroidetesChryseobacterium−0.138−0.224n.d.n.d.
FirmicutesBacillus0.389 *0.3080.541 **0.524 **
FirmicutesLysinibacillus−0.236−0.032n.d.n.d.
FirmicutesNeobacillus0.1360.339−0.0710.017
ProteobacteriaPseudoxanthomonas0.112−0.2620.0820.064
ProteobacteriaVariovorax0,118−0.1820.2910.290
Correlation are significant at the <0.05 level * and 0.01 level **: n.d. (fungi/bacteria not present in soil samples collected in 2019).
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Afonso, S.; Pereira, E.L.; Arrobas, M.; Rodrigues, M.Â.; Choupina, A. Composts Obtained by Mixing Hop Leaves with Wheat Straw or Farmyard Manure Improved Soil Properties and Increased Microbial Communities. Horticulturae 2023, 9, 1304. https://doi.org/10.3390/horticulturae9121304

AMA Style

Afonso S, Pereira EL, Arrobas M, Rodrigues MÂ, Choupina A. Composts Obtained by Mixing Hop Leaves with Wheat Straw or Farmyard Manure Improved Soil Properties and Increased Microbial Communities. Horticulturae. 2023; 9(12):1304. https://doi.org/10.3390/horticulturae9121304

Chicago/Turabian Style

Afonso, Sandra, Ermelinda L. Pereira, Margarida Arrobas, M. Ângelo Rodrigues, and Altino Choupina. 2023. "Composts Obtained by Mixing Hop Leaves with Wheat Straw or Farmyard Manure Improved Soil Properties and Increased Microbial Communities" Horticulturae 9, no. 12: 1304. https://doi.org/10.3390/horticulturae9121304

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop