Next Article in Journal
Does Sodium Nitroprusside Alleviate Water Deficit Stress in Impatiens walleriana Shoots Grown In Vitro?
Previous Article in Journal
Perspectives on Effective Long-Term Management of Carbon Stocks in Chernozem under Future Climate Conditions
Previous Article in Special Issue
Straw Return Decomposition Characteristics and Effects on Soil Nutrients and Maize Yield
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Agriculture
Volume 13
Issue 10
10.3390/agriculture13101902
agriculture-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

Soil Microbial Community Responses to Cyanobacteria versus Traditional Organic Fertilizers

by
Antisar Afkairin
1,*,
Mary Stromberger
1,2,
Heather Storteboom
1,
Allison Wickham
1,
David G. Sterle
1,3 and
Jessica G. Davis
1,4
1
Department of Soil and Crop Sciences, C127 Plant Sciences Building, Colorado State University, Fort Collins, CO 80523-1170, USA
2
The Ohio State University Graduate School, Columbus, OH 43210, USA
3
Western Colorado Research Center, 3168 B 1/2 Rd, Grand Junction, CO 81503, USA
4
Agricultural Experiment Station, Nutrien Ag Sciences Building, Colorado State University, Fort Collins, CO 80523-3001, USA
*
Author to whom correspondence should be addressed.
Agriculture 2023, 13(10), 1902; https://doi.org/10.3390/agriculture13101902
Submission received: 17 August 2023 / Revised: 22 September 2023 / Accepted: 26 September 2023 / Published: 28 September 2023
(This article belongs to the Special Issue Effects of Agricultural Management Practices on Soil Fertility and Microbiome Structure)
Browse Figures
Review Reports Versions Notes

Abstract

:
This study explores the impact of diverse organic fertilizers, including a non-traditional cyanobacteria-based alternative, on soil microbial communities in varying soil types and depths. The research aims to elucidate the effects of these fertilizers on soil microorganisms in certified organic cucumber (Cucumis sativus) field and peach (Prunus persica) orchard settings. Fertilizers were applied either on the soil surface or banded 5 cm below the soil surface, and microbial ester-linked fatty acids (EL-FAMEs) were analyzed in collected soils. Notably, cyanobacteria and Neptune hydrolyzed fish emulsion fertilizers induced significant alterations in the microbial communities of cucumber plots, enhancing microbial biomass and favoring the proliferation of Gram-negative bacteria, Gram-positive bacteria, and actinomycetes compared to other treatments. In the peach orchard, fertilizer choice differentially impacted microbial communities, especially in the first year and at greater soil depths. Notably, the supplementation of poultry manure with cyanobacteria fertilizer resulted in augmented microbial biomass and relative fungal and arbuscular mycorrhizal fungal abundances compared to poultry manure alone. These shifts have promising implications for organic vegetable and fruit cultivation. The study further underscores the potential of cyanobacteria-based fertilizers to reduce reliance on traditional options and minimize manure application, promoting self-sufficiency and benefiting soil microorganisms, plant growth, and the ecosystem. Thus, the research emphasizes the importance of exploring and adopting cyanobacteria-based fertilizers to bolster sustainable agricultural practices.

1. Introduction

Fertilization is a worldwide management practice used to improve nutrient availability to plants. Organic and inorganic fertilizer applications to soil can also alter structure and biomass, enhance soil microbial activity, alter the microbial community structure, and ultimately improve soil health [1,2]. Soil microbial biomass and enzyme activities increase when organic fertilizers are applied to soil [3,4]. Studies in grassland plots have determined that the soil microbial community had been affected by the applications of manure and/or inorganic fertilizers over a long period of time (nearly 100 years) [5]. Commonly, organic fertilizers have a great influence on soil microbial communities. For instance, in an 8-month field trial, manure applications increased microbial biomass C, soil respiration, enzyme activities, and N mineralization [6]. Microbial changes may be due to the addition of organic C to the soil as manure, which benefits saprophytic soil microorganisms, in addition to providing nutrients such as N, P, and K [6]. However, in short-term (less than 1 year) fertilizer experiments, others found no significant effects on soil microbial communities despite the existence of fertilizer inputs [3,7,8].
Organic fertilizers such as fish emulsion, manure, and compost are the primary sources of N fertilizer for organic or low input cropping systems, but they can be expensive and difficult to transport [9]. However, there is another fertilizer option being developed that allows growers to produce organic N fertilizer on-farm, in the form of living cyanobacteria [10]. Cyanobacterial fertilizer, or cyano-fertilizer, can be a good source of N for organic growers due to its low residual NO3-N compared to fish emulsion, its effectiveness compared to other organic fertilizers [9], and its ability to be produced on-farm [10,11].
Cyanobacteria are prokaryotic oxygenic phototrophs found in nearly every habitat on Earth [12]. Cyanobacteria play a major role in agriculture by contributing to different functions such as inoculants for reinforcing soil fertility as well as enhancing soil structure in addition to improving crop yield because they perform oxygenic photosynthesis, and they have the unique ability to fix N from the atmosphere while also solubilizing P and increasing P content in plants [12,13]. The application of cyanobacterial biofertilizers has been shown to reduce N2O and NH3 emissions as compared to solid organic fertilizers, such as blood and feather meals [14,15]. This result can lead to improved soil fertility and plant growth because of increased organic matter content and enzymatic activities (dehydrogenase and nitrogenase) [16,17]. In addition, cyanobacteria produce phytohormones [18], and auxin additions in fertilizers have been shown to be positively correlated with lettuce β-carotene concentrations [19].
Fatty acid methyl ester (FAME) profiling is an effective method to characterize microbial community biomass and composition, including bacterial and fungal components, and to assess rapid changes in microbial communities in response to soil amendments [20]. There are two methods to measure the FAMEs—Phospholipid Fatty Acid Analysis (PLFA) and Ester-Linked Fatty Acid Methyl Ester (EL-FAME), and both provide information on soil microbial community and biomass composition in relation to soil management [21]. Extraction of microbial fatty acids using the EL-FAME method provides a relatively simple, rapid analysis, relative to the PLFA method, and has the ability to distinguish microbial communities that are different in biomass and structure among different soil types, environments, and management practices [22,23]. Since fatty acids and phospholipids are degraded rapidly by endogenous and exogenous phospholipases upon cell death, they are reliable measures of viable cell biomass [24,25]. Additionally, fatty acids and phospholipids can be used to identify broad physiological groups of bacteria [24,26,27]. These methods have detected differences in microbial community composition among rhizospheres of different crop species, cultivars, and soil types [28,29]. When the structure of the microbial community is more stable, there are important effects on the rates of denitrification, nitrification, and N fixation [30]. Also, the structure of the community and soil microbial biomass are good indicators of soil health and quality, and cropland management practices can change those parameters due to their sensitivity [31].
Microbial community fatty acid profiling can provide a good understanding of how microbial biomass, community composition, and potentially their activity and processes are affected by fertilization practices or soil depth, although very few studies have been conducted to date. The PLFA method was employed to determine the effect of inorganic N fertilizer and dairy manure application on microbial communities and biomass composition [24]. They found that manured surface soils were enriched with bacterial mono-unsaturated (Gram-negative) and eukaryotic markers. Microbial communities also differ in composition and biomass as a function of depth within a soil type [32]. For example, Gram-negative bacteria, fungi, and protozoa were less abundant at deeper depths, while Gram-positive bacteria and actinomycetes were more abundant with depth [32]. A better understanding of microbial community responses to diverse organic fertilizers is needed, as studies are scarce and often focused on manures. Very little is known about how organic fertilizers, such as fish emulsion and new fertilizers, including cyano-fertilizer, affect soil microbial communities. In addition, because organic fertilizers are sometimes applied at or below the soil surface, organic fertilizers may have differential effects on soil microbial communities depending on soil depth. Therefore, the aim of this study was to determine the effect of different organic fertilizers on the soil microbial community in two different soil depths within different cropping systems over a 2-year period. We hypothesized that the soil microbial community composition would be impacted by organic fertilizer type. In addition, we predicted that variations would be observed across soil depths and cropping systems.

2. Materials and Methods

2.1. Experimental Design and Soil Sampling

Soil samples were collected from a certified organic cucumber (Cucumis sativus) field and a certified organic peach (Prunus persica) orchard, both in Colorado. The first location was at the Colorado State University Horticultural Research Center (4300 E County Road 50, 80524) in Fort Collins, CO, USA. The second location was at Ela Family Farms in Hotchkiss, CO, USA. Each location represented a unique experiment with different treatments, and both experiments used a randomized complete block design (RCBD).
The first experiment was conducted during the 2015 growing season on certified organic land at the Colorado State University Horticultural Research Center (4300 E County Road 50, 80524) near Fort Collins, CO, USA. A cucumber plot previously described by [33] had a soil classified as fine, smectitic, mesic, Aridic Argiustoll of the Nunn series [34], soil pH of 8.1, and organic matter of 2.7%. The average maximum temperature during the growing season (1 May–30 September) was 26 °C, and the average minimum temperature was 17 °C. There was 6.1 cm of rain during the growing season. Prior to 2015, plots at this location were planted to a buckwheat cover crop and received an annual application of manure which was broadcast and then tilled in, following certified organic practices. The fertilizer treatments applied in 2015 were as follows: unfertilized control, blood meal 12-0-0 (Down to Earth, Inc., Eugene, OR, USA) surface applied or sub-surface applied, feather meal 12-0-0 (Down to Earth, Inc., Eugene, OR, USA) surface applied or sub-surface applied, hydrolyzed fish emulsion 2-4-1 (Neptune’s Harvest, Gloucester, MA, USA), AlaskaTM non-hydrolyzed fish emulsion 5-1-1 (Planet Natural, Bozeman, MT, USA), and cyano-fertilizer. The cyano-fertilizer (Anabaena cylindrica) was originally cultured from sediment collected from Richard’s Lake in Fort Collins, CO, USA [10]. It was grown on-farm in modified Allen–Arnon media using methods described by [10] and had an average 23.3 mg N kg−1 or <1% N by weight (Total Kjeldahl N) and pH of 7.13. This study included eight treatments replicated four times in a randomized complete block. This included a control and three types of fertilizer applications applied at 76 kg N ha−1: solid fertilizers (blood meal, feather meal) surface banded near the drip irrigation line, liquid fertilizers (hydrolyzed fish emulsion, non-hydrolyzed fish emulsion, and cyano-fertilizer) applied through the drip irrigation system, and solid fertilizers (blood meal, feather meal) sub-surface banded 5 cm deep near the drip irrigation line.
The second experiment took place in a peach orchard in Hotchkiss, CO, USA previously described by [35]. The soil had pH 7.6, and soil type was Mesa loam which is classified as a mesic Typic Calciargid [34]. Annual precipitation averages 580 mm, and air temperatures range from −12 °C minimum and −2 °C maximum in January to 10 °C minimum and 25 °C maximum in July www.weather-atlas.com (accessed on 27 September 2023). The following fertilizer treatments were applied for two years during 2014–2015: 112 kg N ha−1 of RichlawnTM 5-3-2 dried poultry manure (HM) (Platteville, CO, USA), 112 kg N ha−1 of dried poultry manure + 4.9 kg N ha−1 of cyano-fertilizer (HM+C), and 84 kg N ha−1 of dried poultry manure + 4.9 kg N ha−1 as cyano-fertilizer (LM+C). Poultry manure was applied to the plots by broadcasting within the tree rows but not in the alleys between the trees. Manure was not incorporated. The cyano-fertilizer was applied separately from manure in liquid form through the micro-sprinkler irrigation system. These treatments were replicated five times in a randomized complete block design.
After harvest of either cucumbers (August 2015) or peaches (August 2014 and 2015), soil samples were collected from each location by compositing ten 0–2.5 cm deep cores and ten 2.5–7.5 cm deep cores from within each plot; samples were not taken within 30 cm of each plot border to minimize edge effects. After sampling, soil samples were stored in coolers and transported to the laboratory, homogenized by hand, placed into 50 mL sterile polypropylene centrifuge tubes, and stored at −80 °C until further use.

2.2. Ester-Linked Fatty Acid Methyl Ester (EL-FAME) Analysis

Prior to EL-FAME extraction, soil moisture was determined gravimetrically. Ten g of soil were weighed, dried in an oven at 105 °C for 24 h, and weighed again. Community structure was determined from frozen soil samples by EL-FAME [21]. Whole-cell fatty acids including phospholipids, glycolipids, and neutral lipids were extracted from the soil. Each soil sample was placed into a 50 mL glass centrifuge tube, and Reagent 1 (0.2 N KOH prepared in methanol) was added to each tube. Then, samples were extracted for one hour in a 37 °C water bath, during which time samples were vortexed for 10 s every 10 min. After that, Reagent 2 (1.0 M acetic acid) was added to neutralize the pH. Then Reagent 3 (10 mL hexane) was added to each tube to extract fatty acids into the organic phase (hexane) and vortexed for 20 s. Then, all tubes were centrifuged at 480× g (Sorvall HS-4 swinging bucket motor) for 20 min at 4 °C to separate the hexane and aqueous phases. By using a Pasteur pipet, two-thirds of clean hexane supernatant was removed from each tube and transferred into new glass tubes (17 × 100 mm). Samples were prepared for gas chromatography (GC) by evaporating the hexane solvent under N2 gas. The dried EL-FAME residue was then re-dissolved by adding100 μL of hexane containing 0.10 μg μL–1 nonadecanoic acid methyl ester (C 19:0) as an internal standard. All samples were stored at –20 °C prior to analysis.
EL-FAME extracts were analyzed by gas chromatography-mass spectrometry (GC-MS) with a Trace GC coupled to a Thermo TSQ8000 Evo mass spectrometer (Thermo Scientific, Waltham, MA, USA), equipped with a Phenomenex ZB-5HT Inferno GC column (30 m × 0.25 mm × 0.25 µm). The GC inlet was set at 285 °C, and the oven temperature was programmed at 60 °C for 2 min, a ramp of 15 °C per min to 330 °C, and then held at 330 °C for 10 min. Peaks were identified based on mass spectral and retention time matching to a 37 FAME mixture (Sigma-Aldrich, Saint Louis, Missouri, USA) and a bacterial acid methyl ester mixture (Sigma-Aldrich). Additional FAMEs were detected based on mass spectral matching to the NIST Mass Spectral Library (v14) [36,37]. According to [21,23], the EL-FAME biomarkers were attributed to the following microbial groups: Gram-positive bacteria (i14:0, i15:0, a15:0, i16:0, i17:0, and a17:0), Gram-negative bacteria (16:1ω7c, 17:1ω7, 17:0cy, 18:1ω7c, 18:1ω8, and 19:0cy), actinomycetes (10Me16:0, 10Me17:0, and 10Me18:0), fungi (18:2ω6,9c), and arbuscular mycorrhizal (AM) fungi (16:1ω5c). Total bacterial biomass was determined from the sum of biomarkers of Gram-positive and Gram-negative bacteria, actinomycetes, as well as the general bacterial fatty acids 15:0 and 17:0. Total microbial biomass was the sum of all bacterial and fungal EL-FAMEs. Stress indicators were signified by the ratio of 17:0cy to its precursor 16:1ω7c (Stress 1), and by the ratio of 19:0 cy to its precursor 18:1ω7c (Stress 2) [38,39].

2.3. Statistical Analysis

Data were analyzed using R Version 3.5.3 and RStudio Version 1.0.153. (The R Foundation for Statistical Computing) [40]. We used a mixed model that considered treatment, depth, and their interaction as fixed effects, and block and block × treatment as random effects to account for the randomized complete block design and the spatial correlation between soil samples taken at consecutive depths in the same plot. Comparisons of soil microbial community groups among organic fertilizer treatments were significant (p < 0.05), and Tukey adjustment pairwise comparisons were used to identify variables that differed significantly among treatments [40]. Soil microbial community EL-FAME data were analyzed by Principal Components Analysis (PCA), after normalizing the data from nmole g−1 soil using the PC-ORD statistical package (MjM Software version 4, Gleneden Beach, OR, USA, 1999). Communities were analyzed separately by location, using the PC-ORD version 6 statistical package [41].

3. Results

3.1. Microbial Community Structure

In the cucumber experiment, cyano-fertilizer and Neptune hydrolyzed fish emulsion resulted in a different soil microbial community structure compared to microbial communities of control soil and soil receiving other organic fertilizer treatments, with the PCA analysis explaining 54.6% of the variance (Figure 1). Changes in microbial community structure were consistent between the two soil depths. Differences in community structure were largely attributed to fertilizer-induced shifts in several EL-FAMEs, as indicated by the PCA analysis. EL-FAMEs with negative eigenvector coefficients for PC 1, and therefore associated with communities from cyanobacteria or fish emulsion fertilized soil, included biomarkers for actinomycetes (10Me17:0 and 10Me18:0) and Gram-negative bacteria (17:1ω7, 17:0cy, 18:1ω8) (Table 1). Conversely, the EL-FAME with the highest positive eigenvector coefficient for PC 1 was 18:1ω9 (fungi). Differences in community structures in soil fertilized with cyanobacteria or fish emulsion versus communities in all other soil treatments were statistically significant, according to multi-response blocked permutation procedure (MRBP) analysis conducted in PC-ORD.
In the peach orchard, soil microbial community structure was affected by the different fertilizers in 2014 but not in 2015. In 2014, microbial community structures in several plots receiving the LM+C treatment (75 kg N ha−1 as dried poultry manure + 25 kg N ha−1 as cyano-fertilizer) differed from the other microbial communities, with the PCA analysis explaining 67.4% of the variance along PC1 and 2 (Figure 2). EL-FAMEs associated with several LM+C communities included 16:1ω5 (AM fungi), 18:1ω9 (fungi), and i14:0 (Gram-positive bacteria), based on positive eigenvector coefficients for PC1 and/or PC2 (Table 2). Pairwise comparisons among the fertilizer treatments in 2014 by MRBP analysis in PC-ORD showed that the lower application rate of dried poultry manure with cyano-fertilizer treatment (LM+C) resulted in a soil microbial community structure significantly different from the other fertilizer treatments.

3.2. Microbial Community Biomass

Total microbial EL-FAME biomass in the top layer of soil (0–2.5 cm depth) of the cucumber experiment ranged from 24 to 73 nmol EL-FAMEs g−1 soil (Table 3). However, microbial EL-FAME biomass was lower in the deeper soil layer (2.5–7.5 cm depth), ranging from 9 to 38 nmol g−1 soil, and there was no significant difference among treatments in the deeper depth. Within the 0–2.5 cm soil depth, cyano-fertilizer and Neptune hydrolyzed fish emulsion were the only organic fertilizers that increased microbial EL-FAME biomass compared to the control. Total microbial, Gram-positive bacterial, and Gram-negative bacterial EL-FAME biomass levels were ~2.5×, ~2.4×, and 3–4× greater in surface soil receiving either cyano-fertilizer or Neptune hydrolyzed fish emulsion than in control soil (Table 3). Neptune hydrolyzed fish emulsion and cyano-fertilizer also increased actinomycete EL-FAME biomass by an order of magnitude in surface soil compared to the control, although the effect was not statistically significant for the cyano-fertilizer.
The addition of cyano-fertilizer to poultry-manure-fertilized peach orchard soil affected microbial EL-FAME biomass in soil at the deeper depth (2.5–7.5 cm) in 2014. Total microbial EL-FAME biomass was 1.6× greater in the LM + C treatment (75 kg N ha−1 dried poultry manure + 25 kg N ha−1 cyano-fertilizer) compared to HM soil receiving 100 kg N ha−1 dried poultry manure (Table 4). Biomass of EL-FAMEs associated with Gram-negative bacteria, fungi, and AM fungi were 1.5×, 1.7×, and 2× greater in subsurface soils fertilized with LM+C compared to the HM treatment, respectively (Table 4). Fertilizer treatment had no significant effect on Gram-positive bacterial or actinomycete EL-FAME biomass. Overall, the peach orchard soil had greater EL-FAME biomass compared to the EL-FAME biomass in the cucumber soil.

4. Discussion

4.1. Microbial Community Responses to Fertilizers

Organic fertilizers can have significant short-term (<1 year) and long-term impacts on soil microbial communities. In this study, the microbial community structure and biomass was sensitive to different organic fertilizers in both annual vegetable and perennial tree fruit systems, although not consistently each year. Short-term increases in microbial biomass have also occurred when vermicompost was applied [42]. In our study, total microbial biomass and relative abundance and biomass of bacteria, actinomycetes, fungi, and AM fungi were influenced in the short term by organic fertilizer type in one or both experiments [43,44]. A greater fungal biomass is generally present under conditions with greater C availability and moisture [31]. In addition, increased fungal biomass has also been recorded in situations with plant residues and greater soil moisture [44,45]. Augmentation of fungal biomass has resulted following higher N application rates due to the fungi being more responsive to N than to C [46]. Compost addition to paddy soils resulted in a shift in microbial community structure, specifically increased amounts of actinomycetes and Gram-positive bacteria [47]. On the other hand, in a conventionally cultivated soil, actinomycete fatty acid biomarkers were in greater abundance than in an organically managed soil, which may be due to the role of actinomycetes as decomposers of nutrient-poor C compounds, and they may thus appear more active in the growing season when N becomes limited in the soil [48,49]. In contrast, others reported low fungal biomass when compared to microbial biomass, and no effect on fungal biomass by different types of organic fertilizer such as vermicompost or manure [50].
Cyanobacteria have been reported to influence the microbial community of tropical alluvial soil under both unsterilized and sterilized conditions [51]. In this study, the cyano-fertilizer and Neptune hydrolyzed fish emulsion increased microbial biomass and altered microbial community structure as compared to other fertilizers and the control. Cyano-fertilizer and Neptune hydrolyzed fish emulsion are liquid treatments that contain abundant soluble C and N; in addition, the cyano-fertilizer contains living cyanobacteria that can fix C and N. Other studies have also reported that organic manure and biofertilizer, organic nutrient management, chemical nutrient management, and integrated nutrient management increased soil microbial biomass shortly after application [52]. Organic fertilizers generally provide C, N, and energy to soil microbes for their growth and reproduction, and organic fertilizer effects are typically longer lasting than chemical fertilizer effects [53], although this study demonstrates that certain organic fertilizers (e.g., cyano-fertilizer and Neptune hydrolyzed fish emulsion) can result in short-term increases in soil bacterial, fungal, and total microbial biomass more so than other organic fertilizers. Heterocyst cells containing characteristic glycolipids produced by N-fixing cyanobacteria were observed, and the chemical structures of major components of Anabaena cylindrica are 3,25-Dihydroxyhexacosanyl α-D-glycopyranoside and 3,25,27-Trihydroxyoctacosanyl α-D-glycopyranoside [54,55]. In general, the fatty acid composition of planktonic Anabaena includes 14:0, 16:0, 16:1(cis-), 18:0, 18:2, 16:1 ω9, and 18:1 ω 7 [56,57,58]. If cyanobacteria survived in soil after irrigation, these organisms may have contributed to EL-FAMEs measured in this study, and subsequently microbial community structural differences among fertilizer treatments (Table 1 and Table 2). However, the presence of cyanobacteria alone does not fully explain microbial community changes, as soil microbial biomass and structure responded similarly between cyano-fertilizer and Neptune hydrolyzed fish emulsion in the cucumber experiment.

4.2. Microbial Community Groups under Two Different Depths in Cucumber and Peach Soils

To our knowledge, we are the first to report that cyano-fertilizer or Neptune hydrolyzed fish emulsion can affect the biomass and structure of soil microbial communities, and this effect was statistically significant at the 0–2.5 cm depth in cucumber field soil and the 2.5–7.5 cm depth in peach orchard soil. Overall, cyano-fertilizer increased total microbial biomass in both soils (cucumber plots and peach orchard), with the increase being attributed mainly to bacteria and actinomycetes in the cucumber soil, and AM fungi and total fungi in the peach orchard soil. The effect on microbial biomass was consistent despite large differences in overall microbial biomass between the cucumber and peach orchard soils, where biomass differences were presumably due to differences in soil type and management (annually cropped, tilled system in cucumber production vs. perennial, no-till system in peach orchard production). The difference in microbial biomass between the two experiments is in agreement with others [59] who found that the microbial biomass in arable soil was mainly influenced by soil type and short-term management. Furthermore, effects of cyano-fertilizer in the peach orchard soil were more heterogenous across plots than in the cucumber soil and inconsistent over time, as evidenced by the less distinguished separation of microbial community EL-FAME profiles in 2014 and lack of significant effects in the second year. This more subtle effect of cyano-fertilizer in the peach orchard soil could be due to the lack of soil mixing (due to no-till management), a relatively high microbial biomass that made smaller changes difficult to detect, and non-uniform growth of fungi and AM fungi in soil.
Our findings that both bacterial abundance and microbial biomass decreased with soil depth, particularly in the cucumber soil, are in agreement with other scientific papers [58]. Microbial biomass and CO2 respiration were greater in the top 5 cm depth of an upland grassland soil and decreased in the deeper 15 cm depth with change in the diversity due to the primary source of nutrient input in grasslands being above ground [58]. In addition, a study in a floodplain found that land use exerts strong effects on soil microbes in the topsoil and that microbial biomass and activity decrease with soil depth [59]. In this study, the shallow soil depth (0–2.5 cm) had greater microbial biomass compared with the deeper depth (2.5–7.5 cm) in the cucumber experiment. Within the no-till peach orchard, however, there was no consistent pattern of microbial biomass being greater in the surface than in the subsurface depth, because the cyano-fertilizer addition in the LM+C treatment stimulated microbial biomass to such an extent in the first year, that the depth effect was mostly negated. This may have occurred if the lower application rate of poultry manure allowed more of the cyano-fertilizer to move more deeply into the soil, so that it had a greater effect on microbial communities at the deeper depth than observed in the HM+C treatment. In another study, manure application increased microbial biomass in the 5–10 cm depth due to the movement of soluble C below the 0–5 cm depth [24]. Similarly, in a grassland soil during the summer, changes in the microbial community in the 5–10 and 10–15 cm depths were found to be influenced by rhizosphere effects, such as increased root exudation at the height of plant growth [59].

5. Conclusions

When possible, it would be beneficial to choose organic fertilizers that not only increase crop yield but also promote soil microbial biomass, as this could promote the sustainable management of agroecosystems. This study demonstrated that organic fertilizer decisions can influence soil microbial communities even in short periods of time (<1 year). Bacteria and actinomycetes were the most sensitive to cyano-fertilizer and Neptune hydrolyzed fish emulsion in the annually cropped cucumber system, whereas fungi and AM fungi responded positively (in the first year) to cyano-fertilizer in the perennial peach orchard. These specific shifts in biomass and community structure could be beneficial to organic vegetable and fruit production, as increased bacterial and actinomycete biomass could stimulate decomposition and nutrient turnover for vegetables, and increased fungal and AM fungal biomass could provide greater soil C storage, nutrient cycling, and symbiotic relationships with orchard trees. Furthermore, this study demonstrated that cyano-fertilizer could reduce grower reliance on commercial organic fertilizers or reduce application rates of manure when used as a supplemental fertilizer. Either scenario benefits growers by reducing dependence on commercial fertilizers, conserving primary fertilizer sources, and reducing environmental impacts associated with organic fertilizer production, transportation, and excessive application. Thus, cyano-fertilizer technology and application should be further explored and used by growers, due to its potential impact on soil microbial biomass and fungi for improved soil health in addition to its benefits to plant production and the environment.

Author Contributions

Conceptualization—A.A. and J.G.D.; writing (original draft preparation)—A.A., H.S. and J.G.D.; writing (review and editing)—A.A., M.S. and J.G.D.; visualization—A.A.; supervision—J.G.D.; technical assistance—H.S., A.W. and D.G.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the USDA Western Sustainable Agriculture Research and Education program project #SW14-023.

Institutional Review Board Statement

Not applicable.

Data Availability Statement

Data are available upon request.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Dixon, M.; Rohrbaugh, C.; Afkairin, A.; Vivanco, J. Impacts of the green revolution on rhizosphere microbiology related to nutrient acquisition. Appl. Microbiol. 2022, 2, 992–1003. [Google Scholar] [CrossRef]
  2. Abed, R.M.M.; Dobretsov, S.; Sudesh, K. Applications of cyanobacteria in biotechnology. J. Appl. Microbiol. 2009, 106, 1–12. [Google Scholar] [CrossRef] [PubMed]
  3. Crecchio, C.; Curci, M.; Mininni, R.; Ricciuti, P.; Ruggiero, P. Short-term effects of municipal solid waste compost amendments on soil carbon and nitrogen content, some enzyme activities and genetic diversity. Biol. Fertil. Soils 2001, 34, 311–318. [Google Scholar] [CrossRef]
  4. Ebhin Masto, R.; Chhonkar, P.K.; Singh, D.; Patra, A.K. Changes in soil biological and biochemical characteristics in a long-term field trial on a sub-tropical inceptisol. Soil Biol. Biochem. 2006, 38, 1577–1582. [Google Scholar] [CrossRef]
  5. Kidd, J.; Manning, P.; Simkin, J.; Peacock, S.; Stockdale, E. Impacts of 120 years of fertilizer addition on a temperate grassland ecosystem. PLoS ONE 2017, 12, e0174632. [Google Scholar] [CrossRef] [PubMed]
  6. Zhang, Y.; Hao, X.; Alexander, T.W.; Thomas, B.W.; Shi, X.; Lupwayi, N.Z. Long-term and legacy effects of manure application on soil microbial community composition. Biol. Fertil. Soils 2018, 54, 269–283. [Google Scholar] [CrossRef]
  7. Franco-Otero, V.G.; Soler-Rovira, P.; Hernández, D.; López-de-Sá, E.G.; Plaza, C. Short-term effects of organic municipal wastes on wheat yield, microbial biomass, microbial activity, and chemical properties of soil. Biol. Fertil. Soils 2011, 48, 205–216. [Google Scholar] [CrossRef]
  8. Marschner, P.; Kandeler, E.; Marschner, B. Structure and function of the soil microbial community in a long-term fertilizer experiment. Soil Biol. Biochem. 2003, 35, 453–461. [Google Scholar] [CrossRef]
  9. Yoder, N.; Davis, J.G. Organic fertilizer comparison on growth and nutrient content of three kale cultivars. HortTechnology 2020, 30, 176–184. [Google Scholar] [CrossRef]
  10. Barminski, R.; Storteboom, H.; Davis, J.G. Development and evaluation of an organically certifiable growth medium for cultivation of cyanobacteria. J. Appl. Phycol. 2016, 28, 2623–2630. [Google Scholar] [CrossRef]
  11. Wolde, G.; Asmamaw, M.; Sido, M.Y.; Yigrem, S.; Wolde-meskel, E.; Chala, A.; Storteboom, H.; Davis, J.G. Optimizing a cyanobacterial biofertilizer manufacturing system for village-level production in Ethiopia. J. Appl. Phycol. 2020, 32, 3983–3994. [Google Scholar] [CrossRef]
  12. Afkairin, A.; Ippolito, J.A.; Stromberger, M.; Davis, J.G. Solubilization of organic phosphorus sources by cyanobacteria and a commercially available bacterial consortium. Appl. Soil Ecol. 2021, 162, 103900. [Google Scholar] [CrossRef]
  13. Asmamaw, M.; Wolde, G.; Yohannes, M.; Yigrem, S.; Woldemeskel, E.; Chala, A.; Davis, J.G. Comparison of cyanobacterial bio-fertilizer with urea on three crops and two soils of Ethiopia. Afr. J. Agric. Res. 2019, 14, 588–596. [Google Scholar] [CrossRef]
  14. Toonsiri, P.; Del Grosso, S.J.; Sukor, A.; Davis, J.G. Greenhouse Gas Emissions from solid and liquid organic fertilizers applied to lettuce. J. Environ. Qual. 2016, 45, 1812–1821. [Google Scholar] [CrossRef] [PubMed]
  15. Erwiha, G.M.; Ham, J.; Sukor, A.; Wickham, A.; Davis, J.G. Organic fertilizer source and application method impact ammonia volatilization. Commun. Soil Sci. Plant Anal. 2020, 51, 1469–1482. [Google Scholar] [CrossRef]
  16. Dhuldhaj, U.; Pandya, U. Implementation of biofortification technology by using PGPR for sustainable agricultural production. Agric. Important Microbes Sustain. Agric. 2017, 2, 63–79. [Google Scholar] [CrossRef]
  17. Kumar, M.; Singh, D.P.; Prabha, R.; Sharma, A.K. Role of Cyanobacteria in nutrient cycle and use efficiency in the soil. In Nutrient Use Efficiency: From Basics to Advances; Springer: New Delhi, India, 2015; pp. 163–171. [Google Scholar] [CrossRef]
  18. Wenz, J.; Davis, J.G.; Storteboom, H. Influence of light on endogenous phytohormone concentrations of a nitrogen-fixing anabaena Sp. cyanobacterium culture in open raceways for use as fertilizer for horticultural crops. J. Appl. Phycol. 2019, 31, 3371–3384. [Google Scholar] [CrossRef]
  19. Sukor, A.; Amer, F.S.M.; Vanamala, J.; Davis, J.G. Phytohormones in organic fertilizers influence β-carotene concentration and marketable yield of lettuce (Lactuca sativa). Acta Hortic. 2022, 1348, 15–22. [Google Scholar] [CrossRef]
  20. Schutter, M.E.; Dick, R.P. Comparison of fatty acid methyl ester (FAME) methods for characterizing microbial communities. Soil Sci. Soc. Am. J. 2000, 64, 1659–1668. [Google Scholar] [CrossRef]
  21. Li, C.; Cano, A.; Acosta-Martinez, V.; Veum, K.S.; Moore-Kucera, J.A. Comparison between fatty acid methyl ester profiling methods (PLFA and EL-FAME) as soil health indicators. Soil Sci. Soc. Am. J. 2020, 84, 1153–1169. [Google Scholar] [CrossRef]
  22. Stromberger, M.; Shah, Z.; Westfall, D. Soil microbial communities of no-till dryland agroecosystems across an evapotranspiration Gradient. Appl. Soil Ecol. 2007, 35, 94–106. [Google Scholar] [CrossRef]
  23. Peacock, A.; Mullen, M.; Ringelberg, D.B.; Tyler, D.D.; Hedrick, D.B.; Gale, P.M.; White, D.C. Soil microbial community responses to dairy manure or ammonium nitrate applications. Soil Biol. Biochem. 2001, 33, 1011–1019. [Google Scholar] [CrossRef]
  24. Balkwill, D.L.; Leach, F.R.; Wilson, J.T.; Mcnabb, J.F.; White, D.C. Equivalence of Microbial biomass measures based on membrane lipid and cell wall components, adenosine triphosphate, and direct counts in subsurface aquifer sediments. Ecotoxicol. Environ. Saf. 1988, 16, 73–84. [Google Scholar] [CrossRef]
  25. Guckert, J.B.; Ringelberg, D.B.; White, D.C.; Hanson, R.S.; Bratina, B.J. Membrane fatty acids as phenotypic markers in the polyphasic taxonomy of methylotrophs within the proteobacteria. Microbiology 1991, 137, 2631–2641. [Google Scholar] [CrossRef] [PubMed]
  26. Vestal, J.R.; White, D.C. Lipid Analysis in Microbial Ecology. BioScience 1989, 39, 535–541. [Google Scholar] [CrossRef]
  27. Steger, K.; Jarvis, Å.; Smårs, S.; Sundh, I. Comparison of signature lipid methods to determine microbial community structure in compost. J. Microbiol. Methods 2003, 55, 371–382. [Google Scholar] [CrossRef]
  28. Schutter, M.; Sandeno, J.; Dick, R. Seasonal, Soil type, and alternative management influences on microbial communities of vegetable cropping systems. Biol. Fertil. Soils 2001, 34, 397–410. [Google Scholar] [CrossRef]
  29. Hsu, S.-F.; Buckley, D.H. Evidence for the functional significance of diazotroph community structure in soil. ISME J. 2008, 3, 124–136. [Google Scholar] [CrossRef]
  30. Bending, G.D.; Turner, M.K.; Jones, J.E. Interactions between crop residue and soil organic matter quality and the functional diversity of soil microbial communities. Soil Biol. Biochem. 2002, 34, 1073–1082. [Google Scholar] [CrossRef]
  31. Amador, J.A.; Atoyan, J.A. Structure and composition of leach field bacterial communities: Role of soil texture, depth and septic tank effluent inputs. Water 2012, 4, 707–719. [Google Scholar] [CrossRef]
  32. Fierer, N.; Schimel, J.P.; Holden, P.A. Variations in microbial community composition through two soil depth Profiles. Soil Biol. Biochem. 2003, 35, 167–176. [Google Scholar] [CrossRef]
  33. Wickham, A.; Davis, G.J. Optimizing organic carrot (Daucus carota var. sativus) yield and quality using fish emulsions, cyanobacterial fertilizer, and seaweed extracts. Agronomy 2023, 13, 1329. [Google Scholar] [CrossRef]
  34. Soil Science|Natural Resources Conservation Service. Available online: https://www.nrcs.usda.gov/conservation-basics/natural-resource-concerns/soil/soil-science (accessed on 16 August 2023).
  35. Sterle, D.G.; Stonaker, F.; Ela, S.; Davis, J.G. Cyanobacterial biofertilizer as a supplemental fertilizer for peaches: Yield, trunk growth, leaf nutrients and chlorosis. J. Am. Pomol. Soc. 2021, 75, 165–175. [Google Scholar]
  36. Frostegard, A.; Tunlid, A.; Baath, E. Phospholipid fatty acid composition, biomass, and activity of microbial communities from two soil types experimentally exposed to different heavy metals. Appl. Environ. Microbiol. 1993, 59, 3605–3617. [Google Scholar] [CrossRef] [PubMed]
  37. Gomez, J.D.; Denef, K.; Stewart, C.E.; Zheng, J.; Cotrufo, M.F. Biochar addition rate influences soil microbial abundance and activity in temperate soils. Eur. J. Soil Sci. 2014, 65, 28–39. [Google Scholar] [CrossRef]
  38. Denef, K.; Roobroeck, D.; Manimel Wadu, M.C.W.; Lootens, P.; Boeckx, P. Microbial community composition and rhizodeposit-carbon assimilation in differently managed temperate grassland soils. Soil Biol. Biochem. 2009, 41, 144–153. [Google Scholar] [CrossRef]
  39. Grogan, D.W.; Cronan, J.E., Jr. Cyclopropane ring formation in membrane lipids of bacteria. Microbiol. Mol. Biol. Rev. 1997, 61, 429–441. [Google Scholar]
  40. Lenth, R.; Singmann, H.; Love, J.; Buerkner, P.; Herve, M. Emmeans: Estimated Marginal Means, Aka Least-Squares Means (Version 1.3.4). Emmeans Estim Marg Means Aka Least-Sq Means. 2019. Available online: https://CRAN.R-project.org/package=emmeans (accessed on 10 April 2019).
  41. Esaki, K.; Ichinose, Y.; Yamada, S. Statistical analysis of process monitoring data for software process improvement and its application. Am. J. Oper. Res. 2012, 2, 43–50. [Google Scholar] [CrossRef]
  42. Frey, S.D.; Elliott, E.T.; Paustian, K. Bacterial and fungal abundance and biomass in conventional and no-tillage agroecosystems along two climatic gradients. Soil Biol. Biochem. 1999, 31, 573–585. [Google Scholar] [CrossRef]
  43. Wienhold, B.; Halvorson, A. Cropping system influences on several soil quality attributes in the northern great Plains. J. Soil Water Conserv. 1998, 53, 254–258. [Google Scholar]
  44. Yevdokimov, I.; Gattinger, A.; Buegger, F.; Munch, J.C.; Schloter, M. Changes in microbial community structure in soil as a result of different amounts of nitrogen fertilization. Biol. Fertil. Soils 2008, 44, 1103–1106. [Google Scholar] [CrossRef]
  45. Kimura, M.; Asakawa, S. Comparison of community structures of microbiota at main habitats in rice field ecosystems based on phospholipid fatty acid analysis. Biol. Fertil. Soils 2005, 43, 20–29. [Google Scholar] [CrossRef]
  46. Lundquist, E.J.; Scow, K.M.; Jackson, L.E.; Uesugi, S.L.; Johnson, C.R. Rapid response of soil microbial communities from conventional, low input, and organic farming systems to a wet/dry cycle. Soil Biol. Biochem. 1999, 31, 1661–1675. [Google Scholar] [CrossRef]
  47. MacKenzie, M.D.; Quideau, S.A. Microbial Community structure and nutrient availability in oil sands reclaimed boreal soils. Appl. Soil Ecol. 2010, 44, 32–41. [Google Scholar] [CrossRef]
  48. Lazcano, C.; Gómez-Brandón, M.; Revilla, P.; Domínguez, J. Short-term effects of organic and inorganic fertilizers on soil microbial community structure and function. Biol. Fertil. Soils 2012, 49, 723–733. [Google Scholar] [CrossRef]
  49. Ranjan, K.; Priya, H.; Ramakrishnan, B.; Prasanna, R.; Venkatachalam, S.; Thapa, S.; Tiwari, R.; Nain, L.; Singh, R.; Shivay, Y.S. Cyanobacterial inoculation modifies the rhizosphere microbiome of rice planted to a tropical alluvial soil. Appl. Soil Ecol. 2016, 108, 195–203. [Google Scholar] [CrossRef]
  50. Dinesh, R.; Srinivasan, V.; Hamza, S.; Manjusha, A. Short-term incorporation of organic manures and biofertilizers influences biochemical and microbial characteristics of soils under an annual crop [Turmeric (Curcuma longa L.)]. Bioresour. Technol. 2010, 101, 4697–4702. [Google Scholar] [CrossRef]
  51. Zeng, L.S.; Liao, M.; Chen, C.L.; Huang, C.Y. Effects of lead contamination on soil enzymatic activities, microbial biomass, and rice physiological indices in soil-lead-rice (Oryza sativa L.) system. Ecotoxicol. Environ. Saf. 2007, 67, 67–74. [Google Scholar] [CrossRef]
  52. Murate, N.; Nishida, I. Lipids of blue-Green algae (Cyanobacteria). Lipids Struct. Funct. 1987, 9, 315–347. [Google Scholar] [CrossRef]
  53. Gunstone, F.D.; Harood, J.L. The Lipid Handbook, 2nd ed.; Chapman & Hall: London, UK, 2021; pp. 1–1273. Available online: https://books.google.com/books?hl=en&lr=&id=INZa6WmqDA8C&oi=fnd&pg=PP1&dq=Gunstone,+F.D.+and+Harwood,+J.L.,+2007.+The+lipid+handbook+with+CD-ROM.+CRC+press.&ots=Sv3z6c8Ay1&sig=ueCDpgrIFv7RluBvF14sfdumHR0#v=onepage&q=Gunstone%2CF.D.andHarwood%2CJ.L.%2C (accessed on 27 July 2021).
  54. Willers, C.; Jansen van Rensburg, P.J.; Claassens, S. Phospholipid fatty acid profiling of microbial communities–a review of interpretations and recent applications. J. Appl. Microbiol. 2015, 119, 1207–1218. [Google Scholar] [CrossRef]
  55. Gugger, M.; Lyra, C.; Suominen, I.; Tsitko, I.; Humbert, J.-F.; Salkinoja-Salonen, M.S.; Sivonen, K. Cellular fatty acids as chemotaxonomic markers of the genera anabaena, aphanizomenon, microcystis, nostoc and planktothrix (cyanobacteria). Int. J. Syst. Evol. Microbiol. 2002, 52, 1007–1015. [Google Scholar] [CrossRef]
  56. Li, R.; Watanabe, M.M. Fatty Acid profiles and their chemotaxonomy in planktonic species of anabaena (cyanobacteria) with straight trichomes. Phytochemistry 2001, 57, 727–731. [Google Scholar] [CrossRef]
  57. Girvan, M.S.; Bullimore, J.; Pretty, J.N.; Osborn, A.M.; Ball, A.S. Soil type is the primary determinant of the composition of the total and active bacterial communities in arable soils. Appl. Environ. Microbiol. 2003, 69, 1800–1809. [Google Scholar] [CrossRef]
  58. Bardgett, R.D.; Leemans, D.K.; Cook, R.; Hobbs, P.J. Seasonality of the soil biota of grazed and ungrazed hill grasslands. Soil Biol. Biochem. 1997, 29, 1285–1294. [Google Scholar] [CrossRef]
  59. Griffiths, R.I.; Whiteley, A.S.; O’Donnell, A.G.; Bailey, M.J. Influence of depth and sampling time on bacterial community structure in an upland grassland soil. FEMS Microbiol. Ecol. 2003, 43, 35–43. [Google Scholar] [CrossRef] [PubMed]
Agriculture 13 01902 g001
Figure 1. Principal component analysis (PCA) of soil microbial community EL-FAMEs from the 2015 cucumber experiment, sampled at two different depths: 0–2.5 cm depth (1) and 2.5–7.5 cm depth (2). Fertilizer treatments are indicated as follows: CY = Cyano-fertilizer, N = Neptune hydrolyzed fish emulsion, A = Alaska non-hydrolyzed fish emulsion, BA = Blood meal surface applied, BB = Blood meal sub-surface applied, C = Control, FA = Feather meal surface applied, and FB = Feather meal sub-surface applied. The percent variance explained by each PC is shown in parentheses.
Figure 1. Principal component analysis (PCA) of soil microbial community EL-FAMEs from the 2015 cucumber experiment, sampled at two different depths: 0–2.5 cm depth (1) and 2.5–7.5 cm depth (2). Fertilizer treatments are indicated as follows: CY = Cyano-fertilizer, N = Neptune hydrolyzed fish emulsion, A = Alaska non-hydrolyzed fish emulsion, BA = Blood meal surface applied, BB = Blood meal sub-surface applied, C = Control, FA = Feather meal surface applied, and FB = Feather meal sub-surface applied. The percent variance explained by each PC is shown in parentheses.
Agriculture 13 01902 g001
Agriculture 13 01902 g002
Figure 2. Principal component analysis (PCA) of soil microbial community EL-FAMEs from the 2014 peach orchard experiment, sampled at two different depths: 0–2.5 cm depth (1) and 2.5–7.5 cm depth (2). Fertilizer treatments are indicated as follows: HM = 100 kg N ha−1 of dried poultry manure, HM+C = 100 kg N ha−1 of dried poultry manure + 25 kg N ha−1 of cyano-fertilizer, LM+C = 75 kg N ha−1 as dried poultry manure + 25 kg N ha−1 of cyano-fertilizer. The percent variance explained by each PC is shown in parentheses.
Figure 2. Principal component analysis (PCA) of soil microbial community EL-FAMEs from the 2014 peach orchard experiment, sampled at two different depths: 0–2.5 cm depth (1) and 2.5–7.5 cm depth (2). Fertilizer treatments are indicated as follows: HM = 100 kg N ha−1 of dried poultry manure, HM+C = 100 kg N ha−1 of dried poultry manure + 25 kg N ha−1 of cyano-fertilizer, LM+C = 75 kg N ha−1 as dried poultry manure + 25 kg N ha−1 of cyano-fertilizer. The percent variance explained by each PC is shown in parentheses.
Agriculture 13 01902 g002
Table 1. Eigenvector coefficients of microbial ester-linked fatty acid methyl esters (EL-FAMEs) for principal component (PC) axes 1 and 2 in the cucumber experiment in 2015, as shown in Figure 1.
Table 1. Eigenvector coefficients of microbial ester-linked fatty acid methyl esters (EL-FAMEs) for principal component (PC) axes 1 and 2 in the cucumber experiment in 2015, as shown in Figure 1.
Eigenvector Coefficient
Microbial GroupEL-FAMEPC1PC2
Actinomycetes10Me16:0−0.574−0.032
10Me17:0−0.9700.025
10Me18:0−0.9340.036
Fungi18:1ω90.6530.570
18:2ω60.200−0.568
AM Fungi16:1ω5−0.169−0.665
Gram-negative Bacteria16:1ω7−0142−0.536
17:1ω7−0.8240.068
17:0cy−0.9040.015
18:1ω7−0.080−0.206
18:1ω8−0.8990.013
19:0cy−0.4120.223
Gram-positive Bacteriai14:00.259−0.368
i15:00.001−0.668
a15:00.279−0.025
i16:0−0.156−0.156
i17:0−0.725−0.001
a17:0−0.755−0.024
Non-specific13:0−0.9370.043
14:0−0.8350.414
15:0−0.9650.021
16:00.8360.107
18:0−0.2450.273
Table 2. Eigenvector coefficients of microbial ester-linked fatty acid methyl esters (EL-FAMEs) for principal component (PC) axes 1 and 2, in the peach experiment in 2014 as shown in Figure 2.
Table 2. Eigenvector coefficients of microbial ester-linked fatty acid methyl esters (EL-FAMEs) for principal component (PC) axes 1 and 2, in the peach experiment in 2014 as shown in Figure 2.
Eigenvector Coefficient
Microbial GroupEL-FAMEPC1PC2
Actinomycetes10Me16:0−0.1030.697
10Me17:0−0.6340.587
10Me18:0−0.970−0.017
Fungi18:1ω90.436−0.300
18:2ω6 −0.235−0.706
AM Fungi16:1ω50.4300.695
Gram-negative Bacteria16:1ω7−0.459−0.607
17:1ω7−0.8830.115
17:0cy−0.101−0.482
18:1ω70.123−0.230
18:1ω8−0.6780.437
19:0cy−0.4420.331
Gram-positive Bacteriai14:00.623−0.688
i15:0−0.427−0.606
a15:00.029−0.948
i16:00.283−0.868
i17:0−0.833−0.181
a17:0−0.436−0.658
Non-specific13:0−0.9650.135
14:0−0.9760.029
15:0−0.959−0.021
16:00.8890.108
18:0−0.737−0.403
Table 3. Soil microbial biomass and community composition in a certified organic cucumber field receiving different organic fertilizer treatments. Within the 0–2.5 cm soil depth, fertilizer treatment means with the same letter are not significantly different at p < 0.05 using Tukey’s honest significant difference (HSD) test. There were no significant differences among fertilizer treatments in the 2.5–7.5 cm depth.
Table 3. Soil microbial biomass and community composition in a certified organic cucumber field receiving different organic fertilizer treatments. Within the 0–2.5 cm soil depth, fertilizer treatment means with the same letter are not significantly different at p < 0.05 using Tukey’s honest significant difference (HSD) test. There were no significant differences among fertilizer treatments in the 2.5–7.5 cm depth.
Treatment 1DepthTotal
Microbial
Biomass		Gram+
Bacteria		Gram−
Bacteria		FungiActinomycetes
nmol g−1
A 0–2.537.3 bc5.02 ab 7.34 bc12.7 a0.60 bc
2.5–7.528.64.214.858.711.46
BA0–2.524.2 c3.34 b3.38 c8.33 a0.74 bc
2.5–7.518.51.862.87 4.900.00
BB0–2.539.8 bc5.73 ab6.61 bc14.0 a0.59 bc
2.5–7.522.63.413.599.210.69
C0–2.527.8 c4.50 ab4.44 c9.80 a0.39 bc
2.5–7.528.54.585.219.030.95
CY0–2.568.1 a10.5 a15.1 ab17.7 a3.12 ab
2.5–7.538.26.418.819.982.35
FA0–2.531.6 c5.29 ab6.16 bc10.9 a0.00 c
2.5–7.59.031.231.523.190.00
FB0–2.526.5 c3.85 b5.33 bc9.74 a0.00 c
2.5–7.516.32.732.435.520.00
N0–2.573.0 a11.1 a17.5 a20.2 a4.23 a
2.5–7.538.56.338.3610.22.59
1 Fertilizer treatments are indicated as follows: A = Alaska non-hydrolyzed fish emulsion, BA = Blood meal surface applied, BB = Blood meal sub-surface applied, C = Control, CY = Cyano-fertilizer, FA = Feather meal surface applied, FB = Feather meal sub-surface applied, N = Neptune hydrolyzed fish emulsion. Treatments were replicated four times.
Table 4. Soil microbial biomass and community composition in a certified organic peach orchard receiving different organic fertilizer treatments in 2014. Within the 2.5–7.5 cm depth, fertilizer treatment means with the same letter are not significantly different at p < 0.05 using Tukey’s honest significant difference (HSD) test. There were no significant differences among treatments in the 0–2.5 cm depth.
Table 4. Soil microbial biomass and community composition in a certified organic peach orchard receiving different organic fertilizer treatments in 2014. Within the 2.5–7.5 cm depth, fertilizer treatment means with the same letter are not significantly different at p < 0.05 using Tukey’s honest significant difference (HSD) test. There were no significant differences among treatments in the 0–2.5 cm depth.
Treatment 1DepthTotal
Microbial Biomass		Gram+
Bacteria		Gram− BacteriaFungiAM FungiActinomycetes
nmol g−1
HM+C0–2.553192.0117175.046.223.9
2.5–7.5368 ab56.0 a78.7ab118 ab43.3 ab27.7 a
LM+C0–2.547292.010615448.824.9
2.5–7.5490 a71.8 a108a162 a64.9 a33.7 a
HM0–2.549987.611716245.222.5
2.5–7.5321 b51.1 a71.2b94.5 b32.5 b25.4 a
1 Fertilizer treatments are indicated as follows: HM+C = 100 kg N ha−1 as dried poultry manure + 25 kg N ha−1 of cyano-fertilizer, LM+C = 75 kg N ha−1 as dried poultry manure + 25 kg N ha−1 as cyano-fertilizer, HM = 100 kg N ha−1 of dried poultry manure. Treatments were replicated five times.
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

Afkairin, A.; Stromberger, M.; Storteboom, H.; Wickham, A.; Sterle, D.G.; Davis, J.G. Soil Microbial Community Responses to Cyanobacteria versus Traditional Organic Fertilizers. Agriculture 2023, 13, 1902. https://doi.org/10.3390/agriculture13101902

AMA Style

Afkairin A, Stromberger M, Storteboom H, Wickham A, Sterle DG, Davis JG. Soil Microbial Community Responses to Cyanobacteria versus Traditional Organic Fertilizers. Agriculture. 2023; 13(10):1902. https://doi.org/10.3390/agriculture13101902

Chicago/Turabian Style

Afkairin, Antisar, Mary Stromberger, Heather Storteboom, Allison Wickham, David G. Sterle, and Jessica G. Davis. 2023. "Soil Microbial Community Responses to Cyanobacteria versus Traditional Organic Fertilizers" Agriculture 13, no. 10: 1902. https://doi.org/10.3390/agriculture13101902

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