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Search Results (391)

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16 pages, 11337 KiB  
Article
Renewal and Iteration Mechanisms of Aged Tea Trees: Insights from Tea Garden Soil Microbial Communities
by Houqiao Wang, Tianyu Wu, Wenxia Yuan, Lijiao Chen, Hongxu Li, Xiujuan Deng, Chun Wang, Weihao Liu, Wei Huang and Baijuan Wang
Agronomy 2024, 14(12), 2955; https://doi.org/10.3390/agronomy14122955 - 12 Dec 2024
Viewed by 457
Abstract
This study focuses on the renewal and iteration mechanisms of aged tea trees in interactions with their soil microbial communities, aiming to elucidate the impact of the planting age of tea trees on the structure and function of soil microbial communities and how [...] Read more.
This study focuses on the renewal and iteration mechanisms of aged tea trees in interactions with their soil microbial communities, aiming to elucidate the impact of the planting age of tea trees on the structure and function of soil microbial communities and how these impacts are linked to the formation of tea quality. By conducting a comparative analysis of the cultivation soil from tea trees with varying planting ages ranging from 30 to 200 years, we employed microbial diversity sequencing, a soil physicochemical property analysis, and tea leaf chemical component detection. We combined these methods with redundancy analysis (RDA) and linear discriminant analysis effect size (LEfSe) to reveal significant correlations between the planting age of tea trees and the soil’s microbial diversity and function. The results indicate that as the planting age of tea trees increases, there are significant changes in the soil’s pH and nutrient content. Concurrently, the components of the tea leaves also change. Most notably, around the 120 years mark of the tea tree planting age, the diversity of the soil microbial community reaches a turning point. Key microbial community analyses revealed shifts in the dominant microbial populations within the soil across the various tea tree planting ages, exemplified by taxa such as Hygrocybe Mycena, Humicola, Bradyrhizobium, and Candidatus Solibacter. These alterations in microbial communities are closely associated with soil nutrient dynamics and the developmental stages of tea trees. These findings not only provide scientific guidance for tea garden management, tea tree cultivation, and tea production but also offer new insights into the impact of tea tree–soil–microbe interactions on tea quality, which is significantly important for enhancing tea quality. Full article
(This article belongs to the Topic Plant-Soil Interactions, 2nd Volume)
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Figure 1
<p>Tea plants at different planting ages and sampling locations.</p>
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<p>Variations in the soil pH (<b>a</b>), total nitrogen (<b>b</b>), total phosphorus (<b>c</b>), total potassium (<b>d</b>), alkali-hydrolyzable nitrogen (<b>e</b>), available phosphorus (<b>f</b>), available potassium (<b>g</b>), and soil organic matter (<b>h</b>) in tea tree cultivation soil with different planting ages.</p>
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<p>Changes in the water extract (WE) (<b>a</b>), tea polyphenol (TP) (<b>b</b>), amino acid (AA) (<b>c</b>), nitrogen (N) (<b>d</b>), phosphorus (P) (<b>e</b>), and potassium (K) (<b>f</b>) content in tea leaves from tea trees with different planting ages.</p>
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<p>The impact of the tea tree planting age on the alpha diversity of soil fungal and bacterial communities (<b>a</b>,<b>b</b>), and the PCoA showing the distribution of soil fungal and bacterial communities across the different planting ages (<b>c</b>,<b>d</b>); * indicates significant difference, <span class="html-italic">p</span> &lt; 0.05, ** indicates significant difference, <span class="html-italic">p</span> &lt; 0.01, no marking indicates no difference.</p>
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<p>Dendrogram of soil fungal and bacterial community structures, based on the UPGMA method, with (<b>a</b>) for fungi and (<b>b</b>) for bacteria.</p>
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<p>Distribution of the top 10 fungal (<b>a</b>) and bacterial (<b>b</b>) phyla in tea garden soil at different planting ages.</p>
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<p>LEfSe analysis depicting the evolutionary cladogram of the abundance changes of characteristic microbial fungi (<b>a</b>) and bacteria (<b>b</b>) in tea garden soil across the different planting ages.</p>
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<p>Functional analysis of key microbial communities in tea garden soil with different tea tree planting ages, with (<b>a</b>) representing fungi and (<b>b</b>) representing bacteria.</p>
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<p>RDA of the relationship between soil microbial community structure and soil properties, with (<b>a</b>) depicting fungi and (<b>b</b>) depicting bacteria.</p>
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18 pages, 11993 KiB  
Article
Evaluating the Impact of Environmental Factors on Bacterial Populations in Riverine, Estuarine, and Coastal Sediments
by Ramganesh Selvarajan, Ming Yang, Henry J. O. Ogola, Timothy Sibanda and Akebe Luther King Abia
Diversity 2024, 16(12), 749; https://doi.org/10.3390/d16120749 - 6 Dec 2024
Viewed by 623
Abstract
Aquatic ecosystems, including rivers, estuaries, and coastal environments, are crucial for maintaining biodiversity, regulating nutrient cycles, and supporting human livelihoods. However, these ecosystems are increasingly being threatened by urbanization, making it essential to understand their microbial communities and their ecological roles. This study [...] Read more.
Aquatic ecosystems, including rivers, estuaries, and coastal environments, are crucial for maintaining biodiversity, regulating nutrient cycles, and supporting human livelihoods. However, these ecosystems are increasingly being threatened by urbanization, making it essential to understand their microbial communities and their ecological roles. This study employed high-throughput 16S rRNA gene sequencing to characterize the bacterial communities within the riverine, estuarine, and coastal sediments of Adyar Creek, Chennai, India. Proteobacteria were the dominant phylum across most samples, with proportions ranging from 39.65% to 72.09%. Notably, the estuarine environment exhibited a distinct taxonomic profile characterized by a significant abundance of Firmicutes (47.09% of the bacterial population). Distinct bacterial classes were observed across sediment types: Alphaproteobacteria (30.07–34.32%) in riverine sediments, Bacilli dominated estuarine sediments (40.17%), and Gammaproteobacteria (15.71–51.94%) in coastal sediments. The most significant environmental factors influencing the bacterial community composition across these samples were pH, salinity, phosphate, and nitrate. LEfSe (Linear discriminant analysis Effect Size) analysis identified specific genera within the estuary, including Bacillus (20.26%), unclassified_Paenibacillus (12.87%), Clostridium (3.81%), Gailella (3.17%), Paenibacillus (3.02%), Massilia (1.70%), Paraburkholderia (1.42%), and Pantoea (1.15%), as potential biomarkers for habitat health. Functional analysis revealed an elevated expression of the genes associated with ABC transporters and carbon metabolism in the estuary, suggesting a heightened nutrient cycling capacity. Furthermore, co-occurrence network analysis indicated that bacterial communities exhibit a strong modular structure with complex species interactions across the three sediment types. These findings highlight bacterial communities’ critical role and the key drivers in estuarine ecosystems, establishing a baseline for further investigations into the functional ecology of these vulnerable ecosystems. Full article
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<p>A map showing the sampling points within the Adyar Estuary Creek.</p>
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<p>Scatter plot analysis of toxic heavy metal concentrations in sediment samples from Adyar Estuary Creek: two-tailed Pearson correlation coefficients and 95% confidence ellipses for data distribution.</p>
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<p>Alpha and beta diversity analysis of sediment samples: (<b>a</b>) ACE, Chao1, Jackknife, and ASVs; (<b>b</b>) Shannon index; (<b>c</b>) Simpson index; (<b>d</b>) principal coordinate analysis (PcoA) of microbial community structure.</p>
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<p>Relative abundance of bacterial community composition in sediment samples: (<b>a</b>) phylum level distribution, (<b>b</b>) class level distribution, and (<b>c</b>) top 20 genus level distribution.</p>
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<p>Redundancy analysis (RDA) shows the first and second ordination axes of RDA explained, respectively. RDA explains the relationships between microbial community composition at the class level across various sediment sample groups and the principal environmental variables.</p>
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<p>Functional prediction between the sample groups: (<b>a</b>) major pathway distribution of collected sediment samples, (<b>b</b>) principal component analysis (PCA), (<b>c</b>) major degradation pathway, and (<b>d</b>) biosynthetic pathways.</p>
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<p>(<b>a</b>) Indicator bacterial members in the three groups (blue—coastal; green—riverine; and orange—estuarine) of sediment samples with LDA values higher than 3.5. (<b>b</b>) Co-network analysis of major bacterial members in the collected samples (red lines show the negative correlation, and green shows the positive correlation between the bacterial members).</p>
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14 pages, 3816 KiB  
Article
Comparative Study of Gut Microbiome in Urban and Rural Eurasian Tree Sparrows
by Shuai Yan, Yu Zhang, Ji Huang, Yingbao Liu and Shaobin Li
Animals 2024, 14(23), 3497; https://doi.org/10.3390/ani14233497 - 4 Dec 2024
Viewed by 542
Abstract
Gut microbiota play a significant role in various physiological functions, including digestion, nutritional metabolism, and host immune function. The composition of these gut microbes is largely influenced by habitats. This study examines the gut microbiota of the Eurasian tree sparrow (Passer montanus [...] Read more.
Gut microbiota play a significant role in various physiological functions, including digestion, nutritional metabolism, and host immune function. The composition of these gut microbes is largely influenced by habitats. This study examines the gut microbiota of the Eurasian tree sparrow (Passer montanus) inhabiting rural and urban environments to understand the effects of habitat variation on microbial composition. We captured 36 rural and 29 urban adult tree sparrows and observed minor differences in body mass but substantial differences in foraging microhabitats between the two groups. Fecal samples from adult males with similar body mass were selected for a gut microbiome analysis to mitigate potential confounding effects, resulting in 20 successfully sequenced samples. The analysis disclosed disparities in gut microbiota diversity and composition between rural and urban sparrows. The urban group demonstrated slightly higher alpha diversity and distinct dominant phyla and genera compared to the rural group. Additionally, differences in the relative abundance of potentially pathogenic bacteria were observed between the groups. Several potentially pathogenic bacteria (e.g., TM7, Staphylococcus, Helicobacter, and Shigella) were more abundant in the urban group, suggesting that tree sparrows may act as transmission vectors and develop stronger immune systems. This could potentially facilitate pathogen dissemination while also contributing to the natural cycling of nutrients and maintaining ecosystem health in urban environments. The beta diversity analysis confirmed structural differences in microbial communities, implicating habitat variation as a contributing factor. Furthermore, the LEfSe analysis emphasized significant differences in gut bacteria abundance (across two phyla, three classes, six orders, seven families, and eight genera) between urban and rural sparrows, with predicted functional differences in metabolic pathways. Notably, lipid metabolism was enriched in urban sparrows, indicating enhanced lipid synthesis and metabolism in urban habitats. In conclusion, this study underscores the profound influence of habitat on the gut microbiota composition and functional potential in tree sparrows. Our findings highlight that urbanization alters the gut microbes and, consequently, the physiological functions of bird species. Full article
(This article belongs to the Special Issue Birds Ecology: Monitoring of Bird Health and Populations, Volume II)
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<p>Sampling sites in satellite image from Baidu Map (left blue circle: rural site; right yellow circle: urban site).</p>
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<p>Frequencies of foraging events across two microhabitats in urban and rural settings (** indicates significant differences between groups).</p>
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<p>Venn diagrams showing number of shared and unique ASVs in rural group (A) and urban group (B).</p>
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<p>Characterization of the gut microbial community composition of each sample (A1–A10 represent rural samples and B1–B10 represent urban samples) at the phylum (<b>A</b>) and genus levels (<b>B</b>).</p>
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<p>The diversity (Shannon and Simpson index) and richness (Chao1 index and observed species) of the gut microbial communities of the <span class="html-italic">Passer montanus</span> (groups were compared using the Mann–Whitney U test).</p>
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<p>PCoA plot of samples using weighted (<b>A</b>) and unweighted (<b>B</b>) UniFrac distances between rural and urban groups.</p>
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<p>The LDA effect size of the gut microbial communities of the <span class="html-italic">Passer montanus</span>. The histogram shows the bacteria that were significantly greater and lower in the fecal samples of urban tree sparrows (<span class="html-italic">n</span> = 10) than rural tree sparrows (<span class="html-italic">n</span> = 10) in red and blue bars (LDA &gt; 2).</p>
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<p>The prediction of function using a PICRUSt analysis. The horizontal coordinates stand for the abundance of functional pathways (per million KOs), the vertical coordinates stand for the functional pathways at the second-level KEGG classification, and the rightmost column stands for the first-level pathways to which these pathways belong. The average abundance of all the samples is presented.</p>
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14 pages, 4490 KiB  
Article
Local Electric Field-Incorporated In-Situ Copper Ions Eliminating Pathogens and Antibiotic Resistance Genes in Drinking Water
by Ruiqing Li, Haojie Dai, Wei Wang, Rulin Peng, Shenbo Yu, Xueying Zhang, Zheng-Yang Huo, Qingbin Yuan and Yi Luo
Antibiotics 2024, 13(12), 1161; https://doi.org/10.3390/antibiotics13121161 - 2 Dec 2024
Viewed by 729
Abstract
Background/Objectives: Pathogen inactivation and harmful gene destruction from water just before drinking is the last line of defense to protect people from waterborne diseases. However, commonly used disinfection methods, such as chlorination, ultraviolet irradiation, and membrane filtration, experience several challenges such as continuous [...] Read more.
Background/Objectives: Pathogen inactivation and harmful gene destruction from water just before drinking is the last line of defense to protect people from waterborne diseases. However, commonly used disinfection methods, such as chlorination, ultraviolet irradiation, and membrane filtration, experience several challenges such as continuous chemical dosing, the spread of antibiotic resistance genes (ARGs), and intensive energy consumption. Methods: Here, we perform a simultaneous elimination of pathogens and ARGs in drinking water using local electric fields and in-situ generated trace copper ions (LEF-Cu) without external chemical dosing. A 100-μm thin copper wire placed in the center of a household water pipe can generate local electric fields and trace copper ions near its surface after an external low voltage is applied. Results: The local electric field rapidly damages the outer structure of microorganisms through electroporation, and the trace copper ions can effectively permeate the electroporated microorganisms, successfully damaging their nucleic acids. The LEF-Cu disinfection system achieved complete inactivation (>6 log removal) of Escherichia coli O157:H7, Pseudomonas aeruginosa PAO1, and bacteriophage MS2 in drinking water at 2 V for 2 min, with low energy consumption (10−2 kWh/m3). Meanwhile, the system effectively damages both intracellular (0.54~0.64 log) and extracellular (0.5~1.09 log) ARGs and blocks horizontal gene transfer. Conclusions: LEF-Cu disinfection holds promise for preventing horizontal gene transfer and providing safe drinking water for household applications. Full article
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<p>Deaths from unsafe sanitation. (<b>a</b>) Estimated annual number of deaths due to unsafe sanitation per 100,000 people. (<b>b</b>) The total death population from unsafe sanitation. Data source: Global Burden of Disease (2024).</p>
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<p>Disinfection performance of the LEF-Cu method. (<b>a</b>) Disinfection efficiency on <span class="html-italic">E. coli</span> O157:H7 at various voltages. (<b>b</b>) Disinfection efficiency on <span class="html-italic">P. aeruginosa</span> PAO1 at various voltages. (<b>c</b>) Disinfection efficiency on the virus MS2 at various voltages. Microorganisms are dosed in filtered tap water with a high concentration of 10<sup>6</sup> CFU/mL (bacteria) or PFU/mL (viruses). The applied voltages range from 0.2 to 3 V and HRT ranges from 0.5 to 3 min. Dashed lines indicate all the microorganisms were inactivated (i.e., live microorganisms were not detected). Error bars represent the standard deviation (<span class="html-italic">n</span> = 3).</p>
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<p>Investigation of disinfection mechanisms. (<b>a</b>) SEM images of <span class="html-italic">E</span>. <span class="html-italic">coli</span> O157:H7 before and after LEF-Cu disinfection. (<b>b</b>) TEM images of <span class="html-italic">E</span>. <span class="html-italic">coli</span> O157:H7 before and after LEF-Cu disinfection. (<b>c</b>) Fluorescence confocal images of PI-stained <span class="html-italic">E</span>. <span class="html-italic">coli</span> O157:H7 before and after LEF-Cu disinfection. Bacteria are dosed in filtered tap water with a high concentration of 10<sup>6</sup> CFU/mL. The HRT was fixed at 2 min. Red circle indicate the place of cell damage.</p>
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<p>Removal of ARGs using LEF-Cu disinfection. (<b>a</b>) Removal performance of iARGs. (<b>b</b>) Removal performance of eARGs. (<b>c</b>) HGT frequency after LEF-Cu disinfection using <span class="html-italic">E. coli</span> (K12) as a donor and <span class="html-italic">E. coli</span> (HB101) as a recipient. Bacteria are dosed in filtered tap water with a high concentration of 10<sup>6</sup> CFU/mL. The applied voltage was set as 2V for pictures (<b>a</b>) and (<b>b</b>). The HRT was fixed at 2 min for all three pictures. Error bars represent the standard deviation (<span class="html-italic">n</span> = 3).</p>
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<p>Contribution of Cu<sup>2+</sup> and local electric field to the LEF-Cu disinfection. (<b>a</b>) Cu<sup>2+</sup> release during LEF-Cu disinfection. (<b>b</b>) Toxicity evaluation of the released Cu<sup>2+</sup>. (<b>c</b>) Contribution of Cu<sup>2+</sup> and local electric field to disinfection. (<b>d</b>) Electric field simulation and schematic summary of the disinfection device mechanism.</p>
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<p>Disinfection efficacy of LEF-Cu method for treating tap water (<b>a</b>), lake water (<b>b</b>), and river water (<b>c</b>). <span class="html-italic">E. coli</span> H7:O157 was dosed in filtered water samples at 10<sup>6</sup> CFU/mL. Error bars represent the standard deviation (<span class="html-italic">n</span> = 3).</p>
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<p>(<b>a</b>) Schematic diagram showing the LEF-Cu disinfection device. (<b>b</b>) View of the LEF-Cu disinfection device.</p>
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15 pages, 4220 KiB  
Article
Temporal Changes in Faecal Microbiota Composition and Diversity in Dairy Cows Supplemented with a Lactobacillus-Based Direct-Fed Microbial
by Bronwyn E. Campbell, Mohammad Mahmudul Hassan, Robert J. Moore, Timothy Olchowy, Shahab Ranjbar, Martin Soust, Orlando Ramirez-Garzon, Rafat Al Jassim and John I. Alawneh
Animals 2024, 14(23), 3437; https://doi.org/10.3390/ani14233437 - 27 Nov 2024
Viewed by 471
Abstract
The rumen microbiota of dairy cows plays a crucial role in fermenting fibrous material, essential for nutrient extraction and overall productivity, detoxification of anti-nutritional toxic compounds, synthesis of vital nutrients, and is essential for optimal animal health. This study investigated the impact of [...] Read more.
The rumen microbiota of dairy cows plays a crucial role in fermenting fibrous material, essential for nutrient extraction and overall productivity, detoxification of anti-nutritional toxic compounds, synthesis of vital nutrients, and is essential for optimal animal health. This study investigated the impact of Lentilactobacillus-, Lactocaseibacillus-, and Lacticaseibacillus-based direct-fed microbial (DFM) supplementation on dairy cows’ faecal microbial composition and diversity. The study was carried out on a commercial dairy farm using 50 Holstein-Friesian cows randomly assigned into control (CON) and treatment (TRT) groups. Faecal samples were collected directly from the rectum every two months from September 2021 to January 2023. The bacterial 16S rRNA gene and fungal ITS-1 regions were amplified, sequenced, and analysed. Microbial diversity was assessed through alpha- and beta-diversity metrics. Linear discriminant analysis effect size (LEfSe) was performed to identify which taxa were driving the changes seen in the microbiota over time and treatment. Bacteroidaceae were the most prevalent bacterial family, followed by Lachnospiraceae and Muribaculaceae in both CON and TRT cows. Ascomycota, Basidiomycota, and Mucoromycota were the dominant three fungal phyla in the faeces of both CON and TRT cows. Bacterial genera Fructilactobacillus was abundant in the CON and Absicoccus in the TRT groups. Fungal taxa Chaetothryriales_incertae_sedis and Pseudomentella were absent in the faeces of TRT cows. Significant temporal and specific taxonomic differences were observed between the CON and TRT groups. The study’s findings underscore the dynamic nature of microbial communities and the importance of targeted dietary interventions. Further research is necessary to elucidate these microbial shifts, long-term impacts, and functional implications, aiming to optimise ruminant nutrition and enhance dairy cow performance. Full article
(This article belongs to the Section Cattle)
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<p>Relative abundance of bacterial families in faeces for control (CON) and DFM-treated (TRT) groups throughout the study period.</p>
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<p>Bacterial alpha diversity analysis (genus level) over time in faeces from Control (CON) and DFM-treated (TRT) experimental groups. Observed (<b>A</b>) and Chao1 (<b>B</b>) for Shannon index (<b>C</b>). Pooled data for overall differences between experimental groups disregarding the effect of the calendar month: Observed and Chao1, <span class="html-italic">p</span>-value: &lt;0.001, <span class="html-italic">t</span>-test: 46.5; Shannon, <span class="html-italic">p</span>-value: &lt;0.001, <span class="html-italic">t</span>-test: −25.2. Asterisk (*) on <span class="html-italic">x</span>-axis below experimental group denotes <span class="html-italic">p</span> &lt; 0.05 between experimental groups for a given calendar month.</p>
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<p>Bacterial beta-diversity (genus level) analysis via (<b>A</b>) principal component analysis and (<b>B</b>) non-metric multidimensional scaling of faeces across eight time points from September 2021 to January 2023. Microbial diversity differed significantly over the 16 months of the trial.</p>
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<p>Linear discriminant analysis effect size (LEfSe) analysis of the top twenty genera from faeces from Control (CON) compared with DFM-supplemented (TRT) experimental group. LDAscore refers to the linear discriminant analysis scores.</p>
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<p>Relative abundance of fungal classes in faeces for control (CON) and treatment (TRT) groups throughout the study period.</p>
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<p>Alpha-diversity analysis (genus-level) of the fungal composition of faeces over time from Control (CON) compared with DFM-treated (TRT) cows. Observed (<b>A</b>) and Chao1 (<b>B</b>) for Shannon index (<b>C</b>). Pooled data for overall differences between experimental groups disregarding the effect of the calendar month: Observed and Chao1, <span class="html-italic">p</span>-value 0.32, <span class="html-italic">t</span>-test: 0.99; Shannon, <span class="html-italic">p</span>-value: 0.23, <span class="html-italic">t</span>-test: −1.2. Asterisk (*) on <span class="html-italic">x</span>-axis below experimental group denotes <span class="html-italic">p</span> &lt; 0.05 between experimental groups for a given calendar month.</p>
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<p>Bacterial beta-diversity (genus level) analysis of fungi of faeces across eight time points from September 2021 to January 2023 (<b>A</b>,<b>B</b>), using Microbial diversity differed significantly over the 16 months of the trial.</p>
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<p>Prevalences of fungal genera found as the core microbiota of the faeces of Control (CON) and DFM-supplemented (TRT) cows.</p>
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17 pages, 4345 KiB  
Article
Seasonal Changes and Age-Related Effects on the Intestinal Microbiota of Captive Chinese Monals (Lophophorus lhuysii)
by Lijing Huang, Yanchu Zheng, Shaohua Feng, Bangyuan Wu, Li Chen, Xiaoqin Xu, Bin Wang, Wanhong Li, Caiquan Zhou and Long Zhang
Animals 2024, 14(23), 3418; https://doi.org/10.3390/ani14233418 - 26 Nov 2024
Viewed by 437
Abstract
The Chinese monal (Lophophorus lhuysii) is a large-sized and vulnerable (VU in IUCN) bird from southwestern China. This study applied 16S rRNA high-throughput sequencing to comprehensively examine the gut microbiota of captive Chinese monals (located in Baoxing, Sichuan, China) across varying [...] Read more.
The Chinese monal (Lophophorus lhuysii) is a large-sized and vulnerable (VU in IUCN) bird from southwestern China. This study applied 16S rRNA high-throughput sequencing to comprehensively examine the gut microbiota of captive Chinese monals (located in Baoxing, Sichuan, China) across varying seasons and life stages. Dominant bacterial phyla identified included Proteobacteria, Firmicutes, Bacteroidetes, and Actinobacteria. Significant seasonal and age-associated shifts were observed within specific bacterial groups, particularly marked by seasonal fluctuations in beta diversity. Moreover, linear discriminant analysis effect size (LEfSe) and functional predictions highlighted distinct winter signatures, indicating possible functional shifts in energy metabolism and disease resistance. In mid-aged adults, an expansion of Gamma-Proteobacteria suggested an elevated susceptibility of the gut microbiota of Chinese monals to chronic disorders and microbial imbalance. Putative pathogenic bacteria exhibited increased abundance in spring and summer, likely driven by temperature, host physiological cycles, interspecies interactions, and competition. These findings imply that the diversity, and structure of the gut microbiota in captive Chinese monals are strongly influenced by seasonal and age-related factors. The insights provided here are essential for improving breeding strategies and preventing gastrointestinal diseases in captivity. Full article
(This article belongs to the Section Birds)
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<p>Distribution of ASV in fecal samples of Chinese monal in different seasons (<b>A</b>) and ages (<b>B</b>).</p>
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<p>The relative abundance of gut microbiota in captive Chinese monals across different seasons (<b>A</b>,<b>B</b>) and age groups (<b>C</b>,<b>D</b>) was presented at both the phylum and genus levels. Panels (<b>A</b>,<b>C</b>) illustrate microbial composition at the phylum level, while panels (<b>B</b>,<b>D</b>) depict the distribution at the genus level.</p>
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<p>Seasonal and age-related variations in gut microbiota abundance in Chinese monals were assessed. LEfSe analysis, incorporating the Kruskal-Wallis test (<span class="html-italic">p</span> &lt; 0.05) and an LDA score threshold of 4.0, was employed to detect significant microbial differences across groups. A cladogram illustrates the seasonal shifts in enriched bacterial taxa (<b>A</b>), while a separate cladogram highlights age-related differences in microbial abundance (<b>B</b>). The letters preceding ASVs denote taxonomic ranks: p = phylum, c = class, o = order, f = family, g = genus, s = species.</p>
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<p>Seasonal and age-related variations in the gut microbiota of the Chinese monal were assessed through α diversity metrics. Panels (<b>A</b>,<b>E</b>) display the Shannon index, (<b>B</b>,<b>F</b>) represent the Simpson index, (<b>C</b>,<b>G</b>) illustrate the Ace index, while (<b>D</b>,<b>H</b>) depict the Chao1 index.</p>
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<p>The beta diversity of the gut microbiota composition in Chinese monals was assessed across different seasons (<b>A</b>,<b>B</b>) and age groups (<b>C</b>,<b>D</b>). NMDS and PCoA were employed to evaluate the variations in gut microbiota communities, with statistical significance denoted by <span class="html-italic">p</span> values (<span class="html-italic">p</span> &lt; 0.05). Each color corresponds to a distinct group, where proximity between samples indicates greater similarity in microbial composition and structure, while greater distance signifies increased dissimilarity. Panels (<b>A</b>,<b>C</b>) display the results from NMDS and panels (<b>B</b>,<b>D</b>) from PCoA.</p>
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<p>Differences in KEGG metabolic pathways of Chinese monal in different seasons (<b>A</b>) and ages (<b>B</b>).</p>
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<p>Seasonal (<b>A</b>,<b>C</b>) and age-related (<b>B</b>,<b>D</b>) variations in the SparCC heatmaps and relative abundance of potential pathogenic bacteria in the gut microbiota of Chinese monals were highlighted.</p>
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25 pages, 8763 KiB  
Article
Root Microbiome and Metabolome Traits Associated with Improved Post-Harvest Root Storage for Sugar Beet Breeding Lines Under Southern Idaho Conditions
by Rajtilak Majumdar, Shyam L. Kandel, Carl A. Strausbaugh, Anuradha Singh, Suresh Pokhrel and Malick Bill
Int. J. Mol. Sci. 2024, 25(23), 12681; https://doi.org/10.3390/ijms252312681 - 26 Nov 2024
Viewed by 477
Abstract
Post-harvest storage loss in sugar beets due to root rot and respiration can cause >20% sugar loss. Breeding strategies focused on factors contributing to improved post-harvest storage quality are of great importance to prevent losses. Using 16S rRNA and ITS sequencing and sugar [...] Read more.
Post-harvest storage loss in sugar beets due to root rot and respiration can cause >20% sugar loss. Breeding strategies focused on factors contributing to improved post-harvest storage quality are of great importance to prevent losses. Using 16S rRNA and ITS sequencing and sugar beet mutational breeding lines with high disease resistance (R), along with a susceptible (S) commercial cultivar, the role of root microbiome and metabolome in storage performance was investigated. The R lines in general showed higher abundances of bacterial phyla, Patescibacteria at the M time point, and Cyanobacteria and Desulfobacterota at the L time point. Amongst fungal phyla, Basidiomycota (including Athelia) and Ascomycota were predominant in diseased samples. Linear discriminant analysis Effect Size (LEfSe) identified bacterial taxa such as Micrococcales, Micrococcaceae, Bacilli, Glutamicibacter, Nesterenkonia, and Paenarthrobacter as putative biomarkers associated with resistance in the R lines. Further functional enrichment analysis showed a higher abundance of bacteria, such as those related to the super pathway of pyrimidine deoxyribonucleoside degradation, L-tryptophan biosynthesis at M and L, and fungi, such as those associated with the biosynthesis of L-iditol 2-dehydrogenase at L in the R lines. Metabolome analysis of the roots revealed higher enrichment of pathways associated with arginine, proline, alanine, aspartate, and glutamate metabolism at M, in addition to beta-alanine and butanoate metabolism at L in the R lines. Correlation analysis between the microbiome and metabolites indicated that the root’s biochemical composition, such as the presence of nitrogen-containing secondary metabolites, may regulate relative abundances of key microbial candidates contributing to better post-harvest storage. Full article
(This article belongs to the Special Issue Advances and New Perspectives in Plant-Microbe Interactions 2.0)
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<p>Sugar beet genotypes relatively resistant to post-harvest storage diseases showed differences in the abundance of bacterial and fungal phyla and genera compared to the susceptible genotype. Mean relative abundance of (<b>A</b>) bacterial phyla; (<b>B</b>) bacterial genera; (<b>C</b>) fungal phyla; and (<b>D</b>) fungal genera in sugar beet roots at mid and late post-harvest storage stages. Susceptible genotype: Sus_Ck; resistant genotypes: KSG2, KSG3, KSG4, and KSG6; M: mid time point; L: late time point. The data are mean ± standard error of 4 biological replicates, each replicate consists of tissues harvested from 2 sugar beet roots. The heat maps (<b>B</b>,<b>D</b>) are plotted using z-score values of the species abundance. ‘0’ means the abundance is at the mean value. Red color means the species abundance is higher than the mean, and blue color means that the species abundance is lower than the mean. Blue to red transition means abundance of the species is transitioning from ‘lower than mean’ to ‘higher than mean’.</p>
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<p>Beta diversity of bacteria and fungi in the resistant (R: KSG2, 3, 4, and 6) and susceptible (S: Sus_Ck) sugar beet genotypes at mid (M) and late (L) storage time points. (<b>A</b>) Cluster dendrogram of bacterial diversity; (<b>B</b>) principal coordinate analysis (PCoA) of bacterial diversity; (<b>C</b>) comparison of weighted UniFrac distances between S and R genotypes (16S); (<b>D</b>) cluster dendrogram of fungal diversity; (<b>E</b>) principal coordinate analysis (PCoA) of fungal diversity; and (<b>F</b>) comparison of weighted UniFrac distances between S and R genotypes (ITS). The data are mean ± standard error of 4 replicates (each replicate consists of 2 sugar beet roots); * <span class="html-italic">p</span> &lt; 0.05. Solid dark circle next to the treatment represents the susceptible genotype.</p>
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<p>Linear discriminant analysis Effect Size (LEfSe) analysis of sugar beet genotypes at different post-harvest storage time points shows putative biomarkers associated with resistant or susceptible genotypes. (<b>A</b>) Bar plot of bacterial taxa at the mid (M) time point; (<b>B</b>) hierarchal taxonomic cladogram of bacterial taxa at the mid time point; (<b>C</b>) bar plot of bacterial taxa at the late (L) time point; and (<b>D</b>) hierarchal taxonomic cladogram of bacterial taxa at the late time point. Susceptible genotype: Sus_Ck; resistant genotypes: KSG2, KSG3, KSG4, and KSG6. Lowercase letters denote d: domain; p: phylum; c: class; o: order; f: family; g: genus.</p>
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<p>KEGG modules were significantly different (<span class="html-italic">p</span> &lt; 0.05 *) between the resistant and susceptible sugar beet genotypes. (<b>A</b>) Mid (M) storage time point (bacteria; 16S); (<b>B</b>) late (L) storage time point (bacteria; 16S); and (<b>C</b>) mid (M) storage time point (fungi; ITS). Data are mean ± standard error of 4 replicates (each replicate consists of samples obtained from two roots). Sus_Ck: susceptible genotype; KSG2, KSG3, KSG4, and KSG6: resistant genotypes.</p>
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<p>KEGG modules were significantly different (<span class="html-italic">p</span> &lt; 0.05 *) between the resistant and susceptible sugar beet genotypes. (<b>A</b>) Mid (M) storage time point (bacteria; 16S); (<b>B</b>) late (L) storage time point (bacteria; 16S); and (<b>C</b>) mid (M) storage time point (fungi; ITS). Data are mean ± standard error of 4 replicates (each replicate consists of samples obtained from two roots). Sus_Ck: susceptible genotype; KSG2, KSG3, KSG4, and KSG6: resistant genotypes.</p>
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<p>Sparse Correlations for Compositional data (SparCC). (<b>A</b>) Correlation heatmap between bacterial communities; (<b>B</b>) correlation network between bacterial communities; (<b>C</b>) correlation heatmap between fungal communities; and (<b>D</b>) correlation network between fungal communities. A solid line between two bacterial/fungal communities indicates a positive correlation and a dotted line indicates a negative correlation between them. The thicker the solid line, the higher the value of the positive correlation between them.</p>
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<p>Untargeted metabolome analysis of sugar beet roots showed distinct differences between the resistant (R) and susceptible (S) lines at mid and late storage time points. Metabolites showing major differences between the R vs. S lines at the (<b>A</b>) mid storage stage and (<b>B</b>) late storage stage. Sus.Ck: susceptible genotype; KSG2, KSG3, KSG4, and KSG6: relatively resistant genotypes; M: mid and L: late storage time points.</p>
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<p>Pathway enrichment analysis of sugar beet roots. (<b>A</b>) Mid storage time point and (<b>B</b>) late storage time point. Data are mean ± standard error of 4 replicates (each replicate consists of samples obtained from two roots).</p>
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<p>Carbohydrate content in the roots showed small changes between the resistant and susceptible lines at the late storage time point. Cellular contents (mg/g FW) of: (<b>A</b>) sucrose and (<b>B</b>) fructose, glucose and galactose, and raffinose. Data are mean ± standard error of 4 replicates (each replicate consists of samples obtained from two roots); <span class="html-italic">p</span> &lt; 0.05 * between the susceptible (Sus_Ck) and resistant genotypes (KSG2, KSG3, KSG4, and KSG6).</p>
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<p>Correlation analysis between the root microbiome and metabolome at the late storage time point reveals a distinct pattern in the two highly resistant genotypes vs. the susceptible genotype. (<b>A</b>) Susceptible genotype (Sus_Ck); (<b>B</b>) resistant genotype, KSG4; and (<b>C</b>) resistant genotype, KSG6. Data are mean ± standard error of 4 replicates (each replicate consists of samples obtained from two roots). An ‘X’ sign inside the rectangular boxes in the heatmap indicates <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>Disease symptoms on sugar beet roots at the late storage time point under indoor storage conditions. Representative samples showing surface coverage with fungal growth in the susceptible genotype (Sus_Ck) and resistant genotypes (KSG2, KSG3, KSG4, and KSG6).</p>
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21 pages, 7032 KiB  
Article
Modulation of Canine Gut Microbiota by Prebiotic and Probiotic Supplements: A Long-Term In Vitro Study Using a Novel Colonic Fermentation Model
by Alessandro Gramenzi, Luana Clerico, Benedetta Belà, Meri Di Leonardo, Isa Fusaro and Giulia Pignataro
Animals 2024, 14(22), 3342; https://doi.org/10.3390/ani14223342 - 20 Nov 2024
Viewed by 618
Abstract
The gut microbiota plays a crucial role in dogs’ health, influencing immune function, digestion, and protection against pathogens. This study evaluates the effects of three canine dietary supplements—Microbiotal (prebiotic), Lactobacillus reuteri (probiotic), and a combination of both—on the gut microbiota composition of a [...] Read more.
The gut microbiota plays a crucial role in dogs’ health, influencing immune function, digestion, and protection against pathogens. This study evaluates the effects of three canine dietary supplements—Microbiotal (prebiotic), Lactobacillus reuteri (probiotic), and a combination of both—on the gut microbiota composition of a healthy canine donor using an in vitro colonic fermentation model. The SCIME™ platform, adapted to simulate the canine gastrointestinal tract, was used to monitor microbial shifts in the luminal and mucosal environments of the proximal and distal colon over a 2-week treatment period. The microbial communities were analyzed using 16S rRNA sequencing to assess changes at various taxonomic levels. Alpha- and beta-diversity indices were calculated, while LEfSe and treeclimbR were employed to identify taxa-driving microbial shifts. Results indicated that all treatments led to significant modulations in key microbial groups, with enrichment of Limosilactobacillus, Bifidobacterium, Prevotella, and Faecalibacterium. These changes suggest improved saccharolytic fermentation and butyrate production, particularly when prebiotics and probiotics were co-administered. This study highlights the promising benefits of combined prebiotic and probiotic supplementation in promoting gut health and microbial diversity, providing a basis for future studies targeting the metabolic activity of the gut microbiota using the same supplements and technology. Full article
(This article belongs to the Section Companion Animals)
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<p>Standard setup of the Simulator of the Canine Intestinal Microbial Ecosystem (SCIME™), consisting of four sequential reactors, simulating the different canine gastrointestinal tract regions.</p>
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<p>Modified version of the SCIME into a Triple-M-SCIME used for the current study. St + SI: vessel serving as stomach and small intestine, PC: proximal colon, and DC: distal colon.</p>
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<p>Effect of treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) on alpha diversity as calculated using four different measures (observed (count of unique taxa in each sample), Chao1, Shannon, and Simpson) in the luminal proximal colon (PC) at the end of the control (CTRL) and treatment (TR) period. Three samples (A, B, C) were collected during each period, represented by different colors.</p>
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<p>Effect of treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) on alpha diversity as calculated using four different measures (observed (count of unique taxa in each sample), Chao1, Shannon, and Simpson) in the luminal distal colon (DC) at the end of the control (CTRL) and treatment (TR) period. Three samples (A, B, C) were collected during each period, represented by different colors.</p>
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<p>Effect of treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) on alpha diversity as calculated using four different measures (observed (count of unique taxa in each sample), Chao1, Shannon, and Simpson) in the mucosal proximal colon (PC) at the end of the control (CTRL) and treatment (TR) period. Three samples (A, B, C) were collected during each period, represented by different colors.</p>
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<p>Effect of treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) on alpha diversity as calculated using four different measures (observed (count of unique taxa in each sample), Chao1, Shannon, and Simpson) in the mucosal distal colon (DC) at the end of the control (CTRL) and treatment (TR) period. Three samples (A, B, C) were collected during each period, represented by different colors.</p>
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<p>Discriminant analysis of principal components (DAPC) to show differences in community composition (beta diversity) in the luminal proximal colon (PC) at the end of the control (CTRL) and treatment (TR) period following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P). Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3). Each dot represents one sample.</p>
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<p>Discriminant analysis of principal components (DAPC) to show differences in community composition (beta diversity) in the luminal distal colon (DC) at the end of the control (CTRL) and treatment (TR) period following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P). Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3). Each dot represents one sample.</p>
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<p>Discriminant analysis of principal components (DAPC) to show differences in community composition (beta diversity) in the mucosal proximal colon (PC) at the end of the control (CTRL) and treatment (TR) period following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P). Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3). Each dot represents one sample.</p>
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<p>Discriminant analysis of principal components (DAPC) to show differences in community composition (beta diversity) in the mucosal distal colon (DC) at the end of the control (CTRL) and treatment (TR) period following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P). Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3). Each dot represents one sample.</p>
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<p>Jitter plots showing abundances of different phyla in the luminal proximal colon (PC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on absolute levels. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of different phyla in the luminal distal colon (DC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on absolute levels. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of different phyla in the mucosal proximal colon (PC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on relative abundances. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of different phyla in the mucosal distal colon (DC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on relative abundances. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant families in the luminal proximal colon (PC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on absolute levels. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant families in the luminal distal colon (DC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on absolute levels. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant families in the mucosal proximal colon (PC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on relative abundances. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant families in the mucosal distal colon (DC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on relative abundances. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant genera in the luminal proximal colon (PC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on absolute levels. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant genera in the luminal distal colon (DC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on absolute levels. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant genera in the mucosal proximal colon (PC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on relative abundances. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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<p>Jitter plots showing abundances of the 20 most abundant genera in the mucosal distal colon (DC) following treatment with the different test products (Microbiotal, M; probiotic, P; and their combination, M + P) at the end of the control (CTRL) and treatment (TR) period based on relative abundances. Each color represents one of six categories (groups), i.e., CTRL_M (n = 3), TR_M (n = 3), CTRL_P (n = 3), TR_P (n = 3), CTRL_M + P (n = 3), and TR_M + P (n = 3).</p>
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18 pages, 6181 KiB  
Article
The Colonization of Synthetic Microbial Communities Carried by Bio-Organic Fertilizers in Continuous Cropping Soil for Potato Plants
by Wenming Zhang, Shiqing Li, Pingliang Zhang, Xuyan Han, Yanhong Xing and Chenxu Yu
Microorganisms 2024, 12(11), 2371; https://doi.org/10.3390/microorganisms12112371 - 20 Nov 2024
Viewed by 555
Abstract
Synthetic microbial communities (SynComs) play significant roles in soil health and sustainable agriculture. In this study, bacterial SynComs (SCBs) and fungal SynComs (SCFs) were constructed by selecting microbial species that could degrade the potato root exudates associated with continuous cropping obstacles. SCBs, SCFs, [...] Read more.
Synthetic microbial communities (SynComs) play significant roles in soil health and sustainable agriculture. In this study, bacterial SynComs (SCBs) and fungal SynComs (SCFs) were constructed by selecting microbial species that could degrade the potato root exudates associated with continuous cropping obstacles. SCBs, SCFs, and SCB + SCF combinations were then inoculated into organic fertilizers (OFs, made from sheep manure) to produce three bio-organic fertilizers (BOFs), denoted by SBFs (BOFs of inoculated SCBs), SFFs (BOFs of inoculated SCFs), and SBFFs (BOFs of inoculated SCB + SCF combinations), respectively. The OF and three BOFs, with a chemical fertilizer (CK) as the control, were then used in pot experiments involving potato growth with soil from a 4-year continuous cropping field. Microbial diversity sequencing was used to investigate the colonization of SCBs and SCFs into the rhizosphere soil and the bulk soil, and their effects on soil microbial diversity were evaluated. Source Tracker analysis showed that SCBs increased bacterial colonization from the SBFs into the rhizosphere soil, but at a relatively low level of 1% of the total soil bacteria, while SCFs increased fungi colonization from the SFF into the bulk soil at a much higher level of 5–18% of the total soil fungi. In combination, SCB + SCF significantly increased fungi colonization from the SBFF into both the bulk soil and the rhizosphere soil. Overall, the soil fungi were more susceptible to the influence of the BOFs than the bacteria. In general, the application of BOFs did not significantly change the soil microbial alpha diversity. Correlation network analysis showed that key species of bacteria were stable in the soils of the different groups, especially in the rhizosphere soil, while the key species of fungi significantly changed among the different groups. LEfSe analysis showed that the application of BOFs activated some rare species, which were correlated with improvements in the function categories of the tolerance of stress, nitrogen fixation, and saprotroph functions. Mantel test analysis showed that the BOFs significantly affected soil physicochemical properties, influencing bacterial key species, and core bacteria, promoting potato growth. It was also noted that the presence of SynCom-inoculated BOFs may lead to a slight increase in plant pathogens, which needs to be considered in the optimization of SynCom applications to overcome continuous cropping obstacles in potato production. Full article
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<p>Source Tracker analysis of SynCom colonization into the BOF (SBF: BOF inoculated with SCBs; SFF: BOF inoculated with SCFs; SBFF: BOF inoculated with SCB + SCF).</p>
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<p>Source Tracker analysis of microbial colonization from BOF into bulk soil (B) and rhizosphere soil (R): (<b>a</b>). soil bacteria; (<b>b</b>). soil fungi (OF: organic fertilize; SBF: BOF inoculated with SCBs; SFF: BOF inoculated with SCFs; SBFF: BOF inoculated with SCB + SCF).</p>
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<p>Effect of application of BOFs on soil microbial alpha diversity and species composition (B: bulk soil; R: rhizosphere soil: (<b>a</b>), bacteria, (<b>b</b>): fungi). CK: no organic fertilizers applied; F: organic fertilizer applied; FSB: SBF applied; FSF: SFF applied; FSBF: SBFF applied.</p>
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<p>Effect of bio-organic fertilizer implication on soil microbial beta diversity (B: bulk soil; R: rhizosphere soil: (<b>a</b>): bacteria, (<b>b</b>): fungi). Ellipses represent a 95% confidence interval. CK: no organic fertilizers applied; F: organic fertilizer applied; FSB: SBF applied; FSF: SFF applied; FSBF: SBFF applied.</p>
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<p>The correlation network diagram of each group (B: bulk soil; R: rhizosphere soil: (<b>a</b>), bacteria, (<b>b</b>): fungi). The correlation network was constructed as Spearman’s correlation, with a genera relative abundance of &gt;0.1%, r &gt; 0.8, and <span class="html-italic">p</span> &lt; 0.05. Red lines represent significant positive relationships, and green lines denote negative relationships. The genera shown on these figures are the module hub (key species) in each network diagram (ZI (within-module connectivity), with &gt;2.5 and PI (among-module connectivity) &lt; 0.62. CK: no organic fertilizers applied; F: organic fertilizer applied; FSB: SBF applied; FSF: SFF applied; FSBF: SBFF applied.</p>
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<p>Line discriminant analysis (LDA) effect size (LEfSe) analysis (B: bulk soil; R: rhizosphere soil; (<b>a</b>), bacteria, (<b>b</b>): fungi). CK: no organic fertilizers applied; F: organic fertilizer applied; FSB: SBF applied; FSF: SFF applied; FSBF: SBFF applied. The figures show the genera with LDA scores greater than 3.0.</p>
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<p>Mantel test analysis of plant growth, environmental factors, and microbial community (biomarkers were analyzed according to <a href="#microorganisms-12-02371-f006" class="html-fig">Figure 6</a>, key species were analyzed according to <a href="#microorganisms-12-02371-f005" class="html-fig">Figure 5</a>, and the fraction of the core bacterial/fungal species analyzed with a Veen plot are shown in <a href="#microorganisms-12-02371-f003" class="html-fig">Figure 3</a>). *, **, and *** represent <span class="html-italic">p</span> &lt; 0.05, <span class="html-italic">p</span> &lt; 0.01, <span class="html-italic">p</span> &lt; 0.001, respectively; a dark green square represents significant positive relationships, and a pink square represents negative relationships. The darker the color or the larger the square area, the greater the absolute value of the correlation coefficient; red lines represent <span class="html-italic">p</span> &lt; 0.01, green lines represent 0.01 &lt; <span class="html-italic">p</span> &lt; 0.05, blue lines represent <span class="html-italic">p</span> ≥  0.05, — represents a positive relationship, --- represents a negative relationship, and the width of the line represents the magnitude of the correlation coefficient.</p>
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<p>Function prediction of microbial community (R: rhizosphere soil; B: bulk soil). CK: no organic fertilizers applied; F: organic fertilizer applied; FSB: SBF applied; FSF: SFF applied; FSBF: SBFF applied.</p>
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26 pages, 5180 KiB  
Article
Sugar Composition of Thai Desserts and Their Impact on the Gut Microbiome in Healthy Volunteers: A Randomized Controlled Trial
by Sayamon Senaprom, Nuttaphat Namjud, Thunnicha Ondee, Akkarach Bumrungpert and Krit Pongpirul
Nutrients 2024, 16(22), 3933; https://doi.org/10.3390/nu16223933 - 18 Nov 2024
Viewed by 754
Abstract
Background: The relationship between consuming Thai desserts—predominantly composed of carbohydrates—and gut microbiome profiles remains unclear. This study aimed to evaluate the effects of consuming various Thai desserts with different GI values on the gut microbiomes of healthy volunteers. Methods: This open-label, parallel randomized [...] Read more.
Background: The relationship between consuming Thai desserts—predominantly composed of carbohydrates—and gut microbiome profiles remains unclear. This study aimed to evaluate the effects of consuming various Thai desserts with different GI values on the gut microbiomes of healthy volunteers. Methods: This open-label, parallel randomized clinical trial involved 30 healthy individuals aged 18 to 45 years. Participants were randomly assigned to one of three groups: Phetchaburi’s Custard Cake (192 g, low-GI group, n = 10), Saraburi’s Curry Puff (98 g, medium-GI group, n = 10), and Lampang’s Crispy Rice Cracker (68 g, high-GI group, n = 10), each consumed alongside their standard breakfast. Fecal samples were collected at baseline and 24 h post-intervention for metagenomic analysis of gut microbiome profiles using 16S rRNA gene sequencing. Results: After 24 h, distinct trends in the relative abundance of various gut microbiota were observed among the dessert groups. In the high-GI dessert group, the abundance of Collinsella and Bifidobacterium decreased compared to the low- and medium-GI groups, while Roseburia and Ruminococcus showed slight increases. Correlation analysis revealed a significant negative relationship between sugar intake and Lactobacillus abundance in the medium- and high-GI groups, but not in the low-GI group. Additionally, a moderately negative association was observed between Akkermansia abundance and sugar intake in the high-GI group. These bacteria are implicated in energy metabolism and insulin regulation. LEfSe analysis identified Porphyromonadaceae and Porphyromonas as core microbiota in the low-GI group, whereas Klebsiella was enriched in the high-GI group, with no predominant bacteria identified in the medium-GI group. Conclusions: The findings suggest that Thai desserts with varying GI levels can influence specific gut bacteria, though these effects may be temporary. Full article
(This article belongs to the Special Issue Nutrition–Microbiome Interaction in Healthy Metabolism)
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<p>Nutritional value per 50 g of available carbohydrates (one serving size) of Thai desserts for testing. Note: Nutritional value data were retrieved from Namjud et al. [<a href="#B21-nutrients-16-03933" class="html-bibr">21</a>], except for total sugar content, which was assessed in this study. The serving size was 50 g of available carbohydrates [<a href="#B26-nutrients-16-03933" class="html-bibr">26</a>]. ND = not detected; GI = glycemic index; low GI ≤ 55, medium GI = 56–69, and high GI ≥ 70 [<a href="#B27-nutrients-16-03933" class="html-bibr">27</a>].</p>
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<p>CONSORT flow chart of the experimental design in this study.</p>
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<p>The relative abundance of the identified phyla, classes, orders, families, and genera of only the top 100 taxa. The relative abundance data of all identified taxa are presented in <a href="#app1-nutrients-16-03933" class="html-app">Supplementary Tables S1–S5</a>.</p>
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<p>Boxplots showing the comparison of changes in relative abundance of the gut microbiota after the 24 h consumption of testing Thai desserts. The gut microbiome profiles were compared at the level of phylum (<b>A</b>), class (<b>B</b>), order (<b>C</b>), family (<b>D</b>), and genus (<b>E</b>). Data were analyzed using the Kruskal–Wallis test and Dunn’s post hoc test, with the Benjamini-Hochberg method. Asterisks (*) indicate significant differences at adjusted <span class="html-italic">p</span> &lt; 0.05. More detailed data on changes in relative abundance are presented in <a href="#app1-nutrients-16-03933" class="html-app">Supplementary Table S6</a>.</p>
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<p>The gut microbial taxa determined as dominant biomarkers for association with the consumption of Thai desserts. After the 24 h intervention in the low-GI and high-GI dessert groups, the three taxa, the family Porphyromonadaceae, and the genera <span class="html-italic">Porphyromonas</span> and <span class="html-italic">Klebsiella</span> were determined as dominant biomarkers as analyzed by LEfSe with an LDA score ≥ 1.0 (<b>A</b>). Cladogram exhibiting the phylogenetic distributions of gut microbiota (<b>B</b>).</p>
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<p>The alpha diversity indices of the dessert groups are shown by boxplots. The indices consist of observed ASVs, Chao1, Shannon, and PD whole tree. Black dots represent 20 samples obtained from combining the index values of both baseline and after the 24 h intervention for each dessert group. Alpha diversity values of each sample and quartiles of the distribution (minimum, first quartile, median, third quartile, and maximum of boxes) are demonstrated. No significant differences between dessert groups were observed by One-Way ANOVA or the Kruskal–Wallis test on any indices.</p>
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<p>The beta diversity indices between dessert groups as shown by 2-dimentional plots. The indices consist of PCoA on weighted (<b>A</b>) and unweighted (<b>B</b>) UniFrac distances, or GUniFrac distances (<b>C</b>), and NMDS based on Bray–Curtis dissimilarity (<b>D</b>). Colored dots represent samples at baseline or after the 24 h intervention for each dessert group. No significant difference between dessert groups was observed by the PERMANOVA test in any indices.</p>
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<p>Correlation between dietary nutrient intake and the relative abundance of gut microbiota at the genus level. The correlation data were analyzed by Spearman’s rank correlation coefficient. Asterisks (*) indicate statistically significant Spearman’s rho (<span class="html-italic">ρ</span>) values at <span class="html-italic">p</span> &lt; 0.05, while daggers (†) indicate the tendency of correlation at 0.05 ≤ <span class="html-italic">p</span> &lt; 0.10.</p>
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11 pages, 1567 KiB  
Article
Leaching Efficiency During Autumn Irrigation in China’s Arid Hetao Plain as Influenced by the Depth of Shallow Saline Groundwater and Irrigation Depth, Using Data from Static Water-Table Lysimeters and the Hydrus-1D and SIMDualKc Models
by Tiago B. Ramos, Meihan Liu, Haibin Shi, Paula Paredes and Luis S. Pereira
Land 2024, 13(11), 1797; https://doi.org/10.3390/land13111797 - 31 Oct 2024
Viewed by 610
Abstract
The need for controlling salinity in arid zones is essential for sustainable agricultural production and irrigation water use. A case study performed for two years in Hetao, Inner Mongolia, China, is used herein to rethink the contradictory issues of arid lands represented by [...] Read more.
The need for controlling salinity in arid zones is essential for sustainable agricultural production and irrigation water use. A case study performed for two years in Hetao, Inner Mongolia, China, is used herein to rethink the contradictory issues of arid lands represented by water saving and controlling soil and water salinity. Two sets of static lysimeters, where water table depths (WTDs) were fixed at 1.25, 150, 2.00, and 2.25 m, were continuously monitored, and soil water and solute data were used to calibrate and validate two models: the soil water balance model SIMDualKc and the deterministic soil water and salt dynamics model HYDRUS-1D. Once accurately calibrated, the models were used to simulate maize water use, percolation, and capillary rise, along with the observed variables for the actual WTD and the autumn irrigation applied. Simulation scenarios also considered agricultural system degradation and dynamic water table behavior. Results have shown that large leaching efficiencies (Lefs) were obtained for large irrigation depths in cases of shallow water tables, but higher Lefs corresponded to high application depths when the water table was deeper. Agricultural system degradation, particularly increased groundwater salinity, lowered Lef, regardless of WTD. Conversely, water savings were minimal and only achievable when considering the dynamic nature of groundwater. These results indicate that there is a need to define different WTDs based on soil characteristics that influence fluxes and root zone storage, as well as the impacts of newly installed drainage systems aimed at salt extraction. Full article
(This article belongs to the Special Issue Salinity Monitoring and Modelling at Different Scales: 2nd Edition)
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<p>Reference evapotranspiration (ETo), precipitation (P), and irrigation (I) during the 2017 and 2018 growing seasons (AI, autumn irrigation).</p>
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<p>The efficiency of autumn irrigation depth for leaching rootzone salts under variable water table depth (WTD) conditions (adapted from Ramos et al. [<a href="#B33-land-13-01797" class="html-bibr">33</a>]).</p>
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<p>The efficiency of autumn irrigation depth for leaching rootzone salts under variable water table depth (WTD) conditions in a scenario where groundwater salinity (EC<sub>gw</sub>) increased 0.5 and 1.0 dS m<sup>−1</sup> compared to present conditions (1.75 dS m<sup>−1</sup> as observed in 2018). (<b>a</b>) WTD = 1.25 m; (<b>b</b>) WTD = 1.5 m; (<b>c</b>) WTD = 2.0 m; (<b>d</b>) WTD = 2.25 m.</p>
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<p>The efficiency of autumn irrigation depth for leaching rootzone salts in scenarios with static and variable water table depths (WTD). For both scenarios, the mean WTD was 1.5 m.</p>
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18 pages, 3455 KiB  
Article
The Rumen Microbiome Composition of Raramuri Criollo and European Cattle in an Extensive System
by Adrian Maynez-Perez, Francisco J. Jahuey-Martínez, José A. Martínez-Quintana, Michael E. Hume, Robin C. Anderson, Agustín Corral-Luna, Felipe A. Rodríguez-Almeida, Yamicela Castillo-Castillo and Monserrath Felix-Portillo
Microorganisms 2024, 12(11), 2203; https://doi.org/10.3390/microorganisms12112203 - 31 Oct 2024
Viewed by 656
Abstract
Understanding the relationship between Raramuri Criollo cattle (RC) and their microbial ruminal ecosystem will help identify advantageous characteristics of adapted cattle as alternatives to achieve sustainable beef production systems. Our objective was to characterize the rumen microbiome of RC in comparison to Angus [...] Read more.
Understanding the relationship between Raramuri Criollo cattle (RC) and their microbial ruminal ecosystem will help identify advantageous characteristics of adapted cattle as alternatives to achieve sustainable beef production systems. Our objective was to characterize the rumen microbiome of RC in comparison to Angus and Hereford breeds (European, E) and the cross between them (E × RC). Ruminal fluid was collected from 63 cows in their second productive cycle after grazing in the same paddock for 45 d, in the dry (n = 28) and rain (n = 35) seasons. DNA from ruminal fluid was isolated for 16s rRNA gene next-generation sequencing. The data were analyzed with QIIME2 and compared against the SILVA 16s rRNA database. Beta diversity was different (p < 0.05) between RC and E in both seasons. A microbial core was represented by the most abundant phyla. Planctomycetes and Spirochaetes represented above 1% in the rain season and below 1% in the dry one, whereas Euryarchaeota was below 1% and around 3%, respectively. LEfSe analysis identified differentiated (p < 0.05) key microbial groups that explain the differences between lineages at different taxonomic levels, reflecting the ability of the rumen ecosystem of RC cattle to adapt to hostile environmental conditions by having microbial groups specialized in the degradation of highly fibrous content. Full article
(This article belongs to the Section Microbiomes)
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<p>Principal coordinates analysis (PCoA) showing the different localization of bovine rumen microorganisms of three lineages. Based on the unweighted unifrac distance ((<b>A</b>,<b>C</b>) for the dry and rain seasons, respectively) and Bray–Curtis dissimilarity matrix ((<b>B</b>,<b>D</b>) for the dry and rain seasons, respectively).</p>
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<p>Heatmap of ruminal core microbiomes of three cattle lineages (Raramuri Criollo = Red, European = Green; E × RC = Blue) in the dry (<b>A</b>) and rainy (<b>B</b>) seasons. The reported microbial groups were identified in at least 50% of samples. The number provided before the microbial taxa represents the taxonomic level (D_1: Phylum; D_2: Class; D_3: Order; D_4: Family; D_5: Genus).</p>
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<p>Linear discriminant analysis effect size (LEfSe) to determine biomarker microorganisms within the rumen microbial community of different lineages for the dry (<b>A</b>) and the rainy (<b>B</b>) seasons. The taxonomic levels of the microbial groups are indicated by the prefixes: D_2: Phylum; D_3: Class; D_4: Order; D_5: Family; D_6: Genus.</p>
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<p>Principal coordinate analysis (PCoA) showing the different localization of seasons based on Bray–Curtis distances of predicted metabolic pathways in ruminal microorganisms, identified by the CowPi tool.</p>
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<p>Boxplot of mismatch repair, pantothenate, CoA biosynthesis, peptidoglycan biosynthesis, ribosome biogenesis, and starch and sucrose metabolism predicted pathways for bacteria isolated from the rumen of different lineage cows during the rainy season.</p>
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19 pages, 7630 KiB  
Article
Investigation into Critical Gut Microbes Influencing Intramuscular Fat Deposition in Min Pigs
by Long Jin, Ke Li, Zhimin Li, Xuankai Huang, Li Wang, Xibiao Wang, Shengwei Di, Shiquan Cui and Yuan Xu
Animals 2024, 14(21), 3123; https://doi.org/10.3390/ani14213123 - 30 Oct 2024
Viewed by 641
Abstract
To determine the pivotal microorganisms affecting intramuscular fat (IMF) accumulation in Min pigs and to discern the extent of the influence exerted by various intestinal segments on IMF-related traits, we sequenced 16S rRNA from the contents of six intestinal segments from a high [...] Read more.
To determine the pivotal microorganisms affecting intramuscular fat (IMF) accumulation in Min pigs and to discern the extent of the influence exerted by various intestinal segments on IMF-related traits, we sequenced 16S rRNA from the contents of six intestinal segments from a high IMF group (Group H) and a low IMF group (Group L) of Min pigs weighing 90 ± 1 kg. We then compared their diversity and disparities in bacterial genera. Group H exhibited considerably higher α diversity in the jejunum and colon than Group L (p < 0.05). When 95% confidence levels were considered, the main β diversity components for the ileum, caecum, and colon within Groups H and L exhibited absolute segregation. Accordingly, 31 differentially abundant genera across Group H were pinpointed via LEfSe and the Wilcoxon test (p < 0.05) and subsequently scrutinised based on their distribution and abundance across distinct intestinal segments and their correlation with IMF phenotypes. The abundances of Terrisporobacter, Acetitomaculum, Bacteroides, Fibrobacter, Treponema, Akkermansia, Blautia, Clostridium sensu stricto 1, Turicibacter, Subdoligranulum, the [Eubacterium] siraeum group, and dgA 11 gut groups were positively correlated with IMF content (p < 0.05), whereas those of Bacillus, the Lachnospiraceae NK4A136 group, Streptococcus, Roseburia, Solobacterium, Veillonella, Lactobacillus, the Rikenellaceae RC9 gut group, Anaerovibrio, and the Lachnospiraceae AC2044 group were negatively associated with IMF content (p < 0.05). Employing PICRUSt2 for predicting intergenic metabolic pathways that differ among intestinal microbial communities revealed that within the 95% confidence interval the colonic microbiome was enriched with the most metabolic pathways, including those related to lipid metabolism. The diversity results, bacterial genus distributions, and metabolic pathway disparities revealed the colonic segment as an influential region for IMF deposition. Full article
(This article belongs to the Section Pigs)
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<p>Sample clustering and comparison of IMF content. (<b>A</b>): Conceptual clustering diagram for the Min pig population on the basis of IMF content. Red lines (M1–M9): individuals in the low IMF group; Blue lines (M10–M20): individuals with intermediate IMF levels; Green lines (M21–M30): individuals in the high IMF group. (<b>B</b>): Difference in IMF contents between Groups L and H, *** <span class="html-italic">p</span>&lt; 0.0001.</p>
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<p>Paraspinal longissimus muscle oil-red-O-stained sections from Groups L and H. (<b>A</b>,<b>B</b>): Section from Group L. (<b>C</b>,<b>D</b>): Section from Group H.</p>
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<p>Sequencing depth chart of the entire sample. (<b>A</b>): Dilution curve. (<b>B</b>): Shannon curve.</p>
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<p>The exclusive and shared OTU taxa among the contents of different intestinal segments in Groups H and L. (<b>A</b>): Duodenum. (<b>B</b>): Jejunum. (<b>C</b>): Ileum. (<b>D</b>): Caecum. (<b>E</b>): Colon. (<b>F</b>): Rectum.</p>
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<p>The α diversity indices of specific intestinal segments from Groups H and L. (<b>A</b>): ACE index. (<b>B</b>): Chao1 index. (<b>C</b>): Shannon index. (<b>D</b>): Simpson index, * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01.</p>
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<p>β diversity indices of the respective intestinal segments in Groups H and L. (<b>A</b>): Duodenum. (<b>B</b>): Jejunum. (<b>C</b>): Ileum. (<b>D</b>): Caecum. (<b>E</b>): Colon. (<b>F</b>): Rectum.</p>
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<p>Distribution of prominent microbial populations at both the phylum and genus levels for each intestinal segment. (<b>A</b>–<b>F</b>): Phylum level. (<b>G</b>–<b>L</b>): Genus level.</p>
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<p>The microbiome profile analysis results for each intestinal segment (using an LDA &gt; 4.0 as a discernment criterion). (<b>A</b>): Duodenum. (<b>B</b>): Jejunum. (<b>C</b>): Ileum. (<b>D</b>): Caecum. (<b>E</b>): Colon. (<b>F</b>): Rectum.</p>
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<p>Significant differences were identified in the Group H and Group L genera across intestinal segments, * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01. (<b>A</b>): Duodenum. (<b>B</b>): Jejunum. (<b>C</b>): Ileum. (<b>D</b>): Caecum. (<b>E</b>): Colon. (<b>F</b>): Rectum.</p>
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<p>Genomic variations in the microbiome communities of Groups H and L revealed significant KEGG pathway enrichment (95% confidence interval), <span class="html-italic">p</span> &lt; 0.05, Red dashed boxes indicate KEGG pathways related to metabolism. The circles show deviations from the 95% confidence interval, indicating inter-group differences. (<b>A</b>): duodenum, (<b>B</b>): ileum, (<b>C</b>): caecum, (<b>D</b>): colon, (<b>E</b>): rectum.</p>
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13 pages, 4189 KiB  
Article
Spatiotemporal Distribution of nirS-Type Denitrifiers in Cascade Reservoir Sediments of the Qinghai Plateau
by Yi Wu, Xufeng Mao, Hongyan Yu, Hongyan Li, Feng Xiao, Yuhua Mo, Haichuan Ji and Yuanjie Ma
Diversity 2024, 16(11), 656; https://doi.org/10.3390/d16110656 - 24 Oct 2024
Viewed by 576
Abstract
Compared to single damming, the impact of cascade damming on nitrogen-related microorganisms in river ecosystems exhibits greater complexity. However, there is still a lack of research on the response of denitrifiers to the construction of cascade reservoirs. A study was conducted on 10 [...] Read more.
Compared to single damming, the impact of cascade damming on nitrogen-related microorganisms in river ecosystems exhibits greater complexity. However, there is still a lack of research on the response of denitrifiers to the construction of cascade reservoirs. A study was conducted on 10 cascade reservoirs in the upper reaches of the Yellow River to investigate the impact of cascade reservoir construction on nirS-type denitrifying bacteria in sediments. Sediment samples were collected in May (dry season) and August (wet season) of 2023. The spatiotemporal characteristics of the nirS-type denitrifying bacterial community and gene abundance were analyzed using Illumina high-throughput sequencing and real-time fluorescence quantification PCR (qPCR). Redundancy analysis (RDA) and variation partitioning (VP) were utilized to assess the impact of environmental factors on these communities. The results showed the following: (1) Proteobacteria was the predominant phylum of nirS-type denitrifying bacteria in cascade reservoir sediments. At the genus level, unclassified Proteobacteria (69.51–95.64%) showed the highest relative abundance, followed by Paracoccus, Rhodanobacter, and Pseudomonas, indicating that unclassified Proteobacteria may dominate denitrification in these reservoir sediments. (2) The α and β diversity indices of nirS-type denitrifying bacteria were higher in the dry season than in the wet season, and also higher in young reservoirs compared to old reservoirs (p < 0.05). (3) Temporally, the abundance of the nirS gene was significantly higher in the wet season (12.71 × 107 copies/g dry sediment) compared to the dry season (66.35 × 105 copies/g dry sediment). Spatially, the abundance of the nirS gene was higher in the central region, while relatively lower at both ends. (4) RDA and VP analysis indicated that the community structure and abundance of nirS-type denitrifying bacteria were significantly influenced by the total nitrogen in sediments (19.31%) and water temperature (14.13%). Spearman correlation analysis showed that organic carbon significantly affected the diversity of nirS-type denitrifying bacteria (p < 0.05). The results contribute to a better understanding of the nitrogen-related microbial community in cascade reservoir sediments of the Yellow River, providing a scientific basis for reservoir management. Full article
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<p>Distribution of cascade reservoirs in the upper reaches of the Yellow River. Longyangxia (LYX), Ninna (NN), Lijiaxia (LJX), Zigonglaka (ZGLK), Kangyang (KY), Gongboxia (GBX), Suzhi (SZ), Huangfeng (HF), Jishixia (JXS), and Dahejia (DHJ).</p>
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<p>Venn diagram of OTUs of reservoirs in the young reservoirs (<b>a</b>) and old reservoirs (<b>b</b>). The first part of the abbreviation represents the reservoir name, while the second part represents the sampling seasons.</p>
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<p>Relative abundance of <span class="html-italic">nirS</span>-type denitrifying bacterial communities in sediment at phylum (<b>a</b>) and genus levels (<b>b</b>) with reservoir abbreviations as described in <a href="#sec2dot1-diversity-16-00656" class="html-sec">Section 2.1</a>.</p>
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<p>LEfSe analysis of <span class="html-italic">nirS</span>-type denitrifying bacteria in different seasons. LDA value histogram (<b>a</b>) and Cladogram diagram (<b>b</b>).</p>
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<p>Alpha diversity of <span class="html-italic">nirS</span>-type denitrifying bacteria in sediments between different seasons (<b>a</b>) and reservoir ages (<b>b</b>).</p>
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<p>PCoA analysis of sediment <span class="html-italic">nirS</span>-type denitrifying bacteria between different seasons (<b>a</b>) and reservoir ages (<b>b</b>).</p>
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<p>Abundance of <span class="html-italic">nirS</span> gene in different seasons and reservoir age. Uppercase and lowercase letters denote statistically significant differences (<span class="html-italic">p</span> &lt; 0.05) in gene abundance between young and old reservoirs, and among 10 cascade reservoirs within the same season, respectively. Error bars represent standard deviations.</p>
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<p>Redundancy analysis (<b>a</b>) and hierarchical partitioning (<b>b</b>) of environmental factors and <span class="html-italic">nirS</span>-type denitrifying bacteria communities. Red arrows indicate dominant bacterial genera, while gray arrows represent environmental factors. Blue and red circles denote the 95% confidence intervals for reservoir ordination in the wet and dry seasons, respectively.</p>
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<p>Correlation heatmap of the dominant genus, diversity, and abundance of <span class="html-italic">nirS</span>-type denitrifying bacteria and environmental factors. The size of the boxes in the lower left corner represents the size of the correlation between the environment factors, and the color represents the positive or negative correlation. The color of lines represents the magnitude of significance, and the thickness represents the magnitude of correlation with environmental factors. “*”, “**”, and “***” indicate statistically significant differences at the significance levels of <span class="html-italic">p</span> &lt; 0.05, 0.01, and 0.001, respectively.</p>
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17 pages, 5466 KiB  
Article
Effects of Anthracnose on the Structure and Diversity of Endophytic Microbial Communities in Postharvest Avocado Fruits
by Xi Chen, Zhuoen Jiang, Peng He, Xiuhua Tang, Haiyun Song, Tao Zhang, Zhejun Wei, Tao Dong, Shufang Zheng, Xinghao Tu, Jian Qin, Jingjing Chen and Wenlin Wang
Agronomy 2024, 14(11), 2487; https://doi.org/10.3390/agronomy14112487 - 24 Oct 2024
Viewed by 602
Abstract
This study aimed to provide foundational research for the biological control of postharvest avocado fruits anthracnose and establish a microbial system of postharvest avocado fruits. The high-throughput sequencing of avocado fruits from the anthracnose-infected and healthy groups was performed using Illumina NovaSeq second-generation [...] Read more.
This study aimed to provide foundational research for the biological control of postharvest avocado fruits anthracnose and establish a microbial system of postharvest avocado fruits. The high-throughput sequencing of avocado fruits from the anthracnose-infected and healthy groups was performed using Illumina NovaSeq second-generation sequencing technology. The results revealed that, except for Colletotrichum sp. strain 38#, there were differences in the bacterial community structure of avocados before and after infection, as determined through alpha and beta diversity analysis. Additionally, there were significant differences in the endophytic fungal community structure, allowing clear differentiation between the infected and healthy avocados. The endophytic bacterial community was primarily composed of 4 phyla and 10 genera, with the Bacteroidota phylum and Chryseobacterium genus demonstrating sensitivity to anthracnose pathogens, as evidenced by a decrease in their relative abundance after infection. The endophytic fungal community was characterized by 3 phyla and 10 genera. After infection, the relative abundance of 2 phyla (Anthophyta and Basidiomycota) and 7 genera (Eucalyptus, Candida, Kluyveromyces, Talaromyces, Oidiodendron, Nigrospora, and Pestalotiopsis) decreased, whereas the relative abundance of the Colletotrichum genus increased dramatically. The LEfSe (Linear discriminant analysis Effect Size) analysis indicated that significant biomarkers were more prevalent in endophytic bacteria than in endophytic fungi in the avocados. In endophytic bacteria, the key biomarkers included the Firmicutes phylum (Bacilli class), Proteobacteria phylum (Gammaproteobacteria class, Pseudomonadales order, Pseudomonadaceae family, and Pseudomonas genus), Flavobacteriales order, Weeksellaceae family, and Chryseobacterium genus. In endophytic fungi, the important biomarkers were Saccharomycetes class (Saccharomycetales order), Glomerellales order (Glomerellaceae family and Colletotrichum genus), and Botryosphaeriales order (Botryosphaeriaceae family and Lasiodiplodia genus). These results may provide a theoretical basis for the development of future biological agents for avocado anthracnose. Full article
(This article belongs to the Section Horticultural and Floricultural Crops)
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<p>Healthy Avocado Groups (CK).</p>
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<p>Infected Avocado Groups. (<b>a</b>) <span class="html-italic">Colletotrichum</span> sp. strain No. 31 (31#). (<b>b</b>) <span class="html-italic">Colletotrichum</span> sp. strain No. 38 (38#). (<b>c</b>) <span class="html-italic">Colletotrichum</span> sp. strain No. 64 (64#).</p>
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<p>Distribution of endophytic sequences. (<b>a</b>) Endophytic Bacteria. (<b>b</b>) Endophytic Fungi. CK represents the healthy group. S31, S38, and S64 represent the groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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<p>Annotation result statistics of ASVs. (<b>a</b>) Endophytic Bacteria. (<b>b</b>) Endophytic Fungi. CK represents the healthy group. S31, S38, and S64 represent infected groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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<p>Rarefaction curve of endophytes. (<b>a</b>) Endophytic Bacteria. (<b>b</b>) Endophytic Fungi. CK represents the healthy group. S31, S38, and S64 represent the groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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<p>Species accumulation boxplot of endophytes. (<b>a</b>) Endophytic Bacteria. (<b>b</b>) Endophytic Fungi.</p>
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<p>NMDS of endophytes. (<b>a</b>) Endophytic Bacteria. (<b>b</b>) Endophytic Fungi. CK represents the healthy group. S31, S38, and S64 represent the groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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<p>Histogram of relative abundance of species in the endophytic bacteria. (<b>a</b>) Phylum Level. (<b>b</b>) Genus Level. CK represents the healthy group. S31, S38, and S64 represent the groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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<p>Histogram of relative abundance of species in the endophytic fungi. (<b>a</b>) Phylum Level. (<b>b</b>) Genus Level. CK represents the healthy group. S31, S38, and S64 represent the groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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<p>LEfSe Cladogram of Endophytes. (<b>a</b>) Endophytic Bacteria. (<b>b</b>) Endophytic Fungi. CK represents the healthy group. S31, S38, and S64 represent the groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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<p>Histogram of LDA score distribution. (<b>a</b>) Endophytic Bacteria. (<b>b</b>) Endophytic Fungi. CK represents the healthy group. S31, S38, and S64 represent the groups infected by anthracnose strains 31#, 38#, and 64# in sequence.</p>
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