INTRODUCTION
Overview
Soil quality is crucial for agricultural production as it underpins the diverse goods and services provided by terrestrial ecosystems (Chee 2004) Contaminants can degrade soil quality, negatively impacting both the quantity and quality of agricultural outputs (Baishya 2015) This has led to an increased need for monitoring and policies to enhance agricultural productivity, with initiatives such as the United Nations' goal of zero net land degradation by 2030 (Lal 2012) Soil biota, including bacteria, protozoa, and nematodes, play essential roles in soil services and are considered effective indicators of soil quality, making them valuable for monitoring systems (Karlen et al 1997) Bio-indicators are particularly sensitive to disturbances from pesticides and topsoil removal (Foissner 1999b; Asif et al 2018), which is why they are useful in assessing soil quality in agricultural ecosystems Ciliated protozoa, known for their single-celled structure and high sensitivity to environmental changes, respond rapidly to soil contamination and exhibit quick population growth, positioning them as potential indicators of soil health (Foissner 1994, 1999b; Esteban and Finlay 2010).
Ciliated protozoa are dikaryotic unicellular organisms that are relatively easy to identify due to their unique characteristics, including locomotion, infraciliature, and nuclei While studies have documented the species richness and abundance of protozoa in certain soil habitats, many species are still undescribed, highlighting the need for further research in this area (Foissner 1997a, 1997b; Foissner et al 2002; Foissner 2016).
Ciliates serve as effective bio-indicators in various environments, particularly in activated-sludge sewage treatment facilities Certain ciliate species can indicate the condition of sludge by assessing factors like Biochemical Oxygen Demand (BOD), dissolved oxygen levels in aeration tanks, and sludge retention time Their application as bio-indicators extends to evaluating water quality, highlighting their significance in environmental monitoring.
Research on the community structure of soil ciliates and their relationship with soil characteristics is limited (Ekelund and Rũnn 1994; Li et al 2010a; Li et al 2010b) Findings indicate a strong correlation between the abundance of soil ciliates and various soil physical and chemical parameters Additionally, numerous studies have demonstrated that ciliates serve as effective bio-indicators of contamination, particularly from trace metals and pesticides, in both natural and agricultural ecosystems (Foissner 1994, 1997c; Díaz et al 2006; Geisen et al 2010).
Soil protozoa, despite their significance, are less researched compared to other soil organisms like nematodes, bacteria, and fungi, with many ciliate species still undescribed To advance the field of soil protistology, it is essential to establish connections with studies focused on other soil organisms.
Soil protists, including amoebae, ciliates, and flagellates, play a vital role in supporting plant health by interacting with beneficial bacteria to mitigate plant diseases They promote the growth of plant-growth promoting bacterial species, which in turn enhances antibiotic production This synergistic relationship between protists and bacteria significantly improves disease suppression in plants, aligning with the objectives of sustainable agricultural development (Geisen et al 2018).
Natural and agricultural soils exhibit distinct properties, such as pH levels, contaminant concentrations, and organic matter content (Trivedi et al 2016) Agricultural ecosystems face more disturbances due to practices like pesticide application, ploughing, and fertilizer use, which can lead to the burial of topsoil and increased trace metals in the soil This raises important questions about how soil conditions influence ciliate diversity in these environments and whether soil ciliates can serve as effective biological indicators of soil quality This PhD research aims to investigate and compare the biodiversity of soil ciliates in both natural and agricultural soils, assessing their potential use as bio-indicators for soil quality monitoring.
Literature review
Soil quality refers to the ability of soil to perform essential functions within ecosystem limits, supporting plant and animal productivity, preserving water and air quality, and promoting human health and habitation (Karlen et al 1997).
Soil quality encompasses key aspects such as soil productivity, fertility, degradation, and environmental quality, integrating physical, chemical, and biological characteristics (De La Rosa and Sobral 2008) While "soil health" emphasizes biodiversity and ecological functions (Bruggen and Semenov 2000; Allen et al 2011), both terms are often used interchangeably (Wolfe 2006) De La Rosa and Sobral (2008) introduced a comprehensive definition that distinguishes between inherent soil quality—factors like texture and cation-exchange capacity—and dynamic soil quality, or soil health For the purpose of this project, the term soil quality will be adopted.
Figure 1.1 Graphic of soil quality concept integrating soil suitability and soil health
(adapted from De La Rosa and Sobral 2008)
1.2.1.2 Physical, chemical, and biological indicators for soil quality
Soil systems serve multiple essential functions, including providing physical, chemical, and biological environments for organisms, regulating water flow, and recycling nutrients They support biological activity and diversity crucial for plant growth and animal productivity, while also filtering and detoxifying substances The assessment of soil quality is influenced by various soil information forms tailored to specific uses, such as agriculture, forestry, and nature conservation.
Dynamic soil quality (Soil health)
Soil quality is crucial for sustainable agricultural production, as emphasized by Nortcliff (2002), who notes the need for conservative measures to maintain agricultural activities long-term The demand for effective soil quality assessment is rising, necessitating the development of standardized soil analysis methods Bio-indicators are proposed as a viable approach for this assessment, serving as measurable attributes that reflect soil quality in relation to specific functions These indicators can encompass physical, chemical, and biological characteristics of soil, making it essential to select appropriate parameters for effective evaluation A comprehensive list of soil parameters is categorized into three main groups, as outlined by De La Rosa and Sobral (2008).
According to Nortcliff (2002), selecting soil attributes involves seven key features: the purpose of land use, the relationship between soil function and indicators, measurement ease and reliability, spatial and temporal variation patterns, sensitivity to changes in soil management, comparability with routine monitoring, and the skills required for indicator interpretation The choice of soil indicators is contingent on the specific soil functions being evaluated; for instance, Gómez et al (1999) identified six soil indicators to assess the sustainability of agricultural production in an olive farm.
Table 1.1 Soil attributes used as indicators of soil quality (from De La Rosa and
Stoniness Soil structure Bulk density Porosity Soil compaction Aggregate strength and stability Soil crusting
Drainage Water retention Infiltration Hydraulic conductivity Topsoil depth
Chemical attributes Color pH Salinity Carbonate content Sodium saturation Cation exchange capacity Plant nutrients
Toxic elements Biological attributes Organic material content
Fractions of organic materials Population of organisms Nematode communities Microbial biomass Respiration rate Mycorrhizal associations Enzyme activities
Fatty acid profiles Bioavailability of pollutants
1 2.2 Protozoa and the diversity of free living ciliated protozoa
Protozoa are unicellular, eukaryotic organisms that exhibit phagotrophic behavior and can be categorized into three main groups: Ciliates, Flagellates, and Amoeboid Their small size, high abundance, and ability to form cysts enable them to disperse easily, making many free-living protozoa cosmopolitan in distribution They inhabit various aquatic and damp environments, including oceans, fresh and brackish waters, soils, and even extreme habitats like snow and ice While protozoa can thrive in diverse settings, many species show preferences for specific habitats, with certain groups, like foraminifera, being exclusive to marine environments Freshwater and marine habitats are particularly rich in free-living protozoan species.
Approximately 36,400 free-living protozoan species exist, with Mora et al (2011) suggesting a similar number may inhabit the world's oceans Despite their global presence, this diversity is relatively small compared to that of animals or plants Protozoa are the most abundant phagotrophic microorganisms in the biosphere, effectively capturing and ingesting food particles Many species in both marine and freshwater environments exhibit mixotrophy, utilizing phagotrophy and phototrophy due to the presence of endosymbiotic algae, chloroplasts, or sequestered chloroplasts, which enable photosynthesis (Stoecker et al 1987; Finlay et al 1988; Stoecker et al 1988; Esteban et al 2010).
Protozoa are vital components of ecosystems, functioning within the microbial loop due to their microscopic size, typically ranging from 20 to 200 micrometers, though some can grow several millimeters (Finlay and Esteban 2013) They primarily feed on smaller microorganisms, effectively regulating bacterial populations in freshwater, marine, and soil environments through active grazing (Fenchel 1982; Berninger et al 1991; Finlay et al 2000) Additionally, protozoa enhance microbial activity in both oxic and anoxic conditions (Finlay and Esteban).
2013) by stimulating decomposition of organic matter, and by increasing the rate of
Protozoa play a vital role in nutrient turnover and carbon cycling, significantly enhancing microbial biomass (Finlay et al 1988; Finlay 1997; Esteban et al 2006; Geisen et al 2018) Their presence in the rhizosphere can increase plant biomass by 30-80% (Bonkowski et al 2000), making them essential for transferring biomass within microbial food webs Additionally, protozoa serve as important indicators of soil and water quality (Esteban et al 1991; Foissner 1994, 1997c; Lee et al 2004; Madoni 2011).
The microbial loop is intricately connected to the grazer food web, highlighting the role of microorganisms in nutrient cycling In soil ecosystems, fungi serve as an essential food source for various organisms, including flagellates, ciliates, and metazoans, thereby enhancing the complexity of these interactions.
Ciliated protozoa are a unique group of protists distinguished by their dikaryotic structure, featuring a micronucleus for reproduction and a macronucleus for vegetative functions They utilize cilia, which originate from kinetosomes and consist of microtubules, microfibrils, and external ciliature, for both locomotion and feeding Additionally, ciliates possess the capability to reproduce both asexually and sexually through conjugation, facilitating genetic diversity.
8 material to be exchanged between two cells (Lee et al 1985; Lynn and Corliss 1991; Finlay and Esteban 2013)
Free-living ciliates are estimated to comprise between 3,000 and 30,000 species, indicating a relatively low global diversity due to their ubiquitous nature Many of these species can be easily cultured in laboratory settings, which enhances research opportunities in the study of ciliates.
There have been many new soil ciliate species described or re-described (Foissner
1995, 1997b, 1998; Foissner et al 2002; Foissner 2016) with species remaining undiscovered (Foissner 1997a, 2016) This demonstrates that more intensive research on the diversity of soil ciliated protozoa is needed
1.2.3 The influence of soil characteristics on ciliated protozoa
1.2.3.1 The influence of soil physical parameters on ciliated protozoa
Research shows that soil physical properties, including texture, moisture, temperature, and compaction, significantly influence the populations of protozoa and ciliates in the soil (Cowling 1994; Ekelund and Rứnn 1994; Foissner 1999b; Li et al 2010a).
Soil texture, structure, and moisture are crucial factors influencing soil protozoa, particularly ciliates, as demonstrated by Ekelund and Rứnn (1994) The presence of distinct protozoan groups can vary within different soil aggregates due to restricted movement between soil particles at low moisture levels (Vargas and Hattori 1990) However, when soil moisture is sufficiently high, it enhances the mobility and dispersal of protozoan communities Consequently, soil moisture is recognized as the most significant factor affecting the abundance of soil ciliates (Li et al 2010a).
Soil temperature significantly influences soil protozoa, as highlighted by Cowling (1994) An experiment by Darby et al (2006) demonstrated that the optimal temperature range for the growth of desert protozoa is crucial for their development.
Research aim and objectives
There is no doubt that ciliates have a strong connection to soil properties However, previous studies only investigated a limited range of soil properties (Li et al 2010a;
Research on the impact of soil properties on soil ciliates is limited, highlighting a need for further studies in this area Additionally, the biodiversity of soil ciliates requires attention, as many species remain unidentified or have not been thoroughly described since their initial discovery.
This study aims to explore the abundance and species richness of ciliated protozoa in both natural and agricultural soils, assessing the relationship between physio-chemical soil properties and ciliate occurrence By investigating soil ciliate biodiversity and analyzing soil characteristics, the research seeks to establish the potential of soil ciliates as bio-indicators of soil quality Controlled experiments were conducted to evaluate the effects of specific contaminants, including copper, glyphosate, and cypermethrin Additionally, the taxonomy of ciliate species was examined to identify rare and new species, thereby enhancing the understanding of ciliate biodiversity in soil ecosystems.
The objectives of the current study were:
1 To determine the soil properties in three study sites, i.e., one natural ecosystem and two agricultural ecosystems (Chapter 3);
2 To investigate the abundance and richness of soil ciliated protozoa in the three study sites and to determine correlations between soil ciliates and soil properties in order to determine if soil ciliates are bio-indicators of soil quality (Chapter 4);
3 To assess the impact of specific contamination factors on soil ciliate abundance and species richness (Chapter 6);
4 To examine the taxonomy of soil ciliate species in the three study sites and to identify rare and new species (Chapter 5).
MATERIALS AND METHODS
Study sites
The present study focused on three selected sites: East Stoke Fen Nature Reserve as a natural site, and Vicarage Farm and Corfe Castle Farm as agricultural locations For each site, two 1 x 2 m plots were established, with their positions determined through a random walk method utilizing distances and bearings generated by the RANDBETWEEN function in Excel.
Figure 2.1 Locations of study sites: East Stoke Fen, Vicarage Farm and Corfe
Castle Farm near Wareham (UK) Inset: approximate location of the study area in the UK
Table 2.1 Experimental Sites in Natural and Agricultural ecosystems
Study site Plot Grid reference Plot size Use
East Stoke Fen Plot 1 SY 86565 86612 1 x 2 m Nature reserve
Vicarage Farm Plot 1 SY 79690 81683 1 x 2 m Agricultural production Plot 2 SY 79680 81660 1 x 2 m
Corfe Castle Farm Plot 1 SY 96425 83342 1 x 2 m Agricultural production Plot 2 SY 96437 83353 1 x 2 m
East Stoke Fen Nature Reserve - Natural site
East Stoke Fen Nature Reserve, managed by the Dorset Wildlife Trust and owned by the Freshwater Biological Association, spans 4.5 hectares in the River Frome floodplain east of Wool, Dorset, United Kingdom The reserve features diverse terrain, including reed-marsh, wet deciduous woodland, and oak copse, with much of the area shaded by trees.
Figure 2.2 Natural study site at East Stoke Fen Nature Reserve (Dorset, UK)
Sampling took place at two farms in the UK: Vicarage Farm in Winfrith Newburgh, which cultivated spring oats and winter wheat, and Corfe Castle Farm in Wareham, which grew spring barley and winter turnip Vicarage Farm is situated at an altitude of 151 meters above sea level, while Corfe Castle Farm is at 12 meters.
Figure 2.3 The agricultural study site at Corfe Castle Farm in Wareham (UK) (Image: author’s own collection)
Figure 2.4 The agricultural study site at Vicarage Farm in Winfrith Newbourgh (UK)
Soil sampling
Soil samples were collected monthly from a natural site between January 2016 and April 2017, while samples from agricultural sites were gathered from May 2016 to April 2017 Following April 2017, soil sampling continued on a monthly basis.
During the 23rd season, specifically in July, October, and December 2017, samples were collected from all study sites Each sampling occurred in the morning, and the samples were promptly transported to the River Laboratory in Wareham, conveniently located near the sampling locations.
Soil samples were collected from both natural and farm sites by randomly designating two 1 x 2 m plots at each location A stainless steel soil corer, 4 cm in diameter, was utilized to extract topsoil samples to a depth of 5 cm, ensuring the inclusion of the litter layer.
During each sampling event, eight core samples were randomly collected from each plot to ensure representation The soil obtained was extracted from the corer and placed in a labeled plastic bag, creating a bulk sample for that specific plot.
Soil and air temperatures in each sampling point at the time of sampling were recorded by using a Checktemp ® LC thermometer (Hanna Instruments, Rhode Island, USA).
Ciliate species richness and abundance
2.3.1 Determination of soil ciliate abundance
Methods for retrieving ciliates from soil samples are categorized into direct and indirect approaches (Foissner 1987) Direct methods are effective for estimating ciliate abundance only when utilizing fresh, moist soil, as active ciliates are scarce and reliant on soil moisture (Adl and Coleman 2005) In contrast, indirect methods enable the activation of ciliate cysts formed under dry environmental conditions, thereby allowing for the assessment of ciliate diversity across various soil types One such indirect method is the Most Probable Number (MPN) technique, which involves adding water, organic substances, mineral elements, and/or bacterial food sources to enrichment media to promote the growth of soil protozoa (Foissner).
MPN methods, while useful for estimating protozoan populations, have significant drawbacks, such as the tendency to favor specific protozoan species based on the type of bacterial food provided and variability in results depending on the enrichment protocol (Foissner 1987) To address these limitations, Finlay et al (2000) introduced a more effective method for estimating the abundance of free-living protozoa in soil, which requires only the addition of filtered rainwater This approach facilitates the comparison of protozoa across different soil types, making it a valuable tool for ecological studies.
(2000) was followed to investigate abundance and richness of soil ciliated protozoa The details of the method are described below:
Upon returning to the River Laboratory, soil samples were extracted from their original plastic bags and thoroughly mixed in a clean 30 cm-diameter glass bowl to achieve homogeneity A sub-sample of approximately 50 grams was then spread in a 15 cm diameter glass Petri dish and left to dry at room temperature (18-22 °C) for six days After drying, the soil was passed through a 4 mm sieve to eliminate larger debris, such as stones, resulting in what is now referred to as "air-dried soil," which was utilized for ciliate-related research The remaining soil was reserved for analyzing soil characteristics.
To facilitate the development of ciliates, 5 g of air-dried soil was placed into a 5 cm diameter sterile plastic Petri dish This was replicated three times for each sample
Filtered rainwater was added to samples to create a slurry, using Whatman syringe filters with a 0.2 µm pore diameter to remove ciliates and other microbes The slurry samples were incubated in the dark at 15°C, with ciliates being examined after 4 and 10 days of incubation.
After 4 days and 10 days of incubation, the wet soil prepared as above was pressed to release water A 50 àl sample from the water runoff was taken and placed on to a glass Sedgewick-Rafter Chamber Ciliate numbers in the 50 àl subsample were determined under a compound microscope at a magnification of 40-125 X This was repeated five times for each Petri dish of soil
To calculate the number of ciliate cells per gram of oven-dried soil, use the formula: g -1 = a x (V1 + V2) / (50 x W) In this equation, 'a' represents the count of ciliate cells from a 50 µL sample of water runoff, while 'V1' is the volume of rainwater added to 5 grams of air-dried soil.
V2: Volume of water determined in 5 g of air-dried soil
W: oven- dried weight of 5 g of air-dried soil
During the counting process, the species richness of ciliates was determined by identifying observed ciliates using taxonomic works such as Kahl (1935); Foissner
In order to enhance the identification of ciliates, samples were subjected to enrichment cultures after counting, allowing for the growth of species not initially observed This method also ensures an adequate supply of cells for detailed observations and silver impregnations Enrichment cultures are created by adding approximately 5 grams of rewetted soil into 50 mL sterile culture flasks containing a specific medium, as outlined in Table 2.2.
Table 2.2 Medium used to prepare enrichment cultures of soil ciliates
Soil from the Petri dish ~ 5 g of rewetted soil
*SES (Soil Extract added Salts) components (http://www.ccap.ac.ukmedia/SES.pdf):
All ciliates that grew in the flask were identified under a compound microscope using a combination of glass micro-pipetting and Silver-staining techniques (see below).
Various impregnation and staining techniques, including vital staining, protargol, silver-carbonate, and the dry/wet silver nitrate method, are employed to highlight the distinct characteristics of ciliates (Fernández-Galiano 1994; Lee and Soldo 1992; Foissner 2014) Among these, the ammoniacal silver carbonate impregnation method is particularly effective in revealing kinetosomes and other essential features such as the kinetodesmal fibre and nuclear apparatus This method is not only efficient but also accelerates the identification process of ciliate species, making it the preferred technique in the current study (Fernández-Galiano 1994).
The procedure followed was adapted from Fernández-Galiano (1994) The following components are added to a glass beaker (Volume 50 mL) * :
2 mL of the sample to be studied
1 mL of ammoniacal silver carbonate solution
* Order of components added as listed from top to bottom
The beaker was then stirred to mix the components, and placed into a water bath at
The beaker was heated to 55 °C in a water bath until the liquid attained a brandy-like color Subsequently, the liquid was transferred into a glass evaporation dish containing approximately 50 mL of distilled water The resulting suspension was allowed to settle, followed by a gentle swirl to concentrate the cells at the center of the capsule Finally, a glass micropipette was utilized to transfer the concentrated cells onto a microscope slide, which was then covered with a coverslip for observation under a compound microscope.
The use of formalin for cell fixation is unsuitable for certain ciliates, such as hypotrichs, as it causes them to burst (Foissner 2014) Consequently, for hypotrichs and other delicate ciliates, the protargol method outlined in Foissner (2014) was employed This procedure was adapted from the original work by Foissner (2014).
Ciliate cells in the sample were fixed in a 1:1 ratio with commercial Bouin’s fluid for 20 minutes The fixed cells were transferred to a 15 mL polypropylene centrifuge tube and concentrated by washing three times with tap water to eliminate Bouin’s fluid, using centrifugation at 2000 rpm for two minutes During this washing step, enough tap water was added to reach a total volume of 10 mL, and the mixture was shaken vigorously After centrifugation, the supernatant was carefully decanted while retaining 1 mL in the tube to avoid disturbing the loosely pelleted ciliates.
At the final washing stage, approximately 0.3 mL of concentrated and fixed cells remained in the tube A small drop of albumin-glycerol was applied to the center of a clean glass slide, followed by the addition of the concentrated cells The mixture was thoroughly blended using a mounted needle and spread across the middle third of the slide Typically, eight clean glass slides were prepared for each sample The slides were then left to dry overnight at room temperature Once dry, they were placed in an eight-slot staining jar filled with ethanol.
To prepare ciliate cells for observation, immerse the slides in 95% ethanol for 20 minutes, followed by rehydration in 70% ethanol and two 5-minute washes with tap water Next, treat the slides with 0.2% potassium permanganate for 2 minutes, then transfer them to 2.5% aqueous oxalic acid for 3 minutes After washing the slides twice with tap water and once with distilled water for 3 minutes each, place them in a warm (60°C) 0.5% protargol solution for 40-60 minutes, depending on the ciliate species Once stained, briefly dip a slide in tap water, add ordinary developer, and observe under a compound microscope When the oral structures of the ciliate cells become faintly visible, remove the slide, pour off excess fluid, and rinse it in a container with 500 mL of tap water.
To halt the development of ciliate infraciliature, the slides were treated with 5% sodium thiosulphate for 5 minutes This procedure was consistently applied to all remaining slides Following this treatment, each slide was washed three times with tap water, with each wash lasting 3 minutes The slides were then sequentially transferred through 70% and 100% ethanol.
The samples were fully dehydrated and underwent two 10-minute transfers through xylene Afterward, the slides were mounted using a synthetic neutral medium and covered with a 24x40 mm cover glass It is essential to allow the slides to harden overnight before conducting any observations.
Determination of soil physical characteristics
Soil texture refers to the relative proportions of sand, silt and clay in the soil (White
Soil texture significantly influences the physio-chemical properties of soil, impacting the communities that inhabit it (White, 1997) This study employed Rowell's (1994) method to analyze soil structure, starting with a wet oxidation process to eliminate organic components that bind mineral particles, especially clays A 10 g sub-sample of fine earth fraction soil was treated with 30 mL of 6% H2O2, heated to 40 °C until frothing subsided, followed by the addition of 70 mL of 6% H2O2 and heating to 60 °C for an hour Finally, the temperature was raised to 100 °C until frothing ceased, ensuring complete oxidation of organic material.
The next step in the process involved using a blender and a chemical anti-flocculent to effectively separate and disperse the mineral fraction of the soil The beaker from the wet oxidation phase was filled to approximately 100 mL, and 10 mL of a 10% sodium hexametaphosphate dispersant was added The mixture was blended for 45 seconds, followed by a 15-second pause, and then blended again for another 45 seconds Finally, any soil residue from the blender was rinsed with distilled water into the beaker.
The sample was then wet sieved through a 63 àm sieve contained in a funnel into a
500 mL cylinder using a spray of water Spraying was stopped when 450 mL of water had collected in the measuring cylinder The volume in the cylinder was filled
29 up to 500 mL by water used to wash out the funnel The cylinder was stoppered and used to determine silt and clay fraction
The sand fraction collected on the sieve was placed into a pre-weighed 250 mL beaker After carefully decanting the excess water, the sand was dried at 105°C for a minimum of 24 hours.
To accurately measure the silt and clay fractions, a sealed measuring cylinder was inverted 10 to 20 times to evenly distribute the particles in the water column Subsequently, 20 mL of the solution was extracted from a depth of 10 cm and transferred to a 100 mL beaker for drying in an oven at 105 °C for a minimum of 24 hours After 8 hours, an additional 20 mL sample was taken from the same depth, which contained only clay, as the silt had already settled This method ensures that the clay content in the sampled solution accurately reflects the overall clay content in the cylinder at that time, and the solution was again dried in the oven at 105 °C for at least 24 hours.
To determine the weight of the sand fraction, the weight of an empty beaker was subtracted from the weight of the beaker containing the dried sand, resulting in the weight denoted as S The weight of the dried silt and clay fractions, recorded as M, was calculated by subtracting the empty beaker's weight from the beaker with the fractions and multiplying by 25 (500 mL/20 mL) Similarly, the mass of the clay fraction, denoted as C, was obtained using the same method The total weight of mineral material in the soil sample, referred to as Wt, was calculated by summing the weights of the sand, silt, and clay fractions (Wt = S + M + C) Additionally, the weight of the silt fraction was determined by subtracting C from M (M - C).
Percent sand, silt and clay can be calculated by % = (fraction weight/Wt) x 100
Determination of chemical characteristic
The pH of a solution is defined as: pH= -log10[H + ]
Soil pH can be measured by mixing air-dried soil with distilled water, 0.1 M CaCl2, or 1 M KCl solution in specific ratios (Hendershot et al 2007; Kabała et al 2016) In this study, a 10 g sub-sample of air-dried fine soil was combined with 25 mL of distilled water in a 50 mL beaker After thorough stirring, the mixture was allowed to stand for 15 minutes The pH meter was calibrated using a pH 7.0 buffer solution prior to measuring the soil pH in the suspension Each soil sample's pH was measured three times with an H-series minilab pH meter (HACH, USA).
2.5.2 Soil oven-dry (OD) weight
To determine the soil oven-dry weight, a precise 5 g sub-sample of fresh fine soil is placed in a 50 mL beaker and then transferred to a laboratory oven set at 105°C After being left in the oven for a minimum of 24 hours, the dry soil is reweighed The oven-dry weight is calculated using the formula: % OD = (dry weight/wet weight) x 100.
This was repeated three times for each soil samples (Rowell 1994)
Soil moisture was also recorded here by: Moisture (%) = 100 % - % OD
2.5.3 Determination of total major and trace elements in soils by aqua regia acid digestion
Aqua regia acid digestion is utilized for the analysis of major and trace elements in soils, including heavy metals, aluminum, calcium, magnesium, potassium, phosphorus, and sulfur This method is adapted from the approach established by McGrath and Cunliffe in 1985.
A 0.300 g (+/- 0.02 g) sub-sample of fine earth fraction soil was placed into a boiling tube The exact weight of the soil was recorded Aqua regia was created by adding
To prepare the soil sample for analysis, 9 mL of 37% HCl and 3 mL of 70% HNO3 were added to a tube containing Primar Plus trace metal grade acids from Fisher Scientific The mixture was swirled to ensure thorough mixing and then placed in a cold heating block, allowing it to stand at room temperature for at least two hours to facilitate the cold digestion of organic matter Given the high organic content of the Fen soil, extended digestion was necessary to prevent frothing during heating Subsequently, the tube was heated overnight at 60°C, followed by an increase to 105°C for one hour to complete the soil digestion Finally, the temperature was raised to 150°C to evaporate the contents, and heating was halted once the tube contents were dried After cooling to room temperature, the dried tube was collected for further processing.
To re-suspend the digest, HNO3 was added and the resulting suspension was filtered through Whatman® No 42 filter paper into a 15 mL polypropylene centrifuge tube This process was repeated three times for each soil sample, and blanks along with standard reference materials (NRC TH-2) were included in each digestion All samples were subsequently analyzed using ICP-OES (Varian Vista-Pro, Varian Pty, Australia).
Concentration in soil (mg kg -1) ) Where:
ICP = value recorded by ICP-OES system
2.5.4 Determination of extractable (bioavailable) elements in soils
This method utilized a neutral pH salt solution, specifically CaCl2, to assess the concentration of metals and other substances like phosphates in soils that are potentially available for uptake by organisms, as outlined by Houba et al (1990) and Novozamsky et al.
Bioavailable metals in soil are present as free metal ions in the soil solution or absorbed to cation exchange sites on clays and organic matter This study utilized an extraction method based on Houba et al (1990), where a 2 g sub-sample of fine earth fraction soil was combined with 20 mL of 0.01 M CaCl2 in a 30 mL polypropylene tube The mixture was shaken horizontally at 125 rpm for 3 hours at room temperature (20-22°C) Following this, the soil suspension was filtered through Whatman® No 42 filter paper into a 15 mL centrifuge tube and stored at -18°C until analysis Each sample was analyzed in triplicate using ICP-OES (Varian Vista-Pro, Varian Pty, Australia).
Concentration in soil (mg kg -1) ) Where:
ICP = value recorded by ICP-OES system
2.5.5 Determination of organic matter by loss on ignition
There are three common methods for determining soil organic carbon: loss on ignition, the Walkley and Black oxidizable carbon method, and dry combustion (Konare et al 2010) Among these, loss on ignition is the simplest and most convenient (Konare et al 2010) This method involves placing approximately 2 grams of fine oven-dried soil in a crucible and heating it to 450°C for 10 hours After ignition, the soil is cooled in a desiccator before being re-weighed to calculate soil organic matter using the loss on ignition technique (Rowell 1994).
2.5.6 Determination of total ammoniacal nitrogen and nitrate nitrogen
Extraction of ammoniacal and nitrate with 2 M potassium chloride
The extraction of soil ammoniacal and nitrate using 2 M KCl is a widely accepted method due to its simplicity and speed (Rowell 1994) Furthermore, the addition of KCl does not alter the levels of exchangeable NH4+.
The NO3 nitrogen determination method follows the procedure outlined by Keeney and Nelson (1982) In this process, 4 grams of fresh soil is combined with 20 mL of 2 M KCl in a sterile tube After stoppering, the mixture is shaken on an orbital shaker for one hour, and the resulting soil suspension is filtered using Whatman® No filter paper.
42 into a clean tube This extraction solution was used for determine concentrations of ammoniacal and nitrate nitrogen Three replications were prepared for each sample
Determination of extractable nitrate nitrogen via vanadium (III) reduction
Several methods exist for determining nitrate-N, including nitrate reductase, specific ion electrode, chemiluminescence, and the copperized cadmium reduction method Among these, the colorimetric method using vanadium reduction and Griess reagents is considered equally valid and is widely used This sensitive procedure is advantageous as it does not necessitate specialized equipment, costly reagents, or lengthy processes.
Vanadium (III) in HCl solution significantly enhances the reduction of nitrate to nitrite and/or nitric oxide, which are then detected using Griess reagents, consisting of sulphanilamide and N-(1-naphthyl)-ethylenediamine, to form a red dye The reagent solution, containing vanadium and Griess reagents, was combined with nitrate samples in a ratio of 4 mL of reagent to 80 µL of extractant in cuvettes Absorbance was measured at 540 nm with a UV/Vis spectrophotometer (Varian Cary 50, Varian Pty, Australia) after a 12-hour color development period at room temperature Calibration was performed using nitrate standards ranging from 0 to 5.
10, 25 and 50 mg L -1 was made up in 2 M KCl to calibrate the UV/vis and allow conversion of absorbance into concentration
Total ammoniacal nitrogen (TAN) by the Salicylate-Hypochlorite method
The indophenol blue method is a widely used technique for quantitatively determining total ammoniacal nitrogen (Ma et al., 2014) However, its use is discouraged due to the requirement of phenol, a highly hazardous chemical (Molins-Legua et al.).
The salicylate-hypochlorite method is a safer alternative to the indophenol method, utilizing the reaction of ammonia with salicylate and hypochlorite in the presence of sodium nitroprusside (Bower and Holm-Hansen, 1980) This method has demonstrated results comparable to the traditional indophenol blue method, with slightly improved accuracy (Le and Boyd, 2012).
The procedure utilized the method established by Le and Boyd (2012), beginning with the dilution of 1 mL of filtered 2 M KCl soil extracts in 4 mL of analytical grade water within a clean tube Following this, 0.6 mL of salicylate catalyst solution and 1 mL of alkaline hypochlorite solution were added and thoroughly mixed with the sample The resulting mixture was then stored in a low light environment for one hour to allow for color development, as the pigments produced in this method are sensitive to light Finally, the absorbance was measured at 640 nm using a UV/vis spectrophotometer.
Soil microbial properties
Glycosidase or glycoside hydrolase is part of a group of hydrolase enzymes that hydrolyse the glycosidic bonds in carbohydrates The general equation of the reaction is (Eivazi and Tabatabai 1988):
β-glucosidase, a crucial enzyme in the glycosidase group, plays a vital role in the soil carbon cycle and serves as a bio-indicator for soil biological fertility (Zhang et al 2011) This enzyme is essential in the final stage of cellulose degradation, where it catalyzes the hydrolysis of β-glycosidic bonds in cellobiose, ultimately releasing glucose (Singhania et al 2013).
To determine activity of β-glucosidase, the assay method described by Tabatabai
(1994) was used This method works by measuring the amount of p-nitrophenol
The release of p-nitrophenol (pNP) occurs when β-glucosidase enzymes cleave the bond between this compound and the carbohydrate moiety of p-nitrophenyl-β-glucoside (pNPG) To conduct the experiment, a 1 g sub-sample of fresh soil, sieved to 2 mm, was placed in a 50 mL conical flask Subsequently, 0.25 mL of toluene and 4 mL of Tabatabai’s modified universal buffer (pH 6) were added After swirling the flask briefly, 1 mL of pNPG was introduced to initiate hydrolysis reactions, and the flask was incubated at 37°C with shaking at 100 rpm for one hour To halt the activity of the β-glucosidase enzyme, 1 mL of 0.5 M CaCl2 and 4 mL of Tris(hydroxymethyl)aminomethane (THAM) buffer (pH 12) were added, followed by filtering the soil suspension through a Whatman® filter.
No 2 filter paper into a 30 mL polypropylene tube This process was repeated three times for each soil sample A blank was prepared as above but which excluded the
To account for background absorbance from soil matter, 35 addition of pNPG was utilized Absorbance measurements of all samples were conducted at a wavelength of 405 nm using a Varian Cary 50 UV/vis spectrophotometer Standards were created from pNP to convert the absorbance readings into micromoles of p-nitrophenol released per gram of soil per hour.
Calculations àmol p- Nitrophenol released g -1 soil h -1 = / 139.11
Fs: concentration of p-nitrophenol in sample
Fb: concentration of p-nitrophenol in the blank
Wts: weight of soil sample
Wtb: weight of blank soil
Sd: dry matter content of soil expressed as a decimal (determined as in 5.2)
Phosphatases are a diverse group of enzymes that catalyze the hydrolysis of phosphate-ester bonds, with phosphomonoesterases playing a crucial role in the mineralization of organic phosphorus (Anna et al 2012) These enzymes are categorized into two types based on their optimal pH levels: acid phosphatases function best at pH 6.5, while alkaline phosphatases operate at pH 11.0 (Eivazi and Tabatabai 1977; Pang and Kolenko 1986) This study focuses on acid phosphomonoesterase, as agricultural soil pH is often adjusted to 6.5 to enhance nutrient availability.
The phosphomonoesterase activity was determined using the method established by Tabatabai (1994), which involves measuring the release of p-nitrophenol (pNP) during the hydrolysis of p-nitrophenyl phosphate (pNPP) by phosphomonoesterases A 1 g sub-sample of fresh soil, sieved to 2 mm, was placed in a 50 mL conical flask, followed by the addition of 0.2 mL of toluene and 4 mL of a modified universal buffer (pH 6.5) as per Tabatabai’s protocol.
In a controlled experiment, 36 mL of soil suspension was combined with 1 mL of pNPP to initiate hydrolysis reactions, followed by thorough swirling and incubation at 37°C with shaking at 100 rpm for one hour To halt the enzyme reactions, 1 mL of 0.5 M CaCl2 and 4 mL of 0.5 M NaOH were added The resulting mixture was then filtered through Whatman® No 2 filter paper into a sterilized 30 mL tube, with this procedure repeated three times for each soil sample A blank sample, prepared without pNPG, accounted for any background absorbance from soil matter Absorbance measurements were conducted at 405 nm using a Varian Cary 50 UV/vis spectrophotometer, and standards were created from pNP to quantify the absorbance in terms of μmol p-nitrophenol released per gram of soil per hour.
Calculations àmol p- Nitrophenol released g -1 soil h -1 = / 139.11 Where:
Fs: concentration of p-nitrophenol in sample
Fb: concentration of p-nitrophenol in the blank
Wts: weight of soil sample
Wtb: weight of blank soil
Sd: dry matter content of soil expressed as a decimal (determined as in 5.2)
Total microbial activity is a crucial indicator of organic matter turnover and soil quality in natural environments A widely used method to assess this activity is fluorescein diacetate (FDA) hydrolysis, which is both accurate and straightforward Various bacterial and fungal extracellular and membrane-bound enzymes can break down the fluorescein-acetate bonds, resulting in the release of fluorescein, a bright yellow dye The concentration of fluorescein produced is measured by its absorbance at 490 nm, with higher levels indicating increased microbial activity in the soil.
This method adapted the assay using FDA developed by Adam and Duncan (2001)
In a 50 mL conical flask containing a 2 g sub-sample of fresh soil (sieved to 2 mm), 15 mL of 60 mM potassium phosphate buffer at pH 7.6 was added, followed by the addition of 0.2 mL of a 1000 μg mL -1 FDA stock solution to initiate the reactions.
The flask was sealed and placed in an orbital incubator shaker at 30°C and 100 rpm for 20 minutes To halt the reactions, 15 mL of a 2:1 chloroform/methanol solution was added The mixture was then transferred to a 50 mL centrifuge tube and centrifuged at 2000 rpm for 3 minutes to collect the supernatant, which was filtered through Whatman® No 2 filter paper into a 30 mL polypropylene tube This process was repeated three times for each soil sample, with a blank prepared without FDA stock to correct for background absorbance from soil matter The fluorescein concentrations in the soil extracts were measured by absorbance at 490 nm using a Varian Cary 50 UV/vis spectrophotometer.
To assess microbial activity in soil, the release of fluorescein is measured in milligrams per gram of dry soil By determining the soil's moisture content, microbial activity can be calculated using the formula: μg fluorescein released g⁻¹ soil h⁻¹ = x 3.
Fs: concentration of fluorescein in sample
Fb: concentration of fluorescein in the blank
Wts: weight of soil sample
Wtb: weight of blank soil
Sd: dry matter content of soil expressed as a decimal (determined as in 5.2).
Data analysis
Statistical analysis was conducted with SPSS and R software
A two-way ANOVA was performed in R version 3.4.2 to investigate the differences between a dependent variable and two independent variables The datasets were assessed for homogeneity of variance using Levene’s test, and when this assumption was violated, appropriate adjustments were made.
38 data was subject to adjustment using the Heteroscedasticity-Consistent Covariance Matrix Estimator HC3 described by MacKinnon and White (1985) using the R car package (Fox and Weisberg 2011)
A one-way ANOVA test was performed using SPSS version 20 to assess the differences between a dependent variable and an independent variable The data sets underwent analysis for homogeneity of variance through Levene’s test In instances where the assumptions were not satisfied, data were either log10 transformed or analyzed using Welch’s robust ANOVA as necessary.
Correlations between variables were determined by Spearman’s rank order method, conducted by using SPSS vs 20