Next Article in Journal
Mathematical Model of the Migration of the CO2-Multicomponent Gases in the Inorganic Nanopores of Shale
Next Article in Special Issue
Performance of CO2 Adsorption on Modified Activated Carbons Derived from Okara Powder Waste: Impacts of Ammonia Impregnation
Previous Article in Journal
Thermal Safety Study of Emulsion Explosive Matrix under the Coupled Effects of Environmental Pressure and Bubble Content with Internal Heat Source
Previous Article in Special Issue
Sustainable Napier Grass (Pennisetum purpureum) Biochar for the Sorptive Removal of Acid Orange 7 (AO7) from Water
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Processes
Volume 12
Issue 8
10.3390/pr12081678
processes-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

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

Effects of Biochar-Amended Composts on Selected Enzyme Activities in Soils

by
Faraj Zaid
1,2,
Nasruddeen Al-Awwal
1,*,
John Yang
1,*,
Stephen H. Anderson
3 and
Bouzeriba T. B. Alsunuse
1,4
1
Department of Agriculture & Environmental Sciences & Cooperative Research, Lincoln University of Missouri, Jefferson City, MO 65102, USA
2
Department of Chemical Engineering, Sirte University, Sirte P.O. Box 674, Libya
3
School of Natural Resources, University of Missouri-Columbia, Columbia, MO 65211, USA
4
Department of Plant Production, Agricultural College, Benghazi University, Alkufra P.O. Box 1308, Libya
*
Authors to whom correspondence should be addressed.
Processes 2024, 12(8), 1678; https://doi.org/10.3390/pr12081678
Submission received: 16 July 2024 / Revised: 5 August 2024 / Accepted: 6 August 2024 / Published: 11 August 2024
(This article belongs to the Special Issue Application of Biochar in Environmental Research)
Browse Figures
Versions Notes

Abstract

:
This study examines the effect of biochar as an agricultural soil supplement on soil quality indicators, specifically enzyme activity in Missouri regions. While the benefits of biochar on soil bulk density, soil organic carbon, and infiltration have been established, its effect on soil enzyme activity has remained underexplored in this region. A three-year field investigation was conducted with six treatments (compost, biochar, compost + biochar, biochar + compost tea, fescue, and control) to evaluate the effects on enzymes such as β-glucosidase (BG), acid and alkaline phosphatases (ACP-ALP), arylsulfatase (ARS), dehydrogenases (DG), arylamidase (AMD), cellulase (CLS), and urease (URS). Furthermore, soil pH, organic matter (OM), and cation exchange capacity (CEC) were determined. The results showed that compost and biochar treatments considerably increased soil enzyme activity compared to other treatments, with nitrogen application further increasing enzyme activity. Soil pH, OM, and CEC were all important determinants in determining enzyme activity, with BG demonstrating strong positive associations with ACP and AMD (99.5%). This study shows that compost and biochar amendments significantly improve soil physicochemical and biological properties, thereby enhancing soil health and assisting farmers’ sustainable soil management practices.

1. Introduction

In recent years, compost and biochar have received wide attention and their effects on soil properties have been studied [1]. Compost and biochar are used as soil amendments to increase soil organic matter content [2] and enhance soil fertility [3]. Additionally, adding compost and biochar to soil can improve its structural stability, nutrient availability, plant nutrient uptake, and bulk density. This will promote soil biological activity, reduce erosion losses, and improve water-holding capacity [4].
Biochar alone may also improve soil physical properties [5], increase soil pH [6] and cation exchange capacity (CEC) [7], enhance nutrient retention, alter soil microbial populations, and augment crop yields [8]. Biochar is produced by thermal decomposition of biomass under oxygen-limited conditions [9]. The initial characteristics of the feedstock utilized, along with the temperatures attained during pyrolysis, might affect the chemical and physical characteristics of the biochar [10,11]. High-temperature pyrolysis produces biochar, which typically has a large surface area, is effective as an adsorbent [12], and is strongly fragrant and resistant to breakdown [13]. In contrast, low-temperature pyrolysis favors greater recovery of C and nutrients that are increasingly lost at higher temperatures [14,15]. Low-temperature biochar, which has a less condensed carbon structure, is expected to have greater reactivity in soils than high-temperature biochar [14,16,17].
Furthermore, when pyrolyzed, plant species with many large-diameter cells in their stem tissues can lead to greater macropore quantities in the biochar particles. The capacity of biochar to adsorb larger molecules such as phenolic compounds can be improved by increasing the number of macropores [18]. Biochar is more resistant to microbial degradation than uncharred organic matter because of its macromolecular structure, which may contain aromatic C [19]. Biochar is thought to have a long residence time in soil [20]. However, its recalcitrance and physical nature are significant obstacles to quantifying long-term stability [21]. Biochar has also been shown to improve soil fertility. Biochar application has been shown to improve soil fertility by increasing the pH of acidic soils [22], retaining certain nutrients due to their surface electrical charges, thereby increasing water retention [23,24], reducing nutrient leaching [25], and retaining cations and natural matter [26]. Applying biochar to soil changes its physicochemical qualities and promotes soil microbial activity, improving soil quality and plant performance. Investigating how soil microbial populations and soil enzymes respond to biochar additions is critical for gaining a better understanding of the interactions between biochar, soil, and plants [27].
Nutrient cycling in soils involves biochemical, chemical, and physiochemical reactions with biochemical processes mediated by microorganisms, plant roots, and soil animals [28,29]. It is well known that enzymes, which are proteins with catalytic capabilities, catalyze biological reactions. Enzymes are catalysts, meaning that they are substances that do not undergo permanent alteration and cause chemical reactions to proceed at faster rates. In addition, they are specific to the chemical reactions in which they participate. Enzymes are denatured by elevated temperatures and extreme conditions (pH < 3 or >9) [30]. Their physiochemical state and influence on chemical reactions are markedly dependent on pH, ionic strength, temperature, and the presence or absence of inhibitors or activators [28]. Biochar addition to the soil is likely to proceed only when sufficient improvements in soil performance and productivity are perceived or assured [31]. Soil enzymatic activity is used to indicate soil health [32]. The objectives of this study were to investigate the effects of compost and biochar treatments on selected soil enzyme activities—BG, ACP and ALP, ARS, DG, AMD, CLS, and URS—as these have not been investigated at a regional level in the State of Missouri. Therefore, this necessitates regional research, and it will assist in translating the research findings into practical guidelines for small farmers. The hypothesis is that the biochar amendment will affect the enzyme activity positively.

2. Materials and Methods

2.1. Study Site

The Big Horn Field, operated by the Lincoln University of Missouri, was selected as the study site. The site is located at 38°34′26″ N and 92°17′19″ W. The soil at the site was a Harvester silt loam (fine-silty, mixed, superactive, and nonacidic). mesic oxyaquic units, with slopes ranging from 3 to 15 percent. The annual precipitation averages 965.2 mm with most occurring from April through August and the mean annual temperature is 12.8 °C [33]. The study consisted of six plots (26 × 76 m each) with a split-plot design (five treatments and one control). Treatments were randomly assigned to the plots in triplicate. The treatments were established in the fall, and tomatoes (Solanum lycopersicum) were grown annually for three years. The treatments included compost, biochar, compost + biochar, biochar + compost tea, fescue, and control. The biochar was prepared by slow pyrolysis of air-dried switchgrass (Panicum virgatum) between 400 °C and 600 °C in a custom-made pyrolizer for 3 h. A 2 mm sieve was used to filter the biochar sample. The switchgrass was grown and harvested at Lincoln University Allen T. Busby Agriculture Farm, an organic-certified farm. The application rate of biochar for the soil in Central Missouri ranged from 2% to 4% by weight. Using 2% biochar, the application rate to a depth of 15 cm on soil with a bulk density of 1.3 g/cm3 is 39 metric tons per hectare. A custom-made water culture system was assembled using the principles of the Actively Aerated Compost Tea (AACT) brewing system to produce composted tea. The test system had two compartments: one housing compost tea brewed from worm castings, and another with castings blended with biochar. The compost tea was brewed for 36 h in 0.038 metric tons of buckets using exposed tap water from the public water supply; the water was allowed to be oxygenated for approximately 24 h to remove chlorine before adding the biochar. A combination of 50% biochar + 50% compost tea was used as the biochar + compost tea treatment. Similarly, 50% biochar and 50% compost were used as the biochar and compost treatments, respectively. The treatments were applied to the topsoil at a depth of 15 to 20 cm using shovels to remove the soil and thoroughly mix it with 1.6 kg of the respective treatment in a wheelbarrow, before backfilling the 4-square-foot area with the soil mixture from the wheelbarrow. The tomatoes were transplanted or seeded afterward. Even though collecting soil samples in the first and second years may have provided insight into the amendments’ initial and intermediate effects, these could be less consistent due to ongoing soil changes and microbial community development. Therefore, soil samples were collected only in April, June, and August of the third year of the study. These months allow for a spread across diverse seasonal conditions and the identification of patterns or changes caused by varying environmental factors, such as early spring, early summer, and late summer, resulting in a comprehensive perspective of enzyme activity throughout time and the ability to compare enzyme activities. All plant experiments/protocols were performed in accordance with relevant institutional, national, and international guidelines and legislation.

2.2. Sample Collection and Soil Enzyme Assays

All samples were collected from the topsoil layer (0–15 cm) of each plot. Soil samples were air-dried at 37 °C for one hour, sieved with a 2 mm sieve, stored at room temperature for immediate enzyme analysis, and stored at 4 °C for later use. Soil pH values were determined using a combination of glass electrodes (soil: water ratio = 1:2.5). All soil samples were homogenized and incubated at a constant temperature for a specific time to allow optimal activity measurements. The activities of the selected enzymes were determined by multiple assays, following the procedures described in the literature. Measurements were calculated based on the quantitative release of the enzymatic product using a colorimetric method [34]. Eight enzymes were selected for this study based on their key roles in carbon, nitrogen, phosphorus, and sulfur cycling: β-glucosidase (C), acid and alkaline phosphatase (P), arylsulfatase (S), dehydrogenases (C), arylamidase (N), cellulase (C), and urease (N). The enzyme activity was determined calorimetrically in triplicate. The method described by Eivazi and Tabatabai [35] was used to assay β-glucosidase activity. Acid and alkaline phosphatases were estimated as described by Eivazi and Tabatabai [36]. The method of Tabatabai and Bremner [37] was used to assay arylsulfatase activity. A 1 g soil sample was used for individual tests, and p-nitrophenol derivative substrates were used. The samples were incubated on a mechanical shaker for an hour at 37 °C in an ideal buffer pH. Using a spectrophotometer, the generated p-nitrophenol (PNP) was then measured at specific enzyme absorbance (410–420 nm) [31,32,33]. The method described by Casida and coworkers [38] was used to assess dehydrogenase activity. Then, 1 mL of 3% aqueous 2, 3, 5 triphenyl tetrazolium chloride (TTC) solution was added to 6 g of soil, and 2.5 mL of DI water was added to each sample. The samples were incubated at 37 °C for 24 h. Then each sample was washed and filtered by adding methanol in 10 mL aliquots until the reddish color disappeared. The filtrate was diluted to 100 mL with methanol and color intensity was measured at 485 nm. Arylamidase activity was determined using the method described by Acosta-Martinez and Tabatabai (2000) [39]. The method described by Deng and Tabatabai (1994) [40] was used to assess cellulase activity. The procedure involved 1 g of soil samples incubated with 50 mM acetate buffer (pH 5.5), carboxymethyl cellulose (CMC), and toluene at 30 °C for 24 h. After incubation, the reducing sugar was measured using the Somogyi-Nelson method. The method of Tabatabai and Bremner [37] was again used to assay urease activity. Then, 5.0 g soil was placed in a 50 mL Erlenmeyer flask, and 0.2 mL of toluene and 9 mL of Tris (hydroxymethyl) aminomethane (THAM) buffer were added into the flask containing the soil sample and swirled for a few seconds to mix the contents. Next, 1 mL of 0.2 M urea solution was added, and the flask was swirled again for a few seconds and placed in an incubator at 37 °C. After 2 h, the stopper was removed and approximately 35 mL of KCl-Ag2SO4 solution was added. The flask was swirled for a few seconds and allowed to stand until the contents had cooled to room temperature (ca. 5 min). The contents were then made up to volume (50 mL) by adding KCl-Ag2SO4 solution and the flask was closed the flask using a stopper and inverted several times to mix the contents. The intensity was then measured with a Klett-Summerson photoelectric colorimeter.
The soil samples were sent to the University of Missouri’s Soil and Plant Testing Laboratory for physical and chemical measurements. The soil organic matter was determined by dry combustion [41].

2.3. Statistical Analysis

Each set of data was averaged in triplicates. All enzyme activity data were statistically analyzed using analysis of variance (ANOVA) assuming a randomized split complete block design, and Tukey’s multiple comparison tests were used to determine significant differences in means under treatments. Statistical differences were tested at the 5% level (p < 0.05) using Statistical Analysis System (SAS) University Edition (V.9.4) software. Correlation among some selected enzymes was analyzed using RStudio’s corplot function.

3. Results and Discussion

Overall, the application of organic amendments had a positive effect on soil enzymatic activity. The highest enzyme activities were generally found in April, which might be because the April samples had better soil water and temperature conditions to stimulate enzyme activity. The total activity decreased for all enzymes in August. This decrease in enzyme activity could indicate a more stable soil condition in which organic matter does not decompose rapidly. This stability may be beneficial for long-term soil health and carbon sequestration. Furthermore, in a stable soil environment, soil organisms are less stressed, which might reduce the need for high enzyme activity as part of the stress response mechanism. Table 1 shows the chemical properties of compost and biochar, which had the highest organic matter and CEC values.
Table 2 shows the mean soil pH values for all treatments on the three sampling occasions, with no statistically significant differences observed between treatments. These pH values influence soil enzyme activity, resulting in changes in nutrient dynamics.
Pearson’s correlation analysis was performed on the combined dataset of some of the selected enzymes to determine significant correlations. A significant positive correlation between r = 0.957 (URS), r = 0.995 (ACP), and 0.995 (AMD) was observed in relation to BG. Similarly, both URS and ACP had a positive correlation with AMD. However, the ARS was negatively and significantly correlated with all the selected enzymes with r ranging from −0.976 to −1.0 with significance indicated with asterisks (see Figure 1).
β-Glucosidase enzymes are essential components of cellulase enzyme complexes that help in the breakdown of cellulose, a complex carbohydrate found in plant cell walls. These enzymes work together to degrade carbohydrates into their constituent glucose molecules and release bioactive compounds from glycosidic precursors, which can then be used as energy sources [42]. β-Glucosidase activity showed no differentiation in the last two sampling periods (June and August) for the treatments (Figure 2). However, in the first sampling period (April), there was a clear increase in the activity of all treatments relative to the other sampling periods. There was an increase in β-glucosidase activity in August for all treatments, except for the control. The control soil consistently had the lowest activity values in the last month of analysis, which can be attributed to low organic matter content, which is directly related to enzyme activity factors that include both oxidative and hydrolytic responses [43] and the inability of the soil to degrade recalcitrant materials left in the soil [44]. de Mora and colleagues [45] found increased β-glucosidase activity in municipal compost compared to biochar-treated soil. Conversely, this study found no significant difference between the compost-treated, biochar, compost biochar, or biochar compost tea at all sampling times except the control, which decreased in the last month. This indicates similar substrate availability in each of the treatments, regardless of the application rate or amendment source. β-Glucosidase plays a major role in microbial metabolism by releasing low-molecular-weight sugars that serve as energy sources and help in global carbon cycling, as well as being involved in the final step of cellulose degradation and is sensitive to residue management [46,47,48]. Carbon cycle transformation, soil organic matter composition, and cycles were all correlated with β-glucosidase. This enzyme serves as a marker for changes in soil agrotechnical parameters as well as for some changes in soil biological activity [49].
Plants, fungi, and bacteria release acid phosphatase enzymes into the soil matrix and are optimal under slightly acidic (pH 6.5) soil conditions [50]. They contribute significantly to cycling phosphorus by breaking down organic forms of phosphorus, such as nucleotides, phytate, and other phosphate esters found in plant residues and soil organic matter in an ecosystem to release inorganic phosphorus for plant utilization [51]. Acid phosphatase is an indicator of the potential of soil to release PO43− from organic matter. Due to the high phosphorus loading rates in all amendments, there was an immediate difference in the phosphatase activities of the biochar compared to the compost and compost biochar treatments. This may be due to the presence of phosphatases in the compost before incorporation into the soil. Activity levels showed a significant increase in the compost treatment compared with the biochar treatment for all three sampling periods. However, the trends showed significant decreases over time (Figure 3), with acid phosphatase activities being the lowest in August. The treatments showed no clear mean separation from that of the control in the final month. This suggests that the lost substrate stimulated the microbial community to produce phosphatase.
Alkaline phosphatase is generated by a wide range of microbes, plants, and even certain soil flora. Inorganic phosphate, which plants can absorb, is released when the enzyme hydrolyzes organic phosphates. This process promotes nutrient availability and boosts plant growth, both of which are crucial for maximizing soil productivity [52]. Activity levels showed a significant increase in the compost biochar treatment compared with the other treatments. However, the trend showed a significant decrease over time (Figure 4). The treatments showed no clear mean separation from the control at all studied times and had the lowest activity compared to the other treatments. Differences in the application rates, and thus the amount of available N and P, may affect the activity of phosphatase enzymes. The overall decrease in phosphatase activity may reflect the loss of P substrate due to microbial utilization as well as inhibition from excess P inputs due to feedback inhibition.
Arylsulfatase is involved in the sulfur cycle and is an essential indicator of microbial activity and nutrient cycling within the soil ecosystem. It breaks down sulfur-containing compounds to release sulfate ions (SO42⁻), which are essential nutrients for plant growth because sulfur is a critical component of amino acids, vitamins, and enzymes in plants, making it crucial for their development and overall soil fertility [53,54]. The control soil had significantly lower arylsulfatase activity than the rest of the treatments. However, two months after incorporation, there were no differences between the treatments, suggesting a loss of stimulation by compost and biochar (Figure 5). Our results may be due to lower levels of organic matter in the control soil than in the organic-amended soils. Arylsulfatase activity was higher in the compost + biochar and compost biochar + compost tea treatments than in all other treatments from April to August. The arylsulfatase activity significantly decreased by approximately 60% from April to August. Although this difference was significant compared to all other treatments, it was the only enzyme that had the highest values in the compost and compost biochar treatments. The control soil treatment from June to August showed no differences. This may be due to the loss of arylsulfatase substrates, which stimulate the activity of this enzyme over time. Arylsulfatase activity was significantly higher during the first sampling period (April) immediately after amendment incorporation. Tejada and colleagues [55] reported similar results with organic amendments to degraded soils. Similar patterns of arylsulfatase activity were found by Bandick and Dick [46] during the given period following the introduction of manure into the soil. Arylsulfatase activity is susceptible to a range of different management approaches [56,57]. As an extracellular enzyme that catalyzes the hydrolysis of organic sulfate esters and releases plant-available sulfate (SO42−), arylsulfatase is also important in that it may be an indirect indicator of fungi because of the ester sulfate bond that is lacking in bacteria [57].
Dehydrogenase is involved in the biological oxidation of organic matter and can be used to evaluate the overall metabolic activity of soil microorganisms. It contributes to the decomposition of organic matter, which improves the soil structure by gluing soil particles for better structure, water retention, and nutrient availability [58]. Figure 6 shows that the highest dehydrogenase activity (0.78 µg of TPF formed g−1 soil h−1) was observed for compost for the first sampling period (April). The highest dehydrogenase activity was observed in all the soil treatments during the same period. The decrease in dehydrogenase activity between 60 and 120 d indicated the absence of active decomposition and confirmed that the compost was mature. Activity levels showed a significant increase in the compost treatment compared to the biochar treatment for all three sampling periods, which was also the same for the control. This may be due to the presence of dehydrogenases in the plots before their incorporation into the soil. Dehydrogenase enzymes play a significant role in the biological oxidation of soil organic matter by transferring protons and electrons from substrates to acceptors [59]. Dehydrogenase provides correlative information on the biological activity and microbial population in soil and is considered to exist in soils as an integral part of intact cells. The measurement of dehydrogenase activity represents the immediate metabolic activity of the soil [60].
Arylamidase plays an important role in the assessment of soil health. It is involved in the breakdown of organic nitrogenous compounds and serves as an indicator of microbial activity and nutrient cycling in soil ecosystems. It works by breaking down amides and releasing ammonia (NH3) and other forms of organic nitrogen. This process is crucial for nitrogen cycling in the soil as it converts organic nitrogen into ammonium, which can be further transformed into forms that are accessible to plants [61]. The arylamidase enzyme demonstrated the highest activity (around 48 µg of p-naphthylamine g−1 soil h−1) for the control in the last sampling period (Figure 7). The highest arylamidase activity was observed in all soil treatments. The decrease in arylamidase activity between days 0 and 60 indicates the absence of active decomposition. Activity levels showed a significant decrease in all treatments in the second sampling period compared with the other treatments. This may have been due to more rain during that period, which resulted in low arylamidase activity in June. The arylamidase enzyme catalyzes one of the initial reactions in N mineralization because it is involved in the release of amino acids from soil organic matter. The arylamidase is an indicator of the N cycle, the hydrolysis of an N-terminal amino acid from peptides, amides, and arylamides, is an index of N mineralization in soils [47].
Cellulase breaks down cellulose and releases sugars, including glucose, into the soil. These sugars can act as food sources for soil microorganisms, fostering their activity and growth. This microbial activity aids in the mineralization of nutrients, releasing them for use by plants [62]. There was no statistical difference in cellulase activity between the first two sampling periods (Figure 8). However, at the end of the sampling period, there was a clear decrease in the activity of all treatments. The control soil consistently had the lowest activity values, except for the last sampling period in the biochar treatment, which can be attributed to the low organic matter content and the inability of the soil to degrade recalcitrant materials left in the soil. Conversely, this study found no significant difference between compost treated with biochar and biochar treated with compost tea in the last two sampling periods. This indicates similar substrate availability in each of the treatments, regardless of the application rate or amendment source. Cellulase is the most abundant organic compound in the biosphere, comprising almost 50% of the biomass that is synthesized by the photosynthetic fixation of CO2 [63]. The growth and activity of soil microorganisms depend on the carbon source that is contained mainly as plant residues in the soil; however, for carbon to be released as an energy source for use by microorganisms, cellulose in plant debris must be degraded into high molecular weight oligosaccharides, cellobiose, and glucose by cellulase enzymes [64]. Since cellulase enzymes play an important role in the global recycling of the most abundant polymer in nature, more research should be conducted to understand the nature of this enzyme better, so that it can be used more regularly as a predictive tool in the assessment of soil fertility.
Urease enzymes catalyze the hydrolysis of urea, releasing ammonium ions (NH4⁺) and carbon dioxide (CO2) as byproducts via the conversion of organic nitrogen. This process is essential for the mineralization of nitrogen in the form of ammonium, which can be taken up by plants, contributing to their growth and overall nitrogen nutrition in soils. Adequate nitrogen availability is crucial for plant health and productivity [65]. Urease activity remained similar between all soil-amended treatments, with slight differences, except for the control, which was different for the entire experiment (Figure 9). Urease is a hydrolase that catalyzes the hydrolysis of urea to CO2 and NH4+, with the latter available for uptake by microorganisms and plants [66].
In general, within the three-month sampling period, we observed an increase in BG activity in August for all treatments except the control. Both acid and alkaline phosphatase activities decreased in August. Arylsulfatase activity decreased in August, except for in biochar and compost tea. DG activity decreased in August, except in the case of biochar. AMD activity increased in August for all treatments, whereas CLS activity decreased in August. urease activity increased in August in all the treatments. Our results demonstrated that the BG, ALP, and ARS indicated no significant difference among the treatments in the mid-month and the last month of sampling. Future research will focus on a long-term study into the effect of these treatments on soil health.

4. Conclusions

The amount of nitrogen applied significantly affected enzyme activity, indicating its crucial role in biochemical processes. Compost biochar also had significant effects on soil enzyme activity; however, there was no fixed pattern of such effects in this study. Owing to their sensitivity to environmental factors, soil enzymes such as BG, ACP-ALP, ARS, DG, AMD, CLS, and URS are suitable indicators of soil quality. Compost and biochar as soil amendments are sensitive to changes in soil properties; therefore, they are recommended for use as soil amendments. Almost all enzyme activities were highest in biochar, compost biochar, and biochar compost tea, except for BG, ACP, and DG in the compost treatment during the first sampling period. This could be due to the presence of active enzyme components in the initial stage of treatment. Assessing the effects of soil management on soil enzyme activities is important for selecting practices to optimize sustainable plant production. Soil management methods affect the physical, chemical, and biological properties of soil in several different ways, and therefore affect enzyme activities. The results indicated that compost was favorable for increasing soil enzyme activities, and the effects of biochar were significant for some soil enzyme activities. The strong association between BG, ACP, and AMD (99%) can be taken into consideration and reduce the cost of analysis for small farmers. Determining one or two of the enzymes can still help farmers to learn about their soil’s biological activity, and hence help them to manage their soil sustainably.

Author Contributions

Conceptualization, F.Z. and N.A.-A.; validation, F.Z. and N.A.-A.; formal analysis, F.Z., N.A.-A., and B.T.B.A.; data curation, F.Z.; writing—original draft preparation, F.Z. and N.A.-A.; writing—review and editing, analyzing and interpreting the data; F.Z., N.A.-A., J.Y., S.H.A., and B.T.B.A.; visualization, F.Z. and N.A.-A.; supervision, J.Y. and S.H.A. All authors have read and agreed to the published version of the manuscript.

Funding

While this project did not receive external funding, the support and encouragement from various individuals were invaluable.

Data Availability Statement

The data supporting the study’s conclusions are available upon reasonable request from the corresponding author [N.A.-A.].

Acknowledgments

We thank the anonymous reviewers for their valuable comments, which helped us substantially improve the manuscript. We would like to express our sincere appreciation to all those who contributed to completing this work, specifically Raymond Bayan.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Thapa, R.B.; Coupal, R.H.; Dangi, M.B.; Stahl, P.D. An Assessment of Plant Growth and Soil Properties Using Coal Char and Biochar as a Soil Amendment. Agronomy 2024, 14, 320. [Google Scholar] [CrossRef]
  2. Thapa, R.B.; Budhathoki, S.; Shilpakar, C.; Panday, D.; Alsunuse, B.T.B.; Tang, S.; Stahl, P.D. Enhancing Corn Yield and Soil Quality in Irrigated Semiarid Region with Coal Char and Biochar Amendments. Soil Syst. 2024, 8, 82. [Google Scholar] [CrossRef]
  3. Kapoor, A.; Sharma, R.; Kumar, A.; Sepehya, S. Biochar as a means to improve soil fertility and crop productivity: A review. J. Plant Nutr. 2022, 45, 2380–2388. [Google Scholar] [CrossRef]
  4. Fischer, D.; Glaser, B. Synergisms between compost and biochar for sustainable soil amelioration. Manag. Org. Waste 2012, 1, 167–198. [Google Scholar]
  5. Blanco-Canqui, H. Biochar and soil physical properties. Soil Sci. Soc. Am. J. 2017, 81, 687–711. [Google Scholar] [CrossRef]
  6. Ghorbani, M.; Amirahmadi, E.; Bernas, J.; Konvalina, P. Testing biochar’s ability to moderate extremely acidic soils in tea-growing areas. Agronomy 2024, 14, 533. [Google Scholar] [CrossRef]
  7. Domingues, R.R.; Sánchez-Monedero, M.A.; Spokas, K.A.; Melo, L.C.; Trugilho, P.F.; Valenciano, M.N.; Silva, C.A. Enhancing cation exchange capacity of weathered soils using biochar: Feedstock, pyrolysis conditions and addition rate. Agronomy 2020, 10, 824. [Google Scholar] [CrossRef]
  8. Zheng, H.; Wang, Z.; Deng, X.; Herbert, S.; Xing, B. Impacts of adding biochar on nitrogen retention and bioavailability in agricultural soil. Geoderma 2013, 206, 32–39. [Google Scholar] [CrossRef]
  9. Oleszczuk, P.; Jośko, I.; Futa, B.; Pasieczna-Patkowska, S.; Pałys, E.; Kraska, P. Effect of pesticides on microorganisms, enzymatic activity and plant in biochar-amended soil. Geoderma 2014, 214, 10–18. [Google Scholar] [CrossRef]
  10. Bailey, V.L.; Fansler, S.J.; Smith, J.L.; Bolton, H., Jr. Reconciling apparent variability in effects of biochar amendment on soil enzyme activities by assay optimization. Soil Biol. Biochem. 2011, 43, 296–301. [Google Scholar] [CrossRef]
  11. Gundale, M.J.; DeLuca, T.H. Charcoal effects on soil solution chemistry and growth of Koeleria macrantha in the ponderosa pine/Douglas-fir ecosystem. Biol. Fertil. Soils 2007, 43, 303–311. [Google Scholar] [CrossRef]
  12. Lima, I.M.; Marshall, W.E. Adsorption of selected environmentally important metals by poultry manure-based granular activated carbons. J. Chem. Technol. Biotechnol. Int. Res. Process Environ. Clean Technol. 2005, 80, 1054–1061. [Google Scholar] [CrossRef]
  13. Singh, B.; Cowie, A. A novel approach, using 13C natural abundance, for measuring decomposition of biochars in soil. In Carbon and Nutrient Management in Agriculture; Massey University, Fertilizer and Lime Research Centre: Palmerston North, New Zealand, 2008. [Google Scholar]
  14. Sun, Z.; Bruun, E.W.; Arthur, E.; de Jonge, L.W.; Moldrup, P.; Hauggaard-Nielsen, H.; Elsgaard, L. Effect of biochar on aerobic processes, enzyme activity, and crop yields in two sandy loam soils. Biol. Fertil. Soils 2014, 50, 1087–1097. [Google Scholar] [CrossRef]
  15. Keiluweit, M.; Nico, P.S.; Johnson, M.G.; Kleber, M. Dynamic molecular structure of plant biomass-derived black carbon (biochar). Environ. Sci. Technol. 2010, 44, 1247–1253. [Google Scholar] [CrossRef]
  16. Demisie, W.; Liu, Z.; Zhang, M. Effect of biochar on carbon fractions and enzyme activity of red soil. Catena 2014, 121, 214–221. [Google Scholar] [CrossRef]
  17. Steinbeiss, S.; Gleixner, G.; Antonietti, M. Effect of biochar amendment on soil carbon balance and soil microbial activity. Soil Biol. Biochem. 2009, 41, 1301–1310. [Google Scholar] [CrossRef]
  18. Keech, O.; Dizengremel, P.; Gardeström, P. Preparation of leaf mitochondria from Arabidopsis thaliana. Physiol. Plant. 2005, 124, 403–409. [Google Scholar] [CrossRef]
  19. Baldock, J.A.; Smernik, R.J. Chemical composition and bioavailability of thermally altered Pinus resinosa (Red pine) wood. Org. Geochem. 2002, 33, 1093–1109. [Google Scholar] [CrossRef]
  20. Krull, E.S.; Baldock, J.A.; Skjemstad, J.O. Importance of mechanisms and processes of the stabilisation of soil organic matter for modelling carbon turnover. Funct. Plant Biol. 2003, 30, 207–222. [Google Scholar] [CrossRef]
  21. Lehmann, J. Bio-energy in the black. Front. Ecol. Environ. 2007, 5, 381–387. [Google Scholar] [CrossRef]
  22. Van Zwieten, L.; Kimber, S.; Morris, S.; Chan, K.Y.; Downie, A.; Rust, J.; Joseph, S.; Cowie, A. Effects of biochar from slow pyrolysis of papermill waste on agronomic performance and soil fertility. Plant Soil 2010, 327, 235–246. [Google Scholar] [CrossRef]
  23. Rondon, M.A.; Lehmann, J.; Ramírez, J.; Hurtado, M. Biological nitrogen fixation by common beans (Phaseolus vulgaris L.) increases with bio-char additions. Biol. Fertil. Soils 2007, 43, 699–708. [Google Scholar] [CrossRef]
  24. Joseph, S.; Peacocke, C.; Lehmann, J.; Munroe, P. Developing a biochar classification and test methods. In Biochar for Environmental Management; Routledge: London, UK, 2012; pp. 139–158. [Google Scholar]
  25. Laird, D.; Fleming, P.; Wang, B.; Horton, R.; Karlen, D. Biochar impact on nutrient leaching from a Midwestern agricultural soil. Geoderma 2010, 158, 436–442. [Google Scholar] [CrossRef]
  26. Liang, Y.S.; Choi, Y.H.; Kim, H.K.; Linthorst, H.J.; Verpoorte, R. Metabolomic analysis of methyl jasmonate treated Brassica rapa leaves by 2-dimensional NMR spectroscopy. Phytochemistry 2006, 672, 503–511. [Google Scholar] [CrossRef] [PubMed]
  27. Palansooriya, K.N.; Wong, J.T.; Hashimoto, Y.; Huang, L.; Rinklebe, J.; Chang, S.X.; Bolan, N.; Wang, H.; Ok, Y.S. Response of microbial communities to biochar-amended soils: A critical review. Biochar 2019, 1, 3–22. [Google Scholar] [CrossRef]
  28. Tabatabai, M.A. Soil enzymes. In Methods of Soil Analysis: Part 2 Microbiological and Biochemical Properties; ACSESS: Sydney, Australia, 1994; Volume 5, pp. 775–833. [Google Scholar]
  29. Alsunuse, B.T.B. Effects of Cover Crops and Crop Rotation on Soil Health in Northwestern Wyoming. Ph.D. Thesis, University of Wyoming, Laramie, WY, USA, December 2023. [Google Scholar]
  30. Longo, M.A.; Combes, D. Analysis of the thermal deactivation kinetics of α-chymotrypsin modified by chemoenzymatic glycosylation. In Progress in Biotechnology; Elsevier: Amsterdam, The Netherlands, 1998; Volume 15, pp. 135–140. [Google Scholar]
  31. Sohi, S.; Lopez-Capel, E.; Krull, E.; Bol, R. Biochar, climate change and soil: A review to guide future research. CSIRO Land Water Sci. Rep. 2009, 5, 17–31. [Google Scholar]
  32. Rahman, G.M.; Rahman, M.M.; Alam, M.S.; Kamal, M.Z.; Mashuk, H.A.; Datta, R.; Meena, R.S. Biochar and organic amendments for sustainable soil carbon and soil health. In Carbon and Nitrogen Cycling in Soil; Springer: Singapore, 2020; pp. 45–85. [Google Scholar]
  33. Available online: https://casoilresource.lawr.ucdavis.edu/gmap/ (accessed on 12 January 2023).
  34. Ansari, J.; Eivazi, F.; Anderson, S.H.; Bardhan, S. Selected Enzyme Activities Under Different Land Use Management in Lower Missouri River Floodplain Soils. Commun. Soil Sci. Plant Anal. 2024, 55, 27–39. [Google Scholar] [CrossRef]
  35. Eivazi, F.; Tabatabai, M.A. Glucosidases and galactosidases in soils. Soil Biol. Biochem. 1988, 20, 601–606. [Google Scholar] [CrossRef]
  36. Eivazi, F.; Tabatabai, M.A. Phosphatases in soils. Soil Biol. Biochem. 1977, 9, 167–172. [Google Scholar] [CrossRef]
  37. Tabatabai, M.A.; Bremner, J.M. Arylsulfatase activity of soils. Soil Sci. Soc. Am. J. 1970, 34, 225–229. [Google Scholar] [CrossRef]
  38. Casida, L.E., Jr.; Klein, D.A.; Santoro, T. Soil dehydrogenase activity. Soil Sci. 1964, 98, 371–376. [Google Scholar] [CrossRef]
  39. Acosta-Martinez, V.; Tabatabai, M.A. Enzyme activities in a limed agricultural soil. Biol. Fertil. Soils 2000, 31, 85–91. [Google Scholar] [CrossRef]
  40. Deng, S.P.; Tabatabai, M.A. Cellulase activity of soils. Soil Biol. Biochem. 1994, 26, 1347–1354. [Google Scholar] [CrossRef]
  41. Nelson, D.W.; Sommers, L.E. Total carbon, organic carbon, and organic matter. In Methods of Soil Analysis: Part 3 Chemical Methods; John Wiley & Sons: Hoboken, NJ, USA, 1996; Volume 5, pp. 961–1010. [Google Scholar]
  42. Ejaz, U.; Sohail, M.; Ghanemi, A. Cellulases: From bioactivity to a variety of industrial applications. Biomimetics 2021, 6, 44. [Google Scholar] [CrossRef] [PubMed]
  43. Sinsabaugh, R.L.; Gallo, M.E.; Lauber, C.; Waldrop, M.P.; Zak, D.R. Extracellular enzyme activities and soil organic matter dynamics for northern hardwood forests receiving simulated nitrogen deposition. Biogeochemistry 2005, 75, 201–215. [Google Scholar] [CrossRef]
  44. Garcia, C.; Hernandez, T.; Barahona, A.; Costa, F. Organic matter characteristics and nutrient content in eroded soils. Environ. Manag. 1996, 20, 133–141. [Google Scholar] [CrossRef]
  45. de Mora, A.P.; Ortega-Calvo, J.J.; Cabrera, F.; Madejón, E. Changes in enzyme activities and microbial biomass after “in situ” remediation of a heavy metal-contaminated soil. Appl. Soil Ecol. 2005, 28, 125–137. [Google Scholar] [CrossRef]
  46. Bandick, A.K.; Dick, R.P. Field management effects on soil enzyme activities. Soil Biol. Biochem. 1999, 31, 1471–1479. [Google Scholar] [CrossRef]
  47. Acosta-Martínez, V.; Tabatabai, M.A. Arylamidase activity in soils: Effect of trace elements and relationships to soil properties and activities of amidohydrolases. Soil Biol. Biochem. 2001, 33, 17–23. [Google Scholar] [CrossRef]
  48. Zang, X.; Liu, M.; Fan, Y.; Xu, J.; Xu, X.; Li, H. The structural and functional contributions of β-glucosidase-producing microbial communities to cellulose degradation in composting. Biotechnol. Biofuels 2018, 11, 51. [Google Scholar] [CrossRef]
  49. Rachwał, K.; Gustaw, K.; Kazimierczak, W.; Waśko, A. Is soil management system really important? comparison of microbial community diversity and structure in soils managed under organic and conventional regimes with some view on soil properties. PLoS ONE 2021, 16, e0256969. [Google Scholar] [CrossRef]
  50. Margalef, O.; Sardans, J.; Fernández-Martínez, M.; Molowny-Horas, R.; Janssens, I.A.; Ciais, P.; Goll, D.; Richter, A.; Obersteiner, M.; Asensio, D.; et al. Global patterns of phosphatase activity in natural soils. Sci. Rep. 2017, 7, 1337. [Google Scholar] [CrossRef] [PubMed]
  51. Park, Y.; Solhtalab, M.; Thongsomboon, W.; Aristilde, L. Strategies of organic phosphorus recycling by soil bacteria: Acquisition, metabolism, and regulation. Environ. Microbiol. Rep. 2022, 14, 3–24. [Google Scholar] [CrossRef] [PubMed]
  52. Timofeeva, A.; Galyamova, M.; Sedykh, S. Prospects for using phosphate-solubilizing microorganisms as natural fertilizers in agriculture. Plants 2022, 11, 2119. [Google Scholar] [CrossRef] [PubMed]
  53. Basta, N.T.; Busalacchi, D.M.; Hundal, L.S.; Kumar, K.; Dick, R.P.; Lanno, R.P.; Carlson, J.; Cox, A.E.; Granato, T.C. Restoring ecosystem function in degraded urban soil using biosolids, biosolids blend, and compost. J. Environ. Qual. 2016, 45, 74–83. [Google Scholar] [CrossRef]
  54. Romillac, N.; Slezack-Deschaumes, S.; Amiaud, B.; Piutti, S. Soil Microbial Communities Involved in Proteolysis and Sulfate-Ester Hydrolysis Are More Influenced by Interannual Variability than by Crop Sequence. Agronomy 2023, 13, 180. [Google Scholar] [CrossRef]
  55. Tejada, M.; Hernandez, M.T.; Garcia, C. Application of two organic amendments on soil restoration: Effects on the soil biological properties. J. Environ. Qual. 2006, 35, 1010–1017. [Google Scholar] [CrossRef] [PubMed]
  56. Ndiaye, E.L.; Sandeno, J.M.; McGrath, D.; Dick, R.P. Integrative biological indicators for detecting change in soil quality. Am. J. Altern. Agric. 2000, 15, 26–36. [Google Scholar] [CrossRef]
  57. Knight, T.R.; Dick, R.P. Differentiating microbial and stabilized β-glucosidase activity relative to soil quality. Soil Biol. Biochem. 2004, 36, 2089–2096. [Google Scholar] [CrossRef]
  58. Wolińska, A.; Stępniewska, Z. Dehydrogenase activity in the soil environment. Dehydrogenases 2012, 10, 183–210. [Google Scholar]
  59. Sánchez-Monedero, M.A.; Mondini, C.; Cayuela, M.L.; Roig, A.; Contin, M.; De Nobili, M. Fluorescein diacetate hydrolysis, respiration and microbial biomass in freshly amended soils. Biol. Fertil. Soils 2008, 44, 885–890. [Google Scholar] [CrossRef]
  60. Gu, Y.; Wang, P.; Kong, C.H. Urease, invertase, dehydrogenase and polyphenoloxidase activities in paddy soil influenced by allelopathic rice variety. Eur. J. Soil Biol. 2009, 45, 436–441. [Google Scholar] [CrossRef]
  61. Ekenler, M.; Tabatabai, M.A. Arylamidase and amidohydrolases in soils as affected by liming and tillage systems. Soil Tillage Res. 2004, 77, 157–168. [Google Scholar] [CrossRef]
  62. Jayasekara, S.; Ratnayake, R. Microbial cellulases: An overview and applications. Cellulose 2019, 22, 10–5772. [Google Scholar]
  63. Geisseler, D.; Horwath, W.R.; Joergensen, R.G.; Ludwig, B. Pathways of nitrogen utilization by soil microorganisms—a review. Soil Biol. Biochem. 2010, 42, 2058–2067. [Google Scholar] [CrossRef]
  64. Eriksson, K.E.; Blanchette, R.A.; Ander, P. Microbial and Enzymatic Degradation of Wood and Wood Components; Springer: Berlin/Heidelberg, Germany, 2012. [Google Scholar]
  65. Cordero, I.; Snell, H.; Bardgett, R.D. High throughput method for measuring urease activity in soil. Soil Biol. Biochem. 2019, 134, 72–77. [Google Scholar] [CrossRef]
  66. Vitolo, M. Notes on urea hydrolysis by urease. J. Pharm. Pharm. Sci. 2022, 11, 96–135. [Google Scholar]
Processes 12 01678 g001
Figure 1. Pearson’s correlation chart between selected soil enzymes: β-glucosidase (BG), urease (URS), acid phosphatase (ACP), arylamidase (AMD), and arylsulfatase (ARS). *** p < 0.0001, ** p < 0.01, * p < 0.05.
Figure 1. Pearson’s correlation chart between selected soil enzymes: β-glucosidase (BG), urease (URS), acid phosphatase (ACP), arylamidase (AMD), and arylsulfatase (ARS). *** p < 0.0001, ** p < 0.01, * p < 0.05.
Processes 12 01678 g001
Processes 12 01678 g002
Figure 2. Effect of different treatments on β-glucosidase (BG) activities. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 2. Effect of different treatments on β-glucosidase (BG) activities. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g002
Processes 12 01678 g003
Figure 3. Effect of different treatments on acid phosphatase (ACP) activities. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 3. Effect of different treatments on acid phosphatase (ACP) activities. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g003
Processes 12 01678 g004
Figure 4. Effect of different treatments on alkaline phosphatase (ALP) activities. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 4. Effect of different treatments on alkaline phosphatase (ALP) activities. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g004
Processes 12 01678 g005
Figure 5. Effect of different treatments on aylsulfatase (ARS) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 5. Effect of different treatments on aylsulfatase (ARS) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g005
Processes 12 01678 g006
Figure 6. Effect of different treatments on dehydrogenase (DG) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 6. Effect of different treatments on dehydrogenase (DG) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g006
Processes 12 01678 g007
Figure 7. Effect of different treatments on arylamidase (AMD) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 7. Effect of different treatments on arylamidase (AMD) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g007
Processes 12 01678 g008
Figure 8. Effect of different treatments on cellulase (CLS) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 8. Effect of different treatments on cellulase (CLS) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g008
Processes 12 01678 g009
Figure 9. Effect of different treatments on urease (URS) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Figure 9. Effect of different treatments on urease (URS) activity. Treatments include the control for April, June, and August. The different lower-case letters show significant differences (p < 0.05) between the treatments.
Processes 12 01678 g009
Table 1. Selected soil chemical properties for the study.
Table 1. Selected soil chemical properties for the study.
TreatmentspHsEC
dS m−1		%OMP Bray I
mgkg−1		Ca
mgkg−1		Mg mgkg−1K
mgkg−1		CEC cmolc kg−1
Compost6.910.3480.62872.0764.6527.312.2
Biochar6.89.234.288.82914.8814.2593.411.8
Compost + Biochar6.912.64.85197.54998.41120.81120.812.4
Biochar + Compost Tea6.913.44.385.14013.0695.6695.611.7
Control6.651.72.7527.75712.2588.5588.511.3
Fescue7.13.33.152.73830.2263.1263.111.3
Table 2. Mean soil pH values of the study treatments for three sampling periods.
Table 2. Mean soil pH values of the study treatments for three sampling periods.
TreatmentsAprilJuneAugust
Compost6.186.246.61
Biochar6.646.616.71
Compost + Biochar6.756.786.77
Biochar + Compost Tea6.516.696.76
Control6.366.566.74
Fescue6.556.666.78
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Zaid, F.; Al-Awwal, N.; Yang, J.; Anderson, S.H.; Alsunuse, B.T.B. Effects of Biochar-Amended Composts on Selected Enzyme Activities in Soils. Processes 2024, 12, 1678. https://doi.org/10.3390/pr12081678

AMA Style

Zaid F, Al-Awwal N, Yang J, Anderson SH, Alsunuse BTB. Effects of Biochar-Amended Composts on Selected Enzyme Activities in Soils. Processes. 2024; 12(8):1678. https://doi.org/10.3390/pr12081678

Chicago/Turabian Style

Zaid, Faraj, Nasruddeen Al-Awwal, John Yang, Stephen H. Anderson, and Bouzeriba T. B. Alsunuse. 2024. "Effects of Biochar-Amended Composts on Selected Enzyme Activities in Soils" Processes 12, no. 8: 1678. https://doi.org/10.3390/pr12081678

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

Article Metrics

Back to TopTop