Molecular Analysis of Methanogen Richness in Landfill and Marshland Targeting 16S rDNA Sequences

Author

Shailendra Yadav, Sharbadeb Kundu, Sankar K. Ghosh, S. S. Maitra 

Published: 1 January 2015

 by Hindawi Publishing Corporation

 in Archaea

Archaea, Volume 2015, pp 1-9; doi:10.1155/2015/563414 

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Abstract: Methanogens, a key contributor in global carbon cycling, methane emission, and alternative energy production, generate methane gas via anaerobic digestion of organic matter. The methane emission potential depends upon methanogenic diversity and activity. Since they are anaerobes and difficult to isolate and culture, their diversity present in the landfill sites of Delhi and marshlands of Southern Assam, India, was analyzed using molecular techniques like 16S rDNA sequencing, DGGE, and qPCR. The sequencing results indicated the presence of methanogens belonging to the seventh order and also the order Methanomicrobiales in the Ghazipur and Bhalsawa landfill sites of Delhi. Sequences, related to the phyla Crenarchaeota (thermophilic) and Thaumarchaeota (mesophilic), were detected from marshland sites of Southern Assam, India. Jaccard analysis of DGGE gel using Gel2K showed three main clusters depending on the number and similarity of band patterns. The copy number analysis of hydrogenotrophic methanogens using qPCR indicates higher abundance in landfill sites of Delhi as compared to the marshlands of Southern Assam. The knowledge about “methanogenic archaea composition” and “abundance” in the contrasting ecosystems like “landfill” and “marshland” may reorient our understanding of the Archaea inhabitants. This study could shed light on the relationship between methane-dynamics and the global warming process.1. IntroductionMethane is an important greenhouse gas because it is 25 times more powerful than CO2 in global warming potential (i.e., the ability of the gas to trap heat in the atmosphere) and thus plays a crucial role in climate change and carbon cycling [1, 2]. Methane emission has contributed approximately 20% to global climate change from preindustrial times [1, 3]. About 500–600 Tg of methane is emitted annually to the atmosphere of which 74% is biogenic, produced by methanogenic Archaea [4].The methanogenic Archaea (methanogens) usually occurs in highly reduced, anoxic environments such as landfills, wetlands, rice fields, rumen, and marine sediments where they serve as a terminal electron sink [5, 6]. Methanogens are strict anaerobes and the presence of oxygen leads to the formation of reactive oxygen species (ROS), which damage their cell membranes, DNA, and proteins [7, 8]. Methanogens are phylogenetically divided into 5 families within the phylum Euryarchaeota and are comprised of 31 known genera [9, 10]. Methanogens can utilize a wide range of compounds for methane production, but, in most natural systems, there are two major pathways for methanogenesis, reduction of CO2 (hydrogenotrophic methanogenesis) and cleavage of acetates (acetoclastic methanogenesis). A third pathway for methane generation is called methylotrophic methanogenesis that occurs in marine sediments and salt lakes where methane is produced from methylated compounds such as trimethylamine [11, 12].Landfill sites are the third largest source of methane. It constitutes about 30 and 24% of the anthropogenic methane production in Europe and US, respectively [4, 13]. In comparison to the western countries, the composition of municipal solid waste (MSW) in developing countries like India is higher (40–60%) in organic waste. This has more potential to emit higher GHGs (Green House Gases) per ton of MSW compared to the developed world [14]. Moreover, landfills in India are neither well planned nor engineered and are often found in low-lying open areas, where municipal waste is haphazardly and indiscriminately disposed. These sites have neither landfill lining to avoid percolation of leachate to groundwater table nor leachate collection facility. The city generates about 6000 tonnes of solid waste per day and the expected quantity of solid waste generation in Delhi would be about 12,750 tonnes per day by 2015 [15]. Due to scarcity of land in big cities, municipal authorities are using the same landfill for nearly 10–20 years. Thus, the possibility of anaerobic emission of GHGs further increases [16].Microbial decomposition, climatic conditions, MSW wastes characteristics, and landfilling operations are among the many factors that contribute to the generation of methane [2, 17]. The migration of gas and leachate away from the landfill boundaries and their release into the surrounding environment present serious environmental threats, including potential health hazards, fires and explosions, damage to vegetation, unpleasant odors, landfill settlement, ground water pollution, air pollution, and global warming [18–20].Wetlands (marshland) are the largest source of natural methane emissions contributing about 10–231 Tg methane per year accounting for 20–39% of annual global CH4 emission [4, 21]. Methanogens in the moist, anoxic (oxygen-free) wetland soil produce CH4 as they decompose dead plant material. The methane emission from wetland was increased by 7% from 2003 to 2007 [2, 19]. Methane production in wetlands is affected by the acetate supply through acetate fermentation or the CO2 reduction potential [22, 23]. The exponential increase in the rate of CH4 production with temperature is due to the availability of more substrates and is not associated with changes in the composition of methanogens [24]. Methanogens belonging to the groups Methanomicrobiales and Methanosarcinales performing acetoclastic and methylotrophic pathway were found to be dominant in landfill sites [25–27]. In acidic conditions, due to the presence of acid tolerant hydrogenotrophic methanogens, H2/CO2 is efficiently converted to methane compared to acetate, and methanogenic activity decreases with decrease in pH regardless of the substrates [28].The prokaryotic diversity in our planet dictates our planet’s ecosystems by acting as key functional drivers [29]. The understanding of the functional potential of the most individual microbial flora residing within the ecosystem is extremely limited because of our inability to isolate and culture them in laboratory conditions [30]. Since the methanogens are anaerobes and are difficult to culture, they are identified by culture independent molecular techniques like PCR amplification, denaturing gradient gel electrophoresis (DGGE), and quantitative real-time PCR, using molecular markers such as 16S rDNA genetic locus [31–34]. Hence, the present study was aimed at detecting the methanogenic Archaea inhabitants (richness) (by DGGE), identification by DNA sequencing, and quantification by qPCR in both the landfill sites of Delhi and marshland sites of Southern Assam, India.2. Material and Methods2.1. Collection of Leachate and Sediment SamplesLeachate samples were collected from three landfill sites (Bhalswa, Okhla, and Ghazipur) in the area of New Delhi, India. These sites are active landfill sites and are still in use. They do not have the leachate collection facility or landfill liner to avoid percolation of leachate to the ground water table (aquifer). Soil, sediment sample was collected from marshlands (Silcoorie Lake (Silchar), Badarpur, and Karimganj) of Southern Assam, India, in sterile falcon tubes. The details of sites along with criteria and physiochemical parameters are shown in Tables 1 and 2.Table 1: Sampling point from Delhi landfill site (Ghazipur, Bhalswa, and Okhla) and Southern Assam marshland (Silcoorie Lake (Silchar), Badarpur, and Karimganj) areas.Table 2: Chemical analysis of leachate samples obtained from three landfill and marshland sites. All parameters are in mg L−1 adapted from Ghosh et al. 2015 and Roy and Gupta 2012 [37, 38].2.2. Nucleic Acid Extraction, PCR Amplification, and CloningDNA from both landfill leachate and marshland sediment samples was extracted on the same day of sampling using Fast DNA Spin Kit for Soil (MP Biomedicals, CA, USA). DNA from the marshlands and landfill leachate was amplified using the primer set 86FWD and 1340REV (Table 3).Table 3: List of primers for PCR amplification of 16S rDNA gene and DGGE used in the present study.The amplification profile was 94°C for 5 min, 94°C for 30 s for 30 cycles, and 58°C for 1 minute, elongation at 72°C for 2 minutes, and final extension at 72°C for 10 minutes followed by a cooling step down to 4°C [35, 36]. Obtained 16S rDNA PCR products were purified by PCR purification kit (Fermentas, UK) as recommended by manufacturer protocol. PCR amplicons of 16S rDNA gene were cloned inside PTZ57R/T vector using the Insta-T/A cloning kit (Fermentas, UK) and transformed into Escherichia coli DH5α. The positive clones were selected using blue-white screening on Luria-Bertani plates containing Ampicillin (100 mg/mL), X-gal (20 mg/mL), and IPTG (100 mM). Then, positive clones were sequenced using M13 FWD primer.2.3. DNA Sequencing and Phylogenetic Analysis of 16S rDNA ClonesSequencing was performed for all the clones with the ABI prism 3130 Genetic Analyzer (Applied Biosystem Inc., CA) at Department of Biochemistry, South Campus, Delhi University. The sequences were edited to exclude the PCR primer-binding site and manually corrected with Sequence Scanner 1.0 (Applied Biosystems) and were checked further for vector contamination using the Vecscreen tool (http://www.ncbi.nlm.nih.gov/tools/vecscreen/). The sequences showing similarity with vector sequences from both ends were trimmed. Sequences were then compared with the available nucleotide database from the NCBI GenBank using the BLAST program [39]. The partial nucleotide sequences of 16S rDNA genes were submitted to NCBI under accession numbers KM041239 to KM041252 (Table 4).Table 4: List of accession numbers of the sequences submitted in NCBI and their percent similarity with database along with the sampling sites.Partial 16S rDNA sequences obtained from this study were used for similarity search in NCBI database using BLAST program. After performing BLAST, sequences showing similarity above 90% were used and aligned in MEGA software version 6.0 [40] using ClustalW. The phylogenetic relatedness among c

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Archaeal Community Changes Associated with Cultivation of Amazon Forest Soil with Oil Palm

Author

Tupinambá, Cantã, Maurí o, cio Egí, dio, Ohana Yonara Assis Costa, Jessica Carvalho Bergmann, Ricardo Henrique Kruger, Cynthia Maria Kyaw, Cristine Chaves Barreto, Betania Ferraz Quirino 

Published: 1 January 2016

 by Hindawi Publishing Corporation

 in Archaea

Archaea, Volume 2016, pp 1-14; doi:10.1155/2016/3762159 

Abstract: This study compared soil archaeal communities of the Amazon forest with that of an adjacent area under oil palm cultivation by 16S ribosomal RNA gene pyrosequencing. Species richness and diversity were greater in native forest soil than in the oil palm-cultivated area, and 130 OTUs (13.7%) were shared between these areas. Among the classified sequences, Thaumarchaeota were predominant in the native forest, whereas Euryarchaeota were predominant in the oil palm-cultivated area. Archaeal species diversity was 1.7 times higher in the native forest soil, according to the Simpson diversity index, and the Chao1 index showed that richness was five times higher in the native forest soil. A phylogenetic tree of unclassified Thaumarchaeota sequences showed that most of the OTUs belong to Miscellaneous Crenarchaeotic Group. Several archaeal genera involved in nutrient cycling (e.g., methanogens and ammonia oxidizers) were identified in both areas, but significant differences were found in the relative abundances of Candidatus Nitrososphaera and unclassified Soil Crenarchaeotic Group (prevalent in the native forest) and Candidatus Nitrosotalea and unclassified Terrestrial Group (prevalent in the oil palm-cultivated area). More studies are needed to culture some of these Archaea in the laboratory so that their metabolism and physiology can be studied.1. IntroductionThe Amazon forest area represents 50% of the world’s remaining rainforests [1]. This biome spreads across Brazil, Bolivia, Peru, Ecuador, Colombia, Venezuela, Republic of Guyana, and French Guyana. The Amazon is the largest Brazilian biome and occupies an area of 4,196,943 km2, corresponding to 67% of the Brazilian territory [2]. The Amazon forest provides important ecosystem services such as hydrological cycles and carbon sequestration and storage. More importantly, it hosts over 20% of all plant and animal species in the world [3], indicating its high species diversity.Amazon’s biodiversity encompasses not only macroflora and macrofauna but also its microorganisms, which are often neglected. Mineral materials and organic compounds present in soil create distinct microhabitats populated by different microbial communities. Microorganisms are crucial to the balance of ecosystems, with soil microbial communities playing important roles in soil fertility, plant health, and essential biogeochemical processes such as nitrification, ammonia oxidation, and methanogenesis [4–7].As of 2010, oil palm was cultivated on 112,500 hectares of land in Brazil [8], primarily in the Amazon region. Oil palm (Elaeis guineensis Jacq.) is a highly productive perennial crop, yielding 2,000–8,000 kg oil/ha [8]. The oil, which is extracted from the fruit, has diverse applications in the food and cosmetic industries and can also be used for biodiesel production [9].Although the Amazon is one of the most species-rich biomes on Earth, little is known about its archaeal diversity. To date only one published microbial ecology study has focused on Amazon soil archaeal diversity using 16S rRNA gene sequencing [10], and only one soil type (i.e., Amazonian dark earth, also called Terra Preta) was studied.The archaeal taxonomy is a matter of constant change since its proposal, in 1977. Initially two phyla were recognized: Crenarchaeota and Euryarchaeota, but in the subsequent years, many new phyla were proposed. One example is the Thaumarchaeota phylum [11], composed predominantly of mesophilic members; it encompasses the ammonia-oxidizing archaea. The phylum Korarchaeota was proposed in 1996, after the identification of DNA sequences from the Obsidian Pool, in Yellowstone National Park [12], composed of one candidate thermophilic species, whose genome was completely sequenced [13]. Recent works have described putative new phyla, such as Nanoarchaeota, Aigarchaeota, Aenigmarchaeota, Parvarchaeota, and Lokiarchaeota, but these phyla are not widely accepted yet, due to the low number of specimens or DNA sequences available. In addition, further analyses have positioned sequences belonging to these phyla in already described phyla, such as Euryarchaeota or Thaumarchaeota [14–17]. There are some archaeal groups, such as MCG (Miscellaneous Crenarchaeotic Group), which are poorly characterized in terms of phylogenetic affiliation; recent data revealed that this group is probably more closely related to the phylum Thaumarchaeota than to the Crenarchaeota.Mesophilic archaea seem to play important roles in the cycling of important nutrients such as nitrogen and carbon. The importance of ammonia-oxidizing archaea (AOAs) has been well documented in different ecosystems, such as soils, marine, and freshwater environments, where they sometimes can be found in higher abundance than the ammonia-oxidizing bacteria (AOBs) [18–20]. On the other hand, their real role in nitrification is not yet well understood due to the scarce number of cultured AOAs and the few physiological studies available for this group. The methanogens are among the nonextremophilic archaea, which are widely distributed in anaerobic environments, such as flooded soils, or marine soils and vents. Methanogens are also found in the gut of termites, in rumen of cows, or even in the mouth and intestine of humans. These organisms play important roles in the carbon cycle, transforming small compounds such as acetate and propionate into methane, and removing the hydrogen, which is potentially hazardous to some bacterial cells. On the other hand, methane production is one of the major gases involved in the planet’s greenhouse effect (reviewed by [21]).There are several studies associating land use with changes in the structure and abundance of soil microbial communities, such as the influence of the land use over the diversity of AOA and AOB in grassland soils [22] and the impacts of edaphic factors on those archaea in tropical soils [23]. Therefore, the conversion of native forest into palm tree culture can be another example of a potential impact of the oil palm cultivation in the archaeal communities of Amazonian soils.This work aimed to improve our understanding of how the soil archaeal community is impacted by oil palm cultivation. To this end, microbial DNA was extracted from soil samples from native forest and an adjacent area under oil palm cultivation. The archaeal 16S ribosomal RNA (rRNA) gene was amplified and sequenced using high-throughput methods for comparative analysis. Here we show for the first time that soil archaeal diversity is reduced in soil under oil palm cultivation compared to native forest soil.2. Materials and Methods2.1. Site Description, Sampling, and ProcessingSoil samples were collected in the State of Pará, Brazil, in an oil palm-cultivated area and an adjacent area of Amazon native forest near the city of Moju (Figure 1). The tropical forest in this region is dense, with trees that are 25–35 m tall [24]. The climate is equatorial, hot, and humid (Ami type according to the Köppen climate classification). Annual temperatures range from 25°C to 27°C, and rainfall is 2,000–3,000 mm per year being irregularly distributed [25]. The soil is predominantly “Latossolo Amarelo” (a type of Oxisol) [26].Figure 1: (a) Native forest and (b) oil palm-cultivated sites.The oil palm cultivation in the sampled farm is not as controlled as other crops (Figure 1(b)). There is no irrigation regime; natural precipitation of the rainforest is the only way these plants are irrigated. In the Amazon, the soil is very moist, due to the high precipitation levels during the year. In the studied area, the annual period of flooding is from February to April. Furthermore, the soil around the palm trees is not fertilized in a homogeneous fashion, since only one side of the plants is directly fertilized.In October 2010, after plant litter was removed, a soil borer was used to obtain four 10 cm deep soil samples from three points in the oil palm-cultivated area (S02°00′28.9′′/W048°37′57.4′′, S02°00′29.2′′/W048°37′56.6′′, and S02°00′31.3′′/W048°37′54.3′′) (Figure 1(b)) and four samples from the native Amazon forest area (S02°00′27.2′′/W048°35′53.0′′) (Figure 1(a)). The samples collected in each area were mixed, ground, and sieved to remove larger particles, yielding one composite sample for each area, with approximately 1 kg each. The samples were stored in plastic bags on dry ice during transportation. A subsample was sent to physicochemical analysis at SoloQuímica Análises de Solo Ltda. (Brasília, DF, Brazil). The rest of the samples were then stored at −80°C until DNA extraction. Initially, the physicochemical characteristics of the soils samples were evaluated individually and a high variation among replicas was observed. This result was due to the heterogeneous fertilization of the palm trees in the cultivation fields in Amazon; therefore, composite samples were necessary to describe the microbiota in oil palm soil.2.2. DNA Extraction, PCR, and Pyrosequencing AnalysisTotal DNA was extracted according to the protocol of Smalla et al. [27], using 2 g soil per sample. To minimize DNA extraction bias, this procedure was performed in quadruplicate. PCR reactions were performed using the following primers specific for Archaea: 340F (5′-CCC TAY GGG GYG CAS CAG-3′) and 1000R (5′-GGC CAT GCA CYW CYT CTC-3′) [28]. Archaea 16S rRNA genes were amplified, yielding 660 bp amplicons. Adapters used as priming sites for both amplification and sequencing (454 Life Sciences, Branford, CT, USA) were ligated to the 5′ end of the primer sequences. Each 20 μL PCR reaction contained 10–30 ng total DNA, 1x reaction buffer, 4 μM dNTP, 10 μM of each primer, 200 μg/mL bovine serum albumin, 0.5 U KAPA2G Robust HotStart polymerase, and Milli-Q water. Amplification was performed in an Applied Biosystems GeneAmp® PCR System 9700 Thermal Cycler (Applied Biosystems, Foster City, CA, USA) using the following program: 2 min at 98°C; followed by 30 cycles of 30 seconds at 95°C, 30 seco

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Ecology of Nitrogen Fixing, Nitrifying, and Denitrifying Microorganisms in Tropical Forest Soils

Author

Silvia Pajares, Brendan J. M. Bohannan 

Published: 5 July 2016

 by Frontiers Media SA

 in Frontiers in Microbiology

Frontiers in Microbiology, Volume 7; doi:10.3389/fmicb.2016.01045 

Abstract: Soil microorganisms play important roles in nitrogen cycling within forest ecosystems. Current research has revealed that a wider variety of microorganisms, with unexpected diversity in their functions and phylogenies, are involved in the nitrogen cycle than previously thought, including nitrogen-fixing bacteria, ammonia-oxidizing bacteria and archaea, heterotrophic nitrifying microorganisms, and anammox bacteria, as well as denitrifying bacteria, archaea, and fungi. However, the vast majority of this research has been focused in temperate regions, and relatively little is known regarding the ecology of nitrogen-cycling microorganisms within tropical and subtropical ecosystems. Tropical forests are characterized by relatively high precipitation, low annual temperature fluctuation, high heterogeneity in plant diversity, large amounts of plant litter, and unique soil chemistry. For these reasons, regulation of the nitrogen cycle in tropical forests may be very different from that of temperate ecosystems. This is of great importance because of growing concerns regarding the effect of land use change and chronic-elevated nitrogen deposition on nitrogen-cycling processes in tropical forests. In the context of global change, it is crucial to understand how environmental factors and land use changes in tropical ecosystems influence the composition, abundance and activity of key players in the nitrogen cycle. In this review, we synthesize the limited currently available information regarding the microbial communities involved in nitrogen fixation, nitrification and denitrification, to provide deeper insight into the mechanisms regulating nitrogen cycling in tropical forest ecosystems. We also highlight the large gaps in our understanding of microbially mediated nitrogen processes in tropical forest soils and identify important areas for future research.

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A Review on Heavy Metals (As, Pb, and Hg) Uptake by Plants through Phytoremediation

Author

Bieby Voijant Tangahu, Siti Rozaimah Sheikh Abdullah, Hassan Basri, Mushrifah Idris, Nurina Anuar, Muhammad Mukhlisin 

Published: 1 January 2011

 by Hindawi Publishing Corporation

 in International Journal of Chemical Engineering

International Journal of Chemical Engineering, Volume 2011, pp 1-31; doi:10.1155/2011/939161 

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Abstract: Heavy metals are among the most important sorts of contaminant in the environment. Several methods already used to clean up the environment from these kinds of contaminants, but most of them are costly and difficult to get optimum results. Currently, phytoremediation is an effective and affordable technological solution used to extract or remove inactive metals and metal pollutants from contaminated soil and water. This technology is environmental friendly and potentially cost effective. This paper aims to compile some information about heavy metals of arsenic, lead, and mercury (As, Pb, and Hg) sources, effects and their treatment. It also reviews deeply about phytoremediation technology, including the heavy metal uptake mechanisms and several research studies associated about the topics. Additionally, it describes several sources and the effects of As, Pb, and Hg on the environment, the advantages of this kind of technology for reducing them, and also heavy metal uptake mechanisms in phytoremediation technology as well as the factors affecting the uptake mechanisms. Some recommended plants which are commonly used in phytoremediation and their capability to reduce the contaminant are also reported.1. IntroductionHeavy metals are among the contaminants in the environment. Beside the natural activities, almost all human activities also have potential contribution to produce heavy metals as side effects. Migration of these contaminants into noncontaminated areas as dust or leachates through the soil and spreading of heavy metals containing sewage sludge are a few examples of events contributing towards contamination of the ecosystems [1]. Several methods are already being used to clean up the environment from these kinds of contaminants, but most of them are costly and far away from their optimum performance. The chemical technologies generate large volumetric sludge and increase the costs [2]; chemical and thermal methods are both technically difficult and expensive that all of these methods can also degrade the valuable component of soils [3]. Conventionally, remediation of heavy-metal-contaminated soils involves either onsite management or excavation and subsequent disposal to a landfill site. This method of disposal solely shifts the contamination problem elsewhere along with the hazards associated with transportation of contaminated soil and migration of contaminants from landfill into an adjacent environment. Soil washing for removing contaminated soil is an alternative way to excavation and disposal to landfill. This method is very costy and produces a residue rich in heavy metals, which will require further treatment. Moreover, these physio-chemical technologies used for soil remediation render the land usage as a medium for plant growth, as they remove all biological activities [1].Recent concerns regarding the environmental contamination have initiated the development of appropriate technologies to assess the presence and mobility of metals in soil [4], water, and wastewater. Presently, phytoremediation has become an effective and affordable technological solution used to extract or remove inactive metals and metal pollutants from contaminated soil. Phytoremediation is the use of plants to clean up a contamination from soils, sediments, and water. This technology is environmental friendly and potentially costeffective. Plants with exceptional metal-accumulating capacity are known as hyperaccumulator plants [5]. Phytoremediation takes the advantage of the unique and selective uptake capabilities of plant root systems, together with the translocation, bioaccumulation, and contaminant degradation abilities of the entire plant body [3].Many species of plants have been successful in absorbing contaminants such as lead, cadmium, chromium, arsenic, and various radionuclides from soils. One of phytoremediation categories, phytoextraction, can be used to remove heavy metals from soil using its ability to uptake metals which are essential for plant growth (Fe, Mn, Zn, Cu, Mg, Mo, and Ni). Some metals with unknown biological function (Cd, Cr, Pb, Co, Ag, Se, Hg) can also be accumulated [5].The objectives of this paper are to discuss the potential of phytoremediation technique on treating heavy metal-contaminated side, to provide a brief view about heavy metals uptake mechanisms by plant, to give some description about the performance of several types of plants to uptake heavy metals and to describe about the fate of heavy metals in plant tissue, especially on arsenic (As), lead (Pb), and mercury (Hg). This study is related to a research project that aims to identify potential plants in tropical country such as Malaysia which can uptake heavy metal contaminants from petrochemical wastewater.2. Heavy Metals: Sources and Effect in theEnvironmentHeavy metals are conventionally defined as elements with metallic properties and an atomic number >20. The most common heavy metal contaminants are Cd, Cr, Cu, Hg, Pb, and Zn. Metals are natural components in soil [6]. Some of these metals are micronutrients necessary for plant growth, such as Zn, Cu, Mn, Ni, and Co, while others have unknown biological function, such as Cd, Pb, and Hg [1].Metal pollution has harmful effect on biological systems and does not undergo biodegradation. Toxic heavy metals such as Pb, Co, Cd can be differentiated from other pollutants, since they cannot be biodegraded but can be accumulated in living organisms, thus causing various diseases and disorders even in relatively lower concentrations [7]. Heavy metals, with soil residence times of thousands of years, pose numerous health dangers to higher organisms. They are also known to have effect on plant growth, ground cover and have a negative impact on soil microflora [8]. It is well known that heavy metals cannot be chemically degraded and need to be physically removed or be transformed into nontoxic compounds [1].2.1. Arsenic (As)Arsenic (atomic number 33) is a silver-grey brittle crystalline solid with atomic weight of 74.9, specific gravity 5.73, melting point 817°C (at 28 atm), boiling point 613°C, and vapor pressure 1 mm Hg at 372°C [9]. Arsenic is a semimetallic element with the chemical symbol “As”. Arsenic is odorless and tasteless. Arsenic can combine with other elements to form inorganic and organic arsenicals [10]. In the environment, arsenic is combined with oxygen, chlorine, and sulfur to form inorganic arsenic compounds. Inorganic arsenic compounds are mainly used to preserve wood. Organic arsenic compounds are used as pesticides, primarily on cotton plants [11].Arsenic exists in the −3, 0, +3, and +5 valence oxidation states [9], and in a variety of chemical forms in natural waters and sediments [12]. Environmental forms include arsenious acids (H3AsO3, H3AsO3, ), arsenic acids (H3AsO4, , ), arsenites, arsenates, methylarsenic acid, dimethylarsinic acid, and arsine. Two most common forms in natural waters arsenite () and inorganic arsenate (), referred as As3+ and As5+ [9]. From both the biological and the toxicological points of view, arsenic compounds can be classified into three major groups. These groups are inorganic arsenic compounds, organic arsenic compounds, and arsine gas [13]. It is a hard acid and preferentially complexes with oxides and nitrogen. Trivalent arsenites predominate in moderately reducing anaerobic environments such as groundwater [9]. The most common trivalent inorganic arsenic compounds are arsenic trioxide, sodium arsenite, and arsenic trichloride [13]. Trivalent (+3) arsenates include As(OH)3, , AsO2OH2−, and [9]. Arsenite (As(OH)3, As3+) is predominant in reduced redox potential conditions [12].Arsenic is one of the contaminants found in the environment which is notoriously toxic to man and other living organisms [14]. It is a highly toxic element that exists in various species, and the toxicity of arsenic depends on its species. The pH, redox conditions, surrounding mineral composition, and microbial activities affect the form (inorganic or organic) and the oxidation state of arsenic. It is generally accepted that the inorganic species, arsenite [As3+] and arsenate [As5+], are the predominant species in most environments, although the organic ones might also be present [15].In general, inorganic compounds of arsenic are regarded as more highly toxic than most organic forms which are less toxic [10, 14, 16, 17]. The trivalent compounds (arsenites) are more toxic than the pentavalent compounds (arsenates) [16, 17]. It has been reported that As3+ is 4 to 10 times more soluble in water than As5+. However, the trivalent methylated arsenic species have been found to be more toxic than inorganic arsenic because they are more efficient at causing DNA breakdown [17]. Although As5+ tends to be less toxic compared to of As3+, it is thermodynamically more stable due to it predominates under normal conditions and becomes the cause of major contaminant in ground water [14]. Arsenate which is in the pentavalent state (As5+) is also considered to be toxic and carcinogenic to human [18].2.2. Lead (Pb)Lead (Pb), with atomic number 82, atomic weight 207.19, and a specific gravity of 11.34, is a bluish or silvery-grey metal with a melting point of 327.5°C and a boiling point at atmospheric pressure of 1740°C. It has four naturally occurring isotopes with atomic weights 208, 206, 207 and 204 (in decreasing order of abundance). Despite the fact that lead has four electrons on its valence shell, its typical oxidation state is +2 rather than +4, since only two of the four electrons ionize easily. Apart from nitrate, chlorate, and chloride, most of the inorganic salts of lead2+ have poor solubility in water [19]. Lead (Pb) exists in many forms in the natural sources throughout the world and is now one of the most widely and evenly distributed trace metals. Soil and plants can be contaminated by lead from car exhaust, dust, and gases from various industrial sources. Pb2+ was f

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Nanoenhanced Materials for Reclamation of Mine Lands and Other Degraded Soils: A Review

Author

Ruiqiang Liu, Rattan Lal 

Published: 1 January 2012

 by Hindawi Publishing Corporation

 in Journal of Nanotechnology

Journal of Nanotechnology, Volume 2012, pp 1-18; doi:10.1155/2012/461468 

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Abstract: Successful mine soil reclamation facilitates ecosystem recovery, minimizes adverse environmental impacts, creates additional lands for agricultural or forestry uses, and enhances the carbon (C) sequestration. Nanoparticles with extremely high reactivity and deliverability can be applied as amendments to improve soil quality, mitigate soil contaminations, ensure safe land–application of the conventional amendment materials (e.g., manures and biosolids), and enhance soil erosion control. However, there is no report on using nanoenhanced materials for mine soil reclamation. Through reviewing the up-to-date research results on using environment-friendly nanoparticles for agricultural soil quality improvement and for contaminated soil remediation, this paper synthesizes that these nanomaterials with high potentials for mine soil reclamation include zeolites, zero-valent iron nanoparticles, iron oxide nanoparticles, phosphate-based nanoparticles, iron sulfide nanoparticles and C nanotubes. Transport of these particles in the environment and their possible ecotoxicological effects are also discussed. Additionally, this article proposes a practical and economical approach to applying nanotechnology for mine soil reclamation: adding small amounts of nanoparticles to the conventional soil amendment materials and then applying the mixtures for soil quality improvements. Hence the cost of using nanoparticles is reduced and the benefits of both nanoparticles and the conventional amendment materials are harnessed.1. IntroductionEver since the commencement of industrial-scale mining of coal and other minerals, drastic environmental impacts have been recorded arising from both the mined lands and from the wastes left behind at the surface [1, 2]. The local landscape and the soil quality are among the most severely disturbed environmental components by the mining processes through directly stripping the vegetation and soil layers (open-pit mining) and/or through depositing the ores and mining wastes on the soils [3]. Dramatic alterations of the geological environment of the coal/ores and the mining wastes significantly reduce the chemical stability of the minerals, resulting in the release of the environmental disruptive chemicals into the soils and creating the “mine soils.” Typical mine soils often refer to the antecedent or original soils which are affected and degraded by the acid drainage and mining wastes. Practically, this type of soils also include the exposed parent materials due to accelerated soil erosion and/or the top soil removal for open pit mining, and the deposition of mining solid wastes. Although the properties vary from location to location depending on the local geochemistry, a mine soil is usually acidic, heavy-metal laden, nutrient depleted, highly compacted, and not favorable to plant growth [4].The strategy for mine soil reclamation is to minimize the environmental impacts of mining by restoring the mine soils and the local ecosystems to the antecedent levels. An adequate reclamation of mine soils not only benefits the local environment but also can contribute to improving the global environment through carbon (C) storage in biomass and in the soils, and thus off-setting the increase of CO2 emissions from industrial activities. The depleted and drastically disturbed mine soils have a larger potential of C storage over agricultural soils due to the fact that intensively cultivated soils contain relatively high soil organic matter (SOM) and further increasing the C sink capacity is difficult. In contrast, mine soils usually contain low soil organic C (SOC) and thus possess high C storage potentials. Taking full advantages of the available C sink capacity by growing vegetation at the abandoned mining sites would increase the atmospheric CO2 adsorption and enhance the terrestrial C sequestration [5–7].Reclamation of mine soils for C sequestration requires high quality of remediation techniques and treatments. It is not enough just to protect against acid mine drainage (AMD) and heavy metals from contaminating the ground and surface waters. The mine soils must be reclaimed in situ so that the grasses, crops, or trees could grow sustainably with limited management. On a long run, a quick establishment of the vegetation and enhancing microbial activities in a mine soil can improve soil quality by accentuating phytoremediation of the contaminants, decreasing soil erosion, and enhancing concentrations of SOM and plant nutrients.1.1. Mine Soil Quality and Amendment MaterialsHarnessing an effective CO2 sink of a mine soil site requires establishment and maintenance of a healthy forest or other vegetation cover for a time scale of at least 25 years. Soils with high quality are indispensable to support the vegetation that can thrive and sustain itself. However, mine soils, especially the gob piles or mining rock wastes, usually have poor soil quality such as low SOM content, low fertility, micronutrient imbalance or toxicity, low N and P availability, soil compaction caused by the grading operations, shallow soil depth, low moisture holding capacity, high electrical conductivity, high heavy metal contents, and extreme pH, which all adversely affect vegetation establishments and SOC sequestration [5]. Therefore, soil amendments and proper management are needed in order to improve the soil physical, chemical, and biological properties at a disturbed site for establishing vegetation and making it an effective atmospheric CO2 sink. Several natural minerals and agricultural, industrial and municipal wastes have been tried for these purposes as soil amendments. For example, manures [8, 9], composts [10], biosolids [9, 11, 12], and paper mill sludge [9] have been successfully applied to increase the SOM content in the mine soils. Limestones, zeolites [13], and coal combustion byproducts [14–16] (e.g., fly ash, bottom ash, and flue gas desulfurization (FGD) gypsum) have also been intensively researched in reducing mine soil acidity and decreasing the heavy metal toxicity and uptake by plants. A range of various commercial N, P, and K fertilizers have been used to provide adequate nutrients for the vegetation establishment at mining sites as well [8, 12, 17]. The land applications of these conventional amendment materials are also encouraged by the increasing demands on disposal and reuse of these industrial by-products and community wastes at low cost. However, various levels of heavy metals (e.g., Hg, Cd, Cr, and Pb) and other toxic elements (B, As, Se, and Mo) often occur in coal combustion by-products [18]. Nuisance odors, the potential of pathogen transmission, and presence of toxic and persistent organic chemicals and metals in biosolids have for the most part limited the use of land applications [19]. A survey study [20] on 9 different biosolids produced by municipal wastewater treatment plants in 7 USA states indicated that some biosolids were highly enriched in organic wastewater contaminants (OWCs), suggesting the land application of the solids might become a potential nonpoint source of OWCs into the environment. The OWCs included pharmaceuticals, hormones, detergent metabolites, fragrances, plasticizers, and pesticides. Therefore, new types of effective and environmentally safe soil amendment materials are urgently needed for mine soil reclamation.1.2. Using Nanoenhanced Materials as Soil AmendmentsNanotechnology is an advanced modern approach. It provides new types of materials which offer the unique and important solutions to the limitations of other conventional materials and have numerous applications [21]. Nanomaterials and nanostructures have nanoscaled dimensions that range from 1 to 100 nm and often exhibit novel and significantly changed physical, chemical, and biological properties as a result of their structure, larger specific surface area, and quantum effects that occur at the nanoscale [21]. Applications of nanotechnology in water treatment and purification have witnessed significant developments in recent years [22–24]. However, little progress has been made regarding the application of nanoparticles to improve agricultural soil quality and to reclaim the drastically disturbed lands. Lal [25] proposed that applying nanotechnology in agricultural sector was one of the available options to increase the agricultural production, solve environmental problems, and feed the world’s growing population. Hence, it is imperative to review the state of the science of nanotechnology that has potentials in mine soil reclamation and mine soil quality improvement and to explore the feasibility of using nanoenhanced materials as replacements for the conventional amendment materials in agriculture. The specific nanotechnology interested in this paper encompass those able to increase soil pH and fertility, improve soil physical structures, reduce mobility, availability, and toxicity of heavy metals and other environmental contaminants and those able to stabilize the soil components and abate soil erosion at a mining site. Therefore, the overall objectives of this paper are to (a) review the available literature on various environmentally-friendly nanoenhanced materials which could be used as in situ soil amendments for mine soil reclamation; (b) briefly discuss the transport and mobility of those nanoparticles in the environment as well as their possible ecotoxicological effects (if any); (c) propose a practical approach to application of the nanomaterials in mine soil reclamation at low cost and in a more environmentally friendly fashion.2. Nanomaterials for Soil Reclamation and Environmental RemediationReclamation of mine soils involves removing soil contaminants and enhancing soil quality and fertility. Nanotechnology is a promising approach for these purposes. Two advantages of nanomaterials over the traditional amendments for soil reclamation include the higher reactivity due to smaller particle size and highe

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Impact of Long-Term Forest Enrichment Planting on the Biological Status of Soil in a Deforested Dipterocarp Forest in Perak, Malaysia

Author

D. S. Karam, A. Arifin, O. Radziah, J. Shamshuddin, N. M. Majid, A. H. Hazandy, I. Zahari, A. H. Nor Halizah, T. X. Rui 

Published: 1 January 2012

 by Hindawi (The Scientific World)

 in The Scientific World JOURNAL

The Scientific World JOURNAL, Volume 2012, pp 1-8; doi:10.1100/2012/641346 

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Abstract: Deforestation leads to the deterioration of soil fertility which occurs rapidly under tropical climates. Forest rehabilitation is one of the approaches to restore soil fertility and increase the productivity of degraded areas. The objective of this study was to evaluate and compare soil biological properties under enrichment planting and secondary forests at Tapah Hill Forest Reserve, Perak after 42 years of planting. Both areas were excessively logged in the 1950s and left idle without any appropriate forest management until 1968 when rehabilitation program was initiated. Six subplots (20 m × 20 m) were established within each enrichment planting (F1) and secondary forest (F2) plots, after which soil was sampled at depths of 0–15 cm (topsoil) and 15–30 cm (subsoil). Results showed that total mean microbial enzymatic activity, as well as biomass C and N content, was significantly higher in F1 compared to F2. The results, despite sample variability, suggest that the rehabilitation program improves the soil biological activities where high rate of soil organic matter, organic C, N, suitable soil acidity range, and abundance of forest litter is believed to be the predisposing factor promoting higher population of microbial in F1 as compared to F2. In conclusion total microbial enzymatic activity, biomass C and biomass N evaluation were higher in enrichment planting plot compared to secondary forest. After 42 years of planting, rehabilitation or enrichment planting helps to restore the productivity of planted forest in terms of biological parameters.1. IntroductionMalaysia is a country rich in biodiversity of which natural forest is a home for thousands of flora and fauna [1]. However, the need for development and urbanization catalysed by the pressure of rising human population has made vast area of natural forests cleared up to cultivate new area for housing and wood productions. Liebig et al. [2] stated that the fertility of soil proportionally change with time catalyzed by natural phenomena and human activities. Hence, deforestation of natural forest leads to soil degradation, which proceeds rapidly under tropical climatic conditions [3, 4]. Forest rehabilitation is believed to be one of the best ways to overcome and lower down the demand for woody and nonwoody products from natural forest. Besides that, forest plantation also supports the shortage of wood supply, while sustaining world ecosystem [3]. In addition, forest plantation is also known as an alternative way to restore degraded sites to its original condition and sustains its soil fertility [5, 6]. Insam [7] found that soil fertility and its management are the most crucial part to evaluate a particular site of soil ecological area which gives a preview of the site’s environmental management and the extent of success for a particular forest rehabilitation program which can only be identified through its soil fertility evaluation.Enrichment planting is one of important technique used in forest rehabilitation [8, 9]. Montagnini et al. [10] defined enrichment planting as the introduction of valuable species to degraded forests without the elimination of valuable individual which already existed at that particular site. Adjers et al. [11] summarized that there are total of 25857 ha of forest plantation had been planted through enrichment planting technique in Peninsular Malaysia. Shorea acuminata, S. leprosula, Dryobalanops aromatica, and D. oblongifolia are among the favorite species planted in Peninsular Malaysia [12]. While for secondary forest, it is a forest area which has regrown trees after major disruption and disturbance such as fire and deforestation. Normally, the regeneration of plants species in secondary forests are done naturally by itself without any forest treatment given for a period of long time till the effect of disturbance is no longer noticed.It is undeniable that soil microorganism is the major agents in promoting nutrient cycling including carbon (C), nitrogen (N), phosphorus (P), and sulphur (S). Furthermore, Gaspar et al. [13] concluded that soil microbial biomass comprises 1–4% and 2–6% of total organic C and N in soil, respectively. Rapid turnover of microbial activities in soil is dependent on the changes occurring in the surrounding environment such as climate change, disturbance, and pollutant toxicity [14, 15] which made microbial activity a good sensitive indicator [16] for soil fertility evaluation. Islam and Weil [17] also stressed the importance of including microbial biomass evaluation to describe the status of fertility and quality of soil at a particular study site.Enzymatic activities are also one of the important evaluation aspects for determining soil fertility. They play a vital role in the organic residues degradation, humic substance synthesis, pollutant degradation, and nutrient cycles in soil [18]. Fluorescein diacetate (FDA) hydrolysis assay provides a reliable estimation of overall microbial activity in soil [19] and is widely used to analyse bacterial and fungal enzymatic activities [20, 21]. In addition, FDA analysis is considered as nonspecific because it is hydrolysed by various types of enzymes which include protease, esterase, and lipase [13, 21]. Heal and Maclean [22] found that approximately 90% of the energy transfer cycle in the soil was via microbial decomposer, and total microbial activity illustrates a general measurement of the organic matter turnover. Behera and Sahani [5] stated the importance of including biological studies, such as the evaluation of microbial biomass in land evaluations, because they provide a better indication of changes or degradation in forest soils than carbon and nitrogen analyses. Vásquez-Murrieta et al. [23] also stated that the key factors regulating and maintaining continuous supplies of nutrients in the soil for plant uptake are circulated by soil microbes. Soil fertility evaluation primarily focuses on the physicochemical properties in order to describe the growth performance of particular tree species at the plantation without taking into account the importance of soil biological properties as sensitive indicator to the changes occurring in the soil [24]. Hence, the objective of this study was to provide information and compare soil biological properties under enrichment planting and secondary forests after 42 years (as for 2010) of planting at Bukit Tapah Forest Reserve, Perak, Malaysia.2. Materials and Methods2.1. Description of the Study SiteThe study was carried out in enrichment planting (N 04.179394° E 101.31998°) and secondary forest (N 04.17336° E 101.31974°) at Bukit Tapah Forest Reserves, Perak (Figure 1) on 21st until 23rd July 2011. The mean annual rainfall and temperature are 2,417 mm and 24.5°C, respectively. The soils in this study area are classified as Ultisols, which are considered as highly weathered due to large amount of low-activity clays associated with high Al saturation [3]. All of the tree species of Shorea leprosula, S. bracteolata, and S. macroptera planted were done on 2nd February 1968, and the age of the trees was 42 years old in 2010, while adjacent secondary forest was left idle to undergo natural regeneration without any reforestation activity. Compartment 13 of Bukit Tapah is one of the 10 compartments that was gazetted for enrichment planting at Perak South District, Malaysia. About 1,185 hectares out of 64,984 hectares of Bukit Tapah Forest Reserve were converted to enrichment planting program of which compartment 13 covers 87.2 hectares of the forest reserves. The purpose of enrichment planting done at this area is to replace and curtail this particular area which had undergone excessive logging before 1968.Figure 1: Enrichment planting (F1) and secondary forest (F2) plots at Tapah Hill Forest Reserve, Perak, Malaysia (Scale 1 : 20 000).The size of the poly bags used to plant the seedlings was 10 cm × 15 cm × 23 cm. Twenty-six thousand five hundred and forty-four saplings were planted with 304 saplings per hectare, and the rates of survival recorded in 1970 found that only 9,158 trees managed to grow well and survive with resulting in 105 saplings per hectare, respectively. Shorea leprosula, S. parvifolia, S. bracteolata, and S. macroptera were the main species of Dipterocarpaceae planted in compartment 13 enrichment planting plot. The trees were planted on a 10 m × 3 m grid.2.2. Experimental Design and Soil SamplingThis study used a completely randomized design. Enrichment planting and secondary forest plots were designated as F1 and F2, respectively. Six subplots were demarcated in each plot in order to serve as replicates. Six soil samples were randomly collected at depths of 0–15 cm and 15–30 cm in each subplot. The samples were then mixed together to form a composite sample for each soil depth range. Hence, 12 composite samples (six from soil depth 0–15 cm and six from soil depth 15–30 cm) were collected from each plot for the analysis. The composite samples were kept in UV-sterilized polyethylene bags at 0°–4°C.2.3. Total Microbial PopulationSpread-plate technique or direct count of colony forming unit was used to evaluate the estimation of microbial population [25, 26]. Nutrient agar was used for bacterial culture. Dilution factor of 10−2, 10−3, and 10−4 was found to be suitable for colony calculation after few pilot test carried out to standardize the dilution factor for every population counts. The number of colony forming units per gram soil was calculated using the following equation: =numberofcolonyformingunits/gofdryweightsoil(meanplatecount)(dilutionfactor),(dryweightsoil,initialdilution)(1) where dry weight soil = (Weight of moist soil, initial dilution blank) × [(1 − % moisture soil sample)/100]. The results were expressed in log10 g−1 soil.2.4. Microbial Enzymatic ActivityFluorescein diacetate (FDA) hydrolysis assay illustrated by Sánchez-Monedero et al. [18] and Gagnon et al. [27] was used to evaluate m

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Arbuscular Mycorrhizal Colonization Enhanced Early Growth of Mallotus paniculatus and Albizia saman under Nursery Conditions in East Kalimantan, Indonesia

Author

Dewi Wulandari, Saridi, Weiguo Cheng, Keitaro Tawaraya 

Published: 1 January 2014

 by Hindawi Publishing Corporation

 in International Journal of Forestry Research

International Journal of Forestry Research, Volume 2014, pp 1-8; doi:10.1155/2014/898494 

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Abstract: Forest over logging, forest fire, forest conversion, and opencast mining have promoted deforestation in Indonesia, and reforestation is needed immediately. However, reforestation is limited by low seedling quality and production, and slow seedling growth in nurseries. Native tropical tree and fast-growing species, Mallotus paniculatus and Albizia saman, are potential to promote the first rotation of reforestation. Arbuscular mycorrhizal (AM) fungi are known to promote nutrient uptake and plant growth. We examined the effects of two native AM fungi, Gigaspora decipiens and Glomus clarum, on the growth of M. paniculatus and A. saman seedlings under nursery conditions. At harvest, after six months, we determined AM colonization, shoot dry weight, and shoot N and P concentration. Approximately 90% and 50% of M. paniculatus and A. saman roots, respectively, were colonized by AM fungi, without any difference between the inoculation treatments. G. decipiens and G. clarum increased shoot height, leaf number, shoot dry weight, and shoot N and P uptake of both species. A positive correlation was observed between N and P uptake and shoot dry weight. These results suggest that AM fungi are effective in accelerating nutrient uptake and plant growth, which will, in turn, promote reforestation and sustainable forest timber production.1. IntroductionIn Indonesia, deforestation is occurring rapidly owing to over logging, forest fire, forest conversion into agricultural land or oil rubber plantation [1], and opencast mining [2], and, therefore, immediate restoration is required by applying a comprehensive and systematic reforestation method. Natural forest recovery, particularly in forestland used for bare opencast mining, requires several hundred years and consists of the initial, middle, and climax stages [3]. Pioneering and light-requiring species, such as leguminous trees, grasses, and shrubs, are established first [4] in the initial stage of forest succession, followed by gap-opportunistic species (Meliaceae, Dipterocarpaceae, and Flindersia spp.) in the middle stage, and finally shade-tolerant species in the mature or climax stage [5]. The preparation of seedlings of native tree species [6] and the selection of fast-growing leguminous species [7] with improved nitrogen (N), phosphorus (P), and potassium (K) uptake and biomass production are vital for the initial stage of reforestation.Mallotus paniculatus (Euphorbiaceae) is distributed throughout the Malesian region, including Indonesia [8]. As an evergreen timber tree [9], this plant is an important pioneer species in Kalimantan, Indonesia, because it contributes to the aboveground biomass in secondary forests [10].Albizia saman (Fabaceae) is native to Northern South America and has become naturalized in the tropics [11]. A. saman is usually planted for agroforestry [11] and timber purposes [12]. As a moderately fast growing species [11], A. saman has a high survival rate [13] and grows in a wide range of climatic conditions [11], making it potentially useful for reforestation.The rapid production of M. paniculatus and A. saman seedlings in nurseries is important for successful reforestation. However, the initial growth of A. saman is slow [11]. Furthermore, poor nutrient uptake due to the low fertility and the high acidity of the tropical soil in Indonesia has made it difficult to improve the seedling growth.Manure or green compost is a substrate usually used together with the amendment of other organic nutrients or fertilizers in nurseries to meet the nutritional requirements for plant growth. As fertilizer application is costly, it is important to adopt an inexpensive and environmentally friendly method to meet the nutritional requirements for plant growth improvement. As for the plants grown in pots in nursery, the possible contact between plants with soil microorganisms in the ground including AM fungi is very low. The application of symbiotic soil microorganisms that may facilitate nutrient transfer from substrate to plant may enhance nutrient uptake efficiency. It is well documented that arbuscular mycorrhizal (AM) fungi can improve seedling growth, tree height, and plant yield [14] by increasing nutrients [15]. Considering these abilities, AM fungi can assist plant nutrient uptake and therefore promote seedling growth under nursery conditions.AM colonization was observed in M. paniculatus [16] and A. saman [17]. Inoculation with AM fungi increased the height and shoot dry weight of Macaranga denticulata (Euphorbiaceae) [18]. Inoculation with AM fungi also changed the chlorophyll, carotenoid, sugar, and protein contents of A. saman in nurseries in India [19]. However, they did not measure the AM fungi colonization and its effect on nutrient uptake and growth. Therefore, to the best of our knowledge there is no information about the effect of AM fungal colonization on the N and P uptake and growth of M. paniculatus and A. saman. We hypothesize that the AM fungal inoculation of M. paniculatus and A. saman leads to AM colonization, thereby improving N and P uptake and growth of these plant species.In order to enhance the growth of M. paniculatus and A. saman in nurseries, two AM fungal species, Gigaspora decipiens Hall & Abbott and Glomus clarum Nicholson & Schenck, were inoculated. Those two AM fungal species were used because they are indigenous to Kalimantan and they have the ability to improve nutrient uptake and growth of tropical peat-swamp plants [20]. The objective of this research was to determine whether inoculation with the two native AM fungal species could improve N and P uptake and growth of native M. paniculatus and A. saman under nursery conditions in Binungan, Tanjung Redeb, Berau Regency, East Kalimantan, Indonesia.2. Materials and Methods2.1. Soil PreparationCompost is one of the most common substrates used to grow seedling in pot in nursery in Indonesia. Compost is used to reduce chemical fertilizer application and to prepare good healthy seedling before transplanting it to the field which is usually covered by compost. Compost was collected from a local area in Binungan (N02° 02′, E117° 27′), Tanjung Redeb, Berau Regency, East Kalimantan, Indonesia. Examination of the compost substrate used in the nursery revealed no spores of AM fungi, indicating that the compost substrate possibly did not support spontaneous AM colonization. To distinguish other soil fungi or soil microorganisms, the compost was sterilized in a drum by heating over wood fire for 3 hours and further stored at room temperature. The compost chemical characteristics before sterilization showed that the available P [21] was 622 mg P2O5 kg−1, total N concentration was 26.5 g kg−1, and C concentration was 372.1 g kg−1 (Sumigraph N-C 220 F). The C : N ratio was 14.04, pH H2O was 5.25, and pH KCl was 4.82.2.2. Inoculum Preparation and AM Fungi InoculationGlomus clarum Nicholson & Schenck and Gigaspora decipiens Hall & Abbott were isolated from peat soil in Kalampangan (S2° 13′, E113° 56′), Palangkaraya, Central Kalimantan, Indonesia [20]. Pueraria javanica Benth was cultivated in zeolite to propagate those two AM fungal species under greenhouse conditions for 90 days. The roots, zeolite, and spores were used as the AM fungal inoculum. Inoculation with AM fungi was accomplished by mixing 20 g of the inoculum with 800 g of the sterilized compost in a polyethylene bag (10 cm diameter × 15 cm height). Noninoculated compost was prepared by adding more 20-gram sterilized compost into 800-gram sterilized compost as control.2.3. Seed GerminationSeeds of M. paniculatus (Lamk.) Müll. Arg. were collected from natural forests in Binungan. Seeds of A. saman (Jacq.) Merr. were purchased from a local seed company, Bogor, West Java, Indonesia. The A. saman seeds were soaked in water at 80°C for 1-2 minutes [11]. No scarification was applied to the seeds of M. paniculatus. Approximately five to seven seeds were sown in the inoculated and noninoculated composts at a depth of 1 cm on 29 September 2011. One seedling per polyethylene bag was grown after germination. Due to the large size of the A. saman seedlings, the seedlings were transferred to larger polyethylene bag (15 cm diameter × 20 cm height) four months after sowing on 30 January 2012 by using the same sterilized compost. No fertilizer was applied. The experimental design used was completely randomized design (CRD) where the seedlings in the polyethylene bag were arranged randomly in the nursery. Tap water was applied once every two days. The seedlings were grown under 50% shade from 29 September 2011 to 8 March 2012 in the nursery in Binungan. This plant growth period was determined to get appropriate size of seedling for transplanting to the field.2.4. Growth Parameters M. paniculatus and A. saman seedlings were subjected to three treatments: (1) control, (2) inoculation with G. clarum, and (3) inoculation with G. decipiens. Each treatment had 20 replications and shoot height and leaf number were measured two, three, and four months after sowing. Six months after sowing, the seedlings were sampled and five replications were harvested for each treatment. The 15 remaining seedlings were left for transplanting to field. Shoots and roots were separately harvested. Shoots were oven dried at 70°C for 72 hours and weighed. Ground shoots were digested with HNO3-HClO4-H2SO4 solution. P concentration in the solution with the digested ground shoots was determined colorimetrically with the vanadomolybdate-yellow assay [22] using a spectrophotometer at 880 nm absorbance (Hitachi, U-2900). Shoot N concentration was determined using a Sumigraph N-C 220 F. Shoot P and N contents were calculated by multiplying the shoot nutrient concentration and the shoot dry weight.2.5. Assessment of AM ColonizationAM colonization was observed by harvesting the roots of M. paniculatus and A. saman. The roots were cleared in KOH (100 g L−1) at 80°C for 15 minutes, acidified ..

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Structure of Fungal Communities in Sub-Irrigated Agricultural Soil from Cerrado Floodplains

Published Date

Diversity 2016, 8(2), 13; doi:10.3390/d8020013

Author

Elainy Cristina A. M. Oliveira ¹

, Acacio A. Navarrete ²

, Joenes M. Peluzio ³

, Waldesse P. de Oliveira Junior ³

, Alana de A. Valadares ⁴

, Siu M. Tsai ²

 and Paula B. de Morais ^5,* 

¹

Department of Agricultural Sciences and Technology, Universidade Federal do Tocantins, Rua Badejos, 69-Jd Sevilha, Campus Universitário, Gurupi 77410-530, TO, Brazil

²

Cell and Molecular Biology Laboratory, Center for Nuclear Energy in Agriculture, University of São Paulo, Avenida Centenario 303, Piracicaba 13400-970, Brazil

³

Biotechnology Laboratory, Universidade Federal do Tocantins, Quadra 109 Norte, Av. NS 15, ALCNO 14 s/n, Campus Universitário, Palmas 77001-090, TO, Brazil

⁴

Department of Environmental Technology and Water Resources, University of Brasília, Brasilia 70040-010, Brazil

⁵

Environmental Microbiology and Biotechnology Laboratory, Universidade Federal do Tocantins, Quadra 109 Norte, Av. NS 15, ALCNO 14 s/n, Campus Universitário, Palmas 77001-090, Brazil

^*

Author to whom correspondence should be addressed. 

Academic Editor: Raymon Shange

Received: 1 November 2015 / Revised: 4 April 2016 / Accepted: 16 May 2016 / Published: 19 May 2016

(This article belongs to the Special Issue Soil Microbes Diversity and Soil Function)

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Abstract 

This study aimed to evaluate the influence of soybean cultivation on the fungal community structure in a tropical floodplain area. Soil samples were collected from two different soybean cropland sites and a control area under native vegetation. The soil samples were collected at a depth of 0–10 cm soil during the off-season in July 2013. The genetic structure of the soil fungal microbial community was analyzed using the automated ribosomal intergenic spacer analysis (ARISA) technique. Among the 26 phylotypes with abundance levels higher than 1% detected in the control area, five were also detected in the area cultivated for five years, and none of them was shared between the control area and the area cultivated for eight years. Analysis of similarity (ANOSIM) revealed differences in fungal community structure between the control area and the soybean cropland sites, and also between the soybean cropland sites. ANOSIM results were confirmed by multivariate statistics, which additionally revealed a nutrient-dependent relation for the fungal community structure in agricultural soil managed for eight consecutive years. The results indicated that land use affects soil chemical properties and richness and structure of the soil fungal microbial community in a tropical floodplain agricultural area, and the effects became more evident to the extent that soil was cultivated for soybean for more time. View Full-Text

Keywords: land use change;  ARISA;  fungi;  tropical soil

This is an open access article distributed under the Creative Commons Attribution License which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. (CC BY 4.0).

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