Next Article in Journal
Methodological Approach to Reconstructing Lost Monuments from Archaeological Findings: The San Francesco di Castelletto Church in Genoa
Previous Article in Journal
Mineralogy of Chub Lake-Type Hematite Deposits in St. Lawrence County, NY
Previous Article in Special Issue
Investigation of Spectral Variation of Pine Needles as an Indicator of Arsenic Content in Soils
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Minerals
Volume 9
Issue 10
10.3390/min9100568
minerals-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

Arsenic (As) Removal Using Talaromyces sp. KM-31 Isolated from As-Contaminated Mine Soil

by
In-Hyun Nam
1,†,
Kumarasamy Murugesan
2,†,
Jungho Ryu
1,* and
Jae Hwan Kim
3,*
1
Geologic Environment Research Division, Korea Institute of Geoscience and Mineral Resources (KIGAM), Daejeon 34132, Korea
2
Department of Environmental Science, Periyar University, Salem 636011, India
3
Department of Human Environment Design, College of Science, Cheongju University, Cheongju 28503, Korea
*
Authors to whom correspondence should be addressed.
†
These authors equally contributed this work.
Minerals 2019, 9(10), 568; https://doi.org/10.3390/min9100568
Submission received: 7 June 2019 / Revised: 16 August 2019 / Accepted: 19 September 2019 / Published: 20 September 2019
(This article belongs to the Special Issue Arsenic Geochemistry and Health)
Browse Figures
Versions Notes

Abstract

:
Bioremediation is an environmentally-benign and cost-effective approach to removing arsenic from contaminated areas. A fungal strain hyper-tolerant to arsenic was isolated from soil from a mine site and used for the removal of arsenic. The isolated fungus was identified as Talaromyces sp., and its growth rate, arsenic tolerance, and removal rates were investigated for As(III) and As(V). Arsenic tolerance tests revealed that the fungus was highly resistant to arsenic, tolerating concentrations up to 1000 mg/L. Robust mycelial growth was observed in potato dextrose broth containing either As(III) or As(V), and there was no difference in growth between that in arsenic-free medium and medium amended with up to 300 mg/L of either arsenic species. The isolate showed relatively low growth rates at As(V) concentrations >500 mg/L, and almost no growth at As(III) concentrations >300 mg/L. Both arsenic species were effectively removed from aqueous medium (>70%) in tests of the biosorption of arsenic onto mycelial biomass. Surface modification of the biomass with Fe(III) (hydr)oxides significantly enhanced arsenic removal efficiency. The findings indicate that this soil fungal strain has promise for use in bioremediation strategies to remove arsenic from highly contaminated aqueous systems.

1. Introduction

Arsenic (As) is a major pollutant in the environment, derived from the weathering of rocks and volcanic activity, and from anthropogenic inputs including industrial activities, mining operations, and agriculture [1,2]. In the natural environment, arsenic is present in a variety of inorganic forms such as As(V) and As(III), and organic forms such as monomethylarsonic acid (MMA), dimethylarsinic acid (DMA), and trimethylarsenic oxide (TMA). Microbes can convert arsenic from one form to another via oxidation/reduction and methylation/demethylation processes [3]. As(V) occurs predominantly under oxidizing conditions, whereas As(III) is mainly found under reducing conditions [4]. In particular, As-contaminated soil is of concern, as human exposure to arsenic can occur through both the food chain and the water supply [2,5]. Various technologies to remediate arsenic-polluted soil and water have been developed, including stabilization and solidification, but the application of these methods has been limited for economic reasons and because of disposal problems [6,7].
Microbial biogeochemical transformation of inorganic elements is a result of assimilatory processes in which an element is incorporated into biomass, and/or dissimilatory processes in which transformation results in energy generation and/or detoxification [8]. Fungi are able to tolerate and/or detoxify metals using several mechanisms, including redox transformation, biosorption, extracellular and intracellular precipitation, and active uptake. As a result, the responses of microorganisms to toxic metals can dramatically alter metal abundance and speciation [9]. Biosorption is considered to be a promising method for the removal of metal ions from aqueous solutions. Algae, fungi, and bacteria have been used as biomass-derived biosorbents for several metals [10,11,12]. Biosorption of toxic metals and metalloids by fungal biomass in both living and non-living forms provides a cost-effective and suitable technology for remediating soil and water contaminated by toxic metals [13,14,15].
Many fungal species have been found to be capable of arsenic removal, and the accumulation of arsenic by microorganisms has been recognized as a potential remediation method because it is environmentally-friendly and cost-effective [7]. Fungal strains capable of arsenic accumulation or biosorption include Phaeolus schweinitzii [16], Fusarium oxysporum [17], Sinorhizobium meliloti [18], Neosartorya fischeri [15], Aspergillus candidus [19], Trichoderma sp., Neocosmospora sp., Rhizopus sp. [20], and Paecilomyces sp. [21]. Given the variety of mechanisms of heavy metal resistance and the reported removal of arsenic by fungi, it is likely that arsenic-tolerant fungi with substantial arsenic removal ability will be present at As-contaminated sites. Therefore, it is important to expand investigations to screen native fungi present at As-contaminated sites for their potential for arsenic remediation.
The surface charge on fungal mycelium is typically negative over pH ranges of 3.0~10.0 [20,22]. Although few fungal species are able to remove or volatilize anionic metal(loid)s, including As(V), their negative surface charge is unfavorable for adsorptive removal of anionic pollutants. Therefore, we hypothesized that surface modification with a cationic agent could enhance arsenic removal ability. The synthesis and utilization of iron oxide materials having specific properties and functions have been widely investigated because of their high surface area to volume ratios [23]. In past studies [24], iron oxide materials have been widely used to remove a range of metalloids and heavy metals, including arsenic, lead, mercury, cadmium, and copper. Moreover, iron oxides have low toxicity, chemical inertness, and biocompatibility, and hence have enormous potential in biotechnology applications [25]. Surface-coating media and preparation methods play an important role in determining surface chemical properties, size distribution, morphology, and magnetic properties; therefore fungi- or protein-mediated biological methods for iron oxide nanomaterials as emerging methods necessitate wide development [26]. Nonetheless, iron oxide-based biotechnologies for heavy metal adsorption are at a relatively early stage in terms of wide application.
In the present study, microorganism in soil from an arsenic-contaminated mine was isolated and screened for arsenic removal ability, yielding in a single culturable fungal isolate. The objectives of this study were: (1) To explore the tolerance and growth of the arsenic-tolerant fungal isolate in the presence of As(III) and As(V); (2) to assess the arsenic removal and transformation activity of the arsenic-tolerant isolate; and (3) to test the removal of arsenic from an aqueous system using mycelial pellets and Fe(III) (hydr)oxides-coated dry fungal biomass.

2. Materials and Methods

2.1. Chemicals

Sodium arsenite (NaAsO2) and sodium arsenate dibasic heptahydrate (Na2HAsO4·7H2O) were obtained from Merck (Darmstadt, Germany). For quantification, an authentic analytical arsenic standard (1000 mg/L) was obtained from Junsei Chemicals (Kyoto, Japan). Potato dextrose broth and agar media were obtained from Fluka (Darmstadt, Germany). All chemicals used were of analytical grade or the highest purity.

2.2. Fungal Isolation and Molecular Identification

An arsenic-resistant filamentous fungus was isolated by selective enrichment of soil suspension (100-fold dilution) with arsenic-polluted soil (5000 mg/kg of total arsenic) collected at a former arsenic mine site located near the city of Cheongyang, Republic of Korea. The isolate was cultured in mineral medium containing 0.25% KH2PO4, 0.20% MgSO4, 0.50% (NH4)2SO4, 0.50% NaCl, and 0.25% glucose supplemented with 500 mg/L NaAsO2. The pH of the medium was maintained at 5.3 using 100 mM citrate phosphate buffer. The cultures were incubated at 28 °C for 7 days. Molecular identification of the arsenic-tolerant isolate was achieved by sequencing the internal transcribed spacers ITS1 and ITS2 of the test fungus. The ITS regions were PCR-amplified using the universal primers ITS1: 5′-TCCGTAGGTGAACCTGCGG-3′ and ITS4: 5′-TCCTCCGCTTATTGATATGC-3′. Extraction of genomic DNA was performed by using the Power Soil DNA extraction Kit (Mo Bio Laboratories Inc., Carlsbad, CA) and the extracts thus obtained were pooled prior to PCR amplification. PCR cycling conditions were as follows: One initial cycle of denaturation at 94 °C for 3 min, followed by 34 cycles of 94 °C for 30 s (denaturation), 60 °C for 30 s (annealing), and at 72 °C for 45 s (elongation), and an extension for 5 min at 72 °C. To identify the isolate, the PCR-amplified ITS DNA was sequenced, and the sequence was used in a BLAST search of the NCBI database. Pairwise nucleotide sequence comparisons, DNA alignments using CLUSTALW, and phylogenetic relationships to the most closely related fungal strains were analyzed using the MEGA 5.0 software package. The obtained sequence was deposited in GenBank (accession No. MK955349).

2.3. Arsenic Tolerance Assay

A fungal isolate was tested for tolerance to arsenic in potato dextrose agar (PDA) medium containing various concentrations (50, 100, 200, 300, 500, 800, and 1000 mg/L) of As(III) or As(V) as sodium arsenite and sodium arsenate, respectively. Mycelial discs (5 mm) from pure 7-day-old cultures grown on PDA were inoculated onto Petri dishes (9 cm) containing As(III) or As(V), and incubated at 30 °C. Mycelial growth was monitored by measuring the radial growth of the mycelial colony. For the determination of dry cell weight, the PDA plate was mixed with 100 mL of distilled water and boiled to dissolve the agar. The agar solution containing the fungal biomass was filtered through dry No. 5 Whatman filter paper, and the filter paper with the retained fungal cells was then dried in an oven at 60 °C for 24 h until a constant dry weight was attained. Arsenic tolerance was calculated by comparing the radial growth of each fungus in control medium containing no arsenic. A tolerance index was used, defined as the change in radius of the fungus in the presence of arsenic divided by its change in radius in the same period in the control. The fungal isolate that grew was regarded as tolerant to arsenic and was used in further experiments.

2.4. Arsenic Biosorption Experiments

An arsenic biosorption experiment was conducted using pre-grown mycelium of Talaromyces sp. KM-31. A culture of the fungus that had been grown in potato dextrose broth for 7 days was filtered onto a glass fiber filter disc (47 mm) and washed with deionized water. A known quantity of mycelial biomass (60 mg/mL of dry weight) was transferred into Erlenmeyer flasks containing deionized water and various concentrations of As(III) or As(V) (1~25 mg/L). Arsenic biosorption/removal was monitored at different time intervals over a period of 7 days. The uptake of arsenic by live and inactivated Talaromyces sp. KM-31 mycelium was analyzed using the Langmuir adsorption isotherm model, and biosorption efficiencies were compared [27]. Inactivated mycelium was obtained from fungal cultures grown in PDB that were dried in an oven at 80 °C for 24 h after harvesting.

2.5. Arsenic Removal by Modified Fungal Biomass

To assess whether arsenic removal is enhanced by coating with Fe(III) (hydr)oxides, the surface of the fungal biomass was modified by addition of a Fe(III) hydroxide or nano-sized Fe(III) oxide coating. For surface coating, wet mycelium was initially suspended in 1 M FeCl3·6H2O Fe(III) chloride solution, then 3 M NaOH or NaBH4 was added and the mixture stirred well to form particles of Fe(III) hydroxide or nFe(III)-oxide, respectively. The mycelium coated with Fe(III) (hydr)oxides was then separated and dried overnight in a vacuum oven at 90 °C. The dried mycelium was ground to a fine powder for use in arsenic removal experiments.

2.6. Analytical Methods

To determine the arsenic content, dried mine soil or treated fungal material was placed in a Teflon vessel and extracted in a microwave oven using concentrated nitric acid (HNO3), following EPA Method 3051A [28]. Following cooling, each sample was filtered and the volume was adjusted using ultra-pure water. Arsenic content was determined using inductively coupled plasma mass spectrometry (ICP-MS) (Thermo Elemental, Franklin, MA, USA). As speciation was conducted by using a solid phase extraction resin (SupelcleanTM LC-SAX SPE tube, Sigma-Aldrich, Seoul, Korea) that can remove anionic As(V) (Astotal – As(III) = As(V)). All samples were analyzed in triplicate to check precision. The analytical precision among replicates (n = 3) was within 5%. Final results were reported as mean values of the triplicate measurements, and were expressed as mg/kg dry weight. To determine the residual concentration of arsenic in the medium, samples of the growth medium in the flasks after incubation were centrifuged at 16,000 rpm for 20 min and the arsenic concentrations of the resulting cell-free supernatant was determined using ICP-MS.

3. Results and Discussion

3.1. Isolation and Identification of anAarsenic-Tolerant Fungal Strain

Only one type of fungus was cultured on both PDA and PDB from As-contaminated mine soil. The ITS DNA sequence of the purified fungal isolate showed maximum identity with the sequence of Talaromyces helicus, and phylogenetic analysis also showed that the isolate was closely related to T. helices (GenBank accession No. FJ430764). The purpose of this investigation was to obtain soil fungi from highly arsenic-contaminated mine soils for assessment of their potential for use in bioremediation of arsenic-contaminated aqueous systems. A wide range of fungi from all major taxonomic groups have been observed in metal-polluted habitats [29], and various fungi have been isolated from soil polluted with heavy metals in various parts of the world, including Aspergillus, Rhizopus, Penicillium, Fusarium, Chaetomium, Geomyces, and Paecilomyces spp. [12,14,29,30]. Talaromyces sp. are known to be tolerant of harsh conditions, making them potentially useful in extreme environments, including acidic and metal(loid)-contaminated mine drainage and soils [31,32]. Past studies have shown metal ion adsorption by Talaromyces sp. Romero et al. used T. helicus to degrade biphenyl treated with high levels of copper (Cu), and showed the detoxification abilities of this species and its adaptation to heavy metals and biacrylic compounds [33]. In addition, T. emersonii has been shown to have uranium (U) biosorption ability [34]. Using metal-resistant fungal strains in bioremediation strategies could increase the detoxification and biosorption of pollutants in contaminated areas.

3.2. Growth and Arsenic Tolerance of Talaromyces sp. KM-31

The isolated Talaromyces sp. KM-31 strain grew well on PDA medium augmented with up to 200 mg/L As(III) or 300 mg/L As(V) (Figure 1).
The isolate showed relatively low growth rates at As(V) concentrations >500 mg/L and almost no growth at As(III) concentrations >300 mg/L, indicating that, although the fungal strain was tolerant to arsenic, high concentrations inhibited growth. The isolate was more tolerant of As(V) than As(III) because the latter has a higher toxicity. After 15 days of incubation, the tolerance index values over a range of As(III) and As(V) concentrations were 1.0 or near that value, indicating that growth was not affected on solid PDA medium (Table 1 and Table S1).
In PDB media, the isolate grew well up to 200 mg/L As(III) or 300 mg/L As(V). The tolerance index results show that growth was completely inhibited at concentrations >300 mg/L As(III), while As(V) did not completely inhibit growth even at concentrations >500 mg/L. However, 30−65% inhibition of mycelial growth occurred at concentrations >300 mg/L, indicating that the isolate was highly tolerant of As(V) in the complex organic medium. For cultures 15 days old, the rate of removal of As(V) was 3- to 5-fold higher than that for As(III). The variation in tolerance of this strain to As(III) or As(V) may be exhibited by redox conditions because of the presence of one or more types of tolerance strategy or resistance mechanisms [35,36]. For different species of metal(loid), or among different fungal strains, differing biological mechanisms are involved in fungal survival [8,10,13,37]. In-vitro evaluation of tolerance of Aspergillus and Rhizopus over a wide range of arsenic concentrations [38] revealed that As-resistant isolates tolerated arsenate concentrations up to 1000 mg/L, and maintained their tolerance index in a range of 0.35 to 0.98 (Table 1). The radial growth patterns of Talaromyces sp. KM-31 in the present study appear to reflect tolerance development in the fungus (Table 1 and Figure 1); this could be used as a basis for selecting tolerant strains in metal(loid) removal assays. Intracellular accumulation of arsenic in vacuoles may play a major role in detoxification of arsenic in tolerant fungi [39], although other explanations for arsenic tolerance include detoxification by arsenic biomethylation and radial growth stimulation in the presence of different concentrations of arsenic in PDA media [40].

3.3. Arsenic Removal by Live Mycelial Pellets

Arsenic removal by live mycelium of Talaromyces sp. KM-31 was assessed in an aqueous system (Figure 2). Effective removal of relatively low concentrations of arsenic (1.0~5.0 mg/L) by live biomass was observed, but as the concentration increased, the extent of removal decreased. The live mycelium preferentially removed As(V) compared with As(III), and showed no reduction or oxidation of either arsenic form. This is consistent with the data on Talaromyces sp. KM-31 growth patterns (Figure 1 and Table 1).
The removal of As(V) was analyzed using adsorption isotherms (Figure 3), which showed that the experimental data fitted well with the Langmuir isotherm model. The isotherm studies revealed that the adsorption capacity increased as the concentration of As(V) increased. According to the Langmuir isotherm, the maximum arsenic absorption capacity was 0.005 mg/g for live mycelium and 0.0008 mg/g for inactivated mycelium, and the isotherm constants (KL) were 0.8948 L/mg and 1.027 L/mg for live and inactivated mycelium, respectively (Figure 3). The mycelial sorption of arsenic was directly estimated by digesting the mycelium after it had been exposed to arsenic, and then comparing the arsenic concentration in solution with that estimated based on removal efficiency. A mass balance difference was observed between the removed and residual arsenic contents, suggesting that the live mycelium may have volatilized the absorbed arsenic, in agreement with previous reports [41].
The inactivated mycelial biomass showed poor removal ability compared with the live mycelium, indicating that active arsenic biosorption mechanisms are involved [14]. The biosorption mechanisms may differ among fungal species depending on redox chemistry and biomass specificity [42,43]. Previous studies have reported that metal tolerant strains of fungi, including Rhizopus nigricans and Aspergillus spp., could accumulate cadmium (Cd) and chromium (Cr) from aqueous solution [38], although the biosorption levels varied among different fungal strains [44,45,46]. Similarly, in this study, we showed that the live mycelium of Talaromyces sp. KM-31 accumulated arsenic, but with greater tolerance of As(V). Biosorption of toxic metal(loid)s involves ionic species on the fungal cell surface, including extracellular polysaccharides, proteins, and chitins [14,38]. Most fungal species have chitin and chitosan as integral parts of their cell wall structure, and hence have potential for effective biosorption of metal(loid)s [47]. The metal biosorption by Rhizopus arrhizus was predominantly associated with cell wall chitin and chitosan [44], but deacetylated amino groups of glucosamine may also act as binding sites for metals [20,48]. Melanins and phenolic polymers also have oxygen-containing moieties, including carboxyl, phenolic, alcoholic hydroxyl, carbonyl, and methoxy groups, that could potentially act as metal-binding sites [29]. It has been proposed that arsenic-tolerant fungi isolated from contaminated soil reduce arsenic uptake by enhanced production of melanin, which may bind the toxic element [49]. Arsenic binding by fungi seems mainly to involve passive biosorption to the cell wall, which promotes arsenic uptake through intracellular accumulation [20,48].

3.4. Arsenic Removal by Modified Biomass of Talaromyces sp. KM-31

The potential to enhance metal(loid) accumulation or sorption by Talaromyces sp. KM-31 was assessed by coating the surface of the mycelium with Fe(III) (hydr)oxides (Table 2).
We evaluated the arsenic removal rate by biomass of Talaromyces sp. KM-31 coated with Fe(III) (hydr)oxides. X-ray diffraction analysis showed that the two modes of coating used in the present work formed different Fe(III) oxide species (Figure 4). From diffraction peaks, the primary particle size of Fe(III) hydroxide and Fe(III) oxide coated on fungi was determined to be approximately 24 and 17 nm, respectively, which confirmed the formation of nano-sized Fe(III) (hydr)oxide on fungi. Arsenic removal data revealed that modification of the biomass enhanced removal of both arsenic species relative to uncoated biomass (Table 2). In the case of As(III), live biomass of Talaromyces sp. KM-31 coated with Fe(III) hydroxide removed 69.0% of the arsenic, while biomass coated with nano-sized Fe(III) oxide removed 91.0%, whereas untreated mycelium removed only 4.0%. In the case of As(V), the removal rates were higher than those for As(III), which is consistent with the growth and tolerance results. The untreated biomass removed 13.0% of the As(V), while the removal rates for treated biomass were 79.0% with the Fe(III) hydroxide coating and 96.0% with the Fe(III) oxide coating. Thus surface modification resulted in better affinity, and hence more effective removal of arsenic. This is arisen from an intrinsic property of Fe(III) oxide species favorable for arsenic adsorption (see Table S2). In past studies, the addition of coatings has been shown to be effective in arsenic removal. For example, sand coated with iron oxide showed greater arsenic removal than sand alone [50], orange juice industrial residue loaded with Fe(III) [51], and Penicillium purpurogenum [52]. In another study, chemical modification of Penicillium chrysogenum biomass increased arsenate removal [53]. Similarly, in the present study, we found that Fe(III) (hydr)oxide modification of Talaromyces sp. KM-31 greatly enhanced its arsenic removal capacity. Fungal cells are inherently fragile, limiting freely-suspended biomass as a good sorbent for metal(loid) removal from aqueous solution. This fragility problem can be reduced by immobilization of fungal biomass in a suitable porous matrix [12]. Use of Fe(III) (hydr)oxides as an immobilization matrix has advantages, including large surface area, absence of toxicity, and ease of handling.

4. Conclusions

Talaromyces sp. KM-31 isolated from highly As-contaminated soil from a mine site showed hyper-tolerance to arsenic, and was able to grow in the presence of arsenic concentrations up to 300 mg/L in a complex organic medium. Although the isolate tolerated arsenic and grew well in its presence, its arsenic removal ability was poor at high arsenic concentrations. At low arsenic concentrations, live mycelium was more effective at removing As(V) than inactivated mycelium. Powder prepared from mycelium of Talaromyces sp. KM-31 that had been coated with Fe(III) (hydr)oxide showed several-fold enhanced removal of both As(III) and As(V), suggesting that the modified biomass could be used for water purification. The present findings indicate that this Fe(III)-modified fungal strain has promise for use as an efficient biosorbent in bioremediation-based approaches to the removal of arsenic from aqueous systems.

Supplementary Materials

The following are available online at https://www.mdpi.com/2075-163X/9/10/568/s1: Table S1: Growth of Talaromyces sp. KM-31 in PDA medium amended with As(III) or As(V).; Table S2: Arsenic removal (%) by Fe(III) oxides as a non-fungal reference.

Author Contributions

I.H.N. and J.H.K. designed the study; I.H.N., K.M., and J.R. performed the experiment and analyzed the data; J.R. and J.H.K. provided critical feedback and helped shape the research; I.H.N. wrote the final version of the manuscript.

Funding

This work was supported by an Institute of Information & Communications Technology Planning & Evaluation(IITP) grant funded by the Korea government (MSIP) (Grant No. B0132-15-1001, Development of Next Imaging System) and the Korea Institute of Geoscience and Mineral Resources (KIGAM) Basic Research Project (No. 19-3426).

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Donahoe-Christiansen, J.; D’Imperio, S.; Jackson, C.R.; Inskeep, W.P.; McDermott, T.R. Arsenite-oxidizing Hydrogenobaculum strain isolated from an acid-sulfate-chloride geothermal spring in Yellowstone National Park. Appl. Environ. Microbiol. 2004, 70, 1865–1868. [Google Scholar] [CrossRef] [PubMed]
  2. Wang, S.; Mulligan, C.N. Occurrence of arsenic contamination in Canada: Sources, behavior and distribution. Sci. Total Environ. 2006, 366, 701–721. [Google Scholar] [CrossRef] [PubMed]
  3. Turpeinen, R.; Kallio, M.P.; Kairesalo, T. Role of microbes in controlling the speciation of arsenic and production of arsines in contaminated soils. Sci. Total Environ. 2002, 285, 133–145. [Google Scholar] [CrossRef]
  4. Tamaki, S.; Frankenberger, W.T. Environmental biochemistry of arsenic. Rev. Environ. Contam. Toxicol. 1992, 124, 79–110. [Google Scholar] [PubMed]
  5. Moreno-Jiménez, E.; Manzano, R.; Esteban, E.; Peñalosa, J. The fate of arsenic in soils adjacent to an old mine site (Bustarviejo, Spain): Mobility and transfer to native flora. J. Soil Sediment 2010, 10, 301–312. [Google Scholar] [CrossRef]
  6. Mulligan, C.N.; Yong, R.N.; Gibbs, B.F. Remediation technologies for metal-contaminated soils and groundwater: An evaluation. Eng. Geol. 2001, 60, 193–207. [Google Scholar] [CrossRef]
  7. Wang, S.L.; Zhao, X.Y. On the potential of biological treatment for arsenic contaminated soils and groundwater. J. Environ. Manag. 2009, 90, 2367–2376. [Google Scholar] [CrossRef]
  8. Gadd, G.M. Bioremedial potential of microbial mechanisms of metal mobilization and immobilization. Curr. Opin. Biotechnol. 2000, 11, 271–279. [Google Scholar] [CrossRef]
  9. Singh, A.; Prasad, S.M. Remediation of heavy metal contaminated ecosystem: An overview on technology advancement. Int. J. Environ. Sci. Technol. 2015, 12, 353–366. [Google Scholar] [CrossRef]
  10. Stolz, J.F.; Oremland, R.S. Bacterial respiration of arsenic and selenium. FEMS Microbiol. Rev. 1999, 23, 615–627. [Google Scholar] [CrossRef]
  11. Brierley, C.L. Bioremediation of metal-contaminated surface and groundwaters. Geomicrobiol. J. 1990, 8, 201–223. [Google Scholar] [CrossRef]
  12. Jaiswal, V.; Saxena, S.; Kaur, I.; Dubey, P.; Nand, S.; Naseem, M.; Singh, S.B.; Srivastava, P.K.; Barik, S.K. Application of four novel fungal strains to remove arsenic from contaminated water in batch and column modes. J. Hazard. Mater. 2018, 356, 98–107. [Google Scholar] [CrossRef] [PubMed]
  13. Pumpel, T.; Schinner, F. Native fungal pellets as biosorbent for heavy metals. FEMS Microbiol. Rev. 1993, 11, 159–164. [Google Scholar] [CrossRef]
  14. Volesky, B.; Holan, Z.R. Biosorption of heavy metals. Biotechnol. Prog. 1995, 11, 235–250. [Google Scholar] [CrossRef] [PubMed]
  15. Cernansky, S.; Urik, M.; Sevc, J.; Khun, K. Biosorption and biovolatilization of arsenic by heat-resistant fungi. Environ. Sci. Pollut. Res. 2007, 14, 31–35. [Google Scholar] [CrossRef]
  16. Pearce, R.B.; Callow, M.E.; Macaskie, L.E. Fungal volatilization of arsenic and antimony and the sudden infant death syndrome. FEMS Microbiol. Lett. 1998, 158, 261–265. [Google Scholar] [CrossRef] [PubMed]
  17. Granchinho, S.C.R.; Franz, C.M.; Polishchuk, E.; Cullen, W.R.; Reimer, K.J. Transformation of arsenic(V) by the fungus Fusarium oxysporum melonis isolated from the alga Fucus gardneri. Appl. Organomet. Chem. 2002, 16, 721–726. [Google Scholar] [CrossRef]
  18. Carrasco, J.A.; Armario, P.; Pajuelo, E.; Burgos, A.; Caviedes, M.A.; Lόpez, R.; Chamber, M.A.; Palomares, A.J. Isolation and characterization of symbiotically effective Rhizobium resistant to arsenic and heavy metals after the toxic spill at the Aznalcóllar pyrite mine. Soil Biol. Biochem. 2005, 37, 1131–1140. [Google Scholar] [CrossRef]
  19. Vala, A.K. Tolerance and removal of arsenic by a facultative marine fungus Aspergillus candidus. Bioresour. Technol. 2010, 101, 2565–2567. [Google Scholar] [CrossRef]
  20. Srivastava, P.K.; Vaish, A.; Dwivedi, S.; Chakrabarty, D.; Singh, N.; Tripathi, R.D. Biological removal of arsenic pollution by soil fungi. Sci. Total Environ. 2011, 409, 2430–2442. [Google Scholar] [CrossRef]
  21. Rodríguez, I.A.; Martínez-Juárez, V.M.; Cárdenas-González, J.F.; Moctezuma-Zárate, M.G. Biosorption of arsenic(III) from aqueous solutions by modified fungal biomass of Paecilomyces sp. Bioinorg. Chem. Appl. 2013, 376780, 1–5. [Google Scholar] [CrossRef] [PubMed]
  22. Murad, F.N.; Kabbashi, N.A.; Alam, M.Z.; Al-Khatib, M.F.R. Immobilization of fungal biomass with multi-walled carbon nanotubes as biosorbent. Malays. J. Microbiol. 2018, 14, 113–119. [Google Scholar]
  23. Afkhami, A.; Saber-Tehrani, M.; Bagheri, H. Modified maghemite nanoparticles as an efficient adsorbent for removing some cationic dyes from aqueous solution. Desalination 2010, 263, 240–248. [Google Scholar] [CrossRef]
  24. Xu, P.; Zeng, G.M.; Huang, D.L.; Feng, C.L.; Hu, S.; Zhao, M.H.; Lai, C.; Wei, Z.; Huang, C.; Xie, G.X.; et al. Use of iron oxide nanomaterials in wastewater treatment: A review. Sci. Total Environ. 2012, 424, 1–10. [Google Scholar] [CrossRef] [PubMed]
  25. Gupta, A.K.; Gupta, M. Synthesis and surface engineering of iron oxide nanoparticles for biomedical applications. Biomaterials 2005, 26, 3995–4021. [Google Scholar] [CrossRef]
  26. Machala, J.; Zboril, R.; Gedanken, A. Amorphous iron(III) oxides: A review. J. Phys. Chem. B 2007, 111, 4003–4018. [Google Scholar] [CrossRef] [PubMed]
  27. Langmuir, I. The adsorption of gases on plane surfaces of glass, mica and platinum. J. Am. Chem. Soc. 1918, 40, 1361–1403. [Google Scholar] [CrossRef]
  28. U.S. Environmental Protection Agency. Method 3051A: Microwave Assisted Acid Digestion of Sediments, Sludges, Soils, and Oils; U.S. Gov. Print Office: Washington, DC, USA, 2007.
  29. Gadd, G.M. Interactions of fungi with toxic metals. New Phytol. 1993, 124, 25–60. [Google Scholar] [CrossRef]
  30. Babich, H.; Stotzky, G. Heavy metal toxicity to microbe-mediated ecologic processes: A review and potential application to regulatory policies. Environ. Res. 1985, 36, 111–137. [Google Scholar] [CrossRef]
  31. Visagie, C.M.; Jacobs, K. Three new additions to the genus Talaromyces isolated from Atlantis sandveld fynbos soils. Persoonia 2012, 28, 14–24. [Google Scholar] [CrossRef]
  32. Hassan, N.; Rafiq, M.; Rehman, M.; Sajjad, W.; Hasan, F.; Abdullah, S. Fungi in acidic fire: A potential source of industrially important enzymes. Fungal Biol. Rev. 2019, 33, 58–71. [Google Scholar] [CrossRef]
  33. Romero, M.C.; Reinoso, E.H.; Urrutia, M.I.; Kiernan, A.M. Biosorption of heavy metals by Talaromyces helicus: A trained fungus for copper and biphenyl detoxification. Elect. J. Biotechnol. 2006, 9, 1–11. [Google Scholar] [CrossRef]
  34. Bengtsson, L.; Johansson, B.; Hackett, T.J.; McHale, L.; Mchale, A.P. Studies on the biosorption of uranium by Talaromyces emersonii CBS 814.70 biomass. Appl. Microbiol. Biotechnol. 1995, 42, 807–811. [Google Scholar] [CrossRef] [PubMed]
  35. Tebo, B.M.; Obraztsova, A.Y. Sulfate-reducing bacterium grows with Cr(VI), U(VI), Mn(IV) and Fe(III) as electron acceptors. FEMS Microbiol. Lett. 1998, 162, 193–198. [Google Scholar] [CrossRef]
  36. Santini, J.M.; Sly, L.I.; Schnagl, R.D.; Macy, J.M. A new chemolithoautotrophic arsenite-oxidizing bacterium isolated from a gold mine: Phylogenetic, physiological, and preliminary biochemical studies. Appl. Environ. Microbiol. 2000, 66, 92–97. [Google Scholar] [CrossRef] [PubMed]
  37. Niggemyer, A.; Spring, S.; Stackebrandt, E.; Rosenzweig, R.F. Isolation and characterization of a novel As(V)-reducing bacterium: Implications for arsenic mobilization and the genus Desulfitobacterium. Appl. Environ. Microbiol. 2001, 67, 5568–5580. [Google Scholar] [CrossRef] [PubMed]
  38. Zafar, S.; Aqil, F.; Ahmad, I. Metal tolerance and biosorption potential of filamentous fungi isolated from metal contaminated agricultural soil. Bioresour. Technol. 2007, 98, 2557–2561. [Google Scholar] [CrossRef] [PubMed]
  39. Canovas, D.; de Lorenzo, V. Osmotic stress limits arsenic hyper-tolerance in Aspergillus sp. P37. FEMS Microbiol. Ecol. 2007, 61, 258–263. [Google Scholar] [CrossRef]
  40. Sakurai, I. Biomethylation of arsenic is essentially detoxicating event. J. Health Sci. 2003, 49, 171–178. [Google Scholar] [CrossRef]
  41. Urík, M.; Čerňanský, S.; Ševc, J.; Šimonovičová, A.; Littera, P. Biovolatilization of arsenic by different fungal strains. Water Air Soil Pollut. 2007, 186, 337–342. [Google Scholar] [CrossRef]
  42. Gadd, G.M.; White, C. Microbial treatment of metal pollution—A working biotechnology? Trends Biotechnol. 1993, 11, 353–359. [Google Scholar] [CrossRef]
  43. Churchill, S.A.; Walters, J.V.; Churchill, P.F. Sorption of heavy metals by prepared bacterial cell surfaces. J. Environ. Eng. 1995, 121, 706–711. [Google Scholar] [CrossRef]
  44. Zhou, J.L. Zn biosorption by Rhizopus arrhizus and other fungi. Appl. Microbiol. Biotechnol. 1999, 51, 686–693. [Google Scholar] [CrossRef]
  45. Bai, R.S.; Abraham, T.E. Studies on chromium(VI) adsorption-desorption using immobilized fungal biomass. Bioresour. Technol. 2003, 87, 17–26. [Google Scholar] [PubMed]
  46. Yan, G.; Viraraghavan, T. Heavy-metal removal from aqueous solution by fungus Mucor rouxii. Water Res. 2003, 37, 4486–4496. [Google Scholar] [CrossRef]
  47. Gadd, G.M. Microbial influence on metal mobility and application for bioremediation. Geoderma 2004, 122, 109–119. [Google Scholar] [CrossRef]
  48. Pillichshammer, M.; Pumpel, T.; Poder, R.; Eller, K.; Klima, J.; Schinner, F. Biosorption of chromium to fungi. Biometals 1995, 8, 117–121. [Google Scholar] [CrossRef]
  49. Fogarty, R.V.; Tobin, J.M. Fungal melanins and their interactions with metals. Enzyme Microbiol. Technol. 1996, 19, 311–317. [Google Scholar] [CrossRef]
  50. Thirunavukkarasu, O.S.; Viraraghavan, T.; Subramanian, K.S. Arsenic removal from drinking water using iron oxide-coated sand. Water Air Soil Pollut. 2003, 142, 95–111. [Google Scholar] [CrossRef]
  51. Ghimire, K.N.; Inoue, K.; Makino, K.; Miyajima, T. Adsorptive removal of arsenic using orange juice residue. Sep. Sci. Technol. 2002, 37, 2785–2799. [Google Scholar] [CrossRef]
  52. Say, R.; Yilmaz, N.; Denizli, A. Biosorption of cadmium, lead, mercury, and arsenic ions by the fungus Penicillium purpurogenum. Sep. Sci. Technol. 2003, 38, 2039–2053. [Google Scholar] [CrossRef]
  53. Loukidou, M.X.; Matis, K.A.; Zouboulis, A.I.; Liakopoulou-Kyriakidou, M. Removal of As(V) from wastewaters by chemically modified fungal biomass. Water Res. 2003, 37, 4544–4552. [Google Scholar] [CrossRef]
Minerals 09 00568 g001 550
Figure 1. Changes in biomass of Talaromyces sp. KM-31 after 15 days of cultivation on PDA medium augmented with arsenic. The As(III) and As(V) concentrations ranged from 0 to 1000 mg/L.
Figure 1. Changes in biomass of Talaromyces sp. KM-31 after 15 days of cultivation on PDA medium augmented with arsenic. The As(III) and As(V) concentrations ranged from 0 to 1000 mg/L.
Minerals 09 00568 g001
Minerals 09 00568 g002 550
Figure 2. Percent change in live mycelial biomass of Talaromyces sp. KM-31 at various initial arsenic concentrations (1.0~25.0 mg/L) in an arsenic removal assay. The experimental conditions were biomass 60 g/L, contact time 72 h, pH 6.0, temperature 30 °C, and shaking at 160 rpm.
Figure 2. Percent change in live mycelial biomass of Talaromyces sp. KM-31 at various initial arsenic concentrations (1.0~25.0 mg/L) in an arsenic removal assay. The experimental conditions were biomass 60 g/L, contact time 72 h, pH 6.0, temperature 30 °C, and shaking at 160 rpm.
Minerals 09 00568 g002
Minerals 09 00568 g003 550
Figure 3. Isotherm for the removal of arsenic. The Langmuir isotherm is shown. The experimental conditions were biomass 60 g/L, contact time 72 h, pH 6.0, temperature 30 °C, and shaking at 160 rpm.
Figure 3. Isotherm for the removal of arsenic. The Langmuir isotherm is shown. The experimental conditions were biomass 60 g/L, contact time 72 h, pH 6.0, temperature 30 °C, and shaking at 160 rpm.
Minerals 09 00568 g003
Minerals 09 00568 g004 550
Figure 4. X-ray diffractograms and images (insets) for untreated Talaromyces sp. KM-31 and Fe(III) hydr(oxide)-coated biomass. (A) untreated biomass; (B) Fe(III) hydroxide-coated biomass; and (C) nFe(III) oxide-coated biomass.
Figure 4. X-ray diffractograms and images (insets) for untreated Talaromyces sp. KM-31 and Fe(III) hydr(oxide)-coated biomass. (A) untreated biomass; (B) Fe(III) hydroxide-coated biomass; and (C) nFe(III) oxide-coated biomass.
Minerals 09 00568 g004
Table
Table 1. Arsenic tolerance index for Talaromyces sp. KM-31 at varying concentrations of As(III) or As(V) in PDB and PDA media after 15 days of cultivation, and arsenic removal rates in PDB medium.
Table 1. Arsenic tolerance index for Talaromyces sp. KM-31 at varying concentrations of As(III) or As(V) in PDB and PDA media after 15 days of cultivation, and arsenic removal rates in PDB medium.
Arsenic Tolerance Index in Solid and Liquid MediaArsenic Removal (%)
As (mg/L)PDAPDBPDB
As(III)As(V)As(III)As(V)As(III)As(V)
01.01.01.01.000
500.980.960.980.985.2426.2
1000.960.970.980.984.4117.25
2000.960.960.970.972.87.53
3000.980.970.130.960.411.24
5000.960.970.0080.6700.78
8000.960.970.0080.500.52
10000.960.980.0040.3500.32
Table
Table 2. Arsenic removal (%) by untreated biomass, Fe(III) hydroxide-coated biomass, and Fe(III) oxide-coated biomass.
Table 2. Arsenic removal (%) by untreated biomass, Fe(III) hydroxide-coated biomass, and Fe(III) oxide-coated biomass.
Treatment aArsenic Removal (mean ± SD b, %)
As(III)As(V)
Untreated biomass4 ± 0.3213 ± 0.97
Fe(III) hydroxide-coated biomass69 ± 2.7979 ± 3.65
nFe(III) oxide-coated biomass91 ± 5.2596 ± 4.18
a Sorbent, 1 g/L; sorbate, 5 mg/L; pH 6.0; temp, 30 °C; contact time, 24 h. b The mean and standard deviation of three replicate determinations.

Share and Cite

MDPI and ACS Style

Nam, I.-H.; Murugesan, K.; Ryu, J.; Kim, J.H. Arsenic (As) Removal Using Talaromyces sp. KM-31 Isolated from As-Contaminated Mine Soil. Minerals 2019, 9, 568. https://doi.org/10.3390/min9100568

AMA Style

Nam I-H, Murugesan K, Ryu J, Kim JH. Arsenic (As) Removal Using Talaromyces sp. KM-31 Isolated from As-Contaminated Mine Soil. Minerals. 2019; 9(10):568. https://doi.org/10.3390/min9100568

Chicago/Turabian Style

Nam, In-Hyun, Kumarasamy Murugesan, Jungho Ryu, and Jae Hwan Kim. 2019. "Arsenic (As) Removal Using Talaromyces sp. KM-31 Isolated from As-Contaminated Mine Soil" Minerals 9, no. 10: 568. https://doi.org/10.3390/min9100568

APA Style

Nam, I. -H., Murugesan, K., Ryu, J., & Kim, J. H. (2019). Arsenic (As) Removal Using Talaromyces sp. KM-31 Isolated from As-Contaminated Mine Soil. Minerals, 9(10), 568. https://doi.org/10.3390/min9100568

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