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Chromatographia

, Volume 82, Issue 1, pp 125–141 | Cite as

How to Deal with Mercury in Sediments? A Critical Review About Used Methods for the Speciation of Mercury in Sediments

  • C. Hellmann
  • R. D. Costa
  • O. J. SchmitzEmail author
Review
  • 477 Downloads
Part of the following topical collections:
  1. 50th Anniversary Commemorative Issue

Abstract

Sediments serve as an indicator of the state of the environment, as they reveal, anthropogenic influences (e.g. industry) over time. The knowledge about the composition of sediments, in particular by the speciation, helps in the assessment of the environmental situation. The speciation of mercury in sediments is still being discussed and continues to pose a great challenge for analytical chemists. Despite a broad number of publications in this area, there is no gold-standard about the speciation of mercury in sediments. The reason for this is the growing interest in new, better methods for the speciation of mercury, which increases the number of publications and the uncertainty among the analysts. Therefore, the methodology of mercury speciation in sediments requires improvement and would benefit from a standardized approach. The goal of this review is to give an overview of the existing methods and to discuss the issues of methodology. Discussed parts in this review article include: (1) available reference material, (2) the methodology of extraction, (3) enrichment procedures, (4) separation and (5) detection.

Graphical Abstract

Keywords

Mercury speciation Sediments Extraction Separation Pre-concentration Detection 

Introduction

Since decades, mercury has been an ongoing topic of discussion because of the environmental situation and has been of particular interest since the occurrence of Minamata in 1956 [1]. Since then the World Health Organization (WHO) categorized mercury as one of ten substances that threatens public health [2]. The occurrence of mercury can basically be classified into three categories by distinguishing between elemental mercury Hg(0), inorganic mercury Hg(I), Hg(II) and various forms of organic mercury CH3Hg+, (CH3)2Hg, EtHg+, PhHg+ [3, 4].

Elemental mercury is one of the elements which has a liquid aggregate state at room temperature. The main intake path for this species works via the lung because Hg(0) evaporates rapidly due to the considerable vapor pressure at room temperature. After resorption, it can pass the blood–brain barrier, as well as the placenta and then spread throughout the entire organism [5, 6].

Inorganic mercury compounds may be present in the oxidation states (I) and (II), whereby Hg(I), which is frequently present as calomel (Hg2Cl2), is poorly water-soluble, thereby being classified as safe [6]. While the intake through the lungs and the digestive tract is variable, the dermal intake is irrelevant [7]. Friberg et al. demonstrated that only 8% of dermally applied mercury chloride could be absorbed in 5 h [8].

Since there is already a very high risk potential in small quantities, organic mercury is of particular interest. While ethyl mercury is very rapidly degraded to Hg2+ [9], CH3Hg+ and (CH3)2Hg have special properties, which makes them among the greatest environmental threats. (CH3)2Hg has a particular influence on the distribution of organic mercury in the environment which can be explained by its volatility, water insolubility and non-existent affinity for thiol groups (–SH) [6]. In contrast, the CH3Hg+ shows a high affinity for thiol groups. Since these are frequent components in proteins, the CH3Hg+ is found throughout the body binding sites. For example, it is passed through the cell membrane by the amino acid l-cysteine [10, 11]. The resorption pathways of CH3Hg+ are manifold, which means that they can be absorbed by the intact skin in addition to oral and inhaled uptake [12].

Through anthropogenic (industrial processes, combustion processes, gold production, etc.) or natural (volcano, etc.) intake, mercury enters the environment and results in a biogeochemical cycle, which describes the distribution of the mercury species in the respective areas of the environment (Fig.1).

While Hg(0) is predominantly found in the atmosphere, the dominant species in water is the inorganic Hg2+. This can be explained by the high concentration of salts (such as chlorides) which supports the oxidation of Hg(0) to Hg2+, as well as the presence of sulfides which interfere with methylation [13, 14]. According to the hard and soft acids and bases principle (HSAB), mercury is preferred in this respect for the formation of complexes with sulfur-containing ligands or organic material [15]. Due to the fact that sediments are a natural source of these components, they are capable of binding heavy metals. A number of species can be found in soils and sediments. However, in contrast to other compartments, a different distribution of the species is present. For the conversion processes in these matrices, mainly the presence of organic material as well as the temperature is responsible. While in fish mainly CH3Hg+ is found [16, 17, 18, 19], this is only about 0.1–1% of the total Hg [20] in soils and sediments. On the contrary, there is a relatively high content of inorganic mercury, which especially leads to problems in the analysis of methyl mercury in sediments. Table 1 illustrates different mercury concentrations worldwide.

Fig. 1

Biogeochemical cycle of mercury

(adapted from [21])

Table 1

Concentration levels of mercury species in sediments worldwide

Country

Total Hg

Methyl mercury

References

Tasmania, Australia

0.004–0.194 mg kg−1

0.02–20.1 µg kg−1

[22]

Negro river, Brazil

70–271 mg kg−1

0.47–1.79%

[23]

Amazon, Brazil

0.069–0.109 mg kg−1

0.62–4.78 µg kg−1

[24]

Ontario, Canada

0.03–0.20 mg kg−1

0.17–2.86 µg kg−1

[25]

South Florida, USA

0.02 mg kg−1

0.08 µg kg−1

[26]

Seine, France

0.3–1 mg kg−1

2%

[27]

Baltic, Poland

0.001–0.2 mg kg−1

< 1%

[28]

Elbe river, Germany

0.5–27 mg kg−1

50–27 µg kg−1

[29]

By altering conditions such as pH or salt concentration, which affect solubility, mobility and bioavailability, the species can be separated from the sediment again [30]. As a result, sediments are a permanent source of mercury in surface water [31]. This leads to the interest in sediments which has increased dramatically in recent years. Figure 2 shows the matrices studied in the last decades [32].

Fig. 2

Matrices of interest in the last few years

(adapted from [32])

Because of the toxic differences and the occurrence of the species in the individual areas, speciation of mercury becomes indispensable, when a state of environment needs to be evaluated [33].

A species is generally understood as the specific form of an element defined as an isotopic composition, electronic or oxidation state, and/or complex or molecular structure [34]. The analysis can be divided into several sub-sections, which are shown in Fig. 3. For the analysis of mercury in sediments, all interesting mercury species have to be extracted with a suitable extraction method, enriched to determine also low concentrated species, chromatographic or electrophoretic separated and detected with an element-specific detector. There are already a number of methods for the speciation of mercury in sediments found in the literature. In the following, the individual sections of a speciation will be discussed and critically evaluated. This should help other analysts when developing a methodology.

Fig. 3

Flow of an analysis, including the common used techniques

(adapted from [21])

Handling of Sediment Samples (Storage and Pretreatment)

For the speciation of mercury, a special procedure with regard to sampling, storage and pretreatment of the samples becomes necessary. The samples are usually stored in containers made of polymer [35, 36], glass [37], PTFE or stainless steel [38]. Yu et al. [39] recommends to use PTFE, FLPE, PET, Pyrex glass and quartz to store samples. They are also pointing out that mercury loss has to be feared using PE containers [40, 41, 42, 43], since they promote species degradation. As a result PTFE containers should be preferred, which reduce adsorption on the surface and enhance the stability of the species [44]. Despite the arguments against the use of PE containers for the speciation of mercury, acid-cleaned PE vessels has become established for the speciation in sediments [45, 46, 47, 48], since it has an stabilizing effect on the mercury species [39, 44, 49]. Basically, it requires a thorough cleaning of all components that come in contact with the sample. Cleaning procedures are performed, including aqua regia or nitric acid. In special cases, working in special clean rooms may be required. To avoid any contamination, the sample should be analyzed as early as possible after the collection or stored in a dark place at low temperatures (refrigerator) [3]. In the literature it is also reported that a pretreatment of the sample with acids can minimize the risk of methylation [50]. Common used acids are HNO3, H2SO4 or HCl, whereby studies show that nitric acid is the most effective additive [39]. Furthermore, it is advisable to cover freshly collected sediment samples with a water layer for transportation [48]. After reviewing the literature, it is evident that little is said about the treatment after sampling and before the analysis (e.g. sieving, drying, homogenization). Since these steps are also of particular importance with regard to the production of reference material, they should be given more attention. Due to the mentioned influencing factors, great care must be taken in choosing the right storage and pretreatment procedure.

Quality Control with Suitable Reference Material

Legislation within the European Union provides that methylmercury has a proven quality in matrices such as food or environmental samples. To determine these, mainly suitable Certified Reference Material (CRM) is used. However, reviewing mercury speciation methods in sediments still poses a major challenge to analytics, which in turn is due to the difficult analytical conditions, the different species concentrations, the comparability of methods, the available reference material, or artifact formation [51]. While a number of reference materials exist for marine samples such as fish or mussels, there is a lack of material especially for sediments. Some reference materials are used in the literature, which differ in their levels of mercury species (Table 2). While ERM CC-580 has very high levels of mercury, the concentrations in IAEA-456 are very low. Depending on the analytical conditions, the appropriate material can be selected. It should be noted, however, that in the case of using a higher concentrated reference material such as ERM CC-580, effects such as artifact formation may be observed, which do not occur with low concentrated real samples. In this context, the choice must be made very thoroughly. Due to possible species transformations also reference materials containing other mercury species are required. There are several methods in the literature using ERM CC-580 as reference material and numerous publications dealing with the origin and characterization of it [52, 53]. Fabbri et al. [52], who has been intensively involved in the characterization of this reference material, mentions that the sediment used for the preparation of the reference material has a high sulfur content. Accordingly, they conclude that the released mercury forms a complex with the available sulfur. HgS is non-mobile and thus remains inactive in the sediment. This suggests that the reference material CRM 580 used contains a low content of extractable mercury [52]. This fact should be taken into account when using different extraction procedures. Since sediments differ from one another to some extent, the question here is whether the analysis of one reference material is sufficient for the statement about the quality of the measurement method. In this context, many working groups use sediment, which is artificially mixed with mercury standards. However, this approach leads to further uncertainty about the quality of the developed extraction method, since mercury-added sediment behaves differently when extracted than naturally occurring sediment [54]. Under these circumstances, these methods should be chosen wisely.

Table 2

Used reference material for the speciation of mercury in sediments

Matrix

Code

Supplier

Certified value mg kg−1 as Hg (dry mass)

Total Hg

CH3Hg+

Estuarine sediment

ERM-CC580

IRMM

132 ± 3

0.0755 ± 0.0037

Marine sediment

IAEA-433

IAEA

0.168 ± 0.017

0.17 ± 0.07

Marine sediment

IAEA-158

IAEA

0.132 ± 0.014

0.00141 ± 0.0004

Coastal sediment

IAEA-456

IAEA

0.077 ± 0.005

0.000125 ± 0.000019

Estuarine sediment

IAEA-405

IAEA

0.81 ± 0.040

0.00549 ± 0.0006

IAEA International Atomic Energy Agency, Vienna (Austria), ERM European Reference Material [Institute for Reference Materials and Measurements (IRMM), Geel (Belgium)]

Extraction

The extraction is the first essential step for the investigation of mercury species. This differs according to the matrix and, above all, poses a great challenge in the investigation of sediments. This is due to the fact that the concentration of methylmercury in sediments is less than 1% of the total mercury content [55]. The challenge is to find an adequate extraction which enables high recoveries without losses or conversion. If sediments are investigated, consideration must be given to the different characteristics of the respective species (e.g. solubility), since this has an influence on the selected extraction method. Figure 4 gives an overview on the different species and their properties. In addition, the mobility, bioavailability and toxicity of mercury are determined by biogeochemical fractionation. For this reason, knowledge of the individual fractions is of particular importance [56]. The behavior of mercury in sediments and the resulting fractions are influenced by factors such as total mercury, sulfur and iron content, pH, and organic matter [57, 58, 59]. Organic material occurring in sediments leads to the formation of complexes due to an interaction between its functional groups (e.g. carboxylic acids, phenols, thiols, alcohols) and Hg [56]. Because of its chemical properties, mercury preferably forms strong ionic bonds with reduced sulfur groups found in dissolved organic carbon (DOC), including humic and fulvic acids [60, 61, 62]. As a result, poorly soluble mercury sulfides are formed [63]. Adsorption effects can be observed in the presence of iron [64]. In addition to these effects, the influence of the pH is often described. According to Manohar et al. [65] low pH values (< 4) lead to a desorption of mercury. The reason for this is the increased positive charge of the organic material. However, it has already been shown in the literature that the influence of this parameter is rather low in case of mercury [57] and that the pH values are often above 4, whereby no desorption is to be expected [56].

Fig. 4

Fractionation of mercury species

(adapted from [66])

To distinguish between these different fractions, a Hg-specific sequential extraction can be used. While in sequential extraction the main aim is to extract the specific species successively, the mobility of all species of an element should be determined within the sample [15]. Bloom et al. [67] who developed a Hg-specific sequential extraction for mercury, presents a method by which mercury compounds can be classified into different classes using different reagents in various steps. The process leads to a distinction between (a) water soluble, (b) human stomach acid soluble, (c) organo-chelated, (d) elemental mercury, (e) mercuric sulfide. Although the use of sequential extraction is frequently used in the speciation of mercury, the general procedure already suggests some disadvantages. Therefore, it is very time-consuming [68] and often characterized by low reproducibility [68, 69, 70]. Furthermore, it is not entirely accurate since a change of the sample during this chemical treatment has to be feared [71, 72]. The literature also criticizes the non-specific removal of species during this procedure [71, 72, 73, 74, 75, 76]. Because of the arguments mentioned, single reagent extraction is preferred over sequential ones [15].

The literature includes a large number of different extraction procedures, which can be categorized in acid leaching [27, 77, 78, 79, 80, 81, 82, 83, 84, 85, 86], alkaline extraction [55, 87, 88, 89] and distillation [55, 79, 90, 91]. When selecting the most suitable extraction method, the generation of artifacts must be considered. Artifacts are the unintentional transformation of one species into another, which can be caused by the sampling, extraction or separation method [92].

Although the use of distillation leads to good recoveries for CH3Hg+ in sediments [91] and reduces matrix effects [93], more artifacts are formed compared to other extraction methods when analyzing samples with an Hg2+ content of ≥ 2 µg g−1 [55, 90]. If the content of the inorganic mercury is below the stated limit, the distillation is still a useful method [90]. Due to the occurrence of a low artifact formation and good recovery [79] the acid extraction is used very frequently and results in many modifications. In addition to microwave-assisted [54, 80, 81] and ultrasound-assisted extraction [77] which are used to increase the speed and efficiency of the extraction [54, 94] the use of organic solvents is also often described [66, 79, 82, 87, 90, 95]. Table 3 shows an excerpt of the extraction methods available in the literature.

Table 3

Extraction of sediments using various leaching procedures

Reagents

Sample

Sample amount

Technique

Recovery for CH3Hg+

References

Acidic extraction

 4 M HNO3

CRM BCR 580a

1.0 g

HPLC-ICP-MS

97.3 ± 2.7%

[80]

 2 M HNO3

Sediment

1.0 g

Et-CT-GC-QFAASb

Mean 97%

[81]

 50% H2SO4, BrCl,10% NaCl

Estuarine sediment

0.2–0.5 g

CV-AFS

80–90%g

[27]

 Conc. CH2O2

Sediment

NA

Et-GC-CVAFSc

Less than 10%

[79]

 10% HCl

Sediment

NA

Et-GC-CVAFSc

61.6%

[79]

 9 M H2SO4; 0.2 mL of 20% KCl

Sediment

0.5–1.0 g

Et-GC-CVAFSc

90.4 ± 11.4%

[79]

 25% CuSO4 in 6 M HCl

Sediment

0.2–0.5 g

GC-AFS

95.5

[84]

 Conc.HCl (37%)

CRM BCR 580a

0.3 g

HPLC-CVAFS

103.7 ± 7.6%

[77]

Acidic extraction using organic solvents

 18% KBr,5% H2SO4, 1 M CuSO4, CH2Cl2

Estuarine sediment

0.5 g

GC-CVAFS

88.1 ± 11.9%

[79]

 5% H2SO4, 1 M CuSO4,CH2Cl2, 18% KBr

Sediment

0.5 g

Et-GC-CVAFSc

98.2 ± 15.3%

[79]

 HCl, toluene

CRM BCR 580a

2.0 g

CGC-ECDd

88 ± 5%

[86]

 0.03 M CuSO4, 0.38 M KBr in 5% H2SO4, toluene

CRM BCR 580a

2–3 g

GC-MIPAESe

98 ± 6%

[82]

 2% HCl + 10% ethanol

CRM BCR 580a

1.0–2.0 g

HPLC-ICP-MS

98.1 ± 8.4%

[66]

 Conc. HCl (37%), CH2Cl2

CRM BCR 580a

0.050 g

HPLC-CVAFS

87.5 ± 6.3%

[77]

Alkaline extraction

 25% KOH/MeOH

Sediment

1.0 g

Et-GC-CVAFSc

118.9 ± 8.6%

[79]

 25% KOH/MeOH, CH2Cl2

Sediment

NA

Et-GC-CVAFSc

101.5 ± 3.0%

[79]

 25% KOH/MeOH, CH2Cl2, conc.HCl

CRM BCR 580a

0.25 g

HPLC-UV-CV-AFS

93.6 ± 1.8%

[96]

Extraction with sulfur containing additives

 HCl, toluene, cysteine acetate

CRM BCR 580a

0.2 g

CGC-ECD

88 ± 5%

[53]

 0.5% 2-mercaptoethanol, 5% methanol

CRM BCR 580a

0.2 g

HPLC-ICP-MS

92 ± 3%

[78]

 Citrate buffer, dithizone in CHCl3, nitrite/acid mixture, Na2S2O3; 0,05 M CH3COONH4

River sediment

5 g

HPLC-AFS

80%

[97]

 H2SO4/NaCl, toluene, thiosulfate

CRM BCR 580a

0.2 mg

CGC-CVAASf

85%

[86]

 7.6% HCl, 10% 2-mercaptoethanol, 10% ammonia

CRM BCR 580a

1 g

HPLC-ICP-MS

102 ± 5%

[98]

aSediment reference material

bEthylation-cryogenic trapping–gas chromatographic quartz furnace atomic absorption spectrometry

cEthylation-cryogenic trapping–gas chromatographic quartz furnace atomic fluorescence spectrometry

dCapillary gas chromatography–electron capture detection

eGas chromatography–microwave induced plasma atomic emission spectrometry

fCapillary gas chromatography–cold vapour atomic absorption spectrometry

gRecovery of spiked sediments

When using acid extraction the right procedure has to be considered. Horvat et al. [91] have found that pure extraction with HCl is not sufficient to quantitatively dissolve CH3Hg+ from the sediment. It was also shown that a decomposition of CH3Hg+ occurs above 4 M HCl. Hintelmann et al. [99] determined that the extraction with HCl also leads to artefact formation. Bloom et al. [79] showed that a combination of H2SO4, KBr, CuSO4 results in a good isolation of methylmercury from sediments, with no formation of artifacts. Some years later it was recommended to choose nitric acid for the extraction, since better recoveries for CH3Hg+ were observed, compared to other acids [80, 90, 92] and it shows little or no interferences to ICP-MS [81]. Rahman et al. also demonstrated the impact of different nitric acid concentrations on the extraction efficiencies of both species. While an increase in the acid concentration results in better recoveries for Hg2+, a decrease was observed in the case of CH3Hg+. They supposed to use a concentration of maximum 4 M to get the best result for both species.

Apart from acidic and distillative extraction, methods for alkaline extraction have also been mentioned in the literature. In particular, the use of KOH–methanol [100] and tetramethylammonium hydroxide [101] for the extraction of methylmercury from sediments and biological samples were described. While this methodology works particularly well for biological specimens, problems occur when extracting sediments, which are associated with the presence of organic material, sulfides and iron [91].

Figure 5 describes potential pathways for the speciation of mercury species with HPLC-ICP-MS. Basically, the speciation of mercury in sediments can take place in two different ways. Whereas in case of the simpler option the sample is directly measured after alkaline or acidic extraction and neutralization (blue line), variant two (red line) uses an additional organic extraction step after the first extraction, which should lead to a higher recovery of the methylmercury.

Fig. 5

Possible approaches of extracting mercury species from sediments followed by the analysis with HPLC-ICP-MS

(adapted from [102])

Another way to analyze the mercury content is to use a direct mercury analyzer (DMA), as often described in the literature [80, 103, 104]. This measuring instrument is able to determine the mercury content of solid, liquid and gaseous samples by thermal decomposition, amalgamation and photometry. Although the main field of application is the total mercury analysis, speciation analysis is also possible. In a method according to Environmental Protection Agency (EPA) [105], speciation in sediments by a previous extraction and separation via SPE with subsequent analysis by means of DMA is described. In addition, this instrument has very low detection limits, down to the nanogram range. Because of its advantages in terms of speed, robustness and redundant sample preparation, it serves as a companion instrument in the speciation of mercury.

Extraction-Free Methods

Due to the above mentioned complications when using different extraction procedures, many researchers try to use extraction-free methods instead of using other procedures. In this context, X-ray absorption [71, 72, 105, 106, 107] and thermodesorption [73, 108, 109, 110, 111, 112, 113, 114] are often mentioned. X-ray absorption spectroscopy (XAS) is a useful tool for the nondestructive analysis of mercury compounds using high-energy X-ray radiation. The analysis does not require any chemical treatment, therefore, no change in speciation has to be feared. When using X-ray absorption, a distinction is principally made between X-ray absorption near-edge structure (XANES) and X-ray absorption fine structure (EXAFS), which produce different information about the analyte. While XANES can provide information about the state of oxidation and the geometry of the absorbing element, EXAFS refers to information about the direct neighboring elements of the absorbing atom [71]. Although it is characterized by the peculiarities and advantages the analysis of mercury species requires at least a concentration of > 100 µg g−1 [72], which makes it useless for the analysis of environmental samples. In addition, due to similar spectra, the species may be misclassified [71]. Thermodesorption is a further common technique for the speciation of mercury. The principle is based on the thermal release of the individual mercury species according to their desorption temperature and thus allows their differentiation and identification [73]. The method is characterized by several advantages, therefore, it is sensitive, allows immediate identification and requires none or little sample preparation [115]. Windmoller et al. could show that in case of Hg(0) concentrations in the µg kg−1 range can be detected [116]. For the analysis of organic Hg species a previous extraction becomes necessary. Fernandez-Martinez et al. has presented a simple extraction consisting of an acidic extraction followed by organic separation using dichloromethane, which leads to satisfactory results. In contrast to conventional methods, the procedure is particularly fast and reduces artifact formation due to fewer steps [117]. As a result this technique provides a simple and inexpensive alternative to the determination of organic mercury species.

Extraction Efficiencies and Artifact Formation

The unintentional generation of species transformations (artifacts) especially in sediments is well-known and results therefore to significant errors [20, 79].The reasons for an unwanted artifact formation are manifold but the literature suggests that artifacts occur during derivatization [91], separation [118] and extraction [20, 79]. Furthermore, it has been reported that formation of artifacts in presence of organic material [20, 85] and due to high amounts of soluble inorganic mercury [92, 119, 120] occurs, which means that the artifact formation must be strictly controlled during the analysis of sediments. For this reason, it is pointed out in the literature that the published results of methylmercury concentrations in soils and sediments must be used carefully [20]. Since there is so far no extraction method without artifact formation, these often lead to a misinterpretation of the results, provided no correction has been undertaken. To check the extraction efficiency and artifact formation, the standard addition is often used. However, Liang et al. show that the addition of Hg(II) standards is not sufficient for the identification of artifacts. They observed that the behavior of Hg(II) standards in the sample and naturally occurring Hg(II) differs in the occurrence of CH3Hg+ artifacts [90]. A much better and already established method for the control of artifacts is the species-specific isotope addition (SSIA) [20]. This method allows correction of the measured methylmercury concentration by means of stable mercury isotopes. However, an ICP-MS is required for this application in addition to the acquisition of isotopic standards. On the basis of the difficult situation, it becomes clear that a prior examination of the measuring method for the analysis of methylmercury, in particular in soils and sediments, is indispensable. Therefore, it would be advisable to find a methodology with which artifact-free measurements can be carried out.

Difficulties of Applying Available Methods for the Speciation of Mercury in Sediments

A commonly used application for the speciation of mercury in sediments is the method according to EPA [121]. This can be used as a guideline for the fractionation and quantification of mercury species and includes two approaches for the extraction of the mobile, extractable fraction and an enrichment method. The first approach describes the use of an acidic (4 M HNO3), microwave-based extraction. In the method it is stated that the extract finally contains the mobile, extractable fraction. For the analysis of the non-extractable fraction or the subdivision into semi-mobile and non-mobile, the remaining solid has to be treated further. Using this extraction procedure in our laboratory we found about 94% recovery for the inorganic mercury when analyzing the reference material CRM 580. These results suggest that the fraction found corresponds to the extractable fraction available in the reference material. Accordingly, in the reference material used, about 90% of the total mercury content should be present as the extractable fraction. Han et al. [66] on which an extraction and enrichment method is based in the method according to EPA,focused on ultrasound-based extraction of mercury species using 2% HCl + 10% ethanol. In addition to suitable reference material (CRM 580 and SRM 2709), a direct mercury analyzer was used to validate the results. He was able to show that after extraction of the reference material CRM 580, only a content of 1.4 mg kg−1 of soluble inorganic mercury could be found, which corresponds approximately to a content of 1.1% of the total mercury content. He mentions that the remaining 133 mg kg−1 represent the non-extractable mercury. In view of the characterization of the reference material and the mentioned high content of sulfur complexes, the results of Han et al. [66] appear to be more reliable.

However, this does not mean that the microwave-based method does not work, but it should be noted in the method description that it can be used for methylmercury and the total mercury content—not only extractable mercury—otherwise, a faulty interpretation of the measured values may occur. This example is intended to illustrate the difficulties of using a method from the literature. Another very frequently used method for speciation of Hg in sediments is a combination of acidic and organic extraction [122, 123, 124], often just acid extraction [47, 125], followed by ethylation and analysis with GC-CVAFS. When transferring a method based on GC-CVAFS to an HPLC–ICP–MS system, special challenges often arise, as many diversions have to be accepted. Nevado et al. presented a procedure based on a microwave extraction including purification with dichloromethane and subsequent measurement by means of GC-AFS [126]. For the transfer of the original method to an HPLC–ICP–MS system, which does not require any derivatization, especially the sample amount and the finally resulting concentration of methylmercury are crucial. Instead of the derivatization step, as mentioned in the original method, evaporation of the dichloromethane must take place with simultaneous uptake in an aqueous solution, to allow a direct injection afterwards. While the original method leads to satisfactory results (94 ± 4%) [126] and is considered as a possible universal method, the adapted approach leads to none separation, due to the high amount of inorganic mercury, and consequently making the quantification challenging. As a result, to simplify quantification when using the coupling of HPLC with ICP–MS a separation or enrichment method should always be used. These challenges should therefore always be remembered when transferring a method.

Pre-concentration

Due to the very low contents of CH3Hg+ in sediments, the use of suitable enrichment methods is necessary despite the use of sensitive measuring instruments. Jagtap et al. were able to show that the high content of Hg2+ can also require a prior separation of both species since otherwise problems with the determination of both species can occur [78].

There are several enrichment methods reported in the literature such as solid phase extraction (SPE) [127], solid phase micro extraction (SPME) [128], liquid phase microextraction (LPME) [38], dispersive liquid–liquid microextraction (DLLME) [129, 130], hollow fiber based liquid–liquid–liquid microextraction [131] and also purge and trap extraction (P&T) [132, 133].

SPE and SPME are techniques which lead to the separation or enrichment of species by the use of different stationary phases. The principle is based on the different affinities of the analytes to the stationary and mobile phases. In the context of mercury speciation in sediments, thiol-containing reagents such as dithizone, l-cysteine, 2-mercaptoethanol or sodium thiosulfate are often used. Table 4 shows a literature section of SPE applications for mercury species from sediments and biological samples. It should be noted, however, that despite the high number of enrichment methods, only a few are available for the analysis of sediments.

Table 4

SPE methods for the analysis of mercury species from sediments and biological samples

Stationary phase

Pretreatment with functional groups

Elution

Detection method

Sample

References

Thiol/thiourea on silica

–SH

75% methanol, 1.5 mM ammonium pyrrolidine dithiocarbamate

LC-CV/AFS

Urine, sediment, biological tissue

[77]

Cotton

–SH

1M HCl, 1M NaCl or 6 M HCl, saturated NaCl, 0.1% CuCl2. 2H2O

LC-ICP/MS

Soil and sediments

[80]

RP18

Methanol, 14 mmolL-1 2-mercaptophenol

LC-CV/AFS

Sediments, zoobenthos, river water

[134]

RP18 with ODS

0.5 mM ammoniumacetat

LC-ICP/MS

Needles, moss, leaves, soil, sediment, foodstuff

[135]

Purge and trap is a further enrichment method, which is mainly used in combination with gas chromatography [50, 132, 133, 136]. This common used and effective method involves purging a sample (liquid/solid) with an inert gas (Ar, He, N2) to trap the volatile analytes on sorbents like Tenax [50, 133, 137]. Since this technique requires a cleaning step and a derivatization, other methods are recommended for the enrichment of mercury species.

In addition to the above-mentioned methods, the LPME or especially the DLLME are frequently reported in this context. The latter makes particular use of chelating agents such as dithizone [130] or l-cysteine [129] when specifying mercury. This enrichment method has some advantages as it is characterized by high enrichment factors, consumption of little extraction agents as well as simplicity. The functional principle is based on the use of an extracting agent in combination with a dispersant, which is added to the aqueous sample and thus results in a satisfactory extraction. Because of the green chemistry thoughts and the consumption of different solvents, vortex [138] or ultrasound [139] is suggested [38].

Hyphenated Techniques

Separation

The usage of hyphenated techniques is very popular when analyzing mercury species. Especially for sediments and the small amount of methylmercury, a powerful separation technique is indispensable. Some methods are described for the separation of mercury species during analysis, with a distinction being made between chromatographic methods like gas chromatography (GC) [140, 141, 142, 143, 144, 145], liquid chromatography, including high performance liquid chromatography (HPLC) [96, 146, 147, 148, 149, 150, 151, 152, 153, 154] and ion chromatography (IC) [155], capillary electrophoresis (CE) [156, 157, 158, 159, 160] and non-chromatographic methods involving cold-vapor (CV) generation [77, 161, 162, 163] or thermal desorption [73, 108, 109, 110, 111, 112, 113, 114]. Figure 6 shows an overview of the used methods.

Fig. 6

Frequently used separation methods for the speciation of mercury in sediments

(adapted from [38])

GC is one of the most well-known methods for the separation of mercury species and represents, especially in combination with ICP–MS or AFS, a powerful coupling for specifying Hg in complex samples. If ICP–MS is used as detection technique it is pointed out in the literature that GC is the best separation technique for this coupling due to its 100% sample introduction [140]. Furthermore, it has a higher sensitivity [141] and better separation [164] compared to HPLC–ICP–MS, which can be of particular importance if sediments have to be analyzed. Table 5 illustrates some methods for specifying mercury with GC. But to utilize GC, a derivatization in form of aqueous phase ethylation [91, 143, 144], propylation [165] or phenylation [165] is necessary. The literature highlights the fact that this additional and complex step can lead to species transformation as well as losses and contamination of the sample [164, 166]. Therefore HPLC as a rapid, efficient and derivatization-free method is more often used. Despite of the lower sensitivity compared to other methods, the use of HPLC for the speciation of mercury offers a wide application range because of a higher variability in stationary and mobile phases [167]. An overview of the broad application with HPLC for the speciation of mercury is given in a review by Harrington in 2000 [168]. Table 6 shows an excerpt of LC based separation techniques.

Table 5

GC methods for the separation of mercury species

(adapted from [38])

Matrix

Pretreatment

Detection

References

Sediment

CuSO4, H2SO4 and KBr

CV-AFS

[169]

Sediment

TSE1

ICP–MS

[50]

Sediment

CuSO4, H2SO4 and KBr

ICP–MS

[170]

Fish and sediment

P & T

MS

[171]

Sediment

P & T

Py2-AFS

[87]

Sediment

Alkaline digestion

ECD

[172]

Sediment

P & T

ICP–MS

[173]

1Thiosulfate extraction

2Pyrolysis

Table 6

HPLC methods for the separation of mercury species

Matrix

Stationary phase

Mobile phase

Detection

References

Sediment

C18

3% acetonitrile, 60 mM NH4Ac-acetic acid (pH 4.5), 0.1% 2-mercaptoethanol

CV-AFS

[138]

Lake water and sediment

C18

Methanol/(2.5 mM l-cysteine,12.5 mM (NH4)2HPO4, 0.05% trimethylamine, pH 7.0)

ICP–MS

[131]

Sediment

C8

0.5% 2-mercaptoethanol in 5% CH3OH, pH 5.3

ICP–MS

[78]

Sediment CRM

C18

Methanol, 2-mercaptoethanol,pH 5

UV-CV-AFS

[96]

Sediment

C8

75% Methanol, 1.5 mM ammoniumpyrrolidine dithiocarbamate

CV-AFS

[77]

Sediment

C18

2.5 mM l-cysteine, 12.5 mM (NH4)2HPO4, 0.05% trimethylamine, pH 7.0

ICP–MS

[131]

Sediment

C18

Acetonitrile/water (65:35), 0.5 mM sodium pyrrolidine dithiocarbamate, ammonium acetat pH 5.5

ICP–MS

[135]

The HPLC methods mentioned in the literature for specifying Hg are based on reversed phase (RP)-stationary phases including C18 [127, 131] or C8 [78, 130].

To achieve good separation in RP-HPLC, methanol is an important component in many mobile phases. It could be shown that the methanol content has a crucial influence on the separation of mercury species [174]. But when using ICP–MS, it has to be considered that the solvent content should not exceed 5%. Without an addition of oxygen to the plasma gas a higher concentration leads to instability of the plasma and deposits on the cones [175, 176]. In consequence there are limited possibilities to separate the species. Therefore the use of sulfur-containing additives, such as 2-mercaptoethanol [78, 98, 177] or l-cysteine [147, 149, 150, 168], plays an important role when specifying mercury. Rai et al. were able to confirm the importance of these substances by showing that adsorption effects can be prevented by the use of l-cysteine in the mobile phase [178]. Depending on the used compounds a change in elution order can be observed. This occurrence can be explained by the different pKa values and the ultimately formed complexes. While neutral complexes are formed during the use of 2-mercaptoethanol, which lead to longer retention times, the use of l-cysteine results in charged complexes which have less interaction with the column and thus a shorter retention time [78]. This can be helpful when analyzing real samples due to the large excess of inorganic mercury, which can lead to overlapping of the smaller CH3Hg+ peak.

Ion chromatography also belongs to the group of liquid chromatography and describes a further effective and fast method for specifying mercury species [38, 179]. It is characterized above all by the use of cation exchange columns and the absence of organic solvents, why IC is often described as a “green” method [155]. Despite the good properties of this separation method, there is so far no application for sediments. The capillary electrophoresis is another separation method which is characterized by good separation efficiency, high resolution, high separation speed, low sample consumption and the lack of interaction between analyte and capillary [38]. Despite the many good features of this separation technique, the main drawback of CE is a lower sensitivity compared to LC and GC. By coupling to element-specific detectors, such as ICP–MS, the sensitivity can be positively influenced, but the coupling may lead to undesired memory effects [180]. As a result, the CE has little use in specifying mercury.

Detection

Numerous detection techniques are discussed in the literature, which are coupled with the above mentioned separation techniques when specifying mercury. Depending on the analyte and the chosen separation method, the appropriate detection method can be selected. For the detection of mercury species element specific detectors like atomic absorption spectroscopy (AAS) [181, 182], cold vapor atomic absorption spectrometry (CV-AAS) [183, 184], atomic fluorescence spectroscopy (AFS) [134, 185, 186, 187], especially cold-vapor atomic fluorescence spectrometry (CV-AFS) [188], mass spectrometry (MS) [146, 150, 189, 190] and inductively coupled plasma mass spectrometry (ICP–MS) [127, 131, 132, 155, 191, 192, 193, 194, 195, 196, 197, 198] are used. Table 7 provides a brief overview of the above-mentioned detection techniques used in many hyphenated techniques. GC and HPLC are the most common separation methods for the hyphenation with an element specific detector. As pointed out before GC has the particular advantage as it allows a quantitative transfer of the sample, without previous nebulization. The detection systems used for the coupling with GC are usually AFS [126, 199, 200], ICP–MS [201, 202, 203] and as alternative a common MS [144, 204]. AFS and ICP–MS are widely used for the detection of several elements, especially of mercury, arsenic and selenium [205] and are characterized by high selectivity and sensitivity. AFS is also characterized by low costs as well as easy handling [38, 205]. Because of the mentioned advantages, the AFS was particularly popular in the 1980s and 1990s and is still a suitable detection method for the speciation of mercury. Especially in combination with the advantages of gas chromatography, it is one of the most popular methods for the speciation of trace metals [199]. Though less common than other combinations, GC-MS offers a simple and inexpensive alternative to GC–ICP–MS and GC–AFS [144, 194]. Nevertheless it is characterized by higher limits of detection, which can be crucial when analyzing trace metals in various matrices. In addition to high sensitivity and selectivity the coupling of GC and ICP–MS (Table 8) offers also the possibility of a multi-element analysis and unlike other methods, it is characterized by the opportunity to show artifacts using speciated isotope dilution mass spectrometry (SID-MS) [3, 20, 66, 206, 207, 208, 209, 210]. However, it must be pointed out that all hyphenated techniques including GC require a derivatization step as it is only suitable for volatile species. To form more easily volatiles sodium tetraethylborate [119] is commonly used. But according to the literature derivatization may be time consuming [3] and artifact formation has to be feared [91].

Table 7

Commonly hyphenated techniques used for the speciation of mercury

Separation

Detection

References

HPLC

(CV)-AAS

HG-AAS

(CV)-AFS

(CV)/(HG)-AFS

API-MS

ICP-MS

DAD

[128]

[78]

[138, 211, 212]

[213]

[214]

[127, 131, 155, 191, 192, 193]

[215]

GC

(CV)-AFS

MS

ICP-MS

AAS

[24, 200]

[194]

[132, 194]

[81]

CE

ICP-MS

UV

DAD

[195, 196, 197]

[216, 217, 218]

[129]

To overcome these challenges usually LC methods are used. LC–CV–AFS is a widely used technique as it allows low detection limits without prior derivatization. Further advantages are lower running costs and simplicity, compared to LC–ICP–MS, by providing higher sensitivity. Using this hyphenation it is essential to imply an additional step for the conversion of organic to inorganic mercury with a suitable reagent, in general sodium borohydride or tin(II)chloride [205]. In addition, no control of artifacts can be achieved using appropriate isotopic standards.

Table 8

Analytical performance of relevant analytical methods for mercury species in sediments

Separation

Detection

Derivatization

Preconcentration

LOD methyl mercury (ng L−1)

LOD inorganic Hg (ng L−1)

References

HPLC

ICP-MS

HF-LLLME

2.9

4.1

[131]

GC

ICP-MS

NaBEt4

0.2

0.1

[170]

HPLC

(CV)-AFS

Thiol–thiourea on silica

1.5

[77]

GC

(CV)-AFS

NaBEt4

LLE

1.12

[169]

Other detection systems used in combination with HPLC are presented by ICP–AES, ICP–MS, CV–AAS and API–MS [152]. Whereas in view of the particularly low detection limits (ppb-range [167]) the use of ICP–MS as detection method is preferred. Despite the higher operating costs and lower sensitivity compared to AFS [205], ICP–MS, especially in combination with HPLC, offers a wide spectrum of analytical methods. Based on its capabilities, ICP–MS is a detection method that is closest to the ideal for optimal detection (low detection limits, wide linear range, ability to measure different isotopes, application of isotope dilution [3, 119, 122, 208]). Therefore the use of ICP-MS has grown enormously, which can be confirmed by the high number of publications [3]. Compared with the other mentioned coupling types, the combination of the HPLC (easy sample pretreatment, no derivatization required, various number of separation methods) with ICP–MS as detection technique leads to a powerful hyphenated technique for the speciation of mercury in sediments.

Conclusion

According to the World Health Organization, mercury is one of the most dangerous substances. The reason for this is the occurrence of mercury in different species, which differ in their degree of toxicity. Due to this fact, a speciation is indispensable for the evaluation of the environmental condition. The speciation can be divided into several sub-sections, which are defined by extraction, enrichment, separation and detection. When choosing the right methods, artifact formation is a crucial factor that must be taken into account, as this can lead to incorrect interpretation of results. For the correct evaluation of speciation methods reference materials are often used. Some reference materials are already available on the market for mercury speciation in sediments. However, these are often certified only for the total and methylmercury content. Since there are also other species available in sediments, it would be useful to offer reference materials that are also suitable for the analysis of these.

Above all, sediments, in which the methylmercury concentration is only 0.1–1% of the total mercury content, must be treated with special caution. For the extraction of sediments mainly acidic, alkaline or distillative methods are used. However, the acid extraction is the most common technique and is used in various modifications. As already mentioned, the concentration of methylmercury in sediments is very low, a suitable enrichment method followed after extraction is necessary. For the enrichment of mercury species solid phase extraction is a very frequently used method and provides a number of different phases, e.g. thiol-containing compounds with a high affinity for mercury. After a successful enrichment, the separation of the species with a suitable separation method takes place. HPLC and GC are the most commonly used methods for this purpose. Both methods have their advantages and disadvantages, but a considerable disadvantage of GC is the necessity of derivatization. This required step leads, beyond additional time, to the formation of artifacts. Due to the lack of a need for derivatization and the enormous variety of separation phases, HPLC has many advantages in specifying mercury in sediments. A number of detection methods are discussed in the literature, while (CV)-AFS and ICP–MS are the most common. Despite the many mentioned advantages of (CV)-AFS for the speciation of mercury, ICP-MS has the essential advantage that it does not need any derivatization. In addition, this technology is characterized by the significant advantage to show artifacts using speciated isotope dilution mass spectrometry. Nevertheless by checking the literature of the last five years, a trend towards the use of GC based methods in combination with CV–AFS becomes obvious. In addition, it is mainly referred to the use of existing methods (e.g. USEPA 1630) and shown little new methods [47, 77, 122, 124, 125, 169, 219, 220, 221, 222, 223, 224, 225, 226, 227, 228, 229, 230]. Based on the ease of use, robustness and low running cost [205], the combination of GC and CVAFS seems to have a firm place in the speciation of Hg. Due to the complexity of the speciation of mercury in sediments and the resulting high number of publications in this field, this review should give an overview of the existing methods and consider them critically. Background is the property of sediments, in particular, to bind heavy metals and release them depending on environmental conditions. Accordingly, they serve as risky storage that needs to be controlled. Therefore, the goal is to find a working, robust standard method that is suitable for different sediments.

Notes

Acknowledgements

Financial support from the Federal Institute of Hydrology is gratefully acknowledged.

Compliance with ethical standards

Conflict of interest

The authors declare that they have no conflict of interest.

References

  1. 1.
    Kudo A, Miyahara S (1991) Water Sci Technol 23:283–290Google Scholar
  2. 2.
    WHO (2010) Public Health and Environment. World Health Organization, Geneva. http://www.who.int/ipcs/features/10chemicals_en.pdf?ua=1
  3. 3.
    Leermakers M, Baeyens W, Quevauviller P, Horvat M (2005) Trac Trend Anal Chem 24:383–393.  https://doi.org/10.1016/j.trac.2004.01.001 Google Scholar
  4. 4.
    Fitzgerald WF, Lamborg CH, Hammerschmidt CR (2007) Chem Rev 107:641–662.  https://doi.org/10.1021/cr050353m Google Scholar
  5. 5.
    Hursh JB, Clarkson TW, Cherian MG, Vostal JJ, Vandermallie R (1976) Arch Environ Health 31:302–309Google Scholar
  6. 6.
    Syversen T, Kaur P (2012) J Trace Elem Med Bio 26:215–226.  https://doi.org/10.1016/j.jtemb.2012.02.004 Google Scholar
  7. 7.
    Gochfeld M (2003) Ecotox Environ Safe 56:174–179.  https://doi.org/10.1016/S0147-6513(03)00060-5 Google Scholar
  8. 8.
    Friberg L, Skog E, Wahlberg JE (1961) Acta Derm Venereol 41:40–52Google Scholar
  9. 9.
    Magos L, Brown AW, Sparrow S, Bailey E, Snowden RT, Skipp WR (1985) Arch Toxicol 57:260–267.  https://doi.org/10.1007/Bf00324789 Google Scholar
  10. 10.
    Clarkson TW (2002) Environ Health Persp 110:11–23Google Scholar
  11. 11.
    Kerper LE, Ballatori N, Clarkson TW (1992) Am J Physiol 262:R761–R765Google Scholar
  12. 12.
    Aberg B, Ekman L, Falk R, Greitz U, Persson G, Snihs JO (1969) Arch Environ Health 19:478–484Google Scholar
  13. 13.
    Ullrich SM, Tanton TW, Abdrashitova SA (2001) Crit Rev Environ Sci Technol 31:241–293.  https://doi.org/10.1080/20016491089226 Google Scholar
  14. 14.
    Li WC, Tse HF (2015) Environ Sci Pollut Res 22:192–201.  https://doi.org/10.1007/s11356-014-3544-x Google Scholar
  15. 15.
    Issaro N, Abi-Ghanem C, Bermond A (2009) Anal Chim Acta 631:1–12.  https://doi.org/10.1016/j.aca.2008.10.020 Google Scholar
  16. 16.
    Aceto M, Foglizzo AM, Mentasti E, Sacchero G, Sarzanini C (1995) Int J Environ Anal Chem 60:1–13.  https://doi.org/10.1080/03067319508027222 Google Scholar
  17. 17.
    Mason RP, Reinfelder JR, Morel FMM (1995) Water Air Soil Pollut 80:915–921.  https://doi.org/10.1007/Bf01189744 Google Scholar
  18. 18.
    Malm O, Branches FJP, Akagi H, Castro MB, Pfeiffer WC, Harada M, Bastos WR, Kato H (1995) Sci Total Environ 175:141–150.  https://doi.org/10.1016/0048-9697(95)04910-X Google Scholar
  19. 19.
    Weber JH, Puk R (1994) Appl Organomet Chem 8:709–713.  https://doi.org/10.1002/aoc.590080723 Google Scholar
  20. 20.
    Hintelmann H, Falter R, Ilgen G, Evans RD (1997) Fresen J Anal Chem 358:363–370.  https://doi.org/10.1007/s002160050431 Google Scholar
  21. 21.
    Leopold K, Foulkes M, Worsfold P (2010) Anal Chim Acta 663:127–138.  https://doi.org/10.1016/j.aca.2010.01.048 Google Scholar
  22. 22.
    Bowles KC, Apte SC, Maher WA, Bluhdorn DR (2003) Water Air Soil Pollut 147:25–38.  https://doi.org/10.1023/A:1024561830113 Google Scholar
  23. 23.
    Bisinoti MC, Junior ES, Jardim WF (2007) J Brazil Chem Soc 18:544–553.  https://doi.org/10.1590/S0103-50532007000300008 Google Scholar
  24. 24.
    Araujo BF, Hintelmann H, Dimock B, Sobrinho RD, Bernardes MC, de Almeida MG, Krusche AV, Rangel TP, Thompson F, de Rezende CE (2018) Limnol Oceanogr 63:1134–1145.  https://doi.org/10.1002/lno.10758 Google Scholar
  25. 25.
    He TR, Lu J, Yang F, Feng XB (2007) Sci Total Environ 386:53–64.  https://doi.org/10.1016/j.scitotenv.2007.07.022 Google Scholar
  26. 26.
    Kannan K, Smith RG, Lee RF, Windom HL, Heitmuller PT, Macauley JM, Summers JK (1998) Arch Environ Contam Toxicol 34:109–118.  https://doi.org/10.1007/s002449900294 Google Scholar
  27. 27.
    Mikac N, Niessen S, Ouddane B, Wartel M (1999) Appl Organomet Chem 13:715–725.  https://doi.org/10.1002/(Sici)1099-0739(199910)13:10%3C715::Aid-Aoc918%3E3.0.Co;2-4 Google Scholar
  28. 28.
    Kannan K, Falandysz J (1998) Water Air Soil Pollut 103:129–136.  https://doi.org/10.1023/A:1004967112178 Google Scholar
  29. 29.
    Wilken RD, Hintelmann H (1991) Water Air Soil Pollut 56:427–437.  https://doi.org/10.1007/Bf00342289 Google Scholar
  30. 30.
    Minganti V, Capelli R, Drava G, De Pellegrini R (2007) Chemosphere 67:1018–1024.  https://doi.org/10.1016/j.chemosphere.2006.10.053 Google Scholar
  31. 31.
    Gabriel MC, Williamson DG (2004) Environ Geochem Health 26:421–434.  https://doi.org/10.1007/s10653-004-1308-0 Google Scholar
  32. 32.
    Ibanez-Palomino C, Lopez-Sanchez JF, Sahuquillo A (2012) Anal Chim Acta 720:9–15.  https://doi.org/10.1016/j.aca.2012.01.015 Google Scholar
  33. 33.
    Le Roux S, Baker P, Crouch A (2016) S Afr J Chem S Afr T 69:124–131.  https://doi.org/10.17159/0379-4350/2016/v69a15 Google Scholar
  34. 34.
    Templeton DM, Ariese F, Cornelis R, Danielsson LG, Muntau H, Van Leeuwen HP, Lobinski R (2000) Pure Appl Chem 72:1453–1470.  https://doi.org/10.1351/pac200072081453 Google Scholar
  35. 35.
    Sarica DY, Turker AR (2012) Clean Soil Air Water 40:523–530.  https://doi.org/10.1002/clen.201100535 Google Scholar
  36. 36.
    Braaten HFV, de Wit HA, Harman C, Hagestrom U, Larssen T (2014) Int J Environ Anal Chem 94:381–384.  https://doi.org/10.1080/03067319.2013.823489 Google Scholar
  37. 37.
    Martinis EM, Wuilloud RG (2010) J Anal Atom Spectrom 25:1432–1439.  https://doi.org/10.1039/c004678g Google Scholar
  38. 38.
    Amde M, Yin YG, Zhang D, Liu JF (2016) Chem Speciat Bioavailab 28:51–65.  https://doi.org/10.1080/09542299.2016.1164019 Google Scholar
  39. 39.
    Yu LP, Yan XP (2003) Trac Trend Anal Chem 22:245–253.  https://doi.org/10.1016/S0165-9936(03)00407-2 Google Scholar
  40. 40.
    Rosain RM, Wai CM (1973) Anal Chim Acta 65:279–284.  https://doi.org/10.1016/S0003-2670(01)82493-4 Google Scholar
  41. 41.
    Leermakers M, Lansens P, Baeyens W (1990) Fresen J Anal Chem 336:655–662.  https://doi.org/10.1007/Bf00331410 Google Scholar
  42. 42.
    Krivan V, Haas HF (1988) Fresen Z Anal Chem 332:1–6.  https://doi.org/10.1007/Bf00487020 Google Scholar
  43. 43.
    Stoeppler M, Matthes W (1978) Anal Chim Acta 98:389–392.  https://doi.org/10.1016/S0003-2670(01)84069-1 Google Scholar
  44. 44.
    Lansens P, Meuleman C, Baeyens W (1990) Anal Chim Acta 229:281–285.  https://doi.org/10.1016/S0003-2670(00)85140-5 Google Scholar
  45. 45.
    Sedlackova L, Kruzikova K, Svobodova Z (2014) Food Chem 150:360–365.  https://doi.org/10.1016/j.foodchem.2013.10.041 Google Scholar
  46. 46.
    Li X, Wang Y, Li BH, Feng CH, Chen YX, Shen ZY (2013) Environ Earth Sci 69:1537–1547.  https://doi.org/10.1007/s12665-012-1988-1 Google Scholar
  47. 47.
    Kim E, Noh S, Lee YG, Kundu SR, Lee BG, Park K, Han S (2014) Mar Chem 158:59–68.  https://doi.org/10.1016/j.marchem.2013.11.004 Google Scholar
  48. 48.
    Zhang T, Kucharzyk KH, Kim B, Deshusses MA, Hsu-Kim H (2014) Environ Sci Technol 48:9133–9141.  https://doi.org/10.1021/es500336j Google Scholar
  49. 49.
    Weiss HV, Shipman WH, Guttman MA (1976) Anal Chim Acta 81:211–217.  https://doi.org/10.1016/S0003-2670(00)89480-5 Google Scholar
  50. 50.
    Avramescu ML, Zhu J, Yumvihoze E, Hintelmann H, Fortin D, Lean DRS (2010) Environ Toxicol Chem 29:1256–1262.  https://doi.org/10.1002/etc.158 Google Scholar
  51. 51.
    Leermakers M, Nguyen HL, Kurunczi S, Vanneste B, Galletti S, Baeyens W (2003) Anal Bioanal Chem 377:327–333.  https://doi.org/10.1007/s00216-003-2116-6 Google Scholar
  52. 52.
    Fabbri D, Felisatti O, Lombardo M, Trombini C, Vassura I (1998) Sci Total Environ 213:121–128.  https://doi.org/10.1016/S0048-9697(98)00083-7 Google Scholar
  53. 53.
    Quevauviller P, Fortunati GU, Filippelli M, Bortoli A, Muntau H (1998) Appl Organomet Chem 12:531–539.  https://doi.org/10.1002/(Sici)1099-0739(199808/09)12:8/9%3C531::Aid-Aoc758%3E3.0.Co;2-I Google Scholar
  54. 54.
    Vazquez MJ, Carro AM, Lorenzo RA, Cela R (1997) Anal Chem 69:221–225.  https://doi.org/10.1021/ac960513h Google Scholar
  55. 55.
    Hintelmann H (1999) Chemosphere 39:1093–1105.  https://doi.org/10.1016/S0045-6535(99)00180-0 Google Scholar
  56. 56.
    Frohne T, Rinklebe J (2013) Water Air Soil Pollut 224:1591.  https://doi.org/10.1007/s11270-013-1591-4 Google Scholar
  57. 57.
    Wallschlager D, Desai MVM, Wilken RD (1996) Water Air Soil Pollut 90:507–520.  https://doi.org/10.1007/Bf00282665 Google Scholar
  58. 58.
    Davis A, Bloom NS, Hee SSQ (1997) Risk Anal 17:557–569.  https://doi.org/10.1111/j.1539-6924.1997.tb00897.x Google Scholar
  59. 59.
    Karlsson T, Skyllberg U (2003) Environ Sci Technol 37:4912–4918.  https://doi.org/10.1021/es034302n Google Scholar
  60. 60.
    Ravichandran M (2004) Chemosphere 55:319–331.  https://doi.org/10.1016/j.chemosphere.2003.11.011 Google Scholar
  61. 61.
    Khwaja AR, Bloom PR, Brezonik PL (2006) Environ Sci Technol 40:844–849.  https://doi.org/10.1021/es051805c Google Scholar
  62. 62.
    Zhong H, Wang WX (2008) Environ Pollut 151:222–230.  https://doi.org/10.1016/j.envpol.2007.01.049 Google Scholar
  63. 63.
    Skyllberg U, Qian J, Frech W, Xia K, Bleam WF (2003) Biogeochemistry 64:53–76.  https://doi.org/10.1023/A:1024904502633 Google Scholar
  64. 64.
    Fiorentino JC, Enzweiler J, Angelica RS (2011) Water Air Soil Pollut 221:63–75.  https://doi.org/10.1007/s11270-011-0769-x Google Scholar
  65. 65.
    Manohar DM, Krishnan KA, Anirudhan TS (2002) Water Res 36:1609–1619Google Scholar
  66. 66.
    Han Y, Kingston HM, Boylan HM, Rahman GMM, Shah S, Richter RC, Link DD, Bhandari S (2003) Anal Bioanal Chem 375:428–436.  https://doi.org/10.1007/s00216-002-1701-4 Google Scholar
  67. 67.
    Bloom NS, Preus E, Katon J, Hiltner M (2003) Anal Chim Acta 479:233–248.  https://doi.org/10.1016/S0003-2670(02)01550-7 Google Scholar
  68. 68.
    Saniewska D, Beldowska M (2017) Talanta 168:152–161.  https://doi.org/10.1016/j.talanta.2017.03.026 Google Scholar
  69. 69.
    Reis AT, Rodrigues SM, Davidson CM, Pereira E, Duarte AC (2010) Chemosphere 81:1369–1377.  https://doi.org/10.1016/j.chemosphere.2010.09.030 Google Scholar
  70. 70.
    Bacon JR, Davidson CM (2008) Analyst 133:25–46.  https://doi.org/10.1039/b711896a Google Scholar
  71. 71.
    Andrews JC (2006) Struct Bond 120:1–35.  https://doi.org/10.1007/430_011 Google Scholar
  72. 72.
    Kim CS, Bloom NS, Rytuba JJ, Brown GE (2003) Environ Sci Technol 37:5102–5108.  https://doi.org/10.1021/es0341485 Google Scholar
  73. 73.
    Reis AT, Coelho JP, Rucandio I, Davidson CM, Duarte AC, Pereira E (2015) Geoderma 237:98–104.  https://doi.org/10.1016/j.geoderma.2014.08.019 Google Scholar
  74. 74.
    Barnett MO, Harris LA, Turner RR, Henson TJ, Melton RE, Stevenson RJ (1995) Water Air Soil Pollut 80:1105–1108.  https://doi.org/10.1007/Bf01189771 Google Scholar
  75. 75.
    Martin JM, Nirel P, Thomas AJ (1987) Mar Chem 22:313–341.  https://doi.org/10.1016/0304-4203(87)90017-X Google Scholar
  76. 76.
    Gleyzes C, Tellier S, Astruc M (2002) Trac Trend Anal Chem 21:451–467.  https://doi.org/10.1016/S0165-9936(02)00603-9 Google Scholar
  77. 77.
    Brombach CC, Gajdosechova Z, Chen B, Brownlow A, Corns WT, Feldmann J, Krupp EM (2015) Anal Bioanal Chem 407:973–981.  https://doi.org/10.1007/s00216-014-8254-1 Google Scholar
  78. 78.
    Jagtap R, Krikowa F, Maher W, Foster S, Ellwood M (2011) Talanta 85:49–55.  https://doi.org/10.1016/j.talanta.2011.03.022 Google Scholar
  79. 79.
    Bloom NS, Colman JA, Barber L (1997) Fresen J Anal Chem 358:371–377.  https://doi.org/10.1007/s002160050432 Google Scholar
  80. 80.
    Rahman GMM, Kingston HM (2005) J Anal Atom Spectrom 20:183–191.  https://doi.org/10.1039/b404581e Google Scholar
  81. 81.
    Tseng CM, deDiego A, Martin FM, Donard OFX (1997) J Anal Atom Spectrom 12:629–635.  https://doi.org/10.1039/a700832e Google Scholar
  82. 82.
    Qian J, Skyllberg U, Tu Q, Bleam WF, Frech W (2000) Fresen J Anal Chem 367:467–473.  https://doi.org/10.1007/s002160000364 Google Scholar
  83. 83.
    Xiang WJ, Liu J, Chang M, Zheng CG (2012) Chem Eng J 200:91–96.  https://doi.org/10.1016/j.cej.2012.06.025 Google Scholar
  84. 84.
    Roulet M, Guimaraes JRD, Lucotte M (2001) Water Air Soil Pollut 128:41–60.  https://doi.org/10.1023/A:1010379103335 Google Scholar
  85. 85.
    Falter R (1999) Chemosphere 39:1051–1073.  https://doi.org/10.1016/S0045-6535(99)00178-2 Google Scholar
  86. 86.
    Quevauviller P (1999) Chemosphere 39:1153–1165.  https://doi.org/10.1016/S0045-6535(99)00184-8 Google Scholar
  87. 87.
    Carrasco L, Vassileva E (2015) Anal Chim Acta 853:167–178.  https://doi.org/10.1016/j.aca.2014.10.026 Google Scholar
  88. 88.
    Canario J, Antunes P, Lavrado J, Vale C (2004) Trac Trend Anal Chem 23:799–806.  https://doi.org/10.1016/j.trac.2004.08.009 Google Scholar
  89. 89.
    Dmytriw R, Mucci A, Lucotte M, Pichet P (1995) Water Air Soil Pollut 80:1099–1103.  https://doi.org/10.1007/Bf01189770 Google Scholar
  90. 90.
    Liang L, Horvat M, Feng XB, Shang LH, Lil H, Pang P (2004) Appl Organomet Chem 18:264–270.  https://doi.org/10.1002/aoc.617 Google Scholar
  91. 91.
    Horvat M, Bloom NS, Liang L (1993) Anal Chim Acta 281:135–152.  https://doi.org/10.1016/0003-2670(93)85348-N Google Scholar
  92. 92.
    Hammerschmidt CR, Fitzgerald WF (2001) Anal Chem 73:5930–5936.  https://doi.org/10.1021/ac010721w Google Scholar
  93. 93.
    Lorenzo RA, Vazquez MJ, Carro AM, Cela R (1999) Trac Trend Anal Chem 18:410–416.  https://doi.org/10.1016/S0165-9936(99)00118-1 Google Scholar
  94. 94.
    Tseng CM, DeDiego A, Martin FM, Amouroux D, Donard OFX (1997) J Anal Atom Spectrom 12:743–750.  https://doi.org/10.1039/a700956i Google Scholar
  95. 95.
    Bowles KC, Apte SC (2000) Anal Chim Acta 419:145–151.  https://doi.org/10.1016/S0003-2670(00)00997-1 Google Scholar
  96. 96.
    Ramalhosa E, Segade SR, Pereira E, Vale C, Duarte A (2001) Anal Chim Acta 448:135–143.  https://doi.org/10.1016/S0003-2670(01)01317-4 Google Scholar
  97. 97.
    Hintelmann H, Wilken RD (1993) Appl Organomet Chem 7:173–180.  https://doi.org/10.1002/aoc.590070303 Google Scholar
  98. 98.
    Cattani I, Spalla S, Beone GM, Del Re AAM, Boccelli R, Trevisan M (2008) Talanta 74:1520–1526.  https://doi.org/10.1016/j.talanta.2007.09.029 Google Scholar
  99. 99.
    Hintelmann H, Evans RD (1997) Fresen J Anal Chem 358:378–385.  https://doi.org/10.1007/s002160050433 Google Scholar
  100. 100.
    Bloom NS (1992) Can J Fish Aquat Sci 49:1010–1017.  https://doi.org/10.1139/F92-113 Google Scholar
  101. 101.
    Tseng CM, de Diego A, Pinaly H, Amouraoux D, Donard OFX (1998) J Anal Atom Spectrom 13:755–764.  https://doi.org/10.1039/A802344a Google Scholar
  102. 102.
    Jagtap R, Maher W (2015) Microchem J 121:65–98.  https://doi.org/10.1016/j.microc.2015.01.010 Google Scholar
  103. 103.
    Maggi C, Berducci MT, Bianchi J, Giani M, Campanella L (2009) Anal Chim Acta 641:32–36.  https://doi.org/10.1016/j.aca.2009.03.033 Google Scholar
  104. 104.
    Rezende PS, Silva NC, Moura WD, Windmoller CC (2018) Microchem J 140:199–206.  https://doi.org/10.1016/j.microc.2018.04.006 Google Scholar
  105. 105.
    Kim CS, Brown GE, Rytuba JJ (2000) Sci Total Environ 261:157–168.  https://doi.org/10.1016/S0048-9697(00)00640-9 Google Scholar
  106. 106.
    Sladek C, Gustin MS (2003) Appl Geochem 18:567–576.  https://doi.org/10.1016/S0883-2927(02)00115-4 Google Scholar
  107. 107.
    Kim CS, Rytuba JJ, Brown GE (2004) J Colloid Interf Sci 271:1–15.  https://doi.org/10.1016/S0021-9797(03)00330-8 Google Scholar
  108. 108.
    Biester H, Scholz C (1997) Environ Sci Technol 31:233–239.  https://doi.org/10.1021/es960369h Google Scholar
  109. 109.
    Bollen A, Wenke A, Biester H (2008) Water Res 42:91–100.  https://doi.org/10.1016/j.watres.2007.07.011 Google Scholar
  110. 110.
    Higueras P, Oyarzun R, Biester H, Lillo J, Lorenzo S (2003) J Geochem Explor 80:95–104.  https://doi.org/10.1016/S0375-6742(03)00185-7 Google Scholar
  111. 111.
    Hojdova M, Navratil T, Rohovec J (2008) Bull Environ Contam Toxicol 80:237–241.  https://doi.org/10.1007/s00128-007-9352-y Google Scholar
  112. 112.
    Piani R, Covelli S, Biester H (2005) Appl Geochem 20:1546–1559.  https://doi.org/10.1016/j.apgeochem.2005.04.003 Google Scholar
  113. 113.
    Rallo M, Lopez-Anton MA, Perry R, Maroto-Valer MM (2010) Fuel 89:2157–2159.  https://doi.org/10.1016/j.fuel.2010.03.037 Google Scholar
  114. 114.
    Rumayor M, Diaz-Somoano M, Lopez-Anton MA, Martinez-Tarazona MR (2013) Talanta 114:318–322.  https://doi.org/10.1016/j.talanta.2013.05.059 Google Scholar
  115. 115.
    Reis AT, Coelho JP, Rodrigues SM, Rocha R, Davidson CM, Duarte AC, Pereira E (2012) Talanta 99:363–368.  https://doi.org/10.1016/j.talanta.2012.05.065 Google Scholar
  116. 116.
    Windmoller CC, Silva NC, Andrade PHM, Mendes LA, do Valle CM (2017) Anal Methods UK 9:2159–2167.  https://doi.org/10.1039/c6ay03041f Google Scholar
  117. 117.
    Fernandez-Martinez R, Rucandio I (2013) Anal Methods UK 5:4131–4137.  https://doi.org/10.1039/c3ay40566d Google Scholar
  118. 118.
    Tseng CM, De Diego A, Wasserman JC, Amouroux D, Donard OFX (1999) Chemosphere 39:1119–1136.  https://doi.org/10.1016/S0045-6535(99)00182-4 Google Scholar
  119. 119.
    Martin-Doimeadios RCR, Monperrus M, Krupp E, Amouroux D, Donard OFX (2003) Anal Chem 75:3202–3211.  https://doi.org/10.1021/ac026411a Google Scholar
  120. 120.
    Delgado A, Prieto A, Zuloaga O, de Diego A, Madariaga JM (2007) Anal Chim Acta 582:109–115.  https://doi.org/10.1016/j.aca.2006.08.051 Google Scholar
  121. 121.
    US EPA SW-846 Update V Mercury species fractionation and quantification by microwave assisted extraction, selective solvent extraction and/or solid phase extraction, method 3200, July 2014. https://www.epa.gov/sites/production/files/2015-12/documents/3200.pdf
  122. 122.
    Liu Y, Chai XL, Hao YX, Gao XF, Lu ZB, Zhao YC, Zhang J, Cai MH (2015) Environ Sci Pollut R 22:8603–8610.  https://doi.org/10.1007/s11356-014-3942-0 Google Scholar
  123. 123.
    He TR, Zhu YZ, Yin DL, Luo GJ, An YL, Yan HY, Qian XL (2015) Environ Sci Pollut R 22:5124–5138.  https://doi.org/10.1007/s11356-014-3864-x Google Scholar
  124. 124.
    Liang P, Lam CL, Chen Z, Wang HS, Shi JB, Wu SC, Wang WX, Zhang J, Wang HL, Wong MH (2013) J Soil Sediment 13:1301–1308.  https://doi.org/10.1007/s11368-013-0719-x Google Scholar
  125. 125.
    Schwartz GE, Redfern LK, Ikuma K, Gunsch CK, Ruhl LS, Vengosh A, Hsu-Kim H (2016) Environ Sci Proc Imp 18:1427–1439.  https://doi.org/10.1039/c6em00458j Google Scholar
  126. 126.
    Nevado JJB, Martin-Doimeadios RCR, Bernardo FJG, Moreno MJ (2008) Anal Chim Acta 608:30–37.  https://doi.org/10.1016/j.aca.2007.12.001 Google Scholar
  127. 127.
    Yin YG, Chen M, Peng JF, Liu JF, Jiang GB (2010) Talanta 81:1788–1792.  https://doi.org/10.1016/j.talanta.2010.03.039 Google Scholar
  128. 128.
    Turker AR, Cabuk D, Yalcinkaya O (2013) Anal Lett 46:1155–1170.  https://doi.org/10.1080/00032719.2012.753608 Google Scholar
  129. 129.
    Yang FF, Li JH, Lu WH, Wen YY, Cai XQ, You JM, Ma JP, Ding YJ, Chen LX (2014) Electrophoresis 35:474–481.  https://doi.org/10.1002/elps.201300409 Google Scholar
  130. 130.
    Gao ZB, Ma XG (2011) Anal Chim Acta 702:50–55.  https://doi.org/10.1016/j.aca.2011.06.019 Google Scholar
  131. 131.
    Chen BB, Wu YL, Guo XQ, He M, Hu B (2015) J Anal Atom Spectrom 30:875–881.  https://doi.org/10.1039/c4ja00312h Google Scholar
  132. 132.
    Pietila H, Peramaki P, Piispanen J, Majuri L, Starr M, Nieminen T, Kantola M, Ukonmaanaho L (2014) Microchem J 112:113–118.  https://doi.org/10.1016/j.microc.2013.10.002 Google Scholar
  133. 133.
    Taylor VF, Carter A, Davies C, Jackson BP (2011) Anal Methods UK 3:1143–1148.  https://doi.org/10.1039/c0ay00528b Google Scholar
  134. 134.
    Margetinova J, Houserova-Pelcova P, Kuban V (2008) Anal Chim Acta 615:115–123.  https://doi.org/10.1016/j.aca.2008.03.061 Google Scholar
  135. 135.
  136. 136.
    Pietila H, Peramaki P, Piispanen J, Starr M, Nieminen T, Kantola M, Ukonmaanaho L (2015) Chemosphere 124:47–53.  https://doi.org/10.1016/j.chemosphere.2014.11.001 Google Scholar
  137. 137.
    Mao YX, Yin YG, Li YB, Liu GL, Feng XB, Jiang GB, Cai Y (2010) Environ Pollut 158:3378–3384.  https://doi.org/10.1016/j.envpol.2010.07.031 Google Scholar
  138. 138.
    Leng G, Yin H, Li SB, Chen Y, Dan DZ (2012) Talanta 99:631–636.  https://doi.org/10.1016/j.talanta.2012.06.051 Google Scholar
  139. 139.
    Stanisz E, Werner J, Matusiewicz H (2013) Microchem J 110:28–35.  https://doi.org/10.1016/j.microc.2013.01.006 Google Scholar
  140. 140.
    Bravo-Sanchez LR, Encinar JR, Martinez JIF, Sanz-Medel A (2004) Spectrochim Acta B 59:59–66.  https://doi.org/10.1016/j.sab.2003.10.001 Google Scholar
  141. 141.
    Krystek P, Ritsema R (2004) Appl Organomet Chem 18:640–645.  https://doi.org/10.1002/aoc.697 Google Scholar
  142. 142.
    Stoichev T, Martin-Doimeadios RCR, Tessier E, Amouroux D, Donard OFX (2004) Talanta 62:433–438.  https://doi.org/10.1016/j.talanta.2003.08.006 Google Scholar
  143. 143.
    Gomez-Ariza JL, Lorenzo F, Garcia-Barrera T, Sanchez-Rodas D (2004) Anal Chim Acta 511:165–173.  https://doi.org/10.1016/j.aca.2004.01.051 Google Scholar
  144. 144.
    Centineo G, Gonzalez EB, Sanz-Medel A (2004) J Chromatogr A 1034:191–197.  https://doi.org/10.1016/j.chroma.2004.01.051 Google Scholar
  145. 145.
    Munoz J, Gallego M, Valcarcel M (2004) J Chromatogr A 1055:185–190.  https://doi.org/10.1016/j.chroma.2004.09.026 Google Scholar
  146. 146.
    Bloxham MJ, Gachanja A, Hill SJ, Worsfold PJ (1996) J Anal Atom Spectrom 11:145–148.  https://doi.org/10.1039/ja9961100145 Google Scholar
  147. 147.
    Ho YS, Uden PC (1994) J Chromatogr A 688:107–116.  https://doi.org/10.1016/S0021-9673(94)89019-6 Google Scholar
  148. 148.
    Sarzanini C, Sacchero G, Aceto M, Abollino O, Mentasti E (1994) Anal Chim Acta 284:661–667.  https://doi.org/10.1016/0003-2670(94)85070-4 Google Scholar
  149. 149.
    Bramanti E, Lomonte C, Onor M, Zamboni R, D’Ulivo A, Raspi G (2005) Talanta 66:762–768.  https://doi.org/10.1016/j.talanta.2004.12.031 Google Scholar
  150. 150.
    Wan CC, Chen CS, Jiang SJ (1997) J Anal Atom Spectrom 12:683–687.  https://doi.org/10.1039/a605765i Google Scholar
  151. 151.
    Ackley KL, Sutton KL, Caruso JA (2000) J Anal Atom Spectrom 15:1069–1073.  https://doi.org/10.1039/b000986p Google Scholar
  152. 152.
    Shum SCK, Pang HM, Houk RS (1992) Anal Chem 64:2444–2450.  https://doi.org/10.1021/ac00044a025 Google Scholar
  153. 153.
    Blanco RM, Villanueva MT, Uria JES, Sanz-Medel A (2000) Anal Chim Acta 419:137–144.  https://doi.org/10.1016/S0003-2670(00)01002-3 Google Scholar
  154. 154.
    Dong LM, Yan XP, Li Y, Jiang Y, Wang SW, Jiang DQ (2004) J Chromatogr A 1036:119–125.  https://doi.org/10.1016/j.chroma.2004.02.070 Google Scholar
  155. 155.
    Chen XP, Han C, Cheng HY, Wang YC, Liu JH, Xu ZG, Hu L (2013) J Chromatogr A 1314:86–93.  https://doi.org/10.1016/j.chroma.2013.08.104 Google Scholar
  156. 156.
    Tu Q, Qvarnstrom J, Frech W (2000) Analyst 125:705–710.  https://doi.org/10.1039/a908880f Google Scholar
  157. 157.
    Lee TH, Jiang SJ (2000) Anal Chim Acta 413:197–205.  https://doi.org/10.1016/S0003-2670(00)00807-2 Google Scholar
  158. 158.
    da Rocha MS, Soldado AB, Blanco-Gonzalez E, Sanz-Medel A (2000) J Anal Atom Spectrom 15:513–518Google Scholar
  159. 159.
    da Rocha MS, Soldado AB, Blanco-Gonzalez E, Sanz-Medel A (2000) Biomed Chromatogr 14:6–63Google Scholar
  160. 160.
    Medina I, Rubi E, Mejuto MC, Cela R (1993) Talanta 40:1631–1636.  https://doi.org/10.1016/0039-9140(93)80077-5 Google Scholar
  161. 161.
    Mercader-Trejo F, de San Miguel ER, de Gyves J (2005) J Anal Atom Spectrom 20:1212–1217.  https://doi.org/10.1039/b505000f Google Scholar
  162. 162.
    Mercader-Trejo F, Herrera-Basurto R, de San Miguel ER, de Gyves J (2011) Int J Environ Anal Chem 91:1062–1076.  https://doi.org/10.1080/03067311003782658 Google Scholar
  163. 163.
    Nguyen TH, Boman J, Leermakers M, Baeyens W (1998) X-ray Spectrom 27:277–282.  https://doi.org/10.1002/(Sici)1097-4539(199807/08)27:4%3C277::Aid-Xrs297%3E3.0.Co;2-U Google Scholar
  164. 164.
    Koplik R, Klimesova I, Malisova K, Mestek O (2014) Czech J Food Sci 32:249–259Google Scholar
  165. 165.
    Bulska E, Baxter DC, Frech W (1991) Anal Chim Acta 249:545–554.  https://doi.org/10.1016/S0003-2670(00)83032-9 Google Scholar
  166. 166.
    Tao H, Murakami T, Tominaga M, Miyazaki A (1998) J Anal Atom Spectrom 13:1085–1093.  https://doi.org/10.1039/a803369b Google Scholar
  167. 167.
    Uria JES, Sanz-Medel A (1998) Talanta 47:509–524Google Scholar
  168. 168.
    Harrington CF (2000) Trac Trend Anal Chem 19:167–179.  https://doi.org/10.1016/S0165-9936(99)00190-9 Google Scholar
  169. 169.
    Kadlecova M, Daye M, Ouddane B (2014) Anal Lett 47:697–706.  https://doi.org/10.1080/00032719.2013.848364 Google Scholar
  170. 170.
    Lambertsson L, Lundberg E, Nilsson M, Frech W (2001) J Anal Atom Spectrom 16:1296–1301.  https://doi.org/10.1039/b106878b Google Scholar
  171. 171.
    Park JS, Lee JS, Kim GB, Cha JS, Shin SK, Kang HG, Hong EJ, Chung GT, Kim YH (2010) Water Air Soil Pollut 207:391–401.  https://doi.org/10.1007/s11270-009-0144-3 Google Scholar
  172. 172.
    Caricchia AM, Minervini G, Soldati P, Chiavarini S, Ubaldi C, Morabito R (1997) Microchem J 55:44–55.  https://doi.org/10.1006/mchj.1996.1357 Google Scholar
  173. 173.
    Hintelmann H, Evans RD, Villeneuve JY (1995) J Anal Atom Spectrom 10:619–624.  https://doi.org/10.1039/ja9951000619 Google Scholar
  174. 174.
    Lin LY, Chang LF, Jiang SJ (2008) J Agric Food Chem 56:6868–6872.  https://doi.org/10.1021/jf801241w Google Scholar
  175. 175.
    de Souza SS, Rodrigues JL, Souza VCD, Barbosa F (2010) J Anal Atom Spectrom 25:79–83.  https://doi.org/10.1039/b911696f Google Scholar
  176. 176.
    Houserova P, Matejicek D, Kuban V (2007) Anal Chim Acta 596:242–250.  https://doi.org/10.1016/j.aca.2007.06.020 Google Scholar
  177. 177.
    Rahman GMM, Kingston HM (2004) Anal Chem 76:3548–3555.  https://doi.org/10.1021/Ac030407x Google Scholar
  178. 178.
    Rai R, Maher W, Kirkowa F (2002) J Anal Atom Spectrom 17:1560–1563.  https://doi.org/10.1039/b208041a Google Scholar
  179. 179.
    Tu Q, Johnson W, Buckley B (2003) J Anal Atom Spectrom 18:696–701.  https://doi.org/10.1039/b300992k Google Scholar
  180. 180.
    Kuban P, Houserova P, Kuban P, Hauser PC, Kuban V (2007) Electrophoresis 28:58–68.  https://doi.org/10.1002/elps.200600457 Google Scholar
  181. 181.
    Soliman EM, Saleh MB, Ahmed SA (2006) Talanta 69:55–60.  https://doi.org/10.1016/j.talanta.2005.08.070 Google Scholar
  182. 182.
    Jiang HM, Hu B, Jiang ZC, Qin YC (2006) Talanta 70:7–13.  https://doi.org/10.1016/j.talanta.2006.02.047 Google Scholar
  183. 183.
    Landi S, Fagioli F, Locatelli C (1992) J AOAC Int 75:1023–1028Google Scholar
  184. 184.
    Oda CE, Ingle JD (1981) Anal Chem 53:2305–2309.  https://doi.org/10.1021/Ac00237a040 Google Scholar
  185. 185.
    Leopold K, Foulkes M, Worsfold PJ (2009) Trac Trend Anal Chem 28:426–435.  https://doi.org/10.1016/j.trac.2009.02.004 Google Scholar
  186. 186.
    Logar M, Horvat M, Akagi H, Pihlar B (2002) Anal Bioanal Chem 374:1015–1021.  https://doi.org/10.1007/s00216-002-1501-x Google Scholar
  187. 187.
    Labatzke T, Schlemmer G (2004) Anal Bioanal Chem 378:1075–1082.  https://doi.org/10.1007/s00216-003-2416-x Google Scholar
  188. 188.
    Campbell MJ, Vermeir G, Dams R, Quevauviller P (1992) J Anal Atom Spectrom 7:617–621.  https://doi.org/10.1039/ja9920700617 Google Scholar
  189. 189.
    Seibert EL, Dressler VL, Pozebon D, Curtius AJ (2001) Spectrochim Acta B 56:1963–1971.  https://doi.org/10.1016/S0584-8547(01)00334-2 Google Scholar
  190. 190.
    Monperrus M, Tessier E, Veschambre S, Amouroux D, Donard O (2005) Anal Bioanal Chem 381:854–862.  https://doi.org/10.1007/s00216-004-2973-7 Google Scholar
  191. 191.
    Jia XY, Han Y, Liu XL, Duan TC, Chen HT (2011) Spectrochim Acta B 66:88–92.  https://doi.org/10.1016/j.sab.2010.12.003 Google Scholar
  192. 192.
    Jia XY, Gong DR, Han Y, Wei C, Duan TC, Chen HT (2012) Talanta 88:724–729.  https://doi.org/10.1016/j.talanta.2011.10.026 Google Scholar
  193. 193.
    Cheng HY, Wu CL, Shen LH, Liu JH, Xu ZG (2014) Anal Chim Acta 828:9–16.  https://doi.org/10.1016/j.aca.2014.04.042 Google Scholar
  194. 194.
    Nevado JJB, Martin-Doimeadios RCR, Krupp EM, Bernardo FJG, Farinas NR, Moreno MJ, Wallace D, Roper MJP (2011) J Chromatogr A 1218:4545–4551.  https://doi.org/10.1016/j.chroma.2011.05.036 Google Scholar
  195. 195.
    Zhao YQ, Zheng JP, Fang L, Lin Q, Wu YN, Xue ZM, Fu FF (2012) Talanta 89:280–285.  https://doi.org/10.1016/j.talanta.2011.12.029 Google Scholar
  196. 196.
    Li BH (2011) Anal Methods UK 3:116–121.  https://doi.org/10.1039/c0ay00480d Google Scholar
  197. 197.
    Trujillo IS, Alonso EV, Pavon JMC, de Torres AG (2015) J Anal Atom Spectrom 30:2429–2440.  https://doi.org/10.1039/c5ja00335k Google Scholar
  198. 198.
    Garcia-Ordiales E, Covelli S, Rico JM, Roqueni N, Fontolan G, Flor-Blanco G, Cienfuegos P, Loredo J (2018) Chemosphere 198:281–289.  https://doi.org/10.1016/j.chemosphere.2018.01.146 Google Scholar
  199. 199.
    Nevado JJB, Martin-Doimeadios RCR, Bernardo FJG, Moreno MJ (2005) J Chromatogr A 1093:21–28.  https://doi.org/10.1016/j.chroma.2005.07.054 Google Scholar
  200. 200.
    Cai Y, Monsalud S, Furton KG, Jaffe R, Jones RD (1998) Appl Organomet Chem 12:565–569.  https://doi.org/10.1002/(Sici)1099-0739(199808/09)12:8/9%3C565::Aid-Aoc762%3E3.0.Co;2-K Google Scholar
  201. 201.
    Nevado JJB, Martin-Doimeadios RCR, Moreno MJ (2009) Sci Total Environ 407:2372–2382.  https://doi.org/10.1016/j.scitotenv.2008.12.006 Google Scholar
  202. 202.
    Martin-Doimeadios RCR, Krupp E, Amouroux D, Donard OFX (2002) Anal Chem 74:2505–2512.  https://doi.org/10.1021/ac011157s Google Scholar
  203. 203.
    Rodrigues JL, Alvarez CR, Farinas NR, Nevado JJB, Barbosa F, Martin-Doimeadios RCR (2011) J Anal Atom Spectrom 26:436–442.  https://doi.org/10.1039/c004931j Google Scholar
  204. 204.
    Moreno MJ, Pacheco-Arjona J, Rodriguez-Gonzalez P, Preud’Homme H, Amouroux D, Donard OFX (2006) J Mass Spectrom 41:1491–1497.  https://doi.org/10.1002/jms.1120 Google Scholar
  205. 205.
    Sanchez-Rodas D, Corns WT, Chen B, Stockwell PB (2010) J Anal Atom Spectrom 25:933–946.  https://doi.org/10.1039/b917755h Google Scholar
  206. 206.
    Cano-Pavon JM, De Torres AG, Sanchez-Rojas F, Canada-Rudner P (1999) Int J Environ Anal Chem 75:93–106.  https://doi.org/10.1080/03067319908047303 Google Scholar
  207. 207.
    Heumann KG, Gallus SM, Radlinger G, Vogl J (1998) Spectrochim Acta B 53:273–287.  https://doi.org/10.1016/S0584-8547(97)00134-1 Google Scholar
  208. 208.
    Castillo A, Roig-Navarro AF, Pozo OJ (2006) Anal Chim Acta 577:18–25.  https://doi.org/10.1016/j.aca.2006.06.024 Google Scholar
  209. 209.
    Guo W, Hu SH, Wang XJ, Zhang JY, Jin LL, Zhu ZL, Zhang HF (2011) J Anal Atom Spectrom 26:1198–1203.  https://doi.org/10.1039/c1ja00005e Google Scholar
  210. 210.
    Jian L, Goessler W, Irgolic KJ (2000) Fresen J Anal Chem 366:48–53.  https://doi.org/10.1007/s002160050010 Google Scholar
  211. 211.
    Brombach CC, Chen B, Corns WT, Feldmann J, Krupp EM (2015) Spectrochim Acta B 105:103–108.  https://doi.org/10.1016/j.sab.2014.09.014 Google Scholar
  212. 212.
    Guzman-Mar JL, Hinojosa-Reyes L, Serra AM, Hernandez-Ramirez A, Cerda V (2011) Anal Chim Acta 708:11–18.  https://doi.org/10.1016/j.aca.2011.09.037 Google Scholar
  213. 213.
    Ai X, Wang Y, Hou XD, Yang L, Zheng CB, Wu L (2013) Analyst 138:3494–3501.  https://doi.org/10.1039/c3an00010a Google Scholar
  214. 214.
    Harrington CF, Romeril J, Catterick T (1998) Rapid Commun Mass Spectrom 12:911–916.  https://doi.org/10.1002/(Sici)1097-0231(19980731)12:14%3C911::Aid-Rcm254%3E3.0.Co;2-X Google Scholar
  215. 215.
    Pena-Pereira F, Lavilla I, Bendicho C, Vidal L, Canals A (2009) Talanta 78:537–541.  https://doi.org/10.1016/j.talanta.2008.12.003 Google Scholar
  216. 216.
    Li PJ, He M, Chen BB, Hu B (2015) J Chromatogr A 1415:48–56.  https://doi.org/10.1016/j.chroma.2015.08.062 Google Scholar
  217. 217.
    Chen C, Peng MT, Hou XD, Zheng CB, Long Z (2013) Anal Methods UK 5:1185–1191.  https://doi.org/10.1039/c2ay26214b Google Scholar
  218. 218.
    Li PJ, Zhang X, Hu B (2011) J Chromatogr A 1218:9414–9421.  https://doi.org/10.1016/j.chroma.2011.10.071 Google Scholar
  219. 219.
    Ting Y, Chen C, Ch’ng BL, Wang YL, Hsi HC (2018) J Hazard Mater 354:116–124.  https://doi.org/10.1016/j.jhazmat.2018.04.074 Google Scholar
  220. 220.
    Gilmour C, Bell JT, Soren AB, Riedel G, Riedel G, Kopec AD, Bodaly RA (2018) Sci Total Environ 640:555–569.  https://doi.org/10.1016/j.scitotenv.2018.05.276 Google Scholar
  221. 221.
    Blum PW, Hershey AE, Tsui MTK, Hammerschmidt CR, Agather AM (2018) Biogeochemistry 137:181–195.  https://doi.org/10.1007/s10533-017-0408-8 Google Scholar
  222. 222.
    Cesario R, Hintelmann H, Mendes R, Eckey K, Dimock B, Araujo B, Mota AM, Canario J (2017) Environ Pollut 226:297–307.  https://doi.org/10.1016/j.envpol.2017.03.075 Google Scholar
  223. 223.
    Valdes C, Black FJ, Stringham B, Collins JN, Goodman JR, Saxton HJ, Mansfield CR, Schmidt JN, Yang S, Johnson WP (2017) Environ Sci Technol 51:4887–4896.  https://doi.org/10.1021/acs.est.6b05790 Google Scholar
  224. 224.
    Kodamatani H, Maeda C, Balogh SJ, Nollet YH, Kanzaki R, Tomiyasu T (2017) Chemosphere 173:380–386.  https://doi.org/10.1016/j.chemosphere.2017.01.053 Google Scholar
  225. 225.
    Cesario R, Monteiro CE, Nogueira M, O’Driscoll NJ, Caetano M, Hintelmann H, Mota AM, Canario J (2016) Water Air Soil Pollut 227:475.  https://doi.org/10.1007/s11270-016-3179-2 Google Scholar
  226. 226.
    Mendes LA, de Lena JC, do Valle CM, Fleming PM, Windmoller CC (2016) Appl Geochem 75:32–43.  https://doi.org/10.1016/j.apgeochem.2016.10.011 Google Scholar
  227. 227.
    Monteiro CE, Cesario R, O’Driscoll NJ, Nogueira M, Valega M, Caetano M, Canario J (2016) Mar Pollut Bull 104:162–170.  https://doi.org/10.1016/j.marpolbul.2016.01.042 Google Scholar
  228. 228.
    Liu B, Schaider LA, Mason RP, Shine JP, Rabalais NN, Senn DB (2015) Estuar Coast Shelf Sci 159:50–59.  https://doi.org/10.1016/j.ecss.2015.03.030 Google Scholar
  229. 229.
    Ma X, Yin YG, Shi JB, Liu JF, Jiang GB (2014) Anal Methods UK 6:164–169.  https://doi.org/10.1039/c3ay41625a Google Scholar
  230. 230.
    Kodamatani H, Tomiyasu T (2013) J Chromatogr A 1288:155–159.  https://doi.org/10.1016/j.chroma.2013.02.004 Google Scholar

Copyright information

© Springer-Verlag GmbH Germany, part of Springer Nature 2018

Authors and Affiliations

  1. 1.Applied Analytical ChemistryUniversity Duisburg-EssenEssenGermany

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