Single Particle-Inductively Coupled Plasma Mass Spectroscopy

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Technical Note

SP-ICP-MS Analysis of Metallic Nanoparticles in Environmental Samples with Large Dissolved Analyte Fractions Dina M. Schwertfeger, Jessica R. Velicogna, Alexander H. Jesmer, Richard P. Scroggins, and Juliska I. Princz Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.6b02716 • Publication Date (Web): 15 Sep 2016 Downloaded from http://pubs.acs.org on September 15, 2016

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Analytical Chemistry

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Title: SP-ICP-MS Analysis of Metallic Nanoparticles in Environmental Samples

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with Large Dissolved Analyte Fractions

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Authors: D. M. Schwertfeger, Jessica R. Velicogna, Alexander H. Jesmer, Richard P. Scroggins,

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Juliska I. Princz*

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Affiliations: Biological Assessment and Standardization, Environment and Climate Change

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Canada, 335 River Road South, Ottawa, Ontario K1V 1C7 Canada

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* To whom correspondence may be addressed ([email protected])

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Juliska Princz, Ph.D.

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Soil Toxicology Laboratory, Biological Assessment and Standardization

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Environment and Climate Change Canada

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335 River Road

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Ottawa (Ontario) K1V 1C7

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Email:

[email protected]

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Telephone:

613-990-9544

ACS Paragon Plus Environment


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Abstract

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There is an increasing interest to use SP-ICP-MS to help quantify exposure to engineered

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nanoparticles, and their transformation products, released into the environment. Hindering the

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use of this analytical technique for environmental samples is the presence of high levels of

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dissolved analyte which impedes resolution of the particle signal from the dissolved. While

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sample dilution is often necessary to achieve the low analyte concentrations necessary for SP-

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ICP-MS analysis, and to reduce the occurrence of matrix effects on the analyte signal, it is used

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here to also reduce the dissolved signal relative to the particulate, while maintaining a matrix

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chemistry that promotes particle stability. We propose a simple, systematic dilution series

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approach where by the first dilution is used to quantify the dissolved analyte, the second is used

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to optimize the particle signal, and the third is used as an analytical quality control. Using simple

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suspensions of well characterized Au and Ag nanoparticles spiked with the dissolved analyte

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form, as well as suspensions of complex environmental media (i.e., extracts from soils

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previously contaminated with engineered silver nanoparticles), we show how this dilution series

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technique improves resolution of the particle signal which in turn improves the accuracy of

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particle counts, quantification of particulate mass and determination of particle size. The

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technique proposed here is meant to offer a systematic and reproducible approach to the SP-ICP-

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MS analysis of environmental samples and improve the quality and consistency of data

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generated from this relatively new analytical tool.

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INTRODUCTION

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In order to evaluate the risk of engineered nanoparticles (ENPs) released into the environment, it

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is necessary to perform quantitative assessments of both the exposure and effects to non-target


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organisms in environmental media.1,2 Single-particle inductively-coupled plasma mass

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spectroscopy (SP-ICP-MS) has been identified as a valuable tool in this regard, given its ability

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to detect and quantify the elemental composition of nanoparticles in suspensions at

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environmentally relevant concentrations (i.e., ng L-1).3 However, SP-ICP-MS is not without its

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shortcomings. One difficulty that arises in SP-ICP-MS, particularly for the analysis of

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environmental media, occurs when dissolved analyte concentrations are large relative to the

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analyte mass in the nanoparticles, which can lead to erroneous particle information.4

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For SP-ICP-MS, samples must be sufficiently dilute such that, ideally, one particle at a time is

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ionized in the plasma, which generates a burst of ions that enter the mass spectrometer and result

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in a spike of the intensity signal, often referred to as pulse intensity. In contrast, the dissolved

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analyte will produce a constant, continuous signal because of the steady stream of ions entering

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the plasma. The intensity of a particle is essentially the pulse intensity after subtracting the

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intensity of the dissolved or background signal, which can then be related to the mass of the

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particle via calibration with well characterized nanoparticles, or via dissolved analyte calibration.

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5

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fraction of the analyte within the particle are known. Particle concentration (i.e., particles per

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mL) is determined by counting the number of pulses detected and accounting for operational

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factors that affect the rate of ions reaching the detector, such as sample flow rate and transport

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efficiency (i.e., the fraction of nebulized particles that reach the plasma). An overview of SP-

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ICP-MS theory and related calculations are provided in Laborda et al.,6 while an evaluation of

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transport efficiency calculations is provided in Pace et al.5

Particle size is calculated from its mass if the analyte density, particle geometry and mass

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One of the key aspects in processing SP-ICP-MS data is distinguishing the pulse intensity of

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the particle from the dissolved signal. The intensity threshold used to discriminate the two



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analyte forms is often determined using an iterative computation.4 An approximate threshold

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limit is established, above which, events will be removed (e.g., events with intensities above the

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mean intensity + 3δ). This initial calculation usually removes the majority of particle events. Of

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the remaining events, the mean intensity is re-calculated and the procedure is repeated until

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convergence (i.e., no more events can be removed). The final intensity threshold can be

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converted to analyte mass, or particle volume, to determine the detection limits for particulate

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mass or particle size, respectively. Some suppliers of SP-ICP-MS instruments are now supplying

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software which performs these computations automatically for each sample.

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A difficulty in the computation used to resolve particle intensity from the dissolved signal

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occurs when there is a high level of dissolved analyte. When this occurs, the dissolved signal

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overlaps the particle signal, resulting in the computation of a threshold intensity that is larger

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than the intensities of some (or potentially all) the particles in a sample. For example, when an

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aqueous suspension of Ag nanoparticles (60 nm nominal size) was spiked with increasing

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concentrations of dissolved Ag, the number of particles detected decreased and the particle size

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detection limit increased.4

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There are many instances where environmental samples are expected to contain high levels of

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the dissolved analyte. This may occur if the analyte of interest naturally occurs in the

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environment in the form of dissolved ionic or complexed species (i.e., geologic or pedogenic in

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origin), or if ENPs released into the environment undergo weathering and transformation to

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dissolved species during their translocation to, or residence in, the media being tested.

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Few researchers have developed reliable techniques to address the problem of analyzing

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environmental samples with large dissolved analyte fractions. Hadioui and colleagues4

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developed a technique whereby samples were first passed through an exchange column to


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remove much of the dissolved Ag+ species from samples prior to introduction into the ICP-MS.

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This technique resulted in more accurate sizing and counting of the nanoparticles in their

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suspensions and was further applied to the analysis of ZnO nanoparticles in wastewater.7 While

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effective, the procedure adds a considerable amount of time to the analysis. This may be a

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reasonable trade-off for suspensions containing NPs that are highly unstable and readily dissolve,

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but may not be practical for all scenarios.

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In this study, we investigate a sample dilution technique, which is ideally suited to the

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analysis of suspensions that are relatively stable, such as soil and biosolids extracts. The

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technique involves diluting samples in order to reduce the dissolved signal, thereby increasing

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the ability to resolve the particle signal. While sample dilution for SP-ICP-MS is not a novel

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concept, it is often done haphazardly from one sample to the next with the focus being to reduce

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the occurrence of analytical coincidence (i.e., two or more particles entering the plasma at once).

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Unlike standard ICP-MS analysis for which well-established technical guidance have been

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published (e.g., EPA 6020A, 200.8), SP-ICP-MS is still in its infancy and lacks supportive

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technical guidance and implementation of consistent quality controls, particularly for the analysis

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of complex environmental matrices. We propose the use of a simple dilution series wherein each

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successful dilution achieves a specific goal: (1) the capture of a dissolved analyte reading above

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the limit of quantitation (LoQ) for the dissolved signal, (2) the capture of an optimized particle

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signal, and (3) a quality control check that the particle analysis is truly optimized and free of

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coincidence and matrix effects.

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For this study, Au and Ag nanomaterials were selected. Citrate-stabilized Au nanoparticles

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are highly stable and thus allowed us to explore the dilution technique without any interfering

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effects of particle dissolution. Silver nanoparticles are one of the most widely used engineered



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nanomaterials and their release from consumer products into the wastewater stream and

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subsequent accumulation in sewage sludge represent a major translocation pathway.8 In North

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America, processed sewage sludge (i.e., biosolids) is applied to agricultural lands representing an

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important ecotoxicological exposure scenario. As such, governing agencies have initiated

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assessments on the fate and effects of ENPs released into the terrestrial environment.9 The

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overall goal of this study is to help further the development of a systematic, standardized

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approach to SP-ICP-MS analysis of environmental samples (e.g., soil and biosolids) which is

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expected to improve the quality of data generated by this relatively new analytical technique.

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EXPERIMENTAL SECTION

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Chemical. All Au and Ag nanoparticles used in this study were commercially purchased and

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are listed in Table 1, along with suppliers’ specifications. Verification of these specifications was

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carried out using dynamic light scattering (DLS), transmission electron microscopy (TEM) with

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energy-dispersive X-ray spectroscopy (EDS), and SP-ICP-MS. All purchased suspensions were

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placed in an ultrasonic bath (Branson 3510, 40 kHz, 130 watts, 6 L capacity) for at least 20 min

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prior to analysis. In the case of the Ag nano-powder, a 15 g L-1 dispersion was prepared using

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NanopureTM water (Barnstead 2-cartridge water filtration system, > 18.3 MΩ·cm, TOC < 10 µg

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L-1) and placed in the ultrasonic bath for 40 min. The DLS analysis was conducted on ca. 20 mg

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L-1 dilutions of initial suspensions using a MalvernTM Nano ZS Zetasizer with NIBS® optics

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arranged at 175° for backscatter detection. Data was analyzed using cumulants analysis.10

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Imaging of the same dispersion was conducted using an FEITM Tecnai G2 F20 TEM (FEI,

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Hillsboro OR) at the Carleton University Nano Imaging Facility (Ottawa, Ontario). Details of the


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SP-ICP-MS analysis are provided in the relevant section below. The Au and Ag nanomaterials

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are hereafter referred to by their nominal sizes.

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Test Samples. Two types of samples were analyzed in this study: (1) suspensions of gold

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nanoparticles (AuNP) or silver nanoparticles (AgNP) prepared in NanopureTM water, and (2) soil

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extracts prepared from AgNP-contaminated soils. The nanoparticle suspensions were prepared

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with 30 nm AuNP (NIST 8012), 60 nm AuNP (8013) or 30 nm AgNP (Ted Pella). For each

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material used, a set of suspensions were prepared for which the particle concentration was kept

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constant at approximately 105 particles mL-1, while increasing the dissolved Au or Ag

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concentrations. Stock solutions of dissolved Au and Ag were prepared with an Au standard

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solution (High-Purity Standards, 1000 µg mL-1 in 2% HCl) and silver nitrate (Sigma Aldrich

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Reagent Plus®), respectively. Dissolved analyte concentrations ranged from 0.1 to 10 µg L-1 for

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the suspensions containing the 30 nm particles, and 0.1 to 40 µg L-1 for suspensions containing

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the 60 nm particles. Dilutions of test samples were always prepared fresh using NanopureTM

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water and analyzed immediately. The soil extracts analyzed in this study were used to

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demonstrate the dilution series approach to SP-ICP-MS analysis on real environmental samples

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and were taken from a soil previously amended with AgNP-contaminated biosolids. Details of

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the method development of the soil extraction technique are not critical to the understanding of

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how to optimize the SP-ICP-MS analysis presented here in, and are outside the scope of this

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study; however, further details have been included in a manuscript being prepared by the authors.

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Soil preparation and the soil extraction procedures are briefly described as follows: An aliquot of

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a 15 g L-1 suspension of the 40 nm AgNP (Nano-composix) was mixed into a slurry of municipal

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biosolids, aged 3 d in sealed containers, then incorporated into an agricultural field soil (sandy

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loam, pH 5.8) at a rate of 1 g biosolids per 100 g soil (dry weights) to obtain various Ag



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concentrations. The soil samples used in this study included a soil sample spiked with 833 µg g-1

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Ag taken from an earthworm assay (nAgEA), a soil sample spiked with 833 µg g-1 Ag taken

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from a microbial assay (nAgMicr), and a soil sample spiked with 145 µg g-1 Ag taken from a

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collembolan assay (nAgFC). Soils were extracted with tetrasodiumpyrophosphate (2.5 mM

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TSPP) using a soil-to-reagent ratio of 1:10 (g:mL) and mixed on a rotary mixer for 30 min

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followed by ultrasonication. Suspensions were allowed to settle overnight to allow large particles

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to settle out. Sample suspensions were initially diluted 100x and all subsequent dilutions were

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maintained with a 0.025 mM TSPP matrix.

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SP-ICP-MS Instrumentation. SP-ICP-MS analysis was performed using a PerkinElmer

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NexION 300D ICP-MS operating in single particle mode using the SyngistixTM Nano-

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Application module (PerkinElmer Inc., Waltham, MA). Instrumental and data acquisition

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parameters of the analysis are provided in Table 2. Time resolved signals were acquired using

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fast-scanning mode, for which settling times have been reduced to negligible. Due to the fast

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data acquisition rate, dwell times as low as 10 µs can be achieved. Previous research has shown

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that dwell times of 50 to 100 µs are optimal for our instrumental set-up and for the analysis of

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nanoparticles similar to ours.11 Transport efficiency was determined by the particle size method5

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using the 60 nm NIST Au standard for reference; all calculations were performed within the

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SyngistixTM module. Dissolved and particle calibrations were performed for

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analytes, with dissolved calibrations ranging from 0.1 to 10 µg L-1. Dissolved Au and Ag

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calibration solutions were prepared in 0.5% HCl and 1% HNO3 matrices, respectively. Gold

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particle calibrations were performed using NIST Au 30 and 60 nm suspensions while for the Ag

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particle calibrations, Ted Pella Ag 30 and 60 nm suspensions were used. For particle

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calibrations, the intensity histograms were fitted to a log normal curve from which the most


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Au and

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Ag

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frequent area intensity was derived and related to particle mass (i.e., size). The dissolved and

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particle calibrations produced comparable particle size data with mean particle intensities

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derived from the dissolved calibration being on average 9% lower. The particle calibration

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method was selected to process the results of this study, as the un-acidified matrix was more

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consistent with that of the samples. SP-ICP-MS analysis requires that samples are sufficiently

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dilute such that analytical coincidence is avoided. Based on our operating parameters, we

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estimated that coincidence could be avoided if sample particle concentrations were ≤ 1.2 x 105

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mL-1; therefore target particle concentrations for calibration suspensions were approximately 105

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particles mL-1. The method detection limit (MDL) and LoQ for the dissolved analysis were

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calculated as the mean reading of six method blanks (i.e., 0.025 mM TSPP solution) plus 3δ and

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10δ, respectively.

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Data Processing. Initial data processing was performed within the SyngistixTM Nano-

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application module. Assuming a particle event in the plasma is approximately 350 µs,11 the 50 µs

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dwell time used allows for multiple readings per particle. An algorithm is used to identify when a

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succession of readings comprise a particle event from which peak area is calculated. In this way,

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particle intensity is related to peak area. The determination of the intensity threshold used to

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discriminate whether a signal is part of a particle event, or the dissolved (i.e., background) signal

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is also performed within the SyngistixTM module using an iterative “mean + 3δ” computation,

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similar to that described by Hadioui et al.4 The software records the number of particle events

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and produces a frequency distribution based on particle intensity (i.e., peak area). Particle

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intensity was related to particle mass (and thus size) using a particle calibration curve derived

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from analyzing the 30 and 60 nm nanoparticle suspensions (Au or Ag) based on the assumption

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that Au and Ag particles were spherical and pure Au0 or Ag0 with a density of 19.32 and 10.49 g



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cm-3, respectively. Particle size distributions (PSD) were fit to a Gaussian smoothing function

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from which mean and mode particle sizes were derived. It must be noted that while the particle

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geometry and mass fraction may be known for well characterized particles (i.e., test samples

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prepared from purchased suspensions), this is usually not the case for particles in environmental

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media, for which a plasma event may include transformation products of various composition, or

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naturally occurring particles. Therefore, for particles detected in environmental media, only the

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mass of the analyte detected in a discrete particle (i.e., particulate Ag) is detectable, the size of

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the actual particle is unknown. However, for the analysis of particulate Ag in our soil extracts,

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we processed the data using a default spherical shape and mass fraction of 1, in order to calculate

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an equivalent mono-metal particle size, denoted with the subscript “eqv” (e.g., PSDeqv), indicative,

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not of actual particle sizes, but reference sizes used to aid in the understanding of the amount of

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mass detected.

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RESULTS AND DISCUSSION

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Particle sizes of the Au and Ag nanomaterials were verified by DLS (Table S1, Figure S1,

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Supporting Information) as well as by TEM/EDS and SP-ICP-MS (selected data, Figures S2 and

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S3, Supporting Information). Measured particle sizes generally fell within the suppliers’ size

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specifications, except for the Nanocomposix 40 nm Ag particles which showed a minor fraction

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of particles smaller than 19 nm.

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To investigate the effects of the SP-ICP-MS particle signal when samples contain large

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concentrations of the dissolved metal, nanoparticle suspensions containing ca. 105 particles mL-1

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were spiked with increasing amounts of dissolved metal and analyzed. Figure S4 (Supporting

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Information) shows a portion of the time-resolved Au signal during analysis of the 30 nm


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suspensions with and without dissolved Au added. When analyzing the suspension spiked with

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the dissolved Au, the steady dissolved signal drowns out most of the particle signals, resulting in

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fewer particles detected of relatively large intensities, likely agglomerates or artifacts of

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coincidence. Particle intensities were converted to particle size for which frequency histograms

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were derived. Figure 1 shows the histograms generated from analyzing the 30 nm Au suspension

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with increasing amounts of dissolved Au. Adding 0.5 µg L-1 dissolved Au had little effect on the

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size distribution, while the addition of 1, 2 and 3 µg L-1 dissolved Au significantly lowered the

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number of detectable particles while skewing particle sizes to increasingly larger sizes. This

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trend is clearly illustrated with both 30 nm and 60 nm Au suspensions, in Figure 2, which shows

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that at low concentrations of dissolved Au, the detected particle sizes and counts were accurate

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and constant, but above a certain threshold, increasing concentrations of dissolved Au led to

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erroneous increases to particle size and decreases in the number of particles detected. For the 30

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nm Au suspensions, this threshold of dissolved Au is between 0.5 and 1.0 µg L-1, while for the

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60 nm Au suspensions, it was between 10 and 20 µg L-1. When suspensions containing the

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highest dissolved Au additions tested, i.e., 30 nm + 3 µg L-1 dissolved Au and 60 nm + 40 µg L-1

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dissolved Au, were diluted such that the dissolved concentrations were below these thresholds,

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particle size and particle counts were measured more accurately (Figure 2). The reason for this

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improvement is demonstrated by comparing the particle histograms of the 30 nm + 3 µg L-1

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dissolved Au sample before, and after dilution (Figures 1e and 1f). For the diluted sample, the

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dissolved signal was sufficiently reduced to allow for the detection of the particles, thereby

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resolving the sample’s particle size histogram. A similar trend was observed for suspensions

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made from the 30 nm Ag nanoparticles for which a dissolved Ag threshold between 0.5 and 1.0

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µg L-1 was also observed. (Figure S5, Supporting Information). These data show that the



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dissolved analyte had a similar effect in magnitude on particle data for these similarly-sized

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particles, despite being different elements.

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When analyzing environmental samples, the particle sizes as well as particulate and dissolved

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concentrations are generally unknown. Because the dissolved analyte fraction is an important

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parameter for environmental assessments, we began with obtaining dissolved analyte readings

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within the calibration range and above the LoQ of 0.1 μg L-1, diluting the samples if necessary.

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For the three environmental samples analyzed in this study, only the nAgMicr extract required an

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initial dilution (i.e., 100x) to achieve a valid dissolved reading (Table 3). With knowledge of the

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dissolved fraction, the next analysis involved diluting the sample to achieve a dissolved analyte

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concentration near the MDL of 0.05 μg L-1, thereby maximizing the ratio of particle signal /

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background (dissolved) signal. This “optimal” dilution also needed to provide a particle

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concentration below the (coincidence) criteria of 105 particles mL-1. When applying this dilution

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technique to the analysis of soil extracts taken from ecotoxicological assays, a third dilution that

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was 2x the optimal dilution proved advantageous as a quality control. If the dilution selected for

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the “optimal” analysis was truly optimal, then diluting it 2x should yield a similar particle

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intensity (i.e., size) with half the number of particles counted. For example, analysis of the soil

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extract from the earthworm assay (nAgEA) showed 5.05 µg L-1 dissolved Ag and generated an

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equivalent particle size distribution (PSDeqv) with a mode of 68.2 nm (Table 3). The optimal

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dilution for the analysis of this sample was 100x, which reduced the dissolved signal to near the

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MDL and provided a particle count < 105 particles mL-1, resulting in a PSDeqv with a mode of

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37.4 nm (Figure 3a). When this optimal dilution was further diluted 2x, the resulting PSDeqv

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showed a similar mode, but approximately half the number of particles, as expected, thus


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providing confidence that the analysis performed on the “optimal” dilution selected was indeed

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not significantly impacted by excess dissolved analyte, analytical coincidence, nor matrix effects.

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For the analysis of the soil extract from the microbial assay, the initial 100x dilution captured

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a dissolved reading of 0.48 µg L-1 (Table 3). While we would not expect this dissolved

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concentration to impact the particle signal (see Figure S5, Supporting Information), the particle

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concentration detected at this dilution exceeded 105 particles mL-1, thus resulting in a high

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probability for analytical coincidence. An optimal dilution of 2500x was determined which

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shifted the PSDeqv left (Figure 3), resulting in the smaller mode of 32.7 nm and a greater overall

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particle concentration, evidence that analytical coincidence likely affected the analysis of the

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100x dilution. A further 2x dilution reduced the number of detectable particles to ca. 700 (data

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not shown). However, we have found that when the event sample size drops much below

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approximately 1000 events, there is greater variability among SP-ICP-MS data from analytical

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replicates. Extending the sample time to 120 s allowed for a larger sample size of ca. 1400

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events, which allowed for more reproducible data, and yielded a PSDeqv which generally

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overlapped that from the optimal dilution, as would be expected given the operational

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parameters. This technique of increasing sample time to increase the number of particle events

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can be particularly useful when analyzing a sample with a large ratio of dissolved analyte /

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particle counts, for which the technique of diluting the sample to reduce the dissolved signal

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yields a sample with a very low particle count.

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Similar to the previous samples, dilution of nAgFC shifted the initial PSDeqv left (Figure 3c),

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resulting in a lower mode of 40.6 nm. However, a further 2x dilution (with the extended sample

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time of 120 s), revealed a bimodal PSDeqv. This sample was re-run several times to verify this

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bimodal pattern and data from duplicate analyses were combined. Given that over 30% more



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particulate Ag mass was detected from the analysis of the most dilute sample compared to the

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next more concentrated sample (Table 3), it is likely that the detection of the bimodal distribution

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was due to an improved signal optimization (i.e., lower particle DL), rather than from dissolution

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of the 40 nm particles. Each of the two distributions was fit to a log-normal curve separately to

293

determine their equivalent mode particle sizes which were 28.7 and 43.4 nm. While other

294

analytical techniques are required to further identify/characterize particles in these two distinct

295

size classes (e.g., Ag2S, AgNP with surface precipitate), the example here illustrates the

296

usefulness of checking the data acquired from what was believed to be the “optimal” dilution by

297

analyzing the sample with a further 2x dilution.

298

The most obvious drawback to analyzing environmental samples using a dilution series is the

299

potential for particle dissolution. While research has shown that AgNPs in distilled water likely

300

undergo fairly rapid dissolution in the short-term (i.e., < 24 h),12 AgNPs contained in more

301

complex matrices, particularly in the presence of natural organic matter, are more stable,13,14 with

302

humic substances likely providing both steric and charge stabilization. Mitrano et al.14 showed

303

that the presence of ligands, such as chloride and sulfide, also had a stabilizing effect on AgNPs,

304

and suggested the formation of relatively insoluble halide-Ag or sulfide-Ag precipitates on the

305

particle surface. Thus environmental samples containing ENPs, and/or their transformation

306

products may very well be stabilized by their solution chemistry enough to carry out dilution and

307

analysis in the very short-term (i.e., minutes) without significant changes to analyte mass of the

308

particulates. It would be useful to perform dilution-stability tests on samples of various complex

309

matrices to gain information regarding the rate of change that dilution may have on particle

310

stability. The dilution approach shown here may be more suited for soil extracts or wastewaters

311

which generally have a large humic component. In the case of soil analysis, all extractions (e.g.,


Page 15 of 23

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15 312

soil pore water) and preparatory procedures are by necessity operationally defined, thus, it would

313

be advantageous to utilize an extraction technique that is amenable to SP-ICP-MS analysis. We

314

have found that the alkaline and dispersing attributes of TSPP result in large particle/dissolved

315

concentration ratios and stable extracts. For some environmental samples, it may be possible to

316

inhibit, or retard particle dissolution by maintaining the ionic strength of the environmental

317

sample during the dilution, (i.e., using a dilute Na solution) and/or adding a dispersant or

318

surfactant which may offer some steric stability. In our case, we maintained the TSPP

319

concentration of 0.025 mM in all dilutions.

320

Conclusions

321

For SP-ICP-MS analysis of environmental samples affected by engineered nanoparticle

322

contamination, quantification of both the dissolved and nano-particulate forms of the metal

323

analyte are of interest; however, it is doubtful that one dilution/analysis can optimally capture

324

both signals. As we have shown, a large dissolved signal can drown-out the particle signal

325

leading to erroneous sizing and particle counts. The degree to which an analysis will be

326

negatively impacted by presence of the dissolved analyte depends on both the particle size and

327

dissolved analyte concentration. The approach demonstrated here relies on analyzing samples in

328

a dilution series such that (1) an analysis captures the dissolved signal within the calibrated range

329

and above the LoQ, (2) an analysis captures an optimal particle signal, and (3) a third analysis is

330

used to verify the optimal particle data. An optimal particle signal can be achieved by diluting

331

the sample such that the dissolved signal is at or below the MDL and the particle concentration is

332

< 105 particles mL-1, to minimize the probability of analytical coincidence. Sampling time may

333

be extended to ensure sufficient number of events (i.e., > 1000) to achieve a suitable level of

334

statistical confidence when generating the particle frequency histograms and for ensuring



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Page 16 of 23

16 335

sufficient reproducibility. For the verification analysis, a 2x dilution of the optimal sample

336

dilution is recommended. The verification analysis ensures that interferences from the dissolved

337

signal, sample matrix effects and analytical coincidence have been sufficiently minimized. While

338

humic substances contained in environmental samples are likely to help stabilize particles and

339

buffer any rapid changes to particles that may be caused by the dilution process, further research

340

on how to preserve and stabilize environmental samples for SP-ICP-MS analysis is required. The

341

dilution technique demonstrated here offers a systematic and reproducible approach to the

342

analysis of environmental samples which is anticipated to improve the quality of data generated

343

by this relatively new analytical tool.

344 345 346 347

Associated Content

348

Additional information as noted in the text.

349

Acknowledgements

350

The research was funded by Environment and Climate Change Canada through the Chemicals

351

Management Plan.

352

References

353 354 355 356 357 358 359 360 361 362 363

(1) von der Kammer, F.; Ferguson, P. L.; Holden, P. A.; Masion, A.; Rogers, K. R.; Klaine, S. J.; Koelmans, A. A.; Horne, N.; Unrine, J. M. Environ. Toxicol. Chem. 2012, 31, 32-49. (2) Montano, M. D.; Ranville, J. F.; Lowry, G. V.; Lowry, G. V.; Von Der Kammer, F.; Blue, J. Environ. Chem. 2014, 11, 351-366. (3) Hassellöv, M.; Readman, J. W.; Ranville, J. F.; Tiede, K. Ecotoxicol. 2008, 17, 344-361. (4) Hadioui, M.; Peyrot, C.; Wilkinson, K. J. Anal. Chem. 2014, 86, 4668-4674. (5) Pace, H. E.; Rogers, N. J.; Jarolimek, C.; Coleman, V. A.; Higgins, C. P.; Ranville, J. F. Anal. Chem. 2011, 83, 9361-9369. (6) Laborda, F.; Jimâenez-Lamana, J.; Bolea, E.; Castillo, J. R. J. Anal. At. Spectrom. 2011, 26, 1362-1371. (7) Hadioui, M.; Merdzan, V.; Wilkinson, K. J. Environ. Sci. Technol. 2015, 49, 6141-6148.

Supporting Information


Page 17 of 23

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17 364 365 366 367 368 369 370 371 372 373 374 375 376 377

(8) Gottschalk, F.; Nowack, B. JEM 2011, 13, 1145-1155. (9) Regulatory Cooperation Council (RCC) Nanotechnology Initiative, Final Report on Work Element 2, Priority Setting: Development of a joint nanomaterial classification scheme.: Available at: Government of Canada NanoPortal (http://nanoportal.gc.ca), Accessed May 28, 2012. (10) Koppel, D. E. J. Chem. Phys. 1972, 57, 4814. (11) Hineman, A.; Stephan, C. J. Anal. At. Spectrom. 2014, 29, 1252. (12) Liu, J.; Hurt, R. H. Environ. Sci. Technol. 2010, 44, 2169-2175. (13) Cumberland, S. A.; Lead, J. R. J.Chrom. A 2009, 1216, 9099-9105. (14) Mitrano, D. M.; Ranville, J. F.; Bednar, A.; Kazor, K.; Hering, A. S.; Higgins, C. P. Environ. Sci.: Nano 2014, 1.



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Page 18 of 23

18 378 379 380 381

382 383 384 385 386 387

TABLES & GRAPHS Table 1. Supplier Information for the Nanomaterials Used Metal

Nominal size

Supplier

Product Name

Surface Coating

Form

Supplier’s size specification

Au

30 nm

NIST*

Citrate

60 nm

NIST

Ag

30 nm

Ted Pella

Ag

60 nm

Ted Pella

Ag

40 nm

Nano-composix

Aqueous dispersion Aqueous dispersion Aqueous dispersion Aqueous dispersion Powder

27.6 ± 2.1 nm

Au

Reference Material 8012 Reference Material 8013 PELCO NanoXact Silver PELCO NanoXact Silver Redispersible silver nanospheres

Citrate Citrate Citrate PVP, 88% by mass

* National Institute of Standards and Technology

Table 2. SP-ICP-MS Instrumental and Analytical Details Parameter Group ICP-MS component

Parameter

Parameter Value

ICP-MS model

Spray chamber

Perkin Elmer NexION 300D Low pressure PFA nebulizer Cyclonic, glass

Neb gas flow

1.04 L/min

Plasma RF power

1600 W

Sample flow rate

0.270 mL/min

Analyte monitored

107

Sample introduction Transport efficiency Instrument dwell time Sample analysis time Readings per sample

Manual, with a 60 s rinse and 20 s read delay 9.2% to 9.8%

Nebulizer

ICP-MS settings

Single Particle Parameter

Ag or 197Au

50 µs 60 s (or more) ~ 1.2 x 106 (or more)

388 389


56.0 ± 0.5 nm 32.7 ± 4.8 nm 62 ± 7 nm 38.6 ± 9.8 nm

Page 19 of 23

19 390 350

(a)

300

Frequency

Frequency

250 200 150 100 50 0 24

391

37

44

49

54

35

58

400 350 300 250 200 150 100 50 0

(b)

24

61

37

44

49

54

10

(c)

30

58

(d)

8

Frequency

Frequency

25 20 15 10 5

4 2

24 37 44 49 54 58 61 64 67

24 37 44 49 54 58 61 64 67 69

392

6

0

0

3

140

(e)

(f)

120 100

2

Frequency

Frequency

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


1

80 60 40 20

0

0 24 37 44 49 54 58 61 64 67 69 72 74 76

393 394 395 396 397

Diameter (nm)

24

37

44

49

54

58

Diameter (nm)

Figure 1. Particle size histograms for the 30 nm Au particle suspension with (a) no dissolved Au added, (b) 0.5 µg L-1 dissolved Au, (c) 1 µg L-1 dissolved Au, (d) 2 µg L-1 dissolved Au, (e) 3 µg L-1 dissolved Au, and (f) 3 µg L-1 dissolved Au following 5x dilution and extending sample analysis time to 120 s.



20 30 nm 30 nm (5x dilute)

90

60 nm 60 nm (10x dilute)

mean size (nm)

80 70 60 50 40 30 20 1.0

2.0

3.0

4.0

5.0

log [dissolved Au] (ng L-1)

398 1000000 100000

particles mL-1

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 23

10000 1000 100 1.0

399 400 401 402 403 404 405 406 407 408 409

2.0

3.0

4.0

5.0

log [dissolved Au] (ng L-1)

Figure 2. Mean particle size (nm) and number of particles (mL-1) measured in 30 nm and 60 nm Au particle suspensions (nominal particle concentrations = 105 particles mL-1) spiked with increasing concentrations of dissolved Au, shown as log concentration (ng L-1) along the x-axis. Included are the results for the 30 nm + 3 µg L-1 dissolved Au sample which was diluted 5x (×), and results for the 60 nm + 40 µg L-1 dissolved Au sample which was diluted 10x (+). Note: The dilution factors have been applied to the particle counts so that values shown represent the final particle concentrations for the original (undiluted) samples.


Page 21 of 23

21 nAgEA (1x) nAgEA (50x) nAgEA (100x)

60 50

Frequency

40 30 20 10 0 0

20

40

60

80

100

120

140

410 nAgMicro (100x) nAgMicro (2500x) nAgMicro (5000x)

140 120

Frequency

100 80 60 40 20 0 0

20

40

60

80

100

120

411 nAgFC (1x) nAgFC (100x) nAgFC (200x) - 1st curve nAgFC (200x) - 2nd curve

60 50 40

Frequency

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


30 20 10 0 0

412 413 414 415 416 417 418 419

20

40

60

80

100

120

140

Diameter (nm)

Figure 3. Equivalent particle size distributions (PSDeqv) for particulate Ag detected in soil extracts using a dilution series technique. Soil samples were taken from: an earthworm assay (nAgEA), a microbial assay (nAgMicro), and a collembolan assay (nAgFC). Extract dilutions are shown in brackets. Note: analysis time for nAgMicro (5000x) and nAgFC (200x) was extended from 60 s to 120 s.



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Page 22 of 23

22 420 421 422 423

Table 3. Analytical information and SP-ICP-MS results for soil extracts analyzed for 107Ag using a dilution series. Sample time for analyses was 60 s, except for nAgMicr (5000x) and nAgFC (200x) which were analyzed for 120 s. Note: DF = dilution factor; part. = particle

Sample

Dissolv ed Ag (µg L-1)

Mode sizeeqv (nm)

# peaks detecte d

Final* part. counts log [part.] (mL-1)

1

5.05

68.2

8763

5.5

1.03

100

0.06

37.4

2542

6.9

6.77

200

< 0.05

36.6

1227

6.9

6.08

100

0.48

39.8

13055

7.7

41.5

2500

Single Particle-Inductively Coupled Plasma Mass Spectroscopy

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