stable isotope labelling of metal/metal oxide nanomaterials for … · 2020. 3. 11. · 1 1 stable...
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Stable isotope labelling of metal/metal oxide nanomaterials for 1
environmental and biological tracing 2
Peng Zhang1,*, Superb Misra2, Zhiling Guo1, Mark Rehkämper3 & Eugenia Valsami-Jones1,* 3
1School of Geography, Earth and Environmental Sciences, University of Birmingham, Birmingham, UK. 4 2Materials Science and Engineering, Indian Institute of Technology Gandhinagar, Gujarat, India. 3Department of 5 Earth Science and Engineering, Imperial College London, London, UK. Correspondence should be addressed to P. 6 Z. ([email protected]) and E.V.J ([email protected]). 7 8
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ABSTRACT 14
Engineered nanomaterials are often compositionally indistinguishable from their natural counterparts 15
and thus their tracking in the environment or within biota requires the development of appropriate 16
labelling tools. Stable isotope labelling has become a well-established such tool, developed to assign 17
“ownership” or “source” to engineered nanomaterial enabling their tracing and quantification, especially 18
in complex environments. A particular methodological challenge for the stable isotope labelling is to 19
ensure the label is traceable in a range of environmental scenarios but without inducing modification of 20
the properties of the nanoamaterial and without loss of signal from the label, thus retaining realism and 21
relevance. This protocol describes the strategy for stable isotope labelling of several widely used metal 22
and metal oxide nanomaterials, namely ZnO, CuO, Ag, and TiO2, using isotopically enriched precursors, 23
namely 67Zn or 68Zn metal, 65CuCl2, 107Ag or 109Ag metal, and 47TiO2 powder. A complete synthesis 24
requires 1 to 8 days depending on the type of nanomaterial, the precursors used and the synthesis 25
methods adopted. The physicochemical properties of the labeled particles are determined by optical, 26
diffraction and spectroscopic techniques for quality control. The procedures for tracing the labels in 27
aquatic (snail and mussel) and terrestrial (earthworm) organisms and monitoring the environmental 28
transformation of labelled silver nanomaterials are also described. We anticipate this labelling strategy 29
can be adopted by industry to facilitate applications such as nanosafety assessments before 30
nanomaterials enter the market and environment as well as product authentication and tracking. 31
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INTRODUCTION 33
Why is labelling required for environmental and biological tracing? 34
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There has been a notable rise in the development and production of nanomaterials in recent years owing 35
to their broad range of applications, from antimicrobials and drug carriers to next generation computer 36
chips and advanced materials, to name but a few. Estimated global production for typical nanomaterials, 37
such as ZnO, Cu, Ag, TiO2, and Fe, were reported as 34,000, 200, 450, 88,000, and 42,000 tons per year, 38
respectively1. The release of engineered nanomaterials into landfills, soil, air, water, and other 39
environmental compartments is therefore inevitable. Substantial research funding and efforts have been 40
devoted to nanosafety studies in the past 15 years and the field has moved forward significantly. 41
However, until a satisfactory regulatory regime is established concerns will remain regarding their 42
potential impact on the environmental and human health2. 43
Evaluating biological responses to nanomaterials at environmentally relevant concentrations is a 44
crucial point in the field of nanosafety3. A particular challenge encountered in such studies is to 45
distinguish and detect the newly introduced nanomaterials in complex natural biological and 46
environmental matrices at low concentrations and against potentially high natural background values of 47
the same element, e.g., zinc, copper, and titanium. Traditional analytical approaches for measuring total 48
metal concentrations of the target elements, such as inductively coupled plasma mass spectrometry 49
(ICP-MS) techniques, have gone a long way in improving their sensitivity and accuracy in recent years, 50
but are challenged when low levels of nanomaterials need to be determined against high natural 51
background levels of the target element. In particular, it was recommended that nanomaterial exposures 52
should generate elevated elemental concentrations in the tested samples that exceed the background 53
levels by a factor of 10 or more4. This recommendation follows from the observation that elemental 54
concentrations can have uncertainties that exceed ± 10% of whilst the background levels of elements in 55
biological tissues and organisms can readily vary by more than 10% even when they grow under 56
essentially identical conditions5,6. Many published exposure scenarios have used elevated concentrations 57
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of nanomaterials as a result of the analytical limitations, which, however, are generally not 58
environmentally realistic. For example, ZnO nanomaterial concentrations of 100 to 6400 mg kg-1 were 59
used in soil exposures of Eisenia fetida and Folsomia candida7,8; these levels are about 2 to 3 orders of 60
magnitude higher than the predicted environmental concentrations (PECs) of ZnO nanomaterials in soil9. 61
An important additional consideration is that nanomaterials are highly dynamic and prone to 62
transformation (physical, chemical, or biological) upon entering the environment or biological tissues10. 63
This applies even to materials that were previously considered “insoluble”, such as CeO2 nanomaterials, 64
which can release Ce3+ and transform into CePO4, Ce carboxylates etc11. Some other metal-based 65
nanomaterials (e.g., Ag, CuO/Cu2O, and ZnO nanomaterials) may, furthermore, dissolve quickly or 66
transform to structurally and/or chemically different phases. These processes further complicate 67
detection10. Introduction of a tracer (“label”) into the nanomaterial makes it possible to distinguish both 68
the original form and their transformation products from any natural background. 69
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Comparison of different labelling methods 71
A label may be an organic fluorescent dye, a foreign rare element of low natural abundance, or a less-72
abundant isotope (stable or radioactive) of the same constituent element(s) as in the nanomaterial. 73
Examples of fluorescent dyes include fluorescein isothiocyanate and rhodamine, which are used to trace 74
the uptake and distribution of nanomaterials (e.g., SiO2, graphene oxide) in cells and organisms12,13. 75
Labelling with a fluorescent dye, however, inevitably modifies the surface properties of the 76
nanomaterial and thus alters their environmental and biological behavior. Another commonly used 77
labelling technique is radioisotope labelling. For example, radioisotopes 141Ce, 59Fe, and 198Au are 78
frequently used for labelling of CeO2, Fe3O4, and Au nanomaterials and tracing in the environment and 79
organisms14-16. However, it is of more limited applicability due to the hazards involved in dealing with a 80
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radioactive substance. Moreover, the short half-life of many radioisotopes prevent their use in long term 81
environmental exposures of months to years. Labels in the form of fluorescent dyes and (if added as a 82
surface layer) radioisotopes can also detach from the core of the nanomaterial and thus may not 83
adequately replicate its real behaviour. For example, graphene oxide (GO) labelled with radioactive 125I 84
can gradually release the 125I in physiological fluids17. 85
Compared with the labelling methods above, stable isotope labelling of the nanomaterial itself (i.e. 86
where the nanomaterial is synthesized to have a unique isotopic composition throughout) is a safer as 87
well as more versatile and robust approach that has been well-established and has a range of essential 88
roles in many research fields such as earth18,19, environmental20, and life sciences21,22. Typical examples 89
in biological investigations are, for example, incorporation of different stable isotopes to support 90
accurate protein quantification in proteomics research and for the study of metabolic fluxes23,24. 91
Additionally, stable isotope labelling has potential for further development in the context of nanosafety 92
and environmental tracing of nanomaterials as well as new applications, such as e.g. quality control 93
(where the label could act as unique identifier) in products. 94
Different stable isotopes of the same element are fractionated during many natural processes such 95
as condensation, thermal diffusion, precipitation and biological reactions, and this leads to small mass-96
dependent variations in the isotopic abundances of the elements25. Tracing of the stable isotope labelled 97
nanomaterial takes place against this natural background and relies on the measurement of isotopic 98
changes in a system or compartment that results from the introduction of nanomaterials, prepared from a 99
highly enriched isotope of a constituent element 26. The isotopic changes that are induced by such 100
labelling are thereby orders of magnitude larger than the natural mass-dependent isotope fractionations 101
and can thus be readily detected even in complex natural samples. Once the nanomaterials are labelled, 102
the label can be detected at high selectivity in a large variety of bulk samples following dissolution using 103
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commonly available mass spectrometric techniques, such as quadrupole, sector-field or multiple-104
collector inductively coupled plasma mass spectrometry (ICP-MS), i.e. ICP-QMS, SF-ICP-MS, and 105
MC-ICP-MS. In situ detection of labelled products is also possible, for example using laser ablation 106
ICP-MS (LA-ICP-MS)27 or secondary ion mass spectrometry (SIMS) /nano-SIMS/time of flight SIMS 107
(TOF-SIMS) instrumentation28,29. 108
Another advantage of stable isotope labelling is its ability to monitor transformation of 109
nanomaterials (e.g., silver nanoparticles) in the environment. A critical question raised in toxicological 110
studies of metal-based nanomaterials is whether any observed toxic effects originate directly from the 111
particulates or the dissolved metal ions released from the nanomaterials.2 Addressing this question is 112
challenging due to the complex environmental and biological systems in which transformations take 113
place. In natural environments, silver nanoparticles can be oxidized to release Ag+ ions, which can then 114
be reduced again, as they move from one environmental compartment to another, to regenerate the silver 115
into a new material. These reactions can take place in limited space and time, so that the processes are 116
difficult to monitor. Using a double stable isotope labelling method, whereby enriched 107Ag 117
nanoparticles and 109AgNO3 were simultaneously employed, it was possible to monitor complex 118
transformation kinetics in aqueous media30, highlighting the capabilities of stable isotope labeling. 119
Although stable isotope labelling has many advantages compared with other labelling techniques, 120
it also has some limitations. Firstly, it can only be employed where more than one stable isotopes of the 121
element exist; notably gold has only one stable isotope and thus cannot be labelled using this technique. 122
Furthermore, the label can most practically be traced using ICP-MS bulk sample analysis, which is 123
destructive, requiring full digestion of the sample material using mineral acids. This in turn implies 124
sample availability in sufficient quantity and the adoption of appropriate health and safety measures and 125
precautions to prevent sample contamination. Additionally, compared with fluorescence labelling, the 126
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cost of stable isotopes is relatively high, and this should be considered in advance. Lastly, whilst in situ 127
imaging of stable isotopes is, in principle, possible by using, e.g. LA-ICP-MS and nano-/TOF-SIMS 128
techniques, such instrumentation is not widely available and the analyses require careful calibration 129
procedures. 130
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Stable isotope labelled metal/metal oxide nanomaterials 132
The authors, along with collaborators, have made major advances in the labelling approaches of 133
several particularly relevant nanomaterials, e.g., ZnO,4,31,32 CuO,33,34 and Ag35,36 nanomaterials. In 134
addition, TiO2 nanomaterials labelled with 47Ti and core/shell structured iron oxide@SiO2 labelled with 135
57Fe were also developed recently37,38. The labelling approaches were shown to be efficient and highly 136
sensitive for detecting the nanomaterials in diverse environmental matrices and biological tissues at low 137
concentrations, similar to the PECs of the nanomaterials. 138
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General concept of stable isotope labelling of nanomaterials 140
In general terms, the procedure of stable isotope labelling involves the synthesis of an nanomaterial 141
using a stable isotope enriched precursor as raw material (Figure 1). Isotopically enriched metal salts 142
may be directly used as precursors for nanomaterial synthesis. For example, isotopically enriched 143
65CuCl2 and 109AgNO3 were used directly as the precursors for synthesis of 65CuO nanomaterials and 144
109Ag nanomaterials 33,35. Enriched materials in the form of elemental metal or metal oxide are usually 145
transformed into soluble metal salts for further nanomaterial synthesis, typically by acid digestion31. 146
Direct milling of the metal/metal oxide powder could also, in principle, transform micron sized powders 147
or metal filings to labelled nanoparticulate material. Such milled particles are however, typically not of 148
uniform size and their size range cannot be controlled or modified precisely. Spark discharge reactions 149
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can alternatively be used to prepare isotopically labelled nanomaterials, but this requires specific 150
instrumentation which is not widely accessible39. In contrast, wet chemistry procedures can produce 151
nanomaterials with controllable size over a wide size range, and other desired properties (e.g. specific 152
structural form, complex composition). As such procedures are also accessible for most laboratories, 153
they are considered most suitable and common to produce stable isotope labeled nanomaterials. 154
The methods used for transforming precursors depend primarily on the procedures that are applied 155
for nanomaterial synthesis. For example, commercially available enriched Zn isotopes are usually in the 156
form of Zn metal or ZnO powder. To produce ZnCl2 or Zn(NO3)2 precursors, hydrochloric acid or nitric 157
acid digestion can be used. In our study, we refluxed the Zn metal powder in acetic acid to produce zinc 158
acetate as the precursor, which was optimal for the synthesis of ZnO nanomaterials by a simple 159
hydrolysis method40,41. 160
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Figure 1 here 162
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EXPERIMENTAL DESIGN 164
Precursor choice and preparation 165
Most metallic elements have more than one stable isotope, and nanomaterials produced from such 166
metals can hence be labelled with an enriched stable isotope. However, there are many considerations 167
for choosing the most suitable isotope for labelling (Figure 2). The primary factors to be considered are 168
possible spectral interferences during the mass spectrometric analyses, although such effects can be 169
reduced or avoided by using instrumentation such as collision/reaction cell ICP-MS42 or a chemical 170
separation of the element prior to the measurements, as is common for analyses by MC-ICP-MS5. For 171
example, 49Ti and 112Cd are unfavorable for labelling because their measurement is hindered by 172
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polyatomic interferences from 32S16O1H and 40Ca216O2 which have the same mass number as 49Ti and 173
112Cd. Elements such as Si, S and Ca are very common in biological and environmental samples, whilst 174
H, C, N, O, S and Cl are present in air or the reagents and acids that are used for sample preparation. 175
Argon-based interferences are also a major problem for the measurements, as polyatomic species are 176
formed with C, H, O, and N, e.g., 36Ar13C and 36Ar12C1H for 49Ti, and 40Ar216O2 for 112Cd. In these cases, 177
alternative isotopes, namely 46Ti or 47Ti and 114Cd or 116Cd, can be chosen for labelling. 178
The required enrichment level and quantity of the isotopes, as well as the cost of the enriched 179
material should be considered together at this stage. The enrichment levels of commercially available 180
isotopes range widely, from less than 30% (e.g., 29% for 116Cd) to more than 99% 181
(http://www.tracesciences.com/). To achieve high sensitivity, the use of highly enriched isotopes with a 182
relatively low natural abundance is always preferable. However, a more precise detection method (e.g, 183
MC-ICP-MS) can deliver similar tracing sensitivities even when an enriched isotope with high natural 184
abundance is used. For examples, MC-ICP-MS can deliver similar tracing sensitivity for more abundant 185
and hence cheaper 68Zn (~19% natural abundance) as less precise analyses by ICP-QMS can provide 186
using less abundant and hence more expensive 67Zn (~4% natural abundance). 187
The quantity of enriched material required depends on the experimental system to be studied as in 188
some cases, such as mesocosm exposures, large amounts of the labelled materials are needed. However, 189
the sensitivity of detection and the quantity used must be balanced against the cost of the isotopes, which 190
increases substantially for those of low natural abundance. For example, it was previously reported that 191
64Zn enriched to 99% and with natural abundance of 48% costs ~ $4.5 per milligram, while the price 192
jumps to ~$250 per milligram for 70Zn enriched to 95% and a natural abundance of 0.6%.4 193
The methods that are employed for the preparation of labeled nanomaterials should be chosen 194
carefully given the high cost of enriched isotopes and procedures with high yield and simple steps are 195
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much preferred. In addition, they are constrained by the precursor form required, which needs to be 196
available either from commercial sources or an “in-house” preparation. In most cases, metal/metal oxide 197
nanomaterials can be synthesized using simple metal salts as precursors, e.g., AgNO3 for synthesis of 198
Ag nanomaterials, CuCl2 for synthesis of CuO nanomaterials, and Zn acetate for ZnO nanomaterials. 199
These precursors can hence be readily prepared in the laboratory. In general, all factors mentioned above 200
should be considered and balanced to choose the isotopes that are most suitable for a given labelling and 201
tracing study. 202
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Figure 2 here 204
205
Validation of synthesis method with no isotope labelling 206
Although there are many synthesis protocols available in the literatures for each type of nanomaterials, 207
many of these are not suitable for nanotoxicity studies. This reflects that such studies aim to evaluate 208
whether the metal core of nanomaterial is toxic or not, without considering the effects of any 209
contaminants or unwanted surface modifications. Many synthesis methods, however, involve toxic 210
chemicals, surfactants, and organic solvents, which may lead to false positive toxicity results for the 211
nanomaterials. A simple robust method that allows synthesis across the widest possible range of 212
parameters (e.g., size, shape, and surface charge) is preferable. For example, synthesizing nanomaterials 213
with different particle sizes using different methods may be associated with variable impurity contents 214
and structural changes, which may impact the results of the research in an unwanted manner. Methods 215
that involve multiple steps and organic reagents may also be not suitable if they are associated with low 216
synthesis yields. Finally, the synthesis should be robust and produce good yield in a reproducible 217
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manner. All these considerations can be readily verified in preliminary synthesis experiments that 218
employ natural (i.e., non-isotopically enriched) raw materials. 219
Taking ZnO nanomaterials as an example, there are two commonly used methods for the synthesis 220
of ZnO nanomaterials that employ a Zn acetate precursors, and these involve either hydrothermal 221
decomposition or forced hydrolysis. Preliminary work demonstrated that hydrothermal decomposition 222
only gave reaction yields of ~25% and the size of the ZnO nanomaterials was not uniform; as such, the 223
technique was deemed unsuitable for producing isotopically labelled nanomaterials. In comparison, 224
forced hydrolysis of the Zn acetate precursor in diethylene glycol (DEG) gives a yield of 75%, and the 225
size distribution of the ZnO nanomaterials is much more uniform. Furthermore, by manipulating the 226
synthesis conditions, specifically the precursor concentrations and reaction temperature, nanospheres 227
with different sizes can be obtained. The latter method is therefore preferred for producing isotopically 228
labelled ZnO nanomaterials31. 229
230
Step 1: Synthesis of nanomaterials with enriched isotopes 231
Synthesis of ZnO nanomaterials labelled with enriched 67Zn or 68Zn (Step 1A) Zinc has five 232
stable isotopes, 64Zn, 66Zn, 67Zn, 68Zn, and 70Zn, with natural abundance of 49.17%, 27.73%, 4.04%, 233
18.45%, and 0.61%, respectively. Isotopically enriched material is available for all five Zn isotopes with 234
enrichment levels that can exceed 95%. Considering the cost and the sensitivity of detection, 67Zn and 235
68Zn are optimal for labelling ZnO. Commercially available isotopically enriched Zn is usually in the 236
form of Zn metal (sheets or filings) or micron sized ZnO powder. To synthesize ZnO nanomaterials, the 237
Zn metal needs to be transformed into a suitable precursor, whereby Zn acetate is the commonly used 238
precursor for synthesis of ZnO nanomaterials. Zn acetate is readily prepared from Zn metal by refluxing 239
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the latter (preferably in powder form) in concentrated acetic acid. After drying, the Zn acetate can be 240
employed for synthesis of ZnO nanomaterials by hydrolysis. 241
242
Synthesis of CuO nanospheres and nanorods labelled with enriched 65Cu Cu has two stable 243
isotopes 63Cu and 65Cu with natural abundance of 69.17% and 30.83%, respectively. Enriched isotopes 244
of 63Cu and 65Cu are both commercially available with enrichments of up to 99%. Given the high 245
background concentrations of Cu in the environment and bio-organisms, enriched 65Cu (which has of 246
lower natural abundance) is recommended for tracing as this will provide better sensitivity. Since 247
65CuCl2 is available, this form can be directly used as precursor for synthesis of CuO nanomaterials by 248
wet chemistry. If the enriched Cu is purchased in the form of Cu metal or CuO powder, these materials 249
can be transformed into CuCl2 by digestion with hydrochloric acid. From CuCl2, CuO nanospheres and 250
nanorods can be synthesized by precipitation in alkaline solutions with or without the presence of glacial 251
acetic acid33. The yields of this method are 90% and 82% for nanospheres and nanorods, respectively, as 252
confirmed previously. 253
254
Synthesis of Ag nanomaterials labelled with enriched 107Ag and 109Ag Since the two isotopes 107Ag 255
and 109Ag have similar natural abundances of about 50%, 107Ag or 109Ag are equally applicable for 256
labelling purposes. These two isotopes are usually available in the form of Ag metal powder. Most 257
commonly, Ag nanomaterials are synthesized from an AgNO3 precursor, which is readily produced by 258
digestion of the Ag metal powders in concentrated nitric. Following this, citrate-coated Ag 259
nanomaterials were synthesized by reaction of AgNO3 with sodium citrate and NaBH4. By changing the 260
synthesis conditions, e.g., different amounts of NaBH4, Ag nanomaterials with different sizes can be 261
obtained. The yield of the isotopically labelled Ag nanomaterials can reach up to 98%. 262
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263
Synthesis of TiO2 nanomaterials labelled with enriched 47Ti There are five stable isotopes of Ti, 264
namely 46Ti, 47Ti, 48Ti, 49Ti, and 50Ti. Except for 48Ti with the highest natural abundance of 73.72%, 46Ti, 265
47Ti, 49Ti, and 50Ti are all of low natural abundance (8.25%, 7.44%, 5.41%, and 5.18%, respectively). Of 266
these, 49Ti and 50Ti are not suitable as tracers due to serious polyatomic interference from 32S16O1H and 267
36Ar14N, respectively. Thus, only 46Ti and 47Ti can be used. Commercially enriched Ti isotopes are 268
usually in metal or metal oxide form, which need to be transformed into a suitable precursor (e.g., TiCl4) 269
that can be used for synthesis of TiO2 nanomaterials. It should be noted that dissolving TiO2 is 270
challenging due to its refractory nature. In particular, hydrochloric or nitric acid do dissolve TiO2 but a 271
fusion approach can be applied to facilitate dissolution of TiO243. The obtained precursor solution can 272
then be used for synthesis of TiO2 nanomaterials with an appropriate method, such as a microwave 273
assisted precipitation procedure. A published procedure that applies such an approach only achieves 274
yields of ~ 14% for labelled TiO2 however, which is not ideal and should to be improved in future 275
studies. 276
277
Step 2: Quality control steps 278
The quality of the labelled nanoparticles can be established in a few key steps. Firstly, an appropriate 279
synthesis method should be chosen, to ensure that a good yield and suitable physicochemical properties 280
are obtained for the synthesis product. In particular, the synthesis procedure and product should be 281
tested initially using natural, non-isotopically enriched precursors. Such materials, however, will never 282
be exactly identical to the labelled precursors, such that caution should be exercised to minimize any 283
differences. Finally, a full physicochemical characterization of the labelled nanoparticles needs to be 284
performed and the results should be compared with data obtained for non-labelled nanomaterials. This 285
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comparison should yield no significant differences in the size, hydrodynamic diameter, morphology, 286
crystal structure, composition and surface chemistry, which are essential characteristics for reproducible 287
tracer studies. At the minimum, the synthesized nanomaterials hence need to be characterized by 288
transmission electron microscopy (TEM, Step 2A) or scanning electron microscopy (SEM) to confirm 289
size and shape, dynamic light scattering (DLS) or flow field-flow fractionation (FIFFF) to ascertain the 290
hydrodynamic diameter, X-ray powder diffraction (XRD) to determined crystallinity, and ICP-MS for 291
measurement of the chemical and isotopic composition. 292
293
Step 3: Tracing of labelled nanomaterials in environmental samples 294
The techniques for tracing nanomaterials in aquatic organisms (snails and mussels) and terrestrial 295
organisms (earthworms) are described below, thus demonstrating the applicability of the stable isotope 296
labelling approach. In addition, a protocol for monitoring the transformation of Ag nanomaterials is also 297
provided. 298
Tracing labelled nanomaterials in aquatic and terrestrial organisms The exposure protocol should 299
be tailored to the organism, exposure method (dietborne vs waterborne) and environment (terrestrial vs 300
aquatic). Accordingly, the total number of individuals that are investigated in a single exposure can vary 301
from less than ten to several hundreds (REFS?). The minimum number of individuals that compromise a 302
single sample for analysis is thereby determined by the detection limit for the determination of the 303
labeled isotope. In many cases, for example when larger organisms such as earthworm and zebra 304
mussels are investigated, this can be a single organism. However, when smaller organisms (e.g., 305
Limnaea), low exposure levels or ICP-QMS detection without prior chemical separation are applied, it 306
may be necessary to pool several individuals to produce a single sample that contains a measurable 307
quantity of labelled nanoparticles. It is, furthermore, generally recommended that at least three replicate 308
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samples are produced for any single time point and experimental line. Conventional biological exposure 309
studies commonly employ more replicate samples but our experience suggests that this is typically not 310
required for stable isotope tracing of nanomaterials. This reflects, in particular, that the stable isotope 311
approach is not compromised by variable biological background levels of the target element. 312
The organisms need to be acclimatized at normal growth condition prior to exposure. For aquatic 313
species, the experiments are carried out in appropriate water tanks. It is essential to use plastic rather 314
than glass tanks, at least in the case of ZnO nanomaterial exposures because the latter have been shown 315
to release significant quantities of Zn44. The nanomaterial suspensions are dispersed by ultrasonication 316
before addition to the tank. During waterborne exposures, continuous stirring is required to reduce 317
aggregation and sedimentation of the nanomaterial , which can have a significant impact on 318
nanomaterial uptake by the organisms. To ensure the organisms are not affected by the stirring, they 319
should be kept suspended (e.g., on a sieve) above the stirring device. An identical number of individuals 320
that are not exposed to the nanomaterial but are otherwise treated in the same manner are used as a 321
control for the exposure. In some cases, an additional control line is required to check for any effects 322
from the nanomaterial coating or suspension matrix. For example, as labeled ZnO NPs are synthesized 323
by forced hydrolysis in DEG, past exposures of earthworms to 68ZnO NPs employed an additional 324
control line whereby DEG was added to the soils. 325
Earthworms of uniform size and weight are chosen for the terrestrial exposure and the organisms are 326
acclimatized to the system prior to the experiment. The earthworms have two pathways for Zn uptake: (i) 327
dermal uptake via direct contact with the soil and (ii) dietary uptake of soil. For dietary exposures, it is 328
important to allow the earthworms to void their guts for 2 h prior to the exposure. For dermal exposures, 329
earthworms are prevented from oral ingestion by sealing their mouth with medical hystacryl glue. The 330
earthworms are placed into soils spiked with the 68ZnO nanomaterial and sampled at suitable time points. 331
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The accumulation of Zn in the earthworms is then determined following the same procedure that is used 332
for snails and mussels described above. 333
After exposure, the soft tissues are dissected, dried, digested and analyzed by ICP-MS. To obtain 334
accurate results, complete digestion of the samples is critical to ensure full recovery of the element 335
constituents. This is particularly challenging for TiO2 nanomaterials due to their refractory nature, and a 336
fusion method is thus used to digest samples exposed to TiO2 nanomaterials. The element concentrations 337
in the solutions obtained after digestion can be determined by ICP-QMS or high precision MC-ICP-MS 338
by external calibration with a series of standard solutions made of the target element. All solutions are 339
also doped with an internal standard, to compensate for matrix suppression and signal drift during the 340
analyses. Publishd ICP-QMS studies thereby employed the elements Sc, Y and Ge as internal standards 341
for Zn and Cu, Ag and Ti measurements45, respectively, whilst Cu was used for the determination of Zn 342
by MC-ICP-MS32. 343
Since ICP-QMS provides sufficient sensitivity and accuracy for many stable isotope tracing studies 344
whilst use of MC-ICP-MS is associated with additional sample separation, the choice of analytical 345
method should be governed primarily by the experimental design (sampling frequency, nanomaterial 346
exposure concentrations, etc) and the data quality required. 347
348
Monitoring the transformation of labelled 107Ag nanomaterials and 109Ag+ ions The 349
effect of environmental factors such as light irradiation, natural organic matter, divalent cations, pH 350
value and temperature on the transformation of Ag nanomaterials are examined. A key step prior to the 351
ICP-MS measurments is the seperation of Ag nanomaterials and Ag+ ions after incubation, as the Ag+ 352
ions may otherwise adsorb on the Ag nanomaterial surfaces46, which will lead to erroneous results. To 353
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achieve this, the samples are treated with an EDTA solution for 10 minutes and ultrafiltrated prior to 354
measurement. 355
356
MATERIALS 357
BIOLOGICAL MATERIALS 358
In our study, we use snails and mussels as representatives of aquatic organisms, and earthworm as 359
representative of terrestrial organism. As suggested in Step 3, 10 of each organism is selected for each 360
treatment. Snails and mussels are kept in moderately hard synthetic water and fed with benthic diatom 361
Nitzschia paleawas and cynaobacteria (Synechocystis PCC6803), respectively31,37. Earthworm is raised 362
in moist soil and fed with horse manure. 363
364
REAGENTS 365
Synthesis of ZnO nanomaterials labelled with enriched 67Zn or 68Zn 366
• 67Zn or 68Zn metal (89% enrichment, Isoflex, Russia) 367
• Glacial acetic acid (CH3COOH; Sigma Aldrich, Cat. no. 1000631011) 368
! CAUTION A flammable and corrosive liquid, which can cause severe skin burns and eye 369
damage. Wear gloves, lab coat, face mask, and goggles and perform experiments in fume hood. 370
• Diethylene glycol (DEG; ReagentPlus 99%, Sigma Aldrich) 371
! CAUTION DEG may cause damage to organs (kidney) if swallowed. 372
• Ultrapure water (18.2 MΩ cm, e.g., Milli-Q) 373
374
Synthesis of CuO nanospheres and nanorods labelled with enriched 65Cu 375
• Copper chloride dihydrate (65CuCl2.2H2O; Trace Sciences, USA, 99% enrichment) 376
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• Glacial acetic acid (Sigma Aldrich, Cat. no. 1000631011) 377
• Sodium hydroxide pellets (NaOH; Sigma Aldrich, Cat. no. 795429) 378
! CAUTION CH3COOH and NaOH may cause severe skin burns and eye damage. Wear 379
protective gloves, protective clothing and eye protection. CH3COOH is flammable. Perform 380
experiments in fume hood. 381
• Ultrapure water 382
383
Synthesis of Ag nanomaterials labelled with enriched 109Ag or 107Ag 384
• Silver nitrate (109AgNO3 or 107AgNO3; Trace Sciences, USA, 99% enrichment) 385
! CAUTION: May be corrosive to metals and intensify fire. May cause severe skin burns and eye 386
damage. Wear gloves, lab coat and goggles, and handle in fume hood. Silver nitrate is toxic to 387
aquatic life and should not be disposed off in the sink. 388
• Trisodium citrate dihydrate (HOC(COONa)(CH2COONa)2.2H2O; Sigma Aldrich, Cat No.: 71320) 389
• Sodium borohydride (NaBH4; Sigma Aldrich, Cat No.: 71320) 390
! CAUTION When in contact with water, flammable gases may be released which can ignite 391
spontaneously. Sodium borohydride is hazardous and is toxic if swallowed. It can cause sever skin 392
burns and eye damage. Wear protective gloves, lab coat, and eye protection and perform 393
experiments in fume hood. 394
• Ultrapure water 395
396
Synthesis of TiO2 nanomaterials labelled with enriched 47Ti 397
• Ultrapure water 398
• Titanium dioxide (Micro-sized 47TiO2; Eurisotop, France, 95.7% enrichment) 399
19
• Hydrochloric acid (HCl; Sigma Aldrich, Cat. no. H1758) 400
• Ammonium hydrogen difluoride (NH5F2; Sigma Aldrich, Cat. no. 224820) 401
• Sodium hydroxide pellets (Sigma Aldrich, Cat. no. 795429) 402
! CAUTION HCl, NH5F2, and NaOH may cause severe skin burn and eye damage. Wear 403
protective gloves, lab coat, face mask, and eye protection. Handle in fume hood. NH5F2 is toxic if 404
swallowed. 405
406
Analytical procedure for tracing of labelled nanomaterials in environmental samples 407
• Hydrogen peroxide (H2O2; Sigma Aldrich, Cat no. 16911) 408
! CAUTION H2O2 cause serious eye damage. Wear protective gloves, face mask and eye 409
protection. Operate in fume hood. 410
• 2-(N-morpholino)ethanesulfonic acid (MES; Sigma Aldrich, Cat no. 163732) 411
• Boric acid (H3BO3; Sigma Aldrich, Cat no. B6768) 412
! CAUTION H3BO3 may damage fertility and the unborn child. Wear protective gloves, clothing, 413
face and eye protection. Operate in fume hood. 414
415
EQUIPMENT 416
• Ultrapure water system (Milli-Q, Merck Millipore) 417
• Electrothermal stirring heating mantle (Electrothermal, UK) 418
• Three neck flasks, condenser, conical flask, and beakers (Scientific Glass Laboratories ltd, UK) 419
• Teflon vessels and stainless steel autoclave (200 mL and 50 mL volume) 420
• Platinum crucible (Cole-Parmer, UK) 421
• Welding torch (Rothenberger, UK) 422
20
• Microwave digestion system (Synthos 3000, Anton Paar) 423
• Centrifuge (Thermo Scientific, Sorvall ST8) 424
• Ultracentrifuge (L8-80M Beckman Coulter, J6-MI) 425
• Amicon Ultra-15 Centrifugal Filter Units (Merck Millipore, Cat no. UFC900308) 426
• Dialysis membranes (Thermo Scientific, 3.5K MWCO, Cat no.: 68035) 427
• Dynamic Light Scattering (Malvern Zeta sizer Nano ZS, UK) 428
• Zeta cells (Malvern Panalytical, Cat no.: DTS1070) 429
• Polystyrene cuvettes (Malvern Panalytical, Cat No.: DTS0012,) 430
• TEM instrument (Hitachi 7100, Japan) 431
• TEM Grids (Cat No.: AGS160, Agar Scientific) 432
• XRD instrument (Bruker D8 Advantage) 433
• ICP-MS instrument (Perkin Elmer, Nexion 2000) 434
• MC-ICP-MS instrument (Nu Plasma HR) 435
• FIFFF instrument (Postnova Analytics, AF 2000) 436
• Solar simulator (Beifanglihui Co., SN-500) 437
REAGENT SETUP 438
Synthetic moderately hard (MOD) water Prepare synthetic moderately hard water according to the 439
standard guideline (US EPA, 2002) for nanomaterial exposure experiment47. Dissolve appropriate 440
amounts of MgSO4, NaHCO3, and KCl solids in ultrapure water and aerate overnight. Dissolve CaSO4 441
solids in ultrapure water separately and aerate overnight. Mix the above two solutions with a volume 442
ratio of 19:1 to achieve MOD water. The final concentrations of MgSO4, NaHCO3, KCl, and CaSO4 in 443
the MOD water are 96, 60, 4 and 60 mg/L, respectively. The MOD water is then used for rearing snails 444
21
or mussels and for the preparation of nanomaterial suspensions. It can be stored in sealed bottles at room 445
temperature (20 ~ 25 oC) for 1 month. 446
447
EQUIPMENT SETUP 448
Transmission electron microscope (TEM) The TEM imaging is performed on a Hitachi 7100 TEM 449
(Japan) operating at 100 kV. 450
X-ray diffraction (XRD) X-ray diffraction is performed using Bruker D8 Advantage with Cu−Kα 451
radiation. 452
Dynamic Light scattering (DLS) The colloidal stability of the nanomaterials is evaluated by 453
measuring the hydrodynamic size and zeta potential (Malvern Instruments, Nano ZS). Hydrodynamic 454
size is also evaluated by FIFFF. 455
Inductive coupled plasma optical mass spectroscopy (ICP-MS) The concentrations and isotope 456
compositions of the metals in samples are measured by ICP-QMS (Perkin Elmer, Nexion 2000) or MC-457
ICP-MS (Nu Plasam HR). 458
459
PROCEDURE 460
Synthesis of nanomaterials with enriched isotopes 461
1 | The protocol below contains the steps to produce five kinds of isotope labelled nanomaterials, i.e. 462
67ZnO nanospheres (8 nm) (Option A), 65CuO nanospheres (7 nm) (Option B), 65CuO nanorods (7 × 40 463
nm) (Option C), 107Ag nanospheres (26 nm) with citrate capping (Option D), and rice grain like TiO2 464
nanomaterials (10 nm)(Option E). 465
(A) Synthesis of 67ZnO nanomaterials ● TIMING 8 d 466
22
(i) Place a 100 mL three-necked round bottom flask into a heating mantle stirrer. Connect a Liebig 467
condenser to the central neck of the flask. Plug the second opening with silicone bung with 468
thermometer inserted. Place an oval shaped magnetic stirring bar in the flask. 469
(ii) Fixe the entire setup using clamps. Connect the water inlet and outlet tubing to the Liebig 470
condenser and the water outlet tubing to the outlet drain sink. Place the setup in a fume hood. 471
CRITICAL STEP The experimental setup must be fixed tightly. The water supply must be 472
switched on prior to heating the three-necked flask. The water flow must be checked thoroughly 473
to ensure that there is no water leaking from the tubing/glassware. 474
(iii) Weigh 500 mg of 67Zn metal powder and add into the flask through the second opening. 475
! CAUTION Zinc powder may catch fire spontaneously if exposed to air or release flammable 476
gases in contact with water. the powder is toxic if inhaled. Wear gloves, lab coat and face mask. 477
(iv) Add 50 mL of glacial acetic acid into the flask through the second opening. Plug the opening 478
with a silicone bung. 479
(v) Heat the flask to 90 °C with the heating mantle. Keep refluxing at 90 °C for 3 d to form a Zn 480
acetate precursor solution. 481
(vi) Turn down the temperature to 50 °C and keep the temperature for 2 d to obtain the dry Zn 482
acetate precursor. The yield is approximately 1 g. The precursor can be stored for at least 1 year 483
in a tightly sealed container at dry place. 484
? TROUBLESHOOTING 485
(vii) Dissolve 100 mg of the precursor in DEG at 60 °C with stirring for 70 h using the same 486
refluxing setup as above. 487
(viii) Heat the resulting mixture to 180 °C. Add 100 µL ultrapure water into the mixture and 488
keep the temperature at 180 °C for 1 h to force hydrolysis of the precursor. 489
23
! CAUTION High temperature. Ensure the tap water connected to the condenser is turned on 490
prior to the heating. 491
(ix) Terminate the reaction and cool the flask down to room temperature (25 °C). Collect the 492
precipitate and centrifuge the suspension at 15,000 g and 4 °C for 15 min. Discard the 493
supernatant and wash the precipitate four times with ultrapure water. 494
(x) Dry the precipitates in a vacuum oven at 50 °C overnight. 495
■ PAUSE POINT The obtained 67ZnO nanomaterial powders are very stable for at least 1 year 496
at room temperature if stored in a desiccator. 497
498
(B) Synthesis of 65CuO nanospheres ● TIMING 1 d 499
(i) Dissolve 0.512 g 65CuCl2.2H2O in 150 mL ultrapure water at room temperature to obtain a 500
0.02 M solution. Use a three-necked round bottom flask to prepare the solution. Wait till the 501
entire copper salt is dissolved in water, giving a light greenish colour. 502
CRITICAL STEP If CuCl2.2H2O sticks to the walls of the original container, to avoid 503
any loss of isotope, dissolve entire bottle of purchased 65CuCl2.2H2O (with given quantity) at 504
once. 505
■ PAUSE POINT The 65CuCl2.2H2O can be stored for at least 3 months in a tightly sealed 506
container at room temperature. 507
(ii) Add 500 µL of glacial acetic acid to the solution and manually shake to ensure it is mixed in 508
the flask. 509
(iii) Transfer the flask into a heating mantle stirrer and attach a Liebig condenser using clamps. 510
The experimental setup is identical to the setup used for the ZnO synthesis (Step 1A(i) and 511
(ii)). 512
24
(iv) Heat the solution to 100 °C under stirring. Keep a check on the inserted thermometer to 513
monitor the the rise in temperature inside the vessel. Once the temperature reaches about 95 514
°C, weigh 0.6 g of NaOH and keep near the set up. 515
! CAUTION NaOH pellets are hygroscopic and should not be left out for long. The pellets 516
must be solid and should not be moist before adding. Wear gloves, goggles and lab coat. 517
(v) At 100°C open the stopper in the top of the condenser, and drop in the NaOH pellets. The 518
solution turns black immediately upon addition of the NaOH as precipitation of 65CuO takes 519
place. 520
! CAUTION Opening of the condenser stopper must be done carefully and slowly. 521
CRITICAL STEP Ensure that all the NaOH pellets are all added at once. Failure to do 522
so will lead to greater inhomogeneity of the particle size and shape. 523
? TROUBLESHOOTING 524
(vi) Keep the temperature at 100 °C for 10 mins. Then switch off the heater and stirrer, and let the 525
suspension cool down naturally to room temperature. 526
? TROUBLESHOOTING 527
(vii) Dismantle the experimental set up and decant the entire suspension into four 50 mL 528
centrifuge tubes. Centrifuge the suspension at 15,000 g and 4 °C for 10 mins. Discard the 529
supernatant and wash the precipitates four times with ultrapure water. 530
(viii) Place the tubes in a heating oven (60 °C) overnight to produce 65CuO nanomaterials in 531
dry powder form. 532
n ■ PAUSE POINT 65CuO nanomaterials are stable in a desiccator filled with inert gas for at 533
least 6 months. 534
535
25
(C) Synthesis of 65CuO nanorods ● TIMING 1 d 536
(i) Prepare rod shaped 65CuO nanomaterials by following the same steps as those used for the 537
preparation of spherical shaped 65CuO nanomaterials, except for no addition of glacial acetic acid (Step 538
2B (ii)). 539
540
(D) Synthesis of 107Ag nanospheres ● TIMING 4 d 541
(i) Dissolve sodium citrate in 100 mL ultrapure water in a glass beaker to obtain a 0.3 mM solution. 542
Allow the solution to cool at 4 °C for 30 min. 543
(ii) Dissolve 107AgNO3 in 100 mL ultrapure water to obtain a 0.25 mM solution. Allow the solution 544
to cool at 4 °C in the dark for 30 min. 545
! CAUTION AgNO3 may cause severe skin burns and eye damage. Wear gloves, lab coat and 546
goggles. 547
CRITICAL STEP Light irradiation may cause reduction of Ag+ to Ag metal. The AgNO3 548
solution must be kept in the dark. 549
(iii) Mix sodium citrate and AgNO3 solutions in a conical flask and stir vigorously. 550
(iv) Dissolve NaBH4 in 10 mL ultrapure water to a concentration of 10 mM. 551
! CAUTION NaBH4 is toxic if swallowed. It may cause severe skin burns and eye damage. It 552
react with water and release extremely flammable H2. Wear gloves, lab coat, face mask and 553
goggle. Operate in fume hood. 554
CRITICAL STEP NaBH4 reacts with water. The NaBH4 solution must be prepared freshly 555
and used immediately. 556
(v) Add 6 mL NaBH4 solution to the mixture (step (iii)) and keep stirring for 10 min. Cover the 557
flask with aluminum foil. 558
26
(vi) Heat the solution slowly to boiling. Keep heating and stirring for 90 min. 559
(vii) Terminate the experiment and cool the solution overnight in the dark at room 560
temperature. 561
(viii) Remove non-reacted reagents by dialysis of the obtained Ag nanomaterial suspension in a 562
1 KDa molecular weight cut-off dialysis bag against 5 L of 0.15 mM sodium citrate solution for 563
72 h. 564
■ PAUSE POINT The concentrated 107Ag nanomaterials stock suspension can be kept in a 565
tightly closed container protected from light at 4 °C for several months. 566
? TROUBLESHOOTING 567
568
(E) Synthesis of 47TiO2 nanomaterials ● TIMING 7 d 569
(i) Digest 1 g of micrometer sized 47TiO2 powder in 200 mL concentrated HCl (37%) in a 500 mL 570
Teflon vessel with lid for 6 days. 571
! CAUTION Concentrated HCl is highly corrosive and volatile. Wear gloves, lab coat, face 572
mask and goggles during operation. Perform experiments in fume hood. Keep lid closed during 573
digestion. 574
(ii) After 6 days, add 11.4 g NH5F2 to the Teflon vessel at once and mix with Teflon stirring rod. 575
Close the lid, seal the vessel in a stainless steel autoclave, and heat at 200 oC in a microwave 576
digestion system for 2 h. The obtained precursor solution will be used for anatase crystallization. 577
! CAUTION NH5F2 is toxic and corrosive. Wear gloves, lab coats and goggles. As NH5F2 578
corrodes glass, avoid use of any glassware for the handling. As high temperatures and pressures 579
are produced during the experiment, the lid of Teflon vessel should be closed and the autoclave 580
must be sealed tightly before heating. 581
27
(iii) Add NaOH pellets to the precursor solution until mildly acidic conditions (pH = 6) are reached. 582
A white precipitate will immediately appear. 583
(iv) Centrifuge the precipitate at 12,000 g and 4 oC for 15 mins and wash the pellet with ultrapure 584
water. Repeat the washing three times. Resuspend the pellet in 100 mL ultrapure water and 585
adjust the pH to 6 with HCl. 586
! CAUTION NaOH and HCl are highly corrosive. Wear gloves, lab coats and goggles. 587
(v) Transfer the suspension into four Teflon vessels (maximum volume 50 mL) and screw the lids 588
tightly. Heat the vessels in a microwave system (Synthos 3000, Anton Paar) at 200 oC for 2 h. 589
! CAUTION High temperature and high pressure is produced, the lid of Teflon line should be 590
screwed tightly. 591
(vi) Switch off the microwave system and allow the vessels to cool down to room temperature. 592
Collect the products by centrifuging at 15,000 g for 15 min and washed four times with 593
ultrapure water. 594
(vii) Dry the product in an oven at 60 oC for 24 h. 595
■ PAUSE POINT The synthesized 47TiO2 nanomaterials are stable at least for 1 year at room 596
temperature if stored in a dry place. 597
598
Quality Control Steps ● TIMING 2 d 599
2 | Before using the labelled nanomaterials in environmental tracing studies, their quality should be 600
checked using a variety of characterization techniques, as feasible/necessary. It is anticipated that the 601
physicochemical properties of the labelled nanomaterials should not deviate from those of the non-602
labelled nanomaterials synthesized first, to ensure the methodology works without wasting costly 603
28
labelled material. The four characterisation techniques listed below provide a minimum level of quality 604
control. Fig. 3 shows typical results obtained using two of the techniques, TEM and DLS. 605
(A) TEM imaging ● TIMING 1 h 606
(i) Sonicate the nanomaterial suspensions for 15 min, and deposit 5 µL of nanomaterial suspension 607
on a carbon-coated copper TEM grid and leave for air-drying. 608
(ii) Image the particles; an appropriate imaging approach may be performed using a Hitachi 7100 609
TEM instrument operating at 100 kV. 610
(iii) Determine the average size of the nanomaterials by measuring 100 or more particles by ImageJ 611
software. 612
(B) XRD determination of crystal structure ● TIMING 1 h 613
(i) Perform XRD analyses of dry nanomaterial powders; this can be done, for example, using a Bruker 614
D8 Advantage XRD instrument with copper Kα radiation. Collect the data at angles (2 theta) from 0 to 615
90 degrees. 616
(ii) Compare the XRD peaks with the standard PDF (powder diffraction file) to assign the structure. 617
Make sure the crystal structure of the sample is identical to the structure of the non-labelled product. 618
(C) DLS analysis ● TIMING 1 h 619
(i) Prepare nanomaterial suspensions with appropriate concentrations, followed by sonication for 15 620
minutes before analysis. The optimal concentration for measuring the particle size of the nanomaterial 621
suspension mainly depends on particle size. It is recommended to make the measurements, at a 622
concentration of 0.5 g/L, if the anticipated particle size is smaller than 10 nm and 0.1 g/L if the 623
anticipated particle size is in the range of 10 to 100 nm 624
(https://www.malvernpanalytical.com/en/learn/knowledge-center/user-manuals/MAN0485EN). 625
29
(ii) Analyse the sample. A suitable instrument for such analysis is the Malvern Zetasizer. The 626
hydrodynamic size measured by DLS of the labelled nanoparticles should be identical with the non-627
labelled particles. 628
(D) Analysis by ICP-MS ● TIMING 6 – 7 d 629
(i) Weigh the dry nanomaterials and add 20 mg of the particles in 10 mL deionized water to prepare 630
stock suspensions of nanomaterials. 631
(ii) Digest the ZnO, CuO and Ag nanomaterials (100 µL, taken from the stock suspensions) in an HNO3 632
- H2O2 mixture with a volume ratio of 4:1 (total volume 10 mL) by heating either on a hot plate 633
(typically 3 hrs) or in microwave digestion system (typically 30 min). For the TiO2 nanomaterials, HCl 634
and NH5F2 are used successively for dissolution with the assistance of a microwave digestion system. 635
(iii) Dry the solutions down, suspend // dissolve the material in 2% HNO3 to a volume of 10 mL. Take 636
100 µL of the solution and dilute to 10 mL by 2% HNO3. 637
(iv) Determine the the trace concentration and isotope composition of the metal by ICP-MS. A suitable 638
instrument can be the PerkinElmer Nexion 2000 ICP-MS. 639
640
Tracing the labelled nanomaterials in environment 641
3 | Using the labelled nanomaterials in tracing experiments. Three examples are described here, 642
involving two aquatic (snail and mussel, (A)) and a terrestrial (earthworm, (B)) organisms, which are 643
tested for the traceability of the labels. The general procedure is to expose the snail, mussel or 644
earthworm to the nanomaterials and determine the accumulated metals in the soft tissues. In the final 645
example described (C), the tracing sensitivity of the labels is exemplified by monitoring the 646
transformation of Ag nanomaterials in aqueous systems. 647
648
30
(A) Tracing the labelled nanomaterials in aquatic organisms ● TIMING 2d - 4d 649
(i) Prepare MOD water for culturing of the snails or mussels according to the standard guideline 650
(US EPA 2002)47. 651
(ii) Prepare nanomaterial stock solutions depending on the final exposure concentrations. For 652
example, to expose the snail to 10 µg L-1 of nanomaterials in a 10 L tank, prepare 10 ml of a 100 653
mg L-1 stock solution. Sonicate the stock suspensions for 10 to 20 mins to disperse the 654
nanomaterials homogeneously. Add 1 mL of stock into the tank and mixed homogeneously with 655
a magnetic stirring bar. 656
? TROUBLESHOOTING 657
(iii) Place the snails or mussels with uniform size and weight on sieves and suspend them in tank 658
containing the nanomaterial suspension. Keep the suspension stirred at a speed of about ~ 600 659
rpm. 660
661 CRITICAL STEP The suspension should be stirred continuously to avoid sedimentation 661
of the nanomaterials and potential interference on their uptake by the organisms. Stirring speed 662
should not be too high to disturb the normal activity of snails or mussels, or too low to let the 663
nanomaterials sink to the bottom. 664
(iv) After completion of the exposure, collect the snails or mussels, and rinse their surfaces with 665
ultrapure water four times. Dissect the soft tissues for measurement of the metal concentrations. 666
(v) Dry the soft tissues in a freeze-drier for 24 h and grind the tissues into fine powders. 667
■ PAUSE POINT Samples are stable once dried. Avoid moisture. 668
! CAUTION Wear gloves and face mask to avoid inhalation of powder. 669
(vi) Weigh appropriate amounts of the dry tissues and digest using mineral acids. For samples 670
treated with ZnO, CuO and Ag nanomaterials, use an HNO3 – H2O2 mixture for digestion 671
31
following standard protocols. Transfer the solutions obtained after digestion into a 15 mL Falcon 672
tubes for analysis. For samples from exposures to TiO2 nanomaterials, place the samples in 673
platinum crucibles, ash at 700 oC in an oven and fuse the ashes with ammonium persulfate with a 674
welding torch for 10 min. Cool the crucible and repeatedly rinse the samples with 2% (v/v) 675
HNO3 to obtain the sample solution. The samples are then ready for analysis by ICP-MS. 676
! CAUTION High temperatures may cause severe skin burn. Wear arc welding gloves and 677
helmet during the welding. The flame of the welding torch may cause fire; work in a cleared-off 678
area away from inflammable substance. 679
CRITICAL STEP Samples need to be digested completely until a clear and transparent 680
solution is obtained. A fusion method using ammonium persulfate should be used for samples 681
containing TiO2 nanomaterials to ensure full recovery of Ti from the samples. 682
■ PAUSE POINT Sample solutions in Falcon tubes can be stored at 4 oC for up to 3 months 683
without impact on the metal concentrations. Longer-term storage of the solutions may be 684
associated with absorption of metals on the container walls. 685
(vii) Measure the metal concentrations of the solution by ICP-MS. If MC-ICP-MS is used to 686
determine the metal concentrations and isotope compositions, the target element needs to be 687
separated from the sample matrix by ion exchange chromatography using a suitable element-688
specific protocol41. 689
690
(B) Tracing the labelled nanomaterials in terrestrial organism ● TIMING 5d – 8d 691
(i) Dry the standard soil Lufa 2.2 soil at 60 oC in an oven overnight to remove soil fauna. 692
(ii) Weigh 0.6 kg dry soil and place into a plastic beaker. 693
32
(iii) Add the nanomaterial stock solution to the soil to obtain a concentration of 5 mg kg-1. 694
Thoroughly mix the soil and nanomaterials with a spatula or rod. Add 0.162 kg deionized water 695
to the soil to achieve a moisture content of 27%; this is equivalent to half of the maximum water 696
holding capacity (WHC) of the soil. 697
CRITICAL STEP The soil and nanomaterials need to be mixed thoroughly to achieve 698
a homogeneous distribution of the nanomaterials. This is critical for the subsequent 699
exposure experiment. 700
(iv) Allow the spiked soil to equilibrate for 2 days before the introduction of the earthworms. 701
(v) Collect earthworms (Lumbricus rubellus) from the field, keep them in moist soil and feed with 702
horse manure. 703
(vi) Choose earthworms of uniform size and weight and with developed clitellum. Transfer 704
earthworms to the clean Lufa 2.2 soli at 50% WHC. Allow acclimation of the earthworms at 15 705
oC for 2 days. 706
(vii) Wash the earthworms with deionized water and blot them dry on filter paper. Place the 707
earthworms on clean moist filter paper, allowing them to void the guts for 2 days. Change the 708
paper after 1 day to avoid coprophagy behavior. 709
CRITICAL STEP Voiding of gut is critical for dietary borne exposures. Two days is 710
sufficient for the earthworm species used in this study; for other species, the duration may 711
vary and should be examined. 712
(viii) For dermal-only exposures, seal the earthworm mouth using medical hystacryl glue after 713
removing mucus around the mouth region. 714
(ix) Add the earthworms to the nanomaterial-spiked soils, and collect samples at desired exposure 715
times. 716
33
(x) Digest and analyze the samples following the procedures described in 3A(iv) - (vii). 717
718
(C) Monitoring transformation of 107Ag nanomaterials and 109Ag+ ions in aquatic environments 719
● TIMING 4d 720
(i) Prepare 1 mg L-1 107Ag nanomaterials suspension by diluting the stock solution with deionized 721
water. Prepare 1 mg L-1 109Ag+ (AgNO3) solution in deionized water. Total volume depends on 722
experimental requirement. 723
! CAUTION Operate in the dark and avoid exposure to light. 724
(ii) Mix 20 mL of each solution in a 50 mL fluorinated ethylene propylene bottle. 725
(iii) To establish the effect of light radiation on the transformation, expose one set of bottles to light 726
in a solar simulator equipped with Xe lamps that produce an irradiation intensity of 550 W m-2 . 727
Another set of bottles is wrapped in two layers of aluminum foils and one layer of black plastic 728
bags as dark controls. 729
! CAUTION As Ag NPs and Ag+ are be sensitive to light irradiation, the bottles for the dark 730
control should be wrapped tightly. 731
(iv) To determine the effect of natural organic matter on the transformation, add Suwannee River 732
humic acid (SRHA) to the mixed solutions in Step (ii) to achieve a final concentration of 5 mg L-733
1. 734
(v) To investigate the effect of pH on the transformation in the presence of SRHA, maintain the pH 735
of the solutions in Step (iv) at 5.6, 7.4 and 8.5 using MES (10 mM, pH 5.6) and borate buffer 736
(1mM, pH 7.4 and 8.5) and adjust by addition of NaOH or HNO3. 737
34
(vi) To study the impact of cations, add stock solution of Ca(NO3)2 (40 g L-1) and Mg(NO3)2 (24 g L-738
1) to the solution in Step (ii) to obtain the final Ca2+ and Mg2+ concentrations of 40 mg L-1 and 24 739
mg L-1. 740
(vii) Take 5 mL samples from the bottles at each desired sampling time (up to 3 days). 741
! CAUTION Shake the bottles to homogenize the solutions prior to the sampling. 742
(viii) Mix the 5 mL samples with 2.5 mL EDTA solution (100 mM) for 10 min. 743
(ix) Separate Ag nanomaterials and Ag+ ions in the mixtures by ultrafiltration (30 kD MWCO). The 744
obtained filtrates are diluted in 2.5% HNO3 (v/v) for measurement by ICP-MS. 745
CRITICAL STEP EDTA effectively removes ions adsorbed on particle surfaces 746
without inducing Ag NM dissolution within 20 min, which is critical for the final measurements. 747
■ PAUSE POINT The solutions obtained in Step (ix) can be stored in Falcon tubes at 4 oC for 748
up to three month. Longer term storage may lead to loss of Ag due to the adsorption onto the 749
container walls. 750
Data reduction 751
4 | There are two approaches for quantifying the concentration of stable isotope labelled nanomaterials: 752
i) by measuring the concentration of the accumulated enriched isotope or ii) by measuring the changes 753
of a diagnostic isotope ratio. Using 67ZnO nanomaterials as an example, the concentration of the 754
accumulated enriched 67Zn in an exposed sample can be calculated from the total metal concentration 755
(p67 × [T67Zn]) minus the background concentration of 67Zn (p67 × [T66Zn]), i.e. the following equations: 756
Δ67Zn = p67 × ([T67Zn] – [T66Zn]) (1) 757
p67 indicates the relative abundance of 67Zn in calibration standards, [T67Zn] and [T66Zn] indicate the 758
total Zn concentrations inferred by the ICP-MS software. 759
p67 = Intensity [67Zn / (64Zn + 66Zn + 67Zn + 68Zn + 70Zn)] (2) 760
35
761
? TROUBLESHOOTING 762
Troubleshooting advice can be found in Table 1. 763
764
Table 1 here 765
766
● TIMING 767
Step 1A, Preparation of 67ZnO nanomaterials: 8 d 768
Step 1B, Preparation of 65CuO nanosphere: 1 d 769
Step 1C, Preparation of 65CuO nanorod: 1 d 770
Step 1D, Preparation of 109Ag nanomaterials: 4 d 771
Step 1E, Preparation of 47TiO2 nanomaterials: 7 d 772
Step 2A, TEM imaging: 1 h 773
Step 2B, XRD: 1 h 774
Step 2C, DLS analysis: 1 h 775
Step 2D, Analysis by ICP-MS: 6 – 7 d 776
Step 3A, Exposure of snails/mussels to nanomaterials and ICP-MS analysis: 2 – 4 d 777
Step 3B, Exposure of earthworms to nanomaterials and ICP-MS analysis: 5 – 8 d 778
Step 3C, Transformation of Ag nanomaterials and Ag+ ions in aquatic environment and ICP-MS 779
analysis: 4 d 780
781
ANTICIPATED RESULTS 782
Characterization of isotopically labelled nanomaterials 783
36
The isotope-labelled nanomaterials should have the same physicochemical properties (e.g., morphology, 784
size, surface charge, and crystal structure, etc.) as the non-labelled nanomaterials synthesized during 785
protocol development. This can be confirmed by quality control investigations of the nanomaterials 786
including using TEM/SEM, DLS, XRD, etc (Step 2). Figure. 3 shows images that characterize the size 787
and shape of labeled Ag and TiO2 nanomaterials and a comparison with the non-labelled counterparts. 788
The Ag NP1 and AgNP2 were synthesized using 10 mM and 1 mM solutions of NaBH4, respectively. 789
The hydrodynamic diameters of the Ag NP1 batch with or without 107Ag labelling were 26.8 ± 2.3 nm 790
and 24.7 ± 3.5 nm, respectively (Figure 3a), whilst the Ag NP2 batch had diameters of 17.4 ± 2.5 nm 791
(labeled) and 18.5 ± 3.3 nm (unlabeled) (Figure 3b). This demonstrated that the sizes of the Ag 792
nanomaterials synthesized from enriched 107Ag and natural Ag are identical. The TiO2 nanomaterials 793
that were prepared from enriched 47Ti and natural Ti show identical morphology (rice grain shaped) and 794
size (10.4 ± 3.3 nm and 12.9 ± 5.4 nm, respectively) (Figure 3c and 3d). A summary of the properties of 795
the stable isotope labelled nanomaterials is provided in Table 2. 796
797
Figure 3 here 798
Table 2 here 799
800
Enhanced tracing ability provided by stable isotope labelling 801
802 Figure 4 here 803
804 805
806
807
37
808
As an example of what results can be obtained in biological studies, snails (Lymnaea stagnalis) 809
and zebra mussels (Dreissena polymorpha) were exposed to a stable isotope labelled nanomaterial to 810
examine the tracing sensitivity of the labeling technique in an aquatic environment using the procedure 811
described in Step 3A. The average background concentrations of Zn and Cu in snails were 54 µg g-1 and 812
34 µg g-1, respectively, which is high relative to the predicted environmental concentrations (PECs) of 813
ZnO and CuO nanomaterials. For example, Gottschalk et al., modeled PECs for ZnO nanomaterials of 814
less than 0.5 µg L-1 for surface water, of 0.5 to 4 µg L-1 for waste water treatment plant effluents, of 10 815
to 16 µg g-1 for biosolids, and of 0.04 ~ 0.5 µg g-1 for sediment9. If stable isotope labeling is not used, Zn 816
in snails from ZnO nanomaterials is detectable only when the ZnO nanomaterial exposure 817
concentrations are higher than 5000 µg g-1 (Figure 4a), which does not constitute a realistic scenario. In 818
contrast, Zn additions of as little as 1 µg g-1 can be detected in snails when exposed to only 15 µg g-1 of 819
dietborne ZnO nanomaterials if an enriched label (67Zn) is used. This demonstrates that stable isotope 820
tracing provides a remarkably enhanced detection sensitivity. 821
For CuO nanomaterials, the newly accumulated Cu in snails exposed to CuO nanomaterials is not 822
detectable over a wide range of exposure concentrations (0.2 to 2000 µg L-1) unless the CuO 823
nanomaterials are prepared from enriched 65Cu (Figure 4b). 824
The predicted environmental concentrations of engineered Ag nanomaterials are 0.1 to 100 ng L-1 825
for surface water and 1 to 10 ng g-1 for sediments, and are thus similar to the respective natural 826
background concentrations of Ag 48. Stable isotope labelling is hence the only safe and reliable way for 827
tracing Ag nanomaterials at low environmentally realistic exposure concentration. Using 109Ag enriched 828
nanomaterials, even only 1 ng of newly accumulated Ag per gram of tissue can be detected (open 829
symbols, Figure 4c) in snails that were exposed to 6 ng L-1 Ag nanomaterials. However, if no tracer 830
38
used, quantifying the presence of the newly accumulated Ag requires exposure concentrations that are 831
two orders of magnitude higher. Similarly, for TiO2 nanomaterials, the newly accumulated Ti in mussels 832
(0.2 µg g-1) that were exposed to 47TiO2 nanomaterials at an environmental relevant concentration of 3.9 833
µg L-1 can be detected (Figure 4d). 834
The detection sensitivity can be further improved by tracing the change of a diagnostic isotope 835
ratios using a high precision MC-ICP-MS5. Taking 68Zn labelled ZnO nanomaterials as an example, the 836
68Zn/66Zn isotope ratio determined for an exposed sample is compared to the natural 68Zn/66Zn ratio 837
measured for a calibration standard (68Zn/66Znstd). The difference in isotope ratio can then be used to 838
calculate the proportion of enriched 68Zn present in the sample relative to the total and natural 839
background content of Zn. By measuring 68Zn/66Zn using MC-ICP-MS, 68Zn additions as low as 5 ng g-1 840
can be detected in natural samples with natural Zn backgrounds of 100 µg g-1. As such, the MC-ICP-MS 841
detection sensitivity is considerably better compared to that attainable by ICP-QMS, which can detect 842
68Zn additions that exceed ~120 ng g-1. However, the limitation of the MC-ICP-MS technique is that the 843
target elements need to be separated from the sample matrix before analysis. Such sample preparation is 844
laborious and requires special facilities, and this should be considered when the preferred instrumental 845
technique is chosen for an exposure experiment. 846
The performance of MC-ICP-MS is well demonstrated by a soil exposure of earthworms to labeled 847
68ZnO nanomaterials. After just 4 h of dermal exposure, an accumulation of 68Zn in the earthworms 848
equivalent to 0.03‰ of the total Zn content could be readily detected41. Using 68Zn isotope labelling, it 849
was also shown that the uptake kinetics of ZnO nanomaterial and ionic Zn by earthworm are similar 850
(Figure 4e and 4f), which probably reflects the rapid and complete dissolution of ZnO nanomaterials at 851
such low concentrations. However, different results were obtained in the study of Heggelund et al49, who 852
also exposed earthworm to ZnO nanomaterials and dissolved Zn but at much higher concentrations of up 853
39
to 2500 µg g-1. In particular, the latter study found lower toxicity when earthworms were exposed to 854
ZnO nanomaterials in comparison to those explosed to dissolved Zn, presumably due to limited ZnO 855
nanomaterials dissolution at the high exposure concentrations41. This demonstrated that stable isotope 856
labelling can provide unique mechanistic understanding on the uptake of nanomaterials at environmental 857
realistic exposure concentrations. 858
859
Transformation of Ag nanomaterials and Ag+ ion in aquatic environment 860
When Ag nanomaterials are released into the environment, they can be oxidized to release Ag+ 861
ions. Subsequently, the released Ag+ and any pre-existing Ag+ present can be reduced to produce a 862
mixed source regenerated Ag nanomaterials. Such reactions can be studied by stable isotope labeling, 863
using 107Ag nanomaterials and ionic109Ag+, thus each form of silver having its own distinct isotopic 864
composition. When 107Ag nanomaterials and 109Ag+ coexist, oxidation and dissolution of 107Ag 865
nanomaterials is the dominant reaction in a system of pure water (Figure 5a). However, in the presence 866
of dissolved organic matter (DOM), reduction of Ag+ and regeneration of Ag nanomaterials is the 867
dominant process (Figure 5b). Temperature, pH value and the presence of other divalent ions have a 868
significant impact on these transformation processes. For example, addition of Ca2+ and Mg2+ at 869
environmentally relevant concentrations had no strong effect on the reduction of 109Ag+ (Figure 5c). 870
However, it induced significant agglomeration of 107Ag nanomaterials, which considerably reduced the 871
effective surface area of the Ag nanomaterials and thus limited their subsequent dissolution in the dark 872
(Figure 5d). 873
These findings further highlight the power of the isotope labelling approach in tracing nanomaterials 874
in complex environment samples and systems. There is also significant potential for expanding the 875
double isotope labelling technique to other nanomaterials, in particular by labelling two different 876
40
elements. For example, this may be applicable to CdS quantum dots, an approach currently being 877
developed at the University of Birmingham. By labelling CdS with both 110Cd and 34S, it can be possible 878
to not only quantify the CdS present, but also detect environmental and biological transformation of CdS. 879
For example, to assess whether the labelled Cd and S remain together as the original particles or separate 880
and transform into distinct more stable forms with time. 881
Figure 5 here 882
Adaptation for industrial applications 883
Stable isotope labelling could find a number of industrial applications, whether to ensure safety or 884
authenticity of a product. In an industrial context, stable isotope labeling could be used to ensure the 885
authenticity of a product. More specifically, an isotopic label can link a material to its source following 886
accidental release, theft or any other scenario requiring unequivocal identification. In such cases, a 887
unique isotopic signature could represent an entire product line, a particular product or even a unique 888
batch. To save on the cost of the stable isotope label, the adaptation for industrial applications does not 889
need to employ full isotopic modification but could involve the synthesis procedures described above 890
(A-E) but with only a limited quantity of labeled material and the remaining synthesis precursor 891
unlabeled. This procedure would introduce an isotopic variation that is sufficient to differentiate a 892
labeled product from other similar materials (or batches) and background isotopic compositions, 893
whereby the proportion of enriched label present in the product is experimentally assessed prior to 894
release to ensure robust traceability. 895
896
ACKOWLEDGEMENTS 897
This work was supported by Marie Skłodowska-Curie Individual Fellowship (NanoLabels 750455 to 898
PZ; NanoBBB 798505 to ZG) under the European Union’s Horizon 2020 research program. Financial 899
41
support from MHRD-IMPRINT funds is appreciated. Partial funding from EU H2020 project ACEnano 900
(grant agreement no. 2016-720952) is also acknowledged. 901
902
AUTHOR CONTRIBUTIONS 903
P.Z., S.M., and Z.G. wrote the paper. E.V.J. and M.R. design part of the experiments and revised the 904
paper. S.M. performed part of the experiments. 905
COMPETING FINANCIAL INTERESTS 906
The authors declare no competing financial interests. 907
DATA AVAILABILITY 908
The data that support the plots within this paper are available from the corresponding author upon 909
reasonable request. 910
REFERENCES 911
1 Keller, A. A. & Lazareva, A. Predicted releases of engineered nanomaterials: from global to 912
regional to local. Environ. Sci. Technol. Lett. 1, 65-70 (2013). 913
2 Valsami-Jones, E. & Lynch, I. How safe are nanomaterials? Science 350, 388-389 (2015). 914
3 Holden, P. A. et al. Considerations of environmentally relevant test conditions for improved 915
evaluation of ecological hazards of engineered nanomaterials. Environ. Sci. Technol. 50, 6124-916
6145 (2016). 917
4 Larner, F. & Rehkämper, M. Evaluation of stable isotope tracing for ZnO nanomaterials —new 918
constraints from high precision isotope analyses and modeling. Environ. Sci. Technol. 46, 4149-919
4158 (2012). 920
5 Leavitt, S., Dueser, R. & Goodell, H. Plant regulation of essential and non-essential heavy 921
metals. J. Appl. Ecol. 16, 203-212 (1979). 922
42
6 Sridhar, B. B. M., Han, F. X., Diehl, S. V., Monts, D. L. & Su, Y. Effects of Zn and Cd 923
accumulation on structural and physiological characteristics of barley plants. Braz. J. Plant 924
Physiol. 19, 15-22 (2007). 925
7. Hu, C. W., Li, M., Cui, Y. B., Li, D. S., Chen, J. & Yang, L. Y. Toxicological effects of TiO2 and 926
ZnO nanoparticles in soil on earthworm Eisenia fetida. Soil Biol. Biochem. 42, 586-591 (2010). 927
8. Kool, P. L., Ortiz, M. D. & van Gestel, G. A. M. Chronic toxicity of ZnO nanoparticles, non-nano 928
ZnO and ZnCl2 to Folsomia candida (Collembola) in relation to bioavailability in soil. Environ. 929
Pollut.159, 2713-2719 (2011). 930
9. Gottschalk, F., Sun, T. & Nowack, B. Environmental concentrations of engineered nanomaterials: 931
review of modeling and analytical studies. Environ. Pollut. 181, 287-300 (2013). 932
10 Lowry, G. V., Gregory, K. B., Apte, S. C. & Lead, J. R. Transformations of nanomaterials in the 933
environment. Environ. Sci. Technol. 46, 6893-6899 (2012). 934
11 Zhang, P. et al. Biotransformation of ceria nanoparticles in cucumber plants. ACS nano 6, 9943-935
9950 (2012). 936
12 Santra, S. et al. TAT conjugated, FITC doped silica nanoparticles for bioimaging applications. 937
Chem. Commun. 24, 2810-2811 (2004). 938
13. Zhi, L., Ren, M., Qu, M., Zhang, H. & Wang, D. Wnt ligands differentially regulate toxicity and 939
translocation of graphene oxide through different mechanisms in Caenorhabditis elegans. Sci. 940
Rep. 6, 39261 (2016). 941
14. Zhang, Z. et al. Uptake and distribution of ceria nanoparticles in cucumber plants. Metallomics 3, 942
816-822 (2011). 943
43
15. Freund, B. et al. A simple and widely applicable method to 59Fe-radiolabel monodisperse 944
superparamagnetic iron oxide nanoparticles for in vivo quantification studies. ACS nano 6 7318-945
7325 (2012). 946
16. Kannan, R. et al. Functionalized radioactive gold nanoparticles in tumor therapy. Wiley Interdiscip 947
Rev Nanomed Nanobiotechnol. 4, 42-51 (2012). 948
17 Yang, K., Feng, L., Hong, H., Cai, W. & Liu, Z. Preparation and functionalization of graphene 949
nanocomposites for biomedical applications. Nat. Protoc. 8, 2392-2403 (2013). 950
18 Wiederhold, J. G. Metal stable isotope signatures as tracers in environmental geochemistry. 951
Environ. Sci. Technol. 49, 2606-2624 (2015). 952
19 Simon, J. et al. Calcium and titanium isotope fractionation in refractory inclusions: tracers of 953
condensation and inheritance in the early solar protoplanetary disk. Earth. Planet. Sci. Lett. 472, 954
277-288 (2017). 955
20 Šillerová, H. et al. Stable isotope tracing of Ni and cu pollution in north-East Norway: potentials 956
and drawbacks. Environ. Pollut. 228, 149-157 (2017). 957
21 Chokkathukalam, A., Kim, D.H., Barrett, M. P., Breitling, R. & Creek, D. J. Stable isotope-958
labeling studies in metabolomics: new insights into structure and dynamics of metabolic 959
networks. Bioanalysis 6, 511-524 (2014). 960
22 Chahrour, O., Cobice, D. & Malone, J. Stable isotope labelling methods in mass spectrometry-961
based quantitative proteomics. J. Pharm. Biomed. Anal. 113, 2-20 (2015). 962
23 Yuan, J., Bennett, B. D. & Rabinowitz, J. D. Kinetic flux profiling for quantitation of cellular 963
metabolic fluxes. Nat. Protoc. 3, 1328 (2008). 964
24 Mugoni, V., Medana, C. & Santoro, M. M. 13C-isotope-based protocol for prenyl lipid metabolic 965
analysis in zebrafish embryos. Nat. Protoc. 8, 2337 (2013). 966
44
25 Sharp, Z. Principles of stable isotope geochemistry. (2017). 967
26 Dybowska, A., Misra, S. K. & Valsami-Jones, E. Labelling nanoparticles with non-radioactive 968
isotopes. In Isotopes in Nanoparticles - Fundamentals and Applications. (eds. Llop., J.,Gomez-969
Vallejo., V. & Gibson., N.) 455-485 (Pan Stanford Publishing Pte. Ltd. Singapore, 2016) 970
27 Becker, J. S., Matusch, A. & Wu, B. Bioimaging mass spectrometry of trace elements–recent 971
advance and applications of LA-ICP-MS: A review. Anal. Chim. Acta 835, 1-18 (2014). 972
28 Proetto, M. T. et al. Cellular delivery of nanoparticles revealed with combined optical and 973
isotopic nanoscopy. ACS nano 10, 4046-4054 (2016). 974
29 Lee, P.L. et al. Development and validation of TOF-SIMS and CLSM imaging method for 975
cytotoxicity study of ZnO nanoparticles in HaCaT cells. J. Hazard. Mater. 277, 3-12 (2014). 976
30 Yu, S., Yin, Y., Zhou, X., Dong, L. & Liu, J. Transformation kinetics of silver nanoparticles and 977
silver ions in aquatic environments revealed by double stable isotope labeling. Environ. Sci.: 978
Nano 3, 883-893 (2016). 979
31 Dybowska, A. D. et al. Synthesis of isotopically modified ZnO nanoparticles and their potential 980
as nanotoxicity tracers. Environ. Pollut. 159, 266-273 (2011). 981
32 Laycock, A. et al. Novel multi-isotope tracer approach to test ZnO nanoparticle and soluble Zn 982
bioavailability in joint soil exposures. Environ. Sci. Technol. 51, 12756-12763 (2017). 983
33 Misra, S. K. et al. Isotopically modified nanoparticles for enhanced detection in bioaccumulation 984
studies. Environ. Sci. Technol. 46, 1216-1222 (2011). 985
34 Croteau, M.-N. l., Misra, S. K., Luoma, S. N. & Valsami-Jones, E. Bioaccumulation and toxicity 986
of CuO nanoparticles by a freshwater invertebrate after waterborne and dietborne exposures. 987
Environ. Sci. Technol. 48, 10929-10937 (2014). 988
45
35 Laycock, A. et al. Synthesis and characterization of isotopically labeled silver nanoparticles for 989
tracing studies. Environ. Sci.: Nano 1, 271-283 (2014). 990
36 Croteau, M.-N., Dybowska, A. D., Luoma, S. N., Misra, S. K. & Valsami-Jones, E. Isotopically 991
modified silver nanoparticles to assess nanosilver bioavailability and toxicity at environmentally 992
relevant exposures. Environ. Chem. 11, 247-256 (2014). 993
37 Bourgeault, A. et al. The challenge of studying TiO2 nanoparticle bioaccumulation at 994
environmental concentrations: crucial use of a stable isotope tracer. Environ. Sci. Technol. 49, 995
2451-2459 (2015). 996
38 Meermann, B., Wichmann, K., Lauer, F., Vanhaecke, F. & Ternes, T. A. Application of stable 997
isotopes and AF4/ICP-SFMS for simultaneous tracing and quantification of iron oxide 998
nanoparticles in a sediment–slurry matrix. J. Anal. At. Spectrom. 31, 890-901 (2016). 999
39. Meuller, B. O. et al. Review of Spark Discharge Generators for Production of Nanoparticle 1000
Aerosols. Aerosol Sci. Technol. 46, 1256-1270 (2012). 1001
40 Croteau, M. N., Dybowska, A. D., Luoma, S. N. & Valsami-Jones, E. A novel approach reveals 1002
that zinc oxide nanoparticles are bioavailable and toxic after dietary exposures. Nanotoxicology 1003
5, 79-90 (2011). 1004
41 Laycock, A. et al. Earthworm uptake routes and rates of ionic Zn and ZnO nanoparticles at 1005
realistic concentrations, traced using stable isotope labeling. Environ. Sci. Technol. 50, 412-419 1006
(2015). 1007
42 Pick, D., Leiterer, M. & Einax, J. W. Reduction of polyatomic interferences in biological 1008
material using dynamic reaction cell ICP-MS. Microchem. J. 95, 315-319 (2010). 1009
46
43 Khosravi, K., Hoque, M. E., Dimock, B., Hintelmann, H. & Metcalfe, C. D. A novel approach 1010
for determining total titanium from titanium dioxide nanoparticles suspended in water and 1011
biosolids by digestion with ammonium persulfate. Anal. Chim. Acta. 713, 86-91 (2012). 1012
44. Kay, A. R. Detecting and minimizing zinc contamination in physiological solutions. BMC 1013
Physiol. 4, 1-9 (2004). 1014
45. Vanhaecke, F., Vanhoe, H., Dams, R., & Vandecasteele, C. The use of internal standards in ICP-1015
MS. Talanta, 39, 737-742 (1992). 1016
46. Liu, J. Y. & Robert, H. H. Ion Release Kinetics and Particle Persistence in Aqueous Nano-Silver 1017
Colloids. Environ. Sci. Technol. 44, 2169-2175 (2010). 1018
47. U.S. Environmental Protection Agency. Methods for measuring the acute toxicity of effluents 1019
and receiving waters to freshwater and marine organisms. Edn 5, EPA-821-R-02-012 (U.S. 1020
Environmental Protection Agency Office of Water, Cincinnati, OH, 1991). 1021
48. Eisler, R. Silver hazards to fish, wildlife and invertebrates: a synoptic review. US National 1022
Biological Service. Biological Science Report 32 (1981). 1023
49. Heggelund, L. R. et al. Soil pH effects on the comparative toxicity of dissolved zinc, non-nano 1024
and nano ZnO to the earthworm Eisenia fetida. Nanotoxicology 8, 559-572 (2014). 1025
1026
1027
1028
1029
1030
1031
1032
47
TABLE 1 | Troubleshooting table. 1033
Step Problem Possible reason Solution 1A(vi) The yield of the precursor is very
low Zinc powder was not dissolved completely
Check the temperature, keep at 90 °C Extend the heating time
1B(v) No formation of black precipitate pH of the solution was not high enough
Add accurate amount of NaOH and add it all at once
1B(vi) Formation of light green precipitates
Copper chloride did not react completely
Keep the solution at 100 °C for a longer duration
1D(viii) The yield of Ag nanomaterials is too low
NaBH4 solution was not freshly prepared
Prepare fresh NaBH4 and use it immediately
3(ii) Nanomaterials not disperse homogeneously
The stock suspension was not sonicated sufficiently Stirring speed was too low
Ensure sufficient sonication of the stock Increase stirring speed up to 600 rpm
1034
1035
TABLE 2 | Summary of the properties of the stable isotope labelled nanomaterials synthesized by the 1036
protocols. 1037
67ZnO 65CuO 65CuO 107Ag 47TiO2 Size (TEM) 8 ± 1 nm 7 ± 1 nm 7 ± 1 nm
width 40 ± 10
nm length
28 ± 8 nm 10 ± 3 nm
Shape (TEM) Spherical Spherical Rod Spherical Rice grain-shaped
Crystalline phase (XRD)
Wurtize (ICDD 36-
1451)
Tenorite (ICDD 48-
1548)
Tenorite (ICDD 48-
1548)
FCC (ICDD 87-
0720)
Anatase (ICDD 21-
1272)
Hydrodynamic size (DLS)
38.6 ± 0.7 nm
82 ± 1 nm NA 26.8 ± 2.3 nm
98 ± 7 nm
1038
NA indicate the value is not available due to the shape of nanorod and its tendency to agglomerate. 1039
48
FIGURE LEGEND 1040
Figure 1. Scheme of stable isotope labelling of NMs. x indicates the atomic mass number of isotope M. 1041
99% indicates the enrichment level of the isotope. 1042
Figure 2. Considerations for the choice of suitable isotope and chemical form. 1043
Figure 3. Hydrodynamic diameters and TEM images. (a) Hydrodynamic diameter distributions of 107Ag 1044
NMs (synthesized in Step 1D) determined by FIFFF. Nat-Ag NP indicate the Ag NMs synthesized 1045
without labelling. (b) Hydrodynamic diameter distributions of 107Ag NMs synthesized with a different 1046
method (all steps are identical to Step 1D except for Step 1D (iv) where 1 mM of NaBH4 is used). (c) 1047
and (d) show the TiO2 NMs with and without 47Ti labelling, respectively. Adapted from previous 1048
publications35,37. 1049
Figure 4. Enhanced detection sensitivity for NMs provided by stable isotope labelling. (a) Zinc 1050
concentrations in snails exposed for 3-4 h to dietborne 67ZnO NMs. Triangles and circles represent two 1051
dietborne exposure modes, i.e. food (diatoms) labelled with 67Zn and diatoms mixed with 67ZnO. (b) 1052
Copper concentrations in snails exposed for 24 h to waterborne 65CuO NMs in synthetic MOD water. (c) 1053
Silver concentrations in snails exposed for 24 h to waterborne 109Ag NMs in synthetic MOD water. (d) 1054
47Ti concentrations in zebra mussel exposed for 1 h to waterborne 47TiO2 in the presence of 1 × 106 1055
c/mL of cyanobacteria as food source in synthetic MOD water. (e) and (f) show the 68Zn concentrations 1056
in unsealed and sealed earthworm, respectively, which were exposed to 68ZnO (red squares and line) and 1057
68ZnCl2 (blue circles and line). The solid red or blue lines across the exposure concentrations in a-d 1058
display the mean natural background concentrations of the metals measured in snails that were not 1059
exposed to any NMs. The shaded areas in a and b represent the error for the averaged concentrations, 1060
given as 1× (pink) and 3× (blue) the standard deviation (SD) of the mean (n = 140 in a; n=200 in b). The 1061
dotted lines in d show the SD of the mean (n=28). The open symbols in a and b represent the detectable 1062
49
67Zn and 65Cu (newly accumulated) after exposure to 67ZnO and 65CuO NMs; each concentration was 1063
derived from the total measured 67Zn and 65Cu concentrations minus background. The closed symbols in 1064
a and b represent the sum of the detectable 67Zn and 65Cu, and the background Cu concentrations. Error 1065
bars show SD, n = 10 samples per group. Adapted from previous publications.31,33,36,37 1066
Figure 5. Transformation of 107Ag NMs and 109AgNO3 in an aqueous system. (a) Change of the 109Ag 1067
concentrations in pure water. Error bars show SD of 4 replicates. (b) Fraction of dissolved 109Ag+ over 1068
time in the presence of DOM. (c) Fraction of dissolved 109Ag+ over time. (d) Release of 107Ag ions from 1069
the 107Ag NMs in dark and light conditions. Adapted from Yu et al.30 1070
1071
1072
1073
Precursors
Enriched isotopes
Metal salts
Elemental metal/metal oxides
NMsxM99%
NP-P180396B Valsami-Jones Fig 1
Stable isotope labeling of
nanomaterials
Polyatomic interference
Enrichment level
Price of isotope
Quantity required
Synthesis method
Detection method
NP-P180396B Valsami-Jones Fig 2
a
b
c
d
NP-P180396B Valsami-Jones Fig 3
TiO2 NP
47TiO2 NP
NP-P180396B Valsami-Jones Fig 4
a b
c d
e f
Ag in
flux
into
L. S
tagn
ails
(μg
g-1
day-
1 )
[47Ti
] in
mus
sels
(μg
g-1 )
Total Ag concentration (μg L-1) [47TiO2] NP exposure concentration (μg L-1)
[68Zn
] in
eart
hwor
m (μ
gg-
1 )
[68Zn
] in
eart
hwor
m (μ
gg-
1 )
Hours Hours
b
c d
a
NP-P180396B Valsami-Jones Fig 5