This project has received funding from the European Union Horizon 2020 Programme (H2020) under grant agreement no. 720952 (ACEnano). The authors would like to thank Sigrid Benemann for SEM measurements, Markus Schneider for ToF-SIMS measurements and PCA, and Philipp Reichardt for assistance with filming.

CAUTION: The toxicological properties of nanoparticles are still under investigation; due to their size they can present unique hazards in humans as well as in the environment even when they consist of intrinsically non-hazardous materials. Before undertaking any work with nanoparticles, a proper risk assessment should be completed, and appropriate engineering controls, lab procedures, and PPE (personal protective equipment) put in place, depending on the hazard level of the materials to be studied29,30,31,32.
1. Preparation of Si wafers
NOTE: These steps are necessary to remove undesired (organic) contamination and increase the surface wettability. All solvents used should be at least ACS grade. A standard sonication bath (35 kH and 120 Watts) is suitable.
<ol>
	<li>Wet chemical cleaning of Si wafers
	<ol>
		<li>Put the Si wafer in a beaker with isopropanol and ultrasonicate for 5 min.</li>
		<li>Transfer the Si wafer to a beaker with an alkali glass cleaning solution and ultrasonicate for 10 min.</li>
		<li>Put the wafer in a beaker with ultrapure water. Change the water 10 times by pouring out the water and refilling the beaker; the Si wafers will remain at the bottom due to the capillary effect.</li>
		<li>Dry the wafer with clean N2 gas. 
		NOTE: Drying with N2 prevents the formation of &#8220;coffee rings&#8221; and other artefacts from water drying.</li>
		<li>Put the wafer in a second beaker with isopropanol and ultrasonicate for 10 min.</li>
		<li>Dry the wafer with clean N2 gas.</li>
		<li>Put the wafer in a beaker with ethanol and ultrasonicate for 10 min.</li>
		<li>Dry the wafer with clean N2 gas. The protocol can be paused here.</li>
	</ol>
	</li>
	<li>Plasma or UV/ozone cleaning of silicon wafers
	<ol>
		<li>Introduce the Si wafer in the plasma or the UV/ozone cleaner and switch on for 30 min. 
		NOTE: Wafers should be plasma- or UV/ozone-cleaned immediately before use.</li>
	</ol>
	</li>
</ol>
2. Nanoparticle deposition from suspension
NOTE: The most common exposure route for nanoparticles is by inhalation. Working with suspensions can minimize exposure hazards.
<ol>
	<li>Preparation of nanoparticle suspension from powder 
	NOTE: All quantities described here are examples. The method should be optimized for the particular nanoparticles used in each case.
	<ol>
		<li>Accurately weigh 15 mg of nanoparticle powder (&#177; 10%) into a 10 mL tube.</li>
		<li>Accurately weigh in approximately 8 mL ultrapure water.</li>
		<li>Close the tube, pack in a 50 mL centrifuge tube with paper towel and place in the vortexer at 3,000 rpm for 15 min.</li>
	</ol>
	</li>
	<li>Drop-casting of electrically conductive nanoparticles from aqueous suspension
	<ol>
		<li>Place the wafer in the UV/ozone cleaner for 30 min.</li>
		<li>Place the wafer in one half of the wafer holder and place a 3 &#181;L drop of nanoparticle suspension in the center of the ring.</li>
		<li>Mount a 6.07 mm diameter Viton O-ring on the wafer around the droplet. Take care that the ring does not touch the droplet.</li>
		<li>Place the wafer in a vacuum desiccator under a vacuum of 4 mbar for 15 min to dry the wafer.</li>
		<li>Remove the wafer from the desiccator and examine using light microscopy and XPS to determine that the particle layer is homogeneous and closed. Repeat steps 2.2.1 and 2.2.2 until analysis shows a closed and homogeneous layer. The protocol can be paused here.</li>
	</ol>
	</li>
	<li>Spin-coating of electrically non-conductive nanoparticles from aqueous suspension
	<ol>
		<li>Place the wafer in the UV/ozone cleaner for 30 min. 
		NOTE: By spin-coating suspensions of different concentrations using the same protocol, different levels of surface coverage can be achieved.</li>
		<li>Program the spin-coater. A suitable sample program is: step 1: 500 rpm/s ramp to 1,000 rpm (5 s); step 2: 1,000 rpm/s ramp to 2,000 rpm (3 min); step 3: deceleration at 2,000 rpm/s to 0 rpm.</li>
		<li>Insert the wafer into the spin-coater and switch on the vacuum for fixation.</li>
		<li>Deposit 80 &#181;L of the suspension on the wafer and start the program.</li>
		<li>Remove the wafer from the spin-coater.</li>
		<li>Store the sample in a new, clean wafertray. The protocol can be paused here.</li>
		<li>Analyze the sample using SEM to confirm gapless coverage of the substrate.</li>
	</ol>
	</li>
</ol>
3. Nanoparticle deposition from powder
<ol>
	<li>Nanoparticle deposition on double-sided adhesive tapes (&#8220;stick and go&#8221;)
	<ol>
		<li>Fix the double-sided adhesive to the sample holder and remove the liner.</li>
		<li>Take a spatula-tip of the nanoparticle powder and dip it onto the adhesive.</li>
		<li>Spread the sample over the adhesive and press into the adhesive with the spatula, until as much of the powder is adhered as possible.</li>
		<li>Check that the powder is fixed on the tap by inverting and tapping the sample holder, and by blowing a stream of gas (e.g., nitrogen) across it. The protocol can be paused here. 
		NOTE: Alternatively, a small amount of powder can be placed on a cleaned surface (Alu foil or glass slide) and pressed from above with the adhesive and double-sided sample holder.</li>
		<li>Place a spatula-tip of the powder onto the cleaned surface. Press the sample holder with the adhesive onto the powder from above.</li>
		<li>Check that the powder is fixed on the tap by inverting and tapping the sample holder, and by blowing a stream of gas (e.g., nitrogen) across it. The protocol can be paused here.</li>
	</ol>
	</li>
	<li>Preparation of pressed powder pellets
	<ol>
		<li>Thoroughly clean all parts of the pellet die, taking care not to scratch the polished surface.</li>
		<li>Invert the pellet die and rest on a small spacer.</li>
		<li>Insert the plunger and one stainless steel pellet, with polished side up, and pull the plunger through until there is enough space to fill with the powdered sample.</li>
		<li>Fill the die with a small amount of sample (1 large spatula tip), and then insert the second stainless steel pellet with the polished side facing the sample.</li>
		<li>Place the base onto the body and carefully invert. If a vacuum is desired and available, attach the vacuum pump to the base of the pellet die.</li>
		<li>Place the die into a press, making sure it is centered.</li>
		<li>Apply a light load (2 kN) for approximately 20 s and release.</li>
		<li>Apply a heavier load (6 kN) for 2 min and release.</li>
		<li>After the load is released, release the vacuum pump. 
		NOTE: Due to the different material properties of various nanoparticles, it may be advantageous to prepare a series of pellets with different loads and load times to determine the optimum pellet pressing conditions.</li>
		<li>Invert the die, place the extractor ring into position, and place a light load (up to 1 kN) between the plunger and the extractor ring.</li>
		<li>Remove the die parts from the press and carefully extract the sample pellet with tweezers.</li>
		<li>Gently mount the sample on a cleaned Si wafer using double-sided adhesive. The protocol can be paused here.</li>
	</ol>
	</li>
</ol>
4. Cryofixation of nanoparticle suspensions
<ol>
	<li>Fill the main chamber of the fast-freeze device with liquid nitrogen.</li>
	<li>Fill the cooled fast-freeze chamber with the cryogen (propane).</li>
	<li>Allow the fast-freeze device to cool to its operating temperature. 
	NOTE: The fast-freeze device requires some time to reach operating temperature prior to the sample preparation, therefore a reasonable timeframe (a few hours) is required for cryofixing the samples.</li>
	<li>Drop-cast 10&#8211;20 &#181;L of NP suspension onto a cleaned Si wafer with a pipette.</li>
	<li>Holding the Si wafer with fixing tweezers, place it inside the plunge freeze device.</li>
	<li>Move the fixing tweezers to the plunge position.</li>
	<li>Press the button to drop the sample inside the cryogen.</li>
	<li>Wait several seconds until the sample is completely frozen.</li>
	<li>Transfer the frozen samples as fast as possible into a cooled environment.</li>
	<li>Place the cryofixed sample (Si wafer) into the sample holder and transfer it inside the instrument. 
	NOTE: For transport, dry ice is recommended and short-term sample storage is possible. The samples can be measured in a frozen state with a cooled instrument or with conventional ToF-SIMS settings after stabilization by freeze-drying the sample.</li>
</ol>

The authors have no competing interests to disclose.

A number of methods have been presented for the preparation of nanoparticles for surface analysis using XPS and ToF-SIMS. We have summarized the advantages and disadvantages of these methods, as well as possible sources of error and suitability for different materials, in Table 2. As shown in the representative results, the preparation of nanoparticles can strongly influence the success of the resulting surface analysis. In addition, not all methods are suitable for all particle types due to factors such as signal interference with the substrate or mounting materials, charging effects in non-conducting thick films, state of the nanoparticles as a powder or suspension, potential damage to sensitive outer layers, destruction of biological structures and information on aggregation and interfaces, or vulnerability of sensitive ultrahigh-vacuum instruments to free nanoparticles.
Because XPS and ToF-SIMS measurements average over an area rather than measuring single particles, it is only possible to obtain reproducible results from homogeneous layers; aggregation or agglomeration of the particles on the substrate should therefore be avoided. Additionally, too-thick layers of non-conductive materials cause charging effects during analysis, which can lead to undesired artefacts in the spectra, especially partial charging which cannot be compensated with a flood gun. On the other hand, incomplete films show strong signals from the substrate or mounting materials (e.g., adhesives), which can interfere with sensitive peaks from the particle surface. The ideal thickness of the film is material dependent and should be determined experimentally by analysis of films of different thicknesses. In particular, samples prepared using spin coating should be analyzed with SEM to ensure completeness of the coating.
Working with NP suspensions presents fewer exposure hazards and safety requirements compared to working with NP powders. Drop-casting is a relatively simple method with low equipment requirements and is particularly suitable for conductive nanoparticles in suspension where film thickness is not a concern. While the samples can easily be dried under atmospheric conditions, the vacuum desiccator serves to reduce the drying time for the droplets as well as protecting the wafers from contamination. The Viton ring is used to modify the evaporation patterns of the droplet and thereby minimize the formation of coffee-rings. Evaporation patterns can also be influenced by varying the substrate hydrophilicity using cleaning protocols or by application of alternative coatings51,52, by evaporating in solvent atmospheres53, or even by heating the substrate54. Spin-coating is recommended for suspensions of non-conductive nanoparticles in suspension because it is capable of generating a homogeneous particle layer that is thin enough to avoid charging effects but still thick enough to prevent the Si substrate from contributing to the XPS and ToF-SIMS spectra. For each individual NP system and concentration, both the centrifuge and spin-coating parameters must be optimized but can then very reliably be reproduced even on different instruments. Because the spin-coated drop is always in the middle of the wafer, the radius of rotation is irrelevant and the unit &#8220;revolutions per minute&#8221; (rpm) can be used. The suspension could alternatively be deposited on the wafer after starting the program; however, this would require different spin-coating parameters and a greater amount of suspension to obtain a thicker coating.
Because of their extremely small size, nanoparticles may detach from the substrate and move freely inside the ultrahigh-vacuum chamber when impacted with an ion or x-ray beam. This is a particular problem for samples prepared with powder. In some cases, the nanoparticles can penetrate into the sensitive components of the instrument requiring expensive and time-consuming maintenance. Due to the applied acceleration voltage, the danger of damaging sensitive parts is larger with ToF-SIMS than with XPS. Powdered samples, particularly those prepared using the &#8220;stick and go&#8221; method, should be carefully checked to ensure the powders are fixed securely enough, especially for ToF-SIMS analysis. This can be confirmed by, for example, holding the sample upside down and blowing a stream of gas (e.g., N2) across it. Prior to analysis, the samples can also be left overnight in the airlock or other initial sample entry chamber of the instrument, where a stable vacuum can indicate that there are no loose particles from the sample. Nanoparticles prepared as pellets, however, can even be sputtered (at low acceleration voltages) without damaging the instrument; this method can eliminate contaminants, particularly hydrocarbons, introduced from the press and can also enable bulk analysis of the particles.
Preparation of NP powders in the sample holder stub allows for the preparation of samples with defined geometry and a macroscopically flat surface. Critical points are the cleanliness of the tool for pressing the sample, and the use of a low pressure to avoid changes in the nanoparticle surface due to this procedure. It has the disadvantages of needing a relatively high amount of material, and potential problems with loss of material in high-vacuum instruments. We do not recommend this method for ToF-SIMS analysis, as the particles are not compressed or secured in any way.
Regarding the NP material, the first consideration for sample preparation is elimination or minimizing of interference between NPs and substrates of similar material; for example, Si wafers are an unsuitable substrate for analysis of SiO2 NPs using XPS and ToF-SIMS, even with sufficient sample coverage. Metallic or inorganic nanoparticles can be readily analyzed as powder on an adhesive (assuming they contain no organic layers or coatings) due to the lack of signal interference between the nanoparticles and the double-sided adhesive, a preparation method which would be unsuitable for polymeric NPs. Metallic nanoparticles have more flexibility in terms of possible film thickness used due to the absence of charging effects, and may be drop-cast with relatively little equipment; however, they are likely to contain large amounts of impurities and stabilizers from their synthesis, which must be carefully removed without damage to the particles. Polymeric nanoparticles may more easily be damaged by die pressing but may also more readily hold together in the pellet, depending on the pressures used. Pellets or soft organic coatings on the NP surface also&#160;can be damage sensitive. Direct deposition from the solution has the potential to damage sensitive coatings either through the suspension or the drying process but is advantageous for analyzing NPs already present in suspension. Cryofixation is a suitable method for the analysis of chemical structures, surfaces or interfaces in suspension which would be damaged or destroyed by various other sample preparation techniques, but requires a specialized cryoequipment for both XPS and ToF-SIMS46&#8217;47.
While this paper describes several exemplary methods that can be used for sample preparation, in every case the method should be optimized and validated using alternative analytical methods. A detailed overview of the influence of different factors was recently published22. Besides the development and validation of suitable preparation methods, the documentation of these steps is also of paramount importance40. This publication presents some easy-to-handle methods and is a guide to modify or develop new methods according to the requirements of the specific task.

Nanomaterials are defined as materials having any external dimension between 1 nm and 100 nm or having an internal or surface structure on this scale1. Due to the unique properties arising from their small scale and correspondingly large surface area (among other factors), they find increasing use in a wide variety of fields including agriculture, chemistry, automotive construction, cosmetics, environment, medicine, printing, energy, and textiles. This increased use means that both humans and the environment will be exposed, on a hitherto unknown scale, to these materials whose toxicological properties are not yet fully known, and whose size enables their facile integration into biological or environmental systems2.
After the fundamental properties of surface area and particle size/size distribution, surface chemistry and coatings were identified as the most crucial property of nanomaterials3; smaller particles have a higher surface area per unit mass, and therefore a higher ratio of surface to bulk atoms. Indeed, for nanoparticles of 1 nm size, over 70% of atoms can be found at corners or edges; this strongly influences surface properties such as chemisorption which is highly dependent on the atomic-scale surface morphology4. Regulations dealing with nanomaterials require accurate data regarding physicochemical properties and reliable estimates of the toxicological properties of these materials. In order to efficiently estimate toxicological properties from physical and chemical properties of nanomaterials, the nanomaterials community requires reliable, standardized, and verified analytical procedures. Projects such as ACEnano5 aim to collect and correlate accurate and verifiable physical data from nanoparticles in a framework allowing better regulation and characterization of nanomaterials. This drive towards standardized analytical procedures has also been supported by the editors of ACS Nano, wishing &#8220;to consolidate and to agree on methods of characterization and minimum levels of analysis of materials6&#8221;. Furthermore, XPS and ToF-SIMS offers new possibilities for elucidating the particle architecture of core-shell nanoparticles7,8.
X-ray photoelectron spectroscopy (XPS) and Time-of-Flight Secondary Ion Mass Spectrometry (ToF-SIMS), compared in Table 1, are well-established methods for investigation of surface atoms. In XPS, the sample is irradiated with x-rays having an energy between 1 and 2 keV, causing emission of electrons due to the photoelectric effect. These emitted electrons, having a kinetic energy in the same range, correlate to the binding energy of the electrons in the solid; the appearance of photoelectrons at these defined binding energies and measurable intensities therefore allows quantitative analysis of the composition. Since the mean free pathway of these photoelectrons is below 10 nm, XPS is a highly surface sensitive technique for quantitative analysis. Furthermore, detailed analysis of the binding energies in highly resolved spectra enables the quantitative determination of the valence states of these electrons.
In ToF-SIMS the surface is sputtered with a focused ion beam (primary ions), with the ions ejected from the material (secondary ions) collected and analyzed in a time-of-flight mass spectrometer. The mass/charge pattern obtained allows the determination of the elemental, isotopic, or molecular composition. Due to the mean free pathway of the secondary ions, this technique is also highly surface sensitive and has an information depth of 1&#8211;2 nm but is at best semi-quantitative, due to the matrix effect whereby the ionization probability (and therefore yield) of secondary ions is strongly influenced by their surrounding matrix. ToF-SIMS can be operated in either static or dynamic mode; the difference between the two is the primary ion flux impacting the surface. Static SIMS keeps the primary ion flux to a level that impacts (i.e., fragments) a maximum of 1%-10% of the surface; the surface remains relatively undisturbed, which allows analysis of the top atomic layers of material. Since even static SIMS causes some destruction to the surface, it is considered to be less &#8220;non-destructive&#8221; of the two methods.
These surface-sensitive techniques allow analysis of the first few nanometers of the material, including intentional or unintentional coatings, which, for nanomaterials, can significantly influence material properties. Examples of intentional coatings are capping layers on quantum dots to improve photoluminescence quantum yields and reduce environmental reactivity9, alumina or silica coatings for the prevention of photocatalytic activity of titania nanoparticles in sun blockers10, surface functionalization to enable bioconjugation and subsequent biological activity11, coatings for diagnostic and drug delivery applications12, and fluorocarbon coatings on magnetic particles for ferrofluids and core-shell metallic systems to enhance catalyst properties13. Unintentional coatings, such as oxidation, surface contamination, or protein coronas in biological systems have a similarly strong influence on nanoparticle properties and it is crucial that experimental preparation procedures ensure that the coating and more generally the surface chemistry of the nanomaterial is not destroyed or transformed. It is also crucial to evaluate the properties of the nanoparticles as they are in-situ, as their properties can be drastically altered by the change2,14,15. In addition, the concentration of stabilizers in the nanoparticle suspension can dramatically influence the analysis and structural integrity of the nanoparticles; the presence of a stabilizer can result in large unwanted signals (for example, C, H, O, and Na) in the analysis, while its removal can result in damage or agglomeration of the nanoparticles.
Due to their size and surface area, the storage conditions of nanoparticles also affect their behavior, both as stored powders/suspensions and as prepared samples. The effect of sub-optimal storage conditions, particularly room-temperature storage and exposure to light, have been shown in various studies to cause degradation of the nanoparticles which has been shown to alter the particles&#8217; physical, chemical, and/or toxicological properties14,15,16,17,18. Smaller nanoparticles have been shown to oxidize more rapidly than larger ones with oxidation/degradation rates dependent on storage conditions15 as well as surface chemistry14. The effects of the nanoparticle degradation during storage have been shown to significantly affect physicochemical properties including toxicity14, while the oxidative growth can proceed inwards at the expense of the core15.
The careful storage and preparation of nanomaterials is therefore essential for an accurate surface analysis, and any factors which could influence the sample surface and/or the quality of the measurements should be carefully considered. It should be noted that due to the relatively low spatial resolution of XPS (in the &#181;m range) and ToF-SIMS (a few hundred nm), only a small subset of the nanoparticles can be investigated; these methods average over an area and do not have the ability to image single particles as is possible with techniques such as electron microscopy. For this reason, any analysis requires deposition of the nanoparticles in a continuous layer to ensure no interference from the substrate. Electron microscopy and XPS/ToF-SIMS are therefore often used together as complementary methods for nanomaterial analysis.
Aside from changes in surface chemistry, the main challenges for preparation of nanoparticle samples for XPS and ToF-SIMS analysis are to prepare a layer that is: homogeneous, to increase reproducibility; gapless, to minimize the contribution of the substrate to the spectra; thin enough to avoid charging effects (for non-conductive samples); and securely fixed to the substrate, to avoid free nanoparticles entering and damaging ultrahigh vacuum instruments
Nanoparticles can be deposited onto the substrate from suspension or as a powder. Firstly, we will discuss the different methods for depositing nanoparticles from suspension. Silicon wafers are a commonly used substrate for suspension deposition, because they are relatively cheap, readily available as a highly pure product consisting of pure or doped silicon (doping avoids charging effects), and for most nanoparticles the spectral peaks do not overlap with peaks typical for nanoparticles. This last point is important; before analysis it should be ensured that the substrate peaks are well separated from the peaks expected from the nanoparticles, otherwise interpretation of the spectra is complicated or impossible and the continuous coverage of the substrate by the nanoparticles cannot be verified. Before using silicon wafers, an extensive cleaning procedure (described in this publication) is necessary to remove (organic) contaminants and to increase the surface wettability. Other suitable substrates such as gold films, highly ordered pyrolytic graphite (HOPG), or indium foils have been successfully used, but a discussion about their preparation is beyond the scope of this work19,20,21,22.
Secondly, we present methods for depositing nanoparticle powders on a substrate for XPS and ToF-SIMS analysis and present the advantages and disadvantages of each method, allowing researchers new to the techniques to find the optimal preparation method for their purposes. Thirdly, we discuss cryofixation, which is a suitable preparation method to conserve features such as the agglomeration behavior, organic corona, solid/aqueous interface23,24 or distribution in biological media25 of NPs. Cryofixation, typically fast freezing of material in a liquid nitrogen&#8211;cooled cryogen and analysis in the frozen-hydrated state, allows the analysis and visualization of nanoparticles directly in complex matrices. This procedure does not cause ice crystal formation but forms amorphous ice that keeps membranes and cellular and tissue structures in their native biological state, avoiding damage caused by water crystallization processes and enabling the exact chemical distribution of all cell metabolites and cell membrane compounds to be maintained26,27,28. This preparation method may be of particular interest for presenting an exact chemical map of the actual NP agglomerate or heteroagglomerate, visualizing the exact chemical space in close proximity to the nanoparticle directly in suspension, or correlating either cell tissue-specific features or intra-cellular compartments within NP agglomerates or heteroagglomerates.
As shown through the results presented in this work, the most suitable procedure in a particular case is dependent on a variety of parameters such as the nanoparticles&#8217; hydrophilicity, stability, conductivity, state (e.g., powder or suspension) and the analytical question at hand (e.g., size, bulk properties, or surface coatings). A variety of methods are presented here that can be used for preparation of NPs for surface analysis, as well as a comparison of their advantages and disadvantages.

<table>
	<tbody>
		<tr>
			<td>4-figure Laboratory balance</td>
			<td>Kern &#38; Sohn GmbH</td>
			<td>ADB200-4A</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mm Pellet die</td>
			<td>Specac</td>
			<td>GS03060</td>
			<td></td>
		</tr>
		<tr>
			<td>Alkali glass cleaning solution</td>
			<td>Sigma-Aldrich</td>
			<td>Hellmanex&#8482; III Z805939</td>
			<td>Special cleaning solution for cuvettes</td>
		</tr>
		<tr>
			<td>Carbon adhesive tabs</td>
			<td>Plano</td>
			<td>&#34;Leit-Tabs&#34; G3347</td>
			<td></td>
		</tr>
		<tr>
			<td>Clean laboratory beakers</td>
			<td>any</td>
			<td></td>
			<td>e.g. 300 mL</td>
		</tr>
		<tr>
			<td>Cryo-freezer</td>
			<td>Electron Microscopy Sciences</td>
			<td>EMS-002 Cryo Workstation</td>
			<td></td>
		</tr>
		<tr>
			<td>Dialysis tube with fasteners</td>
			<td>Medicell Membranees Ltd</td>
			<td>DTV12000.06.30</td>
			<td>Molecular weight cut-off (MWCO) 12-14 kDa</td>
		</tr>
		<tr>
			<td>Die press</td>
			<td>any</td>
			<td></td>
			<td>Capable of 2 kN force</td>
		</tr>
		<tr>
			<td>Disposable syringe, 1 mL, Luer-slip</td>
			<td>TH Geyer</td>
			<td>Labsolute 7657545</td>
			<td>Any appropriate volume can be used</td>
		</tr>
		<tr>
			<td>Double-sided adhesive</td>
			<td>3M</td>
			<td>Removable Repositionable Tape 665</td>
			<td></td>
		</tr>
		<tr>
			<td>Dry ice</td>
			<td>Linde AG</td>
			<td>ICEBITZZZ&#174;</td>
			<td>For short term storage/cooling</td>
		</tr>
		<tr>
			<td>Eppendorf transfer pipette and tips</td>
			<td>Eppendorf</td>
			<td>various</td>
			<td>Check correct size for planned pipetting volume</td>
		</tr>
		<tr>
			<td>Ethanol, ACS grade</td>
			<td>Merck KGaA</td>
			<td>1009832500</td>
			<td></td>
		</tr>
		<tr>
			<td>FFP2 or FFP3 mask</td>
			<td>various</td>
			<td></td>
			<td>For working with nanoparticles from non-hazardous materials, when not in a fume hood or glove box</td>
		</tr>
		<tr>
			<td>Isopropanol, ACS grade</td>
			<td>Merck KGaA</td>
			<td>1096342500</td>
			<td></td>
		</tr>
		<tr>
			<td>Lab coat, gloves and goggles</td>
			<td>any</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory centrifuge</td>
			<td>Eppendorf</td>
			<td>Centrifuge 5430</td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory fume hood</td>
			<td>any</td>
			<td></td>
			<td>necessary for working with nanoparticles</td>
		</tr>
		<tr>
			<td>Laboratory stirrer &#38; stirrer bar</td>
			<td>NeoLab</td>
			<td>D-6010</td>
			<td></td>
		</tr>
		<tr>
			<td>Lint-free wipes</td>
			<td>Kimberley Clark Professional</td>
			<td>Kimtech Science Precision wipes</td>
			<td>Recommended for working with Si wafers</td>
		</tr>
		<tr>
			<td>Liquid Nitrogen</td>
			<td>Linde AG</td>
			<td>Stickstoff fl&#252;ssig 5.0</td>
			<td>Only for cooling of the cryogen.</td>
		</tr>
		<tr>
			<td>Microtube/centrifuge tube 1,5 mL</td>
			<td>T.H. Geyer GmbH &#38; Co. KG</td>
			<td>Labsolute 7696751</td>
			<td></td>
		</tr>
		<tr>
			<td>Nitrogen 5.0</td>
			<td>any</td>
			<td></td>
			<td>99.999% purity</td>
		</tr>
		<tr>
			<td>Pasteur pipette, PE, plastic 3 mL</td>
			<td>TH Geyer</td>
			<td>Labsolute 7 691 203</td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur pipette, PE, plastic 3 mL</td>
			<td>TH Geyer</td>
			<td>Labsolute 7 691 203</td>
			<td></td>
		</tr>
		<tr>
			<td>Powder sample holder</td>
			<td>BAM workshop</td>
			<td></td>
			<td>&#34;Home-made&#34; sample holder</td>
		</tr>
		<tr>
			<td>Propane</td>
			<td>Sigma-Aldrich</td>
			<td>769037</td>
			<td>The cryogen should be of highest possible purity.</td>
		</tr>
		<tr>
			<td>Sample vial or centrifuge tube 1 mL</td>
			<td>Greiner Bio-One GmbH</td>
			<td>Cellstar 188 261</td>
			<td>Should be capable of being fixed in the Vortexer</td>
		</tr>
		<tr>
			<td>Silicon wafers</td>
			<td>any</td>
			<td></td>
			<td>ideally 1cm2 pre-cut</td>
		</tr>
		<tr>
			<td>Spin-coater</td>
			<td>SPS Europe</td>
			<td>SPIN150i-NPP</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe filter 0,45 &#181;m</td>
			<td>Th Geyer</td>
			<td>Labsolute 7699803</td>
			<td>For smaller samples; larger versions exist for larger sample volumes</td>
		</tr>
		<tr>
			<td>ToF-SIMS</td>
			<td>IONTOF GmbH</td>
			<td>ToF-SIMS IV or V, equipped with Bi LMIG and flood gun</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers for handling Si wafers</td>
			<td>any</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ultrapure water</td>
			<td>TKA</td>
			<td>MicroPure 08.1202</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonicator</td>
			<td>Bandelin</td>
			<td>Sonorex Super</td>
			<td></td>
		</tr>
		<tr>
			<td>UV/Ozone cleaner</td>
			<td>NanoBioAnalytics</td>
			<td>UVC-1014</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum dessicator</td>
			<td>any</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum pump (membrane/diaphragm)</td>
			<td>Vacuubrand GmbH</td>
			<td>Type MD-4T</td>
			<td></td>
		</tr>
		<tr>
			<td>Viton O-ring 6.07 x 1.78 mm</td>
			<td>Betech GmbH</td>
			<td>2-010, FKM 80</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortexer</td>
			<td>Heathrow Scientific</td>
			<td>Vortexer HS120212</td>
			<td></td>
		</tr>
		<tr>
			<td>Wafer Holder 25mm coin style</td>
			<td>Semiconductor Production Systems Europe</td>
			<td>eWB0091-ASSY-1</td>
			<td></td>
		</tr>
		<tr>
			<td>XPS</td>
			<td>Kratos</td>
			<td>Kratos Axis Ultra DLD</td>
			<td></td>
		</tr>
	</tbody>
</table>

preparation of nanoparticles for tof-sims and xps analysis

Nanoparticles have gained increasing attention in recent years due to their potential and application in different fields including medicine, cosmetics, chemistry, and their potential to enable advanced materials. To effectively understand and regulate the physico-chemical properties and potential adverse effects of nanoparticles, validated measurement procedures for the various properties of nanoparticles need to be developed. While procedures for measuring nanoparticle size and size distribution are already established, standardized methods for analysis of their surface chemistry are not yet in place, although the influence of the surface chemistry on nanoparticle properties is undisputed. In particular, storage and preparation of nanoparticles for surface analysis strongly influences the analytical results from various methods, and in order to obtain consistent results, sample preparation must be both optimized and standardized. In this contribution, we present, in detail, some standard procedures for preparing nanoparticles for surface analytics. In principle, nanoparticles can be deposited on a suitable substrate from suspension or as a powder. Silicon (Si) wafers are commonly used as substrate, however, their cleaning is critical to the process. For sample preparation from suspension, we will discuss drop-casting and spin-coating, where not only the cleanliness of the substrate and purity of the suspension but also its concentration play important roles for the success of the preparation methodology. For nanoparticles with sensitive ligand shells or coatings, deposition as powders is more suitable, although this method requires particular care in fixing the sample.

Watch this Scientific Journal Video about Preparation of Nanoparticles for ToF-SIMS and XPS Analysis at JoVE.com

Preparation of Nanoparticles for ToF-SIMS and XPS Analysis

A number of different procedures for preparing nanoparticles for surface analysis are presented (drop casting, spin coating, deposition from powders, and cryofixation). We discuss the challenges, opportunities, and possible applications of each method, particularly regarding the changes in the surface properties caused by the different preparation methods.

Nanoparticles have gained increasing attention in recent years due to their potential and application in different fields including medicine, cosmetics, ...

preparation-of-nanoparticles-for-tof-sims-and-xps-analysis

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7.6K Views.  Federal Institute for Material Research and Testing (BAM). A number of different procedures for preparing nanoparticles for surface analysis are presented (drop casting, spin coating, deposition from powders, and cryofixation). We discuss the challenges, opportunities, and possible applications of each method, particularly regarding the changes in the surface properties caused by the different preparation methods.

Video: Preparation of Nanoparticles for ToF-SIMS and XPS Analysis

Preparation of Silica Nanoparticles Through Microwave-assisted Acid-catalysis

Microwave-assisted synthetic techniques were used to quickly and reproducibly produce silica nanoparticle sols using an acid catalyst with nanoparticle diameters ranging from 30-250 nm by varying the reaction conditions. Through the selection of a microwave compatible solvent, silicic acid precursor, catalyst, and microwave irradiation time, these microwave-assisted methods were capable of overcoming the previously reported shortcomings associated with synthesis of silica nanoparticles using microwave reactors. The siloxane precursor was hydrolyzed using the acid catalyst, HCl. Acetone, a low-tan &#948; solvent, mediates the condensation reactions and has minimal interaction with the electromagnetic field. Condensation reactions begin when the silicic acid precursor couples with the microwave radiation, leading to silica nanoparticle sol formation. The silica nanoparticles were characterized by dynamic light scattering data and scanning electron microscopy, which show the materials&#39; morphology and size to be dependent on the reaction conditions. Microwave-assisted reactions produce silica nanoparticles with roughened textured surfaces that are atypical for silica sols produced by St&#246;ber&#39;s methods, which have smooth surfaces.

Silica nanoparticles were prepared using acid-catalysis of a siloxane precursor and microwave-assisted synthetic techniques resulting in the controlled growth of nanomaterials ranging from 30-250 nm in diameter. The growth dynamics can be controlled by varying the initial silicic acid concentration, time of the reaction, and temperature of reaction.

Microwave-assisted synthetic techniques were used to quickly and reproducibly produce silica nanoparticle sols using an acid catalyst with nanoparticle ...

Isolation and Preparation of Bacterial Cell Walls for Compositional Analysis by Ultra Performance Liquid Chromatography

The bacterial cell wall is critical for the determination of cell shape during growth and division, and maintains the mechanical integrity of cells in the face of turgor pressures several atmospheres in magnitude. Across the diverse shapes and sizes of the bacterial kingdom, the cell wall is composed of peptidoglycan, a macromolecular network of sugar strands crosslinked by short peptides. Peptidoglycan&#8217;s central importance to bacterial physiology underlies its use as an antibiotic target and has motivated genetic, structural, and cell biological studies of how it is robustly assembled during growth and division. Nonetheless, extensive investigations are still required to fully characterize the key enzymatic activities in peptidoglycan synthesis and the chemical composition of bacterial cell walls. High Performance Liquid Chromatography (HPLC) is a powerful analytical method for quantifying differences in the chemical composition of the walls of bacteria grown under a variety of environmental and genetic conditions, but its throughput is often limited. Here, we present a straightforward procedure for the isolation and preparation of bacterial cell walls for biological analyses of peptidoglycan via HPLC and Ultra Performance Liquid Chromatography (UPLC), an extension of HPLC that utilizes pumps to deliver ultra-high pressures of up to 15,000 psi, compared with 6,000 psi for HPLC. In combination with the preparation of bacterial cell walls presented here, the low-volume sample injectors, detectors with high sampling rates, smaller sample volumes, and shorter run times of UPLC will enable high resolution and throughput for novel discoveries of peptidoglycan composition and fundamental bacterial cell biology in most biological laboratories with access to an ultracentrifuge and UPLC.

The bacterial cell wall is composed of peptidoglycan, a macromolecular network of sugar strands crosslinked by peptides. Ultra Performance Liquid Chromatography provides high resolution and throughput for novel discoveries of peptidoglycan composition. We present a procedure for the isolation of cell walls (sacculi) and their subsequent preparation for analysis via UPLC.

The bacterial cell wall is critical for the determination of cell shape during growth and division, and maintains the mechanical integrity of cells in the ...

Preparation and Characterization of SDF-1&#945;-Chitosan-Dextran Sulfate Nanoparticles

Chitosan (CS) and dextran sulfate (DS) are charged polysaccharides (glycans), which form polyelectrolyte complex-based nanoparticles when mixed under appropriate conditions. The glycan nanoparticles are useful carriers for protein factors, which facilitate the in vivo delivery of the proteins and sustain their retention in the targeted tissue. The glycan polyelectrolyte complexes are also ideal for protein delivery, as the incorporation is carried out in aqueous solution, which reduces the likelihood of inactivation of the proteins. Proteins with a heparin-binding site adhere to dextran sulfate readily, and are, in turn, stabilized by the binding. These particles are also less inflammatory and toxic when delivered in vivo. In the protocol described below, SDF-1&#945; (Stromal cell-derived factor-1&#945;), a stem cell homing factor, is first mixed and incubated with dextran sulfate. Chitosan is added to the mixture to form polyelectrolyte complexes, followed by zinc sulfate to stabilize the complexes with zinc bridges. The resultant SDF-1&#945;-DS-CS particles are measured for size (diameter) and surface charge (zeta potential). The amount of the incorporated SDF-1&#945; is determined, followed by measurements of its in vitro release rate and its chemotactic activity in a particle-bound form.

Preparation and Characterization of SDF-1&amp;#945;-Chitosan-Dextran Sulfate Nanoparticles

The objective of this protocol is to incorporate SDF-1&#945;, a stem cell homing factor, into dextran sulfate-chitosan nanoparticles. The resultant particles are measured for their size and zeta potential, as well as the content, activity, and in vitro release rate of SDF-1&#945; from the nanoparticles.

Chitosan (CS) and dextran sulfate (DS) are charged polysaccharides (glycans), which form polyelectrolyte complex-based nanoparticles when mixed under ...

Preparation of Mica and Silicon Substrates for DNA Origami Analysis and Experimentation

The designed nature and controlled, one-pot synthesis of DNA origami provides exciting opportunities in many fields, particularly nanoelectronics. Many of these applications require interaction with and adhesion of DNA nanostructures to a substrate. Due to its atomically flat and easily cleaned nature, mica has been the substrate of choice for DNA origami experiments. However, the practical applications of mica are relatively limited compared to those of semiconductor substrates. For this reason, a straightforward, stable, and repeatable process for DNA origami adhesion on derivatized silicon oxide is presented here. To promote the adhesion of DNA nanostructures to silicon oxide surface, a self-assembled monolayer of 3-aminopropyltriethoxysilane (APTES) is deposited from an aqueous solution that is compatible with many photoresists. The substrate must be cleaned of all organic and metal contaminants using Radio Corporation of America (RCA) cleaning processes and the native oxide layer must be etched to ensure a flat, functionalizable surface. Cleanrooms are equipped with facilities for silicon cleaning, however many components of DNA origami buffers and solutions are often not allowed in them due to contamination concerns. This manuscript describes the set-up and protocol for in-lab, small-scale silicon cleaning for researchers who do not have access to a cleanroom or would like to incorporate processes that could cause contamination of a cleanroom CMOS clean bench. Additionally, variables for regulating coverage are discussed and how to recognize and avoid common sample preparation problems is described.

Reproducible cleaning processes for substrates used in DNA origami research are described, including bench-top RCA cleaning and derivatization of silicon oxide. Protocols for surface preparation, DNA origami deposition, drying parameters, and simple experimental set-ups are illustrated.

The designed nature and controlled, one-pot synthesis of DNA origami provides exciting opportunities in many fields, particularly nanoelectronics. Many of ...

In Situ Characterization of Hydrated Proteins in Water by SALVI and ToF-SIMS

This work demonstrates in situ characterization of protein biomolecules in the aqueous solution using the System for Analysis at the Liquid Vacuum Interface (SALVI) and time-of-flight secondary ion mass spectrometry (ToF-SIMS). The fibronectin protein film was immobilized on the silicon nitride (SiN) membrane that forms the SALVI detection area. During ToF-SIMS analysis, three modes of analysis were conducted including high spatial resolution mass spectrometry, two-dimensional (2D) imaging, and depth profiling. Mass spectra were acquired in both positive and negative modes. Deionized water was also analyzed as a reference sample. Our results show that the fibronectin film in water has more distinct and stronger water cluster peaks compared to water alone. Characteristic peaks of amino acid fragments are also observable in the hydrated protein ToF-SIMS spectra. These results illustrate that protein molecule adsorption on a surface can be studied dynamically using SALVI and ToF-SIMS in the liquid environment for the first time.

In Situ Characterization of Hydrated Proteins in Water by SALVI and ToF-SIMS

This work presents a protocol for liquid handling and sample introduction to a microchannel for in situ time-of-flight secondary ion mass spectrometry analysis of protein biomolecules in an aqueous solution.

This work demonstrates in situ characterization of protein biomolecules in the aqueous solution using the System for Analysis at the Liquid Vacuum ...

In Situ Detection and Single Cell Quantification of Metal Oxide Nanoparticles Using Nuclear Microprobe Analysis

Micro-analytical techniques based on chemical element imaging enable the localization and quantification of chemical composition at the cellular level. They offer new possibilities for the characterization of living systems and are particularly appropriate for detecting, localizing and quantifying the presence of metal oxide nanoparticles both in biological specimens and the environment. Indeed, these techniques all meet relevant requirements in terms of (i) sensitivity (from 1 up to 10 &#181;g.g-1 of dry mass), (ii) micrometer range spatial resolution, and (iii) multi-element detection. Given these characteristics, microbeam chemical element imaging can powerfully complement routine imaging techniques such as optical and fluorescence microscopy. This protocol describes how to perform a nuclear microprobe analysis on cultured cells (U2OS) exposed to titanium dioxide nanoparticles. Cells must grow on and be exposed directly in a specially designed sample holder used on the optical microscope and in the nuclear microprobe analysis stages. Plunge-freeze cryogenic fixation of the samples preserves both the cellular organization and the chemical element distribution. Simultaneous nuclear microprobe analysis (scanning transmission ion microscopy, Rutherford backscattering spectrometry and particle induced X-ray emission) performed on the sample provides information about the cellular density, the local distribution of the chemical elements, as well as the cellular content of nanoparticles. There is a growing need for such analytical tools within biology, especially in the emerging context of Nanotoxicology and Nanomedicine for which our comprehension of the interactions between nanoparticles and biological samples must be deepened. In particular, as nuclear microprobe analysis does not require nanoparticles to be labelled, nanoparticle abundances are quantifiable down to the individual cell level in a cell population, independently of their surface state.

In Situ Detection and Single Cell Quantification of Metal Oxide Nanoparticles Using Nuclear Microprobe Analysis

We describe a procedure for the detection of chemical elements present in situ in human cells as well as their in vitro quantification. The method is well-suited to any cell type and is particularly useful for quantitative chemical analyses in single cells following in vitro metal oxide nanoparticles exposure.

Micro-analytical techniques based on chemical element imaging enable the localization and quantification of chemical composition at the cellular level. ...

Determining the Chemical Composition of Corrosion Inhibitor/Metal Interfaces with XPS: Minimizing Post Immersion Oxidation

An approach for acquiring more reliable X-ray photoelectron spectroscopy data from corrosion inhibitor/metal interfaces is described. More specifically, the focus is on metallic substrates immersed in acidic solutions containing organic corrosion inhibitors, as these systems can be particularly sensitive to oxidation following removal from solution. To minimize the likelihood of such degradation, samples are removed from solution within a glove box purged with inert gas, either N2 or Ar. The glove box is directly attached to the load-lock of the ultra-high vacuum X-ray photoelectron spectroscopy instrument, avoiding any exposure to the ambient laboratory atmosphere, and thus reducing the possibility of post immersion substrate oxidation. On this basis, one can be more certain that the X-ray photoelectron spectroscopy features observed are likely to be representative of the in situ submerged scenario, e.g. the oxidation state of the metal is not modified.

A protocol to avoid the oxidation of metallic substrates during sample transfer from an inhibited acidic solution to an X-ray photoelectron spectrometer is presented.

An approach for acquiring more reliable X-ray photoelectron spectroscopy data from corrosion inhibitor/metal interfaces is described. More specifically, ...

In Situ Characterization of Shewanella oneidensis MR1 Biofilms by SALVI and ToF-SIMS

Bacterial biofilms are surface-associated communities that are vastly studied to understand their self-produced extracellular polymeric substances (EPS) and their roles in environmental microbiology. This study outlines a method to cultivate biofilm attachment to the System for Analysis at the Liquid Vacuum Interface (SALVI) and achieve in situ chemical mapping of a living biofilm by time-of-flight secondary ion mass spectrometry (ToF-SIMS). This is done through the culturing of bacteria both outside and within the SALVI channel with our specialized setup, as well as through optical imaging techniques to detect the biofilm presence and thickness before ToF-SIMS analysis. Our results show the characteristic peaks of the Shewanella biofilm in its natural hydrated state, highlighting upon its localized water cluster environment, as well as EPS fragments, which are drastically different from the same biofilm's dehydrated state. These results demonstrate the breakthrough capability of SALVI that allows for in situ biofilm imaging with a vacuum-based chemical imaging instrument.

In Situ Characterization of Shewanella oneidensis MR1 Biofilms by SALVI and ToF-SIMS

This article presents a method for growing a biofilm for in situ time-of-flight secondary ion mass spectrometry for chemical mapping in its hydrated state, enabled by a microfluidic reactor, System for Analysis at the liquid Vacuum Interface. The Shewanella oneidensis MR-1 with green fluorescence protein was used as a model.

Bacterial biofilms are surface-associated communities that are vastly studied to understand their self-produced extracellular polymeric substances (EPS) ...

Ligand-Mediated Nucleation and Growth of Palladium Metal Nanoparticles

The size, size distribution and stability of colloidal nanoparticles are greatly affected by the presence of capping ligands. Despite the key contribution of capping ligands during the synthesis reaction, their role in regulating the nucleation and growth rates of colloidal nanoparticles is not well understood. In this work, we demonstrate a mechanistic investigation of the role of trioctylphosphine (TOP) in Pd nanoparticles in different solvents (toluene and pyridine) using in situ SAXS and ligand-based kinetic modeling. Our results under different synthetic conditions reveal the overlap of nucleation and growth of Pd nanoparticles during the reaction, which contradicts the LaMer-type nucleation and growth model. The model accounts for the kinetics of Pd-TOP binding for both, the precursor and the particle surface, which is essential to capture the size evolution as well as the concentration of particles in situ. In addition, we illustrate the predictive power of our ligand-based model through designing the synthetic conditions to obtain nanoparticles with desired sizes. The proposed methodology can be applied to other synthesis systems and therefore serves as an effective strategy for predictive synthesis of colloidal nanoparticles.

The main goal of this work is to elucidate the role of capping agents in regulating the size of palladium nanoparticles by combining in situ small angle x-ray scattering (SAXS) and ligand-based kinetic modeling.

The size, size distribution and stability of colloidal nanoparticles are greatly affected by the presence of capping ligands. Despite the key contribution ...

Synthesis and Characterization of Amphiphilic Gold Nanoparticles

Gold nanoparticles covered with a mixture of 1-octanethiol (OT) and 11-mercapto-1-undecane sulfonic acid (MUS) have been extensively studied because of their interactions with cell membranes, lipid bilayers, and viruses. The hydrophilic ligands make these particles colloidally stable in aqueous solutions and the combination with hydrophobic ligands creates an amphiphilic particle that can be loaded with hydrophobic drugs, fuse with the lipid membranes, and resist nonspecific protein adsorption. Many of these properties depend on nanoparticle size and the composition of the ligand shell. It is, therefore, crucial to have a reproducible synthetic method and reliable characterization techniques that allow the determination of nanoparticle properties and the ligand shell composition. Here, a one-phase chemical reduction, followed by a thorough purification to synthesize these nanoparticles with diameters below 5 nm, is presented. The ratio between the two ligands on the surface of the nanoparticle can be tuned through their stoichiometric ratio used during synthesis. We demonstrate how various routine techniques, such as transmission electron microscopy (TEM), nuclear magnetic resonance (NMR), thermogravimetric analysis (TGA), and ultraviolet-visible (UV-Vis) spectrometry, are combined to comprehensively characterize the physicochemical parameters of the nanoparticles.

Amphiphilic gold nanoparticles can be used in many biological applications. A protocol to synthesize gold nanoparticles coated by a binary mixture of ligands and a detailed characterization of these particles is presented.

Gold nanoparticles covered with a mixture of 1-octanethiol (OT) and 11-mercapto-1-undecane sulfonic acid (MUS) have been extensively studied because of ...