High Resolution Multi-Isotope Imaging Mass Spectrometry (MIMS) Imaging Applications in Stem Cell Biology

Abstract

Multi-isotope imaging mass spectrometry (MIMS) allows the measurement of turnover of molecules within intracellular compartments with a spatial resolution down to 30 nm. We use molecules enriched in stable isotopes administered to animals by diet or injection, or to cells through the culture media. The stable isotopes used are in general ¹⁵N, ¹³C, ¹⁸O, and ²H. For stem cell studies we essentially use ¹⁵N-thymidine, ¹³C-thymidine and ⁸¹Br from BrdU. This protocol describes the practical use of MIMS with specific reference to applications in stem cell research. This includes choice and administration of stable isotope label(s), sample preparation, best practice for high resolution imaging secondary ion mass spectrometry using the Cameca NanoSIMS 50L and methods for robust statistical analysis of label incorporation in regions of interest (ROI).

INTRODUCTION

Multi-isotope imaging mass spectrometry (MIMS) visualizes the incorporation of non-toxic stable isotopes into cells and tissues by combining a state-of-the-art secondary ion mass spectrometer, the use of stable isotope reporters, and intensive computation on samples which can be prepared by any laboratory experienced with electron microscopy sample preparation of cells and tissues. One may think of MIMS as a major leap over autoradiography, due to lateral imaging resolution similar to usual electron microscopy (EM).

This protocol provides a guide to study stem cells by MIMS. We will discuss choice of DNA stable isotope precursors labels, methods of their administration in cells or live animals and important parameters that differ from standard electron microscopy sample preparation protocols. Best practices for MIMS analysis using a Cameca nanoscale SIMS instrument (NanoSIMS 50L) are provided. Useful tips accumulated over our practical experience are also shared in the Commentary section. The protocol may be easily adapted to other biological samples.

STRATEGIC PLANNING

MIMS uses stable isotopes to identify and quantify newly synthesized strands of DNA and can be used in cell or live animal stem cell experiments. It allows the visualization and measurement of stable isotope labeled thymidine incorporation into newly synthesized DNA. Thymidine labeled with different stable isotopes is commercially available (¹⁵N-thymidine, ¹³C-thymidine, ²H-thymidine) or can be custom made (¹⁸O-thymidine). Also, thymidine analogues BrdU and IdU can be analyzed by MIMS by measuring the natural ⁸¹Br and ¹²⁷I signals. Thus, within the same experiment one can pulse serially using ¹⁵N-thymidine, ¹³C-thymidine, ²H-thymidine, ¹⁸O-thymidine and/or with BrdU or IdU. This allows the researcher to study DNA synthesis in cells or tissue from the same experiments at different times in the cell cycle or at different age of animal. Stem cells can be tagged with several types of probes (Fu & Kraitchman, 2010), many of which contain elements not normally present in biological samples that can be directly identified with MIMS (eg. CdSe, ZnSe quantum dots, fluorescent probes containing I). In this protocol, we assume that each laboratory will have their own general cell culture and animal tissue preparation procedures. We will only present steps that are relevant to MIMS analysis. Basic protocol 1 addresses cultured cells and basic protocol 2 live animals. Basic protocol 3 describes the preparation of Si chip substrates required in all MIMS experiments and basic protocol 4 important parameters for high precision NanoSIMS analysis. Basic protocol 5 describes basic data processing and analysis.

BASIC PROTOCOL 1

STABLE ISOTOPE LABELING OF DNA IN CULTURED CELLS

We will describe the preparation of labeled thymidine solution and which stable isotopes that may be used. The choice of stable isotope label(s) should be carefully considered. Stable isotope distributions are studied by examining isotope ratio images. The sensitivity is the ability to measure small excess of label over the natural abundance. A high sensitivity is easier to achieve with isotopes of low natural abundances. For example, ¹⁵N- or ¹⁸O -thymidine (natural abundances 0.36 and 0.20%, respectively) are preferable to ¹³C-thymidine (1.08% natural abundance).

Materials relevant to MIMS

Stable isotope labeled thymidine and/or analogues (BrdU, IdU)

Si chips (5 mm x 5 mm)

1-1. Preparation of thymidine solution

The solubility of thymidine in water is relatively low (50 mg/ml) giving a maximum concentration of 0.2M. A useful trick is to prepare the mother thymidine solution directly in the bottle containing the powder received from the supplier.

Depending upon the experiment, working concentrations of ¹⁵N-thymidine solution will vary depending upon the organ and duration of chase used in the study.

1-2. Cell culturing

Normal cell culturing protocols may be followed. For MIMS analysis more than enough cells are produced when seeded in 35 mm culture dishes.

Use of culture dishes commercially manufactured for optimal cell attachment must be avoided. The surface layer treatment acts as a “sponge” that can retain the stable isotope label after washing steps. If necessary, dishes can be prepared in the lab with attachment factors such as fibronectin. For similar reasons, ultra-low attachment culture dishes that are treated with a hydrogel must be avoided.

For study of DNA replication, a molecule precursor of DNA (in general thymidine) labeled with a stable isotope is added to the culture during the cell cycle, in the log phase of growth or in the lag phase. One can use serial labeling at different times in the cell cycle in a single experiment by using thymidine tagged with different stable isotopes (²H-thymidine, ¹³C-thymidine, ¹⁵N-thymidine, ¹⁸O-thymidine) or thymidine analogues such as BrdU and IdU. All of these are readily available except for ¹⁸O-thymidine. Radiolabels such as ³H-thymidine and ¹⁴C-thymidine can also be used. The labels incorporated into the replicating DNA will all be identified by the NanoSIMS on the same sample. Note that ²H-thymidine or ³H-thymidine should be used with caution because even at room temperature substantial exchange with unlabeled hydrogen in cellular water can occur (Pinson, 1952).

Cells may be fixed, resin embedded and prepared as thin sections or cultured directly on sterilized Si substrate carriers, freeze dried and analyzed as whole cells in the NanoSIMS.

1-2-1. Thin section preparations

Cells can be prepared as thin sections (as in optical microscopy) but deposited on Si chip supports instead of glass slides. Thin section preparations are also suitable for high resolution SEM imaging. Excellent staining for SEM imaging can be found in the literature (Bushong et al., 2015; Deerinck et al., 2010; Katchalski et al., 2018). In general, the atomic elements contained in the stain will not interfere with signals from stable isotopes incorporated in precursor molecules used for MIMS analysis.

1-2-2. Whole mount preparations

Cells can be cultured directly on the Si chips, freeze dried and analyzed as whole mounts. The chips must be sterilized before use. As NanoSIMS analysis peels away (sputters) the sample in slices as thin as a few atomic monolayers, it produces successive isotopic image slices through the cell at the same x-y location. By analyzing whole mounts, we can reconstruct mass resolved 3D images of, for example, DNA distribution within a single nucleus. It does, however, require long durations (days or weeks) of analysis at the NanoSIMS.

1.3. Fixation and Embedding

Cultured cells are fixed in-situ while adhered to the culture dish and lifted off with propylene oxide to maintain cell structure (trypsinizing cells causes them to lose their original shape).

1-3-1 Detaching and embedding cells so that they retain their shape

1-3-1-1. Add sufficient propylene oxide to cover the cells in the culture dish. Cells will detach and float. They should be removed, transferred to a centrifuge tube and soaked in propylene oxide for 20-30 minutes. Centrifuge the cells to obtain a tight pellet. Resuspend in propylene oxide and repeat the 20-30 minute soak followed by centrifugation to a tight pellet.

1-3-1-2. Infiltrate the cell pellet for several hours to overnight (covered) in equal parts epoxy (containing hardener and accelerator) and propylene oxide. The longer the better for embedding. If leaving overnight, place in the refrigerator to prevent the epoxy from becoming too sticky. The next day, let samples warm to room temperature in the hood before embedding.

1-3-1-3. To embed, warm an appropriate amount of 50/50 epoxy/propylene oxide blend (to make it less viscous) in the 60°C vacuum oven for 10-15 minutes. Pour the warmed epoxy into a suitable mold. Let the epoxy sit in the oven 5 minutes before embedding to let air bubbles rise. Immerse infiltrated cell pellet into the epoxy and let sit in a 60°C oven under vacuum until hardened (at least 8 hours).

Choice of embedding resin is important. For epoxy resins, one has to use a cross-linker for polymerization. These cross-linkers contain nitrogen. This fact has to be taken into account when one needs to measure small increments above the natural ¹⁵N/¹⁴N ratio. Alternatively, one can use the acrylic resin LR White® which is hardened without the use of a cross-linker resulting in a resin with lower nitrogen content.

It is also important to avoid cross linking agents when antibodies are used post fixation. The crosslinking agent may occupy the sites where the antibody would otherwise bind.

Note that the thickness of the sections should be around 500 nm. The usual 80-100 nm thick histological sections may be analyzed if correlative TEM/MIMS is required, but they will be quickly sputtered in the NanoSIMS.

1-4. Whole cells prepared by freeze drying

Cells cultured directly on Si chips and analyzed as whole mounts are prepared by freeze drying (Peteranderl & Lechene, 2004; Tang et al., 2014). Improper freeze drying can lead to growth of large ice crystals which can cause morphological cell damage and redistribution of analyte. Thus, samples need to be quenched through the glass transition temperature (−138 °C) of water to convert cellular water to vitreous ice. This may be obtained by quenching in solid nitrogen rather than liquid nitrogen where nitrogen boiling considerably reduces thermal conductivity (solid nitrogen can be obtained by rapidly evaporating liquid nitrogen, for example by pumping liquid nitrogen in a cylindrical dewar by a high capacity mechanical pump). Store the samples in liquid nitrogen. At −80 °C and above the samples will be altered by growth of large crystals at the expense of smaller crystals because the sublimation pressure is higher with smaller radii of curvature. For the same reasons, freeze drying should not be performed with usual commercial equipment at −40 °C, but at −80 °C or below.

To suppress condensation from air water vapor, sample transfer should be done under a cloud of nitrogen. The cloud can be obtained by immersing laboratory wiping paper in liquid nitrogen and waving over the samples.

BASIC PROTOCOL 2

STABLE ISOTOPE LABELING OF DNA IN ANIMALS

Studies in stem cell research on live animals using MIMS introduces complexities that make defining a single protocol impossible. Each lab will have their own research project, to which each MIMS protocol will have to be tailored. Factors to consider are the length of time the stable isotope DNA precursor (thymidine) is administered, the length of chase period to be used, the length of the cell cycle in the organ studied and its variation with age of animal. Surrogate parenting may also be useful in certain cases. Along with ¹⁵N-thymidine, additional DNA replication markers ¹³C-thymidine, ²H-thymidine, ¹⁸O-thymidine, BrdU and IdU can be given at any time during the experiment.

In cell culture, cell cycles may differ in time scales of the order of days. In live animal experiments, the cell cycles in different organs can be different over a time scale of months to years. Cells in the small intestine will turnover every 2-4 days. Cardiomyocytes turnover at a rate of 1%/year in 25 year old humans (Bergmann et al., 2009). This will affect how a MIMS label is best administered to the live animal. For those experiments where the cell cycle is short, the stable isotope labeled DNA precursor can be administered over a few days by injections (intraperitoneal, subcutaneous, or tail veins in rats and mice) and be observed in the stem and daughter cells. Note, however, that injection into muscle can result in inhomogenous temporal distribution of the nucleoside. For experiments where the cell cycle is much longer, delivery of the label using an osmotic pump is more convenient.

The length of chase period will dictate the dose of the labeled DNA precursor required. For example, if the study incorporates a long chase period and the organ of interest has a short cell cycle, higher doses will be required (while remaining below levels that will modify cell cycle) to ensure measurable label in the last generation of cell is achieved.

Examples of MIMS labeling protocols in mice for stem cell studies in mouse intestinal crypt (Steinhauser et al., 2012) and heart (Senyo et al., 2013) give specifics on labeled DNA precursor administration modes, durations and doses.

The following examples are only rough guidelines for studying DNA replication in intestinal crypt stem cells. All parameters (primary and secondary DNA marker doses and time sequences, duration of chases) can be varied and adjusted to the organ studied.

In this section we will present considerations relevant to MIMS analysis of samples prepared from animals administered stable isotope precursor molecules (through diet or injections). Note that the use of stable isotopes may require the same administrative permissions as radiolabels.

Materials relevant to MIMS

Stable isotope labeled thymidine and/or analogues (BrdU, IdU) or stable isotope containing diet (always perform a bulk analysis of the food to determine its stable isotope concentration)

5 mm x 5mm Si chips

2-1. Examples of Labeled Nucleotide Administration in Mice

We will give specifics of DNA labeling of intestinal cells, which have one of the fastest cell renewal rates in the body. The stable isotope labeled thymidine concentrations given are at least one order of magnitude below doses used to modify the cell cycle. They are, however, sufficient for detection by NanoSIMS even if diluted by several rounds of DNA replication during chase. For organs such as the brain, heart, and lung—which practically do not renew or have much slower rates of renewal—parameters will have to be adjusted accordingly.

Specifics regarding thymidine doses, administration mode and duration and chase duration are tailored to answer specific questions. The following examples are only rough guidelines for studying DNA replication in intestinal crypt stem cells.

2-1-1. Example of double labeling starting in-utero

2-1-1-1. Initiate labeling of the female before conception. Administer 500 μg/day of ¹⁵N-labeled thymidine by subcutaneous injection. Upon impregnation, administer 20 μg/hour by osmotic minipump for variable time during pregnancy.

2-1-1-2. Within 24 hours of birth, transfer pups to a surrogate mother that has not received any labeled thymidine.

2-1-1-3. After the desired duration of chase, one may administer 250 μg BrdU to the pups every 6 hours for 24 hours, then euthanize.

2-1-2. Example of triple labeling starting in 4-day old pups

2-1-2-1. Beginning on day 4 of life, administer ¹⁵N-thymidine (50 μg/g) subcutaneously every 12 hours for as long as desired. Then administer BrdU (50 μg/g) subcutaneously every 12 hours for up to more than 10 days.

2-1-2-2. Provide a chase period as desired.

2-1-2-3. Beginning 24 hours prior to euthanization, administer 250 μg of IdU every 6 hours.

2-1-3. Example of double labeling in adult mice

2-1-3-1. Administer ¹⁵N-thymidine at a dose rate of 20 μg/hour via osmotic minipump for 2 weeks.

2-1-3-2. Provide a 5-week chase period (can be shorter or longer, as desired).

2-1-3-3. Administer BrdU via a single 250-μg bolus injection, followed by 20 μg/hour delivered by osmotic minipump for 24 hours prior to sacrifice.

BASIC PROTOCOL 3

PREPARATION OF Si CHIPS, THE GENERAL SAMPLE SUPPORT FOR NANOSIMS ANALYSIS

Materials

5 mm x 5 mm Si chips

Ultrasonic cell disruptor with ^~1/2” probe (eg. Misonix, Sonicator 3000)

600 ml clean beakers (at least 6 total)

Wire mesh sink strainer

Powder free gloves

Fine-tipped Teflon forceps

Sterile Petri dishes

0.2 μm Bottle Top Filter Unit

Magnetic stirrer, magnetic stir bars

Nitrogen gas

Acetone (spectrophotometric grade ≥99.5%)

Methanol (spectrophotometric grade ≥99%)

Isopropanol (spectrophotometric grade ≥99.5%)

Ethanol (spectrophotometric grade ≥99.5%, 200-Proof)

Water, ultra-pure, spectrophotometric grade

Distilled water

3-1. Obtaining Si Chips

Samples to be analyzed by MIMS are supported by Si chips. The Si chips can be ordered or custom diced from a semiconductor grade silicon wafer. We use wafers with <111> direction, with a resistivity of 0.005-0.020 Ohm·cm.

The Si chips need to be extensively cleaned as described in the following section 3-2.

3-2. Cleaning Si chips

3-2-1. Place Si chips in a wire mesh sink strainer.

3-2-2. Place a magnetic stir bar in a clean, 600-ml glass beaker. Hang the strainer with the chips near the bottom of the beaker

3-2-3. Add 200 ml spectrophotometric grade acetone to the beaker. Stir for ½ hour. Sonicate with an ultrasonic cell disruptor at a power of 20 W for 2 minutes. Pour off the acetone.

3-2-4. Add 200 ml spectrophotometric grade methanol to the beaker. Stir for ½ hour. Sonicate with an ultrasonic cell disruptor at a power of 20 W for 2 minutes. Pour off the methanol.

3-2-5. Add 200 ml spectrophotometric grade isopropanol to the beaker. Stir for ½ hour. Sonicate with an ultrasonic cell disruptor at a power of 20 W for 2 minutes. Pour off the isopropanol.

3-2-6. Gently agitate the chips within the strainer in a total of 2 liters of filtered distilled water so that the chips are all well rinsed.

3-2-7. Place the strainer in 200 ml spectrophotometric grade water and stir moderately for half an hour.

3-2-8. Place the strainer in 200 ml spectrophotometric grade ethanol and stir moderately for half an hour.

3-2-9. Transfer the chips to a clean, sterile culture dish.

3-2-10. Cover the chips completely with filtered ethanol.

3-2-11. Individually dry the chips with a stream of dry, filtered nitrogen gas and transfer them to a culture dish, shiny side up.

BASIC PROTOCOL 4

STABLE ISOTOPE ANALYSIS OF DNA REPLICATION IN SINGLE NUCLEI IN A POPULATION OF CELLS WITH NANOSIMS

Note that Cameca Instruments has recently released (March 2021) to all NanoSIMS laboratories a very detailed user manual describing the use of the NanoSIMS (CAMECA Application Laboratory, 2021). Here we provide information gained by the practice of NanoSIMS analysis and MIMS studies of biological samples in general, and DNA replication in particular.

Materials

NanoSIMS “Harvard” holder

Aluminum foil

Powder free gloves

Fine-tipped Teflon forceps

Screwdriver

Stereoscope with camera

Reflected light microscope with DIC capability and camera

Other types of microscopes (eg. fluorescence, electron) are optional

4.1. Mount the Specimens in the NanoSIMS “Harvard” holder

It is imperative to wear laboratory gloves when handling any part that will go into the NanoSIMS analysis chamber.

In this section we describe the mounting of samples in the “Harvard” holder: it can accommodate up to 16, 5 mm x 5mm Si chip sample supports. It is available from Cameca. Other holders (described in the Cameca manual chapter 5.3.3.2.1 “Load”) that accommodate larger sized Si sample supports, to a maximum of eight, are available from Cameca.

4-1-1. In a clean area, for example a laminar flow hood, place a sample holder front face down on clean aluminum foil.

4-1-2. Using the fine tipped forceps, begin placing the Si chips supporting the samples within the insets of the holder windows. Ensure that all four corners of the chip are contacting the inner lip of the inset (Figure 1).

Figure 1:

Loading the “Harvard”” 16 window sample holder. Chips should be placed such that each corner rests on the insert in each window (arrowed).

4-1-3. Map the location of the Si chips in the holder windows. A template as in Figure 2 is very useful.

Figure 2:

Holder windows seen from above numbered from the side of the insertion rod (black dots).

4-1-4. Carefully align the holder component with the springs to the component with the samples. Make sure the two screw holes are aligned and gently push down while tightening the two screws (Figure 3). There should be no gaps between components when fully tightened.

Figure 3:

Left: Holder component with springs to hold samples in place. Right: Final assembly of Harvard holder is achieved by tightening the (2) screws.

4-1-5. Carefully examine the front face of the holder. It is imperative that no chips protrude from the holder in order to protect the objective lens. If there are, disassemble the holder, adjust positions of offending chips and reassemble.

4-1-6. Until analysis, place the sample holder in a heated vacuum oven.

Always include chip(s) with (i) an unlabeled “control” sample and (ii) dried microdroplets of all stable isotope containing media (mother solution, growth media). Also, a patterned Si grid is also helpful to tune the instrument.

If one wants to perform MIMS analysis of TEM ultrathin sections, one can use the adapter for the Cameca “Biology” holder (Cameca manual section 5.3.3.2.1 “Load”) or a resized version can be custom made to fit the “Harvard” holder.

Samples and sample holders must be perfectly dry in the NanoSIMS analysis chamber. They should be kept above ambient room temperature until ready to be loaded (for example in an oven at 50 °C) and in storage. They should never be breathed upon during transport to the NanoSIMS.

Note once the sample holder is in the NanoSIMS analysis chamber, navigation to specific Si chips and the analysis areas on those chips requires accurate documentation. The next sections describe the minimum documentation required.

4-2. Acquire Low Magnification Optical Image of the Sample Holder and Differential Interference Contrast Images of Each Section

It is most useful to have low magnification optical micrographs of the samples to aid navigation in the NanoSIMS. A low magnification image using the NanoSIMS CCD camera is not possible, and so navigation of the sample to the regions of interest can be time consuming and a waste of valuable instrument time.

4-2-1. Acquire an image of the entire sample holder (such as the one shown in Figure 4).

Figure 4:

Low magnification optical micrograph of Harvard holder showing locations of the sections (eg. white arrows) and a Si grid useful for tuning NanoSIMS (red arrow). Five windows were not used (eg. yellow arrow).

4-2-2. Using a reflected light microscope with an automated stage, acquire differential interference contrast (DIC) images. Samples are not always stained for SEM, so DIC can provide suitable contrast to identify specific sample locations. Record the stage positions. It will allow location of samples in the NanoSIMS chamber by a coordinates transformation and reduce time spent navigating to the regions of interest, leaving more time for actual analysis. An example of a DIC image of a longitudinal section through mouse intestinal crypt is shown in Figure 5.

Figure 5:

DIC showing the location of cells of an intestinal crypt with stem cells, amplification zone and terminally differentiated intestinal epithelial cells.

4-3. Starting Cs Source

The procedure for starting the Cs⁺ source is described in the Cameca manual (section 9.1.2.1 “Cs⁺ primary ion source start).

The source will be ready for use when a stable beam current is obtained, measured off-axis by a Faraday cup (Fcp) before all beam focusing optics. The method for measuring the Fcp beam current is described in the Cameca manual (section 9.1.6 “Fcp and Fco beam current checking”). Target values for Cs⁺ ion currents vary from lab to lab, in general ranging from 20-40 nA. Higher currents will speed up analyses but necessitate more frequent source cleanings and replacement. A high Cs⁺ ion current (eg. ~ 80 nA) is also helpful when operating the NanoSIMS at high spatial resolution allowing one sufficient secondary ion currents to facilitate focusing the Cs⁺ beam on the sample

4-4. Stopping Cs source

The procedure for stopping the Cs source is described in the Cameca manual (section 9.1.3.1 “Cs⁺ source shutdown and restart”).

Note that when stopping the source, if the ionizer filament current is turned off first, the Cs will condense on the cold tungsten frit and effectively plug it—sometimes necessitating a change of source.

Note that we do not stop the source when performing several days of consecutive analyses but leave it in “Standby” mode where the 8kV extraction voltage and a reduced heating of the ionizer tip remain applied. This cleans the tip of Cs and speeds up the re-starting and stabilization of the source.

4-5. Load Sample Holder into NanoSIMS 50 Airlock

Attach holder to shuttle (see Cameca manual, section 9.1.1.1 “Sample mounting”). Load holder/shuttle assembly into the airlock (see Cameca manual section 9.1.1.2 “Sample loading in the airlock). Beware that a faulty attached sample may fall inside the NanoSIMS, causing at least two days recovery time.

Samples can be outgassed using the very hot halogen light source of the NanoSIMS airlock. To avoid burning, it is safer to degas biological samples in a vacuum oven around 50 °C for at least overnight before loading into the NanoSIMS.

4-6. Move Sample Holder into the NanoSIMS 50L Vessel Chamber

The process of moving the sample holder to the vessel chamber is described in the Cameca user manual (section 9.1.1.3 “Sample transfer from airlock to vessel chamber). Always ensure that the sample holder is locked, first to the airlock stage, then to the transfer rod.

When the sample is docked into the vessel chamber carousel, retract the transfer rod slowly while observing through the vessel chamber viewport. Indeed the sample holder may remain attached, but not locked, to the transfer rod and at risk of falling. If this is the case, push the holder forward so it is fully back on the carousel and retry to properly retract the transfer rod.

4-7. Move Sample Holder into the NanoSIMS 50L Analysis Chamber

This process is described in the Cameca user manual (section 9.1.1.4 “Sample transfer from the vessel to the analysis chamber”).

Ensure the sample holder is locked in the carousel by the pin on top of the vessel chamber. If the sample is not locked, the sample transfer rod will push the holder off the carousel into the NanoSIMS, causing significant downtime.

4-8. Set NanoSIMS 50L Stage Z-Position

The Z-position set for focusing the CCD camera observation is also the focal point for the secondary ion immersion lens (EOS). Adjusting the Z-position of the sample is therefore critical. The procedure is described in the Cameca manual (section 9.1.5.1 “Moving the sample under optical microscope (CCD)).

4-9. Measure Primary Ion Beam Current Impacting the Sample

Measurement of Fcp (section 4-3) provides a measure of the state of the Cs source. Far more important is the measurement of the primary beam current impinging on the sample (Fco). This value is necessary for ion dose and ion yield calculations. It is a function of several parameters: the value of Fcp, the diameter of the D1 apertures and the L1 lens voltage. It is measured with a Faraday cup (Fco). The procedure is described in the Cameca manual (section 9.1.6 “Fcp and Fco beam current checking”).

Make note of Fco because it is NOT recorded by the NanoSIMS software.

4-10. Record Regions of Interest (ROI) to be Analyzed

Use the NanoSIMS camera to locate the ROIs (Cameca manual section 5.3.1.3 “Getting Started”) or apply a coordinate transformation to ROI coordinates selected by other methods, eg. optical microscopy, DIC, SEM. (Examples: scope2mims, Cameca manual section 6.2 “Point Logger”).

Any additional ROIs should be manually saved to the .prs file.

After one has saved the sample location positions and selected an area to be analyzed, an area on the sample needs to be implanted with cesium to begin tuning the instrument.

4-11. Implant Cs⁺ in Region near Analytical Area for Tuning the Instrument

The overall implantation procedure is described in the Cameca manual (section 9.1.10 “Pre-implantation”). Pay attention to detector damage prevention.

It is necessary to “implant” or deposit a sufficient amount of Cs onto the surface of the sample to elicit, maximize and stabilize secondary ion emission. On biological samples the CN ion count rate is useful for monitoring implantation. Initially, the count rate will increase very slowly with Cs implantation time, followed by a period of exponential growth and finally reaching a constant rate, typically 100’s of thousands of counts when D1 Aperture 2 (Fco ~2.5 pA) and Entrance Slit 3 are used. The dose of Cs⁺ ions required to reach a constant emission of ions is usually between 5 × 10¹⁶ – 5 × 10¹⁷ ions/cm². This will be dependent on many parameters including embedding resin and these dose values should only be used as a rough guide.

4-12. Set Detector Pulse Height Distributions (PHD)

Secondary ions are counted using electron multipliers (EMs). An immense advantage of EM detectors is that—when calibrated properly—they are noiseless and allow single ion counting. The design and principles of EM operation are described in the Cameca manual section 2.1.4.3 (“Electron Multipliers”).

The entire procedure and an example of a proper PHD curve is described in the Cameca manual (section 9.1.12 “Manual EM pulse height distribution (PHD) adjustment”). Check that the C₄X deflector has been properly set (Cameca manual section 9.2.1.4 “Multi-collection”). The PHD should be measured on a sample that has been implanted until the count rate plateaus at a maximum value; this indicates that the sample has been fully implanted.

The distribution of voltage pulses should be centered around 220 mV and is a function of the voltage gain applied to the EM. This is determined by recording the pulse height distribution (PHD) curve and adjusting the EM voltage gain accordingly. The high number of pulses at very low voltages are due to electronic noise and filtered out by setting a threshold voltage below which all pulses are discarded from counting. This is usually around 50 mV.

To practically measure the PHD’s on all detectors, position all movable detectors 2 amu apart from each other at around 450 mm magnet radius. Move to a Si chip and change the magnetic field so that ²⁸Si is measured on detector 1. Measure the PHD and adjust voltage gain and threshold if necessary. Change the magnetic field so that ²⁸Si is measured on detector 2 and measure the PHD and change gain and threshold. Repeat this process for detectors 3-6. The PHD on the fixed detector will need to be set separately. If the fixed detector (described in 4-13, paragraph 4) measures a significantly heavier ion than Si (for example measuring ¹⁹⁷Au), the PHD is best measured from a standard sample containing the same ion species.

After calibrating the EMs, all conditions being the same, the count rate should be equivalent among detectors.

Detector PHDs should be regularly checked or after measuring samples with high count rate.

4-13. Gross Positioning of the Secondary Ion Detectors

In stem cell studies with MIMS, one can identify replicating/replicated nuclei and can measure the number of divisions by using thymidine labeled with stable isotopes ¹³C and/or ¹⁵N, and/or thymidine analogues containing elements ^{79 or 81}Br (BrdU) and ¹²⁷I (IdU) that do not naturally occur in DNA.

Detectors can be positioned, for example, to count ¹²C and ¹³C to measure incorporation of ¹³C-thymidine, ¹²C¹⁴N and ¹²C¹⁵N to measure incorporation of ¹⁵N-thymidine and ⁸¹Br and/or ¹²⁷I for the thymidine analogues. ³¹P is useful as a nuclear marker. Measuring an isotope of Si is helpful to indicate when the sample has been consumed or the location of any holes in the section. By using a variety of DNA markers, one can study DNA replication serially in the same experiment at time points extending from days to months.

Note that we analyze the nitrogen isotopes as the CN⁻ ion. The secondary N⁻ ion yield is very poor and using the CN⁻ ion improves ion yield by several orders of magnitude.

With 6 moveable and one fixed detector, up to seven isotopes can be measured simultaneously in the NanoSIMS 50L. To measure a set of isotopes at the highest mass resolution one should use the lowest magnetic field that allows the heaviest isotope to be measured on the fixed detector. The remaining isotopes are measured by moving the detectors to their appropriate positions along the magnetic radius.

The procedure for positioning the detectors is described below.

4-13-1 Ensure multicollection is selected in both tuning and detection modes (Figure 6, red arrows)

Figure 6:

Tuning window showing location of multicollection tuning and detection modes (red arrows), mass selection window (eg. black arrow), beam on switch (green arrow), NMR regulation switch (brown arrow) and fine detector position (mass) adjustment controls (orange arrows).

4-13-2. In the Tuning/Counting window, enter the heaviest mass (either the value in amu or symbol) into the “Mass” location for detector 7 (Figure 6, black arrow). This will change the magnetic field to the approximate value required to measure these ions in detector 7 and change the mass values at the current locations of the movable detectors. Note it is not obligatory to use all seven detectors.

4-13-3. Turn the beam ON (Figure 6, green arrow).

4-13-4. If you do not see any counts appearing in the fixed detector (detector 7) window, change the magnetic field using the keyboard thumbwheel until counts appear.

4-13-5 Turn “NMR” on (Figure 6, brown arrow). This starts the high precision magnetic field regulation. At this point you should not change the magnetic field.

4-13-6. Enter the mass values of the other isotopes into the “Mass” location for the other detectors. This will move the detector to that position. Be careful to avoid possible detector collisions. It may be necessary to perform this step out of numerical detector order.

4-13-7 If no counts are observed on detectors 1 to 6 after movement, the fine adjustment arrows (Figure 6, orange arrows) may be used to move the detectors in small increments until signal is found. This is a trial and error process.

At this point you should now have counts on all 7 detectors.

Note that the secondary ion column has not yet been aligned, which can greatly enhance count rates.

4-14. Align the Secondary Column

The goal is to maximize the transmission of the secondary ions to the detectors. The procedure for aligning the secondary column is described in the Cameca manual (section 9.1.13 “Optimizing mass analyzer transmission”).

On biological samples a faster method is to maximize the ¹²C¹⁴N⁻ secondary ion current (count rate) using the thumbwheels by iteratively adjusting the triad [EOS-Cy-P₂/P₃] until no further increase in count rate is observed.

4-15. Precise positioning of secondary ion detectors

The full procedure is described in the Cameca manual (section 9.1.14 “Optimizing mass resolving power”).

The HMR spectrum is acquired by scanning the focused secondary ion beam across the slit in front of each EM detector with a pair of voltage plates. This results in a plot of counts as a function of scanning voltage (Figure 7). Because the resolving power of the instrument is not infinite, there may be a combination of elemental and molecular ions that are not separated. For example, the mass spectrum at mass 27 (Figure 7) shows four plateaus indicating the presence of three masses not completely separable by the mass resolving power of the instrument. From low to high mass these are: ¹²C¹⁵N, ¹³C¹⁴N and ¹³C₂¹H. Note that a greater separation between these mass lines could be achieved by using a narrower exit slit (see Cameca manual section 5.2.2 “EM/FC Detector Panel”), which however may create difficulty in peak stability.

Figure 7:

Mass spectrum (HMR) at mass 27. The spectrum shows a plot of detector deflection plate voltage along the abscissa and secondary ion counts along the ordinate. The arrow indicates setting for deflection voltage to measure ¹²C¹⁵N.

4-16. Optimizing Mass Resolving Power MRP using the Quadrupole Lens (Q)

The procedure for optimizing the mass resolving power (MRP) with the quadrupole lens (Q) is described in the Cameca manual (section 9.1.14 “Optimizing mass resolving power”).

The optimum value for Q is dependent upon the magnetic radius. Thus, the best MRP value found by measuring the HMR on detector 7 will not be the same as that measured on detector 1. This discrepancy will be more pronounced the greater the difference in radii between detectors 1 and 7. Thus, Q should be optimized using the mass that requires the highest MRP, or one within a few amu (eg. optimizing Q for best MRP for ¹²C¹⁵N can be done by optimizing Q using ¹²C¹⁴N). In some cases, a compromise value of Q may be required to span the entire mass range of the experiment.

The quadrupole lens has the largest effect on the MRP, but LF₄ also has an effect and should be checked in a manner similar to Q. If the MRP still appears poor, the hexapole may also need to be checked. These procedures are described in the Cameca manual (section 9.3.1.3.2 “Secondary tuning in multicollection mode”). You should be able to get an MRP for ¹²C¹⁴N of at least 5000 using entrance slit 3 and aperture slit 1. If this cannot be attained, it will be impossible to collect good data for mass ¹²C¹⁵N.

4-17. Prepare for Image Acquisition

Basic image acquisition is described in the Cameca manual (section 9.1.15 “Basic image acquisition”).

Before launching the analysis, several acquisition parameters need to be selected. These are:

1. Pixel resolution (eg. 256 x 256, 512 x 512 etc.)

2. Dwell time per pixel

3. Single image plane or multiple image planes?

Single scan images often provide all desired information and for large fields of view this may be the preferred method. Long, single scan images will, however, be prone to drift effects and more susceptible to sample charging. Multiple plane images can provide the data required for 3D information, imaging depth profiles, and a means for drift correction.

The pixel size of the final image should also be considered. One rule of thumb is that the spot size of the primary ion beam should be approximately twice the pixel size. This way, the tissue section or cell is neither undersampled nor heavily oversampled, which will consume the sample more rapidly with no added benefit.

Rather than randomly selecting a number of planes, the importance of counting statistics in MIMS should be considered. These play a large role in precision, which is given (in %) as 100/√N, where N is the number of counts (in a single pixel or ROI). Thus, in a measurement where the weakest counting isotope yields 100 counts, one can expect no better than 10 percent precision in the ratio value. If only small increases in the isotopic ratio values are expected, then the length of the analysis should be sufficient for a precision based on counting statistics to show that regions are indeed statistically significant in enrichment above the natural ratio value. The data can be processed during acquisition to allow a better judgement of how long it will require to achieve the desired counting statistics.

BASIC PROTOCOL 5

DATA REDUCTION AND PROCESSING

In a single plane analysis, the Cs primary beam dwells at each pixel address once and the intensities of the selected secondary ions measured by the detectors are stored in a single “.im” file. In multi-plane analysis, this procedure is repeated up to a thousand times, with data from successive planes also stored in the single .im file.

The data measured at each secondary ion mass can be displayed as a quantitative image and statistically analyzed.

We analyze data from the NanoSIMS using the OpenMIMS plug-in that we developed for ImageJ. A manual describing the general use of OpenMIMS is available here. Here we will provide additional information using examples from stem cells and DNA replication.

5-1. Processing raw NanoSIMS data files into quantitative MIMS images in a study of mouse intestinal stem cells

In the example shown in Figure 8, the mouse received ¹⁵N-thymidine from day 4 until 8 weeks old (day 4 to 4 weeks: two daily subcutaneous injections (50 ug/g); 4 weeks to 8 weeks: osmotic pump at 20 ug/hour), followed by a chase period of four weeks. Prior to harvesting, BrdU was administered for 24 hours (1mg bolus followed with 20 μg/hour by osmotic pump).

Figure 8:

Processing a .im file from a mouse intestine sample. Mouse was treated with 15N-thymidine from day 4 until 8 weeks old (day 4 to 4 weeks: two daily subcutaneous injections (50 ug/g); 4 weeks to 8 weeks: osmotic pump at 20 ug/hour), followed by a chase period of four weeks. Prior to harvesting, BrdU was administered for 24 hours (1mg as a bolus followed with 20μg/hour by osmotic pump). Image field is 30 um x 30 um recorded at 51 2× 512 pixels. Dwell time is 0.75 ms/pixel. Quantitative image data for seven masses is stored for each plane in an image stack containing 271 total planes. Bar: 5 μm.

First image plane for each selected mass in the stack of a multiplane acquisition: (a) ¹²C (b) ¹³C (c) ¹²C¹⁴N (d) ¹²C¹⁵N (e) ³¹P (f) ³²S and (g) ⁸¹Br.

The image stack (containing 271 planes) is examined for bad images that should be discarded. (h) Example of a good ¹²C¹⁴N image plane (i) bad ¹²C¹⁴N image plane showing non-uniform intensity distribution (j) bad ¹²C¹⁴N image plane where top third of image is absent. Inset shows a zoomed image of the Paneth cells marked by red arrow. (h) bad ¹²C¹⁴N image with a zoomed image of the Paneth cells in (j) where the lower cell has clearly changed shape. All planes such as these should be removed from the stack of images. A total of 24 bad planes were found and discarded. The image planes for each mass are now summed providing higher counts at each pixel and better statistical precision.

If blurred sum images, such as those in (l) - (r) are observed, it indicates some spatial drift during the acquisition. The drift can be corrected by applying any necessary x-y translation to the images in the stack. The Autotrack algorithm in OpenMIMS computes and performs the x-y translations, using either the entire image or a selected smaller region of interest containing an object with high contrast (computationally faster). If the pixels in a single plane do not have a high enough count rate, it may be difficult to apply a drift correction and the “Compress” command may be helpful. Its function is to sum a few adjacent planes in the stack. No drift correction will be applied considering that adjacent images will have practically not drifted compared to the entire stack (acquired over several hours). Thus, for a stack of 1000 images and a compression of 10, the new stack will be of 100 images with ~ 10 times more counts per image and it may then be possible to achieve a good drift correction on the compressed stack.

After drift correction, much sharper sum images are observed: (s) ¹²C sum image (t) ¹³C sum image (u) ¹²C¹⁴N sum image (v) ¹²C ¹⁵N sum image (w) ³¹P sum image (x) ³²S sum image (y) ⁸¹Br sum image.

5-2. Isotope Ratio Images

Deriving isotope ratios from the quantitative image data obtained by the NanoSIMS allows one to determine regions of the sample, if any, that have incorporated the stable isotopes. As the information content of a grayscale image is poor, we developed a method based on a hue saturation intensity transformation of the ratio image, originally conceived by the Tsien lab (Poenie et al., 1986) for expression and fluorescence data. In an HSI image, the hue codes for the ratio value, and the intensity at each hue codes for the statistical precision. As we will show, HSI images provide a wealth of information at a glance.

Here, we had administered the DNA precursor thymidine enriched in ¹⁵N to mice for intestinal stem cell studies. The HSI images allow one to quickly observe which cells within the intestinal crypt have incorporated ¹⁵N into their DNA, how that DNA is distributed within the nuclei, and estimate the number of DNA replications. The process of producing an HIS image is described here and summarized in Figure 9.

Figure 9:

Images from same mouse experiment described in Figure 8. (a) ¹²C¹⁴N image (b) ¹²C¹⁵N image (c) ¹²C¹⁵N/¹²C¹⁴N ratio image obtained by simple pixel by pixel division of counts. We prefer to display ratio images using a Hue-Saturation-Intensity (HSI) transformation (d) where the hue codes for the ratio value, with the minimum (blue) representing the natural value, and the intensity at each hue codes for the statistical precision. Higher intensities reflect areas with higher counting statistics. (e) ¹²C¹⁵N/¹²C¹⁴N HSI ratio image. Two cells within and one outside the crypt are observed to be labeled.

5-3. Region of Interest (ROI) Analysis

Region of interest (ROI) analysis is performed to extract numerical values of the incorporation of the label into tables for statistical analysis. These regions can be individual cells in a population, subcellular structures in cells, and areas within the ratio image where one visually suspects label incorporation (brightness of un-normalized mass images of the label cannot be used because of matrix effects).

Regions of interest are recorded within OpenMIMS using the ROI manager (here) and the numerical parameters are selected using the tomography window (here).

An example of ROI analysis is shown in Figures 10–12. Twelve regions of interest were drawn around chromosomes, guided by the ³¹P image (Figure 10c). We selected the 12 ROIs shown in Figure 10 using the OpenMIMS ROI manager and from each ROI we extracted the values for: (i) the ¹²C¹⁵N/¹²C¹⁴N ratio (mean and standard deviation), (ii) the Σ¹²C¹⁵N counts / Σ¹²C¹⁴N counts (N/D; Numerator / Denominator) and (iii) the Br counts (mean and standard deviation), using the OpenMIMS tomography window. If there is a discrepancy between the mean of the ratios and the ratio of the mean, it suggests insufficient counting statistics or highly inhomegenous labeling within the ROI. In Figure 11 we show the table of output from OpenMIMS. It can then be exported and analyzed using a statistical software like JMP (Figure 12).

Figure 10:

Chromosomes in the nucleus of an intestinal crypt cell of a 12 week old mouse. At 10 weeks the mouse received ¹⁵N-thymidine by osmotic pump at 20 ug/hour, followed by the thymidine analogue BrdU for 24 hours before sacrifice (1mg as bolus followed with 20μg/hour by osmotic pump): (a) HSI of ¹²C¹⁵N/¹²C¹⁴N (b) ¹²C¹⁵N/¹²C¹⁴N (c) ³¹P and (d) ⁸¹Br. Field of analysis: 10 μm x 10 μm.

Figure 12:

Data from Figure 11 is exported into JMP software for further analysis. An inverse linear relationship (R-squared = 0.80) between the mean ⁸¹Br counts and the mean ¹²C¹⁵N/¹²C¹⁴N ratio is observed. Blue shading represents 95% confidence levels for fitted line.

Figure 11:

Table output from the 12 ROIs (red column) drawn in Figure 12 showing data for (Blue) the mean and standard deviation of Br counts measured at each pixel, (Green) the mean and standard deviation of ¹²C¹⁵N/¹²C¹⁴N ratio value measured at each pixel and (Yellow) the Σ¹²C¹⁵N counts / Σ¹²C¹⁴N counts (N/D; Numerator/Denominator).

COMMENTARY

Background Information

Until the middle 1930’s, it was believed that our constituents were permanent until death. Rudolf Schoenheimer left Germany in 1933 and joined the Department of Biochemistry of Columbia University. In 1934, he started collaborating with Harold C. Urey, who had isolated deuterium. Schoenheimer developed the method of isotope tagging of biomolecules. In ‘The Edward K. Dunham Lectures for the Promotion of the Medical Sciences 1941’ at Harvard University, he mentioned “Heavy nitrogen (N¹⁵), generously made available by Dr. Urey, has made possible a new technique for the investigation of protein metabolism.” and concluded that “all constituents of living matter, whether functional or structural, of simple or complex constitution, are in steady state of rapid flux.” (Schoenheimer, 1942)

These studies could not be pursued at the subcellular level because there was no radioactive isotope of nitrogen that could be used for autoradiography. Imaging of stable isotope tracers at the subcellular level only became possible with the development of secondary ion mass spectrometry (SIMS) quantitative imaging by Castaing and Slodzian (Castaing & Slodzian, 1962). Major advances in SIMS capability led by Slodzian and Hillion (Slodzian et al., 1992) included a spatial resolution smaller than 50 nm and parallel collection of secondary ions, allowing one to measure isotope ratios at the submicron level. These developments provided the scientific community with an instrument having all the necessary capabilities to extend the pioneering work of Schoenheimer to the subcellular level (C. Lechene et al., 2006; McMahon et al., 2006).

The development of MIMS provided a powerful new way of viewing and measuring, in all fields of biomedical research, important parameters that were previously impossible or difficult to see or measure. We have used MIMS to show for the first time how symbiotic bacteria convert atmospheric nitrogen to a form suitable for biosynthesis (C. P. Lechene et al., 2007). MIMS allowed us to study protein renewal in sensory hair cell stereocilia (Zhang et al., 2012) and resolve “the immortal strand hypothesis” in the mouse intestinal crypt, where direct evidence against non-random template strand segregation in the small intestine was observed (Steinhauser et al., 2012). It has provided new insight into biological problems in a variety of domains including the source and generation of new heart cells in mammals (Senyo et al., 2013), the antioxidant role of lipid droplets in stem cell niches (Bailey et al., 2015) and led to the proposition that age mosaicism across multiple scales is a fundamental principle of adult tissue, cell, and protein complex organization (Toyama et al., 2019). Other applications include studies in the fields of environmental microbiology and soil science (Eichorst et al., 2015; Finzi-Hart et al., 2009; Heister et al., 2012; Milucka et al., 2015; Pasulka et al., 2018; Wolfe-Simon et al., 2011) and numerous other biological and pharmaceutical related interrogations (Cabin-Flaman et al., 2011; Guerquin-Kern et al., 2005; Legin et al., 2014; Lovrić et al., 2017; Richter et al., 2017; Vujic et al., 2018).

Critical Parameters

Before Analysis

1. Choose an appropriate stable isotope label dose in the culture media for cell preparations or food or injections for live animals. Factors to consider are both the absolute concentration of the label introduced into the cell culture media or into the live animal and the number of atoms in the molecule (eg., thymidine) that have been replaced by the enriching isotopic label (eg., ¹⁵N).

2. Be aware of isotopic effects (eg. relative mass differences much greater for ²H and ¹H compared to ¹³C and ¹²C so biological activity may be affected when ²H is used as a label) and exchange effects between heavy isotope labels of H and O with those atoms in water molecules that can lead to a washing of the label.

3. Ensure drops of the labeled culture media are placed on Si chips using micropipettes and loaded into the sample holder. This is particularly important for elements not commonly used as labels.

4. Always verify that natural ratios are measured after tuning of the NanoSIMS using an unlabeled control sample

5. Having optical microscopy and/or SEM images of regions of interest will be very useful when the sample is in the NanoSIMS.

Tuning of the NanoSIMS 50 Before Analysis

1. It is essential to ensure the pulse height distributions of the detectors are properly set (see section 4.12). It is essential to balance the count rates (section 4.12) on the set of detectors used to measure isotope ratio or ratios.

2. Alignment of the Cs source can be done using the Si signal from any Si chip upon which the samples are deposited.

3. It is essential that the peaks in the HMR mass spectra contain flat tops and sharp shoulders (see section 4.16 and Figure 8).

NanoSIMS Analysis of Samples

1. Do not implant the sample with a high Cs beam current (no diaphragm, D1-0). At least D1-1 must be used. Use of high beam currents will rapidly consume the sample.

2. Be aware that during implantation, and particularly if D1-0 is used, isotope label and cellular material may temporarily attach to the objective lens and possibly redeposit on the sample during analysis, relocating isotope labels. This is known to occur in some dynamic SIMS instruments (sometimes referred to as the memory effect). We have not observed translocation of ¹⁵N-thymidine from the nucleus to the cytoplasm, which would be an indicator that this problem is occurring. When implanting samples we never use the highest beam currents (D1-0), and this precaution seems to be sufficient to avoid this problem.

3. Ensure that the NMR regulation is on during analysis. If it has switched off, stop the analysis, switch the regulation back on, and re-check the HMR mass spectra before continuing analysis.

4. The analysis may be stopped when sufficient statistical data in the regions of interest have been acquired.

After Analysis of Samples

1. After processing the data from the first experiment in an analysis campaign, create a tracking document with the raw image data and ratio images. There are features on OpenMIMS to make this easy. Add to this document after each subsequent analysis. This will greatly facilitate future data analysis and manuscript writing.

Troubleshooting:

NanoSIMS 50L analysis

Problem	Possible Cause	Solution(s)
No secondary ion signals	Primary ion source not on. Primary ion beam badly out of alignment or gonio not in proper position. Problems with voltages on lenses or deflectors. Gate valve at Cs source or multicollector chamber is closed. No voltages applied to the detectors. Secondary ion beam grossly out of alignment.	Ensure proper water circulation in cooling circuit. Verify beam currents both in primary column (Fcp) and at sample (Fco). Move to a bare Si area and attempt to detect 28Si counts. Remove all slits, if necessary. Use “Update Parameters” command to look for large differences in applied and measured voltages at various lenses/deflectors. Look for signal at total ion current detector to narrow down in which part of the instrument the problem may be occurring.
Incorrect value of natural isotopic ratio from standards	Detectors are not properly calibrated and PHDs have not been verified. Secondary ion beam is misaligned, such that the field is non-uniformly balanced with CN signal or C signal across the entire field of view. Control sample was contaminated or improperly labeled during sample mounting. C4x deflector not properly set at the plateau for all masses. Wrong peak selected in the HMR or mass resolution insufficient.	Check PHDs on the electron multipliers and balance the count rate on the detectors. Check secondary alignments. Re-check HMRs to ensure proper peak has been selected and sufficient mass resolving power is attained. Check C4x scans.
Higher than natural ratios are measured on control samples though all NanoSIMS 50L instrument parameters are found to be properly set.	Cross contamination during preparation, possibly through pipette tip.	Casts suspicion on all samples for this culture experiment. Redo culturing and do not waste instrument time on dubious samples.
Sample charging	Sample has picked up moisture. Sample is intrinsically non-conductive.	In our experience, we exceptionally encounter sample charging by taking simple, yet extreme precautions to keep samples ultra-dry. They are stored in a vacuum oven at 50 °C. When transferred from the oven to the analytical chamber of the NanoSIMS, they are never allowed to be at lower than room temperature. We take extreme precaution to avoid breathing upon them. In the rare case we do encounter charging, we found that implanting Cs from the Si chip to the periphery of the tissue section creates a conductive pathway that can help dissipate charge. Apply 5 nm conductive Au or Pt to sample. An evaporator is preferred over a sputter system as there is a possibility that energetic Ar+ ions could sputter the sample and translocate material if the samples are placed too close to the plasma region.

Statistical Analysis:

MIMS provides results that are incredibly rich in statistical data, and the OpenMIMS plugin itself offers many statistical parameters to be analyzed and exported (see section 5.4). The data extracted from the OpenMIMS plugin can also be easily imported to other statistical analysis software packages.

Time Considerations:

The time frame for completing a MIMS analysis can vary immensely. The sample labeling, especially in live models, can take just a few days to months. This would be followed by a period of several days for the actual preparation of the samples. MIMS image data could be acquired as quickly as a few minutes or up to several days. There are many determining factors including required level of spatial resolution in x,y and z dimensions as well as the statistical precision required and level of label incorporation. High spatial resolution acquisitions necessitate use of low primary ion beam currents which reduces the secondary ion signals and sputter rate. The only recourse to retain high levels of statistical precision is to extend the acquisition time accordingly. Considering the experiment will contain a standard control sample, perhaps different label concentrations and/or time points, it is not unreasonable for the entire MIMS analysis campaign to be of 10-14 days duration and perhaps longer. Finally, data processing and analysis can easily (and should) require 2-3 times the analysis time but can be done off-line while other analyses are being acquired.

For correlative microscopy studies (and given the destructive nature of the MIMS technique), the NanoSIMS 50L analysis must be the last imaging modality used. Days to weeks may also be required for the acquisition of the correlative images (SEM, fluorescence, TEM).

DATA AVAILABILITY STATEMENT:

The data that support the findings of this study are available from the corresponding author upon reasonable request.