Sample Preparation for Single Cell Mass Spectrometry Metabolomics Studies: Combined Cell Washing, Quenching, Drying, and Storage

Cell-to-cell heterogeneity has emerged as a fundamental principle of biological systems, especially in the contexts of disease progression, drug resistance, and cellular differentiation^1,2. In multicellular organisms, even genetically identical cells in the same microenvironment can exhibit distinct phenotypes, arising from differences at the transcriptomic³, proteomic⁴, and, crucially, metabolomic levels⁵. While single-cell genomics and transcriptomics have gained substantial traction in the past decade, single-cell metabolomics remains comparatively underdeveloped due to the inherent complexity and rapid dynamics of metabolites⁶. Metabolites are highly sensitive to environmental cues, have diverse chemical structures and polarities⁷, and exist at low concentrations in small cell volumes⁸. These properties make their reliable measurement from individual cells an analytical challenge. As metabolomics offers the closest snapshot of a cell's real-time functional state, advancing methods to robustly study the metabolome at single-cell resolution is essential for a deeper understanding of cellular physiology and pathology⁷.

Traditional bulk metabolomics techniques average signals across large cell populations, masking cellular heterogeneity and potentially obscuring rare but significant metabolic signatures. In contrast, single-cell metabolomics reveals unique biochemical profiles at the individual cell level, making it invaluable for studying processes like stem cell differentiation, drug responses, and disease mechanisms^{6,9,10,11,12,13}. Mass spectrometry (MS)-based methods, such as matrix-assisted laser desorption-ionization (MALDI) MS and secondary ion (SI) MS, have advanced the field but face challenges like complex preparation, vacuum requirements, and destructive ionization that limit live-cell analysis^9,14. Particularly, recent developments in instrumentation resulted in cutting-edge techniques, including transmission-mode MALDI coupled with laser-induced postionization (t-MALDI-2¹⁵) and Orbitrap secondary ion mass spectrometry (OrbiSIMS)^16,17, enabled label-free, high-resolution 3D metabolic imaging at the single-cell and subcellular level, offering exceptional mass accuracy and spatial resolution.

Ambient ionization MS addresses these issues. The representative ambient SCMS techniques include live single-cell video mass spectrometry (Video-MS)¹⁸, DESI¹⁹, and nano-DESI²⁰. Among them, the single-probe SCMS is one of the techniques that stands out for its ability to directly sample and ionize live cells in real time using a dual-capillary system^21,22,23,24. This technique minimizes sample disturbance while enabling high-resolution, in situ metabolic profiling and has been successfully applied to studies of drug uptake and influence on cell metabolism^{22,25,26,27,28,29,30}, environmental responses^31,32, and metabolic heterogeneity^33,34,35,36 among individual cells.

While many ambient SCMS methods enable live cell analysis, they typically suffer from low throughput due to the manual handling required for individual cell selection^23,37,38. Since cell metabolism is highly dynamic, extended sample preparation can alter metabolite profiles. To address this, researchers apply quenching techniques immediately after isolating cells^39,40. Quenching halts metabolic activity either by rapid cooling^41,42,43,44 or enzyme denaturation^43,45,46,47, preserving the cells' biochemical state at a specific moment. This step is vital for ensuring accurate and temporally relevant metabolomic data. An effective quenching protocol must rapidly and thoroughly stop intracellular metabolic activity. Various methods have been evaluated, including cold isotonic saline⁴⁸, chilled acetonitrile⁴⁶, cold methanol^44,47,48,49, ice-cold phosphate buffer solution (PBS)^43,50, LN₂^43,44,51, and even hot air treatments⁵². While many of these were originally developed for bulk cell metabolomics, some, like cold methanol and acetonitrile, have been adapted for single-cell techniques such as Pico-ESI-MS³⁹ and MALDI-MS⁴⁶. Despite their effectiveness, each method has drawbacks: organic solvents may cause metabolite leakage and damage to cell membranes^48,53, and nonvolatile salt solutions can interfere with MS by causing ion suppression⁵⁴, reducing sensitivity, and compromising data accuracy^47,55.

LN₂ snap freezing is commonly used in biological research⁵⁶ because it rapidly halts cellular metabolism without leaving behind nonvolatile salts, making it a good fit for SCMS. However, it can damage cell membranes, which is problematic for SCMS^57,58. To reduce this damage, some studies used a method involving fast filtration, cold NaCl washing, and LN₂ freezing, which helps retain fast-turnover metabolites^53,59. Still, this method isn't ideal for SCMS due to difficulties in isolating individual cells and potential matrix effects from salt residues. Storage at -80 °C is another common preservation method, but its effects on single-cell metabolite profiles remain unclear. We have previously reported studies to address these limitations by combining multiple key steps: cells are first washed with a volatile, MS-compatible AF solution, then quenched with LN₂, vacuum freeze-dried, and stored at low temperature (-80 °C)⁶⁰. The current approach minimizes cell damage and metabolite degradation, enabling more accurate SCMS measurements. This work is focused on experimental details, aiming to promote the adoption of this cell preparation method for robust SCMS techniques.

Carry out every step inside a certified Class II biosafety laboratory and sterile conditions. In this study, the HCT-116 cell line was used as a model system, and cells were cultured in complete culture medium. The regents and the equipment used are listed in the Table of Materials.

1. Cell culture

1. Prepare complete medium. Supplement McCoy's 5A medium with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin. Warm it to 37 °C.
2. Incubate the cells at 37 °C in a humidified incubator with 5% CO₂. Monitor confluency daily and passage when cultures reach ~80% confluency.
3. Passage the cells (every ~2 days).
   1. Aspirate the medium and rinse once with 5 ml of 1x PBS.
   2. Add 2 mL of 0.25% trypsin-EDTA to the culture dish and incubate at 37 °C for 3 min to facilitate cell detachment.
   3. Neutralize trypsin by adding 8 mL of pre-warmed complete medium and gently pipetting to disperse cells.
   4. Transfer 1 mL of the suspension into 9 mL of fresh complete medium.
   5. Count the cells.
      NOTE: Cells were counted using a TC20 Automated Cell Counter according to the manufacturer's instructions. Trypan blue or other staining agents were not used.
4. Seed cells on the coverslips.
   1. Dilute the cells to a final concentration of ~1 × 10⁶ cells/mL.
   2. Place the individual 18 mm glass coverslips into wells of a 12-well plate.
      NOTE: Glass coverslips need to be autoclaved before being used for cell culture.
   3. Add 2 mL of the complete medium and 200 µL of the cell suspension (~2 × 10⁵ cells/well).
   4. Incubate the cells overnight to allow cell attachment.
      NOTE: To ensure cells are in normal growing status, routinely check cells' morphology and their growing status (using a microscope) as well as examine potential mycoplasma contamination (using a mycoplasma detection kit).

2. Cell washing with ammonium formate (AF) solution

NOTE: Prepare fresh AF solution each time.

1. Prepare the AF wash buffer. Weigh 0.45 g of ammonium formate and dissolve in 50 mL of LC/MS-grade water. The final concentration of ammonium formate is 0.14 M (0.9% w/w).
   NOTE: Keep the solution ice-cold (0-4 °C) until use.
2. Add 2 mL of cold AF solution to each well of the 12-well plate.
3. Rinse the coverslips. Using a pair of sterile forceps to gently lift each coverslip containing adherent cells from its well and then briefly (for 2-3 s) place it in a separate well pre-filled with 2 mL of cold 0.9% ammonium formate (AF) solution. Repeat the rinsing process.
   NOTE: To avoid disturbing the cells during the washing step, perform cell rinsing very gently.

3. Cell quenching by liquid nitrogen (LN₂)

1. Place each AF-rinsed coverslip (cells facing up) into the Petri dish inside the aluminum foil container.
   NOTE: Wear cryogenic-rated gloves, safety goggles, and a lab coat. Perform all LN₂ handling in a well-ventilated area. Aluminum foil can be used to fold a container for LN₂ quenching.
2. Slowly pour 10-20 mL of liquid nitrogen over the coverslips, ensuring complete coverage.
3. Immediately tilt the Petri dish with tweezers to decant any remaining liquid nitrogen.
   NOTE: Complete this action within a few seconds to prevent large ice-crystal formation from residual wash moisture.

4. Cell freeze-drying in vacuum

1. Freeze-dry immediately after LN₂ quenching to prevent thawing.
   NOTE: Remove the rotor of the SpeedVac to create a flat platform that accommodates a 100 mm Petri dish.
2. Using cryogenic gloves and insulated tweezers, place the Petri dish containing the LN₂ chilled coverslips onto the vacuum concentrator chamber.
3. Process using its standard vacuum settings and continue for 5-7 min until visible LN₂ has evaporated and coverslips appear dry. Dried coverslips should not contain any residual LN₂ and ice crystals.

5. Cell storage in a -80 °C freezer

1. Wrap the container with a paper towel and transfer it to a −80 °C freezer for 48 h.
2. After storage, immediately transfer the coverslips containing the Petri dish into a room-temperature desiccator for ~10 min. Ensure condensed frost or water on coverslips completely disappears before taking them out of the desiccator.
   NOTE: Rapidly transferring the coverslips to the desiccator minimizes condensation; even minor surface wetting can dissolve dried metabolites.
3. After AF washing, divide the samples into two groups (Group 1 and Group 2). Apply the following processing protocols.
   Group 1: LN₂ quench → freeze-dry → SCMS analysis (no storage).
   Group 2: LN₂ quench → freeze-dry → −80 °C storage 48 h → SCMS analysis.
4. Proceed with SCMS analysis as described below.
   NOTE: More cell groups can be prepared using different conditions. Group 1 and Group 2 are used to illustrate the general protocols.

6. Single-probe fabrication and SCMS setup

NOTE: The Single-probe is fabricated in accordance with established procedures^21,24,60,61, and only brief protocols are provided here.

1. Pull the dual-bore quartz needle using a laser puller and generate a ~10 µm tip. Assemble a Single-probe by inserting the fused silica capillary and nano-ESI emitter into the dual-bore quartz needle. Secure all components to a glass slide with quick-setting epoxy.
2. Mount the Single-probe on the XYZ stage. Affix the Single-probe to a motorized XYZ manipulator positioned beneath a digital microscope.
3. Couple the setup to a mass spectrometer for SCMS analysis.
4. Use acetonitrile (≥99.9%) containing 0.1 % formic acid as an extraction solvent at a flow rate of 150 nL/min delivered by a syringe pump.
5. Set the MS parameters.
   Positive ion mode: m/z 200-1500, +2.9 kV, mass resolution 120 k (at m/z 200).
   Negative ion mode: m/z 70-900, −2.1 kV, mass resolution 120 k (at m/z 200).
   NOTE: Because negative ions are generally detected in the m/z range lower than that of positive ions, different m/z ranges can be used for these two ion modes. Additional MS settings include 100 ms maximum injection time, one microscan, standard automatic gain control (AGC), full scan mode (for MS analysis), higher-energy collisional dissociation (HCD) (for MS/MS analysis) with 10-35 Normalized Collision Energy (NCE), and 320 °C for the ion transfer tube temperature. A total ion chromatogram (TIC) was generated at each scan/time point to monitor signal stability and single-cell detection.

7. Single-probe SCMS experiments

1. Place the two coverslips (Group 1 and Group 2) together on the motorized XYZ stage of the Single-probe SCMS setup^21,24,61.
2. To minimize batch effects, select the cells randomly under the microscope.
3. Acquire positive and negative ion mode mass spectra. Analyze multiple (e.g., ~30) cells per group with the same Single-probe, solvent flow rate, and ionization voltage.
   NOTE: Use the same Single-probe, solvent flow rate, and ionization voltages for every experiment to ensure data comparability. Calibrate the Orbitrap daily before acquisition. Both MS and MS/MS analyses can be performed at the single-cell level. Collect MS data to obtain accurate m/z values for studies of metabolite profiles and tentative labels in non-targeted analysis. Conduct MS/MS analysis of selected ions for molecular identification in targeted analysis. Data were acquired using a compatible data acquisition and interpretation software.

8. Data analysis

1. Preprocess the raw SCMS data using a custom R script. Preprocessing steps include background and noise removal and ion intensity normalization to TIC^11,12,23.
2. Deisotope the SCMS data using the Python package ms_deisotope.
3. Perform peak alignment using an in-house Python script⁶⁰. Use a bin width of 0.01 m/z for binning. Use a mass tolerance of 0.01 m/z or 5 ppm for peak alignment.
4. Perform statistical data analysis using MetaboAnalyst 6.0⁶². Conduct a t-test and generate a heat map (for relative metabolite levels). Select Use top 100 in T-test/ANOVA to generate the top 100 metabolites, ranked by p-values from low to high, with a significant difference (p < 0.05) from the comparison. Plot a heatmap using these top 100 metabolites.
5. Search accurate m/z values and MS/MS spectra against the Human Metabolome Database (HMDB⁶³). Use a mass tolerance of 10 ppm in MS database searching.
   NOTE: Other databases, such as LipidMaps⁶⁴, can be used for database searching.

This report describes a methodology that integrates cell washing, rapid cell quenching, vacuum drying, and storage at -80 °C (Figure 1). Specifically, this approach is designed to preserve the metabolomic state of cells prior to SCMS analysis. The effectiveness of this protocol was tested by preparing cells under two different conditions: freshly quenched and dried cells (Group 1, serving as the control), and quenched, dried, and -80 °C stored cells (Group 2). Each group was analyzed using the Single-probe SCMS method in both positive and negative ion modes. Principal Component Analysis (PCA) and heat map visualization were used to evaluate the differences in metabolomic profiles across groups.

PCA results in the positive ion mode (Figure 2A) revealed that Groups 1 and Group 2 had similar overall metabolite profiles. The nearly identical profiles of Group 1 and Group 2 demonstrate that -80 °C storage for 48 h does not significantly alter the metabolome if cells are properly quenched before drying. In the negative ion mode (Figure 2B), a comparable pattern is observed, although Group 2 appears slightly more distinct-potentially due to ionization differences among specific metabolite classes, for instance, phosphatidylcholines are better detected in positive mode, while fatty acids ionize more efficiently in negative mode60. Despite this variation, the consistent clustering between Groups 1 and 2 across both modes supports the conclusion that the integrated quenching-drying-storage protocol maintains metabolic integrity.

Heat map analysis further validated the PCA findings by depicting relative abundances of the top 100 metabolites across two groups. As shown in Figure 3, the top 100 metabolites display highly consistent abundance patterns between Group 1 and Group 2 in both ion modes. No major shifts or group-specific clustering are observed, although minor variations in a few metabolite clusters may reflect differences in ionization efficiency or metabolite stability. These results collectively demonstrate that the quenching-drying-storage workflow preserves cellular metabolomic profiles with high fidelity, making it well-suited for ambient SCMS applications.

Figure 1: General workflow for single-cell mass spectrometry (SCMS) analysis investigating how LN2 quenching and 48-h storage at -80 °C affect metabolite profiles in single cells. (A) Cells were seeded, incubated, and rinsed using ammonium format (AF) solution. (B) Two cell groups were prepared-Group 1: cells were washed, quenched, and freeze-dried for immediate SCMS experiments without storage; Group 2: cells were washed, quenched, freeze-dried, and stored at -80 °C. Please click here to view a larger version of this figure.

Figure 2: Principal Component Analysis (PCA) of SCMS data from HCT-116 cells in experimental groups in (A) positive ion mode and (B) negative ion mode. Please click here to view a larger version of this figure.

Figure 3: Heat maps showing the relative levels of the top 100 metabolites detected in single HCT-116 cells prepared under various conditions, presented for (A) positive ion mode and (B) negative ion mode. Please click here to view a larger version of this figure.

A critical step in this protocol is the immediate quenching of live cells using LN₂ following washing with AF, which substantially reduces enzymatic activity and halts ongoing metabolic processes. Additionally, washing cells with AF prior to quenching enhanced ion intensity and reduced matrix effects, supporting its inclusion as a preparatory step to improve SCMS sensitivity. In the current studies, cells in different groups (i.e., Group 1 and Group 2) were cultured using the same initial batch to minimize their biological variance. 48-h storage time was chosen because cells with additional growing time (Group 1) can still maintain suitable cell densities (e.g., to avoid cell aggregates) for comparative SCMS analysis with stored cells (Group 2). However, caution is warranted when relying on cold storage, even for 48 h, as significant metabolic changes were still observed in comparisons between freshly dried and stored samples (Group 1 vs. Group 2). Although in actual studies, cell and tissue samples are commonly stored for longer periods (e.g., for weeks and months) in a -80 °C freezer, it is suggested to shorten the storage time to minimize changes in metabolites.

Troubleshooting should therefore focus on minimizing the time between drying and SCMS analysis, ensuring that defrosting conditions avoid condensation or partial rehydration. Residual water content and water condensation during sample transitions can partially rehydrate cells and enable enzymatic and chemical reactions such as hydrolysis, oxidation, and proteolysis that degrade metabolites⁶⁵, leading to artificial changes in metabolite composition. Thoroughly dried samples effectively arrest these processes and preserve molecular integrity⁶⁶. Ensuring a moisture-free desiccation environment and rapid transition to SCMS is key to avoiding these pitfalls.

Despite the success of this method in preserving metabolite profiles, limitations remain. The method requires access to a vacuum drying system and a -80 °C freezer, and it is not fully effective for long-term storage. Furthermore, while rapid room-temperature drying preserves metabolites temporarily, it is not a substitute for LN₂ quenching. Storage-related alterations may still occur due to incomplete water removal and the risk of rehydration during sample handling. These limitations highlight the need for method refinement if longer storage or higher throughput is desired.

Protocols reported in this work offer a simplified and effective approach for preserving the single-cell metabolome for ambient SCMS. Unlike methods requiring elaborate cryoprotectants or in situ freezing, our workflow is readily compatible with existing ambient ionization techniques and allows for flexible sampling and analysis timelines. The integration of cell washing, quenching, drying, and short-term storage without substantial loss of metabolomic fidelity offers a practical solution for labs lacking on-demand access to mass spectrometers. This approach holds strong potential for advancing research in single-cell systems biology, cancer metabolism, drug response heterogeneity, and metabolic phenotyping of rare cell populations. The broad applicability of these protocols, particularly for ambient ionization-based SCMS platforms, makes them an important contribution to the field of single-cell analytical techniques. In future studies, these protocols can be adapted and refined to accommodate a wider variety of cell types. With additional sample preparation steps (e.g., matrix application), the processed cells can be potentially analyzed by other SCMS techniques (e.g., MALDI and SIMS), offering higher spatial resolution and throughput for analysis of larger numbers of cells (e.g., highly confluent cells). The results also suggest that these quenching-drying-storage protocols can be potentially used to prepare cells for shipping (e.g., in dry ice containers), allowing for collaborative SCMS studies at different locations.