Strategies for Enhancing Sensitivity in Nanoelectrospray Mass Spectrometry: A Guide for Biomedical Researchers

Abstract

Nanoelectrospray ionization (nESI) mass spectrometry is a cornerstone technology for analyzing biomolecules, but achieving optimal sensitivity is challenging. This article provides a comprehensive guide for researchers and drug development professionals on improving nESI-MS sensitivity. We explore the foundational principles of nano-electrospray, including emitter technology and droplet dynamics. The article details advanced methodological approaches for analyzing proteins, metabolites, and oligonucleotides from physiologically relevant buffers. A dedicated troubleshooting section offers practical optimization strategies for emitter positioning, voltage settings, and salt adduction mitigation. Finally, we validate these techniques through performance comparisons and applications in pharmaceutical and clinical analyses, providing a holistic framework for maximizing signal quality and data reliability in sensitive biomolecular studies.

Understanding Nanoelectrospray Fundamentals: From Taylor Cones to Gas-Phase Ions

nanoElectrospray Ionization (nESI) is a cornerstone technique in modern mass spectrometry, particularly for the analysis of large biomolecules. Its fundamental principle involves applying a high voltage to a liquid sample at the tip of a very fine emitter, producing a spray of highly charged, tiny droplets. As these droplets evaporate, they undergo a series of Coulombic fissions, eventually leading to the release of gas-phase ions. The key differentiator of nESI from conventional electrospray is its operation at nanoliter per minute flow rates, which generates an initial droplet size well below that of standard ESI. This seemingly simple modification—reducing the flow rate and initial droplet size—confers a suite of analytical advantages that dramatically enhance ionization efficiency, improve sensitivity, and reduce sample consumption. This article details these advantages within the context of improving sensitivity in MS research, providing troubleshooting guides and experimental protocols for researchers and drug development professionals.

The Core Mechanisms: Why Smaller Droplets and Lower Flow Rates Matter

The enhanced performance of nESI is not a single phenomenon but the result of several interconnected physical and chemical mechanisms.

The nESI Droplet Lifecycle and Ion Formation

The journey from a liquid sample to a gas-phase ion in nESI follows a precise pathway. The diagram below illustrates this process and the points at which key advantages are realized.

Quantifiable Benefits of Low Flow Rates

The theoretical advantages of nESI are borne out by concrete experimental data. A systematic study infusing an equimolar mixture of a poorly ionizing oligosaccharide (maltotetraose) and an easily ionized peptide (neurotensin) at different flow rates demonstrated a dramatic reduction in ion suppression at lower flows. The table below summarizes the key quantitative findings from this study.

Table 1: Impact of Flow Rate on Ionization Efficiency and Ion Suppression

Flow Rate (nL/min)	Normalized Signal Intensity (Maltotetraose)	Normalized Signal Intensity (Neurotensin)	Maltotetraose/Neurotensin Signal Ratio
~10 nL/min	High (Saturation regime)	High (Saturation regime)	Highest
~20 nL/min	High (Saturation regime)	High (Saturation regime)	High
100 nL/min	Moderate	Moderate	Moderate
>300 nL/min	Low	Low	Low

Source: Adapted from a study on the effect of flow rate using CESI-MS for biotherapeutic molecules [1].

The exponential increase in the signal ratio for the less-easily-ionized maltotetraose at lower flow rates confirms that ion suppression is significantly minimized. This is because the smaller initial droplets and reduced overall volume limit the number of competing molecules, allowing analytes with poorer ionization efficiency to be more effectively detected [1].

Troubleshooting Guide and FAQs

This section addresses common challenges encountered during nESI-MS experiments.

Frequently Asked Questions

• Q1: Why does my nESI signal rapidly fluctuate or become unstable?

  • A: Instability can stem from several sources:
    • Bubbles: Dissolved gases can precipitate and form bubbles as the liquid passes through the emitter, distorting the flow and meniscus geometry [2]. Ensure your samples are properly degassed and avoid temperature swings.
    • Clogging: Residues can accumulate at the emitter tip, especially with narrow-bore emitters. Using samples that are free of particulate matter and employing emitters with a larger internal diameter can improve robustness [3] [2].
    • Lateral Wetting: The electrospray meniscus can become anchored to the side of the emitter tip, leading to an unstable spray. Using emitters with a sharp, well-defined geometry can help lock the meniscus in place [2].
    • Evaporation: Excessive solvent evaporation at the tip can alter buffer concentration and flow dynamics. Using a conductive, hydrophobic-coated emitter can help stabilize the meniscus [2].

• Q2: I am analyzing proteins in native conditions with high salt. My signal is suppressed and I see extensive salt adduction. What can I do?

  • A: This is a common challenge. Two effective strategies are:
    • Use Nanoscale Emitters: Emitters with inner tip diameters of ~500 nm produce smaller initial droplets that limit the co-solvation of nonvolatile salts, inherently reducing adduction [3].
    • Implement Reduced Pressure Ionization: Placing the nESI emitter inside a low-pressure chamber (200-333 mbar) can enhance desolvation. One study observed signal enhancements of up to 20-fold in high-salt solutions (up to 300 mM NaCl) and enabled the detection of a protein complex at concentrations as low as 50 nM, where ambient pressure ionization failed [3].

• Q3: My collision-induced dissociation (CID) or unfolding (CIU) results are not reproducible between users or days. Why?

  • A: A often-overlooked factor is the precise position of the nESI emitter relative to the MS inlet. Even small variations in position can cause unintended in-source activation, shifting the voltage required for 50% dissociation (CID₅₀) by as much as 8 V on some instruments. For maximum reproducibility, consistently place the emitter at the same "sweet spot" relative to the inlet and document this position in your methods [4].

• Q4: Why is nESI particularly advantageous for analyzing hydrophobic substances or complex mixtures?

  • A: During Coulombic fission, hydrophobic molecules are preferentially transferred to the next generation of droplets because they reside at the droplet-air interface. Starting with smaller droplets reduces the number of fission events required to produce ions, thereby reducing the opportunity for ion suppression of more hydrophilic molecules in the mixture [2].

Troubleshooting Common nESI Problems

Table 2: nESI Troubleshooting Guide

Problem	Potential Causes	Solutions
No Spray / No Signal	• Clogged emitter• No electrical contact• Voltage too low• Large air bubble blocking flow	• Flush or replace emitter• Check platinum wire contact• Increase spray voltage within stable range (e.g., 1.0-1.5 kV) [4]• Apply brief pressure to clear bubble or use reduced pressure to pull solution [3]
Unstable / Fluctuating Signal	• Solvent evaporation at tip• Bubble formation in line• Lateral wetting of emitter tip• Unstable meniscus	• Use a hydrophobic emitter coating [2]• Degas solvents and samples• Use emitters with a sharp, well-defined geometry [2]
High Chemical Noise & Salt Adduction	• Non-volatile buffers/salts• Emitter tip too large• Meniscus too large	• Desalt via spin column or buffer exchange [3]• Use narrower-bore emitters (e.g., <1 µm ID) [3]• Use reduced pressure ionization [3]
Poor Reproducibility in CIU/CID	• Uncontrolled in-source activation• Variable emitter position	• Map the optimal emitter position for your instrument [4]• Keep emitter position and spray voltage consistent between runs [4]

Key Experimental Protocols

Protocol: Enhanced Native MS Analysis Using Reduced Pressure Ionization

This protocol is adapted from a 2025 study that demonstrated significant improvements in analyzing proteins and complexes under challenging buffer conditions [3].

1. Objective: To achieve superior signal-to-noise and reduced salt adduction for native proteins in high-salt buffers by implementing a reduced pressure ionization source.

2. Materials:

• Proteins/Complexes: Recombinant truncated KLHL7 (5 µM) or DDB1:DCAF1 complex (50 nM) in 120 mM ammonium acetate [3].
• nESI Emitters: Nanoscale emitters with inner tip diameter of ~500 nm [3].
• Custom Reduced Pressure Chamber: A 3D-printed chamber capable of maintaining 200-333 mbar, interfacing with the MS inlet [3].
• Mass Spectrometer: Modified LTQ Orbitrap XL or Q Exactive UHMR, though the design can be adapted to other platforms [3].

3. Procedure: 1. Sample Preparation: Prepare your protein complex in a near-physiological, volatile buffer (e.g., 120-200 mM ammonium acetate). If non-volatile salts are necessary for stability, note that signal may still be enhanced. 2. Emitter Loading: Load the sample into the nESI emitter, taking care to minimize air bubble introduction. 3. Chamber Setup: Mount the nESI emitter into the custom reduced pressure chamber and seal it. Attach the chamber to the mass spectrometer inlet. 4. Pressure Reduction: Activate the vacuum. The chamber pressure should drop to the operational range (e.g., ~333 mbar for an LTQ Orbitrap XL, ~200 mbar for a Q Exactive UHMR) within seconds. 5. Mass Spectrometry: Initiate the spray with a typical nESI voltage (0.9-1.5 kV). Acquire data in the appropriate positive or negative ion mode for your analyte. Compare the spectrum with one acquired at ambient pressure to observe the enhancement in signal intensity and peak narrowing.

4. Expected Results: The study reported signal enhancements of up to 20-fold with nanoscale emitters in high-salt solutions. Protein ions remained detectable in solutions containing up to 300 mM NaCl. The DDB1:DCAF1 complex was detectable at 50 nM with high signal-to-noise under reduced pressure, whereas no readily assignable signal was detected at ambient pressure [3].

Protocol: Optimizing nESI Voltage for Native Oligonucleotide Analysis

This protocol is based on a 2025 study using capillary vibrating sharp-edge spray ionization (cVSSI), a voltage-controlled field-free technique, for DNA triplexes [5].

1. Objective: To find the optimal applied voltage for sensitive detection of native oligonucleotides while minimizing cation adducts and preserving structure.

2. Materials:

• Oligonucleotides: DNA triplex (e.g., 36mer formed by (GAA)₁₂ and (TTC)₁₂ in a 1:2 ratio), 50 µM in 400 mM ammonium acetate, pH 5.5 [5].
• cVSSI Device: Fused silica capillary pulled to 15-30 µm I.D., attached to a piezoelectric transducer [5].
• Mass Spectrometer: Q-Exactive hybrid quadrupole-Orbitrap or similar, operated in negative-ion mode [5].

3. Procedure: 1. System Setup: Infuse the DNA triplex sample at a constant flow rate (e.g., 2 µL/min). Set the mass spectrometer's heated inlet transfer tube temperature to 300°C. 2. Voltage Sweep: Perform a series of acquisitions while sweeping the DC bias voltage applied to the solution. A recommended range is from -900 V to -1500 V. 3. Data Analysis: For each voltage, analyze the mass spectra. Monitor the abundances of the desired triplex ions ([Tri]⁸⁻, [Tri]⁹⁻, [Tri]¹⁰⁻) and the corresponding adduct ions (Tri + ad). 4. Determine Optimal Voltage: Calculate the ratio of desired triplex ion abundance to triplex adduct ion abundance for each charge state.

4. Expected Results: The study found that a medium applied voltage of ~ -900 V was optimal. It enhanced the peak intensities of the desired DNA triplex ions by 70 to 260 fold for different charge states, compared to higher voltages (-1100 to -1500 V). The ratio of desired ions to adduct ions increased by approximately 6-fold at the lower voltage, indicating a cleaner spectrum with fewer adducts [5].

The Scientist's Toolkit: Essential Research Reagents & Materials

Successful nESI-MS experiments rely on the right tools. The following table lists key materials and their functions.

Table 3: Essential Materials for nESI-MS Research

Item	Function / Description	Key Consideration
Nanoscale Emitters	Produces the fine spray of small droplets. Can be pulled from glass or fused silica.	Geometry is critical: A sharp, well-defined tip ensures a stable, small meniscus. Inner diameters can range from <1 µm for high salt to 10-30 µm for general use [3] [2].
Hydrophobic Emitter Coatings	A hydrophobic internal coating (e.g., LOTUS coating) locks the meniscus at the emitter's inner diameter.	Results in a smaller, more stable meniscus, leading to less solvent evaporation, lower required voltages, and better ionization efficiency [2].
Volatile Buffers	Maintains biomolecules in a native-like state while being compatible with MS. Ammonium acetate is the most common.	Typical concentrations are 100-200 mM. Avoid non-volatile salts and buffers where possible [3] [5].
Syringe Pumps / Pressure Systems	Provides precise, pulseless flow of sample to the emitter at nL/min rates.	Stability at ultra-low flow rates (< 100 nL/min) is essential for consistent performance [1].
Reduced Pressure Chamber	A custom chamber that lowers pressure around the emitter to enhance desolvation.	Can be 3D-printed. Shown to dramatically improve signal in high-salt and low-concentration samples [3].
Capillary cVSSI Device	A field-free ionization source that uses vibration, not high voltage, to generate spray.	Can be gentler for fragile molecules and reduces corona discharge issues in negative-ion mode [5].

The principles of nESI are being integrated into advanced workflows to solve specific analytical challenges. The diagram below maps the decision process for selecting and applying nESI-based solutions.

Understanding the complete droplet lifecycle in electrospray ionization (ESI) is fundamental to improving sensitivity in nanoelectrospray mass spectrometry (nanoESI-MS) research. The journey from a liquid sample to a detectable gas-phase ion involves precisely orchestrated stages, each presenting potential points of ion loss or signal suppression that directly impact analytical sensitivity. For researchers and drug development professionals, mastering this process enables optimization of experimental parameters to achieve maximum signal intensity, particularly for challenging analyses like native protein complexes and large biomolecules. This technical resource details the droplet pathway from initial formation to final ion emission, providing actionable troubleshooting guidance to address real-world experimental issues encountered in the laboratory.

The Electrospray Droplet Lifecycle: Mechanisms and Transitions

The transformation of a liquid sample into gas-phase ions follows a defined sequence of physical events. The diagram below illustrates the complete pathway and key transition mechanisms.

Diagram 1: The complete electrospray droplet lifecycle from Taylor cone formation to gas-phase ion emission.

Stage 1: Taylor Cone and Initial Droplet Formation

The electrospray process initiates when a high voltage (typically 1.7-2.5 kV for nanoESI [2]) is applied to a liquid protruding from a capillary emitter. This creates electrostatic stress that counteracts the liquid's surface tension, forming a conical meniscus known as a Taylor cone with a characteristic 49.3° angle [7]. At the tip of this cone, the electric field becomes intensely concentrated, leading to the ejection of a liquid jet that breaks up into a fine mist of charged primary droplets [7] [2]. The stability of this cone is paramount for a consistent ion signal.

Stage 2: Droplet Evolution and Coulombic Fission

The initially formed droplets undergo a process of desolvation and disintegration:

• Solvent Evaporation: Neutral solvent molecules evaporate from the charged droplets, causing them to shrink while maintaining their charge [7] [8]. This progressively increases the charge density on the droplet surface.
• Rayleigh Limit and Fission: When the electrostatic repulsion within a droplet nearly overcomes the surface tension holding it together (the Rayleigh limit), the droplet becomes unstable and undergoes Coulombic fission [7] [2]. This disintegration event produces smaller, highly charged "offspring" droplets, each containing approximately 25% of the parent droplet's charge but only about 0.1% of its volume [2].

Stage 3: Final Ion Emission Mechanisms

The final stage of ion release from the highly charged, nanometer-scale droplets is described by two primary models:

• Charge Residue Model (CRM): Proposes that successive solvent evaporation and fission events continue until a droplet contains only a single analyte molecule. The ion is released when the remaining solvent evaporates completely, with the droplet's residual charge transferring to the analyte [7]. This model is often invoked for large biomolecules.
• Ion Evaporation Model (IEM): Suggests that when droplets shrink to a critical size (approximately 10 nm [2]), the electric field at their surface becomes strong enough to directly desorb solvated ions into the gas phase before the solvent fully evaporates [7]. This model may be more relevant for smaller ions.

The entire sequence, from primary droplet to ion-emitting droplet, can occur very rapidly, often in less than a millisecond [8].

Troubleshooting Guide: Experimental Issues and Solutions

Frequently Asked Questions

Q1: Why does my signal intensity fluctuate unpredictably during a nanoESI run? Signal instability commonly stems from an unstable electrospray meniscus. Key causes include:

• Voltage/Fluidics Issues: Operating outside the optimal voltage range (typically 1.7-2.5 kV for proteomics) or having a flow rate that cannot sustain a stable jet against solvent evaporation [2].
• Emitter Condition: A clogged, contaminated, or poorly wetted emitter tip disrupts a consistent Taylor cone. Bubbles forming in the fluid path can also cause major instability [2].
• Solution: Use sharper, hydrophobic-coated emitters to lock the meniscus anchor point and improve stability. Ensure your voltage and flow rates are optimized for your specific solvent and emitter geometry [2] [9].

Q2: My MS sensitivity is low for large protein complexes. How can the droplet lifecycle concept help? Large complexes are sensitive and can be disrupted or lost during the ionization process.

• Cause: Excessive energy from too many Coulombic fissions or harsh desolvation conditions can disrupt non-covalent bonds [7]. Adsorption of proteins to the glass emitter surface also reduces the number of ions reaching the gas phase [9].
• Solution: Utilize emitters with inert inner coatings (e.g., polyethylene-glycol) to minimize analyte adsorption [9]. Optimize source conditions (e.g., lower desolvation gas temperature) to promote the "softer" ionization required for native complexes, potentially via the Charge Residue Model pathway [7].

Q3: I observe contamination and high background in my mass spectra. Could droplets be the cause? Yes. Large, slow-moving charged droplets can aspirate deeply into the vacuum stages of the mass spectrometer. When they eventually break apart, they release a burst of non-volatile contaminants and poorly desolvated ions, causing spectral noise and instrument contamination [8].

• Solution: Ensure efficient droplet desolvation by using appropriate source heating and gas flows. Positioning the emitter closer to the MS inlet can also help, as the stronger field accelerates droplet breakup before they can be aspirated [8] [2].

Q4: Why should I use nanoESI over higher-flow ESI for sensitivity-limited applications? NanoESI (flow rates in the nL/min range) generates a first generation of much smaller droplets (often 200-500 nm) compared to high-flow ESI (which can produce droplets up to 35 μm) [8] [2].

• Benefit: Starting with smaller droplets means fewer fission events are needed to reach the ion-shedding regime (~10 nm). This reduces the creation of "zombie" droplets (low-charge byproducts that waste sample), minimizes the accumulation of contaminants that cause ion suppression, and ultimately improves ionization efficiency [10] [2].

Troubleshooting Table: Common Problems and Fixes

Table 1: A guide to diagnosing and resolving common nanoESI issues related to the droplet lifecycle.

Problem Symptom	Potential Cause	Diagnostic Check	Corrective Action
Unstable or pulsing spray	Meniscus wetting instability; Bubble formation; Clogged emitter.	Check for salt crystallization under microscope; Inspect fluidic connections for leaks.	Use sharper, hydrophobic-coated emitters [2]; Degas solvents; Filter samples.
Low signal intensity for all analytes	Poor ionization efficiency; Large initial droplet size; Droplet aspiration into MS.	Measure spray current; Check emitter-to-inlet distance.	Reduce liquid flow rate to nano-scale [10] [2]; Use multi-emitter array to split flow [10].
High chemical noise & background	Incomplete droplet desolvation; Contaminant burst from aspirated droplets.	Check for signal spikes correlated with large droplets.	Optimize source heating and desolvation gas flow [10]; Clean ion inlet and source region.
Signal loss for large, non-covalent complexes	Complex disruption during ionization; Adsorption to glass emitter.	Compare signal with and without volatile salts.	Use surface-modified, inert emitters (e.g., PEG-coated) [9]; Soften source conditions (lower V, T).
Corona discharge & electrical arcing	Voltage too high for given meniscus size and gas pressure.	Observe spray plume for glow; listen for audible snapping.	Reduce spray voltage; Use emitter with smaller meniscus [2]; Introduce slight CO2 sheath gas [10].

Experimental Protocols for Enhancing Sensitivity

Protocol: Utilizing Multi-Emitter Arrays for High Sensitivity

Objective: To significantly increase MS sensitivity by employing an array of nanoESI emitters, which splits a higher liquid flow (e.g., from LC) into multiple nano-flow electrosprays, thereby generating more ions and improving overall ionization efficiency [10].

Materials:

• Fabricated multi-emitter array with individualized sheath gas capillaries [10].
• Syringe pump or LC system.
• Mass spectrometer with a modified SPIN (Subambient Pressure Ionization with Nanoelectrospray) source or standard API source [10].

Method:

• Emitter Array Fabrication: Construct an array by inserting fused silica emitter capillaries (e.g., 150 μm o.d., 10 μm i.d.) through a larger concentric sheath gas capillary assembly. Arrange them in a circular pattern (e.g., 4, 6, or 10 emitters) using a spacer [10].
• Emitter Etching: Chemically etch the emitter tips in hydrofluoric acid (HF) while pumping water through the emitters to create external tapers and prevent inner wall etching [10].
• Source Coupling: Integrate the emitter array into a SPIN source positioned in the first reduced-pressure region (10-30 Torr) of the mass spectrometer. This configuration minimizes ion losses at the inlet [10].
• Sheath Gas Optimization: Apply a controlled flow of CO2 sheath gas individually to each emitter to stabilize the electrospray and enhance droplet desolvation at subambient pressure [10].
• Performance Evaluation: Infuse a standard peptide mixture (e.g., 1 μM each in 0.1% formic acid). Compare the total ion current and signal-to-noise ratio against a standard single-emitter, atmospheric pressure ESI source.

Expected Outcome: The multi-emitter/SPIN configuration has been shown to provide over an order of magnitude improvement in MS sensitivity compared to a standard single-emitter source, as the total ESI current increases with the number of emitters and losses are minimized [10].

Protocol: Surface Modification of NanoESI Emitters

Objective: To improve signal intensity, particularly for native MS and challenging analytes like viral capsids, by reducing nonspecific adsorption to the glass emitter surface [9].

Materials:

• Pulled borosilicate glass nanoESI needles.
• (3-Glycidyloxypropyl)trimethoxysilane (GPTMS).
• Polyethylene glycol (PEG) solution.
• Standard peptide or protein sample for testing.

Method:

• Emitter Cleaning: Thoroughly clean new emitters with piranha solution to activate surface silanol groups. (Caution: Piranha solution is extremely corrosive and must be handled with care.)
• Silane Functionalization: Vapor-deposit GPTMS onto the emitter surface to create an epoxide-terminated layer.
• PEG Coating: React the epoxide-functionalized surface with a 10 mM PEG solution (in 100 mM NaOH) for 16 hours to create a hydrophilic, protein-repellent coating.
• Capping: Cap any unreacted epoxides by incubating with 1 M ethanolamine.
• Performance Validation: Analyze a standard protein or complex (e.g., AAV capsid for CD-MS) and compare the signal intensity and stability against an unmodified emitter.

Expected Outcome: Surface-modified emitters demonstrate a marked increase in signal intensity for native MS and charge detection-MS experiments. It is hypothesized that this improvement may be linked to an effectively increased flow rate through the coated needles, delivering more analyte to the mass spectrometer [9].

The Scientist's Toolkit: Key Reagents and Materials

Table 2: Essential materials and reagents for optimizing the electrospray droplet lifecycle to improve sensitivity.

Item	Function / Rationale	Application Note
Sharp, Hydrophobic Emitters	A mechanically sharp, hydrophobic tip (e.g., LOTUS coating) locks the meniscus at the inner diameter, creating a smaller, more stable Taylor cone. This reduces solvent evaporation at the tip, allows for lower operating voltages, and minimizes corona discharge, leading to better ionization efficiency [2].	Critical for achieving stable, consistent sprays in nanoESI workflows. The defined geometry improves run-to-run repeatability.
Multi-Emitter Array	Splits a single liquid stream (e.g., from LC) into multiple nano-flow electrosprays. This generates a "brighter" total ion current and produces smaller initial droplets, which improves ionization efficiency and reduces sample waste [10].	Ideal for coupling high-flow LC separations with high-sensitivity nanoESI detection. Requires a specialized source (e.g., SPIN) for optimal ion transmission.
Surface-Modified Emitters	A coating (e.g., silane-PEG) passivates the inner glass surface, reducing nonspecific adsorption of precious analyte molecules. This leads to higher signal intensity, especially for "sticky" analytes like large proteins and complexes in native MS [9].	A simple and inexpensive method to improve sensitivity for challenging applications. The modification may also influence flow dynamics.
Sheath Gas Delivery System	A concentric flow of gas (often CO2 or N2) around the emitter stabilizes the electrospray, particularly in subambient pressure environments. It also aids in the initial desolvation of charged droplets, facilitating the transition to gas-phase ions [10].	Essential for stable operation of the SPIN source and multi-emitter arrays. Can also help stabilize conventional high-flow ESI.
Volatile Buffers & Additives	Using mobile phases with volatile salts (e.g., ammonium acetate) and acids (e.g., formic acid) prevents non-volatile residues from accumulating in droplets. This reduces ion suppression, background noise, and source contamination [2].	A fundamental requirement for robust and sensitive ESI-MS. Non-volatile salts will quickly degrade performance.

Troubleshooting Guide: Nanoelectrospray Performance Issues

Unstable Spray or Signal Fluctuations

• Problem: Inconsistent Taylor cone formation, leading to signal instability during data acquisition.
• Potential Causes & Solutions:
  • Cause 1: Suboptimal spray voltage due to meniscus size variations. The optimal voltage range for nano-ESI in proteomics workflows is typically 1.7 to 2.5 kV [2]. An improperly sized meniscus can necessitate voltage adjustments outside this stable window.
  • Solution: Verify and optimize the spray voltage. If the meniscus is too large, it may require higher voltages that can induce corona discharges, where gas ions reduce droplet charge and ionization efficiency [2]. Ensure emitter geometry promotes a small, stable meniscus.
  • Cause 2: Lateral or longitudinal wetting instabilities, where the meniscus anchor point moves unpredictably on the emitter surface [2].
  • Solution: Use emitters with a sharp, well-defined edge geometry. This abrupt geometry locks the meniscus at a consistent anchorage diameter, eliminating a source of variability and improving result consistency [2].
  • Cause 3: Bubbles in the fluidic path. As liquid flows through the system, dissolved gases can precipitate. Expanding bubbles distort the flow rate, disrupt meniscus geometry, and break electrical conductivity [2].
  • Solution: Ensure proper degassing of solvents and mobile phases prior to use.

Poor Ionization Efficiency and Ion Suppression

• Problem: Reduced signal intensity for analytes of interest, often due to competitive ionization in the source.
• Potential Causes & Solutions:
  • Cause 1: Large initial droplet size. Larger droplets require more coulombic fission events to reach the ion-shedding scale (~10 nm), increasing the chance for ion suppression of hydrophilic molecules and generating more low-charge "zombie" droplets that waste sample [2].
  • Solution: Utilize emitters designed to produce smaller first-generation droplets. This is achieved by controlling flow rate and emitter geometry to create a smaller, more stable meniscus [2].
  • Cause 2: Chemical interactions and binding of analytes to the inner surface of the glass emitter [2].
  • Solution: Use emitters with inert, hydrophobic internal coatings. This minimizes analyte adhesion, stabilizes the meniscus at the emitter's inner diameter, and results in a smaller meniscus with better ionization efficiency and a more consistent spray [2].

Rapid Emitter Clogging

• Problem: Physical blockage of the emitter tip, preventing flow and spray.
• Potential Causes & Solutions:
  • Cause: Accumulation of residues or precipitated samples at the emitter tip, especially when using small inner diameter (ID) emitters [2].
  • Solution: For robustness, select an emitter with the largest practical ID to prevent clogging, while still maintaining performance objectives [2]. Implement rigorous sample cleanup procedures to remove particulate matter.

Excessive Electrical Discharge or Adduct Formation

• Problem: Corona discharge in positive ion mode, or prevalent sodium ([M+Na]+) and potassium ([M+K]+) adducts in spectra.
• Potential Causes & Solutions:
  • Cause 1: Spray voltage set too high for the mobile phase composition, leading to rim emission or corona discharge [11].
  • Solution: Lower the sprayer voltage. The adage "if a little bit works, a little bit less probably works better" often applies. For highly aqueous eluents, consider adding 1-2% v/v of a solvent with lower surface tension (like methanol or isopropanol) to stabilize the Taylor cone and allow for stable operation at a lower voltage [11].
  • Cause 2: Contamination from metal ions leached from glass vials or present in solvents [11].
  • Solution: Use high-purity, LC-MS grade solvents and plastic vials instead of glass to minimize metal ion introduction. Thoroughly flush the system between runs [11].

Frequently Asked Questions (FAQs)

Q1: Why does emitter tip geometry matter if the sample is delivered through the inner diameter?

The meniscus size, which is critical for spray stability, is defined by the meniscus angle and the emitter's outer geometry, not the inner diameter [2]. A sharp, well-defined tip ensures the meniscus anchors consistently at the same point. In contrast, a rounded tip allows the meniscus anchorage point to vary, leading to instability and inconsistent results [2].

Q2: What are the benefits of using nano-ESI over conventional ESI for sensitive proteomics analyses?

Nano-ESI produces smaller initial droplets, which leads to several key advantages [2]:

• Improved Ionization Efficiency: Smaller droplets have a lower absolute amount of contaminants (like salts) that accumulate as the solvent evaporates.
• Reduced Ion Suppression: Fewer fission events are needed to reach the ion-shedding regime, which lessens the preferential transfer of hydrophobic substances that can suppress hydrophilic analytes.
• Better Sample Utilization: The population of low-charge "zombie" droplets that waste sample is reduced.

Q3: How can I analyze proteins from solutions with high concentrations of non-volatile salts, like physiological buffers?

Standard nano-ESI emitters struggle with this, but specialized tools exist. Theta emitters—glass emitters with a septum dividing the capillary into two channels—allow one channel to be loaded with the protein in a biological buffer, while the other contains a volatile salt like ammonium acetate [12]. Rapid mixing at the tip is posited to create a population of droplets relatively depleted of non-volatile salts, enabling analysis of proteins and protein complexes from physiologically relevant solutions [12].

Q4: My spray is stable with pure water/acetonitrile but becomes unstable during a gradient. Why?

The stability of the electrospray is highly dependent on solvent properties like surface tension and conductivity [2] [13]. As the mobile phase composition changes during a gradient, the properties of the liquid at the tip change, which can shift the electrospray out of its optimal "cone-jet" regime and into a pulsating or unstable regime [13]. Monitoring the spray current can help diagnose these regime transitions.

Experimental Protocols & Data

Table 1: Optimizing Critical Nano-ESI Parameters for Stable Spray

Parameter	Typical Optimal Range	Effect on Spray Performance	Troubleshooting Action
Spray Voltage	1.7 - 2.5 kV [2]	Too low: No spray. Too high: Corona discharge, unstable spray [2] [11].	Optimize voltage in small increments while monitoring signal stability.
Flow Rate	< 1 μL/min (nano-ESI)	Defines initial droplet size. Higher flows can destabilize the Taylor cone [2] [11].	Ensure flow rate is compatible with emitter size and stable jet formation.
Emitter MS Inlet Distance	1 - 2 mm [2] [12]	Smaller meniscus requires closer proximity to the MS inlet for efficient ion transport [2].	Adjust distance for maximum signal intensity.
Solvent Composition	Additives (e.g., 0.1% FA)	Modifies conductivity & surface tension for stable Taylor cone formation [11].	Add low-surface-tension solvent (e.g., 1-2% IPA) to highly aqueous buffers [11].
Nebulizing Gas	Instrument-specific	Assists in droplet formation and desolvation; critical at higher flow rates [11].	Optimize gas flow to achieve a stable signal without introducing turbulence.

Table 2: Emitter Characteristics and Their Impact on Performance

Emitter Feature	Impact on Meniscus & Spray	Performance Outcome
Sharp, Well-Defined Tip [2]	Anchors meniscus at a consistent diameter.	Improved spray stability and result reproducibility.
Hydrophobic Inner Coating (e.g., LOTUS) [2]	Locks meniscus at the inner diameter, creating a smaller meniscus.	Better ionization efficiency, less evaporation, lower required voltages.
Large Inner Diameter (where possible) [2]	Does not directly define meniscus size, but affects clogging.	Increased robustness and reduced frequency of clogging.
Circular Array Geometry [14]	Reduces electric field inhomogeneities between neighboring emitters.	Enables all emitters in a multi-emitter setup to operate optimally at the same voltage.

Diagram: Relationship Between Emitter Design and Spray Stability

The Scientist's Toolkit: Essential Research Reagent Solutions

Table 3: Key Materials for Advanced Nanoelectrospray Research

Item	Function / Application	Specific Example / Note
Sharp Singularity Emitters [2]	Mechanically sharpened tips for controlled geometry, optimized for performance and repeatability.	Provides a stable meniscus anchorage for consistent results.
LOTUS Coated Emitters [2]	Hydrophobically coated emitters to lock the meniscus at the inner diameter.	Creates a smaller meniscus, leading to better ionization efficiency and spray stability.
Simple Link-Uno [2]	Connection system that provides voltage and connects the column to the emitter with zero dead volume.	Reduces system complexity and potential for installation errors.
Theta Emitters [12]	Dual-channel emitters for mixing sample with additives immediately prior to spray.	Enables analysis of proteins from physiological buffers with high non-volatile salt concentrations.
Ammonium Acetate with Additives [12]	Volatile salt solution supplemented with anions of low proton affinity (e.g., Br-, I-).	When used in one channel of a theta emitter, can reduce ionization suppression and chemical noise from salts.

Troubleshooting Guides

FAQ 1: How do solvent properties affect my nanoelectrospray signal intensity and why?

The core physical properties of your electrospray solvent—surface tension, conductivity, and permittivity (dielectric constant)—directly control the stability of the Taylor cone, the size of the initial charged droplets, and the efficiency of droplet fission and ion release. An imbalance can lead to spray instability, poor ionization, and severe ion suppression effects.

• Underlying Principle: The successful operation of nanoelectrospray relies on the formation of a stable Taylor cone. The balance between the solvent's surface tension (which holds the droplet together) and the electrostatic forces (influenced by the solution's conductivity and the applied field) is critical. Solvents with high permittivity help sustain the electric field required for electrospray.
• Connection to Sensitivity: Optimizing these properties is foundational to improving sensitivity in nanoelectrospray MS research. Efficient and stable ion production increases the absolute signal of your analyte, which is the first step toward achieving lower limits of detection.

FAQ 2: I observe intense signal suppression for my analyte in a mixture. How can solvent properties help?

Signal suppression often occurs when your analyte is out-competed during the ionization process. Modifying the solvent properties can shift this competitive balance.

• Root Cause: In a multi-analyte solution, compounds with higher surface activity or inherent charge will preferentially occupy the droplet surface and dominate charge emission. This is quantified by the signal intensity ratio between a neutral and a charged species, which can vary exponentially with flow rate (and thus effective droplet size) [1].
• Solution Strategy: Using ultra-low flow rates (e.g., ~20 nL/min) inherently generates smaller initial droplets with a higher surface-to-volume ratio. This configuration reduces ion suppression, allowing less easily ionized species (like neutral sugars) to be detected more effectively [1]. The table below summarizes quantitative data on this effect.

Table 1: Impact of Flow Rate on Ion Suppression

Flow Rate (nL/min)	Observed Effect on Ion Suppression	Quantitative Change
~20 nL/min	Ion suppression becomes "practically negligible" [1].	Signal intensity ratios of neutral/charged analytes converge to a saturation regime [1].
> 300 nL/min	Significant ion suppression is observed [1].	Normalized signal intensity for suppressed analytes decreases exponentially with increasing flow rate [1].

FAQ 3: My electrospray is unstable in a high-aqueous solvent. Which property should I adjust?

High water content typically means high surface tension, which destabilizes the Taylor cone. The most effective adjustment is to modify the surface tension.

• Symptom: The spray flickers, is intermittent, or cannot be initiated at achievable voltages.
• Primary Cause: High surface tension resists the formation of the fine jet required for nanoelectrospray.
• Protocol for Correction:
  • Identify Additives: Add a organic modifier, such as methanol or acetonitrile, to your aqueous solution. A starting point is a 1:1 ratio of water:organic solvent.
  • Optimize Concentration: Titrate the organic modifier percentage while monitoring spray stability and ion signal. These modifiers lower surface tension and enhance desolvation.
  • Consider "Supercharging" Additives: For protein analysis, additives like dimethyl sulfoxide (DMSO) or sulfolane can be used to increase charge states, which also modifies solvent properties [15].

Experimental Protocols

Protocol 1: Systematic Evaluation of Ion Suppression

This methodology allows you to quantitatively measure the degree of ion suppression in your specific experimental setup [1].

Methodology:

• Prepare Solution: Create an equimolar mixture of a neutral, poorly-ionizable compound (e.g., maltotetraose) and a readily-ionized compound (e.g., neurotensin).
• Infuse Sample: Infuse the mixture at a series of defined, low flow rates (e.g., from 10 nL/min to over 300 nL/min).
• Data Acquisition: Acquire mass spectra at each flow rate.
• Data Analysis: For each flow rate, calculate the signal intensity ratio of the neutral analyte (maltotetraose) to the charged analyte (sum of all charge states of neurotensin).
• Interpretation: Plot the ratio against the flow rate. A higher ratio at lower flow rates confirms reduced ion suppression, demonstrating the advantage of ultra-low flow nanoelectrospray.

Protocol 2: Utilizing Theta Emitters for Salty or Buffered Solutions

This protocol is for analyzing proteins and complexes directly from physiologically relevant buffers containing non-volatile salts, which normally suppress ionization [12].

Methodology:

• Emitter Preparation: Use a dual-channel theta emitter pulled to an internal diameter of approximately 1.4 µm.
• Loading Solutions:
  • Channel A: Load with your protein sample dissolved in the biological buffer (e.g., PBS).
  • Channel B: Load with a volatile MS-compatible salt solution (e.g., 200 mM ammonium acetate) potentially supplemented with an additive like sodium iodide or bromide (anions with low proton affinity).
• Initiate Spray: Apply a high voltage (0.8 – 2.0 kV) via platinum wires inserted into each channel.
• Mechanism: Incomplete mixing of the two streams at the emitter tip promotes the formation of a population of droplets relatively depleted of non-volatile salts, enabling the observation of the protein ions [12].
• Gas-Phase Activation: Employ gas-phase collisional heating methods (e.g., beam-type CID and dipolar direct current) to remove residual salt adducts after ionization.

Data Presentation

Table 2: Solvent Properties and Their Impact on the Electrospray Process

Solvent Property	Role in Electrospray	Desired Trend for NanoESI	Common Method for Optimization
Surface Tension	Determines the voltage required to overcome cohesive forces and form a Taylor cone.	Lower	Add organic modifiers (MeOH, ACN).
Conductivity	Influences the current carried by the spray and the charge density on droplets.	Moderate to High (for efficient fission)	Add volatile electrolytes (ammonium acetate, formic acid).
Electrical Permittivity (Dielectric Constant)	Affects the ability of the solvent to sustain the electric field required for electrospray.	Higher	Use solvents with higher dielectric constants (e.g., water, DMSO).

Visualization of Troubleshooting Logic

The following diagram outlines a logical workflow for diagnosing and resolving nanoelectrospray issues related to solvent properties.

The Scientist's Toolkit: Research Reagent Solutions

Table 3: Essential Materials for High-Sensitivity Nanoelectrospray MS

Item	Function in Experiment	Rationale
Theta Emitters	Dual-channel emitters for analyzing samples in non-volatile buffers.	Allows incomplete mixing of sample and volatile buffer streams, creating droplets depleted of suppressing salts [12].
Volatile Buffers	e.g., Ammonium Acetate, Formic Acid.	Provides necessary conductivity for electrospray without leaving non-volatile residues that cause adduction and suppression [12] [1].
Organic Modifiers	Methanol, Acetonitrile.	Lowers solvent surface tension for stable spray and improves desolvation efficiency [1].
Low Proton Affinity Anions	e.g., Iodide or Bromide salts.	When added to the buffer channel of a theta emitter, can help mitigate ionization suppression by sequestering sodium ions [12].
Surface-Modified nESI Needles	e.g., Polyethylene-glycol coated emitters.	Reduces nonspecific adsorption of analytes to the glass surface, which can improve signal intensity for challenging molecules like proteins and viral capsids [9].

Advanced nESI Methodologies for Challenging Biomolecular Analyses

The direct analysis of proteins and protein complexes from physiologically relevant buffers using native mass spectrometry (nMS) is a significant challenge due to the interference of non-volatile salts. These salts can suppress ionization, cause extensive adduction, and generate chemical noise that obscures the signals of biological ions of interest [12]. Theta emitters, a specialized nanoelectrospray ionization (nESI) tool, present a powerful solution to this problem. This technical support guide provides detailed troubleshooting and methodologies for researchers aiming to implement theta emitters to improve sensitivity and enable direct analysis from high-salt solutions in drug discovery and development [16] [17].

Theta Emitter FAQ

Q1: What is a theta emitter and how does it overcome the salt challenge? A theta emitter is a glass nanoelectrospray capillary with an internal septum that divides it into two separate channels [12]. This unique design allows you to load your protein sample, dissolved in a biological buffer (e.g., containing Tris, HEPES, or NaCl), into one channel, while loading a volatile MS-compatible solution (like ammonium acetate) into the other channel [16] [17]. The two streams mix at the very tip of the emitter just as electrospray is initiated. This process promotes the formation of a population of electrospray droplets that are relatively depleted of non-volatile salts, thereby reducing salt adduction and ionization suppression and enabling direct analysis from physiologically relevant conditions [12].

Q2: My protein signal is still suppressed despite using a theta emitter. What can I do? Signal suppression is often linked to the type of anions present in the solution. A proven method to mitigate this is to add anions with low proton affinity, such as bromide or iodide, to the volatile buffer channel (e.g., ammonium acetate) [12]. These anions compete effectively with your analyte for charge and help remove sodium ions from the droplets, significantly reducing chemical noise. Studies have shown that this strategy can increase the signal-to-noise (S/N) ratios of protein ions, enhance method reproducibility, and improve overall robustness [12].

Q3: My theta emitter keeps clogging. How can I prevent this? Clogging is a common issue with narrow-diameter emitters. To minimize this:

• Ensure Sample Purity: Centrifuge your protein samples before loading to remove any particulate matter.
• Use Clean Solutions: Filter all buffers through 0.22 µm filters.
• Optimize Emitter Pulling: If fabricating emitters in-house, fine-tune the pulling parameters on your pipette puller to produce more consistent tip geometries. Commercially sourced emitters may offer higher reproducibility [12].

Q4: Why is my spectrum noisy with broad peaks even with the theta emitter setup? Chemical noise and broad peaks are frequently caused by incomplete desolvation or residual salt adducts. To address this:

• Optimize Gas-Phase Activation: Tune your mass spectrometer's collisional activation settings. Beam-type collision-induced dissociation (BTCID) followed by dipolar direct current (DDC) activation in a linear ion trap can effectively remove weakly-bound salt adducts and solvent molecules without causing protein dissociation [12].
• Fine-tune Voltages: Systematically adjust the applied ESI voltage (typically starting from 0.8 kV up to 2.0 kV) to establish a stable spray, which is crucial for generating clean spectra [12].

Key Experimental Protocols and Data

Protocol: Theta Emitter Setup for Direct Analysis from Physiological Buffers

This protocol details the setup for analyzing proteins from biological buffers using theta emitters, adapted from recent research [12].

Step 1: Theta Emitter Preparation

• Pull borosilicate glass theta capillaries (e.g., 1.5 mm o.d., 1.17 mm i.d.) using a micropipette puller (e.g., Sutter Instrument P-87) with customized heating and pulling parameters to achieve tip inner diameters of approximately ~1.4 µm [12].
• Under a microscope, insert dual platinum wires into the back of the theta emitter, ensuring each wire makes contact with the solution in one of the two channels.

Step 2: Sample and Additive Loading

• Channel 1 (Sample): Load your protein sample (e.g., 5-10 µL) dissolved in its native biological buffer (e.g., PBS, Tris, HEPES) which may contain physiological concentrations of NaCl [12].
• Channel 2 (Additive): Load a volatile buffer, typically 199 mM ammonium acetate, optionally supplemented with an additive salt like 150 mM ammonium bromide (NH₄Br) or ammonium iodide (NH₄I) [12].

Step 3: Mass Spectrometry Analysis

• Position the theta emitter approximately 1-2 mm from the mass spectrometer's curtain plate, orthogonal to the orifice.
• Apply a voltage of 0.80 - 2.0 kV to the platinum wires to initiate the electrospray. Begin at 800 V and increase in 50-100 V increments until a stable spray and analyte ions are observed.
• Employ gas-phase activation techniques in the mass spectrometer. First, use beam-type CID (e.g., in a quadrupole collision cell with N₂ bath gas) for initial desolvation and adduct removal. Follow this with dipolar DDC in a linear ion trap to provide additional activation for final cleaning of the ion signal [12].

The workflow and proposed mechanism of signal enhancement are illustrated below.

Performance Data: Theta Emitters with Solution Additives

The following table summarizes quantitative data on the performance of theta emitters used with different solution additives for analyzing proteins from high-salt solutions. Data is compiled from a 2025 study [12].

Table 1: Performance of Theta Emitters with Different Solution Additives for Protein Analysis

Protein / Complex	Mass (kDa)	Biological Buffer Conditions	Additive in AmAc	Key Improvement (S/N or FWHM)
Lysozyme	14	137 mM NaCl, 50 mM Tris-HCl	150 mM NH₄Br	Significant increase in S/N ratio compared to AmAc alone [12]
β-Lactoglobulin Dimer	~36	137 mM NaCl, 50 mM Tris-HCl	150 mM NH₄Br	Improved S/N and spectral reproducibility [12]
Pyruvate Kinase Tetramer	~236	137 mM NaCl, 50 mM Tris-HCl	150 mM NH₄I	Observable complex with reduced adduction; increased S/N [12]
Streptavidin Tetramer	~53	137 mM NaCl, 50 mM Tris-HCl	150 mM NH₄Br	Robust signal with lower FWHM (narrower peaks) [12]

FWHM: Full Width at Half Maximum, a measure of peak broadening.

The Scientist's Toolkit: Essential Research Reagents & Materials

Successful implementation of the theta emitter technique requires specific reagents and equipment. The table below lists the key components.

Table 2: Essential Materials for Theta Emitter Experiments

Item	Function / Description	Example Specifications / Notes
Theta Capillaries	Dual-channel glass capillaries for separate sample and additive introduction.	Borosilicate glass, 1.5 mm o.d., 1.17 mm i.d.; pulled to ~1.4 µm tip i.d. [12]
Volatile Buffer	MS-compatible buffer in the second channel; maintains native structure.	100-200 mM Ammonium Acetate (AmAc), pH ~6.8-7.2 [16] [17]
Low Proton Affinity Additives	Anionic salts added to volatile buffer to reduce chemical noise and adduction.	Ammonium Bromide (NH₄Br) or Ammonium Iodide (NH₄I), typically 150 mM [12]
Micropipette Puller	Instrument to fabricate nanoESI emitters with consistent tip geometry.	e.g., Sutter Instrument P-87 with optimized heating and pulling parameters [12]
Platinum Wires	Electrodes to apply high voltage to the solutions in each channel.	Dual wires supported by a single holder [12]
Gas-Phase Activation Module	Instrument components for removing residual solvent and salt adducts.	Beam-type CID and Dipolar DC (DDC) capabilities [12]

Direct Infusion Nanoelectrospray High-Resolution Mass Spectrometry (DI-nESI-HRMS) is a fit-for-purpose analytical method that enables rapid, targeted parallel analysis of numerous metabolites in biological samples. This technique is particularly valuable for large-scale epidemiological investigations, as it eliminates chromatographic separation, significantly reducing analysis time to approximately 2 minutes per sample while requiring minimal sample volume (less than 10 μL) [18].

The method generates high-resolution MS profiles in both positive and negative polarities, enabling both targeted quantification and untended data mining for hundreds of metabolites. Its application has been successfully demonstrated in characterizing population-specific metabolic phenotypes, such as differences between U.S. and Japanese populations in the INTERMAP study, and in assessing urinary markers as predictors of type 2 diabetes in the ARIC study [18].

Key Advantages and Limitations

• High Throughput: Enables rapid analysis of large sample cohorts (e.g., >10,000 samples in 12 weeks) [18].
• Minimal Sample Consumption: Requires less than 10 μL of biofluid, ideal for precious biobanked samples [18].
• Broad Metabolite Coverage: Capable of profiling amino acids, TCA cycle metabolites, fatty acids, acylcarnitines, and gut microbial cometabolites [18].
• Ion Suppression: Competition for charge can suppress ionization of some analytes due to the simultaneous infusion of all sample components [18].
• Inability to Separate Isomers: Cannot distinguish structural isomers without chromatographic separation or additional MS/MS experiments [18].
• Spectral Complexity: Spectra contain multiple signal types per metabolite (e.g., adducts, in-source fragments), complicating data interpretation [18].

Optimized Experimental Protocol for DI-nESI-HRMS

Sample Preparation

• Extraction: Use a methanol/chloroform/water solvent system (final ratio 2:2:1.8) for polar metabolite extraction from tissues [19].
• Reconstitution: Reconstitute dried sample extracts in 80:20 methanol/water containing 0.25% formic acid [19].
• Cleanup: Centrifuge reconstituted samples at 15,000×g at 4°C for 10 minutes to remove particulate matter prior to analysis [19].

Instrument Configuration and Data Acquisition

• Ionization Source: Utilize a chip-based nanoelectrospray ionization system (e.g., Advion TriVersa NanoMate) for stable, efficient ionization at low flow rates [18].
• Mass Analyzer: Operate with a high-resolution time-of-flight (TOF) mass spectrometer for flexible scan rates and accurate mass measurement [18].
• Data Acquisition:
  • Acquire data in both positive and negative ionization modes.
  • Employ a 2-minute infusion time per sample.
  • For ultrahigh-resolution instruments (e.g., FT-ICR), consider using the spectral stitching technique to maintain sensitivity and dynamic range [18].

Quality Control Procedures

• Pooled QC Samples: Analyze pooled quality control (QC) samples at the beginning and end of each batch and after every five biological samples to monitor performance [19].
• Blank Samples: Run solvent blanks to identify and remove background peaks [19].
• System Equilibration: Analyze two QC samples at the start of a sequence for system equilibration [19].
• Batch Correction: Apply computational batch correction algorithms (e.g., Quality Control-Robust Spline Correction) to correct for intra- and inter-batch variation using the data from the repeatedly-measured QC samples [19].

Workflow Visualization

Performance and Metabolite Quantification

Key Metabolites and Their Analytical Ranges

The following table summarizes exemplary metabolites quantifiable by DI-nESI-HRMS, their biochemical functions, and typical linear ranges demonstrated in large-scale studies [18].

Metabolite	Biochemical Function	Linear Range (μg/mL)	Applicable Study
Hydroxycinnamic acid	Marker of polyphenols consumption	0.1 – 3.3	INTERMAP
Acetylcarnitine	Fatty acid oxidation	0.05 – 1.7	INTERMAP, ARIC
Ascorbic acid	Vitamin C	0.1 – 3.3	INTERMAP
Benzoic acid	Phenylalanine, Tyrosine metabolism	0.5 – 16.7	INTERMAP, ARIC
Citric acid	TCA cycle	0.3 – 12.5	INTERMAP, ARIC
Creatinine	Cell's energy shuttle	1.6 – 50	INTERMAP, ARIC
Glutamic acid	Urea cycle, Glucose-Alanine cycle	0.2 – 6.7	INTERMAP, ARIC

Sensitivity Enhancement Techniques

• SPIN Source: The Subambient Pressure Ionization with Nanoelectrospray (SPIN) source places the emitter directly in the first reduced-pressure region of the mass spectrometer (10-30 Torr), essentially eliminating ion losses at the inlet capillary and increasing ion transmission efficiency [10].
• Multi-Emitter Arrays: Using an array of emitters, each with an individualized sheath gas capillary, splits the liquid flow into multiple nano-electrosprays. This increases total ion current and improves desolvation, leading to a sensitivity increase proportional to the number of emitters [10].
• Surface-Modified Emitters: Chemical etching and surface modification of nano-electrospray needles can further enhance ionization efficiency and stability, particularly for challenging applications like native mass spectrometry [20].

Troubleshooting FAQs & Guides

Droplet Detection and Dispensing Issues

Problem: High rate of false negatives (liquid is dispensed, but not detected by the system).

• Cause & Solution: This is often due to optical obstruction or insufficient liquid volume.
  • Clean the bottom of the source tray and each DropDetection opening using lint-free swabs and 70% ethanol [21].
  • Ensure source wells are filled with sufficient volume (e.g., 10-20 μL) and are free of air bubbles [21].
  • Acceptance Criterion: After cleaning, the number of non-detected droplets in a test run should not exceed 1% of the total (e.g., ≤10 droplets out of 1056) [21].

Problem: Droplets landing out of position on the target plate.

• Cause & Solution: The target tray position may be misaligned.
  • Visually check the landing position by dispensing to a transparent target plate.
  • In the software's advanced settings, use the "Move To Home" function and manually adjust the target tray position. Restart the software after adjustment [21].

Ion Suppression and Signal Instability

Problem: Significant ion suppression, leading to poor sensitivity for some analytes.

• Cause & Solution: This is a fundamental challenge in DIMS where all analytes compete for charge.
  • Sample Dilution: Dilute the sample to reduce overall matrix complexity.
  • NanoESI Flow Rates: Leverage the inherent reduction in ion suppression achieved by low nanoESI flow rates, which generate smaller charged droplets and improve ionization efficiency [18].
  • Stable Spray: Ensure a stable spray by using a chip-based nanoESI source, which is less prone to contamination and provides consistent flow [18].

Problem: Unstable spray or fluctuating signal intensity during infusion.

• Cause & Solution: This can be caused by a partially clogged emitter, poor electrical contact, or solvent issues.
  • Visually inspect the nanoESI chip nozzle for obstructions.
  • Ensure the sample is properly centrifuged to remove particulates [19].
  • Check that the solvent composition is compatible with the nanoESI process and contains a volatile acid/base modifier (e.g., 0.1% formic acid).

Pressure and Leakage Errors

Problem: "Pressure Leakage/Control Error" message appears.

• Cause & Solution: This indicates a poor seal in the fluidic path.
  • Confirm all source wells are fully inserted into the plate and none are missing [21].
  • Check that the dispense head is correctly aligned and positioned approximately 1 mm above the source plate. A 0.8 mm plastic card can be used to gauge this distance [21].
  • Inspect the head rubber for visible damage, cuts, or rips. Contact technical support if damage is found [21].

System and Software Errors

Problem: The instrument does not start even though the on/off button is green.

• Cause & Solution: The instrument's lid may have been open during power-on.
  • Close the lid, switch off the main power, and toggle the on/off switch again [21].

Problem: Created protocol fails to run or is interrupted.

• Cause & Solution:
  • Verify the air pressure supply is connected and within the 3-10 bar (40-145 psi) range [21].
  • Check that the correct liquid class is assigned to the protocol. Different plates and solvents require specific liquid classes for optimal performance [21].

Research Reagent Solutions

The table below lists essential materials and reagents for establishing a robust DI-nESI-HRMS workflow.

Item	Function / Application	Specification / Notes
TriVersa NanoMate	Chip-based nanoelectrospray ionization source	Provides stable, automated nanoESI with minimal cross-contamination [18] [22].
High-Res Mass Spectrometer	Mass analysis	Q-TOF instruments offer a balance of resolution, scan speed, and accessibility [18].
Methanol, Chloroform, Water	Metabolite extraction from tissues	HPLC grade; used in a biphasic system (2:2:1.8 ratio) for polar metabolite isolation [19].
Formic Acid	Mobile phase additive	HPLC grade; used at 0.1-0.25% in the reconstitution solvent to promote protonation in positive ion mode [18] [19].
Fused Silica Capillaries	Custom emitter fabrication	e.g., 150 μm o.d., 10 μm i.d. for creating multi-emitter arrays for sensitivity enhancement [10].
Ceramic Bead Homogenization Tubes	Tissue homogenization	Ensures efficient and reproducible disruption of tissue samples prior to metabolite extraction [19].

Pulsed nanoelectrospray ionization (pulsed nESI) represents a significant advancement in mass spectrometry (MS) for the analysis of intact proteins. Unlike conventional direct current (DC) nESI, which applies a constant voltage, pulsed nESI rapidly cycles the high voltage on and off. This modulation, occurring at high frequencies (typically 10–350 kHz) with sub-nanosecond rise times, fundamentally changes the electrospray process [15]. The technology addresses a key limitation of conventional DC ESI: the formation of relatively large initial droplet sizes that can limit efficient ion desolvation and overall sensitivity [15]. By applying the voltage in pulses, researchers can generate significantly smaller initial droplets and reduce Coulombic repulsion within the spray plume, leading to enhanced ion abundances and improved signal-to-noise ratios for biomolecular analysis [15].

The relevance of pulsed nESI is particularly pronounced in the context of top-down proteomics and the analysis of intact protein complexes. When proteins are ionized from denaturing solutions, they typically produce broad charge state distributions that disperse the ion signal across multiple detection channels [15]. Pulsed nESI technology helps concentrate this signal, making it especially valuable for applications where sample is limited or when analyzing low-abundance species from complex mixtures. The enhanced sensitivity achieved through pulsed nESI is anticipated to benefit various tandem mass spectrometry measurements, including those involving electron capture dissociation (ECD), electron transfer dissociation (ETD), and ultraviolet photodissociation (UVPD), where the extent of dissociation and sequence coverage often increases with both the charge state and abundance of the precursor ion [15].

Technical FAQs: Optimizing Pulsed nESI Experiments

What are the key advantages of pulsed nESI over conventional DC nESI for intact protein analysis?

Pulsed nESI offers several distinct advantages for analyzing intact proteins. Research demonstrates that implementing pulsed nESI with optimal parameters can increase absolute ion abundances of protonated proteins by up to 82% and boost signal-to-noise ratios by up to 154% compared to conventional DC nESI-MS [15]. These improvements stem from fundamental changes in the electrospray process. The pulsed voltage creates a sharper and longer Taylor cone with a smaller half-angle (~12°) compared to DC ESI (~47°), resulting in smaller initial droplet sizes and less radial dispersion of the aerosol plume [15]. Additionally, the pulsed operation reduces the heating effect on the capillary tip, allowing for the application of higher voltages than conventional DC nano-ESI sources, which further boosts ionization efficiency [23].

What are the optimal pulsed voltage parameters for maximizing protein ion signal?

Extensive parameter optimization has revealed that specific pulsed voltage settings maximize performance for protein analysis. The table below summarizes the key parameters and their optimal ranges based on experimental findings:

Table 1: Optimal Pulsed nESI Parameters for Protein Analysis

Parameter	Optimal Range	Impact on Performance
Repetition Rate	~200 kHz [15]	Maximizes ion abundance and S/N for proteins; 200-250 kHz effective for smaller ions (≤1032 m/z)
Pulse Voltage	0 to ~1.5 kV [15]	Sufficient to initiate and maintain stable nESI with nanoscale emitters
Voltage Rise Time	<1 nanosecond [15]	Ensures rapid and precise pulse transitions
Duty Cycle	10% to 90% [15]	Affects the average current and can be tuned for specific applications

How does emitter tip size affect pulsed nESI performance?

Emitter tip diameter significantly influences the electrospray process and overall system performance in pulsed nESI. Nanoscale emitters with inner diameters of approximately 250-300 nm are commonly used [15]. The equivalent resistance of a nano-ESI source changes with respect to both the emitter tip diameter and the applied high voltage [23]. Smaller tip diameters more effectively concentrate the electric field at the emitter tip, which reduces the voltage required to initiate ESI and produces initial droplets with very high surface-to-volume ratios [15]. This enhances desolvation efficiency and improves ion transfer through the atmospheric pressure interface of the mass spectrometer. Importantly, the use of nanoscale emitters also significantly reduces the adduction of non-volatile salts and molecules to protein ions, which is particularly beneficial for maintaining spectral quality when analyzing proteins or protein-ligand complexes from native-like solutions [15].

My protein signal is unstable with pulsed nESI. What should I check?

Signal instability can arise from several sources in pulsed nESI setups. First, verify the stability of your high-voltage pulses using an oscilloscope to ensure consistent pulse shape, frequency, and amplitude. Next, inspect the nanoESI emitter under a microscope for any damage or partial clogging—even minor imperfections can dramatically affect spray stability [15]. Ensure your emitter is properly positioned relative to the mass spectrometer inlet, typically within 5-10 mm, as this distance affects the electric field and ion transmission efficiency [24]. Finally, evaluate your solution conditions; the use of denaturing solutions (acidified with organic modifiers) facilitates protein elongation and higher charge states, but the addition of supercharging agents like dimethyl sulfoxide (DMSO) or 1,2-butylene carbonate should be optimized as they can significantly alter solution properties and spray stability [15].

Can pulsed nESI be combined with other ionization techniques?

Yes, pulsed nESI can be effectively integrated with other ionization methods to expand analytical capabilities. Researchers have successfully implemented alternately pulsed configurations where nESI and atmospheric pressure chemical ionization (APCI) are operated sequentially using the same atmospheric interface and ion path [25]. This approach is particularly valuable for ion/ion reaction experiments, where one ionization source generates multiply charged protein ions while the other produces singly charged reagent ions of opposite polarity [25]. Such configurations enable important processes like proton transfer reactions for charge reduction of proteins and electron transfer dissociation for peptide sequencing, all without requiring major modifications to commercial mass spectrometer hardware [25]. More recent developments also include dual non-contact nESI/nAPCI sources that allow simultaneous detection of both polar and nonpolar analytes from microliter sample volumes [24].

Troubleshooting Guides

Low Ion Abundance

Table 2: Troubleshooting Low Ion Abundance

Problem	Possible Cause	Solution
Insufficient Ion Signal	Suboptimal pulse frequency	Systematically test repetition rates between 10-350 kHz, focusing on ~200 kHz for proteins [15].
	Low pulse voltage	Ensure voltage amplitude reaches 1.0-1.5 kV for stable spray with nanoscale emitters [15].
	Emitter tip too large	Use emitters with inner diameters ~250 nm for smaller initial droplets [15].
	Improper emitter alignment	Position emitter 5-10 mm from MS inlet and align axially for optimal ion transmission [24].

Electrical and Spray Stability Issues

Table 3: Troubleshooting Spray Stability

Problem	Possible Cause	Solution
Unstable Spray Current	Arc-over or electrical breakdown	For pulsed nESI, the electrical breakdown limit is lower than DC; reduce voltage or use smaller emitter [15].
	Joule heating at tip	Use pulsed HV to reduce heating effect, allowing higher voltages than DC [23].
	Non-volatile salts in solution	Use nanoscale emitters (<1 µm) to reduce salt adduction [15] or implement in-capillary electrophoresis [24].
Rapid Tip Damage	Excessive current	Pulsed operation reduces overall current; if problem persists, further reduce duty cycle [15].

Experimental Protocols

Protocol: Benchmarking Pulsed nESI Performance for Intact Proteins

This protocol describes how to quantitatively compare the performance of pulsed nESI against conventional DC nESI for the analysis of intact proteins, using parameters validated in recent literature [15].

Materials Required:

• Proteins: Ubiquitin, cytochrome C, myoglobin, and carbonic anhydrase II as standard test proteins [15]
• Solvent System: Denaturing solution of 49.5/49.5/1% (v/v) water/methanol/acetic acid [15]
• nESI Emitters: Borosilicate capillaries pulled to inner diameters of ~250 nm [15]
• Mass Spectrometer: ESI-equipped mass spectrometer (any type suitable for intact protein analysis)

Procedure:

• Sample Preparation: Prepare individual protein solutions at 1-10 µM concentration in the denaturing solvent system.
• Emitter Loading: Load approximately 2-5 µL of protein solution into a nanoscale emitter using microloader tips.
• DC nESI Reference Measurement: Apply a conventional DC voltage of 0.9-1.2 kV to establish a stable nanoelectrospray. Record mass spectra for 1-2 minutes to establish baseline performance.
• Pulsed nESI Measurement: Switch to pulsed mode with the following initial parameters:
  • Voltage amplitude: 0 to ~1.5 kV
  • Repetition rate: 200 kHz
  • Duty cycle: 50%
  • Rise time: <1 ns Record mass spectra for 1-2 minutes.
• Parameter Optimization: Systemically vary the repetition rate (10-350 kHz) and duty cycle (10-90%) to identify optimal conditions for your specific protein and instrument configuration.
• Data Analysis: Compare absolute ion abundances (peak heights) and signal-to-noise ratios for the same charge states between DC and pulsed operation.

Expected Outcomes: When optimized, pulsed nESI should increase ion abundances by up to 82% and signal-to-noise ratios by up to 154% compared to DC nESI for the test proteins. For protein mixtures, signals for individual components may increase by up to 184% [15].

Protocol: Implementing Alternately Pulsed nESI/APCI for Ion/Ion Reactions

This protocol describes the setup for alternately pulsed nESI and APCI sources to enable ion/ion reaction experiments, which are valuable for protein charge reduction and electron transfer dissociation [25].

Materials Required:

• nESI Emitters: Borosilicate capillaries (1.55 mm O.D., 0.86 mm I.D.) pulled to tip orifice diameters of 30 µm [25]
• APCI Electrode: Platinum wire (0.1 mm O.D.) [25]
• Power Supplies: One high voltage power supply from the instrument and one fast high voltage pulser (e.g., GRX-1.5K-E) [25]
• Reagent Materials: Proton sponge or perfluoro(methyldecalin) for APCI reagent generation [25]

Procedure:

• Source Arrangement: Position the nano-ESI emitter 10-20 mm from the MS sampling orifice. Place the APCI Pt wire parallel to the nESI emitter, 3-10 mm away, to achieve optimal reagent ion signals [25].
• Electrical Configuration: Connect the high voltage power supply to the nESI emitter and the fast high voltage pulser to the APCI Pt wire. Ensure both can be independently triggered from the instrument control software.
• Pulsing Sequence Setup: Program the pulse sequence with the following timing:
  • Pulse nESI on for 50 ms to generate multiply charged protein ions
  • Switch nESI off and isolate/cool ions in the trap for 50 ms
  • Pulse APCI on for 50 ms to generate singly charged reagent ions of opposite polarity
  • Allow reaction period for ion/ion interactions [25]
• Voltage Optimization: Optimize "on" voltages for each source: ±1.2-2.0 kV for nESI emitter and ±3 kV for APCI Pt wire [25].
• Experiment Execution: Implement ion/ion reaction experiments such as:
  • Partial deprotonation of multiply protonated proteins
  • Electron transfer to multiply protonated peptides

Technical Notes: The pulsed operation enables stable ion production from each source and allows ions of opposite polarity to be generated and injected into the mass spectrometer separately without significant compromise in the performance of either ion source [25].

Research Reagent Solutions

Table 4: Essential Research Reagents for Pulsed nESI Experiments

Reagent/Category	Specific Examples	Function and Application
Standard Test Proteins	Ubiquitin, Cytochrome C, Myoglobin, Carbonic Anhydrase II [15]	Benchmarking pulsed nESI performance and optimizing parameters
Denaturing Solvents	Water/Methanol/Acetic Acid (49.5/49.5/1%) [15]	Elongate protein conformations for higher charge states in top-down MS
Supercharging Additives	1,2-Butylene Carbonate, DMSO, Sulfolane [15]	Increase protein charge states; improve MS/MS efficiency
nESI Emitters	Borosilicate Capillaries (~250 nm i.d.) [15]	Produce smaller initial droplets; enhance desolvation efficiency
APCI Reagents	Proton Sponge, Perfluoro(methyldecalin) [25]	Generate reagent ions for ion/ion proton or electron transfer reactions

Technology Workflow and Relationships

The following diagram illustrates the key decision points in implementing and optimizing a pulsed nESI method for intact protein analysis:

Pulsed nESI Method Workflow

The relationship between electrical parameters and analytical performance in pulsed nESI can be visualized as follows:

Parameter to Performance Relationships

FAQs: Core Technology and Applications

Q1: What is the fundamental advantage of a dual nESI/nAPCI source compared to standard nESI? The primary advantage is its ability to simultaneously ionize and detect both polar and non-polar analytes from a single, small-volume sample without pre-treatment. Standard nESI efficiently ionizes polar molecules but often fails for non-polar compounds. The dual source combines electrospray for polar analytes and corona discharge-induced chemical ionization for non-polar ones, providing a more universal detection method for complex mixtures [26] [27] [24].

Q2: How does this technology improve sensitivity in MS analysis? It improves sensitivity through several mechanisms:

• Microsample Analysis: It is optimized for analyzing microliter-volume samples (≤ 5 µL) with minimal dilution, concentrating the analytes [24].
• Reduced Matrix Effects: The non-contact setup and potential for in-capillary extraction help reduce ion suppression from complex matrices like blood [24].
• Direct Analysis: It enables ambient ionization, integrating analyte extraction and ionization into a single step. This eliminates sample preparation steps that can lead to analyte loss [27] [24].

Q3: Can this source analyze proteins in physiological buffers with high salt? Yes. The platform can be activated for electrophoretic separation spray mode. By applying alternating high voltages (e.g., from -5 kV to 2 kV), it enables the efficient detection of proteins and protein complexes even in buffers containing high concentrations of non-volatile salts, mimicking a physiologically relevant environment [12] [24].

Q4: What is the operational principle behind the simultaneous nESI and nAPCI? A single high-voltage power supply is used. At lower voltages (≤ 3 kV), classic nESI occurs, generating protonated ions for polar compounds. When the voltage is ramped above a threshold (> 4 kV), it induces a corona discharge from an auxiliary electrode, activating the nAPCI process. This discharge ionizes gas-phase species and generates molecular ions for non-polar analytes, all from the same emitter [27] [24].

Troubleshooting Guide

The following table outlines common experimental issues, their potential causes, and recommended solutions.

Problem	Possible Cause	Recommended Solution
Signal Suppression for Non-polar Analytes	Insufficient voltage for corona discharge; nAPCI mode not activated.	Ensure the spray voltage is ramped above 4 kV to initiate the corona discharge. Verify the placement and integrity of the auxiliary electrode [27] [24].
Unstable Spray or No Spray	Emitter tip damage; improper electrode alignment; insufficient voltage for the given tip size.	Inspect the glass emitter tip for damage or clogging under a microscope. In a non-contact setup, ensure the high-voltage electrode is correctly positioned with a ~1 cm air gap [24].
Excessive Chemical Noise & Salt Adduction	High concentration of non-volatile salts in the sample matrix.	For protein analysis, use theta emitters to mix the sample with an ammonium acetate solution containing additives like bromide or iodide anions, which help mitigate salt adduction [12].
Poor Sensitivity & Low Signal-to-Noise	Sample matrix effects; emitter tip damage; suboptimal interface parameters.	Utilize in-capillary liquid/liquid extraction to pre-concentrate analytes and remove matrix interferents. Check the MS inlet capillary temperature and lens voltages [24].
Burning or Breakage of Glass Emitter Tip	Excessive Joule heating from high voltage in contact-mode operation.	Switch to a non-contact charging mode. This uses electrostatic induction to charge the solution, allowing the application of high voltages (e.g., 6 kV) without physical contact, thereby preventing tip damage [24].

Experimental Protocol: Direct Analysis from a Complex Matrix

This protocol details the analysis of small molecules from untreated whole blood using the dual nESI/nAPCI source with in-capillary extraction [24].

1. Apparatus Setup:

• Ion Source: Configure the dual non-contact nESI/nAPCI setup.
• Emitter: Use a disposable borosilicate glass capillary (e.g., 1.17 mm i.d.) pulled to a tip diameter of ≤ 5 µm.
• Mass Spectrometer: The method is demonstrated on a Velos Pro LTQ mass spectrometer but can be adapted to others.

2. Reagent and Sample Preparation:

• Extraction Solvent: Prepare a mixture of ethyl acetate and dichloromethane.
• Internal Standard: Prepare a solution of cocaine-D3 in methanol (e.g., 100 µg/mL).
• Sample: Spike the target analyte (e.g., cocaine) and internal standard into 5 µL of untreated whole blood.

3. In-Capillary Extraction Procedure:

• Load the 5 µL spiked blood sample into the open end of the glass capillary.
• Introduce approximately 5 µL of the organic extraction solvent behind the blood sample. A small air gap can be used to separate the two phases.
• Gently tap or centrifuge the capillary to combine the phases within the capillary body. The organic solvent will extract the analytes from the blood matrix.

4. Mass Spectrometry Analysis:

• Spray Voltage: Apply a dynamic voltage ramp from 2 kV to 6 kV.
• Capillary Temperature: Set to 400 °C.
• S-Lens RF Level: Set to 60%.
• Distance: Position the emitter tip ~5 mm from the MS inlet.
• Data Acquisition: Record spectra over a 30-second period. At lower voltages, polar analytes are detected via nESI. As the voltage surpasses 4 kV, nAPCI activates, enabling the detection of non-polar compounds like β-estradiol from the same run.

Workflow and Signaling Pathways

The following diagram illustrates the logical workflow and operational modes of the dual nESI/nAPCI source.

The Scientist's Toolkit: Research Reagent Solutions

The table below lists key reagents and materials used in experiments with dual nESI/nAPCI sources and their specific functions.

Reagent / Material	Function / Application
Borosilicate Capillaries (1.17-1.2 mm i.d.)	Used to fabricate nanoelectrospray emitters. Pulled to fine tips (≤5 µm) to enable stable spray at low flow rates [24].
Ammonium Acetate (AmAc)	A volatile MS-compatible salt. Used for buffer exchange to reduce non-volatile salt adducts, often in the second channel of a theta emitter [12].
Bromide (Br⁻) or Iodide (I⁻) Anions	Added to AmAc as solution additives. Their low proton affinity helps reduce sodium adduction to proteins and chemical noise in salty solutions [12].
Theta Emitters (Septum-divided capillaries)	Allow simultaneous loading of a sample (in biological buffer) and a conditioning solution (e.g., AmAc with additives). Incomplete mixing creates droplets favorable for ionization [12].
Ethyl Acetate / Dichloromethane	Organic solvents used for in-capillary liquid/liquid extraction to isolate analytes from complex biological matrices like whole blood directly within the spray emitter [24].
Auxiliary Electrode (e.g., Silver Wire)	Placed coaxially near the emitter tip to generate a stable corona discharge at high voltages, which is essential for the nAPCI process [24].

Practical Optimization and Troubleshooting for Maximum nESI Sensitivity

Troubleshooting Guides

FAQ: How does emitter position affect my mass spectrometry results, and how can I optimize it?

The position of the nanoelectrospray ionization (nESI) emitter relative to the instrument inlet is a critical parameter that influences both signal stability and the degree of in-source activation, which can inadvertently induce collision-induced dissociation (CID) or unfolding (CIU) [4].

• Problem: Unintended in-source activation causing protein dissociation or unfolding.
• Root Cause: Emitter positions too close to the instrument inlet can subject ions to stronger electric fields and collisional heating before they are fully desolvated, leading to increased internal energy [4].
• Solution: For analyses requiring the preservation of native-like structures or noncovalent complexes (e.g., native IM-MS), position the emitter further from the inlet. One study systematically mapped emitter position and found that "emitter positions closer to the instrument inlet can result in significantly greater in-source activation," shifting the CID50 potential for the loss of heme from holomyoglobin by as much as 8 V on a Waters Synapt G2-Si instrument [4]. Reproducibility is enhanced by consistently reporting the emitter's coordinates relative to the inlet.

FAQ: My electrospray is unstable, especially with high-conductivity buffers. What can I do?

Instability often arises from an inability to maintain a stable Taylor cone, particularly with solvents containing high concentrations of salts or buffers, which increase solution conductivity [28] [29].

• Problem: Unstable spray, fluctuating current, and poor signal intensity with salty samples.
• Root Cause: High conductivity can lead to erratic discharging and difficulty in forming a stable Taylor cone [29].
• Solution: Implement electrokinetic desalting techniques directly within the nESI emitter.
  • Polarity Reversing nESI (PR-nESI): Prior to ionization, apply a high voltage of opposite polarity for a short duration (<10 s). For positive ionization, a negative high voltage (-3 to -5 kV) drives small, highly mobile cations (like Na⁺) away from the emitter tip. When the voltage is switched to the normal positive ionization polarity, the lower mobility analytes (peptides, proteins) are carried back to the tip for stable ionization, providing several minutes of desalted signal [28].
  • Step Voltage nESI: Apply a voltage well above the onset potential (+5.2 to 5.6 kV) for 10-60 seconds, causing discharge and allowing salts to migrate away from the tip. Then, lower the voltage to a stable electrospray level (e.g., +2.4 kV). This method has been shown to improve the signal-to-noise ratio for angiotensin II in tris buffer by 50-fold [28].

FAQ: Where should I apply the high voltage for nanoESI, and why does it matter?

The point of voltage application in a nanoESI setup using nonconducting capillaries (e.g., fused silica) significantly impacts the required voltage, spray stability, and the extent of electrochemical reactions [30].

• Problem: Difficulty finding the optimal spray voltage, or observing oxidized analyte species.
• Root Cause & Solution:
  • Application at a Metal Union (close to emitter): This is a common setup. However, it can induce electrochemical reactions at the metal-solution interface, potentially leading to analyte oxidation [30].
  • Application at the Sample Reservoir (far from emitter): This method requires the application of a higher voltage to achieve a stable spray but significantly reduces the appearance of oxidized analyte peaks. It provides a "cleaner" electrochemical environment [30].
• Optimization Tip: The optimum voltage is also dependent on sample conductivity. Lower potentials must be used with highly conductive electrolytes. Therefore, performing a DC voltage scan is necessary to determine the optimum potential for your specific sample and setup [30].

Sensitivity is limited by the efficiency of both ion production (ionization) and ion transmission into the mass spectrometer [10].

• Problem: Low signal intensity for target analytes.
• Root Cause: In standard atmospheric pressure ESI, the electrospray plume is often larger than the MS inlet orifice, so only a fraction of the generated ions are sampled [10].
• Solution:
  • Utilize Multi-Emitter Arrays: Using an array of emitters, rather than a single emitter, increases the total ion current generated. One study demonstrated that sensitivity increased with the number of emitters in the array, with a over an order of magnitude improvement observed using a multi-emitter configuration compared to a standard single emitter [10].
  • Operate at Subambient Pressures (SPIN source): Placing the ESI emitter in the first reduced-pressure region (10-30 Torr) of the mass spectrometer eliminates the inlet orifice bottleneck. This allows the entire spray plume to be sampled by the subsequent ion funnel. When combined with a multi-emitter array, this configuration provides the highest reported sensitivity [10].

Table 1: Quantitative Effects of Critical Parameter Adjustments

Parameter	Condition/Variable	Quantitative Outcome	Experimental Context
Emitter Position [4]	Close vs. Far from inlet	Shift in CID50 by up to 8 V	Holomyoglobin on Waters Synapt G2-Si
Electrokinetic Desalting [28]	PR-nESI or Step Voltage	50-fold S/N improvement for Angiotensin II	Peptide in tris HCl buffer
Emitter Number [10]	Multi (7) vs. Single Emitter	>10x MS sensitivity increase	9-peptide mixture, SPIN source
Ion Utilization [10]	Single Emitter SPIN source	Up to 50% efficiency	~50 nL/min flow rate

Experimental Protocols

Protocol: Systematic Optimization of ESI Source Parameters Using Design of Experiments (DOE)

This protocol provides a framework for multi-parameter optimization to preserve protein-ligand complexes, based on a study of Plasmodium vivax guanylate kinase (PvGK) with its ligands [31].

• Define Response Variable: Calculate the relative ion abundance of the protein-ligand complex to free protein (PL/P) from the mass spectra. Sum the intensities of all charge states for each species [31].
• Select Critical Parameters: Choose ESI source parameters for optimization. Common factors include [31]:
  • Capillary voltage
  • Nebulizer gas pressure
  • Drying gas flow rate and temperature
  • Skimmer voltages
• Implement Experimental Design: Use an inscribed central composite design (CCI). This statistical design efficiently explores the parameter space by studying each factor at five levels, allowing for the estimation of linear, interaction, and quadratic effects [31].
• Data Analysis and Optimization: Employ Response Surface Methodology (RSM) to analyze the results. The "rsm" package in R software can be used to model the relationship between the parameters and the PL/P response, and to predict the optimal factor settings that maximize the complex's relative abundance [31].
• Validation: Validate the predicted optimal conditions by performing a binding titration or competition experiment to determine the equilibrium dissociation constant (K_D) [31].

Protocol: Fabrication of Multi-Emitter Arrays with Individualized Sheath Gas

This protocol details the creation of emitter arrays for enhanced sensitivity in subambient pressure ionization [10].

• Capillary Assembly:
  • Obtain fused silica capillaries: larger ones for sheath gas (e.g., 360 μm o.d., 200 μm i.d.) and smaller ones for emitters (e.g., 150 μm o.d., 10 μm i.d.) [10].
  • Construct a sheath gas capillary preform by inserting the larger capillaries through a PEEK sleeve. Use a PEEK disk spacer with precisely sized holes to arrange the capillaries into a circular array (e.g., 4, 6, or 10 emitters) [10].
  • Secure the assembly by applying epoxy at both the interior and distal ends behind the spacer. Once cured, cut the interior end flat [10].
• Integration and Sealing:
  • Insert the preform into a T-junction and secure it. Thread the emitter capillaries through the preform so they protrude 1-2 cm. Seal the emitters in place with epoxy at the second seal and cut off the residual ends [10].
• Emitter Tapering:
  • Remove the polyimide coating from the emitter tips using a solution like Nanostrip at 100°C [10].
  • Chemically etch the emitters in hydrofluoric acid (HF) to form external tapers. To prevent etching of the inner wall, pump water through the emitter array at a low flow rate (e.g., 100 nL/min per emitter) during the etching process [10].

Workflow Visualization

Optimization Workflow: A troubleshooting flowchart for nESI parameter optimization.

The Scientist's Toolkit

Table 2: Essential Research Reagent Solutions

Item	Function / Rationale
Ammonium Acetate Buffer (Volatile)	A volatile buffer salt (e.g., 10-200 mM, pH 6.8-7.4) used in "native" MS to maintain protein structure and noncovalent interactions without leaving solid residues [4] [31].
Formic Acid / Acetonitrile with 0.1% FA	Common mobile phase additives for positive-ion mode LC-ESI-MS. Formic acid promotes protonation; acetonitrile aids solubility and desolvation [10].
Pulled Borosilicate Glass nESI Emitters	The standard emitter for nanoESI. Tips are pulled to a small internal diameter (e.g., ~2 μm) to enable stable low-flow-rate electrospray, which improves ionization efficiency [4].
Triethylammonium Formate (TEAF)	A volatile salt used in mobility spectrometry and as a conductivity standard for characterizing electrospray behavior in different setups [29].
Tetraalkylammonium Salts (e.g., C7, C16)	Sparingly soluble mobility standards used to study transport phenomena and peak broadening in ionization sources like paper spray and nanoESI [29].

What is the "Low Proton Affinity Anion" strategy? In native electrospray ionization mass spectrometry (nESI-MS), the presence of non-volatile salts like sodium chloride is a major cause of ion suppression and peak broadening. This occurs as salts condense onto analyte ions, distributing signal across many adducted species and drastically reducing sensitivity [12]. A powerful method to counteract this is the use of solution additives containing anions with relatively low proton affinity (PA).

The core principle is that anions with low PA (e.g., bromide, iodide) are less likely to deprotonate acidic sites on the protein. This reduces the formation of strong binding sites for sodium cations, thereby minimizing nonspecific Na+ adduction. Instead, these anions can facilitate the removal of sodium ions from the electrospray droplet, leading to a significant increase in the signal-to-noise ratio (S/N) of the desired protein ions [32].

Key Research & Quantitative Data

Efficacy of Low Proton Affinity Additives

The table below summarizes data from key experiments demonstrating the effectiveness of various additives in reducing sodium ion adduction.

Table 1: Effectiveness of Additives in Mitigating Sodium Adduction

Protein Analyte	Salt Challenge	Additive (Concentration)	Key Improvement	Source
Ubiquitin (8.6 kDa)	1.0 mM NaCl	25 mM Ammonium Bromide	72-fold increase in abundance of fully protonated ions [32]
Ubiquitin (8.6 kDa)	1.0 mM NaCl	25 mM Ammonium Iodide	56-fold increase in abundance of fully protonated ions [32]
Bovine Serum Albumin (66 kDa)	Not Specified	10 mM L-Serine	~4-fold increase in S/N; ~10-fold peak narrowing [33]
Various Proteins & Complexes (14 - 466 kDa)	Biological buffers at physiologically relevant concentrations	Ammonium Acetate with Bromide/Iodide (via theta emitters)	Significant increase in S/N, method reproducibility, and robustness [12]

The Proton Affinity Scale of Anions

The effectiveness of an anion is inversely related to its proton affinity. The following table lists common anions in order of increasing proton affinity, illustrating why bromide and iodide are particularly effective.

Table 2: Proton Affinity of Selected Anions and Their Utility as Additives

Anion	Proton Affinity (kcal·mol⁻¹)	Effectiveness as Additive	Rationale
Perchlorate (ClO₄⁻)	~306	Not Commonly Reported	Very low PA minimizes both deprotonation and Na+ adduction.
Iodide (I⁻)	314	High	Low PA prevents creation of strong Na+ binding sites on the protein [12] [32].
Bromide (Br⁻)	325	High	Intermediate PA effectively balances Na+ removal and minimizes anion adduction [12].
Chloride (Cl⁻)	333	Moderate
Acetate (CH₃COO⁻)	348	Lower (Baseline)	High PA favors deprotonation of protein acidic sites, creating strong Na+ binding sites and worsening adduction [12].

Experimental Protocols

This is a standard method for analyzing purified protein samples [32].

• Sample Preparation:

  • Prepare your protein sample in a volatile buffer, typically 100-200 mM ammonium acetate, at the desired pH.
  • Introduce the salt challenge (e.g., NaCl) if modeling a specific matrix.
  • Add the low PA additive (e.g., Ammonium Bromide or Ammonium Iodide) directly to the sample solution from a concentrated stock. A final concentration of 25 mM has been shown to be effective [32].

• nESI-MS Analysis:

  • Load the solution into a nESI emitter (pulled borosilicate glass capillary with a tip i.d. of ~1-2 μm).
  • Initiate the spray by applying a voltage of 0.8 - 1.2 kV to a platinum wire inserted into the solution.
  • Position the emitter ~1-2 mm from the mass spectrometer inlet [12].
  • Optimize source conditions and apply collisional activation in the interface region to remove residual solvent and weakly bound adducts.

Protocol: Theta Emitter Strategy for Complex Buffers

This advanced method is suitable for analyzing proteins directly from physiologically relevant buffers without prior desalting [12].

• Emitter and Setup:

  • Use a theta emitter, a specialized glass capillary with an internal septum dividing it into two channels.
  • Insert a dual platinum wire holder, with each wire making contact with the solution in one channel.

• Solution Loading:

  • Channel 1: Load the protein sample dissolved in its native biological buffer (e.g., PBS, Tris) containing non-volatile salts.
  • Channel 2: Load a solution of 200 mM ammonium acetate containing the low PA additive (e.g., bromide or iodide).

• nESI-MS Analysis:

  • Apply a voltage of 0.8 - 2.0 kV to the platinum wires to generate a mixed electrospray.
  • Theta emitter use posits that incomplete mixing of the two streams promotes formation of droplets depleted of non-volatile salts, enabling analysis [12].
  • Employ gas-phase collisional activation methods (e.g., beam-type CID) to aid in the final removal of salt adducts.

Theta Emitter Experimental Workflow

Troubleshooting Guide

Problem: Inefficient Adduct Removal

• Cause: Additive concentration may be too low to compete effectively with native salts.
• Solution: Titrate the additive concentration. Increase the concentration of ammonium bromide/iodide up to 25-50 mM while monitoring S/N [32].
• Cause: Insufficient gas-phase activation.
• Solution: Systematically optimize collisional energies in the source interface and collision cell. Beam-type CID is often the major contributor to adduct removal [12].

Problem: Signal Suppression or Unstable Spray

• Cause: The overall ionic strength of the solution is too high.
• Solution: Ensure the use of nano-flow rates and appropriately sized emitters (∼1-2 μm i.d.). Theta emitters are specifically designed to handle higher ionic strength solutions by creating a localized mixing zone [12].
• Cause: Incorrect emitter positioning.
• Solution: Reposition the nESI emitter. Emitters placed closer to the instrument inlet can cause greater in-source activation, potentially leading to dissociation. Positions farther away can improve desalting but may reduce signal intensity [4].

Problem: Formation of New Adducts or Side Reactions

• Cause: The low PA anion is adducting to the protein as a neutral acid molecule (e.g., HBr, HI).
• Solution: These acid adducts are typically weakly bound and can be readily removed with mild collisional activation in the mass spectrometer's interface region without fragmenting the protein [32].
• Cause: Redox reactions or electrochemical degradation.
• Solution: Try using a lower electrospray voltage to minimize unwanted electrochemical side reactions at the emitter [11].

FAQs

Q1: Can I use this strategy for all proteins and protein complexes? The strategy has been successfully demonstrated for a wide range of systems, from small proteins like ubiquitin (8.6 kDa) to large complexes like alcohol dehydrogenase (148 kDa) [32] [33]. However, the optimal additive and its concentration should be determined empirically for each new system, as the surface composition and number of charge sites can influence effectiveness.

Q2: Are there any risks of protein denaturation when using bromide or iodide? The use of bromide and iodide additives at millimolar concentrations (e.g., 25 mM) is generally considered a "soft" technique that preserves non-covalent protein-protein interactions and native-like structures [32]. However, some anions like perchlorate are known to be denaturing and should be used with caution if native structure is a priority.

Q3: How do low PA anions compare to other desalting methods like buffer exchange? Buffer exchange is a pre-ESI solution, while low PA additives work during the ESI process itself. The additive strategy is simple, requires no extra preparation steps or sample loss, and can be applied to samples where complete salt removal would disrupt complex integrity [12] [32]. It is often used as a complementary technique to, not a replacement for, good sample preparation.

Q4: I work with biological tissue extracts. Will this work for me? Yes. The theta emitter approach with low PA additives was developed for this specific challenge. It allows for the mass analysis of protein complexes extracted from biological tissues where the starting material is limited and the sample contains biological buffers and non-volatile salts at physiologically relevant concentrations [12].

The Scientist's Toolkit

Table 3: Essential Research Reagents & Materials

Item	Function / Rationale
Ammonium Bromide (NH₄Br)	A source of Br⁻ anions (PA = 325 kcal·mol⁻¹). Effectively reduces Na+ adduction with minimal protein denaturation [32].
Ammonium Iodide (NH₄I)	A source of I⁻ anions (PA = 314 kcal·mol⁻¹). Slightly more effective than bromide due to its lower PA [32].
Theta Emitters	Dual-channel glass emitters enabling on-line mixing of sample and additive solutions, crucial for analyzing samples in non-volatile buffers [12].
Ammonium Acetate (NH₄OAc)	The standard volatile buffer for native MS. Serves as the base for creating additive solutions in the theta emitter channel [12] [33].
Pulled Borosilicate Capillaries	Standard nESI emitters (1-2 μm i.d.) for introducing sample-additive mixtures. Smaller diameters produce smaller initial droplets, reducing the number of adducts per droplet [32].

Technical Troubleshooting Guides

FAQ 1: Why is my protein spectrum broad and noisy with poor resolution, and how can I fix it?

Problem: The mass spectrum of your protein or protein complex shows broad, unresolved peaks with a high baseline, making mass determination difficult or impossible.

Diagnosis: This is a classic symptom of salt adduction. Non-volatile salts (e.g., NaCl) condense onto your analyte during the final stages of the electrospray process via the Charged Residue Mechanism (CRM) [34] [35]. Each molecule ends up with a different number of salt adducts (e.g., Na+, K+), distributing the signal over multiple mass-to-charge (m/z) peaks and broadening the spectral features [36] [37].

Solution: Apply controlled in-source collisional activation.

• Action: Systematically increase the declustering potential (DP) or orifice voltage [11]. This parameter accelerates ions into the interface region containing a bath gas (e.g., N2). The collisions transfer energy, heating the ions and facilitating the removal of adducted salt molecules and solvent molecules [12] [35].
• Optimization Tip: Start with a low voltage and increase incrementally. The goal is to find a "sweet spot" where adducts are removed without causing dissociation of non-covalent protein complexes or inducing unwanted in-source fragmentation [11]. Excessive voltage will lead to protein unfolding or dissociation.

FAQ 2: My analyte signal is very weak or completely suppressed, even though it's present in the solution. What is happening?

Problem: The signal for the biological ion of interest is absent or severely suppressed, often accompanied by intense signals for salt clusters.

Diagnosis: This is ion suppression due to high salt concentrations. Salts can remove excess charge from ESI droplets, suppressing the liberation of analyte ions into the gas phase [12] [37]. In extreme cases, the formation of salt clusters outcompetes the formation of analyte ions [35].

Solution: A multi-pronged approach is needed.

• Source Conditions: Ensure your source desolvation gas temperature and flow rates are optimized to aid in droplet fission and solvent evaporation, which can help release analyte ions [11].
• Sample Introduction: Consider using submicron or theta emitters. These smaller-diameter emitters produce smaller initial droplets, which contain fewer non-volatile salts and are more efficient at producing "clean" bio-ions [12] [35].
• Solution Additives: Introduce anions with low proton affinity (e.g., bromide, iodide) to the spray solution. These anions can help displace non-volatile salts like NaCl, reducing both adduction and suppression [12].

Understanding the Mechanisms: Declustering and Activation

The process of in-source cleanup relies on imparting controlled internal energy to ions to disrupt weak non-covalent bonds between the analyte and salt adducts without fracturing the analyte itself.

Workflow for In-Source Desalting via Collisional Activation

The following diagram illustrates the sequential stages of ion desolvation and declustering as ions travel from the atmospheric pressure source into the high-vacuum mass analyzer.

Key Processes in the Workflow:

• Ion Formation and Desolvation: Charged droplets containing the protein and salts undergo desolvation in the source region through heated gas and evaporative cooling in the vacuum interface [38] [35].
• Collisional Activation: A declustering potential (a voltage gradient between the orifice and the skimmer) accelerates the ions. Collisions with neutral gas molecules convert this kinetic energy into internal energy (vibrational excitation) [11] [39].
• Adduct Removal: The internal energy causes the weakly bound salt ions and solvent molecules to be vibrationally stripped from the protein analyte, resulting in a "cleaner" ion beam entering the mass analyzer [12] [37].

Quantitative Guide to Declustering Potentials and Gas Flow Parameters

The table below summarizes key parameters that can be optimized to combat salt adduction, based on experimental data.

Table 1: Optimization Parameters for In-Source Cleanup

Parameter	Typical Function	Role in Combating Salt Adduction	Experimental Range & Examples
Declustering / Cone Voltage	Extracts ions; induces declustering	Primary method for removing salt and solvent adducts via Collision-Induced Dissociation (CID) [12] [11].	Typically 10-60 V for general LC-ESI-MS [11]. Must be optimized for each analyte to avoid dissociation [35].
Collision Gas Pressure	Provides target atoms for CID	Higher pressure (~6-10 mTorr) increases collision frequency, improving desolvation and adduct removal efficiency [12].	Optimized in collision cell (q2) with N₂ gas [12].
Source / Desolvation Temperature	Aids droplet evaporation	Higher temperatures help evaporate solvent from droplets, a prerequisite for effective salt adduct removal [11].	Commonly set to ~100-200°C [11] [39].
Nebulizing / Desolvation Gas Flow	Assists droplet formation and desolvation	Restricts initial droplet size and helps with solvent stripping, reducing the number of salts per droplet [11].	Flow rates must be optimized for specific source designs [11].

Detailed Experimental Protocols

Protocol: Beam-Type Collision-Induced Dissociation (BTCID) for Salt Removal

This protocol is adapted from methods used to analyze proteins and protein complexes directly from solutions with high salt concentrations [12].

Objective: To remove sodium chloride adducts from a protein ion beam using collisional activation in a linear quadrupole collision cell.

Materials:

• Mass spectrometer equipped with a linear ion trap or collision cell (e.g., modified Q-TOF system).
• Protein sample in a volatile buffer (e.g., ammonium acetate) or a biological buffer with salts.
• Nitrogen collision gas.

Method:

• Introduction: Introduce the protein sample via nano-electrospray ionization (nESI). Theta emitters can be used to mix the sample with a volatile salt solution online, promoting the formation of a small population of droplets depleted of non-volatile salts [12].
• Activation:
  • Ions are accelerated into the quadrupole collision cell (q2) pressurized with N₂ bath gas to 6-10 mTorr.
  • The acceleration voltage (BTCID energy) is optimized to provide sufficient collisional energy to disrupt protein-salt interactions without causing protein unfolding or dissociation.
  • This step is the major contributor to salt adduct removal [12].
• Additional RF-Heating (Optional): For maximal removal of weakly-bound adducts, accumulated ions can be displaced from the center of the ion trap using a Dipolar Direct Current (DDC) offset. This shifts ions into regions of higher radiofrequency field, increasing their kinetic energy and subsequent collision energy with the bath gas [12].
• Analysis: Subject the "cleaned" ions to mass analysis (e.g., time-of-flight).

Troubleshooting:

• Insufficient Cleaning: If adducts remain, incrementally increase the BTCID energy or the DDC offset.
• Protein Dissociation: If the protein complex dissociates or the charge state distribution shifts to higher charges (indicating unfolding), reduce the applied activation energy [12] [36].

The Scientist's Toolkit: Research Reagent Solutions

This table catalogs key reagents and materials cited in research for mitigating salt adduction.

Table 2: Essential Research Reagents and Materials for Salt Adduction Mitigation

Item	Function & Rationale	Example Use & Context
Ammonium Acetate (Volatile Buffer)	The standard volatile buffer for native MS. It preserves non-covalent interactions and, unlike NaCl, evaporates readily, minimizing adduct formation [35].	Used for buffer exchange via dialysis or centrifugal filters to replace non-volatile biological buffers [35].
Anions of Low Proton Affinity (e.g., Br⁻, I⁻, NO₃⁻)	Solution additives that compete with adduction. Their low proton affinity promotes the removal of sodium ions from the analyte rather than protons, reducing Na⁺ adduction [34] [12].	Added to the spray solution (e.g., 199 mM AmAc with additive) to reduce ionization suppression and chemical noise from salts like NaCl [12].
Submicron/Theta Emitters	Nano-ESI emitters with very small internal diameters (< 1 µm). They produce smaller initial ESI droplets, which contain fewer non-volatile species, leading to fewer adducts in the final gas-phase ions [12] [35].	Used for direct analysis of proteins from physiologically relevant salt concentrations where conventional emitters fail [12].
CsI or NaI Cluster Ions	Mass calibrants for high m/z range. Salt clusters like [Csₙ(I)ₙ₋₁]⁺ or [Naₙ(I)ₙ₋₁]⁺ provide evenly spaced peaks for accurate mass calibration in the high mass range typical for native MS [40] [39].	A calibrant solution is infused, and the observed cluster ions are used to generate a calibration curve for the mass spectrometer [40].

Troubleshooting Guides

Guide 1: Troubleshooting Sample Loss from Surface Adsorption

Problem Area	Specific Symptom	Likely Cause	Solution	Key Experimental Parameters to Check
Sample Preparation Surfaces	Low signal for lipid vesicles; inconsistent MS results.	Non-specific binding to container walls; vesicle aggregation [41].	Use low-adsorption plasticware; implement a mechanical lysis method (e.g., Surface Acoustic Waves) to replace chemical methods that require transfer [41].	SAW device frequency (9-50 MHz), input RF power, LiNbO3 substrate material [41].
Nano-ESI Emitter	Signal decay over time; high salt adduction; variable in-source activation.	Sample adsorption to emitter inner wall; unstable spray due to clogging or positioning [4].	Use pulled borosilicate emitters with ~2 µm i.d. [4]; optimize emitter position relative to MS inlet; ensure emitter is clean and free of debris.	Emitter tip position (x,y,z coordinates relative to inlet); emitter inner diameter (confirm with SEM) [4].
Sample Transfer	Significant volume loss between preparation and analysis.	Dead volume in transfer lines; adsorption to surfaces of syringes or tubing [42].	Use an integrated platform that combines lysis and ionization in a single device [43]; use narrow-bore, surface-passivated transfer tubing.	Length and internal diameter of transfer lines; use of integrated SCEL-nS platforms [43].

Guide 2: Troubleshooting Container and Surface Selection

Problem Area	Specific Symptom	Likely Cause	Solution	Key Experimental Parameters to Check
Container Material	Unrecoverable sample from stock solutions; unexpected contaminants in spectrum.	Sample adsorption to container polymer; leachates from plastic [42].	Use glass vials when possible; for plastics, use certified low-binding polymers; use integrated sampling/ionization to avoid containers altogether [42].	Container material composition (e.g., polypropylene vs. glass); use of single-cell nano-ESI with live cell sampling [42].
Surface Chemistry	Inefficient nebulization or lysis on a SAW device.	Incorrect surface energy of the SAW substrate, leading to poor droplet control [41].	Apply appropriate surface treatments (e.g., hydrophobic/hydrophilic coatings) to the LiNbO3 substrate to direct jetting dynamics [41].	Surface contact angle; SAW wavelength (80-414 µm) and resultant nebulization efficiency [41].
Ionization Source Environment	Matrix effects causing ion suppression or enhancement; poor reproducibility.	Interferents in the sample (salts, lipids) affecting ionization efficiency in the ESI process [44].	Consider techniques like electron ionization (EI-MS) which are less susceptible to matrix effects [44]; use extensive sample purification/desalting.	Mobile phase composition; source temperature; use of nanoLC-EI-MS for matrix-effect-free analysis [44].

Frequently Asked Questions (FAQs)

Q1: How does the physical position of my nano-ESI emitter affect my sample and data? The position of your nano-ESI emitter is a critical, yet often overlooked, parameter. Bringing the emitter closer to the instrument inlet can lead to significant in-source activation, unintentionally unfolding or dissociating your ions before analysis. This can shift collision-induced dissociation midpoints (CID50) by as much as 8 V [4]. For the most reproducible native MS and CIU data, consistently maintain the emitter at a standardized, optimal position that balances signal intensity with minimal activation [4].

Q2: Are there alternatives to traditional containers to completely avoid sample loss? Yes, innovative "container-less" sample preparation and introduction methods are being developed. For instance, vacuum Matrix-Assisted Ionization (vMAI) allows a matrix:analyte sample to be placed on a metal probe or plate and introduced directly to the mass spectrometer's vacuum, spontaneously generating ions without a traditional solvent container or electrospray emitter [45]. This can drastically reduce surface interactions and sample loss.

Q3: What is the most effective way to prevent sample loss when analyzing delicate lipid vesicles? A highly effective strategy is to minimize sample transfer steps by using an integrated platform. Research shows that a single-chip surface acoustic wave (SAW) device can perform simultaneous mechanical disruption and nebulization of lipid vesicles, directly feeding them into the mass spectrometer [41]. This approach eliminates the transfer and associated volume loss between separate lysis and ionization devices, preserving sample and improving sensitivity [41].

Q4: How can I improve the sensitivity of my nano-ESI setup for limited samples? Beyond container selection, fundamental changes to the ionization source geometry can yield dramatic gains. The Subambient Pressure Ionization with Nanoelectrospray (SPIN) source places the emitter in the first low-pressure region of the mass spectrometer (~30 Torr). This configuration allows the entire electrospray plume to be captured by the ion funnel, eliminating losses at the atmospheric pressure inlet. Coupling this with a multi-emitter array has been shown to improve MS sensitivity by over an order of magnitude compared to standard atmospheric pressure ESI [10].

Table 1. Performance Characteristics of Sample Loss Mitigation Technologies

Technology / Parameter	Key Metric	Performance / Value	Relevance to Sample Loss
Single-Chip SAW Device [41]	Frequency Range	9.24 - 49.89 MHz	Higher frequencies enhance liposome disruption in a single device, preventing transfer loss.
	Substrate Material	128° YX-cut LiNbO3	Chosen for high coupling efficiency and wave stability, ensuring consistent sample processing.
SPIN Source & Emitter Array [10]	Sensitivity Gain	>10x improvement	Multi-emitter array in low-pressure environment drastically improves ion utilization.
	Ion Utilization Efficiency	Up to 50%	Achieved at low nL/min flow rates, meaning 1 in 2 analyte molecules is converted to a detectable ion.
nano-ESI Emitter Positioning [4]	CID50 Shift	Up to 8 V	Emitter too close to inlet causes pre-analysis activation, a form of "information loss."
	Recommended i.d.	~2 µm	Standardized emitter dimensions ensure reproducible spray and minimize clogging.

Experimental Protocols

Protocol 1: Integrated Single-Cell Electrical Lysis and Nanoelectrospray (SCEL-nS)

This protocol details the use of an integrated platform for live single-cell mass spectrometry, which bypasses multiple sample containers to minimize loss [43].

• Platform Setup: Couple the integrated SCEL-nS platform to a high-resolution mass spectrometer (e.g., Orbitrap MS).
• Cell Introduction: Introduce a suspension of live, individual cells to the platform.
• Electrical Lysis: Apply an electrical field within the integrated probe to rapidly lyse a single cell. This occurs within the probe itself, eliminating the need for a separate collection tube.
• irect Ionization: The lysate is immediately mixed with solvent at a T-junction and delivered to the nano-ESI emitter for ionization [42].
• Data Acquisition: Commence MS data acquisition. The entire process from lysis to ionization is dilution-free and occurs within a unified, miniaturized system [43].

Protocol 2: Surface Acoustic Wave (SAW) Nebulization for Direct Vesicle Analysis

This protocol describes using a SAW device to simultaneously disrupt and nebulize lipid vesicles for direct MS analysis, preventing loss from transfer between steps [41].

• Device Fabrication:
  • Start with a cleaned 1 mm thick, 128° YX-cut LiNbO3 wafer.
  • Spin-coat with a negative photoresist (e.g., NR9-1500py) and pattern via UV lithography to define interdigital transducers (IDTs).
  • Deposit 10/100 nm Cr/Au layer via e-beam evaporation and perform a lift-off in acetone to complete the electrode structure [41].
• Surface Treatment: Apply an appropriate surface treatment (hydrophobic/hydrophilic) to the LiNbO3 substrate in the sample region to control droplet jetting dynamics.
• Sample Preparation: Prepare a solution of DOPC liposomes or other extracellular vesicle models in a compatible aqueous buffer.
• Device Operation:
  • Place a small droplet (e.g., 2-10 µL) of the liposome solution onto the sample region between the IDTs.
  • Apply a radio frequency (RF) signal at the device's resonant frequency (e.g., 9-50 MHz) and optimal power to initiate simultaneous disruption and nebulization.
• MS Analysis: Direct the generated aerosol into the mass spectrometer inlet. For enhanced ionization of non-polar lipids, couple with corona discharge ionization [41].

Workflow Visualization

The Scientist's Toolkit: Research Reagent Solutions

Table 2. Essential Materials for Preventing Sample Loss in nano-ESI MS

Item	Function / Application	Technical Specification
Pulled Borosilicate Glass Emitters	Nano-electrospray ionization for minimal flow rates and high sensitivity [4] [42].	~2 µm inner diameter at tip; 1.0/0.78 mm o.d./i.d. [4].
Lithium Niobate (LiNbO3) SAW Substrate	Piezoelectric substrate for integrated mechanical lysis and nebulization; reduces need for chemical lysis and transfers [41].	128° YX-cut; 1 mm thickness; high electromechanical coupling coefficient [41].
Low-Binding Micro-Tubes	Storage and handling of precious samples to minimize adsorption to container walls.	Certified polymer (e.g., PCR-clean, non-stick) for protein/lipid samples.
Surface Treatment Reagents	Modifying surface energy of substrates (e.g., SAW chips) to control fluidic behavior and improve process efficiency [41].	Hydrophobic/hydrophilic coatings (e.g., silanes, fluorinated coatings).
Volatile MS-Compatible Buffers	Creating a native-like environment for biomolecules without causing ion suppression or source fouling [4] [44].	200 mM Ammonium Acetate, pH 7.4 [4].
Nanospray Ion Source	Enabling high-sensitivity analysis at low flow rates, providing longer analysis time and improved detection [42].	Capable of stable ionization at 25 nL/min flow rates [42].

In the pursuit of high-sensitivity analyses within nanoelectrospray mass spectrometry (nESI-MS), spray stability is a foundational prerequisite. It directly influences data quality, reproducibility, and the reliable detection of trace-level analytes in applications ranging from proteomics to single-cell metabolomics [2] [43]. Despite its critical importance, researchers frequently encounter technical instabilities—such as clogging, bubble formation, and wetting issues—that can compromise sensitivity and halt experimental progress. This technical support center article is designed to diagnose these common problems, provide evidence-based troubleshooting guidelines, and present advanced methodologies to enhance the robustness of your nESI-MS workflows, thereby unlocking the full potential of your sensitive analyses.

Troubleshooting Guides & FAQs

Frequently Asked Questions

Q1: What are the most common sources of instability in a nano-electrospray? The most prevalent issues affecting spray stability can be categorized into three areas:

• Clogging: Caused by the accumulation of nonvolatile salts or residues at the very narrow emitter tip [2] [46].
• Bubbles: Formed as the liquid flows through the system; dissolved gases can precipitate, expanding and distorting the flow rate, disrupting the meniscus geometry, and breaking electrical conductivity [2].
• Wetting Instabilities: These include lateral instabilities (where the meniscus sprays sideways) and longitudinal instabilities (where the meniscus size changes unpredictably), often related to emitter geometry and surface chemistry [2].

Q2: How does emitter position affect my MS data, particularly in native MS or CIU/CID experiments? The position of the nESI emitter relative to the instrument inlet is a critical but often overlooked parameter. Recent studies demonstrate that even small variations in emitter position can induce significant in-source activation. On some instrument platforms, positioning the emitter closer to the inlet can shift the mid-point potential (CID~50~ or CIU~50~) for collision-induced dissociation or unfolding by as much as 8 V. This unintended activation can lead to premature unfolding or dissociation of fragile complexes, compromising data interpretation and reproducibility in native MS experiments [4].

Q3: My emitter keeps clogging when I'm analyzing biological samples with nonvolatile buffers. Is there a solution? Yes. A method using induced alternative voltage has been developed specifically for this challenge. By applying an alternating voltage to the emitter, a re-dissolution effect on salt crystals at the tip is induced. This approach has been shown to extend emitter lifetime by 1–2 orders of magnitude compared to conventional nESI when analyzing high-concentration salt solutions that mimic extracellular or intracellular fluid [46].

Q4: Are there any additives that can actually improve spray stability and signal? Emerging research indicates that nanobubbles (NBs) can serve as a beneficial additive. Introducing CO~2~ or N~2~ nanobubbles into the spray solvent has been shown to significantly improve signal responses for both small molecules and proteins. The proposed mechanism involves the nanobubbles increasing the total area of the hydrophobic gas-liquid interface, which can improve analyte transport to the droplet surface. For proteins, this can result in increased signal intensities and higher charge states [47].

Troubleshooting Common Instabilities

The following table summarizes the core issues, their root causes, and practical solutions.

Table 1: Troubleshooting Guide for Common Nano-Electrospray Instabilities

Problem	Root Causes	Solutions & Mitigation Strategies
Clogging [2] [46]	- Accumulation of nonvolatile salts/residues.- Very small emitter inner diameter (ID).	- Use induced alternative voltage to re-dissolve crystals [46].- Use emitters with the largest feasible ID to prevent clogging [2].- Ensure sample is free of particulate matter.
Bubbles [2]	- Precipitation of dissolved gases in the flow path.- Expanding bubbles disrupt flow and electrical contact.	- Degas solvents thoroughly before use.- Inspect the fluidic path for nucleation sites.- Ensure tight connections to prevent gas ingress.
Wetting Instabilities [2]	- Poorly defined emitter tip geometry.- Suboptimal emitter surface chemistry.	- Use emitters with a sharp, well-defined edge for stable meniscus anchorage [2].- Employ hydrophobically coated emitters (e.g., LOTUS) to stabilize the meniscus at the inner diameter [2].
Spray Instability [2]	- Incorrect spray voltage or flow rate.- Solvent evaporation at the meniscus.	- Optimize voltage (typically 1.7–2.5 kV for proteomics) and ensure flow rate is compatible with emitter ID [2].- Position the emitter close to the MS inlet to compensate for weaker electric fields from small menisci [2].
Unintended In-Source Activation [4]	- nESI emitter positioned too close to the mass spectrometer inlet.	- Systemically map and record the emitter position (x, y, z coordinates) for critical experiments.- For native MS and CIU/CID, adopt a standardized "far" position to minimize uncontrolled collisional heating.

Experimental Protocols

Protocol: Mitigating Clogging with Induced Alternative Voltage

This protocol is adapted from methods developed to handle high concentrations of nonvolatile buffers, such as those found in biological samples [46].

1. Emitter Preparation: Use nanoemitters with an inner diameter of less than 1 µm. 2. Solution Preparation: Prepare your sample in the required high-concentration salt solution (e.g., mimicking extracellular fluid). 3. Voltage Application: Instead of a standard DC voltage, apply an induced alternative voltage to the nanoemitter. The periodic change in the electric field direction prevents the stable accumulation of salt crystals by promoting their re-dissolution. 4. Data Acquisition: Infuse the sample and begin MS data acquisition. The signal should remain stable and sensitive for extended periods (e.g., ~10 minutes), significantly longer than with conventional nESI.

Protocol: Standardizing Emitter Position for Reproducible CIU/CID Data

This protocol is crucial for ensuring reproducibility in native MS and collision-induced unfolding/dissociation experiments [4].

1. Emitter Fabrication: Pull borosilicate glass nESI emitters to ~2-micrometer i.d. openings using a Flaming-Brown micropipette puller. 2. Define "Close" Position: Carefully position the emitter tip as close to the instrument inlet as possible without physical contact. On a Waters Synapt G2-Si, this is defined as coordinates (0, 0.5, 0 mm) relative to the inlet center. 3. Define "Far" Position: Systematically retract the emitter to a predetermined distance. Precisely record the x, y, and z coordinates. Note: The "far" position is generally recommended for CIU/CID to minimize unintended in-source activation. 4. Documentation: For every critical experiment, document the exact emitter position coordinates in the experimental metadata. This ensures the same conditions can be replicated in future studies.

The Scientist's Toolkit: Research Reagent Solutions

Table 2: Essential Materials for Stable and Sensitive Nano-Electrospray

Item	Function & Rationale
Hydrophobic Coated Emitters (e.g., LOTUS) [2]	A hydrophobic coating locks the electrospray meniscus at the emitter's inner diameter, resulting in a smaller and more stable meniscus. This improves ionization efficiency, allows for lower spray voltages, and produces a more consistent spray.
Sharp Singularity Emitters [2]	Emitters that are mechanically sharpened to a precise, acute angle with a well-defined edge. This geometry provides a stable anchorage point for the meniscus, eliminating a major source of variability and improving spray consistency.
Nanobubble-Enriched Solvents [47]	Solvents infused with CO~2~ or N~2~ nanobubbles act as a novel additive. They increase the gas-liquid interface area, which can improve signal intensity and charge states for proteins, and mitigate ion suppression for small molecules.
SiO₂ Seeds [48]	While primarily used in membrane distillation, the principle of using inert seeds (30–60 µm) to control crystallization at surfaces is a promising concept for mitigating scale-related clogging in fluidic systems.
Induced Alternative Voltage Source [46]	A power supply capable of providing an alternating voltage waveform, essential for implementing the anti-clogging protocol with high-concentration salt solutions.

Workflow & Visual Guides

The following diagram illustrates a systematic decision-making workflow for diagnosing and resolving common nESI stability issues.

Validation, Performance Benchmarks, and Comparative Analysis of nESI Techniques

FAQ: Troubleshooting Guide for Nanoelectrospray MS Experiments

1. How can I quantitatively measure sensitivity improvements in my nanoESI-MS setup?

Sensitivity is quantified by the ratio of ions successfully transmitted into the mass spectrometer to the number of molecules entering the spray emitter. At nanoflow rates (50-500 nL/min), sampling efficiencies can exceed 70% in optimized systems [49]. To compare configurations, measure the signal intensity (peak height or area) for a standard analyte at a fixed concentration. For example, the Subambient Pressure Ionization with Nanoelectrospray (SPIN) source has demonstrated a 5- to 12-fold improvement in peptide signal intensity compared to standard atmospheric pressure sources [50]. When using multi-emitter arrays, sensitivity increases with the number of emitters, providing over an order of magnitude (>10x) improvement compared to a single emitter with a standard interface [10].

2. What are the best practices for reporting Signal-to-Noise (S/N) ratios and why is my S/N low in physiological buffers?

The S/N ratio should be calculated using the peak intensity of the ion of interest divided by the root-mean-square (RMS) noise in a nearby blank region of the spectrum [12]. Low S/N in physiological buffers is often caused by ionization suppression and extensive chemical noise from non-volatile salts.

A proven method to improve S/N is using theta emitters (~1.4 µm inner diameter) with an solution additive strategy. Load your protein sample in a biological buffer (e.g., PBS) into one channel, and a volatile salt solution like 199 mM ammonium acetate, spiked with anions of low proton affinity (e.g., bromide or iodide), into the other channel. This setup can significantly reduce chemical noise and increase S/N ratios and method reproducibility compared to using ammonium acetate alone [12].

3. How can I improve the reproducibility of my spectra, especially for collision-induced unfolding/dissociation (CIU/D) experiments?

Spectral reproducibility can be severely affected by seemingly minor variations in the nanoESI emitter's position relative to the MS inlet. Studies show that shifting the emitter can alter the Collision-Induced Dissociation 50 (CID50) value—the energy required to fragment 50% of precursor ions—by as much as 8 V on some commercial instruments [4].

To enhance reproducibility:

• Standardize Emitter Position: Carefully document the spatial coordinates (x, y, z) of your emitter tip relative to the MS inlet and consistently use the same position, especially for CIU/D workflows [4].
• Use Fixed-Source Geometries: Employ source designs with fixed, non-articulated positions for the sprayer. This eliminates the "user's touch" variability and has been shown to provide performance equivalent to a fully tunable source but without the need for extensive optimization [49].
• Control Emitter Dimensions: Use emitters with consistent internal diameters, as even subtle differences can change droplet formation and affect ion charge state distributions [4].

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Quantitative Performance Metrics for NanoESI-MS Improvements

Table 1: Measured Improvements from Advanced NanoESI Source Geometries

Source Configuration	Key Metric	Reported Improvement	Experimental Context
SPIN Source [50]	Sensitivity (Peptide Signal)	5- to 12-fold increase	Gradient reversed-phase LC-MS analysis of protein tryptic digests.
Multi-Emitter Array + SPIN [10]	MS Sensitivity	>10x (Order of magnitude) increase	Infusion of a 1 µM equimolar solution of 9 peptides.
Theta Emitters + Additives [12]	Signal-to-Noise (S/N)	Significant increase	Analysis of proteins from physiologically relevant salt solutions.
Fixed NanoESI Source [49]	Reproducibility	Eliminates need for re-tuning; equivalent performance	LC-MS analysis using a fixed, non-articulated sprayer position.

Table 2: Impact of Emitter Position on Spectral Reproducibility [4]

Analyte	Instrument	Observed Effect of Emitter Position
Holomyoglobin	Waters Synapt G2-Si	CID50 value for heme loss shifted by up to 8 V.
Leucine Enkephalin	Waters Synapt G2-Si	CID50 value for dissociation was significantly affected.
Protein Complexes (e.g., BSA)	Waters Synapt G2-Si	CIU50 values and fingerprint RMSD were altered.
Various Ions	Agilent 6545XT	Different, less pronounced effects were observed.

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Detailed Experimental Protocols

Protocol 1: Implementing Theta Emitters for High-Salt Samples

This protocol is designed to achieve robust protein mass analysis directly from physiologically relevant buffers [12].

• Emitter Preparation: Pull borosilicate theta capillaries (1.5 mm o.d., 1.17 mm i.d.) to a final tip inner diameter of ~1.4 µm using a programmable micropipette puller.
• Sample Loading:
  • Channel 1: Load your protein sample dissolved in its native biological buffer (e.g., PBS).
  • Channel 2: Load a solution of 199 mM ammonium acetate, supplemented with a low proton affinity anion additive like sodium bromide or sodium iodide.
• Electrical Contact: Insert dual platinum wires into the open ends of the theta emitter, ensuring each wire contacts only one solution channel.
• Mass Spectrometry:
  • Apply a spray voltage between 0.80 – 2.0 kV, incrementally increasing until a stable spray is observed.
  • Utilize gas-phase collisional activation methods (e.g., beam-type CID and RF-heating in an ion trap) to remove residual solvent and salt adducts without causing dissociation.

Protocol 2: Systematically Mapping Emitter Position for CIU/D Reproducibility

Use this method to document and standardize your emitter position for highly reproducible activation experiments [4].

• Define the "Close" Position: Carefully move the emitter tip to be as close as possible to the instrument inlet without physical contact. Define this as your coordinate system's origin (0, 0, 0), where the axes are vertical, horizontal along the inlet axis, and horizontal perpendicular to the inlet axis.
• Create a Spatial Map: Acquire CID breakdown curves or CIU fingerprints for a standard protein (e.g., holomyoglobin) at multiple defined positions around the "close" position (e.g., varying by 1-2 mm in each direction).
• Quantify the Effect: For each position, calculate the CID50 or CIU50 value. Plot these values against the spatial coordinates to create a map of in-source activation.
• Select Standard Position: Choose a position that provides a strong signal with minimal unintended in-source activation (evidenced by higher CID50/CIU50 values) and use this documented position for all subsequent comparable experiments.

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Experimental Workflow for NanoESI Method Optimization

The diagram below outlines a logical pathway for diagnosing issues and implementing solutions to improve key metrics in nanoESI-MS.

The Scientist's Toolkit: Key Research Reagent Solutions

Table 3: Essential Materials for Advanced NanoESI-MS Experiments

Item	Specification / Example	Function / Rationale
Theta Emitters [12]	Borosilicate, ~1.4 µm i.d., two channels.	Allows simultaneous introduction of sample and additive solution; promotes droplets depleted of non-volatile salts.
Low Proton Affinity Anions [12]	Bromide (Br⁻) or Iodide (I⁻) salts.	Reduces sodium adduction and chemical noise by competing for charge during droplet formation.
Volatile Buffer [12]	200 mM Ammonium Acetate.	Standard MS-compatible buffer for desalting; serves as a base for solution additives.
Fixed-Geometry NanoESI Source [49]	Non-articulated, pre-aligned source.	Eliminates variability from manual sprayer positioning, enhancing robustness and reproducibility.
Pulled NanoESI Emitters [4]	Borosilicate, ~2 µm i.d. opening.	Standard emitters for native MS; consistent dimensions are critical for reproducible CIU/D data.

Technical Troubleshooting Guide

FAQ 1: My analyte signal is weak or unstable with nanoESI. What could be the cause and how can I fix it?

Weak or unstable spray in nanoESI is often related to the emitter tip, sample preparation, or electrical contact.

• Cause 1: Emitter Tip Blockage or Damage. The very fine capillaries used in nanoESI are prone to clogging.
• Solution: Use disposable nanoESI capillaries (e.g., with a 1 µm tip pore size) to eliminate cross-contamination and carry-over effects [51]. Visually inspect the tip under a microscope if possible. Prepare samples in clean, particulate-free solvents and centrifuge if necessary.
• Cause 2: Suboptimal Ionization Solvent/Sample Composition. The solvent must facilitate both droplet formation and efficient ionization.
• Solution: Use an ionization solvent consisting of methanol:water:formic acid in a ratio of 80:20:0.3 to promote protonation and stable spray formation [51]. Ensure the sample is dissolved in a solvent compatible with the running buffer (e.g., high organic content for easy evaporation).
• Cause 3: Poor Electrical Connection. A stable spray requires a complete electrical circuit.
• Solution: For offline nanoESI capillaries, ensure the wire or conductive coating is properly making contact with the sample solution. Check for bubbles at the tip or connection point.

FAQ 2: I am seeing high background noise and unexpected peaks in my spectra. What is happening?

Unexpected peaks can often be attributed to in-source fragmentation (ISF) or system contamination.

• Cause 1: In-Source Fragmentation (ISF). High voltages in the ion source region can impart excess internal energy to ions, causing them to fragment before reaching the mass analyzer. These fragments can be misannotated as real compounds [52].
• Solution: Systematically optimize source parameters to reduce fragmentation. Key parameters to lower include the skimmer voltage and tube lens voltage [52]. For example, reducing the skimmer voltage from 50V to 5-20V can significantly reduce unintended fragmentation [52].
• Cause 2: Carry-over or Contamination.
• Solution: Use disposable nanoESI capillaries to eliminate a major source of carry-over [51]. For LC-ESI-MS systems, implement rigorous washing steps with strong solvents between runs. Run blank samples to identify the source of contamination.

FAQ 3: My chromatographic retention times are not reproducible. How can I improve this?

Poor retention time reproducibility undermines confident identification and quantification.

• Cause 1: Inconsistent Nano-LC System Performance.
• Solution: Ensure your LC system is well-maintained and calibrated. A benchmarking study found that systems like the Proxeon EASY-nLC, Waters nanoACQUITY UPLC, and Eksigent nanoLC-Ultra demonstrated excellent retention time reproducibility (0.7–0.9% RSD), outperforming others like the Eksigent nanoLC-2D (~2% RSD) [53].
• Cause 2: Ionization Source Instability.
• Solution: The ionization source itself can impact chromatography. One study found that simply changing from a traditional nanoESI source to a nebulized nanoFlow ESI source improved retention time reproducibility dramatically (from 14% RSD to 0.5% RSD) [53]. Ensure a stable spray position and optimize gas flows.

FAQ 4: How do I choose the right ESI method to accurately measure protein-ligand interactions?

The choice of ionization method can influence whether non-covalent interactions observed in solution are preserved in the gas phase.

• Recommendation: For determining dissociation constants (K_D), Electrosonic Spray Ionization (ESSI) has been shown to be the gentlest method, providing K_D values closest to those found in solution [54]. ESSI was less sensitive to instrumental parameters and showed weaker charge state dependence of K_D compared to standard ESI and chip-based nanoESI [54]. If sample amount is the limiting factor, nanoESI is a viable alternative, but be aware of potential charge state-dependent K_D variations [54].

Performance Data Comparison

The table below summarizes key performance metrics for different ESI techniques and LC systems based on experimental data.

Table 1: Quantitative Performance Comparison of ESI Techniques and LC Systems

Technique / System	Key Performance Metric	Result / Observation	Application Context
ESSI [54]	Closeness of measured KD to solution value	Best agreement (e.g., 19.4 ± 3.6 µM for HEWL/NAG3)	Protein-ligand interaction studies
nanoESI [54]	Closeness of measured KD to solution value	Shows charge state dependence; less accurate than ESSI	Protein-ligand interaction studies (when sample is limited)
Nebulized NanoFlow ESI [53]	Retention Time Reproducibility (RSD)	~0.5% RSD	Proteomics, Metabolomics
Traditional NanoESI [53]	Retention Time Reproducibility (RSD)	~1-14% RSD	Proteomics, Metabolomics
Proxeon/Waters/Eksigent Ultra LC [53]	Retention Time Reproducibility (RSD)	0.7 - 0.9% RSD	Nanoflow Chromatography
Direct nanoESI-MS/MS [51]	Linear Quantification Range	2.5 - 25,000 ng/mL (for Metronidazole)	Pharmaceutical analysis, direct sample analysis

Detailed Experimental Protocols

Protocol 1: Direct nanoESI-MS/MS for Metabolite/Pharmaceutical Analysis

This protocol allows for rapid analysis without liquid chromatography, ideal for high-throughput screening or when sample is limited [51].

• Sample Preparation: Prepare the sample solution in an ionization solvent (e.g., Methanol:Water:Formic Acid, 80:20:0.3 v/v/v). Include an internal standard if performing quantification (e.g., 1,3,6-polytyrosine) [51].
• Capillary Loading: Load a small volume (e.g., 2 µL) of the sample solution into the back of a disposable nanoESI capillary (e.g., 1 µm tip pore size) using a fine pipette tip [51].
• Instrument Setup: Mount the nanoESI capillary onto the ion source holder. Use a manual XYZ stage and camera to position the capillary tip approximately 3 mm from the mass spectrometer inlet [51].
• MS Data Acquisition: Apply a spray voltage (e.g., 1000 V). Acquire data in full MS scan mode for identification, and MS/MS mode (Collision-Induced Dissociation, CID) for structural confirmation. The signal is typically sustainable for about 30 seconds, allowing for multiple data points [51].
• Data Analysis: For quantification, use the ratio of the [M+H]⁺ peak areas of the analyte and the internal standard. For identification, compare the MS/MS fragments against a database or standard [51].

Protocol 2: Systematic Optimization of ESI Source Parameters using Design of Experiments (DoE)

A multivariate DoE approach is more efficient than one-variable-at-a-time (OVAT) for finding optimal source settings [55].

• Factor Selection: Identify the key source parameters to optimize. Common factors are Capillary Voltage, Nebulizer Gas Pressure, Drying Gas Flow Rate, and Drying Gas Temperature [55].
• Experimental Design:
  • Screening Phase: Use a Fractional Factorial Design (FFD) to identify which factors have a significant impact on your response (e.g., signal intensity of a low-abundance metabolite) [55].
  • Optimization Phase: For the significant factors (e.g., 2-3), apply a response surface design like a Central Composite Design (CCD) or Box-Behnken Design (BBD) to model the response and find the optimum [55].
• Execution and Analysis: Run the experiments as per the design table. Use statistical software (e.g., JMP, Design-Expert) to analyze the data, build a mathematical model, and generate response surface plots [55].
• Verification: Confirm the predicted optimum by running the method with the suggested parameters and verify the improvement in sensitivity or signal-to-noise ratio [55].

Experimental Workflow and Pathway Diagrams

Research Reagent Solutions

Table 2: Essential Materials for nanoESI and LC-ESI-MS Experiments

Item	Function / Application	Example / Specification
Disposable NanoESI Capillaries	Sample emitter for nanoESI; reduces carry-over and contamination.	Humanix capillaries (1 µm tip pore size) [51]
Ionization Solvent	Liquid matrix for creating charged droplets; critical for stable spray and ionization.	Methanol:Water:Formic Acid (80:20:0.3) [51]
LC-MS Grade Solvents	Mobile phase preparation; minimizes chemical noise and ion suppression.	J.T. Baker LC-MS grade Acetonitrile [55]
Volatile Additives	Modifies mobile phase pH to enhance [M+H]+ or [M-H]- ion formation.	0.1% Formic Acid, 0.06% Acetic Acid [53] [55]
Tuning/Calibration Solution	Mass accuracy calibration and instrument performance verification.	Agilent ESI-L Low Concentration Tuning Mix [55]
Collision Gas	Gas used in the collision cell (Q2) for fragmenting precursor ions (CID).	High-purity (99.999%) Argon or Nitrogen [56] [51]

nanoelectrospray Ionization (nESI) has become a cornerstone technique in modern analytical science, prized for its exceptional sensitivity and efficiency. This technical support center is designed to help researchers, scientists, and drug development professionals overcome common experimental challenges and leverage validated nESI methods to achieve superior results in two critical application areas: pharmaceutical quality control and clinical urine analysis. The guidance provided herein is framed within the overarching thesis of improving sensitivity in nanoelectrospray MS research, with a focus on practical, actionable troubleshooting and protocols.

Troubleshooting Guides for nESI Methods

nESI Signal Instability and Loss

• Problem: Signal fluctuation, complete loss of MS signal, or unstable baseline during nESI operation.
• Possible Causes & Solutions:
  • Spray Voltage Issues: Excessively high voltage can cause rim emission or corona discharge, leading to instability. In negative ion mode, high voltages make discharge more likely. Solution: Systematically reduce the sprayer voltage. The optimal range for nESI in proteomics workflows is typically 1.7 to 2.5 kV [2] [11].
  • Unstable Meniscus: The formation of the Taylor cone and jet is unstable. Solution: Ensure you are using emitters with a sharp, well-defined geometry and hydrophobic internal coating to lock the meniscus in place and stabilize its size [2].
  • Flow Rate Issues: The flow rate may be too low to sustain a stable electrospray or is fluctuating. Solution: Remember that solvent evaporation at the tip is a key factor; the theoretical minimum flow is below 1 nL/min, but evaporated flow in online systems is typically 10-20 nL/min. Ensure your system can deliver a stable flow within this range [2].
  • Clogging or Bubbles: Residues can accumulate at the emitter tip, or dissolved gases can precipitate, forming bubbles that disrupt flow. Solution: Use emitters with a sufficiently large inner diameter to prevent clogging. Ensure mobile phases are properly degassed [2].

Poor Ionization Efficiency and Ion Suppression

• Problem: Low signal intensity for the target analyte, often due to competitive ionization in the source.
• Possible Causes & Solutions:
  • Large Initial Droplets: Larger droplets require more evaporation/fission events to shed ions, increasing the opportunity for contaminants to cause ion suppression. Solution: nESI inherently produces smaller droplets (~200-500 nm) than conventional ESI, but ensuring optimal spray conditions (voltage, flow rate, emitter geometry) minimizes initial droplet size [2] [57].
  • High Salt Contamination: Salts promote the formation of metal adducts (e.g., [M+Na]⁺) and suppress the formation of protonated molecules. Solution: nESI exhibits a higher tolerance to salt contamination than conventional ESI [57]. For extreme cases, use rigorous sample preparation (SPE, liquid-liquid extraction). Use plastic vials instead of glass to avoid leaching of metal ions, and use high-purity MS-grade solvents [11].
  • Hydrophilic Analytes: Coulombic fission events preferentially transfer hydrophobic substances to the next droplet generation. Solution: The reduced number of fission events needed in nESI (due to smaller starting droplets) lessens this discrimination against hydrophilic molecules [2].
  • Suboptimal Solvent: The surface tension of the solvent affects stable Taylor cone formation. Solution: Use solvents with lower surface tension (e.g., methanol, isopropanol). For highly aqueous mobile phases, adding 1-2% v/v of isopropanol can lower surface tension and significantly improve signal response and stability [11].

nESI Method Validation for Pharmaceutical QC

• Problem: Uncertainty in how to validate nESI methods for nonclinical dose formulation analysis, as they fall under GLP regulations without specific formal guidance.
• Possible Causes & Solutions:
  • Lack of Regulatory Guidance: There is no specific guidance for dose formulation analysis method validation. Solution: Follow a framework adapted from bioanalytical guidance and industry white papers. The fundamental validation parameters include recovery, accuracy, precision, specificity, selectivity, carryover, sensitivity, and stability [58].
  • Inappropriate Acceptance Criteria: Applying standard bioanalytical QC acceptance criteria directly may not be applicable. Solution: Since dose formulation samples are not true "unknowns," the requirement for QC samples across the entire standard curve may not always be necessary. Define acceptance criteria prior to validation based on the intended use of the method [58].

Frequently Asked Questions (FAQs)

Q1: What are the key advantages of nESI over conventional ESI for sensitive applications like pharmaceutical QC?

nESI offers several critical advantages for sensitivity-driven work. Its primary benefit is extremely high ionization efficiency due to the production of very small initial droplets. This leads to improved sample utilization and reduced ion suppression from matrix components and salts, making it more robust for analyzing complex biological matrices or formulated drugs [2] [57].

Q2: How should I collect and handle urine samples to ensure accurate results in clinical urinalysis?

Proper collection and handling are paramount. First-morning urine is ideal as it is more concentrated. Use a clean-catch method to minimize contamination: patients should clean the urethral area, void a small amount into the toilet, then collect the mid-stream portion into a sterile container [59]. The sample must be analyzed within one hour of collection; if not possible, it should be refrigerated at 4°C for no longer than 24 hours. Delay causes changes in pH, dissolution of casts, and bacterial proliferation [60] [59].

Q3: What is the difference between a full, partial, and early-phase method validation in pharmaceutical QC?

• Full Validation: Required for chronic toxicity studies (>3 months). It encompasses all validation elements with multiple runs for accuracy and precision [58].
• Partial Validation: Conducted when a validated method undergoes a significant change (e.g., vehicle composition, instrument parameters). It requires a minimum of one set of accuracy and precision data [58].
• Early Phase Validation: For acute toxicity studies (≤3 months) with time/API constraints. It may be limited to a single validation run assessing core parameters like accuracy, linearity, and specificity [58].

Q4: What are common sources of false-positive or false-negative results in chemical urinalysis using dipsticks?

Many factors can interfere with dipstick results [59]:

• pH: Highly alkaline urine can cause false-positive protein results.
• Drugs/Contrast Agents: Phenazopyridine can change urine color; iodinated contrast media can elevate specific gravity.
• Preservatives: Formaldehyde can cause false-positive results for leukocyte esterase and glucose.
• Bacteria: Their presence can consume glucose, leading to a false-negative reading.

Q5: How can I optimize the sensitivity of my nESI method for a new chemical entity?

Start with the fundamental parameters [11]:

• Sprayer Voltage: Optimize between 1.7–2.5 kV, seeking the lowest voltage that provides a stable spray.
• Sprayer Position: For small, polar analytes, position the sprayer farther from the MS inlet; for larger, hydrophobic molecules, position it closer.
• Solvent Composition: Favor low-surface-tension solvents (methanol, acetonitrile) and consider a small additive (1-2% isopropanol) for aqueous mobile phases.
• Gas Flow/Temperature: Optimize desolvation gas temperature and flow rates to efficiently evaporate solvents from the tiny nESI droplets.

Validated Experimental Protocols

Protocol 1: nESI-MS Method for Nonclinical Dose Formulation Analysis

This protocol outlines a validated approach for confirming test article concentration and homogeneity in pharmaceutical formulations [58].

1. Method Development and Scope

• Define the intended use: to confirm dosage concentration, homogeneity, and stability.
• Ensure the method covers the entire anticipated dosage concentration range.
• During development, assess critical parameters: container composition, filter bias, solubility, and stability in the vehicle.

2. System Suitability Test (SST)

• Before each analytical run, perform SST to ensure instrument readiness.
• Parameters include: injection precision (retention time and peak area), theoretical plates (N), tailing factor (T), and resolution [58].

3. Stock Standard Comparison

• Prepare two separate stock solutions by weighing the test article (API) independently.
• Dilute both to the same concentration within the linear range.
• The response of the two solutions should compare within a 5% difference to demonstrate weighing accuracy [58].

4. Validation Experiments

• Accuracy and Precision: Perform at least multiple runs (for full validation) using quality control samples prepared in the vehicle. Report % relative error (accuracy) and %RSD (precision).
• Specificity: Demonstrate that the method can unequivocally quantify the analyte in the presence of vehicle components.
• Linearity and Range: Prepare a series of standard solutions across the concentration range. The standard curve should demonstrate a linear relationship with a correlation coefficient (r) of >0.99.
• Stability: Establish analyte stability in the solution and in the formulated vehicle under storage and handling conditions (e.g., benchtop, refrigerated).

Protocol 2: Comprehensive Clinical Urinalysis

This protocol details the standard three-part examination of urine [59].

1. Physical Examination

• Color and Clarity: Note the color (from pale yellow to amber) and clarity (clear or cloudy). Cloudy fresh urine suggests pus (infection) or red blood cells [60] [61].
• Specific Gravity: Measures urine concentration. Use a refractometer or reagent strip. Correct for high glucose or protein levels [60].

2. Chemical Examination (Dipstick)

• Use a reagent strip, following manufacturer instructions precisely.
• Test for: pH, protein, glucose, ketones, bilirubin, urobilinogen, blood, nitrites, and leukocyte esterase.
• Note: A positive nitrite test and positive leukocyte esterase are strong indicators of a urinary tract infection [60] [59].

3. Microscopic Examination

• Centrifuge 10 mL of well-mixed urine at 2000-3000 rpm for 3-5 minutes [60].
• Discard the supernatant and resuspend the sediment in the remaining liquid.
• Place a drop on a slide, cover with a coverslip, and examine under microscope.
• Scan under low power (100x) to identify areas of interest, then use high power (400x) to identify and quantify:
  • Cells: Red blood cells, white blood cells, epithelial cells.
  • Casts: Cylindrical structures formed in the renal tubules.
  • Crystals: Uric acid, calcium oxalate, etc.
  • Bacteria or Yeast.

Optimized nESI Parameters and Data

Table 1: Optimized nESI Source Parameters for Sensitive Detection

This table summarizes key parameters for optimizing nESI sensitivity based on the cited literature and application needs [2] [11].

Parameter	Recommended Setting	Technical Rationale & Impact on Sensitivity
Spray Voltage	1.7 - 2.5 kV	Prevents corona discharge and rim emission. Lower voltages often yield more stable signals.
Flow Rate	~10-20 nL/min (online)	Matches the evaporated flow rate at the emitter, ensuring a stable meniscus and small droplet size.
Emitter Geometry	Sharp tip, hydrophobic coating	Defines a small, stable meniscus, reducing solvent evaporation and ion evaporation for higher efficiency [2].
Solvent Additive	1-2% IPA in aqueous phases	Lowers surface tension, facilitating stable Taylor cone formation at lower voltages and improving signal [11].
Source Temperature	~100 °C (desolvation gas)	Aids in the evaporation of solvent from charged droplets. Must be optimized for specific flow rates.

Table 2: Key Validation Parameters for Nonclinical Dose Formulation Analysis

This table outlines the core experiments required for validating an nESI method for pharmaceutical QC [58].

Validation Parameter	Experimental Goal	Recommended Acceptance Criteria
Accuracy & Precision	Determine closeness and repeatability of measurements.	Defined pre-validation; typically within ±15% of nominal concentration for accuracy and <15% RSD for precision.
Specificity/Selectivity	Demonstrate no interference from vehicle/excipients.	Chromatogram shows clean baseline at analyte retention time for blank vehicle.
Linearity & Range	Establish proportional response to analyte concentration.	Correlation coefficient (r) > 0.99 over the specified range.
Solution Stability	Assess analyte integrity under storage/analysis conditions.	Concentration within acceptable deviation (e.g., ±15%) of fresh sample.

Essential Research Reagent Solutions

Table 3: The Scientist's Toolkit: Essential Materials for nESI and Urinalysis

A list of key reagents, materials, and equipment required for the experiments described in this guide.

Item	Function & Application
Sharp Singularity nESI Emitter	Specially designed emitter with controlled, sharp geometry and hydrophobic coating to stabilize the meniscus and improve ionization efficiency and repeatability [2].
High-Purity MS-Grade Solvents	Solvents (water, acetonitrile, methanol) with minimal metal ion content to prevent adduct formation and background noise [11].
Sterile Urine Collection Kit	Includes a sterile container and cleansing towels for obtaining a clean-catch mid-stream urine sample, minimizing contamination [59].
Chemical Reagent Strips (Dipsticks)	Impregnated strips for rapid, semi-quantitative chemical analysis of urine (pH, protein, glucose, blood, etc.) [60] [61].
Certified Reference Standard (API)	A well-characterized analyte with a certificate of analysis (COA) documenting purity, used for preparing calibration standards and QC samples in method validation [58].

Workflow and Process Diagrams

Diagram 1: nESI Troubleshooting Logic Pathway

Diagram 2: Method Validation Pathways for Pharmaceutical QC

Troubleshooting Guides

Guide 1: Diagnosing and Correcting In-Source Activation Caused by Emitter Positioning

Problem: Collision-Induced Dissociation (CID) breakdown curves or Collision-Induced Unfolding (CIU) transitions are shifting unexpectedly between experiments, showing inconsistent midpoint potentials (CID50/CIU50).

Root Cause: Unintended in-source activation is occurring due to suboptimal nano-electrospray ionization (nESI) emitter position relative to the instrument inlet. The position affects the electric field and collisional heating ions experience before controlled activation experiments [4].

Diagnosis and Solutions:

• Symptom: CID50/CIU50 values shift to lower voltages when the emitter is retracted or positioned farther from the inlet on some instruments.

  • Investigation: Systematically map the CID50 for a standard protein, such as holomyoglobin, against emitter position to create a spatial "map" for your specific instrument [4].
  • Solution: Reproduce the "sweet spot" position that provides optimal signal with minimal in-source activation and standardize this position for all experiments. Document the coordinates relative to the inlet center.

• Symptom: Significant variation in data between different users or days.

  • Investigation: Check for inconsistencies in how emitters are positioned. A "close" position is often defined as the emitter tip being as close as possible without touching the inlet cone (e.g., within 0.2 mm vertically and 1 mm horizontally of the inlet center) [4].
  • Solution: Implement a standardized positioning protocol. Use a fixed alignment jig or microscope to ensure consistent placement. Avoid tuning the signal solely by moving the emitter, as this can introduce unintended activation [4].

• Symptom: Poor signal-to-noise and broad peaks, especially in solutions with non-volatile salts.

  • Investigation: Confirm the emitter geometry and surface chemistry. Standard pulled-glass emitters can clog and exhibit wetting instabilities [2].
  • Solution: Consider using specialized emitters. Theta emitters (with two channels) allow mixing sample in biological buffer with a stream of volatile salt like ammonium acetate directly at the tip, improving signal in challenging matrices [12]. Hydrophobically coated emitters can stabilize the spray meniscus [2].

Guide 2: Addressing Reproducibility Issues in CIU Fingerprints

Problem: High root-mean-square deviation (RMSD) between CIU fingerprints of the same protein acquired under supposedly identical conditions.

Root Cause: Inconsistent ion activation history prior to the IM-MS separation and activation cell, often caused by variable emitter geometry or position.

Diagnosis and Solutions:

• Symptom: Additive shift in CIU50 values for all structural transitions of a protein.

  • Investigation: This is a classic sign of pre-activation in the ion path before the collision cell, such as in the StepWave region on a Waters Synapt G2-Si instrument [4].
  • Solution: Ensure the emitter position is fixed and replicated precisely. For the Synapt G2-Si, positions closer to the instrument inlet can cause significantly greater in-source activation [4].

• Symptom: Unstable spray leading to fluctuating ion signal and noisy CIU data.

  • Investigation: Instability can arise from the emitter geometry. A poorly defined or rounded emitter tip leads to a variable meniscus size, which directly impacts droplet size and ion emission stability [2].
  • Solution: Use emitters with a sharp, well-defined edge. This ensures the meniscus anchors at a consistent diameter, leading to a more stable spray and consistent results [2]. Monitor the spray voltage, as the stable range for proteomics workflows is typically 1.7 to 2.5 kV [2].

Frequently Asked Questions (FAQs)

Q1: Why is emitter position so critical for CIU/CID reproducibility, and how does it cause variation? The nESI emitter position determines the electric field strength and the path ions travel before entering the mass spectrometer. Even small variations can change the amount of collisional activation ions experience in the source region (in-source activation). This unintended activation adds to the deliberate energy applied during CIU/CID experiments, effectively shifting the observed CID50 or CIU50 values. One study showed this shift can be as large as 8 V on a Waters Synapt G2-Si instrument simply by changing the emitter position [4].

Q2: My protocol requires tuning for signal intensity. How can I optimize signal without compromising CIU/CID data with bad emitter positioning? First, optimize for signal using a "far" position that is documented and standardized. Once a position with good signal is found, keep it fixed. Avoid the practice of frequently re-tuning the emitter position for maximum signal, as this is a major source of irreproducibility. Signal can also be improved by other means, such as using high-pressure nanoESI (HP-nanoESI), which allows for higher ion inlet temperatures and applied potentials, improving desolvation and signal intensity without electrical discharge [62].

Q3: Are some instrument types more susceptible to emitter position effects than others? Yes, the effect is instrument-dependent. Research has shown that a Waters Synapt G2-Si instrument exhibits significant shifts in CID50 when the emitter position is changed, with closer positions causing greater activation. In contrast, an Agilent 6545XT instrument showed different, less pronounced effects for the ions studied [4]. You should characterize the effect on your specific instrument.

Q4: What is the single most important factor in emitter design for achieving reproducible spray? A sharp, well-defined emitter geometry is critical. The meniscus size, which dictates initial droplet size and spray stability, is defined by the emitter's outer geometry and sharpness. A sharp, acute angle with a well-defined edge provides a stable anchorage point for the meniscus, eliminating a major source of variability [2].

Q5: How can I analyze proteins from solutions with biological buffers and non-volatile salts without extensive desalting? Theta emitters are a powerful solution. These emitters have a septum dividing the capillary into two channels. You can load your protein sample, dissolved in a physiological buffer, into one channel, and a volatile salt solution (like ammonium acetate) into the other. Incomplete mixing at the tip promotes the formation of droplets depleted of non-volatile salts, allowing analysis of proteins in their native buffer [12].

Experimental Protocols for Key Investigations

Protocol 1: Mapping the Effect of Emitter Position on CID50

Objective: To quantitatively determine how nESI emitter position affects the observed CID50 value on your specific instrument.

Materials:

• Standard protein solution (e.g., 5-15 µM holomyoglobin in 200 mM ammonium acetate) [4].
• Pulled borosilicate glass nESI emitters (e.g., ~2 µm i.d. opening) [4].
• Mass spectrometer with CID capability.

Method:

• Define Coordinates: Establish a coordinate system relative to the center of the instrument's inlet cone (e.g., x: vertical, y: horizontal along inlet axis, z: horizontal perpendicular to inlet).
• Set Initial Position: Place the emitter at a "close" reference position (e.g., x=0, y=0.5, z=0 mm) [4].
• Acquire CID Data: For a selected precursor ion (e.g., holomyoglobin), acquire a CID breakdown curve by stepping the collision energy and plotting the fraction of remaining precursor ion.
• Calculate CID50: Determine the injection potential at which 50% of the precursor ion has dissociated.
• Reposition and Repeat: Systematically move the emitter to new (x,y,z) positions and repeat steps 3-4. Record the CID50 at each point.
• Create Spatial Map: Plot the shift in CID50 as a function of emitter position to identify zones of high and low in-source activation.

Protocol 2: Standardized CIU Fingerprint Acquisition with Fixed Emitter Geometry

Objective: To acquire highly reproducible CIU fingerprints by controlling for emitter geometry and position.

Materials:

• Protein of interest (e.g., Bovine Serum Albumin or NIST monoclonal antibody) in 200 mM ammonium acetate [4].
• Commercially available emitters with controlled, sharp geometry (e.g., "The Sharp Singularity" emitters) [2].
• IM-MS instrument with CIU capability.

Method:

• Emitter Selection: Use a batch of emitters with a verified, consistent tip geometry to minimize variability.
• Fixed Positioning: Use a mechanical stage or jig to place the emitter at a pre-determined "sweet spot" position identified in Protocol 1. Document this position.
• Data Acquisition: Acquire CIU data for your protein standard. For each experiment, record the number of scans integrated and ensure the electrospray is stable throughout [12].
• Data Analysis: Process all data using consistent software and settings (e.g., in CIUSuite 2) [4]. Calculate the RMSD between fingerprints from different days or emitters to validate reproducibility.

Table 1: Quantitative Impact of Emitter Position on CID/CIU Metrics

Analyte	Instrument	Emitter Position Change	Observed Effect on CID50/CIU50	Citation
Holomyoglobin	Waters Synapt G2-Si	Spatial variation across positions	Shift of up to 8 V	[4]
Leucine Enkephalin	Waters Synapt G2-Si	Closer vs. farther from inlet	Significant shift in CID midpoint	[4]
Shiga Toxin Subunit B	Waters Synapt G2-Si	Closer vs. farther from inlet	Significant shift in CID midpoint	[4]
BSA / NIST mAb	Waters Synapt G2-Si	Spatial variation	Shifts in CIU50 and increased RMSD between fingerprints	[4]

Table 2: Research Reagent Solutions for Improved Reproducibility

Item	Function/Explanation	Reference
Sharp Singularity Emitters	Mechanically sharpened emitters with controlled geometry to stabilize the meniscus and improve spray consistency.	[2]
LOTUS Coated Emitters	Hydrophobically coated emitters that lock the meniscus at the inner diameter, resulting in a smaller, more stable spray.	[2]
Theta Emitters	Dual-channel emitters for analyzing samples in non-volatile salts by mixing with volatile buffers at the tip.	[12]
Ammonium Bromide/Iodide	Additives with low proton affinity anions that help reduce sodium adduction and chemical noise in complex matrices.	[12]

Workflow and Relationship Diagrams

Factors Affecting CIU/CID Data

Emitter Positioning Optimization Workflow

Troubleshooting Guides

Troubleshooting Signal Suppression from Complex Matrices

Problem: Weak or absent analyte signal when analyzing samples in biological matrices like whole blood, serum, or buffers containing high concentrations of non-volatile salts.

Explanation: Non-volatile salts and matrix components can crystallize and clog the nanoESI emitter, suppress ionization efficiency through competitive processes, and cause significant signal deterioration [24] [63].

Solutions:

• Implement In-Capillary Extraction: For complex biofluids like whole blood, use an in-capillary liquid/liquid extraction technique. This involves preparing your sample in an organic solvent within the glass capillary to selectively extract analytes while leaving salts and proteins behind [24].
• Utilize Automated Online Desalting: Employ a microfluidic device with a molecular weight cutoff (MWCO) membrane for automated online sample concentration, desalting, and buffer exchange. This has been proven effective for analyzing adeno-associated viruses (AAVs) directly from cell culture media [64].
• Optimize Source Conditions: Lower the flow rates, temperatures, and spray voltages to softer conditions to preserve labile complexes and reduce in-source fragmentation [63].
• Apply Voltage Pulsing: Use electrophoretic separation mode by applying alternating high voltage (e.g., from -5 kV to +2 kV). This can enable the detection of multiply-charged protein ions in buffers with high concentrations of nonvolatile salts by periodically disrupting salt buildup [24].

Troubleshooting Ionization of Nonpolar Molecules

Problem: Inability to detect nonpolar or low-polarity molecules (e.g., PAHs, steroids) using standard nanoESI.

Explanation: Conventional nanoESI primarily ionizes polar molecules that can be easily protonated or deprotonated. Nonpolar analytes lack these functional groups and are therefore largely invisible to standard ESI-MS [65].

Solutions:

• Integrate a Plasma-Based Post-Ionization Source: Construct a simple Dielectric Barrier Discharge Ionization (DBDI) source using a commercial ozone generator power supply. This generates a low-temperature plasma in the ion transfer region that ionizes gas-phase nonpolar molecules via charge transfer or Penning ionization, producing intact molecular ions (M•+) [65].
• Activate Dual nESI/nAPCI Mode: If using a dual nESI/nAPCI source, apply high voltage (e.g., >4 kV) to simultaneously generate charged droplets and corona discharge plasma. This allows for the concurrent detection of polar (via nESI) and nonpolar (via nAPCI) analytes from the same sample [24].
• Switch Ionization Techniques: For less polar molecules where nESI fails, consider using dedicated Atmospheric Pressure Chemical Ionization (APCI) or Atmospheric Pressure Photoionization (APPI) sources, which are more effective for such compounds [66].

Troubleshooting Emitter Clogging and Instability

Problem: Frequent clogging or physical damage (burning/breakage) of the nanoESI emitter tip, especially with high-salt samples or at high voltages.

Explanation: High concentrations of analyte or non-volatile salts can lead to crystallization at the emitter tip [63]. Furthermore, in contact-mode nESI, applying high voltage (e.g., 5-8 kV) directly to the solution causes significant Joule heating, which can boil the solution and damage the glass tip [24].

Solutions:

• Adopt a Non-Contact nESI Configuration: Charge the analyte solution through electrostatic induction by placing the high-voltage electrode near, but not in contact with, the liquid. This setup allows the application of high voltages (>4 kV) without damaging the emitter tip, significantly improving its stability and lifetime [24].
• Optimize Analyte Concentration: Avoid excessively high sample concentrations. For modern MS, low micromolar concentrations are often sufficient. High concentrations promote clustering and crystallization [63].
• Use Appropriate Emitters: For robust, unattended operation with complex samples, consider using a heated electrospray ionization source (HESI) with a larger inner diameter emitter (~70 μm) instead of manually-pulled nanospray tips, which are more prone to clogging [64].

Frequently Asked Questions (FAQs)

Q1: What is the simplest way to analyze both polar and nonpolar compounds in a single, complex microsample without pre-treatment? A1: The most straightforward approach is to use a dual non-contact nESI/nAPCI source. This integrated platform allows simultaneous detection of polar analytes (via nESI) and nonpolar analytes (via corona discharge nAPCI) from microliter volumes of untreated samples, such as raw biofluids [24].

Q2: How can I accurately determine the sensitivity and performance of my method in complex matrices? A2: You should establish key quantitative figures of merit by spiking analytes into your matrix of interest. The table below summarizes the exemplary performance achieved by advanced nanoESI techniques in complex matrices.

Table 1: Exemplary Analytical Performance in Complex Matrices Using Advanced nanoESI Techniques

Analyte	Sample Matrix	Technique	Limit of Detection (LOD)	Key Performance Feature
Cocaine	Untreated Whole Human Blood	Non-contact nESI/nAPCI with in-capillary extraction	Part-per-trillion (pg/mL)	High sensitivity for polar analyte [24]
β-Estradiol	Untreated Whole Human Blood	Non-contact nESI/nAPCI with in-capillary extraction	Part-per-billion (ng/mL)	Efficient detection of nonpolar analyte [24]
Polycyclic Aromatic Hydrocarbons (PAHs)	Methanolic Solution / Fish Tissue	Plasma-assisted nanoESI (DBDI)	~10 ng/mL	Enables detection of nonpolar molecules [65]
AAV Capsids	Cell Culture Media	Automated Online CDMS (SS-CDMS)	<2×10⁹ capsids required	Robust analysis in complex media without clogging [64]

Q3: My protein sample is in a high-salt buffer. What is a quick method to desalt it for nanoESI-MS? A3: For a rapid, online clean-up, you can use a microfluidic device like SampleStream, which is equipped with a 100-kDa MWCO membrane. It performs automated buffer exchange into a volatile buffer like ammonium acetate and concentrates the sample directly before MS analysis, completing the process in under 15 minutes [64]. Alternatively, for manual preparation, multiple rounds of buffer exchange using 100-kDa centrifugal filters can be effective [64].

Q4: Why does my nanoESI signal change when I adjust the position of the emitter, and how can I ensure reproducibility? A4: The position of the nESI emitter relative to the instrument inlet significantly affects the extent of in-source activation and desolvation. Emitter positions closer to the inlet can result in greater unintentional activation, shifting CID50 and CIU50 values and affecting spectral reproducibility [4]. To ensure consistent results, carefully document and standardize the emitter's x, y, and z coordinates for all experiments, and use a defined "far" position unless closer positioning is necessary for signal stability [4].

Experimental Protocols

Protocol: Direct Analysis of Complex Microsamples Using Dual nESI/nAPCI

Purpose: To enable the simultaneous detection of polar and nonpolar analytes from untreated complex matrices (e.g., biofluids) with high sensitivity [24].

Materials:

• Disposable borosilicate glass capillaries (ID 1.17 mm, pulled tip ≤5 μm)
• Dual non-contact nESI/nAPCI source
• Mass spectrometer (e.g., Thermo Fisher Velos Pro)
• Ag auxiliary electrode
• High-voltage power supply (capable of >4 kV)

Workflow:

Procedure:

• Sample Loading: Load a small volume (≤5 μL) of the untreated sample (e.g., whole blood) into a pulled borosilicate glass capillary [24].
• Source Configuration: Place the capillary in the source. Ensure the high-voltage Ag electrode is positioned with an approximately 1 cm air gap from the capillary tip, operating in non-contact mode [24].
• Voltage Application:
  • For analyzing polar compounds only, apply a lower spray voltage (e.g., ≤3 kV). This operates in pure nESI mode [24].
  • For analyzing both polar and nonpolar compounds, apply a high voltage above the breakdown threshold of air (e.g., 6 kV). This activates the simultaneous nESI/nAPCI mode, where the high voltage induces a corona discharge from the auxiliary electrode [24].
• Data Acquisition: Acquire mass spectra with instrument parameters optimized for sensitivity. For the described setup, typical parameters include a capillary temperature of 400°C and a short distance (e.g., 5 mm) from the ion source to the MS inlet [24].

Protocol: Plasma-Assisted nanoESI for Nonpolar Molecules

Purpose: To detect nonpolar molecules that are difficult to ionize with conventional nanoESI by using a low-cost dielectric barrier discharge (DBD) plasma source [65].

Materials:

• Ozone generator power supply (Input: DC 12 V, Output: AC 5 kV, 50 kHz)
• Teflon tube (i.d. 1/16 inch), stainless-steel capillary, copper ring
• Standard nanoESI source and pulled quartz emitters
• Mass spectrometer

Workflow:

Procedure:

• Source Assembly: Construct the DBDI source by coupling a Teflon tube (acting as the dielectric) to the extended ion transfer tube of the mass spectrometer. Insert a stainless-steel capillary to serve as the grounded electrode. Wrap a copper ring tightly around the Teflon tube to act as the high-voltage electrode [65].
• Power Connection: Connect the copper ring and stainless-steel capillary to the output terminals of the commercial ozone generator power supply [65].
• Sample Introduction: Introduce the sample containing nonpolar analytes (e.g., PAHs dissolved in a 1:1 methylene chloride/methanol mixture) using a standard nanoESI setup with a typical spray voltage (e.g., 1.6 kV) [65].
• Plasma Activation: Turn on the ozone generator power supply to generate low-temperature plasma inside the Teflon tube. Allow approximately 10 seconds for the plasma to stabilize. The plasma will ionize the gas-phase nonpolar molecules, allowing their detection as molecular ions (M•+) [65].

The Scientist's Toolkit: Essential Research Reagents & Materials

Table 2: Key Reagents and Materials for Robust Nanoelectrospray MS

Item	Function / Application	Specific Example / Note
Pulled Borosilicate Capillaries	NanoESI emitter for sample introduction.	~2 μm i.d. tip for standard nESI; ≤5 μm for non-contact nESI/nAPCI [24] [4].
Ammonium Acetate Solution	A volatile buffer for native MS and buffer exchange.	Preferred over non-volatile salts to prevent source contamination and signal suppression [63] [4].
Volatile Organic Solvents	Sample dissolution and liquid/liquid extraction.	Methanol, acetonitrile, ethyl acetate. Used for in-capillary extraction from biofluids [24].
Centrifugal Filters (100-kDa MWCO)	Offline buffer exchange and desalting of large molecules (e.g., proteins, AAVs).	Effective for removing salts and small impurities prior to MS analysis [64].
Microfluidic Device with MWCO Membrane	Automated online sample clean-up and concentration.	Enables robust, high-throughput analysis of samples in complex matrices like cell culture media [64].
Ozone Generator Power Supply	Powers a Dielectric Barrier Discharge (DBD) plasma source.	Low-cost solution (Input DC 12V, Output AC 5kV) for ionizing nonpolar molecules [65].
Inert Metal Wires	Applying high voltage to the sample solution in nanoESI emitters.	Platinum wires are commonly used for this purpose [63].

Conclusion

Enhancing sensitivity in nanoelectrospray MS requires a multifaceted approach that integrates fundamental understanding with advanced methodologies and meticulous optimization. The strategies outlined—from employing novel emitter designs like theta tips and pulsed nESI to optimizing operational parameters and using strategic additives—collectively push the boundaries of what is detectable. These advancements are crucial for biomedical research, enabling the analysis of proteins and complexes directly from physiologically relevant buffers, high-throughput metabolic phenotyping of large cohorts, and sensitive detection of therapeutics in biological fluids. Future directions will likely focus on increasing automation and robustness for clinical translation, further miniaturization for single-cell analyses, and developing even more effective methods for analyzing samples in their native state. By systematically applying these principles, researchers can significantly improve the sensitivity, reproducibility, and scope of their nESI-MS analyses, driving discoveries in drug development and clinical diagnostics.

References

• 1. Ionization Efficiency, Ion Suppression and Detection ... [https://sciex.com/tech-notes/ce/ionization-efficiency-ion-suppression-and-detection-sensitivity]
• 2. nano-ElectroSpray Technology - Fossiliontech [https://www.fossiliontech.com/nano-esi]
• 3. Reduced Pressure Ionization Enhances Native Mass ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12559447/]
• 4. Effects of Nano-Electrospray Ionization Emitter Position on ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC11315166/]
• 5. Optimization of Capillary Vibrating Sharp-Edge Spray ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC11983206/]
• 6. Fast and sensitive flow-injection mass spectrometry ... [https://www.nature.com/articles/s41467-020-17026-6]
• 7. Principles of Electrospray Ionization - PMC - PubMed Central [https://pmc.ncbi.nlm.nih.gov/articles/PMC3134074/]
• 8. Observation of charged droplets from electrospray ionization ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC8410725/]
• 9. Surface Modified Nano-Electrospray Needles Improve ... [https://experts.arizona.edu/en/publications/surface-modified-nano-electrospray-needles-improve-sensitivity-fo/]
• 10. Improving the Sensitivity of Mass Spectrometry by Using a ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC4177967/]
• 11. 10 Great Tips for Electrospray Ionization LC–MS [https://www.chromatographyonline.com/view/the-lcgc-blog-10-great-tips-for-electrospray-ionization-lc-ms]
• 12. Improved Mass Analysis of Proteins and ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12453605/]
• 13. Electrospray characteristic curves: in pursuit of improved ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC2625288/]
• 14. Nanoelectrospray Emitter Arrays Providing Inter- ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC2587423/]
• 15. Pulsed Nanoelectrospray Ionization Boosts Ion Signal in ... [https://www.mdpi.com/2076-3417/11/22/10883]
• 16. Toward routine utilisation of native mass spectrometry as ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12319726/]
• 17. Toward routine utilisation of native mass spectrometry as an ... [https://pubs.rsc.org/en/content/articlehtml/2025/md/d5md00617a]
• 18. Optimization and Application of Direct Infusion ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC5387673/]
• 19. Direct infusion mass spectrometry metabolomics dataset [https://www.nature.com/articles/sdata201412]
• 20. Surface Modified Nano-Electrospray Needles Improve ... [https://zenodo.org/records/14640201]
• 21. FAQ & Troubleshooting [https://dispendix.com/knowledge-center]
• 22. Optimization and Application of Direct Infusion ... - Advion, Inc. [https://www.advion.com/rsc-publication/optimization-and-application-of-direct-infusion-nanoelectrospray-hrms-method-for-large-scale-urinary-metabolic-phenotyping-in-molecular-epidemiology/]
• 23. Electric modeling and characterization of pulsed high-voltage nanoelectrospray ionization sources by a miniature ion trap mass spectrometer - PubMed [https://pubmed.ncbi.nlm.nih.gov/30989769/]
• 24. Direct Mass Spectrometry Analysis of Complex Mixtures by Nano-Electrospray with Simultaneous Atmospheric Pressure Chemical Ionization and Electrophoretic Separation Capabilities - PMC [https://pmc.ncbi.nlm.nih.gov/articles/PMC6822699/]
• 25. Alternately Pulsed Nano-electrospray Ionization ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC2575744/]
• 26. Direct Mass Spectrometry Analysis of Complex Mixtures by ... [https://pubmed.ncbi.nlm.nih.gov/31449396/]
• 27. Dynamic Spray Mass Spectrometry for Direct Analysis ... [https://pubmed.ncbi.nlm.nih.gov/41104880/]
• 28. Electrokinetic Manipulations Combined with Direct and ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12272651/]
• 29. Characterization and Analysis of Paper Spray Ionization ... [https://link.springer.com/article/10.1007/s13361-017-1826-5]
• 30. Critical Role of Voltage Application Points in “Analog” and “ ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12063163/]
• 31. Optimization of Electrospray Ionization by Statistical Design of ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC4972871/]
• 32. Solution Additives that Desalt Protein Ions in Native Mass ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC3433631/]
• 33. Desalting large protein complexes during native electrospray ... [https://pubs.rsc.org/en/content/articlehtml/2015/an/c4an02334j]
• 34. Probing Metal Ion Adduction in the ESI Charged Residue ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC11756706/]
• 35. Desalting strategies for native mass spectrometry [https://www.sciencedirect.com/science/article/abs/pii/S0039914024012037]
• 36. Direct Analysis of Proteins from Solutions with High Salt ... [https://link.springer.com/article/10.1007/s13361-018-1893-2]
• 37. Exploring the mechanism of salt-induced signal ... [https://pubmed.ncbi.nlm.nih.gov/25594702/]
• 38. Two decades of studying non-covalent biomolecular ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC3306659/]
• 39. Collision-Induced Dissociation of Electrosprayed NaCl ... [https://link.springer.com/article/10.1007/s13361-016-1468-z]
• 40. Mass calibration options for accurate electrospray ... [https://www.sciencedirect.com/science/article/pii/S1387380621000993]
• 41. Disruption and nebulization of lipid vesicles using surface ... [https://pubs.rsc.org/en/content/articlehtml/2025/lc/d5lc00692a]
• 42. Recent developments in ionization techniques for single- ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC10733462/]
• 43. Enabling high-sensitivity live single-cell mass spectrometry ... [https://www.sciencedirect.com/science/article/pii/S0003267024008699]
• 44. NanoLC-EI-MS: Perspectives in Biochemical Analysis [https://www.mdpi.com/1422-0067/24/14/11746]
• 45. New Ionization Processes for Ionizing Nonvolatile ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12156021/]
• 46. Mitigating clogging of nESI with nonvolatile buffers that ... [https://just.ustc.edu.cn/article/doi/10.3969/j.issn.0253-2778.2020.11.005]
• 47. Nanobubbles in Electrospray Ionization Mass Spectrometry [https://pmc.ncbi.nlm.nih.gov/articles/PMC11948179/]
• 48. Mitigating Wetting and Scaling in Air Gap Membrane ... [https://www.mdpi.com/2077-0375/15/10/321]
• 49. Design Characteristics to Eliminate the Need for Parameter ... [https://link.springer.com/article/10.1007/s13361-019-02301-8]
• 50. Improving Liquid Chromatography-Mass Spectrometry ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC3187566/]
• 51. Direct nano-electrospray ionization tandem mass ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC6894584/]
• 52. Optimization of Electrospray Ionization Source Parameters ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC6297073/]
• 53. Performance Comparisons of Nano-LC Systems, Electrospray ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC3886563/]
• 54. Which electrospray-based ionization method best reflects ... [https://pubmed.ncbi.nlm.nih.gov/18083584/]
• 55. Optimization of the Electrospray Ionization Source with ... [https://www.spectroscopyonline.com/view/optimization-electrospray-ionization-esi-source-use-design-experiments-doe-approach-lc-ms-ms-determi]
• 56. Electrospray Ionisation Mass Spectrometry: Principles and ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC1853331/]
• 57. Nanoelectrospray—more than just a minimized-flow ... [https://www.sciencedirect.com/science/article/pii/S1044030598001573]
• 58. NonClinical Dose Formulation Analysis Method Validation ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC2976985/]
• 59. Urinalysis - StatPearls - NCBI Bookshelf - NIH [https://www.ncbi.nlm.nih.gov/books/NBK557685/]
• 60. Urinalysis - Clinical Methods - NCBI Bookshelf - NIH [https://www.ncbi.nlm.nih.gov/books/NBK302/]
• 61. Urinalysis Information | Mount Sinai - New York [https://www.mountsinai.org/health-library/tests/urinalysis]
• 62. Analytical characteristics of nano-electrospray operated ... [https://www.sciencedirect.com/science/article/abs/pii/S0003267018304057]
• 63. Modern Electrospray Ionization Mass Spectrometry ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC11099892/]
• 64. A Robust and Automated Platform for Charge Detection ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12323853/]
• 65. Simultaneous Detection of Polar and Nonpolar Molecules ... [https://pmc.ncbi.nlm.nih.gov/articles/PMC12195691/]
• 66. Ion Sources [https://www.bruker.com/en/products-and-solutions/mass-spectrometry/ion-sources.html]