<p><strong>🎤Alex Brody</strong></p><p><strong>In this episode of Concentrating on Chromatography, we sit down with Alex Brody, an organic chemist at Aqua America, to discuss the rigorous analytical methods required for regulated drinking water testing.</strong></p><p>Alex walks us through the multi-step PFAS detection process using EPA Method 537.1, including extraction, nitrogen blowdown concentration, and LC-MS analysis. He explains why sample preparation and quality control are critical for achieving trace-level detection—and why these methods can't be rushed or simplified, even with new technologies available.</p><p>We also explore taste and odor investigations, disinfection byproducts, and volatile organic compounds, plus the surprising reality of why regulated labs move slowly when adopting new instrumentation. You'll learn about the quality control checkpoints, peer review processes, and the misconceptions surrounding analytical turnaround times in compliance labs.</p><h5><strong>Topics Covered:</strong></h5><ul><li>Role of organic chemistry in regulated drinking water labs</li><li>EPA compliance requirements and regulatory bodies (EPA, PA-DEP)</li><li>PFAS detection using Method 537.1 and LC-MS</li><li>The critical importance of nitrogen blowdown sample concentration</li><li>Taste and odor analysis: MIB, Geosmin, and unknown compound identification</li><li>Why regulated methods evolve slowly (validation timelines, approval processes)</li><li>Quality control procedures: calibration checks, matrix spikes, surrogate standards, internal standards</li><li>Automated sample preparation and lab efficiency</li><li>Future PFAS regulations and Method 533 expansion</li></ul><h5><strong>Ideal For:</strong></h5><ul><li>Analytical chemists and environmental lab professionals</li><li>Water utilities and compliance officers</li><li>Chromatography practitioners interested in regulated workflows</li><li>Anyone curious about how drinking water safety is ensured</li></ul><h2><strong>Video Transcription</strong></h2><h3><strong>Ensuring Safe Drinking Water Through Advanced Analytical Chemistry</strong></h3><h4><strong>Insights from a Regulated Water Testing Laboratory</strong></h4><p>Analytical laboratories responsible for regulated drinking water testing operate under strict compliance frameworks to ensure the delivery of safe and reliable water supplies. In this interview, Alex, an organic chemist at Aqua America, outlines the scope of analytical work conducted in a modern water quality laboratory. The team primarily focuses on the determination of taste- and odor-causing compounds, disinfection byproducts, volatile organic compounds (VOCs), and per- and polyfluoroalkyl substances (PFAS). These analyses rely heavily on <strong>GC-MS and LC-MS instrumentation</strong>, supported by rigorous maintenance programs to ensure continuous operational performance and data reliability.</p><h4><strong>Regulatory Compliance and Quality Control in Drinking Water Analysis</strong></h4><p>Operating as a regulated drinking water laboratory requires adherence to stringent requirements established by agencies such as the <strong>U.S. Environmental Protection Agency (EPA)</strong> and state-level regulatory bodies. Compliance is driven by extensive quality control protocols integrated into every analytical batch. Typical workflows include calibration verification, matrix spike recovery assessments, internal standard monitoring, and surrogate performance checks. Notably, analytical batches are often limited in sample number to accommodate a substantial proportion of quality control samples, reflecting the critical importance of data defensibility in regulatory reporting.</p><p>One common misconception is the perceived speed at which results can be delivered. In reality, comprehensive sample preparation, including extraction procedures and validation steps, can extend turnaround times to a full working day or longer. Additionally, validated regulatory workflows impose limitations on the rapid adoption of new analytical technologies, as method modifications must undergo formal evaluation and approval before implementation.</p><h4><strong>Analytical Strategies for Taste and Odor Investigations</strong></h4><p>Taste and odor events represent a significant operational challenge for water utilities. Standard targeted methods are routinely applied for compounds such as <strong>2-methylisoborneol (MIB)</strong> and <strong>geosmin</strong>, which are commonly associated with earthy or musty sensory characteristics. When targeted analysis does not reveal the source, laboratories employ <strong>broad-scan GC-MS screening</strong> combined with spectral library interpretation to identify potential causative compounds. In some cases, sensory evaluation panels complement instrumental analysis, providing preliminary insight into potential treatment-related or environmental influences affecting water quality.</p><h4><strong>PFAS Monitoring: Methodological Complexity and Trace-Level Sensitivity</strong></h4><p>PFAS determination represents a rapidly evolving priority in drinking water monitoring. The laboratory currently applies <strong>EPA Method 537.1</strong>, with efforts underway to expand capabilities through certification for <strong>Method 533</strong>, thereby broadening analyte coverage. Analytical workflows typically involve solid-phase extraction followed by solvent evaporation using nitrogen-assisted blow-down techniques to achieve the trace-level sensitivity required for regulatory compliance. This concentration step is essential because PFAS compounds remain non-volatile under controlled evaporation conditions, enabling enhanced detection performance upon reconstitution and LC-MS analysis.</p><h4><strong>Future Directions in Drinking Water Analytics</strong></h4><p>Looking ahead, the primary technological driver in regulated water analysis is the demand for <strong>increasingly lower detection limits</strong> and expanded compound coverage. While regulatory frameworks can slow methodological innovation, laboratories continue to pursue efficiency improvements, such as automated sample preparation systems and optimized analytical workflows. These advances support the growing need to process high sample volumes while maintaining the stringent data quality standards required for public health protection.</p><p>Ultimately, the work performed in regulated drinking water laboratories represents a critical intersection of analytical chemistry, environmental monitoring, and public safety. Continuous methodological refinement, coupled with advances in mass spectrometry and sample preparation technologies, will play a central role in addressing emerging contaminants and ensuring long-term water quality assurance.</p><blockquote><p><strong>This text has been automatically transcribed from a video presentation using AI technology. It may contain inaccuracies and is not guaranteed to be 100% correct.</strong></p></blockquote><blockquote><h4><strong>Concentrating on Chromatography Podcast</strong></h4><p>Dive into the frontiers of chromatography, mass spectrometry, and sample preparation with host David Oliva. Each episode features candid conversations with leading researchers, industry innovators, and passionate scientists who are shaping the future of analytical chemistry. From decoding PFAS detection challenges to exploring the latest in AI-assisted liquid chromatography, this show uncovers practical workflows, sustainability breakthroughs, and the real-world impact of separation science. Whether you’re a chromatographer, lab professional, or researcher you'll discover inspiring content!</p><p>You can find <strong>Concentrating on Chromatography Podcast </strong>in podcast apps:</p><ul><li><a target="_blank" rel="noopener noreferrer" href="https://open.spotify.com/show/1wZg67rCjoGju4yhqxI3tL"><strong>Spotify</strong></a></li><li><a target="_blank" rel="noopener noreferrer" href="https://podcasts.apple.com/tz/podcast/concentrating-on-chromatography/id1788429185"><strong>Apple podcast</strong></a></li><li><a target="_blank" rel="noopener noreferrer" href="https://podscan.fm/podcasts/concentrating-on-chromatography"><strong>podscan</strong></a></li></ul><p>and on <a target="_blank" rel="noopener noreferrer" href="https://www.youtube.com/@ChromatographyTalk/videos"><strong>YouTube channel</strong></a></p></blockquote>

Regulated Drinking Water Testing: PFAS Detection, Sample Prep & Real Lab Insights

<p><strong>This study investigates how ionization polarity influences the gas-phase stability of native-like proteins analyzed by ion mobility spectrometry–mass spectrometry. While proteins generally retain folded structures in both positive and negative electrospray ionization modes, significant differences in unfolding behavior were observed.</strong></p><p>Proteins exhibited greater gas-phase stability when analyzed in the ionization polarity corresponding to their solution-phase charge, requiring higher energy for unfolding. These findings emphasize the importance of considering protein isoelectric points and ionization polarity in native IMS-MS experiments to ensure accurate structural characterization.</p><blockquote><h4><strong>The original article</strong></h4><h4><a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/10.1021/acs.analchem.5c04295"><strong>Investigating the Effect of Isoelectric Points on the Gas-Phase Stability of Native-like Proteins Analyzed in Positive- versus Negative-Ion Mode by IMS-MS</strong></a></h4><p>Alexis N. Edwards, Madeline G. Bannon, Michael S. Cordes, and Elyssia S. Gallagher*</p><p>Anal. Chem. 2026, 98, 6, 4518–4527</p><p><a target="_blank" rel="noopener noreferrer" href="https://doi.org/10.1021/acs.analchem.5c04295">https://doi.org/10.1021/acs.analchem.5c04295</a></p><h6>licensed under <a target="_blank" rel="noopener noreferrer" href="https://creativecommons.org/licenses/by/4.0/"><strong>CC-BY 4.0</strong></a></h6></blockquote><p><strong>Selected sections from the article follow. Formats and hyperlinks were adapted from the original.</strong></p><div><p>In native mass spectrometry (nMS), the native-like structures of proteins are maintained in the gas phase by preserving the secondary, tertiary, and quaternary structures. <a href="javascript:void(0);">(1−7)</a> The preservation of native-like protein structures is primarily accomplished via the use of electrospray ionization (ESI), a soft ionization method that forms a distribution of multiply charged protein ions. These multiply charged ions can then be investigated to characterize stoichiometry, oligomeric composition, and ligand binding. <a href="javascript:void(0);">(8−10)</a></p><p>During ESI, a potential is applied to the sample-containing solution. In an aqueous spray solvent, H<sub>3</sub>O<sup>+</sup> and OH<sup>–</sup> are produced in positive-ion mode (+ESI) and negative-ion mode (−ESI), respectively. <a href="javascript:void(0);">(11)</a> The buildup of like-charges within the solution results in the formation of a Taylor Cone at the capillary tip, which ejects charged droplets that travel toward the inlet of the mass spectrometer. Droplets undergo evaporation and fission until complete analyte desolvation, leading to the release of folded, gas-phase protein ions. As the droplets shrink, H<sub>3</sub>O<sup>+</sup> or OH<sup>–</sup> can become concentrated, causing the pH of the droplet to decrease or increase, respectively. Shifts in pH can denature proteins. Therefore, to maintain native-like protein structures, nMS is conducted with solutions containing volatile salts, usually ammonium acetate, that provide ionic strength and minimize pH changes compared to spraying from pure water. <a href="javascript:void(0);">(11,12)</a> Droplet pH in ESI is influenced by flow rate, solvent composition, and applied voltage, <a href="javascript:void(0);">(13)</a> but often approaches values near the p<i>K</i><sub>a</sub> of acetate (4.76) or ammonium (9.25) in +ESI or −ESI, <a href="javascript:void(0);">(12,14)</a> respectively. During ESI, changes in overall protein charge take place via proton transfer at chargeable amino acid residues, resulting in the formation of protonated or deprotonated protein ions with the charged sites being dependent on the relative proton affinities of ammonium, acetate, and chargeable amino acids as proteins near complete desolvation. <a href="javascript:void(0);">(15−19)</a></p><p>Ion mobility spectrometry (IMS) measures an ion’s mobility through an inert gas under the influence of an electric field. Thus, mobility is directly influenced by the ion’s size, shape, and charge. <a href="javascript:void(0);">(20)</a> A protein’s mobility can be used to estimate its structure by calculating an average momentum-transfer collisional cross section (CCS). In addition to structural characterization, IMS-MS can investigate the gas-phase stability of protein complexes when paired with collision-induced unfolding (CIU). Specifically, in CIU experiments, protein ions are accelerated into a cell filled with inert gas molecules and undergo multiple collisions with higher frequency and energy relative to IMS. This converts kinetic energy to internal, vibrational energy, disrupting noncovalent interactions and inducing unfolding of the protein ions. <a href="javascript:void(0);">(21)</a> Unfolding alters the protein structure, which can then be quantified by monitoring the protein’s mobility during subsequent IMS-MS measurement. <a href="javascript:void(0);">(22)</a> Prior work with mutant proteins has shown that proteins with similar intramolecular interactions have similar CIU fingerprints, while proteins with different intramolecular interactions have different CIU fingerprints. <a href="javascript:void(0);">(23)</a> Therefore, gas-phase protein stability is correlated with intramolecular interactions.</p><p>Past native IMS-MS work has primarily been conducted in +ESI given that -ESI has the reputation of being unreliable for native-like protein analysis due to the occurrence of corona discharge, <a href="javascript:void(0);">(24)</a> and charge-state distributions that may not be representative of folded proteins. <a href="javascript:void(0);">(25)</a> In recent years, −ESI has been increasingly used in CIU and IMS-MS, with recent work demonstrating that −ESI-MS is capable of discerning denatured and native-like protein structures. <a href="javascript:void(0);">(26,27)</a> Prior work showed that charge manipulation of folded proteins, such as charge reduction, to achieve similar absolute charge states in both ion modes, results in CCS values of protein cations that are comparable to their anion counterparts. <a href="javascript:void(0);">(26,28)</a> This suggests that gas-phase structures are weakly dependent on protein charge and ionization polarity. <a href="javascript:void(0);">(23,26,28)</a> Yet, Hong et al. showed that, in the absence of charge reduction, the gas-phase stability of two proteins (avidin and β-lactoglobulin) is sensitive to ionization polarity, highlighting that gas-phase stability is not consistent between +ESI and −ESI. <a href="javascript:void(0);">(29)</a></p><h4><strong>Materials and Methods</strong></h4><h5><strong>Native IMS-MS Analysis</strong></h5><p>Glass capillaries were pulled using a P-1000 micropipette puller (Sutter Instrument Company, Novato, CA). Samples and calibrants were sprayed from pulled capillaries (inner diameter ∼1 μm) and analyzed with a <a target="_blank" rel="noopener noreferrer" href="https://lcms.labrulez.com/products/152"><strong>Synapt G2-S High-Definition MS </strong></a>(Waters Corporation, Milford, MA). The trap and transfer cells contained Ar, the helium cell contained He, and the IMS cell contained N<sub>2</sub>. IMS wave velocity was 350 m/s, while IMS wave height was either 7 or 9 V, depending on the protein size (see <a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/suppl/10.1021/acs.analchem.5c04295/suppl_file/ac5c04295_si_001.pdf">Table S2</a>). Proteins with larger mass required a wave height of 9 V to ensure that all the ions migrated out of the IMS cell prior to the next ion injection. Mass spectra were collected and signal averaged over 30 s. Additional instrument details can be found in the Supporting Information (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/suppl/10.1021/acs.analchem.5c04295/suppl_file/ac5c04295_si_001.pdf">Table S2</a>).</p><p>To limit systematic error due to repositioning of the ESI tip between runs, <a href="javascript:void(0);">(48)</a> the <i>XYZ</i>-stage position was optimized and kept consistent for all replicates. Approximately 5 μL of sample was loaded into a glass capillary, which was then mounted on the <i>XYZ</i>-stage. Electric potential was delivered to the sample by a platinum wire inserted into the capillary and placed in contact with the solution. The voltage applied to the tip holder ranged from 0.60 to 1.2 kV for native samples, and 0.80 to 1.40 kV for denatured samples. Nine replicates in each ion mode were collected for all proteins with measurements spread across 8 days over 3 weeks. These replicates were conducted in pairs by collecting data in both ion modes consecutively. To accomplish this, a CIU experiment was collected in +ESI before the instrument polarity was changed to −ESI. A second CIU experiment was performed, beginning with −ESI to counteract any changes in protein stability due to pH changes during ESI. <a href="javascript:void(0);">(14)</a> This allowed for use of the same capillary, at the same position in relation to the instrument inlet, for each +ESI/–ESI replicate pair. A new tip was used for each +ESI/–ESI replicate pair. CIU experiments were accomplished by increasing collision voltage (CV) to the trap cell in 5 V steps, from 0 V to a maximum of 120 V.</p><h4><strong>Results and Discussion</strong></h4><p>Ten proteins were chosen for analysis in +ESI and −ESI in this work (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/10.1021/acs.analchem.5c04295#tbl1">Table 1</a>). These proteins can be broken into three groups depending on the net-charge in solution at pH 7: net-negative (with pI &lt; 6.5), net-neutral (with 6.5 ≤ pI ≤ 7.5), and net-positive (with pI &gt; 7.5). Five of the ten proteins have a net-negative charge at pH 7, though four of these proteins (con A, β-lactoglobulin, ovalbumin, and α-lactalbumin) are routinely analyzed via +ESI-IMS-MS. <a href="javascript:void(0);">(18,28)</a> Additionally, each protein had charge-state distributions in +ESI and −ESI that resulted in at least one overlapping charge state (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/10.1021/acs.analchem.5c04295#fig1">Figure 1</a>), allowing for direct comparison of <sup>TW</sup>CCS<sub>N2→He</sub> and CIU50 in both ion modes.</p><p><img src="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/Anal_Chem_2026_98_6_4518_4527_Figure_1_Representative_mass_spectra_in_ESI_top_and_ESI_bottom_cb81976247.jpg" alt="Anal. Chem. 2026, 98, 6, 4518–4527: Figure 1. Representative mass spectra in +ESI (top) and −ESI (bottom) for (A) ovalbumin, (B) GNA, (C) β-lactoglobulin, (D) α-lactalbumin, (E) con A, (F) streptavidin, (G) ubiquitin, (H) WGA, (I) cytochrome c, and (J) lysozyme. Proteins that have blue or red charge states indicate additional oligomeric states that were detected. Outlines in red boxes are overlapping charge states that are compared herein. *Indicates contaminant peaks." srcset="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/thumbnail_Anal_Chem_2026_98_6_4518_4527_Figure_1_Representative_mass_spectra_in_ESI_top_and_ESI_bottom_cb81976247.jpg 245w," sizes="100vw" width="1600" height="865"></p><h5><strong>Proteins with Disulfide Bridges are Stable in Both Polarities</strong></h5><p>For both lysozyme and α-lactalbumin, the average <sup>TW</sup>CCS<sub>N2→He</sub> were within experimental uncertainty in +ESI and −ESI for both the initial, folded structures and the final, extended structures (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/10.1021/acs.analchem.5c04295#tbl2">Table 2</a>). However, the 6+ and 6- ions of lysozyme and α-lactalbumin showed a high resistance to unfolding with &gt;90% of the ions retaining their folded <sup>TW</sup>CCS<sub>N2→He</sub> values even at high CV (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/10.1021/acs.analchem.5c04295#fig2">Figure 2</a>). Replicate data showed similar trends (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/suppl/10.1021/acs.analchem.5c04295/suppl_file/ac5c04295_si_001.pdf">Figure S2</a>). While CCS values provide a description of gas-phase structures, CIU experiments provide information on the stability of the gas-phase protein structures. Furthermore, the number and pattern of disulfide bridges present in antibodies has been characterized by CIU. <a href="javascript:void(0);">(63)</a> Lysozyme and α-lactalbumin both contain the glycosyl hydrolases family 22 (GH22) domain, which has four disulfide bridges that span ∼94% and ∼93% of the amino acid sequence, respectively (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/suppl/10.1021/acs.analchem.5c04295/suppl_file/ac5c04295_si_001.pdf">Figure S3</a>). Thus, we hypothesize that rather than effects from protein pI and ionization polarity, the unfolding behavior of these proteins is controlled to a greater extent by the presence of stabilizing disulfide bridges that prevent routine unfolding during collisional activation.</p><p><img src="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/Anal_Chem_2026_98_6_4518_4527_Figure_2_Representative_mobiligrams_showing_arrival_times_for_lysozyme_A_C_and_a_lactalbumin_B_D_0f3d23c3c1.jpg" alt="Anal. Chem. 2026, 98, 6, 4518–4527 - Figure 2. Representative mobiligrams showing arrival times for lysozyme (A, C) and α-lactalbumin (B, D) at low and high CV (blue solid and orange dashed lines, respectively) for +ESI (A, B) and −ESI (C, D). For lysozyme, different values were chosen for the high CV (e.g., 540 eV in +ESI and 480 eV in −ESI) because these were the last analyzed CV where the signal-to-noise ratio (S/N) for the 6 ± ions was greater than 3." srcset="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/thumbnail_Anal_Chem_2026_98_6_4518_4527_Figure_2_Representative_mobiligrams_showing_arrival_times_for_lysozyme_A_C_and_a_lactalbumin_B_D_0f3d23c3c1.jpg 212w," sizes="100vw" width="1600" height="1178"></p><p>To characterize the stabilizing effect of disulfides, MD simulations were conducted that heat denatured 6+ ions of lysozyme and α-lactalbumin with disulfide bridges either present or removed with the resulting cysteine residues being protonated prior to heating (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/suppl/10.1021/acs.analchem.5c04295/suppl_file/ac5c04295_si_001.pdf">Figure S4</a>). Structural changes associated with heat denaturation were characterized by taking the root-mean-square deviation (RMSD) of the heated structures relative to initial, crystal structures. Over ten trials, lysozyme ions possessed an average RMSD of (1.6 ± 0.2) nm when disulfides were present which increased to (3.5 ± 0.5) nm when disulfides were removed. Similarly, the RMSD of α-lactalbumin ions increased from (1.8 ± 0.1) nm when disulfides were present to (3.7 ± 0.3) nm when disulfides were removed, suggesting that the disulfides serve as a barrier to unfolding. This is visually represented in <a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/suppl/10.1021/acs.analchem.5c04295/suppl_file/ac5c04295_si_001.pdf">Figure S4C,D</a> where simulated structures of heated lysozyme and α-lactalbumin ions with intact disulfides adopt more condensed configurations relative to those with disulfides removed. In essence, the disulfides “tie” disparate regions of protein together, which serves as a barrier to more thorough unfolding even at a maximum simulated temperature of 900 K, explaining the similarity of <sup>TW</sup>CCS<sub>N2→He</sub> values in +ESI versus −ESI as well as the limited percentage of ions that undergo a change in <sup>TW</sup>CCS<sub>N2→He</sub> upon additional collisional activation.</p><p>WGA has 16 disulfide bridges per monomeric subunit (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/10.1021/acs.analchem.5c04295#tbl1">Table 1</a>). On average, these disulfides encompass seven amino acids each, with ∼78% the total amino acids in WGA being encompassed by at least one disulfide bridge (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/suppl/10.1021/acs.analchem.5c04295/suppl_file/ac5c04295_si_001.pdf">Figure S3</a>). Compared to lysozyme and α-lactalbumin, the presence of disulfide bridges across small numbers of amino acids enables WGA to unfold with ΔCCS of (370 ± 60) Å<sup>2</sup> and (390 ± 120) Å<sup>2</sup> in +ESI and −ESI, respectively (<a target="_blank" rel="noopener noreferrer" href="https://pubs.acs.org/doi/10.1021/acs.analchem.5c04295#tbl2">Table 2</a>). However, in comparison to the other lectins, GNA and con A, we hypothesize that these disulfide bridges in WGA limit the extent of unfolding, resulting in the extended, <sup>TW</sup>CCS<sub>N2→He</sub> values being within experimental uncertainty in both ion modes.</p><p>To summarize, lysozyme and α-lactalbumin exhibit similar gas-phase behavior in both +ESI and −ESI, with most ions retaining folded structures even under high CV. We hypothesize this resistance to unfolding is largely attributed to their four disulfide bridges, which span over 90% of their sequences and act as stabilizing constraints. MD simulations confirm that removing these disulfides significantly increases structural deviation during heat denaturation, indicating their critical role in maintaining compact configurations. Consequently, the presence of disulfide bridges explains the minimal changes in <sup>TW</sup>CCS<sub>N2→He</sub> and the proteins’ high stability under activation.</p><h4><strong>Conclusions</strong></h4><p>IMS-MS allows for detailed analysis of the structure and stability of native-like proteins in the gas phase. The structure of a protein is dictated by its amino acid sequence and the resulting noncovalent interactions that form the secondary, tertiary, and quaternary structures. Chargeable amino acid residues that are solvent exposed can carry charges leading to the protein having a net-positive, net-negative, or net-neutral charge in solution at physiological pH. Yet, most native IMS-MS analysis is conducted in +ESI, leading to proteins that are net-negative in solution experiencing large swings in their net-charge during the ESI process.</p><p>This work highlights that gas-phase stability of native-like proteins varies with ionization polarity. While CCS values for folded structures are generally comparable between +ESI and −ESI, unfolding energetics differ─net-negative proteins are more stable in −ESI, whereas net-positive proteins are more stable in +ESI. We hypothesize that these differences are due to changes in the intramolecular interactions within cationic and anionic proteins. Therefore, proteins analyzed in the mode opposite to their solution-phase charge may have different gas-phase structures compared to biological systems. For example, proteins that are net-negatively charged in physiological environments (pI &lt; 6.5) are more likely to represent their biological counterparts as gas-phase anions when analyzed in −ESI compared to +ESI, enabling nMS experiments to more closely characterize the structures present in biological systems.</p></div>

Investigating the Effect of Isoelectric Points on the Gas-Phase Stability of Native-like Proteins Analyzed in Positive- versus Negative-Ion Mode by IMS-MS

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<header><p><strong>Join Us for an Engaging Live Webinar on Integrating UV/Vis Spectroscopy Into Your Laboratory Workflows</strong></p></header><main><div><div><p><strong>Program&nbsp;</strong></p><ul><li>Introduction to UV/Vis Spectroscopy</li><li>The Power of Integration</li><li>Live Demonstration</li><li>Customer Use Cases</li><li>Engage &amp; Discuss</li></ul></div></div><div><div><p>Discover how connecting your UV/Vis spectrophotometer with instruments like balances can transform your laboratory processes. By transmitting dissolution data directly to the spectrophotometer, you can streamline workflows and enhance data quality, leading to improved productivity and accuracy.</p></div></div><div><div><div><p>In this informative session, we’ll showcase the power of instrument integration and how it can revolutionize your daily routines. Using real-world case studies from various industries, including battery manufacturing, pharmaceuticals, and food and beverage, you’ll gain valuable insights into optimizing your laboratory workflows.</p><p><strong>Program Highlights</strong></p><ul><li><strong>Introduction to UV/Vis Spectroscopy</strong><br>Understand the basics and applications of UV/Vis spectroscopy in laboratories.</li><li><strong>The Power of Integration</strong><br>Explore the benefits of connecting your UV/Vis spectrophotometer with balances and other instruments.</li><li><strong>Live Demonstration</strong><br>Witness a live demonstration of data transmission from a balance to a UV/Vis spectrophotometer.</li><li><strong>Customer Use Cases</strong><br>Learn about integrated processes used in the battery, pharma, and food &amp; beverage industries.</li><li><strong>Engage &amp; Discuss</strong><br>Participate in a Q&amp;A session where we address your questions and discuss potential applications in your own lab.</li></ul><p>Don’t miss this opportunity to explore innovative solutions that can help you think critically about your own processes and enhance your laboratory's efficiency!</p></div></div></div></main><div><div><p><strong>Target Audience</strong></p><div><p>This webinar is designed for laboratory professionals, scientists, and managers across various industries, including:</p><ul><li>Pharmaceuticals: Quality control and research &amp; development teams looking to improve analytical processes</li><li>Food &amp; Beverage: Quality assurance and product development personnel aiming to enhance product testing efficiency</li><li>Battery Manufacturing: Engineers and technicians interested in optimizing material testing workflows</li><li>Laboratory Managers: Professionals seeking to streamline operations and improve data quality through instrument integration</li></ul><p>Whether you are just starting with UV/Vis spectroscopy or looking to enhance your existing laboratory workflows, this webinar is perfect for you. Register now to secure your spot and take the first step towards a more efficient laboratory.</p><ul><li><span style="padding-right:0px;" id="webinar-session-1"><strong>1. 10. 2025 9:00,&nbsp; English&nbsp;</strong></span><span id="webinar-session-2"><strong><svg width="16" height="16" viewBox="0 0 16 16" fill="none" xmlns="http://www.w3.org/2000/svg"></svg></strong></span></li><li><span id="webinar-session-2"><strong>1. 10. 2025 17:00,&nbsp; English&nbsp;</strong></span></li></ul><blockquote><p><span id="webinar-session-2"><strong>Presenter: </strong></span>Anika Adams (Product Manager, SPG UV/VIS)</p><div><p>Anika Adams is Product Manager for UV/Vis Spectrophotometers at METTLER TOLEDO, where she focuses on driving innovation and optimizing customer workflows. With a degree in business administration and experience spanning software development, mechanical engineering, and production, Anika brings a multidisciplinary perspective to the laboratory business. Her expertise lies in automating and enhancing customer processes, ensuring both product excellence and customer satisfaction.</p><p><strong>Presenter:</strong> Dr. Fabian Müller (Product Manager, SPG UV/VIS)</p><div><div><div><div><p>Dr. Fabian Müller, Ph.D., is an accomplished chemist from the ETH Zürich in Switzerland. After completing his doctoral studies, Dr. Müller joined the team at METTLER TOLEDO as an Application Specialist for UV/Vis spectroscopy, where he quickly established himself as a skilled and knowledgeable professional. His expertise in this area led to his promotion to the position of Product Manager for UV/Vis spectrophotometers.</p></div></div></div></div></div></blockquote></div></div></div>

Mettler Toledo: UV/Vis Integration in Laboratory Workflows 

UV/Vis Integration in Laboratory Workflows 

Join Us for an Engaging Live Webinar on Integrating UV/Vis Spectroscopy Into Your Laboratory Workflows

<div><h4><strong>Start Your Journey to ASTRA Expertise</strong></h4><p>Join us for the last <strong>5 series</strong> on building the perfect method for light scattering analysis.</p><p>Learn advanced tools and techniques to master ASTRA software, from normalization to DLS with flowing systems.</p><h4><strong>Session Overview:</strong></h4><ul><li><strong>Part 6 of 10</strong> Baselines &amp; Peaks - Best Practices &amp; Navigating Unexpected Chromatograms<ul><li>Thursday, September 11</li></ul></li><li><strong>Part 7 of 10 </strong>Light Scattering Formalisms - Zimm, Mie, Berry, Debye, oh my! When and Why to Use Them<ul><li>Thursday, September 25</li></ul></li><li><h5><strong>Part 8 of 10: Calculation Uncertainty and Error Analysis: Establishing Meaningful MALS Results</strong></h5><ul><li>Thursday, October 2</li></ul></li><li><strong>Part 9 of 10: </strong>Distribution Analysis &amp; Results Fitting: Advanced Peak Analysis Tools<ul><li>Thursday, October 9</li></ul></li><li><strong>Part 10 of 10:</strong> Mass Recovery &amp; Data Exporting: Effective Reporting of Results and Findings<ul><li>Thursday, October 23</li></ul></li></ul><h4><strong>Why Attend:</strong></h4><ul><li>Deepen your understanding of light scattering and ASTRA.</li><li>Gain actionable insights for your lab.</li><li>Interact with our expert, Dr. Christian Ackerschott.<br>&nbsp;</li></ul><blockquote><p><strong>Hosted by:</strong> Dr. Christian Ackerschott</p><p>Dr. Christian Ackerschott studied Chemistry at the University of Cologne and finished with his diploma thesis in Biochemistry. He completed his studies with the PhD thesis “Characterization of Recombinant Flagelliform Spider Silk Proteins” at the Technische Universität München, where AF4-MALS was used as an essential characterization method.</p><p>Since 2007, Christian works at Wyatt Technology as Application Scientist and is now Head of Customer Service. His tasks also include customer trainings in Dernbach and onsite as well as education and training of the local Field Application Scientists.</p></blockquote></div>

Waters Corporation: Interactive Web Series EU – Becoming a SEC-MALS & ASTRA Expert

Becoming a SEC-MALS & ASTRA Expert: Part 8

Improve the reliability of your results by understanding error sources and uncertainty in MALS. Learn how to establish meaningful conclusions.

Becoming SEC-MALS & ASTRA Expert Part 8

<div><p><strong>Analytical methods are essential to ensuring pharmaceutical product quality, regulatory compliance, and patient safety. However, their reliability and reproducibility can be compromised by multiple sources of variability—including reagent quality, instrument performance, operator technique, sample preparation, and data processing. Understanding and controlling these sources is critical, especially when justifying post-approval changes to validated methods.</strong></p><p>This three-part webinar series features expert Peter Baker (Live Oak QA, Austin, Texas) and focuses on the importance of identifying, monitoring, and documenting variability within analytical workflows. Through open discussion and real-world insights, participants will gain a deeper understanding of how to build a scientifically sound case for method adjustments while maintaining regulatory compliance. Each session includes a live Q&amp;A to encourage interaction and address specific challenges.</p><h4><strong>Session 3: Analytical Control in Action – Real-World Case Studies</strong></h4><p>The final session presents real-world examples of method variability and how companies have successfully justified changes due to shifts in materials, instruments, or processes. Emphasis will be placed on documentation, regulatory communication, and lessons learned.</p><p><i><strong>Goal: </strong>Apply insights from case studies to your own workflows and gain practical strategies for managing change.</i><br>&nbsp;</p><blockquote><p><strong>Presenter:</strong> Peter Baker (President, Live Oak Quality Assurance LLC)</p><p>Peter E. Baker, President, Live Oak Quality Assurance LLC, has been consulting nationally and internationally since April 2019. Prior to consulting, Peter spent 11 years as a U.S. FDA Drug Investigator, with most of those years spent working in FDA’s overseas offices located in India, China and Chile. Peter was named FDA Investigator of the Year in 2013 for his work uncovering serious breaches in data integrity and has special interest in data governance and compliance with 21 CFR Part 11. Live Oak focuses on training and employee empowerment in critical thinking, with the goal of improving the quality of medicines through good data management.</p><p><strong>Presenter: </strong>Dr. Stephanie Harden (Senior Manager, Small Molecule Core Solutions – LC-MS, Waters Corporation)</p><p>Stephanie Harden, Senior Manager at Waters Corporation, leads the Product Marketing and Scientific teams for Waters small molecule HPLC portfolio. With over 25 years of experience in product and segment marketing, she has a proven track record in strategic program development, cross-functional leadership, and market-driven innovation. Her work connects scientific insight with commercial strategy to drive customer-focused solutions and sustained business growth. Stephanie holds a PhD in Chemistry from the University of Bristol, UK, and is a widely cited author.</p></blockquote></div>

Waters Corporation: Managing Method Variability: A Foundation for Risk-Based Change

Managing Method Variability: A Foundation for Risk-Based Change - Session 3

Discover real case studies on managing analytical variability, documenting changes, and maintaining compliance in pharma workflows.

Analytical Control in Action – Real-World Case Studies

<p><strong>Dr. Michaela Nebel, Product Manager at Sensolytics GmbH, gives an overview of the theory of localized electrochemical techniques with a focus on Scanning Electrochemical Microscopy (SECM), Scanning Droplet Cell technique (SDC), and Scanning Electrochemical Cell Microscopy (SECCM). Then, application examples from different fields in electrochemistry, including electrocatalysis, corrosion research, and sensors are highlighted.</strong></p><p>In recent years, the rising inclusion of localized electrochemical analysis into classical electrochemical methods has allowed for more comprehensive characterization of sample surfaces from various fields of application.</p><h5><strong>Key learnings of this webinar:</strong></h5><ul><li><strong>Theoretical background of localized electrochemical techniques</strong> with a focus on Scanning Electrochemical Microscopy (SECM), Scanning Droplet Cell technique (SDC), and Scanning Electrochemical Cell Microscopy (SECCM)</li><li><strong>Guidelines for the selection</strong> of a localized technique fitting your individual research application</li><li><strong>Practical considerations</strong> for the use of the respective analysis method</li><li><strong>Selected applications</strong> from different fields of research, including electrocatalysis, corrosion research, and sensors</li></ul><blockquote><p><strong>Presenter: </strong>Dr. Michaela Nebel (Product Manager at Sensolytics GmbH)</p></blockquote>

Metrohm: Introduction to Localized Electrochemical Techniques

Introduction to Localized Electrochemical Techniques

Learn SECM, SDC, and SECCM fundamentals and applications in electrocatalysis, corrosion research, and sensors. Practical guidance from experts.

<div><div container-id="cq-colctrl-lt0" comp-id="A1"><div><p><strong>As global food safety regulations become increasingly stringent and consumer scrutiny intensifies, laboratories are under pressure to deliver rapid, accurate results while managing complex sample matrices.</strong></p><p>Analyzing emerging contaminants such as PFAS, pesticides, veterinary drugs and other residues in processed or high-fat foods poses significant challenges in method robustness, instrument uptime and data reliability.</p><p>This webinar will explore how the QSight 500 LC-MS/MS is addressing these evolving demands to provide high-quality data for even the most demanding food matrices.</p><h5><strong>Attend this webinar to:</strong></h5><ul><li>Gain insights into how to optimize your LC-MS/MS workflows for better efficiency and confidence in your results</li><li>Understand how to reliably detect contaminants across complex matrices&nbsp;</li><li>Learn how the QSight 500 LC-MS/MS system brings greater operational efficiency with reduced instrument cleaning&nbsp;</li><li>Explore how enhancing system robustness reduces cost per sample&nbsp;</li></ul><blockquote><p><strong>Presenter: </strong>Jamie Foss (Product Manager, Liquid Chromatography &amp; Mass Spectrometry at PerkinElmer)</p><p>Jamie Foss is the Liquid Chromatography and Mass Spectrometry Product Manager at PerkinElmer. Prior to this role, he was a Sr. Application Scientist developing strategic applications in support PerkinElmer’s LC and LC/MS product portfolio. He has developed a wide variety of applications across food, environmental, industrial, forensics and cannabis. Prior to joining PerkinElmer in 2016, he spent 4 years as a forensic chemist for the State of Maine where he focused on the analysis of controlled substances and clandestine laboratory investigation.</p></blockquote></div></div></div>

Technology Networks: Addressing Evolving Demands in Food Analysis

Addressing Evolving Demands in Food Analysis

Optimize food contaminant analysis with QSight 500 LC-MS/MS. Robust, efficient workflows for PFAS, pesticides, and residues in demanding food matrices.

<div><div container-id="cq-colctrl-lt0" comp-id="A1"><div><p><strong>Prof. Thalassinos's research focuses on understanding how cells respond to DNA damage using single-cell proteomics. The study employs a compound called 60G to induce DNA damage, which activates DNA repair mechanisms such as mismatch repair, base excision repair, and nucleotide excision repair. By analyzing single cells, the research aims to uncover the variability in protein expression and the biological processes involved in DNA repair and apoptosis.</strong></p><p>The team uses advanced techniques like the Orbitrap Eclipse mass spectrometer and the SLN1 single-cell isolation system to isolate and analyze individual cells. This approach allows them to observe the upregulation of proteins involved in DNA repair and the arrest of cells in the S phase of the cell cycle. The findings highlight the importance of single-cell proteomics in revealing cellular heterogeneity and the dynamic response to DNA damage, which could have implications for understanding diseases like cancer and neurodegenerative disorders.&nbsp;</p><h5><strong>Learning points:</strong></h5><ul><li><strong>Single-Cell Proteomics Advantage: </strong>Single-cell proteomics allows for the observation of individual cellular responses to DNA damage, revealing heterogeneity that would be masked in bulk proteomics. This approach helps in understanding the specific activation of DNA repair mechanisms and cell cycle arrest at the S phase.&nbsp;</li><li><strong>DNA Repair Mechanisms: </strong>The study highlights the activation of various DNA repair pathways, such as mismatch repair, base excision repair, and nucleotide excision repair, in response to DNA damage induced by the compound 60G. This understanding is crucial for insights into diseases like cancer and neurodegenerative disorders.&nbsp;</li><li><strong>Technological Integration: </strong>The use of advanced technologies like the <strong>Orbitrap Eclipse</strong> mass spectrometer and the SLN1 single-cell isolation system enables precise isolation and analysis of single cells, facilitating detailed proteomic studies and enhancing the understanding of cellular processes at a granular level.&nbsp;</li></ul><h5><strong>Who should attend:</strong></h5><ul><li><strong>Cancer Researchers: </strong>Understanding the variability in DNA repair mechanisms at the single-cell level can provide insights into tumor heterogeneity and the development of targeted cancer therapies.</li><li><strong>Neurodegenerative Disease Scientists</strong>: Insights into DNA damage response mechanisms can aid in understanding the molecular basis of neurodegenerative diseases and potentially lead to new therapeutic strategies.&nbsp;</li><li><strong>Proteomics and Genomics Technologists:</strong> Professionals working with advanced proteomics and genomics technologies can benefit from the presentation's focus on cutting-edge techniques for single-cell analysis, enhancing their research capabilities and methodologies.&nbsp;</li></ul><p>If you cannot attend this webinar please register to receive a link to the On Demand version the following day.&nbsp;</p><blockquote><p><strong>Presenter: </strong>Prof. Konstantinos Thalassinos (Professor, University College London)</p><p>Kostas moved to the UK in 1998 to study for a BSc in Genetics at the University of Leicester, having completed his high school education in his native Greece. After graduation he went on to the University of York to undertake a Masters in Bioinformatics, which included a three-month industrial placement writing software to process mass spectrometric data. Having gained an interest in mass spectrometry, Kostas then went to the University of Warwick in order to pursue a PhD that combined experimental and computational studies in mass spectrometry-based proteomics. After being awarded his PhD in 2006, Kostas was the successful recipient of a one-year Wellcome Trust Value in People Fellowship. Post-doctoral training followed and in 2010 he moved to the Institute of Structural and Molecular Biology at University College London / Birkbeck to take up a lectureship in biophysical mass spectrometry and has been there ever since. In 2015 he was promoted to Senior Lecturer and in 2019 to Professor of mass spectrometry.</p><p>His lab is using structural mass spectrometry approaches, especially ion mobility and crosslinking, to study the structure and dynamics of proteins and protein complexes, particularly those involved in protein misfolding diseases.</p></blockquote></div></div></div>

Thermo Fisher Scientific: ASMS 2025 Workshops

Probing DNA-damage response at the single cell level

Explore single-cell proteomics in DNA damage research. See how Orbitrap Eclipse and SLN1 reveal repair mechanisms, cell cycle arrest, and disease insights.

<div><div container-id="cq-colctrl-lt0" comp-id="A1"><div><p><strong>Dr. Hale's research integrates native ambient mass spectrometry (NAMS) to analyze intact proteoform complexes in tissue samples under non-denaturing conditions. Utilizing nano-DESI (nano spray desorption electrospray ionization), this technique enables the spatial mapping of proteoforms within tissues, revealing their distribution and functional implications. The method involves home-built ion sources that provide high-resolution imaging compatible with advanced mass spectrometers like the Orbitrap Eclipse.</strong></p><p>The presentation highlights the ability to analyze proteoforms and their post-translational modifications (PTMs) directly from tissue samples without extensive preparation, preserving the native state of protein complexes. This approach is particularly useful for studying neurodegenerative diseases, where understanding the spatial distribution and interactions of proteoforms can provide insights into disease mechanisms. Dr. Hale emphasizes the importance of this technology in advancing pathology research and improving the detection and identification of high molecular weight proteins and membrane protein complexes.</p><h5><strong>Learning points:</strong></h5><ul><li><strong>Spatial Distribution Analysis: </strong>Nano-DESI mass spectrometry enables the spatial mapping of proteoforms within tissues, providing insights into their distribution and functional roles, which is crucial for understanding disease mechanisms, particularly in neurodegenerative diseases.&nbsp;</li><li><strong>Non-Denaturing Conditions: </strong>The technique preserves the native state of protein complexes during analysis, allowing for the study of intact proteoforms and their post-translational modifications (PTMs) directly from tissue samples without extensive preparation.&nbsp;</li><li><strong>Advanced Mass Spectrometry Techniques: </strong>Utilizing high-resolution mass spectrometers like the Orbitrap Eclipse, the method allows for detailed analysis of high molecular weight proteins and membrane protein complexes, enhancing the detection and identification capabilities for complex biological samples</li></ul><h5><strong>Who should attend:</strong></h5><ul><li><strong>Pathology Researchers: </strong>This technique provides valuable insights into the spatial distribution and interactions of proteoforms within tissues, aiding in the understanding of disease mechanisms and pathology.&nbsp;</li><li><strong>Neurodegenerative Disease Scientists: </strong>The ability to analyze intact proteoform complexes in their native state is particularly beneficial for studying neurodegenerative diseases, offering potential breakthroughs in understanding disease progression and identifying therapeutic targets.&nbsp;</li><li><strong>Mass Spectrometry Technologists:</strong> Professionals working with advanced mass spectrometry techniques can benefit from the innovative application of nano-DESI for high-resolution imaging, enhancing their analytical capabilities and methodologies for complex biological samples. &nbsp;</li></ul><p>If you cannot attend this webinar please register to receive a link to the On Demand version the following day.&nbsp;</p><blockquote><p><strong>Presenter: </strong>Dr. Oliver Hale (Postdoctoral Research Fellow, University of Birmingham)</p><p>Dr Oliver Hale is a postdoctoral research fellow who specializes in mass spectrometry. His current focus is on combing native mass spectrometry analysis of intact proteins and mass spectrometry imaging, as part of the EPSRC-funded NAMS project.</p></blockquote></div></div></div>

Imaging of intact proteoform complexes in tissue

Learn how nano-DESI native ambient MS maps proteoforms in tissues under non-denaturing conditions, advancing insights into neurodegeneration and pathology.

<p><strong>New functions of the improved UV Workstation Software will be described and demonstrated.</strong></p><blockquote><p><strong>Presenter:&nbsp;</strong> Scott Melis, PhD (Application Scientist, Agilent Technologies, Inc.)</p><p>Scott Melis has a PHD in Physics from Georgetown University in Washington, DC and started with Agilent in 2021 as a Molecular Spectroscopy Application Scientist. In his graduate work, Scott studied nanoparticle formation and growth for a variety of applications. Projects that he worked with include developing processes to deposit and characterize semiconducting molecular nano-cocrystals for use in optoelectronic devices as well as controlling polymer nanoparticle size through rapid mixing conditions.</p></blockquote>

Agilent Technologies: New Software Features for UV Work Station Software

New Software Features for UV Work Station Software

New functions of the improved UV Workstation Software will be described and demonstrated.

<h4><strong>Our newest HPLC columns are game changing</strong></h4><h5><strong>Liquid chromatography is evolving</strong></h5><p>With growing demands for sensitivity, reproducibility, and speed, today’s workflows need column technology that can keep up. Whether you are analyzing trace-level compounds or navigating complex bioseparations, the right column makes all the difference.</p><p>A new generation of LC column technology is setting the bar higher—delivering cleaner separations, sharper peaks, faster equilibration, and more reliable results for even the most metal-sensitive analytes.</p><h5><strong>Want to see it in action?</strong></h5><p>Join our webinar to explore how this new column technology transforms oligonucleotide separations for characterization, sequencing, and impurity profiling.</p><blockquote><p><strong>Presenter:&nbsp;</strong> Dr. Anne Blackwell (Product Manager Bio LC Columns, Agilent Technologies)</p><p>Anne is a product manger for bio LC columns at Agilent, with previous experience as a product support scientist for bio LC columns and an LC/MS applications scientist focused on metabolomics. &nbsp;She has a Ph.D. in analytical chemistry from the University of Arizona where she studied mass spectrometry of non-covalent protein assemblies.</p><p><strong>Presenter: </strong>Dr. Andrew Coffey (Senior Application Scientist, Agilent Technologies)</p><p>Andrew Coffey joined Agilent Technologies Biocolumns Applications Team in 2011. &nbsp;He has over 35 years experience in method development, technical support and troubleshooting of reversed phase, ion exchange, size exclusion and many other liquid chromatographic techniques.</p></blockquote>

Agilent Technologies: Inert Separations for Oligo LC-MS Workflows

Inert Separations for Oligo LC-MS Workflows

Explore Agilent’s new HPLC columns in a webinar on advanced oligonucleotide separations, sequencing, and impurity profiling.

<h4><strong>Are you navigating ICH M10 requirements for your regulated bioanalysis workflows?</strong></h4><p>Join our experts as we break down the globally harmonized ICH M10 guidelines, show how labs are applying them in practice, and explore how SCIEX OS software helps ensure quality, efficiency, and compliance.</p><h5><strong>Key highlights:</strong></h5><ul><li>What the ICH M10 guideline is and why it matters for regulated bioanalysis</li><li>How pharmaceutical labs are approaching method validation and study sample analysis</li><li>Common workflow challenges and inefficiencies in meeting regulatory expectations</li><li>How SCIEX OS software simplifies data management, calculations, and reporting</li><li>A live demonstration of current calculation workflows using SCIEX OS software</li><li>A vision for the future of streamlined bioanalytical validation</li></ul><blockquote><p><strong>Presenter: </strong>Henri Meijering, Ardena</p><p>Henri Meijering is a seasoned bioanalytical scientist with over 20 years of experience in drug development and pharmacokinetics. He specializes in developing and validating LC-MS/MS and HPLC methods for both small and large molecules. At Ardena Bioanalytical Laboratory, Henri leads scientific strategy and supports complex studies as Business Support Manager. A former core member of the EBF during the ICH M10 rollout, he is passionate about advancing scientific rigor and compliance in regulated bioanalysis.</p><p><br><strong>Presenter: </strong>Heather Hughes, SCIEX</p><p>Heather Hughes holds a degree in chemistry from Durham University and has built a career spanning clinical and pharmaceutical science. They began in NHS hospital laboratories, specializing in clinical chemistry, before moving into the pharmaceutical industry with experience at Pfizer and a contract research organization. With a background in both regulated and non-regulated environments, they joined SCIEX nine years ago as an application specialist. In this role, they support customers through training, troubleshooting, and showcasing SCIEX mass spectrometry solutions for small molecule workflows.</p></blockquote>

Streamline your ICH M10 workflows

Webinar on ICH M10 bioanalysis guidelines—learn method validation, sample analysis, and how SCIEX OS software ensures compliance and efficiency.