<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

<p><strong>Anna Vanluchene&nbsp;from UCT Prague’s&nbsp;Department of Chemical Engineering has received a&nbsp;prestigious start-up grant from the Dagmar Procházková Fund. However, she did not receive funding right away. In this interview, she describes how she developed ten versions of her&nbsp;</strong><i><strong>Illuminating Flow</strong></i><strong>&nbsp;project idea (combining photocatalysis with flow reactors for pharmaceutical synthesis). She explains why she considers perseverance to be crucial when applying for grants, how international experience has influenced her approach to team management, and how her research can reduce the chemical industry’s&nbsp;dependence on fossil fuels and imports from China.</strong></p><p><img src="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/VSCHT_Praha_Na_startovaci_grant_jsem_cekala_dva_roky_a_napsala_deset_verzi_projektu_2_5b7d63b8e9.jpg" alt="VŠCHT Prague/Tomáš Balický: I waited two years for a start-up grant and wrote ten versions of my project idea" srcset="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/thumbnail_VSCHT_Praha_Na_startovaci_grant_jsem_cekala_dva_roky_a_napsala_deset_verzi_projektu_2_5b7d63b8e9.jpg 127w," sizes="100vw" width="1128" height="1387"></p><h5><strong>What did the Dagmar Procházková Fund grant mean to you?</strong></h5><p>I&nbsp;consider it to be a&nbsp;great success. It is the first injection of funding that will allow me to ensure full independence for my research endeavour, to start my own research group, and to fund students so I&nbsp;can get started properly.</p><h5><strong>What have been your previous experiences with grants?</strong></h5><p>During my doctorate, a <i>cotutelle</i> (international doctorate under dual supervision), I&nbsp;was funded by a&nbsp;grant from the French government. Then I&nbsp;worked at the Academy of Sciences of the Czech Republic (CAS) and received a&nbsp;Marie Skłodowska-Curie Action (MSCA) Postdoctoral Fellowship to go abroad. The idea for my now-funded project was developed about two years ago and, since then, I&nbsp;have gradually developed and adapted the idea for various grant calls. I&nbsp;submitted about ten different versions of the proposal. I&nbsp;did not receive a&nbsp;grant from the Dagmar Procházková Fund last year, but this year, my application was successful.</p><h5><strong>This probably requires a&nbsp;great deal of patience and perseverance.</strong></h5><p>Certainly, because the success rates for most grants have been declining for some time. But as a&nbsp;motivational example, I&nbsp;will tell the story of Timothy Noel, a&nbsp;Belgian professor of chemistry living in the Netherlands. He applied for the prestigious European Research Council (ERC) grant, which is among the most sought-after in Europe, seven times. But no failure deterred him and eventually he succeeded and opened the door to potentially massive grant funding. He was already one of the world’s&nbsp;top scientists in the field at that time and openly described his journey on his blog <a target="_blank" rel="noopener noreferrer" href="https://www.noelresearchgroup.com/2022/03/17/my-personal-story-to-obtain-an-erc-grant-7-submissions-4-interviews-and-6-heartbreaks/"><strong>blog</strong></a>.</p><h5><strong>What advice does this convey to other grant applicants and early career researchers?</strong></h5><p>Start early, read a&nbsp;lot, don’t&nbsp;give up, and—most importantly—apply repeatedly. Learn from rejections and overcome possible frustration from failures. Read blogs and books by successful people, where you will find not only the glorious results of their work, but also the hardships they faced over their entire journeys. Of course, building your team and laboratory is impossible without finances. I&nbsp;myself still hope that I&nbsp;will be able to receive even bigger grants in the future.</p><h5><strong>How has your international experience influenced your scientific approach?</strong></h5><p>A&nbsp;lot and on several levels. Different laboratories teach you how to do&nbsp;things differently. You gain varied experience about how different institutions function. You see other people’s&nbsp;positive motivations, their ability to be ambitious without negativity, their involvement in institutional matters, their openness to cooperation with companies (e.g., tech transfers, spin-offs). The most important thing is to broaden your horizons. I&nbsp;see going abroad as a&nbsp;necessity. What I&nbsp;have taken away the most is drive, great support, and the idea that scientific output is not just about metrics, but also about dedication to one’s&nbsp;topic.</p><h5><strong>What is your </strong><i><strong>Illuminating Flow: Transport Phenomena in Photocatalysis</strong></i><strong> about, in layperson’s&nbsp;terms, and how is it unique?</strong></h5><p>Most of today’s&nbsp;chemical industry is still based on thermal processes and fossil fuels. Photochemistry enables electrification, where the energy source is photons delivered by LEDs, which can generate electricity from renewable sources. Current research focuses mainly on chemical reactions and their mechanisms, but as a&nbsp;chemical engineer, I&nbsp;focus on designing processes and reactors suitable for industrial use. In batch reactors, light fades rapidly with distance from a&nbsp;source, so much of a&nbsp;reaction mixture is not illuminated. Flow reactors solve this problem, but they bring new challenges such as working with solid catalysts and subsequently scaling up production and achieving higher production rates of the desired chemicals at a&nbsp;given scale.</p><p>One can imagine a&nbsp;transparent tube with a&nbsp;catalyst on the walls, upon which light shines and through which reactants flow. This project has a&nbsp;unique emphasis on engineering solutions and overall process efficiency.</p><p><img src="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/VSCHT_Praha_Na_startovaci_grant_jsem_cekala_dva_roky_a_napsala_deset_verzi_projektu_3_b978abf25c.jpg" alt="VŠCHT Prague/Tomáš Balický: I waited two years for a start-up grant and wrote ten versions of my project idea" srcset="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/thumbnail_VSCHT_Praha_Na_startovaci_grant_jsem_cekala_dva_roky_a_napsala_deset_verzi_projektu_3_b978abf25c.jpg 241w," sizes="100vw" width="1228" height="799"></p><h5><strong>So, were you inspired by a&nbsp;gap in the market to connect photocatalysis with transport phenomena?</strong></h5><p>Exactly. I&nbsp;combine my areas of expertise: microfluidics, photocatalytic degradation of organic substances (CAS), and synthesis (MSCA Postdoctoral Fellowship). The idea itself is not new, and pharmaceutical companies are very interested in “greener” photochemistry, but in practice, there is often a&nbsp;lack of appropriate engineering solutions and apparatus for industrial use.</p><h5><strong>What are the practical applications of your work and what substances are you focusing on?</strong></h5><p>Mainly on pharmaceuticals and on the synthesis of so-called fine chemicals, which are often the building blocks for the production of pharmaceuticals. There is also a&nbsp;geopolitical dimension to this, i.e. less dependence on imports from China and sustainable synthesis in Europe where it makes economic sense.</p><h5><strong>And what chemicals are you specifically talking about?</strong></h5><p>For example, building blocks for advanced synthesis, compounds prepared by trifluoromethylation reactions, or products of oxidation reactions with singlet oxygen. At the same time, we focus on reactions that are, in the classical design process, less safe or technologically demanding, where new approaches, such as photochemistry in flow reactors, could be safer and have better process control. Of course, this is not a&nbsp;universal solution for all types of reactions.</p><h5><strong>The goal of this project?</strong></h5><p>I&nbsp;plan to start a&nbsp;fully functional laboratory and research team, create high-quality experimental data on selected molecules and based on them, try to build cooperation with industrial partners. Without data, it is impossible to come to companies with just an idea. Within three years, I&nbsp;would like to have results that can be realistically presented and further developed.</p><h5><strong>What are the main challenges in transport phenomena during photocatalysis?</strong></h5><p>The methodology for repeatability of experiments in photochemistry is relatively demanding and there are unforeseen obstacles in building apparatus. Also, the scale-up from microliters to a&nbsp;larger scale.</p><h5><strong>What methodology will you use, and how big are the microreactors and their flow rates?</strong></h5><p>I&nbsp;have two different directions in methodological design. The first direction involves microreactors the size of a&nbsp;laboratory slide. You make a&nbsp;structure using a&nbsp;laser, and then you have to cast it and make a&nbsp;microreactor. This allows you to quickly adjust the geometry of the reactor. This method is, of course, not applicable in the commercial setting, but it is ideal for studying transport phenomena and precise measurements.</p><p>The second direction involves reactors that are flow cells. These are already larger, approximately 15 x 15 centimetres, and their flow rates are in the order of millilitres per hour. This is the initial scale that we will further increase during the project.</p><h5><strong>There is a&nbsp;big difference between a&nbsp;flow rate of microlitres per hour and industrial needs. Will it not work differently in reality?</strong></h5><p>Our research will focus on studying the phenomena themselves for now, and scaling will be addressed later. In addition, we will combine the experimental data obtained with computer simulations that will help design suitable conditions for a&nbsp;larger scale. For some high-value fine chemicals, these scales, combined with continuous operation or parallelization of reactors, may already be practically usable.</p><h5><strong>What catalysts will you use and how will you attach them to the reactor walls?</strong></h5><p>I&nbsp;am focusing on heterogeneous photocatalysis, and we will mainly use graphitic carbon nitride and its more advanced modifications. These are semiconductor materials that can be synthesized relatively easily and are suitable for immobilization on glass reactor walls. A&nbsp;key part of this project is the creation of a&nbsp;stable catalytic layer for long-term flow-through operations. To increase the contact of reactants with the catalyst, we will also consider multiphase modes, for example, gas bubbling, in order to disrupt the otherwise laminar flow.</p><h5><strong>Why did you choose this field and the Department of Chemical Engineering?</strong></h5><p>During my postdoc in Belgium, the department had an opening for an Assistant Professor position that caught my attention and I&nbsp;applied for it. I&nbsp;did my undergraduate, graduate, and initial doctoral studies here, and I&nbsp;love the department. I&nbsp;have a&nbsp;Bachelor’s&nbsp;degree in Chemistry.</p><h5><strong>Are you collaborating with anyone?</strong></h5><p>We collaborate with BUT Brno focusing on creating layers and immobilizing catalysts on glass, as well as with the Institute of Chemical Process Fundamentals of the Academy of Sciences of the Czech Republic. And as for organic synthesis, we are connected with Ghent University via postdoc internships. At the same time, my colleague Viola Tokárová and I&nbsp;received a&nbsp;Czech Science Foundation grant in cooperation with BUT Brno and Masaryk University, with our part of the project focusing on the use of flow reactors for photochemical reactions and testing the antibacterial properties of photocatalytic materials. And I&nbsp;am writing other grant proposals in which I&nbsp;am trying to build new collaborations.</p><h5><strong>What does the creation of an independent research group at UCT Prague look like?</strong></h5><p>I&nbsp;started by renovating the premises that were assigned to me and gradually building my team. It is important for research work to have Ph.D. students; I&nbsp;have already hired one and we will recruit more. In addition to building the group, I&nbsp;will engage in securing further funding for the group, so I&nbsp;will be writing more grant proposals. It is important to think far enough in advance when it comes to prospective funding for a&nbsp;research group.</p><h5><strong>What type of students are you looking for?</strong></h5><p>The laboratory and team I&nbsp;are building will be very multidisciplinary, just like the area I&nbsp;am working on. That is why I&nbsp;am looking for students who are attracted to true multidisciplinary research, those who, for example, have knowledge of engineering, but who don’t&nbsp;mind a&nbsp;little bit of pure chemistry. One student in their own right does not have to (and cannot) know every subject, so I&nbsp;will ideally have a&nbsp;team consisting of different people with different research backgrounds. Motivated, DIY-type students who would rather enthusiastically build or invent something themselves than conduct repetitive measurements. Organic or analytical chemists will also find their place here. If this topic is interesting to students in any way, they can definitely contact me. I&nbsp;am also open to interdepartmental cooperation with other UCT Prague Faculties.</p><h5><strong>How would you describe your path to photocatalysis/chemical engineering?</strong></h5><p>Studying Chemistry as an undergrad gave me a&nbsp;broad perspective, but I&nbsp;was gradually narrowing my focus and eventually chose engineering. Chemical engineering intrigued me with its logic and was the easiest for me to grasp at the time. But I&nbsp;have always enjoyed working on projects more than the studies themselves. Even during my Bachelor studies, I&nbsp;really enjoyed and was interested in the Professor Štěpánek Group’s&nbsp;projects, where I&nbsp;was doing something completely different than I&nbsp;do&nbsp;now. I&nbsp;only got into photochemistry during my time at CAS and then, during my postdoctoral fellowship, I&nbsp;devoted myself to electrochemistry. My current focus is, thus, a&nbsp;natural interlinking of all these areas. Consequentially, I&nbsp;developed an awareness of the necessity to become independent and return to the department with my own, different research agenda, which, in my opinion, is an important and correct basis for establishing my own research group.</p><h5><strong>What is the biggest challenge of this project for you?</strong></h5><p>The transition from being a&nbsp;postdoc to a&nbsp;leadership position is a&nbsp;big change for me and I&nbsp;think it is not talked about enough. It means a&nbsp;shift from my own experimental work to leading a&nbsp;team, planning and delegating tasks, and motivating students. From a&nbsp;practical point of view, one of the main challenges is the transfer of reactions from the microscale to functional systems, the construction of reactors, the immobilization of catalysts, and generally confirming that this direction makes sense in the long term and that it has future potential.</p><p><img src="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/VSCHT_Praha_Na_startovaci_grant_jsem_cekala_dva_roky_a_napsala_deset_verzi_projektu_1_66ac267e06.jpg" alt="VŠCHT Prague/Tomáš Balický: I waited two years for a start-up grant and wrote ten versions of my project idea" srcset="https://storage.googleapis.com/labrulez-bucket-strapi-h3hsga3/thumbnail_VSCHT_Praha_Na_startovaci_grant_jsem_cekala_dva_roky_a_napsala_deset_verzi_projektu_1_66ac267e06.jpg 155w," sizes="100vw" width="830" height="839"></p>

I waited two years for a start-up grant and wrote ten versions of my project idea

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The University of Chemistry and Technology, Prague is a natural centre of first-rate study and research in the area of chemistry in Czechia and is one of the country's largest educational and research institutions focused on technical chemistry, chemical and biochemical technologies, material and chemical engineering, food chemistry, and environmental studies.

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<div><p><strong>Discover new perspectives on the complexity of high‑grade serous ovarian cancer in our upcoming webinar, where cutting‑edge spatial multiomics meets real clinical impact. Learn how MALDI Imaging, combined with LMD‑based proteomics, spatial transcriptomics, and MALDI HiPLEX-IHC, is revealing previously unseen cellular subpopulations linked to treatment response and patient survival. This integrated strategy uncovers critical interactions between tumor and microenvironment that may drive chemoresistance and could pave the way for improved patient stratification and more personalized therapy. Join us to explore these groundbreaking results and see how spatial biology is transforming the future of cancer research.</strong></p><h4><strong>Key Learning Objectives</strong></h4><ul><li>Learn how combining multimodal technologies can accelerate advanced biomarker discovery in oncology research</li><li>Understand how MALDI Imaging adds valuable, untargeted spatial context to existing transcriptomics, proteomics, or imaging workflows</li><li>Identify molecular subtypes and chemoresistance‑related interactions using spatial multiomics</li></ul><h4><strong>Who Should Attend</strong></h4><ul><li>Researchers in oncology, spatial biology, and multiomics</li><li>Clinical and translational scientists seeking new approaches for biomarker discovery and therapy prediction.</li><li>Pharma and biotech professionals developing targeted therapies or exploring tumor–microenvironment interactions.</li><li>Biologists working with transcriptomics, single‑cell, or microscopy methods who want to expand their capabilities into spatial and discover what MALDI Imaging can add</li><li>Anyone interested in cutting‑edge imaging mass spectrometry and its applications in cancer research</li></ul><blockquote><p><strong>Presenter:</strong> Dr. Oliver Klein, Group Leader, Berlin Institute of Health at Charité, Berlin, Germany</p><p>Dr. Oliver Klein received his master degree in Pharmaceutical/Chemical Engineering at the Beuth University of Applied Sciences and was subsequently focusing on the expression of protein domains to structure elucidation via NMR and mass spectrometry at the Leibniz Institute for Molecular Pharmacology (FMP) Berlin. Since 2007 he is working at the Charité, leading the Proteomics Unit at Berlin-Brandenburg Center for Regenerative Therapies (today BIH Center for Regenerative Therapies) since 2009. 2015 Dr. Klein earned his doctorate in the field of regenerative medicine, focusing on the detection of pathophysiological processes in tissue following cell therapy using Imaging MS and LC MS. From 2022 until today Dr. Oliver Klein leads the “Imaging mass spectrometry Unit at BIH” and is involved in several clinically driven projects including BMBFTR, DFG, EU and industry cooperations.</p></blockquote></div>

Bruker: Making the Invisible Visible: Unlocking the Potential of Imaging Mass Spectrometry in Ovarian Carcinoma and Beyond

Making the Invisible Visible: Unlocking the Potential of Imaging Mass Spectrometry in Ovarian Carcinoma and Beyond

Spatial multiomics in ovarian cancer showing how MALDI Imaging uncovers tumor heterogeneity, chemoresistance mechanisms, and clinically relevant biomarkers.

Imaging Mass Spectrometry in Ovarian Carcinoma

<p><strong>The microbiology laboratory plays a vital role in selecting antimicrobial susceptibility testing (AST) methods to guide treatment of severe infections in critically ill patients. Delayed or inaccurate results, especially in multi-drug-resistant cases, can lead to poor outcomes and higher mortality.</strong></p><p>In this SelectScience webinar Dr. James A. McKinnell, M.D. an infectious disease specialist, will describe how accurate minimum inhibitory concentration (MIC) data can improve turnaround times, optimize therapeutic decisions, and support the use of new treatments. The session will explore how reliable broth microdilution-based MICs for novel antibiotics enhance patient outcomes while reducing length of stay, adverse effects, and costs.</p><p>Join the webinar to learn how antimicrobial stewardship efforts are most effective when supported by timely diagnostics and collaboration among microbiologists, clinicians, stewardship teams, and policymakers, helping to address the global challenge of antibiotic resistance in the post-Covid era.</p><h4><strong>Key learning objectives:</strong></h4><ul><li>Describe the role of the microbiology laboratory in guiding AST for critically ill patients.</li><li>Recognize the value of accurate MIC data in improving turnaround times, optimizing therapy, and supporting new antibiotic use.</li><li>Identify how antimicrobial stewardship programs rely on timely diagnostic data and collaboration to improve patient outcomes.</li><li>Discuss the importance of multidisciplinary collaboration among microbiologists, clinicians, stewardship teams, and policymakers to combat antibiotic resistance in a post-Covid world.</li></ul><h4><strong>Who should attend?</strong></h4><ul><li>Clinical microbiologists, laboratory technicians, lab managers, lab directors, infectious disease specialists, and infectious disease clinicians.</li></ul><h4><strong>Accreditation statement</strong></h4><p>SelectScience® is approved as a provider of continuing education programs in the clinical laboratory sciences by the ASCLS P.A.C.E.® Program. Attendees of these webinars may be entitled to 1.0 ACCENT Continuing Education Credits from the ADLM. This activity was planned in accordance with ACCENT® Standards and Policies. Verification of Participation certificates are provided to registered participants based on completion of the activity, in its entirety, and the activity evaluation. For questions regarding continuing education, please email education@myadlm.org.</p><blockquote><p><strong>Presenter: </strong>Dr. James A. McKinnell (M.D. and Infectious Disease Specialist, Milefchik-Rand Medical Group, Torrance and Memorial Medical Center, Torrance CA)</p><p>Dr. James A. McKinnell is an infectious disease specialist whose research focuses on healthcare-associated infections, particularly VRE, MRSA, and carbapenem-resistant organisms. He has authored over 40 publications on antibiotic optimization and antimicrobial resistance. Dr. McKinnell works closely with skilled nursing facilities in Los Angeles and Orange County (USA) to develop practical infection prevention and stewardship strategies. He also contributes as an invited speaker for the Infectious Disease Association of California and as a committee member for Los Angeles County’s HAI and antimicrobial resistance initiatives.</p><p><strong>Moderator: </strong>Sarah Thomas (Associate Editor, SelectScience)</p></blockquote>

SelectScience: Critical antimicrobial susceptibility testing results for critical patients: A 2025 perspective

Critical antimicrobial susceptibility testing results for critical patients: A 2025 perspective

Antimicrobial susceptibility testing highlighting role of accurate MIC data in optimizing therapy, stewardship, patient outcomes in critical infections.

Antimicrobial susceptibility testing results

<h3><strong>A five-part webinar series unlocking advanced insights across the Peptide therapeutics workflows</strong></h3><p><strong>Peptide therapeutics are reshaping modern medicine, with GLP-1 receptor agonists driving rapid advances in metabolic disease treatment. As peptide and GLP-1 programs scale, scientists face increasing pressure to generate data that is accurate, defensible, and decision-ready across development stages.</strong></p><p>Peptide 360 is designed to guide scientists through this complexity, following the peptide workflow from sequence insight to confident development decisions.</p><h5><strong>Who Should Attend?</strong></h5><p>This series is designed for scientists and technical leaders working in peptide and GLP-1 discovery and development, analytical development, characterization, Bioanalysis, DMPK, translational research, CMC, QA/QC, and regulatory-facing analytical roles</p><h5><strong>How the Series Works?</strong></h5><ul><li>Register once for the full series</li><li>Pre-submit questions for live expert responses</li><li>Access recordings anytime after each session</li><li>Certificate of attendance provided</li></ul><h4><strong>Episode 2: Designing Integrated Analytical Workflows for GLP-1 and Peptide Therapeutics</strong></h4><p>The rapid expansion of peptide therapeutics, led by GLP-1 agonists, is increasing the need for analytical workflows that extend beyond individual techniques. Successful peptide development depends on designing integrated workflows that connect method development, characterization, and quality control across the lifecycle.</p><p>This session explores how such workflows are enabled through the combination of complementary separation technologies, mass spectrometry, and informatics within Agilent’s integrated analytical ecosystem. Attendees will gain insight into how end-to-end integration, from data generation to data management, supports consistent, compliant, and decision-ready peptide analysis as programs scale from research to manufacturing.</p><blockquote><p><strong>Presenter: </strong>Lauren Skrajewski-Schuler, Ph.D. (Biopharma Workflow Specialist, Agilent Technologies, Inc.)</p><p><strong>Presenter:&nbsp;</strong>Ashish Pargaonkar, Ph.D. (Biopharma Applications Specialist, Agilent Technologies, Inc.)</p></blockquote>

Agilent Technologies: Peptide 360: Mastering the Invisible from Sequence to Confidence

Designing Integrated Analytical Workflows for GLP-1 and Peptide Therapeutics

Peptide 360 Episode 2 webinar on integrated LC-MS and informatics workflows for GLP-1 and peptide therapeutics development and lifecycle control.

Analytical Workflows for GLP-1 and Peptide Therapeutics

<div><p><strong>Join the growing community of Andrew+ laboratory automation users and discover real research, practical insights, and peer‑to‑peer knowledge sharing.</strong></p><h4><strong>WHAT WILL YOU LEARN</strong></h4><ul><li>Real-world case studies from academic labs using Andrew+ today</li><li>Best-practice approaches shared directly by experienced users</li><li>Tips for integrating Andrew+ into your sample preparation workflows</li><li>Ways to automate tasks to increase efficiency and reduce variability</li><li>Opportunities to connect with peers facing similar research challenges</li></ul><h4><strong>WHO SHOULD ATTEND</strong></h4><ul><li>Academic researchers using or evaluating Andrew+</li><li>Lab members involved in sample preparation or workflow optimization</li><li>Anyone looking to introduce automation into their daily lab activities</li></ul><p><strong>REGISTER NOW!</strong></p><p>This event offers a unique chance to hear directly from the academic user community and discover practical insights that you can immediately apply in your own lab.</p><p>You’ll leave with new ideas, shared experiences, and a clearer understanding of how Andrew+ can support and streamline your research.</p><blockquote><p><strong>Presenter:</strong> Alex Dickens, Turku University</p><p><strong>Presenter: </strong>Otto Kauko, Turku University</p><p><strong>Presenter: </strong>Samira Salihovic, Örebro University</p><p><strong>Chairman: </strong>Paul Heads, Nordics Automation Specialist - Waters</p></blockquote></div>

Waters Corporation: Nordics Lab Automation Online Users Meeting

Nordics Lab Automation Online Users Meeting

Join the growing community of Andrew+ laboratory automation users and discover real research, practical insights, and peer‑to‑peer knowledge sharing.

<p><strong>In Part 3, the final webinar of our Single-Cell Proteomics Master Class Series, we turn foundational knowledge and real-world applications into practical, lab-ready strategies. This final session focuses on the critical steps that determine success in single-cell proteomics workflows—sample preparation and high-quality data analysis.</strong></p><p>Attendees will gain actionable guidance from experts, including our partner Cellenion, who will share best practices for achieving reliable, reproducible single-cell sample preparation. You will also explore how to leverage Thermo Scientific™ Proteome Discoverer™ software to extract deeper insights, streamline data interpretation, and enhance confidence in your results.</p><h4><strong>Why attend?</strong></h4><p>Participants will learn how to:</p><ul><li>Apply proven sample preparation practices to increase workflow robustness and data quality.</li><li>Reduce sources of variability and ensure consistency across single-cell experiments.</li><li>Use Proteome Discoverer software to reveal more meaningful biological insights from complex datasets.</li><li>Integrate lessons from all three webinars to build or refine a high-performing single-cell proteomics pipeline.</li></ul><p>You will also have the opportunity to ask the experts your questions during the live Q&amp;A.</p><p>Whether you are refining an established workflow or preparing to scale your single-cell proteomics efforts, this session delivers the practical expertise needed to achieve reliable, high-impact results.</p><blockquote><p><span><strong>Presenter:</strong> </span>Anjali Seth, PhD (Head of Single Cell Proteomics, Cellenion)</p><p>Anjali Seth leads Single Cell Proteomics at Cellenion. She holds a Master’s from ENS Paris and a PhD from Pierre &amp; Marie Curie University on magnetic nanoparticles for drug delivery. After postdoctoral work in Oxford and Lyon, she joined Cellenion in 2019, advancing single-cell proteomics workflows for mass spectrometry.</p><p><span><strong>Presenter:</strong>&nbsp;</span>Philip Loziuk PhD (Product Manager, Chromatography and Mass Spectrometry Division, Thermo Fisher Scientific)</p><p>Phil has spent 8 years at Thermo Fisher Scientific, driving innovation in proteomics. As Product Manager for Proteome Discoverer, he shapes the software’s vision and impact. Previously, he supported scientists globally as a Field Application Scientist and provided expert product support for the Tribrid platform.</p><p><strong>Moderator:</strong> Bernard Delanghe (Scientific Lead, Proteomics Workflow Applications, Thermo Fisher Scientific)</p><p>Bernard Delanghe specializes in single-cell proteomics workflows, bringing together advanced mass spectrometry and data analysis approaches. With over 20 years of experience in mass spectrometry and bioinformatics, Bernard is based in Bremen, Germany and has held multiple leadership roles across proteomics hardware and software. He was recognized with the 2018 European Proteomics Association Industry Award for his contributions to Thermo Scientific™ Proteome Discoverer™ software.</p></blockquote>

Thermo Fisher Scientific: Single-cell proteomics

Optimizing for Impact: Expert Strategies for Single-Cell Preparation and Data Analysis

Single-cell proteomics webinar: practical sample prep strategies, data analysis with Proteome Discoverer and expert tips for robust, reproducible workflows.

Strategies for Single-Cell Preparation and Data Analysis

<p><strong>Long-read sequencing technologies have enabled the sequencing of DNA fragments exceeding 100 kb, significantly advancing the analysis of highly repetitive genomic regions and enabling accurate sequencing of full-length mRNA transcripts—applications that remain challenging for short-read sequencing platforms.</strong></p><p>At the same time, hybridization-based target enrichment using complementary probes has proven to be a highly effective strategy for achieving high-depth sequencing of specific genomic regions while reducing overall sequencing costs.</p><p>In this webinar, Dr. Masahide Seki will introduce newly developed long-read sequencing methods leveraging target enrichment, with a focus on advanced workflows for long-read DNA methylation analysis and full-length cDNA sequencing. These methods are designed to improve data quality, enhance sequencing efficiency, and expand the range of biological applications accessible through long-read technologies.</p><h4><strong>What You Will Learn:</strong></h4><ul><li>Recent advances in long-read sequencing and remaining technical challenges</li><li>Principles and advantages of target enrichment for long-read applications</li><li>Development of long-read–compatible DNA methylation analysis methods</li><li>Target enrichment strategies for full-length cDNA sequencing</li><li>Applications in genomics, epigenomics, and transcriptome research</li></ul><h4><strong>Who Should Attend:</strong></h4><ul><li>Researchers using or evaluating long-read sequencing technologies</li><li>Scientists studying DNA methylation and epigenetic regulation</li><li>Transcriptomics researchers requiring full-length isoform information</li><li>Core facility managers and sequencing specialists</li><li>Bioinformatics and genomics platform teams</li></ul><blockquote><p><strong>Presenter: </strong>Dr. Masahide Seki (Project Associate Professor, The University of Tokyo, Japan)</p><p>Dr. Seki obtained his Ph.D. from the Graduate School of Frontier Sciences, The University of Tokyo, where he researched stem cells using next-generation sequencing in the laboratory of Professors Sumio Sugano and Yutaka Suzuki. He subsequently served as a Project Researcher and Project Assistant Professor in Prof. Yutaka Suzuki's laboratory, contributing to the Genome Support and Advanced Genome Support projects. Currently, as a Project Associate Professor, he is contributing to the Data Science Training and Education Program for Life Science (DSTEP) while developing advanced sequencing methodologies and conducting applied research on diverse biological systems, such as cancer, using state-of-the-art genomic analysis technologies.</p></blockquote>

Agilent Technologies: Development Of Long-Read Sequencing Methods Leveraging Target Enrichment

Development Of Long-Read Sequencing Methods Leveraging Target Enrichment

Webinar on long-read sequencing with target enrichment: DNA methylation analysis, full-length cDNA sequencing and advanced genomics workflows.

Development Of Long-Read Sequencing Methods

<p><strong>Join us for an in-depth webinar on a robust and ultra-fast method for the analysis of trifluoroacetic acid (TFA) in multiple water sources using the SCIEX 7500+ system, together with a Phenomenex Luna Omega column, enabling high-throughput workflows with minimal sample handling. The method, which includes confirmation with both a quantifier and qualifier transition for confirmation, achieves a rapid turnaround time of just 4 minutes and 30 seconds between samples, allowing for the processing of 40 samples—including blanks and standards—in under four hours.</strong></p><h4><strong>What to expect:</strong></h4><ul><li>Deep understanding of TFA’s environmental relevance</li><li>Skills to run validated LC–MS/MS methods for TFA</li><li>Knowledge of instrument setup and troubleshooting</li><li>Experience with calibration, quantification, and QC</li><li>Best practices for sample handling, data interpretation, and reporting</li><li>Awareness of regulatory frameworks driving TFA monitoring</li></ul><h4><strong>Key learning points:</strong></h4><ul><li>Highly sensitive and robust analytical method for TFA analysis in water samples</li><li>Fast, high‑throughput workflow</li><li>How to address TFA contaminations</li><li>Excellent robustness across long sequences</li><li>Reliable performance across complex water matrices</li><li>Regulatory context highlights growing importance of TFA monitoring</li></ul><h4><strong>Who should attend:</strong></h4><ul><li>Environmental chemists</li><li>Water‑quality laboratories</li><li>LC‑MS/MS analysts and technicians</li><li>Regulatory and compliance professionals</li><li>PFAS researchers</li><li>Industrial chemists (chemical, agrochemical, materials)</li><li>Technical marketing and application specialists</li></ul><p><i>We will send you the recording after the event.</i></p><blockquote><p><strong>Speaker:</strong> Carlos Bueno, Senior Application Support Specialist</p><p>Carlos finished his Bachelor Degree in Chemistry in 2001. Before starting working as applications specialist he worked 4 years in Madrid doping control laboratory. Working in SCIEX since 2023 as Senior Application Support Specialist for EMEA. More than 20 years of experience in Chromatography and Mass spectrometry mostly focused on the development of application in food and environmental markets.</p></blockquote>

SCIEX: A fast and ultra-sensitive method for analysis of TFA in multiple water sources

A fast and ultra-sensitive method for analysis of TFA in multiple water sources

Webinar on ultra-fast LC-MS/MS analysis of TFA in water with SCIEX 7500+, covering validation, QC, and regulatory drivers.

Ultra-sensitive method for analysis of TFA in water sources

<p><strong>Join our upcoming webinar on extractables and leachables and explore the complete extractables and leachables workflow using the SCIEX X500R QTOF system. Learn how to achieve precise screening, confident quantitation, and streamlined data interpretation with SCIEX OS software. The session will also cover best practices and practical considerations for implementing robust extractables and leachables workflows in regulated environments.</strong></p><h4><strong>Key highlights:</strong></h4><ul><li>Overview of extractables and leachables</li><li>Precision screening and quantitation with the SCIEX X500R QTOF system</li><li>Streamlined acquisition and data interpretation using SCIEX OS software</li></ul><blockquote><p><strong>Speaker:</strong> Kiran Maan, Application Support Scientist</p></blockquote>

SCIEX: Extractables and leachables: the complete workflow using the SCIEX X500R QTOF system

Extractables and leachables: the complete workflow using the SCIEX X500R QTOF system

Explore the complete extractables and leachables workflow using the SCIEX X500R QTOF system.

Extractables and leachables workflow using SCIEX X500R QTOF

<h4><strong>Key Learning Points</strong></h4><ul><li>Understand why GLP-1 analogues are difficult to purify (structural similarity, truncations, aggregates).</li><li>Learn how to design purification strategies using a single stationary phase.</li><li>Understand regulatory and CMC implications when scaling peptide purification processes.</li></ul><h4><strong>Who Should Attend</strong></h4><ul><li>Any scientists, industry leaders, and researchers driving the future of peptide purification and GLP‑1 therapeutics.</li></ul><blockquote><p><strong>Speaker:</strong> Dhanunjaya Naidu (Business Development Manager, Phenomenex)</p><p>Dhanunjaya Naidu has earned a master’s degree in Organic Chemistry from Andhra University, India and brings 19 years of experience in synthesis, analytical method development and Purifications. In the past 7 years at Phenomenex, he has given many technical presentations, Analytical method development support, Column positioning and troubleshooting assistance. Currently, he is working as a Business Development Manager at Phenomenex India, and responsible for Pan India Purification Technologies Products. He has publications in reputed international journals in organic synthesis.</p></blockquote>

Phenomenex: Purification Strategies for GLP-1 Analogues: Two-Step RP-LC Method Development with Case Studies

Purification Strategies for GLP-1 Analogues: Two-Step RP-LC Method Development with Case Studies

Webinar on GLP-1 peptide purification—strategy design, impurity control, and regulatory/CMC insights for scalable processes.

Purification Strategies for GLP-1 Analogues

<div><div container-id="cq-colctrl-lt0" comp-id="A1"><div><h4><strong>Optimizing ion analysis: Ion chromatography with mass spectrometry</strong></h4><p><strong>Ion Chromatography–Mass Spectrometry (IC-MS) offers powerful analytical capabilities, but optimal sensitivity and long-term performance require thoughtful system setup. This webinar covers practical strategies for optimizing IC-MS, focusing on the components and decisions that most strongly impact routine analysis.</strong></p><h5><strong>Topics that will be covered include:</strong></h5><ul><li>The importance of high-quality eluent</li><li>How the suppressor can affect instrument lifetime</li><li>How to select the right mass spectrometer for your sensitivity needs</li><li>Eluent management strategies to minimize stress on the MS, avoid contamination, and maintain stable performance</li></ul><p>Designed for both new and experienced users, this webinar delivers actionable tips to improve data quality, instrument reliability, and overall workflow efficiency.</p><blockquote><p><strong>Presenter: </strong>Wai-Chi Man (Product Marketing Manager, Thermo Fisher Scientific)</p><p>Wai-Chi has been with Thermo Fisher Scientific for more than eight years working with a wide range of customers in various industries. She provides ion chromatography (IC) support in method development, troubleshooting, sample preparation, training, seminars, events, and workshops. During this time, she has gained deep knowledge, skill, and understanding in areas such as IC-MS, IC-ICP-MS, carbohydrate analysis, metabolomics, and polar pesticides. Prior to joining the company, she worked for Wellcome Research and GlaxoSmithKline. Wai is passionate about promoting STEM to future scientists by encouraging, nurturing, and supporting them.</p><p><strong>Moderator: </strong>David Moore (Senior Product Manager, Ion Chromatography and Sample Preparation, Thermo Fisher Scientific)</p><p>David Moore is a Senior Product Manager at Thermo Fisher Scientific, responsible for the company’s high-end ion chromatography systems portfolio. With more than 20 years of experience in scientific instrumentation, he focuses on translating complex customer needs into practical, high-performance solutions for advanced analytical laboratories worldwide.</p><p>He holds an MChem in Medicinal Chemistry and works closely with global customers, applications teams, and R&amp;D to shape solutions for demanding workflows, including IC-MS and other hyphenated techniques.</p></blockquote></div></div></div>

Thermo Fisher Scientific: Optimizing ion analysis: Ion chromatography with mass spectrometry

Optimizing ion analysis: Ion chromatography with mass spectrometry

Webinar on optimizing IC-MS workflows: learn how eluent quality, suppressors, and MS selection impact sensitivity, stability, and long-term performance.

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