John Chasse

In your paper (1), you state that single-cell inductively coupled plasma mass spectrometry (SC-ICP-MS) is an emerging methodology that holds promise to enable multielemental analysis of low-volume samples in the future. Can you explain why you believe this?

In my view, SC-ICP-MS has become a hot topic in the field of elemental analysis, and rightly so: Scientist working in the field of bioanalytics applied this approach to look into the smallest units of life. As a matter of fact, most biochemical processes take place in cells. Elemental bioanalysis in cells makes it possible to obtain deep insights into cellular processes and the biological status of specific cells. In comparison, the analysis of bioliquids mainly provides insights on a systemic level of the corresponding organism. Over the last decades, many scientists applied ICP-MS to study the elemental composition in all kinds of biofluids and tissues. Due to the latest technical developments it is now possible to achieve ever lower detection limits. Thus, it is now possible to examine individual cells and their elemental composition—at least partially. This may open a completely new field of application for ICP-MS and there are many scientists who are about to prove this.

Why is ICP-MS generally considered being the method of choice for multielemental analysis as compared to other ICP techniques (AES or OES) or more traditional atomic absorption (F-AAS or GF-AAS)?

ICP-MS is unarguably the gold standard for elemental and especially multielemental analysis. This is mainly due to the fact that this technique is extremely versatile compared to ICP-OES/AES and atomic absorption spectroscopy. Most AAS devices apply element-specific radiation sources (hollow cathode lamps) that enable the detection of a single element at the same time. Although there are some commercially available AAS instruments that allow the detection of 10-20 elements simultaneously, sensitivity limitations hamper the analysis of most trace and ultratrace elements. ICP-OES allows multielemental analysis, is relatively easy to setup, and well-suited for high-throughput analysis—however, spectral interferences can be challenging. ICP-MS allows high-throughput, multielemental analysis with the lowest-possible detection limits. It offers a linear detection range of up to 10 orders of magnitude, enables isotopic analysis (isotope dilution analysis for quantification, control of spectral interferences), and it is possible to analyze any type of sample material (gases, liquids, solids).Furthermore, high-resolution separation techniques such as HPLC, GC, CE etc. can be hyphenated to the ICP-MS. Overall, this makes ICP-MS the most powerful and versatile technique for elemental analysis.

What motivated you to focus on this specific application?

Several times, we have encountered situations, where limited sample volume was available. For example, limited volume of biofluids (blood serum or plasma) might be available in human sample cohorts for interdisciplinary clinical studies. In such cases, the principal investigator often selects the panel of analytical tests according to the required sample volume. Sample volume restrictions can also arise from the donor itself, even if there is no competing analysis. This is the case for animal models, especially for studies with mice where only a few µL of blood serum or plasma are available per animal. Since most of the published multielemental profiling methodologies require relatively large sample volumes, our aim was to provide a tool that overcomes this challenge. For this reason, in 2016 we have developed an ICP-MS based method, which allows the analysis of 29 elements in a 150 µL blood serum sample. However, we soon realized that there is a need for multielement analysis in even smaller sample volumes. This is for example the case when only small amounts of serum and sample are available (for example, in precious clinical cohorts or preclinical studies with rodents). We accepted the challenge and found that it is possible to detect 20 to 30 elements in 3–10 µL of biological fluids. These positive results have motivated us to optimize and validate the method. Finally, we have successfully applied the method in a preclinical model. The method turned out to be very easy to use, it does not require expensive or complicated changes in the experimental setup, and, above all, it has shown very good analytical performance. We then decided to publish the results of our study as well as the experimental setup in order to encourage other readers to perform multi-element analyses in the smallest sample volumes in a straightforward way.

What are some cancer-specific multielemental profiles that you have identified? What type of data analysis techniques were used to determine these profiles? Are there a few predominant elements that seem to be most often present in the mice cancer cells?

In the mouse cancer modes under evaluation, the presence of the tumor influences the serum concentration levels of various elements. Some elements, such as sulfur, calcium, manganese, zinc, strontium and molybdenum, were not affected significantly by the presence of the tumor. On the other hand, elements such as phosphorous and copper were increased in the serum samples of tumor bearing mice, independently of the tumor type. Iron, on the other hand, was depleted. These results seem to indicate that the presence of tumor systemically influences the concentration levels of some serum elements, independently from the tumor type. On the contrary, elements such as vanadium, chromium, bromine, rubidium, cesium and cobalt show trends for tumor-specific concentration changes. Furthermore, our results show that different immune T cells (CD4+ and CD8+) have specific mineral profiles reflecting their specific immune function. Moreover, the presence of a tumor seems to have an impact in the mineral contents of T cells. Indeed, we observed elemental modulations within CD8 cells that could be associated with presence of cancer, such as higher levels of magnesium and manganese and lower levels of iron, cobalt and rubidium. These findings might indicate an increased metabolic need of immune cells for Mn and Mg in the presence of a tumor.

What are the biggest challenges that you encounter in this analysis method? Are there any limitations to using this technique? What options or alternatives are available to overcome these challenges?

While the analysis of small amounts of biofluids is quite straightforward (centrifugation to remove particles and a simple dilution step), the analysis of cells is more challenging. In fact, one hurdle we faced during method development was to remove the cell sorting buffer (containing a high concentration of various elements of interest) from the cells. As large volumes of this buffer are required for a single cell-sorting experiment (roughly 10 liters in our case), it was not possible to prepare a buffer solution based on reagents which are suitable for trace elemental analysis. For a washing step, we applied filters or considered osmosis for buffer removal, but none of these sample preparation strategies proved to be suitable. Finally, applying a mild centrifugation step to form a cell pellet, followed by removing the supernatant with a pipette, provided satisfying results. However, the removal of a few microliters from an Eppendorf tube, and at the same time maintaining the integrity of a tiny cell pellet, remains the Achilles heel of the method and requires some finesse and experience.

With such small samples, is contamination an issue, and how have you overcome this potential problem?

Contamination is always a challenge in ultratrace elemental analysis. Acid-washing, working in a clean environment and under a fume hood are the minimum requirements to obtain high quality results. However, contamination is not more challenging in this method, as compared to most other ICP-MS based methods dealing with trace and ultratrace analysis. This is mainly due to the fact that a relatively moderate sample dilution of (1:10) and few sample preparation steps are applied. Contamination could arise, for example, from plastic materials. We mitigate this risk by applying an acid-washing step up front. Another, often underestimated source of contaminants are chemicals and reagents applied during sample preparation. They often contain elements (for example, iron) that enter the sample in this way and thus mix with the analytes. Due to the inherent properties of elemental mass spectrometry, it is then no longer possible to distinguish easily between endogenous and exogenous elements. This can lead to an overestimation of the element concentration. In our study we only apply buffer solutions and nitric acid during sample preparation. Thus, the overall contamination risk is relatively low.

Can you briefly summarize the results that you have achieved so far?

We have developed a quantitative analytical strategy that enables the analysis of 20 biologically relevant elements in low-volume samples (10 µL) of serum. By applying an optimized workflow, the method is also applicable for the quantification of 12 elements in a low number of sorted cells (250,000 cells). In our paper, we give detailed descriptions of the experimental setup. This enables scientist to apply the approach to study biological-relevant elements (major trace and ultratrace elements) in very small sample volumes. Applied to tumor cells, we could demonstrate that the presence of certain cancer types influences differentially the elemental composition of serum concentrations. Furthermore, we could show that different immune T cells (CD4+ and CD8+) have specific mineral profiles reflecting their specific immune function. Last but not least, the presence of a tumor effects the mineral composition of T cells.

Is there anything that you have done in your study that is different from what other researchers have done as they have examined the same problem?

Most of the ICP-MS-based approaches aiming to detect major trace and ultratrace elements in cells are challenged by the media or solvent in which the cells are transported into the plasma ionization source. If the cells are present in Milli-Q water, the “elemental background” is very low. This facilitates the detection of very small quantities of cellular elements. However, cells are prone to lyse in Milli-Q-water. This makes the cellular membranes permeable for ions and small molecules, which are eventually leaking out of the cell and changing the elemental composition of the cell. Fixation strategies, aiming to stabilize cellular membranes, can prevent lysis. Unfortunately, fixation methods tend to increase permeability of the cell membranes and consequently alter the endogenous elemental composition of the cell. Moreover, aggregation of cells is often seen after fixation and needs to be monitored. In fact, the presence of cells in the monodisperse state has to be confirmed by complementary analysis. Many of the reported SC-ICP-MS methods are overcoming this issue by using yeast cells. The use of yeast cells has two advantages: first, the cells are relatively big, meaning higher absolute quantity of elements per cell. Second, these cells are extremely robust compared to immune cells. Re-suspended in Milli-Q water, yeast cells can stay alive for a couple of hours without undergoing lysis. This timespan allows conducting SC-ICP-MS experiments with intact cells while maintaining an extremely low elemental background of the solvent. Suspensions of cells in buffer solutions, on the other hand, allow keeping the cells alive and preserve the integrity of the cell. However, such buffer solutions contain “contaminants”; for example, major or trace elements which are present in the raw materials. This makes it difficult to distinguish between “cellular” elements and elements from the buffer solution. We were able to overcome this limitation by using a sample preparation strategy that includes a washing step with a suitable buffer solution (containing a relatively low elemental background).

What are your next steps in this work? Do you know how well the discoveries of multielemental profiles in mice cancer cells may extrapolate to human cancer cells?

I am now working in a company which is rather focused on elemental analysis is food and pharmaceutical products. However, my former colleagues are continuing our research work and other scientist are working on different approaches to study trace- and ultratrace elements in cells. I am confident that rather sooner than later we will see new findings in this field, both in pre-clinical and clinical studies.

Tobias Konz is currently leading the Elemental Analysis group at UFAG Laboratorien (in Sursee, Switzerland), specializing in quantitative analysis of elements in food and pharmaceutical products (GMP and ISO environment). He studied biomedical chemistry at the Johannes-Gutenberg University (2000-2007), and completed his diploma thesis and PhD in Analytical Chemistry in the research group of Alfredo Sanz-Medel, under the supervision of Maria Montes-Bayón (University of Oviedo, Spain). In 2014, he started a professional career as ICP-MS specialist at the Nestlé Institute of Health Sciences (Lausanne, Switzerland) in the Nestlé Research organization. The main focus of his work has been to develop quantitative methods for multielemental analysis in low volume samples (biofluids) in a GLP and ISO regulated environment.

(1) T. Konz, C. Monnard, M. Rincon Restrepo, J. Laval, F. Sizzano, M. Girotra, G. Dammone, A. Palini, G. Coukos, S. Rezzi, J.-P. Godin, and N. Vannini, 

https://dx.doi.org/10.1021/acs.analchem.9b05643

Articles

Tobias Konz of Nestlé Research, Lausanne, Switzerland and various associates have developed and validated what they describe as a reliable, robust, and easy-to-implement quantitative method for multielemental analysis of low-volume samples. The ICP-MS-based method comprises the analysis of 20 elements (Mg, P, S, K, Ca, V, Cr, Mn, Fe, Co, Cu, Zn, Se, Br, Rb, Sr, Mo, I, Cs, and Ba) in 10 μL of serum and 12 elements (Mg, S, Mn, Fe, Co, Cu, Zn Se, Br, Rb, Mo, and Cs) in less than 250,000 cells, and involved the analysis of elemental profiles of serum and sorted immune T cells derived from naıv̈e and tumor-bearing mice. The results indicate a tumor systemic effect on the elemental profiles of both serum and T cells. Konz and his colleagues believe their approach highlights promising applications of multielemental analysis in precious samples such as rare cell populations or limited volumes of biofluids that could provide a deeper understanding of the essential role of elements as cofactors in biological and pathological processes. Konz spoke to us about this work.

Quantitative Methods for Multielemental Analysis in Low Volume Biofluids 

The Clara Craver Award, awarded annually by the Coblentz Society, was named after Craver in recognition of her pioneering efforts in promoting the practice of infrared vibrational spectroscopy, and recognizes the efforts and contributions of young professional spectroscopists in applied analytical vibrational spectroscopy. The 2019 recipient, Xiaoyun (Shawn) Chen, a senior research scientist working in the Core R&D Analytical Sciences department of the Dow Chemical Company,has been leading Dow’s global optical spectroscopy technology network since 2013, and also the molecular structure capability since 2016, solving a broad range of problems and improving many different types of R&D and production processes, introducing many colleagues to the benefits of spectroscopy for their applications. Chen recently spoke to 

You have over a decade of experience in spectroscopic and chemometric method development for industrial research and development (R&D) and manufacturing projects, focusing on in situ reaction monitoring. What motivated you to focus on this area??

	Let me begin with how I got started. It was probably around my third year at Dow, and I was working on whatever projects that happened to come my way, when my mentor, Anne Leugers, handed me two attenuated total reflectance (ATR) dipper probes, which were no longer needed by another group, and told me to explore whether I could make use of them. By chance, several colleagues in our engineering and process science group reached out to us to monitor a low-temperature reaction that they could not seem to grasp with conventional off-line methods such as gas chromatography (GC), liquid chromatography (LC), or nuclear magnetic resonance (NMR) spectroscopy. We borrowed another Fourier-transform infrared (FT-IR) instrument from our process analytical group, so that we could use the ATR dipper probe to monitor that reaction. In situ monitoring immediately helped us to detect and prove the existence of a highly labile intermediate, which explained why earlier off-line analysis failed. The focus of the project then shifted to characterizing the reaction kinetics for process optimization, and, as a result, we needed to find a way to quantify the concentration of all reaction species. I was very fortunate that Randy Pell, a world-renowned chemometrician, became my mentor in chemometrics, and together we were able to apply several chemometric tools such as classical least squares (CLS) and partial least squares (PLS) to quantify all the reaction species within the complex mixture. This work was later published externally (1)

	So, that was how I got started-as a confluence of instrument and probe availability and project need. I was truly amazed by how useful and enabling in situ spectroscopy turned out to be. The results from in situ infrared (IR) substantially impacted the direction of that particular project and the work of multiple PhD-level process chemists and engineers. As encouraged by my mentors and my leaders, I started to make an intentional effort to further explore and identify similar opportunities. Very quickly, it became apparent to me that the in situ spectroscopy project was not a lucky one-off, but that there were actually many projects that would benefit. Even though people might not know how they could benefit from in situ spectroscopy, I took it upon myself to champion the application of in situ spectroscopy throughout Dow, and even outside. Since then, I just became naturally drawn to that method, and often had to force myself not to be solely focused on in situ spectroscopy. I truly believe that in situ spectroscopy can be a revolutionary tool for process monitoring.

How broad is the range of projects that you work on? Are the projects very similar to each other, or diverse?

	Extremely broad. Let me start with types of projects. Currently, I have several projects dealing with silicone, a few dealing with catalysts ranging from zeolites to MOF, one related to radical polymerization, one related to polyurethane foaming, several related to ethylene polymerization, and several related to customer applications, such as paper coating and airbag coating, among others. In terms of commercialization stages of these projects, they may range from early stage concept-shaping, to process development, to customer applications. There is never a dull moment as far as projects are concerned. The only common theme among them all is that they all can benefit from the use of optical spectroscopy.

What are the biggest challenges you tend to encounter in your work?

	Time management, motivating others, and delegation. There are many things you need to do well as an industrial analytical scientist. First, you need to make sure that you are working on important and relevant research topics. In my current role, I also need to make sure that we keep abreast of the latest analytical instrument development, so that we can solve the toughest analytical problems with the newest tools. We also need to deliver on existing projects, look out for new projects, document past projects, and mentor new people. Whether you can manage your time well among all the above-mentioned tasks is critical in determining how fulfilled, productive, and successful you can be. I forgot to mention that additionally you may also need to maintain an external presence, and I do that by serving as the newsletter editor for the Society for Applied Spectroscopy (SAS), and on the board of the Coblentz Society, and by continuing to publish in peer-reviewed journals. Motivating others and delegation are things that aren’t necessarily taught at graduate school, but become increasingly important as you progress through your career. They are often easier said than done, and I am fortunate to have several great leaders and mentors who have been coaching me throughout this process. I don’t feel I am an expert in them yet, but am trying very hard to be better at them.

You recently published a study about monitoring polyimide production using a combination of in situ infrared and Raman spectroscopy (2). Why did you use both techniques in the study? What was different about what the two techniques enabled you to see? Can the results of that study guide your choice for IR or Raman spectroscopy for future studies?

	The short answer to your first question is “because we can.” I routinely take an IR, Raman, and near-infrared (NIR) spectrometer with me when I visit my colleagues at their labs. Sometimes there’s a clear winner regarding which type of spectroscopy will be the best fit, and we’ll just use that. Just as often, we cannot really predict which technique will work best, and then we try all of them. Which technique is the most appropriate also depends on the focus of the project, which may continue to shift as we make progress.  Often optical spectroscopy may not be the right tool, and then we’ll bring in other analytical tools as well, though that’s probably outside the scope for this interview. You’ll have to read that paper to see our exact findings. For this particular study, I would say both techniques would eventually lead to similar findings, though your perspective into the reaction progress would be very different, due to the different selection rules. Another advantage of using such a broad range of techniques lies in the insights potentially offered by them. There have also been numerous times when we originally set out to solve one problem, but discovered something totally unexpected, which led the whole project to take on a new direction.
	
	

What do you consider to be the most satisfying aspects of mentoring junior spectroscopists, as well as introducing those unfamiliar with spectroscopy to the techniques?
	

As I am starting my 12th year with Dow, I am also starting to take on new roles, and one of the most important ones is to mentor young people coming to our group. I feel very lucky that I have the chance to interact with the brightest minds coming out of graduate school, and I feel especially privileged to see that such bright young people actually take on my advice as they start to navigate their first “real world” job. Seeing their maturing into driven and motivated researcher is the most satisfying part. As for introducing spectroscopy to those outside the field of spectroscopy (which is the case for ~98% of the colleagues I interact with), that’s almost an infinite source of joy for me. I can still vividly remember the first time when my friend Doug Bland told me that he didn’t believe my in situ IR results, because they didn’t line up with the earlier GC results he had been getting for two years, and his subsequent “conversion” and full embracement of in situ spectroscopy. Changing people’s minds and attitudes is often the most challenging part, but definitely the most rewarding part when you manage to do so. I now have many colleagues who no longer treat in situ spectroscopy as the last resort after they cannot use GC or LC, but rather first try to monitor their reactions with IR or  |||||||||||Raman, and only move on to GC or LC |when IR and Raman don’t work sufficiently well (which does happen from time to time). I’m very proud of this.

What do you think are the aspect of working in industry that scientists in academia don’t understand or appreciate?

	I am not sure whether it is true that academic scientists don’t understand or appreciate industrial scientists. Everyone has his or her own focus and perspective. Even within my own group, I don’t think I always understand or appreciate the importance of another fellow spectroscopist’s work. For example, I have fellow spectroscopists who spend all their time in QC labs or QC method development, and colleagues who devote themselves to process analytical, and still others whose focus was on the data management side, such as library, data communication, and chemometrics. I think we all need to put ourselves into others’ shoes to really appreciate each other more. Along the same lines, maybe it’s really the diverse type of work we do in industry that may be something not fully understood or appreciated by outsiders. We also face very different types of pressure in industry. While we usually do not get stressed out by funding or teaching, there are many other things that you have to keep abreast of to do your job well, such as keeping up with the latest technology development, ensuring a robust project pipeline, creating a roadmap to ensure that we have the most relevant capabilities matching the company’s direction, and last, but not the least, meeting the ever-changing customer and market needs.

 28(5), 319-489 (2014). DOI: 10.1002/cem.2507

 71(9), 1-8, (2017). DOI: 10.1177/0003702817700427

Xiaoyun (Shawn) Chen of Dow Chemical solves a range of analytical problems in R&D and production processes using in situ spectroscopy and chemometrics.

Advancing In Situ Applications of Spectroscopy in an Industrial Setting

In recent years, consumers are increasingly opting for natural and organic food. This has led many food producers to switch from using artificial ingredients to using natural ones. In some cases, however, questions arise about the authenticity of those ingredients. Lili He, an associate professor in the Department of Food Science at the University of Massachusetts, Amherst, focuses on developing and applying advanced analytical techniques to solve critical and emerging issues in food science. Recently, that focus has turned to using surface enhanced Raman spectroscopy (SERS) in the detection, analysis, and identification of both natural and artificial food colorants. Dr. He recently spoke to 

You have developed a SERS-based method that is capable of detecting and identifying common food colorants (1). What motivated you to undertake this work?

	Currently in the United States, there are seven main artificial colorants, and about 25 natural colorants, that are approved for use in food. There has been growing interest in the food industry in replacing artificial colorants with natural alternatives, mainly because consumers are increasingly demanding products with natural ingredients. However, the authenticity of colorants used in food products has raised increasing concerns. Natural and artificial coloring agents can be visually identical, so any detection method would need to be able to provide information about the chemical composition of the colorants that are present. We believed that SERS could be an effective method for colorant detection and identification.

Why is SERS a good technique for analyzing food colorants?

	Colorants with conjugated bond systems are inherently Raman active, and given that Raman spectroscopy measures molecular signatures as a basis to identify the components of a mixture, it can be used to differentiate between visually similar colorants with different molecular structures. In SERS, the detection sensitivity of Raman spectroscopy is enhanced by a nanoparticle substrate. In addition, since each colorant produces a unique spectrum, it is feasible to detect multiple colorants in food. Detecting compounds in food matrices with SERS also has the advantage of being simple and rapid, with minimum sample preparation.

What factors did you consider in your choice of SERS substrate?

	In developing the method, we considering the perspective of potential end-users of the method-people who work in the food industry or in food regulation. Therefore, we chose a simple silver nanoparticle substrate that is commercially available, so that the method and database can be easily transferred to the end users, who should not have any difficulty obtaining the substrate.

	We successfully demonstrated the feasibility of SERS to rapidly detect colorant adulteration, as well as to identify colorants in real commercial food products. Also, we have established a database of all the artificial colorants, and most of the natural colorants, and even some banned colorants. We believe that it is important to include banned colorants in the database, because banned colorants pose great safety concerns to public health. Each colorant has a unique SERS pattern based on its molecular vibrations.

How well did the technique perform on actual food products, compared with its use on aqueous solutions of colorants?

	We successfully detected and identified colorants in several real food matrices, including breakfast cereals, candies, crackers, and juices which used artificial or natural colorants. This technique works very well on actual food products, due to its high sensitivity to colorants; there is no significant interference from the background matrices.

	We will further expand and validate the database to cover all FDA-approved food colorants and banned colorants of concern. We will also explore the use of SERS to study interactions between natural colorants and food ingredients that can affect the stability of colorants. In addition, we will carry out a market survey of a variety of commercial food products claiming to contain natural colorants to identify possible cases of adulteration and mislabeling of colorants and to assess the prevalence of such cases.

	We hope that, after successful completion of that study, we will be able to establish a complete SERS spectral database for certified and exempt colorants regulated by the FDA, as well as for selected banned or unapproved colorants. We also hope that we will gain knowledge about the stability of natural colorants in different food matrices by studying interactions between colorants and food ingredients. In addition, we hope to generate information about the current levels and nature of colorant adulteration and misleading labeling in the food market. The developed method and database could be used by industry to assess the quality and authenticity of purchased colorants, as well as to study their degradation kinetics in food and beverage products. Government agencies could also use the methods for rapid screening of any illegal use of food colorants in food products, whether that screening occurs on site or in a laboratory. Information obtained from this project would help industry improve food quality and safety, as well as assist government agencies in making decisions about regulatory considerations and actions regarding food colorants. These actions would lead to a sustainable food colorant market of high quality, reliability, and safety. Therefore, the outcome of this project will benefit the long-term stability of the U.S. food system.

	(1) J.C. Gukowsky, T. Xie, S. Gao, Y. Qu, and L. He, 

 is Associate Professor of Food Science at University of Massachusetts, Amherst. She received her PhD degree from University of Missouri-Columbia in 2009 and did her postdoctorate training in University of Minnesota between 2009 and 2012. Then she joined the faculty in the Food Science Department at the University of Massachusetts, Amherst as an Assistant Professor in 2012. Dr. He's major research focus is to develop and apply the most advanced and innovative analytical techniques to help solve critical and emerging issues in food science. Her group has developed various surface enhanced Raman scattering based techniques for food safety and food chemistry applications. Dr. He has successfully obtained external funding (over $1.8 M as PI) from federal and industry sources, and published 82 papers. Her excellence has been recognized by receipt of the 2012 Young Scientist Award from the International Union of Food Science and Technology, 2015 ACS-AGFD Young Scientist Award, 2016 Young Investigator Award from Eastern Analytical Symposium, 2016 IFT Samuel Cate Prescott Award for Research, and was selected as one of the Talented 12 by 

SERS is a method that is receiving new attention in the detection, analysis, and identification of both natural and artificial food colorants. Lili He, at the University of Massachusetts, Amherst, recently spoke to Spectroscopy about this important analytical work.

Detecting and Identifying Food Colorants with SERS

Spectroscopy can be difficult to carry out outside a controlled laboratory environment. Imagine, then, the hurdles that would accompany performing spectroscopy in the extreme conditions of deep space or the ocean floor. Mike Angel, a professor of chemistry at the University of South Carolina, has taken on those challenges, working on new types of instruments for remote and in- situ laser spectroscopy, with a focus on deep-ocean, planetary, and homeland security applications of deep ultraviolet (UV), Raman and laser-induced breakdown spectroscopy (LIBS), to develop the tools necessary to work within these extreme environments. A key development is the spatial heterodyne Raman spectrometer, a fixed grating Fourier transform Raman spectrometer. For this development and other work, Angel has been awarded the Lester W. Strock Award from The New England section of the Society of Applied Spectroscopy, in recognition of a selected publication of substantive research in, or application of, analytical atomic spectrochemistry in the fields of earth science, life sciences, or stellar and cosmic sciences.

A past awardee of the 2012 and 2018 Society of Applied Spectroscopy William F. Meggers Award (for published work on spatial heterodyne Raman spectroscopy) as well as their 2018 NASLIBS Award (for remote LIBS using spatial heterodyne spectrometer), Angel, who will receive the Strock award this fall at SciX 2019 in Palm Springs, California, took the time to speak to us about his recent work. This interview is part of a series of interviews with the winners of awards that are presented at the SciX conference.

A recent paper you co-authored (1) discusses the initial demonstration of a deep-ultraviolet (UV) 244 nm excitation spatial heterodyne Raman spectrometer (SHRS), and you mentioned that one of the motivations for developing such a device is planetary exploration using a lander or rover. Later in the paper, you discuss the limitations of Raman spectroscopy in such an environment. Would you briefly summarize these limitations, and how this device will overcome them?  

	Raman issues include poor sensitivity, 0.1% in many cases, and luminescence interference. Deep UV excitation (<~250 nm) gets around the fluorescence issue by shifting the Raman spectrum to a wavelength range where most things don’t fluoresce. Using 244 nm excitation, for example; even CH stretch bands are below 265 nm, and most things don’t fluoresce at such short wavelengths. You can also avoid a lot of fluorescence using near-infrared excitation, at say 785 nm, but in this case, the Raman signal can be much lower, because of the smaller Raman cross section at red wavelengths. The Raman scattering efficiency goes as the inverse 4

 Raman band, using 244 nm excitation, has more than two orders of magnitude larger cross section than the same band using 244 nm excitation. This doesn’t mean you will necessarily get 100 times more signal using 244 nm excitation, because there are other considerations, such as sample penetration depth. Deep-UV also offers another signal advantage, resonance enhancement. In this case, resonance can provide another several orders of magnitude larger Raman cross section. So, the potential is there to get much higher sensitivity using deep-UV excitation. Traditionally, UV spectrometers are large, because of the need for very high spectral resolution, which typically requires a high dispersion path. The SHRS technique allows high spectral resolution in the deep-UV, in a relatively small footprint. 

Another paper you co-authored (2) focuses on combining remote LIBS and Raman spatial heterodyne spectrometers to measure a variety of organic and mineral samples, including materials from deep ocean hydrothermal vents, at a distance of 10 m, using a ~100 mm diameter Fresnel lens as the collection optic. What would be the expected discoveries? Who do you feel would be the beneficiaries of this work? 

	The discovery of organisms that can survive in environments like deep-ocean hydrothermal vents has changed our way of thinking about where life can evolve. These kinds of places might be similar to where life got started on Earth, or might have started on other planets. Studying these places is important to understanding ocean chemistry, but also to understanding the extremes in which life can thrive. Hydrothermal vents are believed to have existed once on Mars, and such locations would be good candidates to explore in the search for signs of past life there. Making chemical measurements at deep-ocean vents is hard, because of the depth, and extreme temperature and pressure, and because chemical sensors that can be used under these extreme conditions are rare. The ability to use remote LIBS and Raman to analyze the elemental composition of hydrothermal vent fluids would greatly expand the amount of chemical information that could be obtained in a limited time. 

You recently co-authored a paper on coupling a spatial heterodyne Raman spectrometer (SHRS) with millimeter-sized optics with a standard cell phone camera as a detector for Raman measurements (3). What prompted you to address this development? Would you explain what is unique about this development and its purpose?

	The driver for this study was planetary exploration, to places like the outer Jovian moons, where spacecraft instruments size and weight are at a premium. I wanted to demonstrate that a truly miniature Raman SHRS spectrometer was feasible. It occurred to me that a cell phone already has a high resolution CMOS CCD detector, though uncooled, and a miniature collection lens of very high quality, built in. So, all we had to do to make a miniature Raman spectrometer was mask down the gratings on our existing SHRS, to a couple mm in this case, and add a relatively low power laser. It worked so well in large part because the throughput of the SHRS was very high, and CMOS detector technology is really good and relatively low noise, even uncooled. I found out when doing background research for the paper that a cell-phone Raman spectrometer had not yet been published, so that was fortunate for us.

I’m struck by the range of research work your group is involved with. Are the projects you are working on very similar to each other, or diverse? How do you choose the projects? What are the biggest challenges you tend to encounter when working within these topics?

	To me, all of our work is very closely related. I’ve been working for years to make spectrometers and chemical sensors that can be used to make measurements in hostile environments. Whether it's the deep ocean, the surface of Mars, or a battlefield, the challenges are similar. Most projects come about by talking to people about their measurement needs and problems. It almost always starts with the need. The biggest challenge is long term funding. I have been very lucky to have had long term support from NSF and NASA. 

What does being awarded the Lester W. Strock Award mean to you?

	I feel incredibly honored, especially looking at the list of previous award winners. This award is traditionally for work in atomic spectroscopy, and even though I do a lot of work using LIBS, an atomic spectroscopy technique, I think of myself mostly a molecular spectroscopist. So, I was very surprised by the award, and I am very thankful.

Mike Angel is a Professor of Chemistry at the University of South Carolina where he has held the Fred M. Weissman Palmetto Chair in Chemical Ecology since 2005 and was named a Carolina Trustee Professor in 2013. He received his PhD from North Carolina State University in 1985 and carried out postdoctoral work with Tomas Hirschfeld at Lawrence Livermore National Laboratory. Angel’s research group’s recent work includes developing the spatial heterodyne Raman spectrometer (SHRS), and exploring the SHRS for deep UV Raman, remote Raman, and for use on future planetary landers and SmallSats. Angel is a member of the Mars 2020 SuperCam science team (2014) and is an elected Fellow of AAAS and an SAS Fellow.  He has been a SAS Tour speaker, an A-Page Advisory Panel member for 

 and an editorial advisory board member of 

, and a member of the scientific committee of NASLIBS and the International LIBS conference.  Other honors include the 2018 and 2012 SAS William F. Meggers Award, 2018 NASLIBS/SAS Award, 2016 Elected Fellow of the Society of Applied Spectroscopy (SAS), 2015 Southern Chemist Award, 2014 Science Team member for Mars 2020 SuperCam instrument, 2013 Carolina Trustee Professor, 2012 ACS South Carolina Chemist of the Year Award, 2011 Federation of Analytical Chemistry & Spectroscopy Societies (FACSS) Innovation Technology Award, 2011 

and 2006 Lawrence Livermore National Laboratory Physics and Advanced Technologies Directorate Award.

Spectroscopy can be difficult to carry out outside a controlled laboratory environment. Imagine, then, the hurdles that would accompany performing spectroscopy in the extreme conditions of deep space or the ocean floor. Mike Angel, a professor of chemistry at the University of South Carolina, has taken on those challenges, working on new types of instruments for remote and in- situ laser spectroscopy, with a focus on deep-ocean, planetary, and homeland security applications of deep ultraviolet Raman, and laser-induced breakdown spectroscopy to develop the tools necessary to work within these extreme environments. 

Author | John Chasse

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