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Terpene Analysis Approaches – Part I

Welcome to the newest blog series that totally stinks! In this series we’re going to look at different approaches for analyzing terpenes in cannabis. To quickly brief those who may not be familiar with terpenes, this class of compounds is responsible for many of the flavors and aromas in cannabis. Not only are cannabis growers and manufacturers interested in terpene content though; researchers also have interest in cannabis terpenes because of their potential therapeutic benefits. It is important to classify terpenes in cannabis to better understand the Entourage Effect. This term refers to the synergistic interactions between cannabinoids and terpenes with respect to physical ailments.1

A common approach to analyzing these compounds is by static headspace (HS) – gas chromatography (GC) – mass spectrometry (MS). This can either be done with a HS Autosampler or a gas-tight syringe, but today, we are going to compare this technique to solid phase microextraction (SPME) with the SPME Arrow (new to SPME Arrow? Click here!). For our initial comparison, we diluted our Terpene Standard Mix 1 & 2 (cat# 34-95 & 34096) to 1 µg/mL, added 1 mL of this to a 20 mL HS vial, then analyzed samples via HS-Syringe-GC-MS and HS-SPME-GC-MS. You’ll notice that the parameters are very different for each approach, because we used the optimal parameters for each technique.

 

 

 

Figure. Analysis of 23 component terpene standard.

 

Results from each approach can be seen above. As you can tell from the data collected, both approaches were far from awesome. We were only able to identify 13 of the 23 terpenes in the standard. You’ll notice that we were able to collect data for many of the monoterpenes, but not so much for the sesquiterpenes. For the compounds that were able to be identified; the HS-Syringe performed slightly better than the HS-SPME Arrow. However, the HS-SPME Arrow had better % RSDs than the HS-Syringe. So, why were we only able to identify 13 of the 23 terpenes and how can we improve this? Tune in next time!

 

 

  1. Russo, E. B. (2011), Taming THC: potential cannabis synergy and phytocannabinoid‐terpenoid entourage effects. British Journal of Pharmacology, 163: 1344-1364.

 

Thrive with the New Rxi-65…TG!

Introducing our new and improved GC stationary phase: The Rxi-65TG! This column is essential for analyzing triglycerides in edible oils! You may be asking yourself, why the need for a new 65-type phase? Well, we certainly were not happy with the way our current Rtx-65TG was performing. After redeveloping the polymer and using Rxi technology, we really think we have something that you will enjoy using! Forget the old days of high bleed and poor thermal stability. With the new Rxi-65TG, you will experience run-to-run/column-to-column reproducibility, excellent thermal stability, and super low bleed! Don’t just take my word for it though, check out the data here!

 

Photo Credit: Roberta Sorge

How Can You Make Food Homogenization Less of a Grind?

Sample preparation of a food matrix is only as good as the initial homogenization of your sample.  This determines how representative and effective your extraction will be.  There are multiple ways to homogenize, grind or comminute your sample.  Some involve a simple grind in a food processor.  Other methods require expensive instrumentation to cryogenically freeze your sample.

In the past, when I only used an industrial food processor, samples were often messy to handle, and after freezing, they required thawing before they could be used.  This resulted in needless down time.  After the blog post, A Hoppy Little Story, where I introduced Colton to a new way of homogenizing food samples, I wondered what other foods could be processed this way.  As it turns out, adding dry ice to your sample while grinding, makes every food matrix I tried not only easier to handle initially, but also easier to handle after freezing.

To share this tip with our blog readers, I decided to make a video that brings you into our innovations lab, and shows how we use the dry ice to assist in grinding and what results you get before and after freezing.  With just the addition of dry ice, homogenization, handling and storage becomes so much easier.  I hope the tips in this video are helpful to you too!

Pro EZGC Terpene Libraries Update

Whether you’re doing testing for cannabis, hops, or other food and fragrance analysis, you probably have an interest in terpenes. Now, to help out with that analysis we’ve updated our Pro EZGC libraries with 60 terpenes and terpenoids for the Rxi-17 and Rtx-200 columns.

Compound CAS# Compound CAS#
alpha-Pinene 80-56-8 Menthone 89-80-5
Isovaleric acid 503-74-2 Geraniol 106-24-1
Camphene 79-92-5 Thymol 89-83-8
b-Pinene 127-91-3 Camphor 76-22-2
Sabinene 3387-41-5 Carvacrol 499-75-2
b-Myrcene 123-35-3 α-Cedrene 469-61-4
3-Carene 13466-78-9 (+)-Pulegone 89-82-7
1-phellandrene 4221-98-1 D-Carvone 2244-16-8
alpha-Terpinene 99-86-5 trans-Caryophyllene 87-44-5
D-Limonene 5989-27-5 trans-β-Farnesene 18794-84-8
Ocimene (2 isomers) 29714-87-2 iso-Bornyl acetate 125-12-2
gamma-Terpinene 99-85-4 Citral (2 isomers) 5392-40-5
p-Cymene 99-87-6 Humulene 6753-98-6
Eucalyptol 470-82-6 Piperitone 89-81-6
Terpinolene 586-62-9 Valencene 4630-07-3
Sabinene hydrate trans 15537-55-0 Geranyl acetate 105-87-3
Linalool 78-70-6 (-)-Verbenone 1196-01-6
(1R)-Endo-(+)-Fenchyl Alcohol 2217-02-9 2-Piperidone 675-20-7
Isoborneol 124-76-5 cis-Nerolidol 142-50-7
(-)-Isopulegol 89-79-2 trans-Nerolidol 40716-66-3
1-Borneol 507-70-0 Guaiol 489-86-1
Hexahydrothymol 89-78-1 Cedrol 77-53-2
Fenchone 1195-79-5 α-(-)-Bisabolol 23089-26-1
alpha-Terpineol 98-55-5 β-Eudesmol (2 isomers) 473-15-4
Dihydrocarveol 619-01-2 Farnesol (2 isomers) 4602-84-0
Thujone (2 isomers) 546-80-5 Caryophyllene oxide 1139-30-6
Citronellol 106-22-9 Nootkatone 4674-50-4
Nerol 106-25-2

 


Try it out at https://www.restek.com/proezgc. And if you need standards for your terpene analysis don’t forget our cannabis terpene standards as well.

https://www.restek.com/catalog/view/45361

https://www.restek.com/catalog/view/45362

Can’t find the perfect terpene standard, we’ll make it for you.

https://www.restek.com/Forms/Reference-Standards-Search-Select-and-Custom-Request

 

GC Inlet Liner Selection, Part II: Split Liners

In the previous installment of this blog series, I discussed liner selection for splitless analyses (GC Inlet Liner Selection, Part I: Splitless Liner Selection).  Today I’d like to discuss liners for split analyses.  During a split injection, the split vent is open and the majority of the flow is vented.  The split ratio, set by the user, determines the total flow exiting the split vent vs the total flow going to the column.  Split injections are essentially dilutions occurring within the inlet and require optimization to assure proper sample transfer since the split can affect sensitivity.

Split injections use high inlet flows versus splitless injections. These high flows lead to narrow, sharp peaks and reduced time in the inlet, which is advantageous for thermally labile compounds. At the same time, because of these high flows, choosing a proper inlet liner is essential, since analytes spend such a small amount of time in the inlet, yet must still be thoroughly vaporized and transferred to the column, uniformly.

To test liner performance for split injections of liquids, I set up a similar experiment to my approach in Part I, this time utilizing a split injection method.  Once again, I wanted to compare liners based on recoveries across a wide molecular weight range, as well as reproducibility from injection to injection.  Hydrocarbons ranging from C8 to C40 were evaluated for response, as well as injection to injection reproducibility.

The following liner configurations were compared using the split conditions listed in Table 1 below.

Table 1: Instrument conditions for liner comparisons.

Figure 1 shows how some common liner configurations compare for peak area response across a wide molecular weight range when performing split injections (20:1 split):

Figure 1: Comparison of peak area response across a wide molecular weight range for various liner configurations used in split mode.

As with splitless injections, the use of wool helps to significantly improve response, by providing extra surface area and homogenization of the sample, promoting thorough vaporization and transfer to the analytical column.  Wool also helps to catch the sample, preventing it from going straight to the bottom of the inlet, potentially condensing out or getting swept out of the split vent.  This is especially critical with fast autosampler injections, where the sample is expelled from the syringe so quickly it may not have time to fully evaporate before reaching the bottom of the inlet.  The straight liner with no wool was added to show the drastic effects of performing split injections without some kind of obstruction to catch the sample, mix, and enhance vaporization.  While the cyclo liner was more effective than a straight liner without wool, it did not meet the same performance as the liners that had wool.

Figure 2a shows injection to injection reproducibility for the liners compared under split conditions.  The straight liner with wool, precision liner with wool, and the low pressure drop liner showed the best reproducibility.  The single taper liner with wool had acceptable performance, albeit slightly worse reproducibility.  As expected, the straight liner without wool showed variability due to lack of an obstruction to catch and homogenize the sample.

Figure 2a: Liner injection to injection reproducibility comparison across wide molecular weight range.

Figure 2b shows a zoomed in view of Figure 2a to more clearly see results for the liners with acceptable reproducibility.  Notice the straight liner with wool, precision liner, and low pressure drop liner all performed very similarly.

Figure 2b: Zoomed in view to show performance of liners with best reproducibility.

Conclusions

Based on performance and cost, the straight liner with wool and the Precision liner appear to be the best choices for split analyses.  The straight liner with wool has a cost advantage over the Precision, however, the Precision ensures that the wool cannot move in the liner, causing variability.  Wool movement should generally not be an issue, unless rapid pressure changes are made.  The low pressure drop liner also performed similarly, but is the most expensive of the wool containing liners shown.  A single taper liner with wool will also work, though some slight performance sacrifices are made with regards to reproducibility in split mode.  The cyclo liner unfortunately did not show reproducible or complete vaporization and transfer of compounds compared to the liners with wool.

For those concerned about the use of wool, activity is much less of an issue when using split mode (compared to splitless), as analytes have very little residence time in the liner to interact with active sites.  When using a good inert deactivation, such as Topaz, wool will seldom be an issue in split mode.

You can see from the data on the straight liner (with no wool), that not having some kind of surface to promote mixing and volatilization of analytes results in low responses and very high injection to injection %RSD’s.

Got Fast Columns? Clean Up Your 8081 Samples Using CarboPrep Plus as a Substitute for Florisil

My colleague Linx Waclaski blogged about a tale of two columns that started with the introduction of a pair of stationary phases specifically tuned for the resolution of the 20 legacy pesticides found in US EPA Method 8081(1). The story examined the gains in speed over the years as measured by the last eluting compound decachlorobiphenyl (DCB). When choosing the right columns, conditions, and assisting with oven ramp rates, DCB elutes in under 6 minutes (2,3,4,5).

These fast run times allow more samples in less time which can put additional demands on the analytical system (i.e. more samples more contamination). Unfortunately, its far more likely labs will receive highly contaminated extracts rather than those rare clean samples we commonly see when analyzing drinking water. A standard cleanup procedure uses Florisil which is comprised of synthetic magnesium silicate (US EPA Method 3620C) and is an excellent option for removing polar contamination. High molecular weight and non-polar contamination can make it through the Florisil cartridge and ultimately cause instruments to fail their calibration checks, requiring maintenance.

Restek’s CarboPrep Plus SPE cartridge can be used in place of Florisil for extract clean up, since this carbon removes more matrix interferences that cause instrument calibration failures.  This new product has been specifically designed to directly replace Florisil, using the same glassware, solvents and even elution solvent volumes.  Other carbon material has variability in cleanliness and reproducibility and has not been specifically tested to assure elution of the common 20 chlorinated pesticides including the planar pesticides; for example, hexachlorobenzenes (EPA 8081). Our initial experiments using off-the-shelf carbon produced clean extracts with high background interferences, and low recoveries of planar compounds. Restek’s carbon is manufactured and treated under controlled conditions in an isolated room and packed in foil to assure low background and long-shelf life.

Visually there is a difference in the final color of the extracts after passing through each of the materials. Jason Thomas, who helped with the development and testing of this new product took these photographs illustrating the differences in extract color following cleanup. Figure 1 is the extract before any cleanup is performed. Running the soil extract through the Florisil leaves a colored extract, which will cause active sites in the GC inlet and column (figure 2).  The soil extract eluted through Carboprep Plus is clear, indicating more of the matrix interferences have been removed (figure 3). The same extracted soil sample was passed through Florisil and Carboprep Plus and the analytical results are shown in figure 4. This material has potential advantages in a variety of other applications which we will be exploring in the future.

Figure 1: Soil extract with no cleanup. Dark color is typical of contamination.

Figure 2: Soil extract after cleanup using Florisil cartridge.

Figure 3: Soil extract after cleanup using CarboPrep Plus cartridge.

Figure 4: Comparison of extracts cleaned using CarboPrep Plus and Florisil. Cleaner extracts result in less contamination reaching the inlet and analytical column keeping the instrument running longer (click to enlarge).

  1. https://blog.restek.com/?p=58300
  2. https://blog.restek.com/?p=58822
  3. https://www.restek.com/Technical-Resources/Technical-Library/Environmental/clp7/Seven-EPA-Methods-on-One-Column-Pair-Using-a-micro-ECD
  4. https://blog.restek.com/?p=59100
  5. https://blog.restek.com/?p=60182
  6. https://www.restek.com/Technical-Resources/Technical-Library/Environmental/env_EVSS2976-UNV
  7. https://www.restek.com/catalog/view/53873

Florisil® is a registered trademark  of U.S. Silica Company

My fitting nut has broken off in the end of my LC column. What to do?

If excessive force is applied at an angle to a fitting or column connector, the ferrule could become stuck in the column end fitting. Likewise, a PEEK connector, column nut or plug can also break and leave part of it in the end fitting. Stainless steel nuts are much harder to break, although a stainless steel ferrule could become stuck if severely over-tightened or cross-threaded.

Whether PEEK or stainless steel, if it is a column nut or PEEK connector that is stuck in the column and there is significant portion still sticking out from the column, I would try first to find a tool to grab hold of this part of the nut to see if it can be loosened by unscrewing. You may be able to use a pair of needle nose pliers or pointy-ended vice grips for this. If the nut does not move after a little effort, this is probably not going to work, since the pieces will likely break further. I would move on to try something else.

If it is a PEEK ferrule that is stuck or a nut/connector that has broken off and is still stuck, try using our PEEK fitting extractor tool, catalog # 25325.

 

Below is a complete listing of all of the PEEK connectors, ferrules and similar products which Restek sells that you may be able to remove using the tool above, catalog #25325.  (Note: this is not a guarantee, but our best suggestion. Also, please note that these are all for 10-32, 1/16” fitting size.) Most of our connectors are a design that incorporates a nut and ferrule into one piece. We offer several varieties of these connectors now and also something similar in PPS (polyphenylene sulfide) material. The list also includes a PEEK coupler and a PEEK ferrule for a Secure-fit fitting.

 

Universal PEEK connectors and plug- 25015. 25016, 27710, 27711

 

 

 

PEEK Hex-head Fittings- 27712

 

 

 

 

 

 

 

PPS Hex-Head Fittings- 27714

 

 

 

 

 

 

 

 

 

 

 

PEEK Column Couplers- 27724, 27725, 27726

 

 

 

 

 

 

1/16” (PEEK) Ferrules for Secure-Fit- 20568

 

 

 

 

 

 

 

 

Please note that for the Secure-Fit fittings, the tool can only be used to remove the PEEK ferrule shown above. It cannot be used for the metal ferrule that goes with this design.

 

If you are using a fitting composed of a PEEK nut and a PEEK ferrule that are separate pieces, you can use the same tool for this also. (Restek does not sell this type of fitting, but the tool can still be used.)  Usually it is less likely for the nut to break in these cases, and the tool would mostly be used to remove the ferrule. If the PEEK nut does break and is stuck also, I would suggest trying the tool to remove it.

 

Instructions for PEEK fitting extractor tool, catalog # 25325 can be found on our website here:  http://www.restek.com/pdfs/730-60-001.pdf  and are shown below. Please note that one should never apply excessive force to the tool, as it can break.

Instructions for Use:

  • Drill an opening approximately 1/4″ deep in the center of the PEEK fitting by turning the narrow drill bit in a clockwise direction. To prevent damage to the head of the column, do not drill deeper than 1/4″.
  • Once the opening in the fitting has been created, remove the drill bit.
  • Insert the extraction tool into the opening. The extraction tool has a tapered bit and reverse threading.
  • Turn the extraction tool in a counterclockwise direction, the normal direction used when removing a PEEK fitting. The tool should grab the PEEK fitting and begin to unscrew the fitting from the column

If you are not able to remove the nut or plug by using the tool, we recommend replacing the column. Please note that we do not recommend disassembling a column yourself to attempt another means of fitting removal.  Removing the column end fitting may disturb the packed bed which will likely reduce column performance.

Also, if a stainless steel ferrule is stuck in the column end fitting, it is very unlikely that this can be removed successfully without damaging the column end fitting.  In this case, the best solution is to replace the column. Again, Restek does not recommend disassembling the column and cannot guarantee performance, accept the return of, or replace a column that has been disassembled once it leaves our facility.

 

I hope this post answered some of your questions. Thank you for reading!

 

 

 

TO-15 + PAMS + TO-11A = China’s HJ759 + PAMS + HJ683 Part 3: Formaldehyde Sampling in Air Canisters

In my previous blog (TO-15 + PAMS + TO-11A = China’s HJ759 + PAMS + HJ683 part 2: Deans switching and TO-15/PAMS) I covered the combination of the TO-15 and PAMS (or HJ759 and PAMS) methods into a single run using a Deans Switch and FID/MS detector set up. I said that I’d revisit integrating TO-11A (i.e., HJ683 in China) methods, but first let’s talk about the methods a bit.

TO-11A differs from TO-15 since it involves the capture and derivitization of formaldehyde and other carbonyls on a coated adsorbent tube rather than air canister sampling. The carbonyls react with 2,4-dinitrophenylhydrazine (DNPH) to form a hydrazone derivative in the adsorbent tube. The DNPH derivative is then eluted off of the tube using a solvent and the resulting solution is analyzed by  HPLC/UV. Having two completely different sampling systems and instruments can be a rather large investment in both time and money, so it’s easy to see why there is a push to use canister sampling and GC/MS analysis instead.

So why the separate setup? Formaldehyde is capable of being analyzed directly by GC with no dervitization, although there are some issues with it being low mass and sharing ions with carbon dioxide. See Fig. 1 below illustrating our combined HJ759/PAMS method, using both air and helium as a fill gas for the canister.

Fig. 1 – Formaldehyde in helium (top) and air (bottom), showing interferences from air/CO2 in the bottom chromatogram.

Even if the chromatography can be made acceptable the issue is that formaldehyde is unstable in canisters, creating the risk of false negatives and low response. Canister coating has advanced since the TO methods were originally written though, and one maker of sampling canisters claims they can collect formaldehyde with no losses. If true that would be a game changer for air analysis, so we decided to put it to the test using their canisters. Six canisters were spiked with 100 ppbv of formaldehyde. Three were filled with dry air and three with air at 80% relative humidity to see if water content had any effect. The canisters were then tested every 12 hours for several days and we were not able to replicate their results, with the data showing a quick and drastic drop in formaldehyde response.

 

Fig. 2 – Average formaldehyde loss in dry and humid competitor canisters. N=3 for each data point and error bars show the standard deviation of the replicate canisters.

How bad was the formaldehyde stability? As seen in Fig. 2 in half a day the formaldehyde had dropped to less than 80% of the original amount in the dry canisters, and less than 60% in the humidified ones.  The humidified canisters held steady at near 50% afterwards, but the dry canisters continued to drop until they were less than 40% after 4 days. The water in the humidified canisters likely traps the formaldehyde initially (Day 0 to Day 3) then slowly releases it later (Day 3 on out), causing the initial lower and higher ending results. Given the quick initial drop in what is the simplest scenario of just formaldehyde in air it’s hard to see this as a viable sampling method, even if the samples are rushed to the lab as quick as possible.

While the push for a universal air method is understandable, at this point the sampling techniques don’t support it and it appears that TO-11A will live on as an independent method. And for those of you interested in aldehyde and ketone in air analysis, Restek has you covered with standards and HPLC columns.

https://www.restek.com/chromatogram/view/LC_EV0532

https://www.restek.com/Technical-Resources/Technical-Library/Air-Sampling/env_EVSS2393A-UNV

SPME Fundamentals: Don’t forget the salt for HS VOCs!

Long story short: We were comparing head space (HS)-SPME data with some colleagues, when they asked us “how are we using a 4 minute extraction time on our brewed coffee, when they need 10 minutes to achieve comparable results?” I told you coffee was on the horizon in my last blog. After comparing the 12 or so HS-SPME extraction and desorption parameters from our method and our colleagues’ method, we could not find anything very divergent. Of course, you know where this story goes based on the title of the blog… Our colleagues were not adding salt to their HS samples. In fact, they were surprised to hear that we were doing this.

Why would we add salt to our HS-SPME samples? Long answer: be sure to check out pages 3 and 4 of “A Technical Guide for Static Headspace Analysis Using GC.” Short answer: it does not matter if we are using HS-Syringe or HS-SPME; the addition of salt to the sample matrix (coffee in this example) will often lower the partitioning coefficient (K) for some target analytes, in particular polars. So, for all of the HS-SPME samples we run, we add sodium chloride. How much salt? Several articles have indicated ~20 – 30% wt/wt salt is optimum [1], but of course you should determine the optimum amount for your particular application.

In an attempt to show our colleagues the power of salt, we analyzed the headspace of brewed coffee samples with and without NaCl, all other variables being equal. We added 30% wt/wt NaCl to 10 mL to achieve saturation, which would help ensure consistency across samples. In addition, all samples were incubated (2 min) and extracted (various times) with a shaker speed of 250 rpm (with and without salt); but we threw in a wild card of 1000 rpm. The results of all this may be found in the following figure:

As you saw in the previous teaser, we observed numerous VOCs in the HS of brewed coffee; however, more on this in a future blog. For today, we are looking 2-furanmethanol, which has been shown to be an excellent differentiator between coffee bean roasts [2]. Here are the take-away messages:

  1. As you may see by the red trace, using a shaker speed of 250 rpm on samples with no salt correlates to 2-Furanmethanol (and other VOCs not shown) not reaching equilibrium until perhaps 960 seconds. I say perhaps, because it is hard to say when equilibrium was reached, as we did not extract longer.
  2. You will also see by the green trace, using a shaker speed of 1000 rpm on samples with no salt indicates equilibrium was reached at 480 seconds.
  3. Finally, the purple trace, using a shaker speed of 250 rpm on samples with salt clearly shows equilibrium was achieved at 240 seconds and the overall response was higher than the other two scenarios

Hopefully it is clear why our colleagues were extracting for 10 minutes and still not able to achieve what we observed in 4 min extractions (it was clear to them). It was all about the salt! Now, you could say something like “weighing out salt is time consuming, messy, and the juice is not worth the squeeze.” To which I would say “stay tuned for our Life Hack on Weighing Out Salt for HS Samples in an up-coming blog.” Till next time…

References

  1. S. W. Myung, H. K. Min, S. Kim, M. Kim, J. B. Cho and T. J. Kim, “Determination of amphetamine, methamphetamine and dimethamphetamine in hman urine by solid-phase microextraction (SPME)-gas chromatography/mass spectrometry,” J Chromatogr B Biomed Sci Appl, vol. 716, no. 1-2, pp. 359-65, 1998.
  2. C. I. I. Rodrigues, C. M. Hanson and J. M. F. Nogueira, “Coffees and Industrial Blends Aroma Profile Discrimination According to the Chromatic Value,” Coffee Science, pp. 167-176, 2012.

SPME Arrow for Acrylamide in Potato Chips

I love potato chips and I love the SPME Arrow. So, after talking with my colleague Joe Konschnik about taking advantage of the SPME Arrow to analyze acrylamide and other off-flavor compounds in potato chips; I immediately grabbed a bag of chips from my file cabinet (yes, I keep snacks readily at hand and I already confessed my love) and headed to the lab.

We originally set out to see if we could use the SPME Arrow to analyze acrylamide, as this compound has had a recent resurgence in the media. Acrylamide is a by-product of high temperature (i.e., >120 °C) cooking of specific food products [1]. Acrylamide is formed during the frying, baking, and roasting of starch-rich foods, such as potatoes and grains, in addition, acrylamide is often found in tobacco smoke. Therefore, acrylamide exposure is mostly inhalation and ingestion of certain foods. The International Agency for Research on Cancer (IARC) has classified acrylamide as a Group 2A (probably carcinogenic to humans) compound [2]. Therefore, the European Commission published their Recommendation on the monitoring of acrylamide levels in food [3]; and the US Food and Drug Administration (FDA) published their guidance to minimize human exposures to acrylamide [4].

The following work was carried out as a proof of concept experiment to evaluate the efficacy of utilizing SPME for acrylamide in various matrices. In particular, a headspace (HS)-SPME approach was desired, so as to minimize the sample preparation steps and solvents; and minimize the amount of matrix interference. Of course the SPME Arrow was utilized, as Colton and I have already demonstrated increased sensitivity with the SPME Arrow over traditional SPME fibers. The following figure and table provides the results and details (respectively) of analyzing the HS of ground potato chips.

So, my potato chips did not have a whole lot going on in the HS. In fact, ultimately, we had to fortify them with acrylamide at 0.5 µg/g. As you may have also seen and as expected, the SPME Arrow continues to give us more information over the traditional SPME fiber (no surprise there). Yes, a lot more work has to be done, but we were happy to see we could recover the spiked acrylamide from the chips as a proof of concept. Additionally, we were able to find 2,5-dimethylpyrazine, which is known for it’s off-flavor characteristics.

This story does not end here, as I happen to really love coffee and espresso even more than potato chips (yes, I have this in my office too. It really is quite nice). We have also started to look at ground coffee and brewed coffee headspace with the SPME Arrow as well. I will leave you with the following teaser c-gram generated with our new triple-phase SPME Arrow and brewed coffee HS, which is loaded with compounds. In particular, next time we will look at the furans we found. Till next time…

 

References

  1. WHO, Consultations and Workshops: Health Implications of Acrylamide in Food. 2002.
  2. IARC, IARC Monographs on the Evaluation of the Carcinogenic Risks to Humans, Vol. 60, Some Industrial Chemicals 1994.
  3. EU, COMMISSION RECOMMENDATION on the monitoring of acrylamide levels in food. Official Journal of the European Union. 3.6.2010.
  4. FDA, Guidance for Industry Acrylamide in Foods. 2016.