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Preserve your SRM transition windows by replacing the same amount of column you remove during maintenance

I’ve been working on a detailed PCB congener analysis using the TSQ-9000 with the AEI. Typically this analysis is done by EPA Method 1668C, which uses high resolution MS. We get some specificity from the selected reaction monitoring, but there are some shortcomings when compared to high resolution data. Where the TSQ-9000 really shines is the sensitivity. We were able to achieve very linear calibrations (avg RF RSD% < 5.5%) for all target compounds using 6 levels (analyzed in triplicate) ranging from 40 ppt to 400 ppb. The 6 point calibration suggested by EPA Method 1668C ranges from 200 ppb (for sensitive instruments) to 2000 ppb.

After 6 weeks of constant running, it was time to perform some maintenance. We needed a new liner and septum, and we needed to trim the column to restore symmetrical peak shapes to the analytes. The SRM method we’re using has well over 500 transitions, many are close together with narrow windows, and the prospect of shifting multiple compounds out of their windows was alarming. Figure 1 shows the pentachlorobiphenyl homologue group. The comprehensive chromatogram would have some congeners from the 3-chloro, 4-chloro, 6-chloro, and 7-chloro homologue groups overlapping, as well as stable isotope labeled internal standards and surrogates.

Figure 1 – The complete pentachlorobiphenyl homologue group. Window defining and WHO toxic congeners are labeled with their ID in blue.

 

It’s possible to use the speed factor calculated by the EZGC method translator to calculate the new retention times, but it’s a manual process involving measuring a deadtime and calculating a new effective length, translating the method, and manually calculating and updating the new transition windows.

The more prudent course of action seemed to be removing the need to update the transition windows. Replacing the segment of column removed with the same length of the same column should result in virtually no change in elution times, eliminating the need to update the transition table. This is exactly what we saw when we replaced 1 loop of the 0.18 mm x 0.18 µm Rxi-PCB column with a fresh segment of the very same length, using a SilTite µ-Union to make the column connection. The before and after chromatograms for the WHO toxic congeners in the EPA 1668C calibration CCV are figures 2 and 3 respectively.

Figure 2 – Retention times for select toxic and window defining PCB congeners straddling the center third of the chromatogram

 

Figure 3 – Retention times for the same select toxic and window defining PCB congeners straddling the center third of the chromatogram following the replacement of the head of the analytical column with a fresh segment of Rtx-PCB.

Aaron Lamb from Thermo Fisher and I will be discussing persistent organic pollutant analysis this afternoon using the TSQ-9000. Following us, there will be a discussion on the alternative method of analysis – High resolution MS, specifically the Orbitrap. The webinar is titled ‘ELEMENTary Environmental Analysis: How to Overcome Interferences to Simplify the ICP-OES and ICP-MS Analyses of Challenging Samples From Wastewater to Electronic Waste’. Click the link if you are interested. The webinar should be available for on demand viewing within a few days. There are many other interesting on demand lectures to pick from.

 

The Foundation of Separation is Preparation

Restek is known as a leader in the field of chromatographic column technology.  The idea is built into our very identity – resolution technology is at the heart of Restek.  We also believe that proper sample preparation sets chromatography up for success, though, by reducing the incidence of confounding results and prolonging the lifetime of your chromatographic investments.  From dilute-and-shoot techniques to multi-step solid phase extraction methods, Restek is just as dedicated to helping you find the best preparation method and product for your sample.

While it is unlikely that we will change our name, we are nevertheless committed to providing analytical solutions to today’s toughest chromatographic challenges, from preparation through separation.  Check out Restek’s growing line of sample preparation products, and let us know how we can help.

 

 

Getting Your LC Up and Running Again

Welcome Back!  You and your lab have been through a lot these last few months and now it’s time to get your LC back up and running.  After it has been down for a few months, it likely needs a little TLC before it’s performing optimally again.  Just like a car that sits around for a long time without being used needs its fluids, filters, and tires checked and maintained, an LC has similar needs.

Let’s walk through the system and get you back up and running.  Before starting anything, you should consider performing preventative maintenance (PM) according to your manufacturer’s specifications.  Another excellent resource that can help is “Routine LC Maintenance: Simple Steps to Preventing Unexpected Downtime.”  After any PM or other maintenance that you decide to do is completed, you can move on to getting the rest of the system ready.

If there are old solvents sitting on the solvent rack, dispose of them all in the proper waste streams and replace them with fresh solvents.  This includes solvents for seal washes and needle washes etc. ALL solvents should be replaced.  At this point you should also replace your column with a union and divert your flow stream directly to waste, making sure the detector is disconnected from the flow path. You should also note right now whether any buffer was in the system.  If so, you’ll need to flush all pumps and lines containing buffer with water to prevent any salt precipitation in the system.  Your pumps will need to be primed and purged according to the LC manufacturer’s specifications. After this, start the pumps with low flow rates (0.2-0.4 mL/min) and purge any buffer from the lines with water for 10-20 minutes.  Also, be sure to rotate any valves in the system that might contain buffer.

Next, flush a strong organic solvent, like isopropyl alcohol, through the system by priming and purging the pumps.  Flushing the system for 10-20 minutes at 0.2-0.4 mL per minute should adequately flush the system.  Finally, you can load your desired mobile phase solvents onto your LC, though you should not incorporate buffers yet.  After all of these steps are completed, all buffers and organic contaminants should be flushed from your LC.

Now you are ready to reconnect the detector into the flow path.  Follow the same order of solvent flushing as above.  Start with aqueous and transition to organic.  With your entire system now flushed, you can begin running diagnostics.

At this point, installing an LC column will help you to identify any problems that might exist as many can show up when there is back pressure on the system.  Observing not only your baseline, but also your back pressure can help you determine if anything needs to be fixed or replaced.

Erratic baselines can indicate the presence of air bubbles.  To fix this, further purging of your pumps or entire system can fix this problem.  If you see systematic fluctuations in pressure profile or detector, this can be more indicative of a pump problem.  The culprit is likely a faulty check valve or a worn pump seal that requires cleaning or replacement.  For all these PM tasks or repairs, make sure to follow your LC system manufacturer’s guidelines.  Many standard parts are available on our website.

If everything checks out, then a diagnostic/system suitability tests with a column in place and an appropriate system suitability mix should be performed to see how well the instrument is performing.  Comparing the current results with previous tests can also be useful to make sure your instrument is performing optimally.

Good luck on getting your systems back up and running and if you have any questions, we are always here and happy to help.

 

Other Useful Resources:

LC Column Cleaning and Regeneration

Connecting Your Column

Preventing LC Column Clogs

LC Troubleshooting-Retention Time Shift

LC Troubleshooting-Baseline Problems

How can analyte protectants and matrix help improve peak shapes?

In my last blog, I presented a new technique called low pressure gas chromatography (LPGC, Figure 1). Just to recap, the LPGC system consists of a relatively short analytical column (10 – 15 m) with large ID and thick film (e.g. 0.53 mm and 1.0 µm, respectively) which is restricted with a narrow guard column (e.g. 5 m x 0.18 mm). The restrictor (guard column) allows for maintaining head pressure on the inlet, while the analytical column is under near-vacuum pressure.

Figure 1: LPGC schematics


During my analysis, I’ve run into issues with early eluting peaks. At the initial 80 °C starting temperature the first two compounds, methamidophos and dichlorvos, showed up as distorted, split peaks. I’ve found that the optimal initial temperature was 70 °C, but the question remains: what if the sample is in the matrix? Or what if we use analyte protectants?
I’ve decided to investigate so I compared the original (solvent) analysis of the QuEChERS Performance Mix (#31152 ) to the analysis of added analyte protectant (0.1 mg/mL shikimic acid) and finally compared those to the celery matrix (Figure 2).

Figure 2: Comparison of pesticide residues’ runs with no matrix or analyte protectant (black trace), celery matrix (red trace) and with analyte protectant (green trace)


Figure 2 shows that the matrix (red trace) distorts the peak shape even further at both tested temperatures. Celery isn’t a very “dirty” matrix (after using dSPE), therefore, it doesn’t act as an analyte protectant for these compounds. On the other hand, the analyte protectant (green trace) helps significantly with the peak shape at both 80°C and 70 °C. At 80 °C, the effect is more pronounced while at 70 °C it helps reduce the tail of methamidophos and narrows the dichlorvos peak.
In conclusion, the analyte protectant (shikimic acid) can help with the peak shape at the original temperature, however, it is still preferable to lower the initial temperature to achieve a good solvent trapping.

The New U.S. EPA Method TO-15A Blog Series-Part 5: Humidification

In the previous blog (https://blog.restek.com/the-new-u-s-epa-method-to-15a-blog-series-part4-clean-lines-for-clean-air/) I left off with our system at or under the 20pptv limit required by the new TO-15A method after replacing and moving our air fill line. Once you’re sure of your air quality and the cleanliness of your lines the next step is filling and humidifying your canisters, and if you’ve done water analysis you may be seeing the issue already. The TO-15A analyte list shares a lot of overlap with common volatile contaminants in water, so now that we’ve got our lab air clean it’s time to work on our lab water.

Before I get into what I did here at Restek, let me start with a story. Years ago, I used to work at a lab doing VOC testing in water (methods 8260, 624, and 524), and our quest for clean water was very similar to what we went through to get clean air here. When I started, the DI water system was next to the sample prep lab and was stored in a large tank to keep enough on hand to supply the entire lab. That meant that after being cleaned by the DI system the water sat around and the solvent fumes from the prep lab caused contamination. It was fine for use in the inorganic and semivolatile labs, but to get the blank levels low enough for the VOC lab we had to boil the water down to ¼ of its original volume. Eventually the lab was remodeled and the VOC lab got its own DI system. They also put the VOC lab under positive pressure to keep solvent vapors out from neighboring labs. We could finally stop boiling our water like we were living in the wilderness. We solved our issues by isolating our water from solvent sources, just like I had to with the air here at Restek.

Fortunately for me, Restek does not have a large prep lab going through many liters of solvent per day. Unfortunately, our lab is shared between volatile GC, semivolatile GC, and LC instruments, so I expected some level of contamination to be present. If your lab does VOC testing in water it’s possible you already have good baseline data for your water. If not, then humidifying a canister you know is relatively clean (i.e., used for standards and blanks only, not for field samples, and has been blank tested with dry air) can be a way to evaluate the quality of your water. Using the canister RH calculator that Jason Herrington posted on this blog years ago (https://blog.restek.com/to-15-canister-relative-humidity-part-ii-examples-and-calculations/) I humidified some canisters to 50% RH and tested them. In the same way as the last blog, testing was done on the full TO-15A/NJLL list, but for the sake of simplicity I’m only showing the problematic compounds. The results shown below compare the final results (seven days after initial fill and humidification) for the dry air and humidified air canisters.

Compound Dry (pptv) Humid (pptv)
n-Pentane ND 24
Ethanol ND ND
Acetonitrile 16 25
Carbon disulfide ND ND
Isopropyl alcohol ND ND
Methylene chloride 20 35
Acetone ND 38
Hexane ND 103
Tertiary butanol ND ND
Tetrahydrofuran (THF) 10 889
2-Butanone (MEK) ND 25
Toluene ND 11
4-Methyl-2-2pentanone (MIBK) ND ND

Table 1: Dry vs. Humid air results in pptv

As expected, a number of common solvents are now present. For the most part the levels were lower than I feared, but I was surprised at the high levels of THF as it is not heavily used in our LC applications. However, it is also used as a solvent for resins so it could be coming from our DI system resin cartridges.
Going back to my previous experience, I decided to try boiling the water to see if I could get the results down below 20 pptv. I took some of the DI water and boiled it on a hot plate down to ½ of the original volume, then used it to humidify canisters to 50% RH. Below the table shows the results of the original and boiled water.

Compound Original (pptv) Boiled (pptv)
n-Pentane 24 35
Ethanol ND ND
Acetonitrile 25 24
Carbon disulfide ND ND
Isopropyl alcohol ND ND
Methylene chloride 35 28
Acetone 38 20
Hexane 103 198
Tertiary butanol ND ND
Tetrahydrofuran (THF) 889 39
2-Butanone (MEK) 25 31
Toluene 11 10
4-Methyl-2-2pentanone (MIBK) ND ND

Table 2: Original vs. Boiled water results for humid air in pptv

The boiled water showed great improvement for THF, but mixed results otherwise. Hexane, pentane, and MEK actually seemed to be concentrated during the boiling. Why didn’t it work this time, when I’ve had luck with this technique in the past? Before I was boiling the water further away from the source of solvent contamination, so it’s possible that by boiling it in the same lab as I did here just isn’t practical at low levels. There are a few other things I could try, such as boiling the water in another location or purchasing DI water from a chemical supplier, but those aren’t guaranteed to work and may not be practical for a lot of labs. There’s also the possibility that some of the compounds have always been present in the canisters and they are being displaced from the canister walls by the water vapor, which would mean further chasing clean water would be a red herring.

Despite the handful of compounds above 20 pptv I’m actually rather happy with the result. 20 pptv is a very low limit that I think many labs will have trouble with. It’s likely that in a shared laboratory setting like ours (with ubiquitous solvent use) some level of blank contamination is inevitable, so how can this be handled? Stay tuned for the next blog in the series, where I’ll cover calibrations and how at low levels it’s easy to end up with high biases.

Modifying QuEChERS for complicated matrices- High Sugar & Starch Samples

In previous blog posts for this series on QuEChERS, we discussed QuECHERS dSPE Selection, Modifying QuECHERS..-Dry Samples and Modifying QuEChERS..-High Fat Samples

Once again, I suggest to use the following from the official QuEChERS website (QuEChERS.com), maintained by CVUA Stuttgart, these specific documents:

For Extraction (Stage 1): https://www.QuEChERS.com/pdf/reality.pdf

For dSPE Cleanup (Stage 2): https://www.QuEChERS.com/pdf/cleanup.pdf

References may be made to information from a table that indicates dSPE Primary and Secondary Actions, which originated in previous posts from this series. You may find it useful as you read the discussion below.

 

High Sugar and Starch Samples

Samples containing high amounts of carbohydrates may present unique challenges for an analysis. Molecular size of these interfering molecules can vary considerably, ranging from small monosaccharides like glucose to very large polysaccharides like starches. Although size and some of their properties vary, they are all composed of repeating units of one or more monosaccharides. Medium size polysaccharides (3-10 sugar units), also called oligosaccharides, can exist as glycans, where they are linked to a lipid, amino acid, or protein. The largest polysaccharides (>10 sugar units), include starch and cellulose, which more closely resemble a polymer because they are very long chains of repeating monosaccharide units. All of these molecules have the potential to wreak havoc with GC and LC analyses for various reasons. They may produce visible chromatographic interference and often they can also inflict damage to LC and GC systems by accumulating residue in injection ports, valves, tubing and sometimes in columns. To reduce the potential of instrument downtime and to reduce the cost of column replacement, it is always best to eliminate these types of materials from sample matrices as much as possible. Here are some techniques that have been used with Quechers to accomplish this.

 

dSPE with PSA/MgSO4 and possibly C18-EC – This is a demonstration of the classic QuEChERS technique, where the analyst would choose MgSO4 and PSA to remove the simple sugar molecules. PSA is indeed powerful in its interaction with small sugar molecules due to its ion exchange properties. However, it is limited in its interaction with larger and more complex molecules containing sugar, since they are less reactive, less polar and sometimes have little or no solubility in water. In the table shown above, the interaction with longer chain carbohydrate molecules and starches is noted as a secondary action for the C18-EC. Consequently, when more complex sugar molecules and starches are present, C18-EC sorbent can be the perfect complement to PSA to remove this matrix. For some sugary matrices that are dry, such as honey, adjustments to water content made in Stage 1 extraction also help to make the dSPE step go more smoothly. Here are some examples of successful use of these techniques. (Please note that some of these also use GCB, but its target for removal is pigment. Also, a quick reminder to use caution with GCB for samples containing planar compounds.)

 

Cooling/Freezing the QuEChERS extract– Similar to what we discussed in the previous blog post about high fat samples, the same approach mentioned on the QuEChERS website (https://www.QuEChERS.com/pdf/cleanup.pdf) can sometimes be used to help remove sugar-related or  starchy sample matrix.  Unfortunately, I was unable to find any examples where this was done successfully, other than the Quechers site itself.  I would be very interested to hear some readers share their experiences in this regard.

 

SPE cartridge (cSPE) cleanup– In some cases, the amount of sugars or carbohydrates is too high to efficiently remove with the above mentioned techniques.  Using cartridge SPE gives the analyst an option to use more sorbent to more effectively remove this interference. Sometimes alternate sorbents such as silica or florisil may be used as well.

Here are some examples of this technique.

 

Matrix matched standards– Sometimes despite extensive measures to remove matrix from a sample, there may be enough residue remaining to affect the analysis. This is often the case with more sensitive analytes, such daminozide. In order to mimic the same effect for calibration standards, they can be prepared to contain the same sample matrix.

Here are some examples of this technique.

Additional Resources:

 

Thanks for reading our discussion of high sugar and starch sample matrices. Please feel free to add your comments and share your experiences. We look forward to further discussion.  Also, please look for the next post in this series on soil samples.

 

FDA Issues Second Warning on Methanol Based Hand Sanitizers

The U.S. Food and Drug Administration has placed methanol containing hand sanitizers on an import alert (1). These products do not list this ingredient and some are incorrectly labeled “FDA approved.” Currently, 87 hand sanitizers have been found to contain methanol (2). Methanol (methyl alcohol Cas# 67-56-1), known as wood alcohol, is commonly found in solvents, and windshield washer fluid (3). Ingestion of 10 mL can cause permanent blindness and 15 mL is considered a lethal dose (4,5). Transdermal methanol poisoning has been well documented and may result in optic nerve necrosis with permanent eye damage (6,7,8,9,10,11). Denatured alcohol is ethanol mixed with other alcohols and can contain 50% methanol. Other additives include isopropyl alcohol, acetone, methyl ethyl ketone, ethyl acetate and methyl isobutyl ketone.

Figure 1: Pro EZGC Chromatogram Model of a denatured alcohol sample illustrating the ability of this column / conditions to determine percent levels of methanol in samples, specifically hand sanitizers. Measured retention times are also indicated on the far-right column as a comparison to the predicted retention times.

 

This is the first part in a series addressing the different aspects of hand sanitizers. Our first goal is to determine a suitable column that can resolve methanol from water and ethanol. We accessed the safety data sheet (SDS) for our denatured alcohol sample and entered those compounds into our Pro EZGC Chromatogram Modeler (12). The program provides 5 different solutions / columns. Our column choice will be discussed in future blogs. Figure 1 is a model of the compounds with the expected retention time compared to the measured retention time using a mass spectrometer. While the model suggests resolution of the compounds, the sample has been diluted 50:1 in distilled water and the solvent peak width is not calculated by Pro EZGC.

Figure 2: Denatured alcohol sample analyzed using a Rtx-VMS by GC/MS with an adjusted scan range to accommodate water and methanol detection. This method is suitable for the analysis of hand sanitizers containing methanol.

We followed USP 611 Method II (13) as a starting point. The MS scan range started at m/z 10 to detect water (m/z 18) and methanol (m/z 31). We also changed the GC program from the conditions suggested by Pro EZGC by adding a 3-minute hold time for better solvent focusing and slowed the initial ramp rate down to 3°C a minute. Future work will rely on our method translator to determine an equivalent solution using a Flame Ionization Detector (FID). Figure 2 is a total ion chromatogram (TIC) showing good resolution between water and methanol. One way to optimize the resolution of early eluting compounds from the solvent peak in the split mode is the use of the 4.0mm ID Precision Inlet Liner w/ Wool, based on Linx Waclaski’s work (14). Using a rinse solvent other than water could result in extraneous peaks, ghost peaks and carryover. The challenge with using 100% water as a rinse solvent is the potential that residue will build up in the syringe barrel. My colleague Corby Hilliard developed a method that incorporated a cosolvent to prevent syringe damage (15). In this case we used a prewash with 100% water and three post washes with 90% water and 10% n-propanol. This method is suitable for measuring percent levels of methanol in samples. Our next blog will present gel and liquid hand sanitizers using this method.

  1. https://www.accessdata.fda.gov/cms_ia/importalert_1166.html
  2. https://www.fda.gov/news-events/press-announcements/coronavirus-covid-19-update-fda-reiterates-warning-about-dangerous-alcohol-based-hand-sanitizers
  3. https://www.vercounty.org/MSDS/SDS-EMA/88-RainEx%20Deicer%20Washer%20Fluid_SDS.pdf
  4. https://www.sciencedirect.com/science/article/pii/S0379073818303037
  5. https://www.msdsonline.com/2014/07/22/methanol-safety-tips-from-msds-experts/
  6. https://pubmed.ncbi.nlm.nih.gov/19628396/
  7. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3617539/
  8. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5292112/
  9. https://pubmed.ncbi.nlm.nih.gov/26196361/
  10. https://pubmed.ncbi.nlm.nih.gov/29908646/
  11. https://www.sciencedirect.com/science/article/pii/S2173579420300530
  12. http://archpdfs.lps.org/Chemicals/Denatured-Alcohol_Ace.pdf
  13. http://www.uspbpep.com/usp29/v29240/usp29nf24s0_c611.html
  14. https://blog.restek.com/gc-inlet-liner-selection-part-ii-split-liners/
  15. http://www.chromatographyonline.com/injecting-water-gc-column-solving-mystery-poor-chromatography-0?pageID=3

LPGC – Fast way to your pesticide analysis!

Throughput is one of the most important parameters in the lab. The more samples we can analyze in a day, the sooner we can get home. Enter Low Pressure GC (LPGC) – this is an invention from our brilliant Jaap de Zeeuw [1-2], where a relatively short analytical column (10 – 15 m) with large ID and thick film (e.g. 0.53 mm and 1.0 µm, respectively) where the flow is restricted with a narrow guard column (e.g. 5 m x 0.18 mm). The restrictor (guard column) allows a normal head pressure at the inlet, while the analytical column is operated under near-vacuum conditions. The low pressure inside the 0.53mm column, shifts the optimum linear velocity about a factor 7 higher, which allows for faster analysis without a total loss in efficiency. The wider ID and thicker film provides also higher capacity, robustness and inertness. In addition, an integrated transfer line adds additional robustness to the method as the absence of phase in the heated transfer line to the MS, helps to reduce background and make the system stabilize faster. Figure 1 shows the LPGC system schematics.

Figure 1: LPGC schematics

Read the rest of this entry »

Water, can’t live without it so how can we deal with it (and adsorbent columns)

Often the gas samples contain some water vapor. Although we are usually not interested in the amount of moisture present in the sample, if we don’t dry it, the water will be injected onto the column with our sample. Alan, in his blog, has previously discussed the effects of water on capillary columns with liquid stationary phases and adsorbent (PLOT) columns. But as they say: a picture is worth a thousand words, so I want to share a few examples I gathered while working with adsorbent columns and the impact of  water/water vapor.

Molecular Sieve 5A

Molecular Sieves 5A are zeolites that are part of the family of hydrated aluminosilicate minerals. They are known adsorbents with a unique, tunnel-like crystalline structure and a well-defined pore size. The analytes separate on molecular sieves based on two adsorption mechanisms. First, how well the molecules fit into the pores of the material, a separation based on the size of the molecules. Second, the physical interactions between the molecules and the MSieve 5A crystal, a separation based on the polarity.  For example, nitrogen and oxygen are both small enough to fit in the pores of 5A mineral, but oxygen, a smaller molecule than nitrogen, will navigate through the “tunnels” faster because it has less interactions with the pore surface and elutes from the column before nitrogen. In the case of carbon monoxide, polarity plays a more important role and the MSieve 5A strongly retains these molecules. Adsorption on molecular sieves is reversible and the adsorption/desorption process is easily regulated with the temperature.
We know that we can’t analyze water using the MSieve 5A columns because of the high temperature required to desorb polar water from the pores of the molecular sieve. What happens if we are injecting water onto the column? How much water can we inject before it’s “too much”? How does “too much” look like? Can we regenerate the MSieve 5A column? And how long does it take to regenerate it to its original performance?

To find the answers, I have performed a quick study where I injected water onto a Rt-MSieve 5A column, 30m x 0.53mm x 50µm (Cat#19723) in between the injections of permanent gases. I kept the system at 40°C the entire time, simulating isothermal analysis of gases. Right after 1µl injection of water I noticed carbon monoxide’s retention time shifted, eluting earlier (Figure 1).

 

Figure 1: Overlay of chromatograms of permanent gases, black overlay – initial analysis, blue chromatogram – analysis after injecting 1µl of water onto the column.

I finally stopped injecting water after carbon monoxide’s capacity factor (k’) was at half of its original value. Until then, 100µl of water was injected onto the column under the described conditions. Analysis of permanent gases at that point shows that adsorption sites are packed with water, analytes start to tail indicating sample loading capacity is decreased, and gases elute faster or even co-elute (Figure 2, Chromatogram B). 100µl of water on the column may not seem like that much until we put it into perspective. Let’s say our sample has 50% relative humidity. With every 1ml injection of gas, we are introducing ~0.01µl of water onto the column. That means, to lose 50% of retention for carbon monoxide we can make 10,000 injections.

 

Figure 2: Analysis of permanent gases (in order of elution: Argon, Oxygen, Nitrogen, Methane, Carbon Monoxide, concentration 3-5mol%, 100µl split 60:1 injection)), 40°C Isothermal, flow He 4 ml/min Chromatogram A: Analysis using a new column, Chromatogram B: Analysis of permanent gases after 200 injections of 0.5µl water at 40°C Isothermal, Chromatogram C: Analysis of gases after fast 20 min conditioning, Chromatogram D: After 2h conditioning, the water was removed from the column.

To regenerate the column the water can be desorbed by conditioning at the column’s maximum temperature, 300°C.  I wanted to track the time required for the column to recover by monitoring the capacity factor of carbon monoxide every 20 min.  After 20 min of conditioning, the water “distributed” through the column, and the peak shape of the analytes improved (Figure 2, Chromatogram C). The original column performance was restored after 2 h of conditioning (Figure 2, Chromatogram D and Chart Figure 3).

 

Figure 3: Graph of water desorption process relative to the k’ of carbon monoxide. The orange plot is the capacity factor of CO at the beginning, and the blue plot is the capacity factor of CO after conditioning at 300°C relative to the time.

Continuous injections of water on the MSieve column without elevating the temperature of the analysis will affect the chromatography, resulting in peak tailing, loss of retention, and resolution. Just remember, even when 50% of the column performance is lost the column can be restored to the original performance by conditioning it at the maximum temperature.

Porous Polymer Columns (Rt-Q, QS, S and U BOND)

Water does not affect the porous polymers and will elute from the column as a peak. Depending on the polarity of the polymer water will be more or less adsorbed and elution time will change accordingly. Figure 4 are chromatograms of permanent and hydrocarbon gases containing water vapor. On the nonpolar divinylbenzene Rt-Q BOND column, water elutes faster, and naturally, water is more retained on the most polar column, Rt-U- BOND. Bear in mind that water will not be detected with the FID detector.

 

Figure 4: Analysis of permanent and hydrocarbon gases on 30m x 0.53 mm x 20µm Rt-Q and Rt-U BOND column (Carrier gas: He@5ml/min, Oven: 40°C (3min) then 10°C/min to 190°C, Detector: TCD)

ShinCarbon Column

ShinCarbon column can adsorb a very limited amount of water. The amount of adsorbed water is dependent on the column dimensions (amount of the material in the column). The adsorbed water has no impact on the retention times of the compounds. However, eventually, if the water is not conditioned out of the column or if the injected concentration of the water is high, it will show up on the chromatogram as an overloaded peak with an almost “never-ending” tail and interfere with the integration of the analytes. Water will always show as “overloaded”. Therefore, depending on the injected concentration retention time of the water peak will shift.

 

Figure 5: Overlay of three chromatograms on a 2m x 1mm 100/120mesh ShinCarbon column– 30µg (green), 80µg (red) of water injection, and permanent gas with C1-C2 hydrocarbon standard (black) – showing where the water will elute from the column. (Carrier gas: He@10ml/min, Oven: 40°C (2min) then 20°C/min to 200°C, Detector: TCD)

Moisture present in the carrier gas will over time have a similar effect on the column performance. Mainly MSieve and ShinCarbon columns act as a carrier gas scrubber, trapping the moisture present in the carrier gas (Figure 6). Note that these materials are hygroscopic and will also attract water when the column is not sealed upon storage.

 

Figure 6: Overlay of the first and second instrument blank injection after the instrument has set idle with the ShinCarbon column installed at 40°C for 48 h. (I think it was time to change my carrier gas filter.)

Don’t forget, carrier gas filters, which will remove the moisture and other impurities, are essential accessory with this type of analysis.

Thermo Trace 1310 Inlet Temperature Profile vs Agilent 7890 for Split/Splitless Injectors

Several years ago, my colleague Scott Grossman wrote an excellent article entitled “It’s a Matter of Degrees, but Do Degrees Really Matter?”  He measured the temperature profile across various Agilent inlets, demonstrating different gradients in temperature exist across inlets, depending on the type of inlet and even among the same inlet type.  One consistent finding is the top and bottom of the inlet are always cooler than the middle. The middle is closest to the actual temperature setpoint.  Depending on the inlet, some had more significant drops at the top or bottom vs others and this may affect chromatography.

I wanted to repeat this experiment on a Thermo Trace 1310 split/splitless inlet to see how it compares to an Agilent 7890 split/splitless inlet.  Please note that this was not to prove that one is better or worse, but rather to understand differences. This information will help when making decisions about inlet temperature setpoints, ultimately affecting things like vaporization potential, compound degradation, septa temperature, etc.  This is especially important in a lab operating both instruments, running high molecular weight compounds.  The fix in this case could be as simple as running one instrument at a slightly higher inlet temperature, since a difference in temperature at the bottom of the inlet will affect performance.

To obtain this data, I took a thermocouple probe and inserted it through the septum until it reached the bottom of the inlet.  After recording this temperature, I would mark and pull up the thermocouple probe in 5 mm increments, recording the temperature at each location, until I reached the top of the inlet.  I plotted the inlet temperature gradient from bottom to top for each inlet (see below).  For these experiments, I used an inlet temperature setpoint of 250 ⁰C, as this is a relatively common setting.  I also checked the Thermo inlet profile at 300 and found the same relative percent error at each inlet location compared to the 250-degree experiments.

Here’s what I found:

Comparison of inlet temperature profiles of Thermo Trace 1310 split/splitless inlet with Agilent 7890 split/splitless inlet.

What you may immediately notice is that the top of the Thermo Trace 1310 inlet is significantly cooler than Agilent’s 7890 inlet.  There is a drastic drop in temperature over the top 20 mm.  This should not affect your compound volatilization, since the sample is injected towards the center, but it will affect the temperature of the septum.  For the Thermo Trace 1300/1310 inlet, I recommend Thermolite Plus septa, which are softer than BTO septa.  BTO septa are ideal for high temperatures and will be less pliable at lower temperatures and also more susceptible to coring.  Because of the lower temperature of the septum, bleed should not be a major issue, either.

You may also notice that the bottom of the Thermo inlet is slightly cooler than the Agilent inlet.  This could affect the vaporization of analytes.  If you are using both instruments in your lab, operate the Thermo Trace 1300/1310 at a higher inlet temperature compared to the Agilent to get comparable results.  This would be most evident with response of compounds with high boiling points.

As you can see, different instruments may have different temperature gradients across their inlets.  This does not make one instrument superior to another, but can affect performance and lead to perceived differences between the instruments, if operated under the same conditions.  Also keep in mind that this data was acquired on just one inlet from each manufacturer.  It’s possible for variation to exist between inlets from the same manufacturer, as Scott Grossman demonstrated in his work.