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Recently one of the designers of the Super-Clean® Gas Filters spent some time with technical service and we were able to ask him some of common questions we get asked by customers. I thought I would share this information with you.
A. The fittings used on the baseplates (and those for the inline filters) are not NPT (National Pipe Thread). He said the NPT threads do not seal well, so they decided to use fittings with parallel threads. If you need replacement fittings for your baseplate, we sell them as custom products (custom numbers are below in bold print). Contact Customer Service for pricing and availability. Note: PTFE tape is not recommended for use on parallel threads.
Restek # 558626 : Brass, for 1/4-inch OD (outside diameter) copper tubing
Restek # 560824 : Brass, for 1/8-inch OD copper tubing
Restek # 556389 : Stainless Steel, for 1/4-inch OD stainless steel tubing
Restek # 560702 : Stainless Steel, for 1/8-inch OD stainless steel tubing
B. Listed maximum flow rates were determined by measuring the outlet flow at the maximum pressure, and not the recommended gas flow through the trap/filter. To achieve the best results, keep the flow rate at or less than 2L/min (liters per minute) for the baseplate filters & inline filters, 8L/min or less for the Big Trap Filters, and 20L/min or less for the LC/MS Filter Systems.
C. A common reason that a new trap will become quickly spent (saturated) is because the check-valve O-rings are cracked/split and are leaking because they are not inspected and/or replaced when the filter is replaced.
You may also want to review What are these O-rings for that I received with my baseplate trap?
D. The lifetime of the trap/filter is largely dependent on the quality of the gas going into it. Keep in mind that a filter/trap does not remove 100% of a particular contaminant, it cleans the gas (generally speaking) an order of magnitude. For example, if the gas quality going in contains 10ppm (parts-per-million) of a particular contaminant, the quality of the gas leaving the filter/trap will contain approximately 1ppm of that contaminant if the gas flow rate is within recommended specifications (see B above).
E. Restek baseplate filters/traps may not work with another manufacturer’s baseplate. In some cases, the metal and/or plastic ring which connects the filter to the baseplate may not have a hole large enough to fit over a Restek filter. To check if our filters will fit, search the Restek website using the competitor’s part number. If found, the trap should fit onto the baseplate. If no search results are found, it likely will not.
To look at this another way, the opening of the Restek gray plastic sealing ring is approximately 38mm inside diameter; some of our competitor’s sealing rings have a smaller ID, and our traps/filters will not fit into a competitor’s sealing ring.
F. All of these traps/filters are designed to be installed right next to the instrument (point-of-use) for best performance. Using one trap/filter for multiple instruments is discouraged because this will increase the number of fittings downstream of the filter which will increase the possibility of leaks.
I hope you have found this information helpful and that this clarifies some of the more common questions we are asked by our customers. Feel free to email firstname.lastname@example.org for any additional questions you may have on these filters/traps.
Dan Li, Rebecca Stevens, and Chris English
Phthalates are a class of compounds that have gained global notoriety. Most investigations focus on determination of phthalates in environmental matrices, baby products or demonstrate the adverse effect on human health. However, the presence of EPA regulated phthalates as impurities in technical phthalate mixtures is underestimated. Regulated phthalate impurities are formed as byproducts of synthesis and are difficult to separate from the technical materials.
The use of non-regulated phthalates should take into consideration regulated phthalate impurities. There are few reports on the analysis of technical-grade phthalate mixtures, indicating that the level of regulated phthalates may be underestimated. In this study, non-regulated technical-grade phthalates from different vendors were investigated using gas chromatography-mass spectrometry (GC-MS). Relatively high levels of regulated phthalates were found as impurities in non-regulated phthalates.
The standard EPA Method 8061A Phthalates Mixture (cat.#: 33227) consists of 15 components, each at a concentration of 1,000 μg/mL. Benzyl benzoate (cat.#: 31847) was used as the internal standard. Technical-grade phthalates (Table 1) were purchased from two vendors.
The GC-MS analysis was performed on a Shimadzu GC-MS QP2010 Plus instrument. An Rtx®-440 capillary column of 30 m x 0.25 mm x 0.25 μm dimension (cat.# 12923) and 3.5 mm Precision Liner w/wool (cat. # 23320.1) were employed for analysis. See Table 2 for experimental details.
Standard solutions were prepared in a concentration range of 1-80 μg/mL (2-100 μg/mL for di-n-octyl phthalate) for calibration curves. The final concentration of the internal standard (benzyl benzoate) in both calibration curves and standards is 20 μg/mL. The technical-grade standards were prepared at 200 μg/mL, except for benzyl 2-ethylhexyl phthalate from Vendor 1, which was prepared at 300 μg/mL to get the impurities in the range of the calibration curve. All standards and tested samples were dissolved and diluted with methylene chloride. During sample preparation plastics were strictly avoided; all preparation work was done using glassware (volumetric flask, syringe, vial, etc).
The Rtx®-440 column (cat.# 12923) provides excellent separation and quantification of the EPA method 8061A phthalate targets (Figure 1). In this mixture, only five components (bis(2-ethylhexyl) phthalate, di-n-butyl phthalate, di-n-hexyl phthalate, dicyclohexyl phthalate and di-n-octyl phthalate) were calibrated since they were present as impurities in technical-grade phthalates. A seven-point internal standard calibration was conducted for each monitored phthalate. The calibration curves were obtained by linear regression of the peak area ratio against concentration ratio of analytes to internal standard. Each sample was injected twice. The linearity (R2) was greater than 0.999 for all five components calibrated (Table 3).
Basically, there are two types of technical-grade phthalates. One type of mixture, such as diisononyl phthalate and diisodecyl phthate, consists of different isomers. The quantification of this type is difficult because the total amount is spread out across many isomer peaks . The other type contains synthetic byproducts that are not removed from main products. For instance, two impurities were detected from hexyl 2-ethylhexyl phthalate, including di-n-hexyl phthalate and bis(2-ethylhexyl) phthalate, which are obviously yielded from synthesis due to the structural relationship (Figure 2).
Among the impurities, only EPA regulated phthalates were measured. The obtained results are shown in Table 1. Impurities levels from two different vendors are consistent with each other except for benzyl 2-ethylhexyl phthalate (highlighted in Table 1), where the concentration is lower for Vendor 1 (1.1%) compared with Vendor 2 (18.1%). This suggests that Vendor 1 and 2 may have a different synthesis process or a different raw material was purchased for this compound. The concentration of regulated phthalate as impurities ranges from 1% to 25%.
The non-regulated technical-grade phthalates are commonly found in industry. Butyl benzyl phthalate, butyl cyclohexyl phthalate and butyl octyl phthalate are widely used as plasticizer, because they are highly compatible or share similar solubility with various polymers, such as polyvinyl acetate, polyvinyl chloride, cellulose nitrate, epoxy, urethane, polyamide, acrylic and so on .
Bis(2-ethylhexyl) phthalate is present in a variety of technical-grade phthalates at relatively high concentrations. Therefore, the actual concentration of bis(2-ethylhexyl) phthalate in plastic products may exceed the regulation limit, even if the manufacturer only uses non-regulated phthalates.
In summary, the quantification of technical-grade phthalate mixtures revealed regulated phthalates as impurities and underscores the importance of this study. The Rtx®-440 column (cat.# 12923) is a good choice for phthalate separation and quantification. Impurity levels in phthalates provided by two different vendors were consistent, except for benzyl 2-ethylhexyl phthalate. The use of these technical mixtures tested in this study for manufacturing of plastics may be cause for concern.
Acknowledgements: The authors would like to thank Shimadzu Corporation for their consultation with the operation of the QP2010 Plus GC-MS Instrument.
 Lin, Z.-P., M. G. Ikonomou, H. Jing, C. Mackintosh, F. A. P. C. Gobas, Determination of phthalate ester congeners and mixtures by LC/ESI-MS in sediments and biota of an urbanized marine inlet. Environ. Sci. Technol. 37, 2100-2108 (2003)
 Lokensgard E, Industrial Plastics: Theory and applications. 5-th Edition, 112-113.
If you have an Agilent 5973, 5975, or 5977 mass spec, there is a simple change to the configuration you can make to optimize its performance. By default, these systems come with a 3 mm drawout plate (or extraction lens) in their source assemblies. According to the manufacturer, this 3 mm lens is ideally suited for trace level (low pg to low ng) analysis. There are two other draw out plate options offered by Agilent, the 6 mm ID and the 9 mm ID. Whether you are using the standard Inert Source or the Extractor source, the draw out plate (or extractor lens) aperture (internal diameter) determines the initial diameter of your ion beam, as shown below (Figure 1).
The areas of the apertures for the 3, 6, and 9 mm ID lenses are 7.07, 28.27, and 63.62 mm² respectively (Figure 2). This extra surface area on the lenses with the smaller apertures can get fairly dirty. Figure 3 shows how the majority of the material buildup on the 3 mm extraction lens occurs inside the 6 mm aperture area. This is important, because low volatility compounds can adsorb onto metal surfaces in the ion source, and after a brief delay, diffuse back into the ion beam. This is most likely the cause of the “source tailing” seen with high molecular PAHs.
Figures 4, 5, and 6 are comparisons of PAH peak shapes when collected with 3, 6, and 9 mm aperture extraction lenses when holding the source temperature, column, and acquisition parameters constant. Figure 4 shows an overlay of m/z 252 acquired on the 30 m x 0.25 mm x 0.10 µm Rxi-PAH column. Benzo[b]fluoranthene is well resolved from the k and j benzofluoranthene isomers, and the peak shapes are similar for all three extractor lenses.Figure 5 shows a significant difference in peak tailing between the 3 mm extractor lens and the other two. While the 9 mm extractor lens is showing a higher response, the peak shapes for the 6 and 9 mm extractor lens runs are still rather symetrical. It isn’t until we look at compounds with a m/z = 50 higher than the benzofluoranthenes that we start to see tailing with the 6 and 9 mm extractor lenses.
Figure 6 is an overlay of three dibenzopyrenes. The 6 and 9 mm extractor lenses show moderate tailing while the 3 mm extractor lens has tailing so bad it is difficult to determine where the final peak ends.
Recommendations from the manufacturer are to use the 3 mm lens for trace analysis applications, the 6 mm lens with purge and trap (e.g. EPA 8260), dynamic headspace, and standard semivolatiles applications (e.g. EPA 8270). The inert 9 mm drawout plate is a custom order, but the 9 mm extractor lens is a standard part (and ships as the standard lens in the PAH analyzer). The first thing I do when I’m operating a new MSD is replace the 3 mm lens with the 6 mm option. That being said, I’ve been experimenting with the 9 mm extractor lens and EPA 8270 using split injection and haven’t encountered any sensitivity or tuning issues (more on that to follow in a different blog).
This post is an extension of a series of posts pertaining to “Red Flags” my colleague has written, pertaining to GC analysis. These are situations and symptoms that tell us in the Tech Service group that something is just not right. So what are examples of some “Red Flags” that we commonly see for HPLC? Below are some of the most common ones that we encounter:
HPLC column high pressure:
Customer reports rapidly increasing pressure with or without a guard column attached. With a guard, the complaint may be that it needs to be replaced frequently.
Our first recommendation will be to isolate and confirm the source of pressure, as described in the blog post Building up pressure on HPLC. First and foremost, it is critical to distinguish between the pressure of the LC system itself versus the columns and guard holders attached.
If using a guard, we always ask if the pressure returns to normal when you remove guard cartridge and/or cap frit inside the holder. If it does, then that confirms that pressure is coming from components of the sample that are being injected. In rare cases, harmful components may also come from mobile phase.
We typically ask what sample prep procedures are used. Samples must be filtered to avoid such issues and if mobile phases contain buffer salts, they must be filtered also. If samples are not filtered with either a syringe filter or Thomson Single Step filter vial, we cannot guarantee the lifetime of a column, guard cartridge, or inlet filter.
If you have purchased a column that can be flushed in a backward direction (3 µm and 5 µm particle size columns), then you may be able to help repair the damage that has occurred to the column. Please follow the instructions located here on our website:
HPLC column degrading peak shape, often with pressure increase:
Customer reports rapidly deteriorating peak shape, usually with increased tailing and broadening peaks, resulting in loss of resolution. Increased pressure is sometimes, but not always reported as well. Shifting retention times are also sometimes observed.
Similar to the previous scenario, if using a guard, we always ask if peak shape returns to normal when the guard cartridge is replaced or when the cartridge and holder are removed. If it does, then that confirms that materials from the sample are likely affecting the packing material. This can be a physical change due to particulates or a chemical change due to fully dissolved sample components (such as in urine matrix). If the initial issue was accompanied by a pressure increase, it is likely a combination of both factors. In either case, please DO continue to use a guard cartridge as it is protecting the analytical column which is much more expensive to replace. Also, please do try replacing the guard cartridge more often. Once a guard cartridge is saturated with chemically active species, it will begin to spill over onto the analytical column. Thus, replacing the cartridge BEFORE you notice a decline in peak shape will protect the analytical column much better.
Again, we typically ask what the sample prep procedures are. Filtration of samples will prevent damage due to particulates, but does not prevent damage from chemically active components in the sample. These must either be captured by a guard cartridge and/or removed by a cleanup procedure. Techniques such as liquid-liquid extraction, Quechers or SPE are all useful tools for this. Selection of the best technique depends on your specific application. The most efficient procedure will be the one with that accomplishes the objective with the most ease and fewer steps.
Please note that although you may be able to determine that sample components are adversely affecting your column, often the damage due to chemical changes is irreversible. Sometimes flushing the column as described above will help and is worth a try. The reason it helps is because of the introduction of solvents with varying polarity, which may remove some bound substances that would not normally be removed with the gradient you usually use.
HPLC column high pressure with Raptor™ 2.7 µm:
Customer reports excessive pressure starting from when the column was first installed on an LC system.
Some HPLC systems have a 400 Bar pressure limit and are not appropriate for use with 2.7 µm particle size Raptor™ columns. This includes some models that are very common in the field, most of which are more than a year or two old. This is discussed in the post: Should I use a 2.7 or 5 µm Raptor™™ column?
If you do not have an LC system with a pressure limit of 600 Bar or higher, we suggest you try using a 5 µm particle size Raptor™ column.
I hope you found this information useful. Thank you for reading!
The range of reference standards available from Restek is very wide, which is great. But finding a particular formulation among all those options can be a challenge. And, let’s face it, filling out a form to inquire about a custom mix can be a pain in the neck.
That’s why we built our new Reference Standards — Search, Select, and Custom Request tool.
We know it’s all about the compounds, so we’ve done a lot to make specifying them easy. The input mechanism verifies compound names and looks up the CAS numbers for you. Better yet, you can paste in whole lists and enter your compounds in bulk.
Once you’ve told us your compounds, our new tool will suggest catalog products that might meet your needs. The matching components in each suggested mix are highlighted in bold. Easily move products back and forth to pick what works best for you.
If you choose to add one or more of the suggested products to your cart, their compounds are filtered out of the list. Any remaining compounds pass through to be requested as a custom. And you always have total freedom to disregard the product suggestions and send our chemists your entire list.
Save yourself time and effort sourcing reference standards from Restek!
As mentioned in my previous post What is Different about Prep LC?, we ask that you start by developing your method on an analytical scale column. Once you have accomplished this, and wish to start using prep scale HPLC for purification, you will need to determine what size preparative scale (“prep”) column would be most appropriate. Below I have presented a possible scenario for selecting a prep column, based on method development done on an analogous analytical scale column. The following is intended for experienced LC users. Please note that these are my thoughts on how this could be done, based on information that is currently available on the topic. Some laboratories that use prep LC on a regular basis may have other practices that work for their specific application.
Column IDs for our prep columns range from 10 -30 mm, whereas the maximum ID for analytical scale is 4.6 mm. The primary consideration for determining the size of a prep column is the desired capacity for sample loading. Columns have a capacity for sample volume and sample analyte mass, so both should be considered. While some instrument manufacturers address sample load in terms of total sample weight, I will be discussing this in terms of total sample volume, since that translates more directly in practical terms. Theoretical sample volume capacity for our prep columns can be anywhere from 40 µL to about 2 mL, but the practical limit should be determined on a case by case basis for each application.
Load Capacity at Analytical Scale
I would start by experimentally determining a practical limit for injection volume on the analytical scale column, with an analytical scale detector in line. For hints on estimating sample injection volumes, please see our FAQs on the subject here: http://www.restek.com/Pages/faq_lc#pkd2. To do this, make a series of injections of a representative sample with volume increasing in increments. (Please see below for an example.) If you are not pleased with results for these injections, try a lower range of volumes. Make sure your representative sample contains the same impurities that you wish to remove from your production scale sample. It will be important also to use a detector that allows you to see both the compound of interest as well as the impurities you wish to remove. Then, come up with a volume that represents an acceptable maximum load volume. Use that number to calculate your scale-up later.
If your prep/production scale sample is expected to be in higher concentrations (in mass of analyte compound per volume units) than you used for the volume test, or perhaps expected to be variable, you should also determine a limit for the mass load at analytical scale. The same technique of doing successive injections in increments will work for this. Examine the peak shape and separation to see what mass of analyte you would be comfortable with loading to get acceptable results for your application. Please keep in mind that practical limits for one analyte may be different from other analytes, based on its unique properties and peak symmetry.
Example at Analytical Scale
The analyst has a 150 x 3.0 mm column, and makes successive injections at 2, 5, 10, 20 and 30 µL.
In this case, 20 µL is determined to be the largest acceptable volume, using a sample at concentration of 15 µg/20 µL = 0.75 µg/µL = 750 µg/mL.
Suppose that the sample given to process at prep scale will vary from time to time. The analyst performs some test injections to determine the maximum mass for the compound of interest per injection. For the 150mm x 3.0 mm column, a series of injections is made containing 10, 15, 30 and 60 µg of analyte at the maximum injection volume of 20 µL. Test injections contain 500, 750, 1500 and 3000 µg/mL, respectively.
Suppose, for this example, that a maximum mass load of 3000 µg/mL is established for the injected sample (60 µg per 20 µL). This information will be used in the example following the next section to help determine the size for a prep column.
Determining Prep Scale Column Dimensions
The next question to ask is “How much sample do you want to be able to load per injection?” As shown in the FAQ mentioned earlier, optimal injection volumes are directly related to cylinder volume of the column. If you are keeping the same column length, the change in optimal injection volume is proportional to the cross-sectional area, A= πR2, where R is the radius. This would be represented mathematically by this equation:
Please note that some sources suggest overall capacity is equally influenced by column length, but opinions on this vary. For our calculation purposes here, we will assume column length stays the same, so length is not a consideration.
Example of Conversion to Prep Scale
Suppose you wish to load a sample volume that is 100 times greater than the maximum volume you determined at analytical scale and your analytical column was 150 mm x 3 mm, then your calculations would look like this:
According to the calculation, you should select a column with radius of 15 mm or larger. A prep column with inner diameter of 30 mm should work for this.
Applying this calculation to the previous example where the maximum volume was 20 µL for a 3 mm ID column, scaling up to a 30 mm ID column would allow you to inject up to 2 mL on the prep column. Suppose that the maximum mass load was 60 µg, or a concentration of 3000 µg/mL for the injected sample. In this case, scaling up to a 30 mm ID column would allow you to inject up to 6000 µg or 6 mg for your compound of interest.
Determining Flow Rate
Another significant difference with prep LC is the flow rate. In order to get the best efficiency for a larger ID column, the flow rate must be increased. As was the case for injection volume, the flow rate also is directly proportional to the cross sectional area of the column, so if you’re converting from analytical scale to prep size, you would multiply the initial (analytical scale) flow rate by the ratio of the radius for the prep column squared over the radius of the analytical column squared. (It is equivalent and sometimes easier to use the diameters rather than the radii.) See formula below:
Where F = flow rate, R = radius and D = diameter. A subscript of “1” indicates analytical scale, while subscript “2” indicates prep scale.
As an example, if your flow rate with a 3 mm ID analytical column was 0.4 mL/minute and you are scaling up to a 30 mm ID column, an equivalent flow rate would be calculated this way:
If you are not certain whether your LC system can accommodate this increased flow rate, please consult your operating manual or contact the manufacturer to ensure it can handle the desired flow rate.
I hope you have found the suggestions in this post useful. Thank you for reading.
Dan Li, Chris English, Jack Cochran, Jason Thomas, and Rebecca Stevens
The alkyl and aryl esters of 1,2-benzenedicarboxylic acid, better known as phthalates, are widely used as plasticizers. Mainly added to polyvinyl chloride (PVC), phthalates make plastic products more durable and flexible. Phthalates are everywhere: plastic toys, PVC water pipes, wallpaper, artificial leather, electrical wire insulation, glue, nail polish, lipsticks, hair spray, plastic water bottles, paints and printing ink are all formulated using phthalate additives. These seemingly endless applications create a big marketplace. Health concerns bubbled up more than a decade ago over their potential to disrupt endocrine signaling .
Aside from health concerns, their ubiquity can cause frustration for chromatographers. Phthalates leached from laboratory consumables, such as rubber tubing, plastic syringes, pipette tips, plastic filters, plastic beakers, plastic stir bars, plastic vials, 96-well plates or other plastic labware can interfere with a lot of chromatographic analysis (both GC and LC). Common lab interference phthalates include diethylhexyl phthalate, diisononyl phthalate, di-n-butyl phthalate (from polytetrafluoroethylene), butyl benzyl phthalate, di-n-octyl phthalate and dimethyl phthalate (from cellulose acetate and Parafilm) . In addition, lab gloves used in sample handling processes are another source for phthalates contamination.
The Rtx-CLPesticides/Rtx-CLPesticides2 GC column set provides unique selectivity and rapid determination of pesticides by GC- μECD (micro Electron Capture Detector) in the low pg range. The advantages of the μECD are clear; low detection of halogenated compounds with minimal interference from hydrocarbons and a variety of other sample matrix. The conjugate electrophore gives phthalate molecule good sensitivity on ECD detector as well. EPA methods 8061A and 606 also recommended ECD for phthalate analysis. Therefore, phthalates can interfere with target compound identification and quantification. This study was designed to determine where the phthalates elute on these pesticide columns relative to the US EPA 8081 chlorinated pesticides.
A significant number of phthalates were observed interfering with pesticides analysis (see the chromatogram and the table). Among contaminants, di-n-butyl phthalate and diethylhexyl phthalate are the most notorious in envirionment due to their low molecular weight, easy partition from polymer matrix, solubility in water, and high usage in polyvinyl chloride production.
Need suggestions to avoid phthalate contaminants? Sometimes avoiding the problem is better than solving the problem. Plastic is the most common source of phthalates. Use glass labware instead of plastic. Although micropipette tips and centrifuge tubes are made from polypropylene, which should be phthalate-free, phthalates can still leach from plastic pipette box, plastic pakage or plastic caps. When using glass, baking-out is a simple way to remove contaminants. In addition, rinsing glassware with redistilled solvents was found to be effective in eliminating phthalates contamination during sample preparation.
Columns: Rtx®-CLPesticides 30 m, 0.32 mm ID, 0.32 μm (cat.# 11141) and Rtx®-CLPesticides2 30 m, 0.32 mm ID, 0.25 μm (cat.# 11324) using Rxi® guard column 5 m, 0.32 mm ID (cat.# 10039) with deactivated universal “Y” Press-Tight® connector (cat.# 20405-261); Sample: Organochlorine pesticide mix AB #2 (cat.# 32292), Pesticide surrogate mix, EPA 8080, 8081 (cat.# 32000); Injection: Inj. Vol.: 2 μL splitless (hold 0.3 min), Liner: Splitless taper (4 mm) (cat.# 20799), Inj. Temp.: 250 °C; Oven: Oven Temp: 120 °C to 200 °C at 45 °C/min to 230 °C at 15 °C/min to 330 °C at 30 °C/min (hold 2 min); Carrier Gas: He; Detector: μ-ECD @ 330 °C; Notes: Instrument was operated in constant flow mode. Linear velocity: 60 cm/sec @ 120 °C. This chromatogram was obtained using an Agilent μ-ECD. To obtain comparable results, you will need to employ a μ-ECD in addition to dual columns connected to a 5-meter guard column using a “Y” Press-Tight® connector. Concentrations are 8-80 ppb for pesticides and 10 ppm for phthalates.
 J. Annamalai, V. Namasivayam. Endocrine disrupting chemicals in the atmosphere: their effects on humans and wildlife. Environ. Int. 76 (2015),78-97.
 A. Reid, C. Brougham, A. Fogarty and J. Roche. An investigation into possible sources of phthalate contamination in the environmental analytical laboratory. Intern. J. Environ. Anal. Chem. 87 (2007) 125-133.
The goal of prep LC is not to produce the best looking chromatogram.
Prep LC can be used to purify a material for manufacturing or research purposes, or it can also be used for sample cleanup prior to analytical measurement, such as in gel permeation chromatography. Most Restek customers are more involved with using it for purification processes, so I will stick to that for this discussion. Products purified by prep LC might include pharmaceutical products, test materials for research or industrial chemicals, to name a few. What surprises some analysts is that the separation criteria are often not as stringent for prep LC as compared to its analytical counterpart. Perfectly symmetric peak shape and baseline resolution are often not required. The goal is not to produce an aesthetically pleasing graph, but to produce a purified product in a way that is overall the most economical and most expedient.
Column eluent is at least partially recovered and used, not sent to waste.
A very big difference in prep LC is that usually the column eluent is separated into timed intervals to make “cuts” of the sample for purification. This is typically done by attaching a fraction collector immediately following the detector and a non-destructive detector, such as UV. Often a switching valve is used to direct flow either to a waste container or to the fraction collector, or the fraction collector itself may contain the switching valve. The system or its components are programmed to begin collecting fractions as soon as analyte peaks become visible on the detector. The collected fractions are usually concentrated/ solvent exchanged to produce a final product or to perhaps to allow analysis by HPLC, GC, or other techniques for an intermediate product. Often multiple aliquots of the same sample are injected repeatedly to purify more of the starting material. Under this scenario, the desired fractions from each chromatographic run would likely be combined to produce the final product. It would be quite common to analyze aliquots from intermediate products or perhaps from individual fractions to monitor progress prior to combining them at the end.
Requires larger column IDs.
The size of the column in prep LC is obviously much larger than analytical LC. For these applications, this is needed to accommodate a larger sample size and also to make using it most cost effective. Column IDs for our prep columns are from 10 -30 mm, whereas the maximum ID for analytical scale is 4.6 mm. Our selection of prep columns can be found here on our website:
Requires method development on analytical scale first.
Since prep HPLC columns are more expensive and have limited availability, we ask that you first develop your method on an analytical column of the same phase. It makes sense to confirm that your compounds of interest can be separated sufficiently before you “graduate” to prep scale. Often analysts will start with a 4.6 mm ID column first. Please do make sure you do this with the exact same column phase to make it worthwhile. In other words, if you plan to use an Ultra C18 prep column, perform your method development on an Ultra C18 analytical column with an ID of 4.6 mm or smaller.
Requires specialized hardware.
Using a prep LC column brings with it certain requirements in terms of LC system hardware. For that reason, many analysts will use a system specifically designed for prep chromatography. Here are some examples of components that would be different:
- Solvent pumps capable of higher flow rates, up to 150 mL/min
- Larger sample loops
- Higher pressure rating valves and components
- Detectors with prep size flow cells
For example, here are links to information on systems available from Shimadzu and Waters:
I hope this post has helped to give you a better idea of what is involved with prep LC and how it is different. Please stay tuned for the next post, where we will discuss scaling up from analytical scale to prep scale LC. Thank you for reading!
My third installment of Technical Service Red Flags contain six more situations that cause us to pay attention and ask questions. I hope you find these helpful.
Using a liquid leak detector to locate gas leaks instead of using an electronic leak detector. Or worse yet, using both at the same time. We have, unfortunately, communicated with many customers who have accidently “sucked” liquids into their electronic leak detector. If this happens, turn off the leak detector immediately. We suggest allowing it to “dry” in a desiccator or bag of rice for several days. If it no longer works (or it does not work well), contact Customer Service or your distributor and order 22655-R (Leak Detector Repair Service).
Leaving your GC oven temperature at or just above ambient temperature overnight with carrier gas flowing. Moisture condensation can become an issue.
Trying to analyze light gases and/or water vapor using a GC/MS. When NOT to use (GC) Mass Spec
Using our products for applications other than those they were designed for. While I am sure many of our products have multiple uses, we are a chromatography company and our products were designed for our chromatography customers. We will unlikely know if our product will work for your non-chromatography application, and we cannot be held responsible if it does not work. In summary, use at your own risk.
Using a TCD with helium as the carrier gas to quantitate hydrogen. Experiments have shown that obtaining a linear calibration curve can be difficult, if not impossible.
Trying to use our Restek ProFLOW flow meter (# 22656) to measure negative (vacuum) gas flows. This item is only designed to measure flows of clean, dry, non-corrosive gases under positive pressure. If you need to use a flow meter for vacuum gas flows, select one of our Alicat M-Series Flow Calibrators
To read my previous Red Flags posts, the links are below. Thanks.
Derivatization is a widely-used technique for GC sample preparation across many industries and in widely varied matrices from soil to plastics to blood that is used to make polar and active compounds more amenable to good GC analysis. If you’re careful about testing your derivatization procedure during method development, you can be confident that you’ll have a reproducible method that can vastly improve the quality of your GC results. While derivatization does require some extra sample handling, the procedure I developed for cannabis plant matrix is very straightforward and easy to perform:
Derivatization Procedure for Cannabis and Hemp Plant Matrices:
- Place 100µL of plant extract into a 1mL Micro-Vial
- Evaporate to dryness at 50°C under a gentle stream of nitrogen
- Add 50µL ethyl acetate and 50µL BSTFA + 1% TMCS
- Incubate at 70°C for 30 minutes
- Cool and dilute with ethyl acetate if desired
In my last blog, I introduced the concept of derivatization for use in cannabis or hemp cannabinoid testing. Once acidic cannabinoids are derivatized, they no longer break down in the GC inlet and can be quantified separately from the neutral cannabinoids. I demonstrated this through derivatization of high-level solvent standards, but work with solvent standards is a far cry from matrix work, which means the procedure needed to be tested in matrix. To kick off the matrix test, I spiked an extract with the most common cannabinoids of interest, derivatized it using the procedure listed above, and my colleague, Jack Cochran, analyzed it via GC-FID with our Rxi-35Sil MS GC column. We can see that we have a beautiful chromatogram with all of the derivatized cannabinoids separated, and very little matrix interference.
In addition to confirming that all derivatization sites are indeed derivatized by analyzing the standards with GC-MS (this is shown in my last blog), we also tested derivatization efficiency using a cannabis extract previously generated at Penn State University with the help of Professor Frank Dorman and a Police Officer Specialist. Because derivatization is a chemical reaction, the derivatization reagent gets used up during the derivatization reaction. Because plant matrix contains many other derivatizable compounds like sugars and sterols, these other compounds may compete for the derivatizing reagent, possibly resulting in the reagent getting used up before all of our analytes of interest can be derivatized.
So how can we be sure our derivatization is going to completion in the presence of matrix? There are a couple things we can do, the first of which is really simple. We can see in our procedure that we use a hefty 50µL of derivatizing reagent per 100µL of cannabis extract. We know that our extract contains a lot less than 50mg of plant matrix, not all of which is derivatizable. This means that by adding 50mg of BSTFA per 100µL of sample, we can be confident that we have a significant excess of derivatizing reagent as compared to derivatizable groups in our sample. Excess derivatizing reagent means that it will never be completely used up, ensuring the reaction will go to completion no matter what.
A more quantitative way to test derivatization efficiency in a matrix where you can’t get blanks is to evaluate analyte linearity with differing amounts of matrix. For example, if you derivatize four THCA-containing samples prepared using 10, 20, 50, and 100µL of cannabis extract and plot the area of THCA versus sample amount, you should end up with a straight line if your derivatization is going to completion. If it’s not, then you’ll likely see THCA area fall off for the samples containing more matrix since the derivatization reagent is being used up before all the analyte in the higher matrix level sample is derivatized. To test our procedure, we did just that. We can see that our linearity looks beautiful for all of the cannabinoids, indicating the derivatization does indeed go to completion.
In addition to verifying that the derivatization reaction goes to completion in the presence of plant matrix, we also verified the procedure using several different samples which were generated at the same time as the sample shown in the figure above. Our preliminary work is still looking good, which is exciting, but what about all of the other matrices cannabis chemists have to work with? Well, we’re planning on moving the work forward into edible matrices next, so stay tuned for an update!