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Rxi-624Sil MS Pro-EZGC Library: 591 Compounds to Choose From

Five years ago Chris English showed in his blog post “I Can’t Drive 55” — The Pure Power of EZGC all 233 compounds that were in the Rxi-624Sil library. While that is a lot, we didn’t want to stop there. This weekend we added over 350 new compounds with various functionalities – e.g. alcohols, aldehydes (first time on EZGC!), amines, esters and more!

Just to show you the power of EZGC, here are few real chromatograms with their modeled counterparts. Let’s start with the hot topic of the day, cannabis terpenes:

The second example is one of the completely new compound sets in the EZGC library – aldehydes:

And last (but not least) the EPA 8260 method:

Volatiles by EPA Method 8260 on Rxi-624Sil MS (30m, 0.25mm ID, 1.40µm)
http://www.restek.com/chromatogram/view/GC_EV1169/GC_EV1169

This huge update was made possible in collaboration with Becca Stevens, Amanda Rigdon, and Megan Burger.

A better way to configure your EZ No-Vent GC-MS Connector Part II

Last time, I wrote a blog (here) that showed a better way to configure the EZ No-Vent in the software so that the column length didn’t need to be manipulated. I kept the volumetric flow the same, and showed different ways to minimize the negative chromatographic effects of the reduced linear velocity. Today, I’m going to show you what happens when you keep the linear velocity the same after installing the EZ No-Vent.

Figure 1: 8270 Chromatogram on a 30 m x 0.25 mm x 0.25 µm Rxi-5Sil MS (cat# 13623) by Split Injection

 

Figure 2 – 30 m column configured as a composite column in MassHunter

 

Figure 3: 8270 Chromatogram on a 30 m x 0.25 mm x 0.25 µm Rxi-5Sil MS (cat# 13623) by Split Injection with the carrier gas linear velocity matched to that of the chromatogram with no EZ No-Vent installed.

 

Figure 4: 30 m column configured as a composite column in MassHunter. Note the volumetric flow and average linear velocity.

As you can see, the tailing that plagued the EZ No-Vent chromatograms in the previous blog (here) has been eliminated by increasing the speed of the analytes through the column. When a GC-MS is run without an EZ No-Vent (Figures 1 & 2), the vacuum extends quite a long way into the analytical column, causing an increase in linear velocity as analytes approach the end of the column. When the EZ No-Vent is installed, the 100 µm restrictor greatly reduces this effect, causing an overall drop in average linear velocity under the same volumetric flow, reducing efficiency. Restoring the original average linear velocity appears to be the solution to the negative chromatographic effects (Figures 3 & 4).

Dilute, Shoot, and Elute – am I missing anything? YES!

 

Everyone’s lab is different in terms of how many samples per day are processed, but they all share the common pain point of sample preparation. Some samples like blood and plasma need a significant amount of prep to remove proteins, phospholipids, and salts, whereas labs running urine or drinking water samples can “get by” with a 5x or greater dilution.

No matter how limited or extensive your sample prep, the one thing that is critical to prolonging the lifetime of both your column and instrument is particulate removal, and you know what that means: filtering.

We’ve blogged about filtering mobile phase before, and recently gave you a behind-the-scenes tour of column construction in the Clog Blog to emphasize how important it is to remove particulates from samples. You also want to keep your HPLC or UHPLC up and running, and the downtime plus parts and labor expense for replacing any or all of the sample needle, injection port, valve rotor, and outlet tubing of your autosampler is far greater than sample filtration supplies.

I like to use “the paint example” when talking about sample prep. Chances are you or a colleague have done some home remodeling that includes painting, so you know you don’t just go get the color you like, roll it on, and you’re done. For best results, you have to fill in any holes or scratches, sand, tape, prime, and finish with your carefully chosen color from the selection of 500 shades of beige. Sure it takes extra time, but it turns out looking great. It’s the same with HPLC sample prep: the more care you take with filtering, the longer your column and instrument will last. My colleague and frequent blogger Nancy from Tech Services has a great post about making your HPLC columns last longer and filtering is high on her list too!

The easiest method for manual sample filtration is to use a Thomson Filter Vial. Anyone who knows me will tell you that this is one of my favorite products EVER. There are different membranes depending on whether your sample has a high aqueous or high organic content, and the 0.2µm membrane is ideal for small ID UHPLC tubing, which is typically 0.1mm ID or less and is prone to clogging. There’s even an eXtreme version of the Thomson vial that has a multi-layer filter for samples with 10-30% particulates. Both vial types are very simple to use with a lot less mess, hassle, and waste compared to a separate syringe, disc filter, and collection vial. Here’s an example using a 0.2µm PVDF filter vial for the analysis of fentanyl in urine.

Another effective way of removing particulates is through centrifugation. After a protein crash or dilution, you can place your vials or multi-well plate into a centrifuge and spin away to pull particulates into a pellet at the bottom of the vial or well. This analysis of immunosuppressive drugs used a precipitation solution vortexed with whole blood, then 4,300 rpm in the centrifuge to remove particulates.

After filtering or centrifugation you can put your vial or plate into the autosampler for analysis. Make sure you adjust the needle setting so it stops 3-5mm above the pellet so it doesn’t suck up any of the particles you just pulled out of the sample solution!

A simple filtration or centrifugation step will allow you to make the most of dilute-and-shoot sample prep’s high-throughput capabilities while helping to keep particulates out of your instrument and column. This reduces instrument downtime and prolongs column lifetime so it’s a win-win for your lab’s productivity. You can also double up on column protection with good sample prep and the use of a guard column, especially if the sample has fine particles that can pass through a filter membrane or not form a good solid pellet. Here is a great starting point for guard column selection.

Thanks for reading!

 

 

 

 

 

 

 

 

 

EXP Fittings: Which Nut Do I Need?

Our brilliant friends at Optimize Technologies are continually innovating and making our lab lives easier. We love their patented EXP Titanium Hybrid Ferrule technology that is truly universal. You can use it on any tubing, any 10-32-port, any instrument—wrench-tight to 20,000 psi. This fully UHPLC-capable EXP ferrule is used with all EXP nuts. Restek offers three EXP nuts: hand-tight, hex-head, and EXP2.

The question is, which nut do I need? I’ve tried them all and found that it is truly situational. Ask yourself: how much space do I have, how often will I make-and-break the connection, and am I connecting PEEK or stainless steel (SS) tubing? Here are two examples.

The EXP hand-tight nut is ideal for column connections, especially on systems operating </=600 bar. Most column ovens in HPLC or intermediate-HPLC instruments are large enough to allow the hand-tight nut. Its larger and more-ergonomic head allows easy hand-tightening for repeated connections up to 8,700 psi. No wrenches, just fingers. While you certainly can wrench-tighten the hand-tight nut to provide leak-tight seals up to 20,000psi, the EXP2 nut is perfect for UHPLC applications. The torque driver allows a wrench-tight seal that’s as easy as finger-tight.

Alternatively, in the super-tight space of a 6-port injection valve, the EXP hand-tight nut won’t fit. The EXP2 nut is ideal in this situation. Its slim profile and torque driver allow flawless, fast, zero-dead-volume connections in places that traditional wrenches don’t fit.

If you’re using PEEK tubing, move to a hex-head nut. EXP2 fittings aren’t recommended for use with PEEK tubing. This is because the torque driver that’s included with the fittings is so effective, that’s it’s easy to crush your tubing. So if you’re using PEEK tubing and have minimal access or space, use the EXP hex-head fittings. If you just can’t fit a traditional wrench in there, try using the socket side of the ValvTool Wrench (find it here http://www.restek.com/catalog/view/960 ). It’s slotted to fit over the tubing and slimmer than traditional wrenches.

Also, consider staggering the height of the nuts around your 6-port injection valve. EXP Hex-Head fittings come in three lengths: short, standard, and long. Varying the height allows breathing room for your wrench. An example of that is shown below.

Most analysts who try EXP fittings keep using EXP fittings, because they’re truly universal, extremely well-designed and durable, and easy. Please let us know what you think of them!

A better way to configure your EZ No-Vent GC-MS Connector

If you are using the EZ No-Vent for Agilent mass spectrometers (cat.# 21323), you are probably tricking your instrument into working properly by inflating the length of your column in your acquisition software. If you have MassHunter, or a recent version of MSD ChemStation  (G1701EA), you don’t have to do this. Instead of configuring your 30 m x 0.25 mm ID column as a 112 m column, you should configure a composite column (see Figure 1). Even if you aren’t using the EZ No-Vent with your mass spectrometer, you should be configuring your column as a composite column because the 17 cm transfer line is heated by the transfer line heater, not the oven.

Figure 1 – 30 m column configured as a composite column in MassHunter (without the EZ No-Vent)

Figure 2 shows a composite column configured with the 17 cm x 0.10 mm ID vacuum restrictor, which is the part of the EZ No-Vent that allows you to disconnect your analytical column without turning off your MSD. You’ll notice the average linear velocity is significantly reduced when the EZ No-Vent is used (39 cm/sec drops to 26.8 cm/sec) even though the volumetric flow stays the same. This has some negative chromatographic effects.

Figure 2 – 30 m column configured as a composite column in MassHunter (with the EZ No-Vent)

Figure 3 is an example chromatogram collected on a 30 m x 0.25 mm x 0.25 µm column without using the EZ No-Vent. Figure 4 shows the chromatographic effect of installing the EZ No-Vent while keeping all acquisition parameters the same (aside from configuring the restrictor in the composite column).

Figure 3 – Example Chromatogram collected without the EZ No-Vent

Figure 4 – Same acquisition parameters as Figure 3, but collected with the EZ No-Vent installed

The good resolution of the first eluting compound (1,4-Dioxane) and the dichloromethane solvent peak has collapsed, in Figure 4, and Aniline and bis(2-Chloroethyl)ether are now coeluting. The resolution for the two PAH pairs has also degraded. Lowering starting oven temperature 20°C improves the separation of 1,4-dioxane and solvent peak, though 1,4-dioxane is still on the solvent peak’s tail (see Figure 5). The PAH resolutions have also improved, almost to the level of performance when the EZ No-Vent isn’t installed.

Figure 5 – Chromatography is improved by reducing the starting oven temperature.

 

The fix to the peak tailing can be found in the follow-up blog: A better way to configure your EZ No-Vent GC-MS Connector Part II

355 compounds have been added to the Rxi-XLB library for the Pro EZGC Chromatogram Modeler

The Rxi-XLB is a popular column for the analysis of PAHs and persistent environmental pollutants (POPs) such as PCBs and Pesticides. We’ve updated the library of 209 PCB congeners, expanded the PAH library to 47 compounds (including 6 deuterated isotopologues commonly used as internal standards or surrogates), and added the 203 compounds in the GC Multiresidue Pesticide Kit, 35 of the 75 polychlorinated naphthalene congeners (PCNs), and 43 phthalates.

Click here to explore the Pro EZGC Chromatogram Modeler. A free website account is required.

 

How do I Replace my LC Tubing?

For those that are experienced with LC, you probably have this all figured out.  If that is the case, this might serve as a good introduction to Restek products that you can use for this.  For those that are newbies to liquid chromatography, welcome and I hope you find this useful!

The best way to replace tubing in an LC system largely depends on where the tubing is located, how it is used and the material it is made from.  In all cases, tubing must be cut straight and square on the end to achieve a good seal.  This is even more critical when you are operating at higher pressure ranges.

PEEK (polyether ether ketone) tubing is used for lower pressure applications.  We recommend using PEEK tubing with ID of ≤0.007 inches for applications up to 7,000 psi and the tubing with an ID of ≥0.010 inches for applications that go up to 5,000 psi.  Our product listing for PEEK tubing is located at this link (click here).  These are color coded with a stripe so you can tell what the ID (internal diameter) is.  Here are the codes:

Clear= 0.0025”

Red= 0.005”

Yellow=0.007”

Blue= 0.010”

Orange= 0.020”

 

We suggest to use our Clean-Cut Tubing Cutter catalog #25069 to cut PEEK (and other plastic) tubing. Here’s what it looks like:

 

Stainless steel tubing can be used at just about any pressure range, including the higher ranges for UHPLC.  It is also considered more permanent, so most internal LC connections are usually stainless steel. Although most durable, stainless steel is harder to work with, in general.  I mentioned earlier that it is critical to have tubing cut straight and square on the ends to achieve a good seal.  This is very true with stainless steel, especially since it is used at higher pressure ranges.  It is also more difficult to ensure the tubing is smooth and very straight on the end, so for these reasons, we do not recommend for customers to cut their own stainless steel tubing for LC.  It is much better to order the tubing that is already professionally cut to the desired length.  We offer stainless steel tubing in pre-cut lengths of 5, 10, 20 and 30 cm and ID sizes ranging from 0.005” – 0.020”.  These are shown at this link (click here) on our website.  We use the same color code for these as we do for the PEEK.  Instead of a stripe along the tubing, there is narrow band around the outside of the tubing.

 

While it is nice to have generic lengths of tubing available with no fittings attached, you may be looking for something more specific.  In some cases, you may need to obtain tubing with pre-swaged end fittings attached or you may be looking for specific dimensions to fit a certain location for replacement within your instrument.  Some instrument parts may also contain a specialized end fitting to serve a certain purpose that is specific for that instrument model.  This is why we often suggest that customers replace the entire tubing assembly that comes with the proper fittings, as specified by the instrument manufacturer.  The only reliable way to do this is to identify the manufacturer’s catalog or part number and seek a direct replacement for that.  Fortunately, for our customers using Agilent LC models, we do sell a few of the more popular tubing assemblies to fit Agilent LC models, located here on our website:   Capillary Stainless Steel Tubing Assemblies

If you select an individual item from the link above on our website, you can see which Agilent part number the item corresponds to.  Feel free also to contact our Tech Services department to see if we have a replacement, if you know the part number and cannot find it.

To help confirm your selection, I have listed the primary location of usage and dimensions for each of these tubing assemblies in the table below.  Please note that some of these may be used for general purposes in other instrument models also, but the location I have specified here in the indicated model does require these particular dimensions.  Generally for most Agilent HPLCs, the tubing used in the system flowpath between the solvent module and the autosampler sample injection valve has an ID of 0.25 mm.  Likewise, the tubing downstream from the sample injection valve to the detector has an ID of 0.17 mm or smaller to minimize band spreading.

 

Here are some links for additional reading on related topics:

How do I know what the internal diameter of my LC tubing is?

Which fitting do I use for which tubing?

How are stainless steel fittings for HPLC different from polymer-based fittings?

Finding the right fitting for HPLC or UHPLC

How can I use Raptor Columns and the EXP Direct Connect holder with stainless steel fittings?

Need parts for your Agilent 1100? HP 1050? Restek has you covered.

Also, be sure to check out our Youtube video “LC Tubing and Fitting Choices”.

I hope you found this post helpful.  Thank you for reading!

 

 

Optimizing Mass on Column to Balance Sensitivity Requirements and Calibration Range with Split Injection

This is a continuation of the EPA Method 8270 blog series started in January of 2016. Previous posts: 1, 2, 3, 4, and 5.

We’ve been focusing on the advantages of split injection analysis, while highlighting the weaknesses of splitless injection. This blog is going to revisit the topic of column overload in more detail – focusing on optimizing a split injection method to maximize sensitivity while maintaining and extended calibration range. This is especially critical if you are migrating your method to a 20 m x 0.15 mm x 0.15 µm or 20 m x 0.18 mm x 0.18 µm column for fast analysis with the new GC Acclerator.

Typical semivolatiles calibrations on a 30 m x 0.25 mm x 0.25 µm 5-type column range from 0.5 µg/mL or 1.0 µg/mL to over 100 µg/mL (many analysts target 160 or 200 µg/mL). However, this column dimension (as well as the 0.32 mm ID format) will usually show signs of peak overload with less than 10 ng of any individual component on column. Isobars that elute close together quickly become coelutions as mass on column increases. Figure 1 highlights the most popular example of this, the benzo[b]fluoranthene and benzo[k]fluoranthene isomeric pair. The three highest concentration calibration standards (120, 80, and 40 µg/mL) do not meet the 50% valley resolution criteria under splitless conditions [technically, the resolution criteria are only evaluated at the midpoint used for the CCV evaluation, but they are good indicators of accurate integration potential]. The extreme peak fronting resulting from column overload makes it impossible to accurately integrate and generate a linear calibration including these points. Additionally, the peak apex of benzo[b]fluoranthene shifts more than 0.2 minutes (12 seconds) across the calibration standards, another symptom of severe column overload.

Figure 1 – Benzo Fluoranthene isomer resolution ( B and K ) at increasing mass on column

The concentration-dependent retention time shift requires wide windows in the data processing software, increasing the likelihood of identification errors when the automated integrator processes the data. This results in more time required for manual data review and integration [which can be a headache for those of you manually recording before and after chromatograms in compliance with your manual integration policies] with an elevated risk of error.

Under split conditions, the isomers meet resolution criteria (50% valley) in each of the 9 calibration standards, and the peak apices shift by at most 0.04 minutes (2.4 seconds) from 0.1 µg/mL to 120 µg/mL, indicating only minor peak overload at the high end of the calibration range. Figure 2 shows a comparison of splitless and split benzo fluoranthene isomer chromatography over the same calibration range.

Figure 2 – Comparison of benzo fluoranthene isomer retention time variation as injection concentration increases using splitless (top) and split 10:1 (bottom) injection techniques

The minimal shift of concentration-dependent retention times allows for a much more narrow integration window and a greater confidence in peak IDs. This is important because compound concentration isn’t the only factor that can cause retention time shifts. Complex sample matrices with an excess of co-extracted material can have the same effect on splitless injection by greatly broadening the initial sample band at the head of the column. Split injection minimizes this by transferring a fraction of the injection to the head of the column.

Figure 2 is a good illustration of how column overload is managed by split injection – but it introduces a new problem that needs to be dealt with: detector overload. Initially, when collecting the splitless injection data, we dropped the gain factor to 0.3 (from the default of 1.0), which reduces the voltage applied to the multiplier below the tune optimized level. This reduces instrument sensitivity, preventing detector overload at the high end of the calibration curve (120 ng on column). For the 10:1 split injection, we increased the gain factor to 3.0 (adding almost 250 volts to the tune optimized level) to make sure sensitivity wasn’t an issue at the low end of the calibration because the split injection delivers 1/11th of the sample to the analytical column. This was a gross over-correction, causing compounds with strong molecular ion responses (such as PAHs) to overload the detector at concentrations as low as 40 µg/mL. Through trial and error, we determined that the best balance between low-end sensitivity and high-end overload occurred at a gain factor of 0.8 (see Figure 3). This appears to be instrument dependent, as we operate a similarly configured 5977a with an optimized gain factor of 1.0 and see similar performance.

Figure 3 – Comparison of benzo fluoranthene isomer peak shapes demonstrating the effect of reducing the gain factor on detector overload.

Once you have established your linear mass on column range, you can adjust standard concentrations and the split ratio (or injection volume) to maximize your calibration range while maintaining sensitivity to meet method required LOQs.

Cannabis Concentrates Part II: We’re Heading to Space!

Welcome back! It’s been a little while since my last blog, but in that time, we’ve been doing some interesting things regarding cannabis research. Last time, I discussed that we would be analyzing residual solvents in cannabis concentrates and today, I’m going to show one of the methods that we’ve been working on. If you missed the previous blog, be sure to check it out here!

Before we get into the sample preparation techniques that we’ve been focusing on, let’s go over some initial things. First, we need to select a column that has the capability of resolving these light weight, volatile compounds (see Table 1). An easy way to do this is through Restek’s Pro EZGC Chromatogram Modeler. After placing our compound list into EZGC, we saw that the program gave us numerous selections with the first column being the Rtx-502.2. However, since our future work includes terpene analysis, we decided to go with the Rxi-624Sil MS, 30 m x 0.25 mm x 1.40 µm (cat. # 13868). This column has better high temperature stability and it also lines up perfectly with what Amanda chose for her method using Full Evaporation Technique (FET).

Now that we have our column selected, we can move on to the main event! When thinking about how to approach this problem (analyzing residual solvents) we started out by using Jason Herrington’s favorite method: KISS. For those who haven’t heard of KISS, it stands for Keep It Simple, Stupid (sorry, no Gene Simmons here). We wanted to keep everything as simple as possible to make your life in the lab as stress free as possible. To do that, we thought that the easiest approach to this analysis would be through headspace (HS). Headspace samples are prepared by taking a sample and adding it to a dilution solvent in a HS vial (we’re using a 20 mL vial). Depending on the analytes of interest, an inorganic salt (Sodium Chloride, Sodium Sulfate, etc.) can be added to the vial as well to help lower the partitioning coefficient of the more polar compounds. For more information on HS analysis, please check out Restek’s Headspace Technical Guide!

In our current method, which we will call HS-Syringe-GC-MS, we are taking the headspace from a 6 mL sample. All standards (STDs) were prepared as follows: 3 g sodium chloride (NaCl) was measured into a 20 mL amber headspace vial (cat. # 23086) with screw top cap (cat. # 23090). 6 mL of deionized (DI) water was then added to the vial. All STDs then received internal standards. Lastly, STDs were capped and vortexed at 3000 rpm for 10 seconds, inverted, then vortexed again for 10 seconds at 3000 rpm. Luckily for us, we are using the CTC PAL RTC rail system, so the vial equilibration and injection are automated. Once equilibrated, 500 µL of the headspace is drawn into a gastight syringe and injected into the gas chromatograph.

 

*Note: If you are not using a rail system, don’t fret! This method can easily be done with a manual setup.

 

For more detailed specs, please refer to Table 2 & 3, which has our rail and GC-MS parameters.

 

Now that we’ve gone over how we are preparing our samples, let’s take a look at the chromatography (Figures 1 & 2):

 

 

This total ion chromatogram (TIC, Figure 1) shows that we are able to resolve all 19 compounds of interest. Some of the polar compounds that have higher partitioning coefficients are a little more difficult to see. However, in the extracted ion chromatogram (EIC, Figure 2), you can see that we were in-fact able to drive them out of the water and into the headspace. The only compounds that are not baseline resolved are the Toluene-d8 and Toluene. The resolution value for these two compounds is 1.38, which is resolved enough for accurate integration. Since Toluene-d8 is our internal standard, we’ll probably be changing our internal standard in the near future, so that Toluene is completely baseline resolved.

 

 

I know we threw a lot at you here, so I think this is a good place to take a break! We do have more to show you, but we don’t want to give it to you all at once. We want to leave you hungry for more! To recap, we were able to take a very simple approach to this analysis by using the HS-Syringe technique and with the help of EZGC, we were able to resolve all of our compounds with hardly any difficulty. In addition, this method can also be done with a manual HS-Syringe and run on a GC-FID. Like I said, we have much more to cover, so stay tuned for our other HS approach in our next blog!

Alicat Flow Meters for Your TO-11/13/15/17 (Anything Air) Laboratory: Let’s Talk About Errors and Ranges

If you already own an Alicat flow meter, then I will soon be preaching to the choir. However, if you do not have an Alicat flow meter and you work in a toxics organics (TO) air laboratory, whether it be U.S. EPA Method TO-11A or TO-17, etc… then you do not know what you are missing out on. Short story: instantaneous pressure and vacuum flow measurement you can fit in your back pocket. ‘nuff said! However, if you are currently in the market for one or soon to be after that sales pitch I just laid on you, then read on…

We offer 2 different flow ranges for the Alicat: low flow (0-50 mL/min) and high flow (0-500 mL/min). Sometimes we hear the question “can I use only the high flow meter to set the flow on my Passive Air Sampling Kit at 0.5 mL/min and 250 mL/min or do I need to have the low flow meter as well? After all, the high flow meter does have a range of 0 – 500 mL/min?”

Here is my answer: Both meters will work, but there will be a lot more error associated with the high flow meter when measuring down to 0.5 mL/min. I say this, because the flow meter accuracy is defined as: ± (0.8% of Reading + 0.2% of Full Scale). The following table shows you 3 possible scenarios that could play out with the two different flow meters when trying to measure flow.


For scenario 1 we see how using a low flow (0 – 50 mL/min) meter to measure an actual flow of 1 mL/min could result in a potential reading ranging from 0.892 – 1.108 mL/min, which is perfectly acceptable for most people. However, for scenario 2 we see how the errors add up on the high flow (0 – 500 mL/min) meter when trying to measure 1 mL/min. In fact, you could be reading 2 mL/min when in reality it is 1 mL/min. That is not to say you will, as these errors/ranges are the maximum (i.e., worst case scenarios). You could also read 1 mL/min for 1 mL/min. The point is, you will not have the confidence in the low flow measurements (e.g., 1 mL/min) when using the high flow meter. However, you can be confident with the low flow meter. Scenario 3 is here to demonstrate: 1) that you may not use a low flow meter to measure 100 mL/min, as it is above the meter’s operational range and 2) to show you that the high flow meter is accurate when applied appropriately.

Sorry for the long-winded answer. Since I love to work in analogies, my shortcut/analogy response is often as follows: would you make a 1 µL injection on your 500 µL syringe or do you own several syringes? Again, ‘nuff said! So, the short answer is that most of us have to use both flow meters.